Advances in Microbial Physiology Volume 40

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Advances in

MICROBIAL PHYSIOLOGY VOLUME 40

This Page Intentionally Left Blank

Advances in

MICROBIAL PHYSIOLOGY Edited by

R. K. POOLE Department of Molecular Biology and Biotechnology The Krebs Institute for Biomolecular Research The University of Sheffield Firth Court, Western Bank ShefJield SIO 2TN, UK

Volume 40

ACADEMIC PRESS San Diego London Boston New York Sydney Tokyo Toronto

This book is printed on acid-free paper. Copyright

0 1998 by ACADEMIC PRESS

All Rights Reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the publisher. Academic Press 525 B Street, Suite 1900, San Diego, California 92101-4495, USA http://www.apnet.com Academic Press 24-28 Oval Road, London N W l 7DX, UK

ISBN 0-12-027740-9 A catalogue record for this book is available from the British Library

Typeset by Keyword Typesetting Services Ltd, Wallington, UK Printed in Great Britain by MPG Books Ltd, Bodmin, Cornwall 98 99 00 01 02 03 M P 9 8 7 6 5 4 3 2 1

Contents

CONTRIBUTORS TO VOLUME 4 0 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ix

The Biochemistry, Physiology and Genetics of PQQ and PQQ-containing Enzymes Pat M. Goodwin and Chris Anthony 1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 2. PQQ in bacteria. . . . , . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6 3. The quinoprotein dehydrogenases containing PQQ , . . . . . . . . . . . . 7 4. The importance of divalent metal ions in the structure and function of PQQ-containing quinoproteins . . . . . . . . . . . , . . . . . . 20 5. The structure and mechanism of PQQ-containing quinoproteins . . 26 6. Quinoproteins in energy transduction . . . . . . . . . . . . . . . . . . . . , 35 7. The physiological functions of the quinoprotein dehydrogenases . . 42 8. Synthesis of PQQ . . . . . . . . . . . . . . . . . . . . . . . . . . . . , . . . . . . 5 1 9. Regulation of synthesis of PQQ and quinoprotein dehydrogenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 9 10. Concluding remarks. . . . . . . . . . . . . . . . . . , . . . . . . . . , . . . . . . 6 6 Acknowledgements . . . . . . . . . . . . . . . . . , . . . . . . . . . . . . . . . . 6 7 References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . , . . . . . . 6 7

Molecular Phylogeny as a Basis for the Classification of Transport Proteins from Bacteria, Archaea and Eukarya Milton H. Saier, Jr 1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8 3 2. Considerations for the systematic classification of transmembrane solute permeases. . . . . , . . . . . . . . . . . . . . . . . . . 84

vi

CONTENTS

3. Proposed transport protein classification system . . . . . . . . . . . . . . 86 4 . Diverse evolutionary origins of integral membrane transport protein families . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 95 5 . The major facilitator superfamily (MFS) . . . . . . . . . . . . . . . . . .107 6. The ATP-binding cassette (ABC) superfamily. . . . . . . . . . . . . . . 109 7. Prokaryotic genome sequence analyses. . . . . . . . . . . . . . . . . . . . 121 8. Independent evolution of distinct transport modes and energy-coupling mechanisms . . . . . . . . . . . . . . . . . . . . . . . . . . . 124 9 . Proposed independent evolution of different channel and carrier 127 families . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 131 10. Conclusions and perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgement . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 131 Note added in proof . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 131 References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 131

The Physiology and Metabolism of the Human Gastric Pathogen Helicobacier pylori David J . Kelly 1 . Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

2. 3. 4. 5. 6.

7. 8. 9.

139 Helicobacter pylori as a gastric pathogen . . . . . . . . . . . . . . . . . . 140 Characteristics of Helicobacter pylori . . . . . . . . . . . . . . . . . . . . . 144 Solute transport. ion movements and acid tolerance in H . pylori . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 149 The microaerophilic nature of H . pylori . . . . . . . . . . . . . . . . . . . 152 Current knowledge of H . pylori carbon metabolism and substrate utilization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 155 The respiratory chain of H. pylori . . . . . . . . . . . . . . . . . . . . . . . 169 Nitrogen metabolism in H . pylori . . . . . . . . . . . . . . . . . . . . . . . 176 Concluding remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 179 Note added in proof . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 180 Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 180 References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 180

Genes Involved in the Formation and Assembly of Rhizobial Cytochromes and their Role in Symbiotic Nitrogen Fixation Maria J . Delgado. Eulogio J . Bedmar and J . Allan Downie 1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 . Haem-copper respiratory oxidases . . . . . . . . . . . . . . . . . . . . . . . 3. Respiratory chains of free-living rhizobia . . . . . . . . . . . . . . . .

193 195 . . 198

vii

CONTENTS

4. Other terminal oxidases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5. 6. 7. 8.

205 209 Symbiosis-specific cytochromes . . . . . . . . . . . . . . . . . . . . . . . . . OtherfixNOQP genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 214 Genes involved in cytochrome c biogenesis . . . . . . . . . . . . . . . . 218 Rhizobial mutants with altered oxidase activity and improved symbiotic nitrogen fixation . . . . . . . . . . . . . . . . . . . . . . . . . . . . 221 Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 222 References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 222

The Starvation-Stress Response (SSR) of Salmonella Michael P . Spector 1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 235 237 2. The starvation-stress response (SSR) . . . . . . . . . . . . . . . . . . . . . 3. The SSR and long-term starvation survival . . . . . . . . . . . . . . . . 264 4 . The SSR and resistance to other environmental stresses . . . . . . . 266 271 5. The SSR and Salmonella virulence . . . . . . . . . . . . . . . . . . . . . . 272 6. Concluding remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 273 273 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Iron Storage in Bacteria Simon C. Andrews 1 . Biologically relevant features of iron . . . . . . . . . . . . . . . . . . . . . 283 2. Ferritins. rubrerythrins and bacterioferritins . . . . . . . . . . . . . . . . 288 3. Primary structures and evolution of iron-storage proteins . . . . . . 305 4 . Structures of bacterioferritin and bacterial ferritin . . . . . . . . . . . 316 5 . Core formation and the iron core . . . . . . . . . . . . . . . . . . . . . . . 323 329 6. Bacterioferritin-associatedferredoxin . . . . . . . . . . . . . . . . . . . . . 7. Intracellular iron metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . 333 8. Perspective . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 339 Note added in proof . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 340 Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 340 References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 341

How Did Bacteria Come to Be? Arthur L . Koch 1. Introduction

...................................... 2 . Evolution of Domains - a scenario . . . . . . . . . . . . . . . . . . . . . .

355 356

viii

CONTENTS

3. 4. 5. 6. 7.

Bacterial wall formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 367 The function of the bacterial wall - non-growth aspects . . . . . . . 374 The function of the bacterial wall - growth aspects. . . . . . . . . . . 382 The wall of the first bacterium . . . . . . . . . . . . . . . . . . . . . . . . . 388 Conclusions.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 393 395 Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 395 Energetics of Alkaliphilic Bacillus Species: Physiology and Molecules Terry Ann Krulwich. Masahiro Ito. Raymond Gilmour. David B . Hicks and Arthur A . Guffanti

1 . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 402 2 . Energetics of pH homeostasis . . . . . . . . . . . . . . . . . . . . . . . . . . 404 3. Energetics of oxidative phosphorylation . . . . . . . . . . . . . . . . . . . 420 Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 432 432 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Author Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Subject Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

439 465

Contributors to Volume 40

Simon C. ANDREWS, School of Animal and Microbial Sciences, The University of Reading, Whiteknights, PO Box 228, Reading RG6 6AH. UK Chris ANTHONY, Division of Biochemistry and Molecular Biology, School of Biological Sciences, University of Southampton, Southampton SO16 7PX, UK ([email protected]) Eulogio J. BEDMAR, Departamento de Microbiologia del Suelo y Sistemas Simbioticos, Estacion Experimental del Zaidin, CSIC, PO Box 419, 1808O-Granada, Spain ([email protected]) Departamento de Microbiologia del Suelo y Sistemas Maria J. DELGADO, Simbiuticos, Estacion Experimental del Zaidin, CSIC, PO Box 419, 1808O-Granada, Spain ([email protected])

J. Allan DOWNIE, Department of Genetics, John Innes Centre, Norwich Research Park, Colney, Norwich NR4 7UH, UK (allan.downie(ujbbsrc.ac.uk) Department of Biochemistry, Mount Sinai School Raymond GILMOUR, of Medicine of CUNY, 1 Gustave L. Levy Place, New York, New York 10029, USA Pat M. GOODWIN, Division of Biochemistry and Molecular Biology, School of Biological Sciences, University of Southampton, Southampton SO 16 7PX, UK (p. goodwin (4wellcome.ac.uk)

X

CONTRIBUTORS TO VOLUME 40

Arthur A. GUFFANTI. Department of Biochemistry, Mount Sinai School of Medicine of CUNY, 1 Gustave L. Levy Place, New York, New York 10029, USA David B. HICKS,Department of Biochemistry, Mount Sinai School of Medicine of CUNY, 1 Gustave L. Levy Place, New York, New York 10029, USA Masahiro ITO. Department of Life Sciences, Toyo University, Gunma 374-0 1 , Japan David J. KELLY,Department of Molecular Biology and Biotechnology, University of Sheffield, Western Bank, Sheffield S 10 2TN, UK (d.kelly (2jsheffield.ac.uk) Arthur L. KOCH,Department of Biology, Indiana University, Bloomington, Indiana 47405-6801, USA ([email protected]) Terry Ann KRULWICH, Box 1020, Department of Biochemistry, Mount Sinai School of Medicine of CUNY, 1 Gustave L. Levy Place, New York, New York 10029, USA ([email protected]) Milton H. SAIER, Jr, Department of Biology, University of California at San Diego, La Jolla, CA 92093-01 16, USA (msaier(aiucsd.edu) Department of Biomedical Sciences, University of Michael P. SPECTOR, South Alabama, Mobile, Alabama 36688, USA (mspector(QJusamail. usouthal.edu)

The Biochemistry, Physiology and Genetics of PQQ and PQQ-containing Enzymes Pat M. Goodwin and Chris Anthony Division of Biochemistry and Molecular Biology, School of Biological Sciences, University of Southampton, Southampton SO16 7 P X , U K

ABSTRACT

Pyrrolo-quinoline quinone (PQQ) is the non-covalently bound prosthetic group of many quinoproteins catalysing reactions in the periplasm of Gram-negative bacteria. Most of these involve the oxidation of alcohols or aldose sugars. PQQ is formed by fusion of glutamate and tyrosine, but details of the biosynthetic pathway are not known; a polypeptide precursor in the cytoplasm is probably involved, the completed PQQ being transported into the periplasm. In addition to the soluble methanol dehydrogenase of methylotrophs, there are three classes of alcohol dehydrogenases; type I is similar to methanol dehydrogenase; type I1 is a soluble quinohaemoprotein, having a C-terminal extension containing haem C ; type I11 is similar but it has two additional subunits (one of which is a multihaem cytochrome c), bound in an unusual way to the periplasmic membrane. There are two types of glucose dehydrogenase; one is an atypical soluble quinoprotein which is probably not involved in energy transduction. The more widely distributed glucose dehydrogenases are integral membrane proteins, bound to the membrane by transmembrane helices at the N-terminus. The structures of the catalytic domains of type 111 alcohol dehydrogenase and membrane glucose dehydrogenase have been modelled successfully on the methanol dehydrogenase structure (determined by X-ray crystallography). Their mechanisms are likely to be similar in many ways and probably always involve a calcium ion (or other divalent cation) at the active site. The electron ADVANCES IN MICROBIAL PHYSIOLOGY VOL 40

ISBN 0-12-027740-9

Copyright 0 1998 Academic Press All rights of reproduction in any form reserved

2

PAT M. GOODWIN AND CHRIS ANTHONY

transport chains involving the soluble alcohol dehydrogenases usually consist only of soluble c-type cytochromes and the appropriate terminal oxidases. The membrane-bound quinohaemoprotein alcohol dehydrogenases pass electrons to membrane ubiquinone which is then oxidized directly by ubiquinol oxidases. The electron acceptor for membrane glucose dehydrogenase is ubiquinone which is subsequently oxidized directly by ubiquinol oxidases or by electron transfer chains involving cytochrome hc,, cytochrome c and cytochrome c oxidases. The function of most of these systems is to produce energy for growth on alcohol or aldose substrates, but there is some debate about the function of glucose dehydrogenases in those bacteria which contain one or more alternative pathways for glucose utilization. Synthesis of the quinoprotein respiratory systems requires production of PQQ, haem and the dehydrogenase subunits, transport of these into the periplasm, and incorporation together with divalent cations, into active quinoproteins and quinohaemoproteins. Six genes required for regulation of synthesis of methanol dehydrogenase have been identified in Merhjdohacterium, and there is evidence that two, twocomponent regulatory systems are involved. 1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 6 2. PQQ in bacteria.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Identification of PQQ as the prosthetic group of quinoproteins . . . . . 2.2. The effect of exogenous PQQ on bacterial growth. . . . . . . . . . . . . . . 3. The quinoprotein dehydrogenases containing PQQ. . . . . . . . . . . . . . . . . . . . . . . 7 3.1. Soluble quinoprotein alcohol dehydrogenases . . . . . . . . . . . . . . . . . . . . . 10 3.2. Membrane-associated quinohaemoprotein alcohol dehydrogenases 13 (type 111 alcohol dehydrogenases). . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3. Glucose dehydrogenases. . . . . . . . . . .. . . . . . . . . . . . . . . . . 16 . . . . . . . . . . . . . . . . . . . . . . . . 19 3.4. Aldehyde dehydrogenases. . . . . . . . . 4. The importance of divalent metal ions in the structure and function of 20 PQQ-containing quinoproteins. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1. Calcium in methanol dehydrogenase and other alcohol dehydrogenases . 20 4.2. The role of divalent metal ions in glucose dehydrogenase . . . . . . . . . . . . 25 5. The structure and mechanism of PQQ-containing quinoproteins. . . . . 5.1. The structure and mechanism of methanol dehydrogenase . . . . 5.2. The structures and mechanisms of alcohol and glucose dehydrogenases. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

5.3. The conserved sequence that was wrongly identified as the

........................ ........................

PQQ-binding domain in quinoproteins

6. Quinoproteins in energy transduction

6.1. Electron transport chains involving soluble alcohol dehydrogenases

....

35 35 37

6.2. Electron transport chains involving membrane-bound quinohaemoprotein alcohol dehydrogenase (type 111) in acetic acid bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

40

PQQ AND QUINOPROTEINS

7.

8.

9.

10.

3

6.3. Electron transport chains involving membrane-bound glucose dehydrogenase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 41 The physiological functions of the quinoprotein dehydrogenases. . . . . . . . . . . 42 7.1. The role of periplasmic quinoproteins that oxidize alcohols . . . . . . . . . . . 43 7.2. The roles of glucose dehydrogenase in Acinetobacter, pseudomonads and enteric bacteria. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 44 7.3. The roles of alcohol and glucose dehydrogenases in the membranes of acetic acid bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 50 SynthesisofPQQ . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 51 8.1. Origin of PQQ backbone . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 51 8.2. The genetics of PQQ biosynthesis ............................... 52 8.3. Does Escherichia coli contain pqq genes?. ......................... 57 Regulation of synthesis of PQQ and quinoprotein dehydrogenases . . . . . . . . . 59 9.1. Synthesis of PQQ and apoenzymes is not coordinated . . . . . . . . . . . . . . .59 9.2. Regulation of PQQ synthesis.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 59 9.3. Factors affecting the synthesis of the quinoprotein dehydrogenases. . . . . 60 66 Concluding remarks. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 67 67 References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1. INTRODUCTION

Pyrrolo-quinoline quinone (PQQ) (Fig. 1) is the prosthetic group of most of the quinoprotein dehydrogenases that catalyse reactions in the periplasm of Gram-negative bacteria. It is synthesized independently of the apoenzyme dehydrogenase and transported to the periplasm where it is incorporated into the structure of the enzyme, forming active holoenzyme. It is always non-covalently bound to the enzyme. This contrasts with the prosthetic groups of other types of bacterial quinoprotein; the amine dehydrogenases which contain tryptophan-tryptophylquinoneand the amine oxidases which contain topa quinone (Fig. 1). A fourth type of quinoprotein is the mammalian lysyl oxidase which contains lysine tyrosylquinone. In all quinoproteins except for those containing PQQ the prosthetic groups are covalently bonded to the enzyme, being derived by modification of tryptophan, tyrosine or lysine residues in the amino acid backbone of the protein (Anthony, 1996, 1998). PQQ was first isolated from glucose and methanol dehydrogenases (Hauge, 1964; Anthony and Zatman, 1967). It was readily released by denaturation of the enzymes, purified, and shown to be a red, highly polar, acidic compound with a very characteristic green fluorescence. At that time it was concluded that it might be a novel flavin or pteridine derivative. Its structure was subsequently elucidated by X-ray crystallography (Salisbury et al., 1979) and its chemistry described in detail by Frank and Duine and their co-workers (Duine et al., 1987; Duine, 1991). They showed that a key

4

PAT M. GOODWIN A N D CHRIS ANTHONY

PQQ

TPQ

TTQ

Figure I The prosthetic groups of quinoproteins. PQQ (pyrroloquinoline quinone) is the prosthetic group of the dehydrogenases discussed in this review. TTQ (tryptophan tryptophylquinone) is the prosthetic group of amine dehydrogenases. TPQ (6-hydroxyphenylalanine or topa quinone) is the prosthetic group of the copper-containing amine oxidases.

feature of the structure of PQQ is the ortho quinone at the C4 and C5 positions of the quinoline ring, which becomes reduced to the quinol during catalysis. The C5 carbonyl in the oxidized form is very reactive towards nucleophiles such as alcohols, ammonia, amines, cyanide and amino acids (Fig. 2) and this reactivity must be taken into account when interpreting mechanisms of enzyme action, when devising assay systems and when measuring PQQ in complex growth media. The midpoint redox potential at pH 7 of the isolated PQQ is about +90 mV but this is likely to be influenced by its environment in the dehydrogenases. An important feature of PQQ, only appreciated more recently, is its ability to complex Ca2+ or Mg2+ in solution. This was first shown by Mutzel and Gorisch (1991) and exploited by Itoh e l al. (1997) in their chetnical model systems in which a Ca2+-PQQ complex is able to catalyse the oxidation of alcohols in organic solvents. It is now generally appreciated that the PQQ-containing enzymes probably all contain a divalent metal ion in their active sites (Section 4). Although PQQ is not covalently bound to the apoenzyme dehydrogenases and can be readily dissociated from some holoenzymes, this dissociation is never important for its metabolic function. In this way PQQ is like FMN and FAD in flavoprotein dehydrogenases and, like them, quinoprotein dehydrogenases are most conveniently assayed by using artificial electron acceptors such as phenazine ethosulphate. This review discusses the structure and function of the PQQ-containing dehydrogenases, the electron transport systems of which they form a part,

COOH

H

HOOC

d

o

o

H

5

H@OoH

HOOC

0

'

OH OH

PQQ

NH

PQQH2

COOH

ammonia adduct (Schiff base) GOOH H&OOH

HOOC

HOOC

HOOC

HO OR

ammonia adduct (aminocarbinoI)

alcohol adduct (hemiketal)

HO CN

cyanide adduct

Figure 2 Adducts of PQQ. The ease of formation of these adducts has considerable importance in discussion of mechanisms, determination of the prosthetic group, and measurements of PQQ in biological samples.

6

PAT M. GOODWIN AND CHRIS ANTHONY

their physiological functions and the genetics and regulation of synthesis of the dehydrogenases and of PQQ itself.

2. P a a IN BACTERIA 2.1. Identification of PQQ as the Prosthetic Group of Quinoproteins

Early in the history of the discovery of quinoproteins a number of enzymes were erroneously claimed to contain covalently bound PQQ. It is now accepted that there are no known examples of such quinoproteins and it has been shown that enzymes previously thought to have covalently bound PQQ contain the other types of quinone prosthetic group, tryptophan tryptophylquinone. topa quinone or lysine tyrosylquinone (for reviews of these other quinoproteins see Duine, 1991; Davidson, 1993; Klinman. 1995; Anthony, 1996, 1998). It is not straightforward to achieve a definitive identification of PQQ as the prosthetic group of an enzyme so we have included here a brief summary of the main types of method that are appropriate. Although the spectra of PQQ-containing quinoproteins have a characteristic absorption band between 300 nm and 420 nm, due to the bound PQQ. this is markedly affected by the environment and cannot be relied upon for definitive identification. It is first necessary to release the PQQ from the protein. The characteristic fluorescence spectrum of the isolated prosthetic group provides a further indication of the identity as PQQ (Anthony and Zatman, 1967; Dekker et al.. 1982) but it is advisable to confirm this by chromatographic methods, with appropriate standards. This is best done by HPLC using a reversed-phase column in combination with absorbance or fluorescence detection; because adduct formation can lead to negative results it may be advisable to produce a known adduct prior to chromatography or biological analysis (Duine el al., 1983, 1987; van der Meer a/., 1990; Duine, 1991). A second valuable method for confirmation of the presence of PQQ is to use its ability to reconstitute active holoenzyme (alcohol or glucose dehydrogenases) from apoenzyme, produced by removal of PQQ, or isolated from bacteria that are unable to synthesize PQQ (Duine et a/., 1983; Groen ef al., 1986; Geiger and Gorisch, 1987; Adachi e t a/., 1988, 1990b). A redox cycling method may be useful to indicate the possible presence of PQQ (Fluckiger et al., 1995), but this is inappropriate as the only way of demonstrating that PQQ is the prosthetic group of a dehydrogenase. A modified form of the assay is useful as a ‘stain’ for quinoproteins after polyacrylamide gel electrophoresis. This non-specific method has been t p f

PQQ AND QUINOPROTEINS

7

used in attempts to demonstrate the presence of very low concentrations of PQQ in animal tissues and fluids, but although it appeared from such experiments that PQQ occurs in eukaryotic organisms, this conclusion has not yet achieved general acceptance (van der Meer et al., 1990; Duine, 1991). 2.2. The Effect of Exogenous PQQ on Bacterial Growth

Many of the bacteria which can synthesize PQQ excrete it into the medium under certain growth conditions (Ameyama et al., 1988; van Kleef and Duine, 1989). This may be important in microbial communities in natural environments because in some circumstances PQQ can stimulate bacterial growth. Some organisms which cannot synthesize PQQ make an apoenzyme dehydrogenase which can be converted to the active holoenzyme if PQQ is present in the growth medium (Groen et al., 1986; van Schie et al., 1987~). These bacteria may then be able to use additional substrates as energy sources, and possibly also as carbon sources. For example, a Pseudomonas sp. which makes the apoenzyme of polyvinyl alcohol dehydrogenase is only able to grow on polyvinyl alcohol when exogenous PQQ is supplied (Matsushita and Adachi, 1993b). The presence of PQQ can also enable more efficient use of a growth substrate, as in the case of Comamonas testosteroni which grows slowly on ethanol and some other alcohols, probably by using an NAD+-dependent alcohol dehydrogenase. This organism also synthesizes an apo-alcohol dehydrogenase; addition of PQQ to the growth medium leads to formation of an active quinohaemoprotein and there is a concomitant increase in growth rate (Groen et al., 1986). Synthesis of PQQ is 'expensive' in energy terms and organisms which can take advantage of exogenous PQQ to convert their apoenzymes into active enzymes may be at an advantage under natural conditions. In this context it is interesting to note that PQQ can act as a chemoattractant to Escherichiu coli (de Jonge et al., 1996), Rhizobium meliloti and Bradyrhizohium japonicum (Boiardi et al., 1996). PQQ can also stimulate growth of bacteria which synthesize active PQQdependent dehydrogenases. In this case, there is no effect on the growth rate or growth yield, but there is a decrease in the lag time on inoculation into fresh medium (Ameyama et al., 1988).

3. THE QUINOPROTEIN DEHYDROGENASESCONTAINING PQQ There is considerable diversity in the nature of the quinoprotein dehydrogenases containing PQQ as their prosthetic group. They may contain only

8

PAT M. GOODWIN AND CHRIS ANTHONY

PQQ (the quinoproteins) or they may also contain haem as a second prosthetic group (the quinohaemoproteins); they may be monomeric or multimeric; freely soluble in the periplasm or bound to membranes; some are firmly associated with other redox components while others are not; their electron acceptors may be cytochromes, blue copper proteins or membrane ubiquinone; the PQQ may be tightly bound or easily dissociated; the enzyme may sometimes be produced in the apo form, requiring addition of PQQ, either during or after growth, for activity. The quinoproteins may be assayed using their physiological electron acceptors, for example, cytochromes, blue copper proteins or soluble ubiquinone analogues. It is more usual to assay them using artificial dye electron acceptor systems such as phenazine methosulphate, phenazine ethosulphate or Wurster’s blue. It is assumed that these either interact directly with the PQQ or accept electrons from the surface of the enzyme near the PQQ. In addition to using these electron acceptors, the quinohaemoproteins may also be assayed with ferricyanide, which accepts electrons from the higher potential haem groups. For references to assay systems see reviews of Anthony (1986, 1996, 1998), Duine et al. (1987) and Duine (1991). The best characterized enzymes are described below; key references are given in Table 1 and their structures, roles in electron transport and physiological functions discussed in Sections 5, 6 and 7 respectively. In addition to those listed in Table 1, the enzymes described below have also been identified as PQQ-containing quinoproteins. A membrane-bound polyvinyl alcohol dehydrogenase (a type I1 alcohol dehydrogenase - a quinohaemoprotein) occurs in a Pseudomonas species during growth on polyvinyl alcohol; this organism cannot synthesize PQQ, which must be provided in the growth medium, or added to the apoenzyme after its isolation in order to obtain active holoenzyme (Shimao et a/., 1989, 1996). A soluble alcohol dehydrogenase which is very similar to methanol dehydrogenase is produced by Rhodopseudomonas acidophila during growth on ethanol (Bamforth and Quayle, 1978). This organism also produces a soluble quinohaemoprotein aromatic alcohol dehydrogenase (Yamanaka, 1989); and a polyethylene glycol dehydrogenase which is likely to be a quinoprotein and is responsible for growth on this substrate (Kawai et a/., 1985; Yamanaka, 1991). A number of other possible PQQ-containing dehydrogenases have been described in the membranes of acetic acid bacteria, in addition to the wellestablished alcohol and glucose dehydrogenases; these include dehydrogenases for aldehydes (see Section 4), glycerol and fructose (Matsushita cr d.,1994).

Table 1 PQQ-containing quinoproteins that oxidize alcohols.'

Enzyme

Prosthetic groups Location

Organism

References

Methanol PQQ dehydrogenase; soluble quinoprotein

Periplasm

Methylotrophs

Anthony and Zatman (1967); Anthony (1986, 1993b, 1996, 1997); Frank er at. (1989)

Alcohol dehydrogenase PQQ (type I); soluble quinoprotein

Periplasm

Pseudomonas aeruginosa

Groen ei at. (1984); Gorisch and Rupp (1989); Mutzel and Gorisch (1991); Schrover et at. (1993)

Pseudomonas putida

Gorisch and Rupp (1989); Toyama e f al. (1995)

Comamonas iesiosteroni

Groen et al. (1986); de Jong et al. (1995a,b); Geerlof et al. (1994); Stoorvogel et at. (1996)

Pseudomonas puiida (two types)

Toyama et al. (1995)

Gtuconobacter

Ameyama and Adachi (1982); Shinagawa et at. (1989); Kondo and Horinouchi (1997); Matsushita and Adachi (1 993a); Matsushita et at. (1994, 1996) Ameyama and Adachi (1982); Inoue et al. (1989, 1990, 1992); Kondo ei at. (1995); Kondo and Horinouchi (1997); Matsushita and Adachi (1993a); Matsushita et al. (1994); Takemura ei at. (1993); Tamaki et at. (1991); Tayama et al. (1989)

Alcohol dehydrogenase PQQ, haem C (type 11); soluble quinohaemoprotein

Alcohol dehydrogenase PQQ, haem C (type 111); membrane quinohaemoprotein

Periplasm

Membrane

Acetobacter

'Only those enzymes that have been studied extensively are included here; see text for mention of others.

10

PAT M. GOODWIN AND CHRIS ANTHONY

Finally, a soluble haemoquinoprotein lupanine hydroxylase which is used during growth on alkaloids has been purified from a Pseudomonas species (Hopper et al.. 1991).

3.1. Soluble Quinoprotein Alcohol Dehydrogenases

3.1.I. Methanol Dehydrogenases The most fully described alcohol dehydrogenases are the methanol dehydrogenases of methylotrophic bacteria which oxidize methanol to formaldehyde during growth of bacteria on methane or methanol (for reviews of the many examples of this enzyme see Anthony, 1986; for reviews of the work of the Delft group on mechanism see Duine er al., 1987; Frank et ul., 1989; Duine, 1991; for reviews of structure and mechanism see Anthony, 1993b, 1996, 1998). Methanol dehydrogenase is a soluble periplasmic enzyme having an a& tetrameric structure; each a subunit (66kDa in Methylohacteriuni extorquens) contains one molecule of PQQ and one Ca2+ ion. The p subunit is very small (8.5 kDa in Methylobacterium extorquens); it has no known function and, like PQQ, it cannot be reversibly dissociated from the a subunit. The structure of this enzyme has several important novel features, including eight ‘tryptophan-docking motifs’ that maintain the structure of the a subunit, the presence in the active site of an unusual disulphide ring structure formed from adjacent cysteine residues, and a Ca” ion coordinated to PQQ (see Sections 4 and 5 for further discussion of its structure and function). In Methylohacterizrm extorqueris the genes encoding the a and p subunits (nixaF and ntsaf) are located in a cluster of 12 methanol oxidation genes (nixaFJGIR(S)A C K L D B ) (Nunn and Lidstrom, 1986b; Amaratunga et al., 1997a,b). MxaG is cytochrome cL, the electron acceptor for the dehydrogenase. Between mxuF and mxaG is nisaJ which codes for a periplasmic protein (30 kDa in Methylobacterium e_utoryuens)of unknown function. Following studies with the enzyme from Acetohacter merhunolicus it has been suggested that this might be a third enzyme subunit (Matsushita et al., 1993), but this has not been confirmed, either for this enzyme or for other methanol dehydrogenases. The msaFJGl genes are cotranscribed from a promoter upstream of mxuF. They have been sequenced and each of the predicted polypeptide products contains a typical signal sequence, which is cleaved on translocation into the periplasm (Nunn and Anthony, 1988; Nunn et al., 1989; Anderson et al., 1990). Methanol dehydrogenase oxidizes a wide range of primary alcohols, having a high affinity for these substrates; for example, the K,, for methanol ~. alcohols are rarely oxidized. The enzyme can be is 5 - 2 0 ~ Secondary

PQQ AND QUJNOPROTEINS

11

assayed with phenazine ethosulphate or Wurster’s blue, but not with ferricyanide. Using phenazine ethosulphate in the dye-linked assay system the pH optimum is about 9 and ammonia or methylamine is required as activator. Its physiological electron acceptor is a novel acidic cytochrome c (cytochrome cL). Apart from its haem-binding site it has little homology with other cytochromes and in particular it lacks the conserved lysine residues which, in most c-type cytochromes, interact with cytochrome oxidase. This is not surprising in view of the fact that cytochrome cL mediates electron transfer between methanol dehydrogenase and a typical Class I c-type cytochrome and it has been shown that the reaction of methanol dehydrogenase and cytochrome cL is initially by way of electrostatic interactions between lysine residues on the basic dehydrogenase and carboxylate residues on the acidic cytochrome (Chan and Anthony, 1991b; Cox et at., 1992); after initial ‘docking’ it is possible that an electron transfer complex is subsequently formed in which there is a hydrophobic component to the bonding (Harris et al., 1994; Dales and Anthony, 1995).

3.1.2. Ethanol Dehydrogenases (Type I Alcohol Dehydrogenases) An ethanol dehydrogenase similar to methanol dehydrogenase has been described in Pseudomonas aeruginosa and Pseudomonas putida (Groen et al., 1984; Gorisch and Rupp, 1989; Mutzel and Gorisch, 1991; Toyama et al., 1995). Like methanol dehydrogenase, it has a high pH optimum, requires ammonia or alkylamines as activator in the dye-linked assay system (ferricyanide is not used as electron acceptor), and is able to oxidize a wide range of alcohol substrates including secondary alcohols. Its absorption spectrum is very similar to that of methanol dehydrogenase but it differs in its very low affinity for methanol; the K,,, for ethanol is about 15 PM and that for methanol about 1000 times higher. In the first description, the enzyme from P . aeruginosa was said to be monomeric with two molecules of PQQ per monomer (101 kDa) (Groen et at., 1984). Subsequently it has been claimed to have the same a2P2 tetrameric structure as in methanol dehydrogenase (Schrover et at., 1993); no P subunit could be seen, however, in the pure enzyme from P . putida (Toyama et al., 1995). This enzyme is unusual in being inhibited by EDTA (measured in the dye-linked assay), which leads to release of PQQ and formation of inactive monomers, as seen by gel filtration (Toyama et a/., 1995). This type I alcohol dehydrogenase uses a specific c-type cytochrome (cytochrome cQEDH)as electron acceptor (Schrover et at.. 1993).

12

PAT M. GOODWIN AND CHRIS ANTHONY

3.1.3. Soluble Quinohaemoprotein Alcohol Dehydrogenases (Type II Alcohol Dehydrogenases) A periplasmic alcohol dehydrogenase has been described in Comamonas testosteroni. It is a monomer (71 kDa) containing two prosthetic groups one molecule of PQQ and a single haem C (Groen et al., 1986; de Jong et al., 1995a,b). In the dye-linked assay system the pH optimum is 7.7 and there is no requirement for an amine activator. Because electron transfer from PQQ is by way of haem C this enzyme can also be assayed using ferricyanide. It has a wide specificity for primary and secondary alcohols, although it is unable to oxidize methanol; it also oxidizes aldehydes and can accept large molecules such as steroids as substrates. This has been exploited for enantiospecific oxidation of industrially important precursor molecules (synthons) (Geerlof et al., 1994). It was first isolated from C . testosteroni as the apoenzyme, containing the haem, which is covalently bound, but lacking PQQ. Reconstitution to the active holoenzyme requires one molecule of PQQ and calcium ions, indicating that its structure and function at the active site might be similar to that of methanol dehydrogenase. EPR spectroscopy has been used to demonstrate the presence of the semiquinone form of PQQ in the active site; haem C has also been detected by EPR spectroscopy and this, together with the absorption spectra, indicates that the iron is similar to that in other low-spin cytochromes c in being coordinated by histidine and methionine (de Jong et al., 1995a). Because this quinohaemoprotein enzyme is soluble and requires addition of PQQ for activity it is likely to provide an excellent system for the study of intraprotein electron transport. Studies with NMR and Raman resonance spectroscopy have shown that binding of PQQ induces a conformational change in the protein, a reorientation of the methionine ligand of haem C , an increase of electron density on one of the pyrrole rings, and an increase in midpoint redox potential of the haem. Although this clearly indicates that the presence of PQQ in the enzyme affects the properties of the haem, it is unclear whether the interactions between the two cofactors are direct or indirect (de Jong et al., 199%). All the available evidence is consistent with the obvious interpretation. that electrons pass from the reduced form of PQQ to the haem (midpoint redox potential, 140mV) and thence to an external electron acceptor. The nature of the electron acceptor for this enzyme has not been reported but, because it is a periplasmic enzyme, this is likely to be a high potential c-type cytochrome or a blue copper protein. The gene encoding this dehydrogenase has been recently isolated, sequenced and expressed in E. coli to give the apoenzyme, lacking PQQ (Stoorvogel et al., 1996); it was necessary to grow the host E. coli in a low concentration of oxygen to obtain synthesis of the haem required for the haem component of the quinohaemoprotein. The DNA sequence

PQQ AND QUINOPROTEINS

13

indicates that the preprotein has a signal peptide typical of periplasmic enzymes. Part of the deduced amino acid sequence is similar to that of methanol dehydrogenase and shows conservation of the tryptophan docking motifs, the adjacent cysteine residues involved in formation of the disulphide ring, and residues involved in binding PQQ and coordination of the Ca2+in the active site. A type I1 alcohol dehydrogenase (soluble quinohaemoprotein) has also been described in P. purida, in which it is formed as the holoenzyme (Toyama et al., 1995). Remarkably, this organism has two immunologically distinct forms of the enzyme; one is induced during growth on mediumchain length alcohols (e.g. butanol) and the other during growth only on 1,2-propanediol or glycerol. The same organism also has a type I alcohol dehydrogenase which is present during growth on all substrates but induced to the highest level during growth on ethanol.

3.2. Membrane-associated Quinohaemoprotein Alcohol Dehydrogenases (Type 111 Alcohol Dehydrogenases)

The type 111 alcohol dehydrogenase is a quinohaemoprotein-cytochrome c complex and has only been described in the acetic acid bacteria Acetobacler and Gluconobacter (for reviews see Beppu, 1993; Matsushita and Adachi, 1993a; Matsushita et al., 1994). Together with the membrane-bound aldehyde dehydrogenase (see below), it is responsible for the characteristic oxidation of alcohol to acetic acid in vinegar production. It does not require ammonia as activator and has a pH optimum of 4-6. Its substrate specificity is relatively restricted compared with other quinoprotein alcohol dehydrogenases; it oxidizes primary alcohols (chain length, C2-C6) but does not oxidize methanol or secondary alcohols and has some activity with formaldehyde and acetaldehyde. It is distinguished from other alcohol dehydrogenases in usually having three subunits and in being tightly bound to the periplasmic membrane, requiring detergent for its isolation. Its natural electron acceptor is ubiquinone in the membrane. Subunit I (72-80 kDa) is a quinohaemoprotein similar to the soluble (type 11) quinohaemoprotein alcohol dehydrogenase, in that it has a single molecule of PQQ and a single haem C. The structural genes of several acetic acid bacteria have been sequenced and the genes encoding subunits I and I1 are adjacent on the genome with the same polarity and in the same reading frame (Tamaki et a f . , 1991; Kondo and Horinouchi, 1997). Translation of the gene sequences shows that all the subunits have N-terminal signal peptides typical of periplasmic proteins. The predicted amino acid sequence of subunit I indicates that it is a typical soluble protein, the first part of which has sequence similarity to the soluble

14

PAT M. GOODWIN AND CHRIS ANTHONY

methanol dehydrogenase but with a C-terminal extension having a single haem binding site (Inoue et ul., 1989, 1990; Tamaki et al., 1991). The predicted amino acid sequence of subunit I1 (48-53 kDa) (Tamaki et al., 1991; Inoue et al., 1992) indicates that it has three haem C binding motifs corresponding to the three haems that can be distinguished by biochemical techniques in the pure protein (Matsushita et ul., 1996). Subunits I + I1 therefore have a total of four haems. Most of these enzymes from acetic acid bacteria have a third subunit (subunit 111, 14-17 kDa), although this is absent from Acetohucter polyoxogmes (Tayama et ul., 1989). The gene coding for this small subunit is not linked to the genes encoding the other two subunits (Takemura et ul., 1993; Kondo rt [JI., 1995; Kondo and Horinouchi, 1997). The predicted amino acid sequence for subunit 111 indicates that its processed size is greater (about 20 kDa) than that estimated by SDS-PAGE (14 kDa). As with all other quinoprotein dehydrogenases, this enzyme may be assayed with phenazine methosulphate, which presumably reacts at the level of PQQ; in addition it can be assayed with ferricyanide which reacts at the level of one or more of the haem C prosthetic groups on subunits I and 11. The rate of reaction with ferricyanide is about 10 times greater than observed with the soluble quniohaemoprotein and it has therefore been suggested that the cytochrome subunit I1 is needed for this high rate of electron transport. The quinohaemoprotein of acetic acid bacteria differs from all other alcohol dehydrogenases in using short-chain ubiquinone homologues (QI and Q2) as electron acceptors and native ubiquinone (Qu and Qlo) when reconstituted in membrane vesicles (Matsushita c’t d., 1992b). I t appears to be unique in a number of ways; it requires detergent for its isolation from membranes and so seems to be a typical integral membrane protein, although none of the subunits appears to have characteristic membrane protein structural domains. Furthermore, the electron acceptor for the quinohaemoproteinxytochrome c complex is membrane ubiquinone, so we have the unusual situation where a c-type cytochrome precedes ubiquinone in the electron transport chain. It has recently proved possible to isolate subunit I1 (the tri-haem cytochrome c) and a separate complex containing subunits I and 111 from Gluconohuctrr .suhoxy/alunLs(Matsushita et al., 1996). Subunits I + 111 had activity in the dye-linked assay, and some activity (only at low pH) with ferricyanide, presumably by way of the haem C of the quinohaemoprotein subunit I. There was no activity using coenzyme Q I but this activity was restored by addition of subunit 11, leading to the conclusion that electron transfer to the quinone takes place by way of the haems on this cytochrome component. As expected, subunit I1 had no catalytic activity alone. Kinetic analysis of ferricyanide reduction at various pH values indicated that only two of the haenis in subunit I1 may be involved in electron transfer from

15

PQQ AND QUINOPROTEINS

reduced PQQ in subunit I to the ubiquinone, but further work on this complex system is needed to be certain about this. The work described above has recently been complemented by a different experimental approach using the enzymes from Gluconobacter suhoxydans and Acrtohacter pasteurianus (Kondo et al., 1995; Kondo and Horinouchi, 1997). Previous work with the enzyme from Acetohacter pol-voxogenes, which has no small subunit 111, had indicated that this subunit is not essential for activity (Tamaki et al., 1991). Subsequently it was shown that spontaneous mutants of A . pasteurianus, lacking subunit 111, contained inactive enzyme and could be complemented by the gene encoding subunit 111. Thus, in this organism at least, subunit I11 is essential for activity (Kondo et al., 1995). Analysis of the location of the subunits in G . suboxydans and A . pasteurianus, and in the mutants led to the suggestion that the cytochrome subunit I1 is firmly embedded in the membrane, that subunits I and 111 are firmly attached to each other and that this attachment helps the dehydrogenase subunit I couple with the cytochrome c (subunit 11), thereby keeping the correct conformation for electron transport of the alcohol dehydrogenase complex on the periplasmic surface of the membrane (Fig. 3) (Kondo

Cytoplasm

Figure 3 The arrangement of the quinohaemoprotein type 111 alcohol dehydrogenase (ADH), and glucose dehydrogenase (GDH) in the membrane of acetic acid bacteria. The interaction of subunit I1 of the alcohol dehydrogenase with the membrane is unusual; although firmly attached, it does not have typical transmembrane helices. The membrane glucose dehydrogenase is likely to be organized in the same way in all bacteria.

16

PAT M. GOODWIN AND CHRIS ANTHONY

and Horinouchi, 1997). This is consistent with the conclusions of Matsushita et al. (1996). Internal electron transport is presumably from PQQH2 to the haem in subunit I and hence by way of two or three of the haems in the cytochrome subunit I1 to the membrane ubiquinone. This raises the question of how the ubiquinone in the membrane reacts with subunit I1 to accept electrons from its haem. Clearly part of the protein must be embedded in the membrane for this to occur but subunit I1 does not appear to have typical hydrophobic transmembrane helices. In some growth conditions (high aeration, low pH), the arrangement of the subunits appears to become altered so that an inactive form of the enzyme is produced; a change in growth conditions (low aeration, neutral pH) then leads to an alteration to the active form, a process that requires expenditure of energy (Matsushita et al., 1995b). Acetobacter methanolicus is an unusual type of acetic acid bacterium, being able to grow on methanol as well as on glycerol and sugars. During growth on glycerol a membrane quinohaemoprotein (type 111 alcohol dehydrogenase) is produced which is similar to that in other species of Acetobacter, together with a ubiquinol oxidase, cytochrome bo (Matsushita et al., 1992a). During growth on methanol, by contrast, a typical methanol dehydrogenase and cytochrome cL are induced, which are unusual only in the optimum pH (pH 4) at which they react (Elliott and Anthony, 1988; Chan and Anthony, 1991b). For oxidation of the cytochrome cL a second periplasmic cytochrome c and a new terminal cytochrome c oxidase (cytochrome co) are also induced, to complete the electron transfer chain from cytochrome cL (Chan and Anthony, 1991a). This is probably the only example of a species of acetic acid bacteria having soluble c-type cytochromes and the appropriate cytochrome oxidase for their oxidation.

3.3. Glucose Dehydrogenases The first description of glucose dehydrogenase and its prosthetic group was by Hauge (1964), PQQ being subsequently identified by Duine and colleagues in a paper in which they first introduced the term quinoproteins (Duine et al., 1979). The organism used for this work was Acinetobacter calcoaceficus which was later shown to be unique in having two different quinoprotein glucose dehydrogenases, one periplasmic and the other an integral membrane protein. 3.3.1. The Membrane-bound Glucose Dehydrogenase This glucose dehydrogenase has been described in a wide range of bacteria including A . calcoaceticus, enteric bacteria, pseudomonads and acetic acid

PQQ AND QUINOPROTEINS

17

bacteria (Table 2). Although the enzymes differ slightly in some properties such as substrate specificity and stability, they are similar in most essential respects (for reviews see Duine et al., 1987; Matsushita and Adachi, 1993a; Matsushita et al., 1994; Anthony, 1996, 1998). Glucose dehydrogenase catalyses the oxidation, to the lactone, of the pyranose form of D-glucose and other monosaccharides, usually including mannose, galactose, rhamnose and xylose. It differs from the soluble enzyme in that it is unable to oxidize disaccharides but, remarkably, a mutation in a single amino acid (histidine to asparagine) was sufficient to confer on the enzyme from G . suboxydans the ability to oxidize maltose (Cleton-Jansen et al., 1991); modelling studies indicate that this change increases the width of the entrance to the active site region of the enzyme (Cozier and Anthony, 1995). The oxidation of glucose to the lactone occurs in the periplasm and the electron acceptor is ubiquinone in the membrane (Beardmore-Gray and Anthony, 1986; Matsushita et al., 1982a, 1987, 1989a,b). After solubilization from the membrane, the enzyme is isolated as a monomer of about 87 kDa, containing one PQQ molecule. The structural gene has been sequenced from four different bacteria; the predicted protein sequences are similar to each other and also have important similarities to other quinoproteins (Cleton-Jansen et al., 1988a, 1989, 1990; Anthony, 1992a, 1996; Cha et al., 1997). Topological and sequence analysis of the protein reveals that it is likely to have five membrane-spanning regions in the N-terminal region and this region is likely to contain the ubiquinone binding site (Yamada et al., 1993b) (Fig. 3). The enzymes from different bacteria differ with respect to their stability and the ease with which PQQ may be dissociated from them; it has been suggested that they can be considered to be in two classes, depending on their stability with respect to EDTA (Dokter et al., 1986; Sode et al., 1995a,b). The type I enzyme is easily denatured and occurs in E. coli and Pseudornonas sp., whereas the more stable type I1 enzyme occurs in Acinetobacter and Gluconobacter. The stability of the E. coli enzyme has been modified by a single amino acid substitution and by formation of a chimeric enzyme using the glucose dehydrogenase structural genes from E. coli and A . calcoaceticus (Sode et al., 1995a,b).

3.3.2. The Soluble Glucose Dehydrogenase of Acinetobacter calcoaceticus Besides the typical membrane-bound glucose dehydrogenase, A . calcoaceticus also contains a completely different soluble enzyme which, to date, has only been described in this organism (Dokter et al., 1986; Geiger and Gorisch, 1986, 1989; Gorisch et al., 1989). It is a dimer of identical subunits of about 50 kDa, each containing one molecule of PQQ, and the sequence of the structural gene indicates that it is a periplasmic protein (Cleton-Jansen

Table 2

PQQ-containing quinoproteins that oxidize glucose.'

Enzyme

Location

Organism

References

Glucose dehydrogenase (soluble)

periplasm

Acinetobacter calcoaceticus

Hauge (1964); Cleton-Jansen et at. (1989); Dokter et at. (1986, 1987); Geiger and Gorisch (1986, 1989); Matsushita et at. (1989b, 1995a); Schlunegger et at. (1993)

Glucose dehydrogenase (membrane)

membrane

Acinetobacter calcoaceticus

Cleton-Jansen et al. (1988a,b); Duine et at. (1979); Matsustuta er at. (1989b, 1995a)

Pseudomonas

Imanaga (1989); Matsushita et al. (1980, 1982a); van Schie et at. (1984)

Gluconobacter suboxydans

Ameyama et at. (1981); Cleton-Jansen et al. (1991); Matsushita et al. (1989~)

Escherichia coli

Ameyama et at. (1986); Beardmore-Gray and Anthony (1986); Cleton-Jansen et al. (1990); Hommes et at. (1984); Matsushita et at. (1986, 1987); Shinagawa er at. (1986); Yamada et at. (1993a,b)

Ktebsietla pneumoniae

Neijssel et at. (1983. 1989); Buurman et at. (1990, 1994)

'The references given are to the enzymes about which most information is available; other bacteria with membrane glucose dehydrogenase are mentioned in the text. No glucose dehydrogenase has been described in which there is any prosthetic group in addition to the PQQ.

PQQ AND QUINOPROTEINS

19

et al., 1989). It catalyses the oxidation of D-glucose, arabinose, galactose, xylose and also the disaccharides lactose, cellobiose and maltose. It is active with Wurster’s blue (pH optimum 9) and 2,6-dichlorophenolindophenol (optimum pH 6 ) , but not with ferricyanide. The soluble enzyme does not react with ubiquinone (Matsushita et al., 1989b), and although it slowly reduces a soluble cytochrome h there is no evidence that this cytochrome interacts with the electron transport chain (Dokter et al., 1988). Although its predicted amino acid sequence has a small degree of similarity to the other quinoprotein dehydrogenases (Cleton-Jansen et al., 1989; Anthony, 1992a), it lacks all their characteristic features including the eight 1 1-residue tryptophan docking motifs which are typical of all the other PQQ-containing quinoprotein dehydrogenases (Anthony, 1996). Its structure has not yet been determined although it has been crystallized (Geiger and Gorisch, 1986; Schlunegger et al., 1993).

3.4. Aldehyde Dehydrogenases

At first glance there appear to be almost as many PQQ-dependent aldehyde dehydrogenases as there are alcohol dehydrogenases, but many of the first descriptions are not sufficient for their definitive identification, and considerable doubt has now been cast on their status as PQQ-containing dehydrogenases. There is a further confusion in that many alcohol dehydrogenases are also able to oxidize aldehydes and it is not certain to what extent this indicates a physiological function. The enzymes for which most information is available are the aldehyde dehydrogenases isolated with detergent from the membranes of acetic acid bacteria (Matsushita and Adachi, 1993~).These usually have a low pH optimum (about pH 4) and oxidize aldehydes of carbon chain length C2-C4. Their function is to catalyse the oxidation, in the periplasm, of aldehydes produced by the action of the membrane-bound quinohaemoprotein alcohol dehydrogenase. Free aldehydes have not been detected during the oxidation of ethanol to acetic acid and these two enzymes are responsible for the production of acetic acid from ethanol which is characteristic of acetic acid bacteria. The aldehyde dehydrogenase has two or three subunits, one being a cytochrome c component and the other originally thought to contain PQQ. However, mutants of one strain of Acetobacter, which were unable to produce PQQ and therefore produced inactive alcohol and glucose dehydrogenases, had the same level of aldehyde dehydrogenase activity as the parent strain, suggesting that this enzyme cannot be a PQQ-containing quinoprotein (Takemura er al., 1994). Furthermore, the gene encoding this enzyme has been isolated and its predicted amino acid sequence does not show similarity to other quinoprotein dehydrogenases (Tamaki et al., 1989). The

20

PAT M. GOODWIN AND CHRIS ANTHONY

electron acceptor for these membrane aldehyde dehydrogenases has not been identified, but as there is usually no cytochrome c oxidase in these bacteria it is probable that electrons are passed to the cytochrome subunit and thence to ubiquinone in the membrane, as shown in the alcohol dehydrogenases of these bacteria.

4. THE IMPORTANCE OF DIVALENT METAL IONS IN THE

STRUCTURE AND FUNCTION OF PQQ-CONTAINING QUINOPROTEINS

That divalent metal ions are important in the structure or function of these quinoproteins was first indicated by the early work on the membrane-bound enzymes (Duine et ul., 1983; Ameyama et ul., 1985) (Tables 3 and 4). PQQ can be removed from these enzymes by treatment with EDTA, heat, low pH or high salt concentrations and in some conditions they are produced as the apo-form, lacking PQQ. Reconstitution with PQQ then requires the presence of a divalent metal ion, which is most commonly Ca2+ or Mg", but other divalent ions are sometimes as good or better (Table 4). Although these observations demonstrated a requirement for divalent metal ions for insertion of PQQ into the enzymes, they provided no indication of whether or not the metal ions had been incorporated as structural or functional components of the active enzymes. 4.1. Calcium in Methanol Dehydrogenase and Other Alcohol Dehydrogenases

The only enzyme for which a structure is available is methanol dehydrogenase, which has a Ca2+ ion tightly coordinated to the PQQ in the active site (Fig. 4; Section 5). The presence of Ca2+ in methanol dehydrogenase was first demonstrated in Methylobucillus glycogenes by Adachi et al. (1990a), and this was subsequently confirmed in the enzymes from other methylotrophs (Richardson and Anthony, 1992) (Table 3). It is not usually possible to remove the Ca2+from methanol dehydrogenase by any treatment, including dialysis against chelating agents, but this has been achieved using the enzyme from Methylophaga marina, a methylotroph that is able to grow at high salt concentrations (Chan and Anthony, 1992). That Ca2+might play some catalytic role in methanol dehydrogenase was indicated by work using some unusual mutants of Methylobacterium extorp e n s . These mutants (defective in the mxuA, K or L genes) synthesize normal a and /3 subunits, and PQQ, but they produce an inactive enzyme

Table 3 Divalent metal ions in PQQcontaining quinoproteins that oxidize alcohols.

Enzyme

Organism

Metal determined

Metal for reconstitution

Methanol dehydrogenase

Methylobacillus

Ca

Ca Sr (during growth)

Adachi et al. (1990a)

Methylobacterium Methylobacterium (mxaA mutant) Methylophilus Paracoccus

Ca None

-

Richardson and Anthony (1992) Goodwin et al. (1996); Goodwin and Anthony (1996)

Ca Ca (Sr)

-

Hyphomicrobium Methylophaga marina

Ca Ca

-

Alcohol dehydrogenase (type I); quinoprotein

Pseudomonm aeruginosa

Ca

Ca Sr (not Mg Mn Cd)

Mutzel and Gorisch (1991); Schrover et al. (1993)

Alcohol dehydrogenase (type 11) (soluble quinohaemoprotein)

Comamonas testosteroni

Ca (not Mg)

Groen et al. (1986); de Jong et al. (1995a,b)

Alcohol dehydrogenase (type 111) (membrane quinohaemoprotein)

Gluconobacter suboxydans

Ca (not Mg)

Shinagawa et a/. (1989)

Ca = Sr = Ba (not Mg) Ca Sr (during growth) Ca

References

Richardson and Anthony (1992) H a m s and Davidson (1994a,b); Richardson and Anthony (1992); Richardson and Anthony (1992) Chan and Anthony (1992)

Table 4 Divalent metal ions in relation to glucose dehydrogenase.

Enzyme

Organism

Metal active in reconstitution

References

G1IlCCS.Z

A cinetobacrer calcoaceticus

Ca(100) Mn(67) Cd(60) (not Mg)

Geiger and Gorisch (1989); Olsthoorn and Duine (1996)

Acinetobacter calcoacericus (mutant)

Cd(127) Ca( 100) Sr(68) Mn(63) Co(l0) Ba(7) (not Mg)

Matsushita et at. (1995a)

Acinetobacter calcoaceticus

Mg > Ca

Duine er at. (1983); Ameyama et at. (1985)

Acinetobacter calcoaceticus (mutant)

Mg( 115) Ca( 100) Zn( 100) Sr(70) Co(33) Cd(27) Ba( 10)

Matsushita et at. (1995a)

Escherichia coli

M g > Ca > Co

Ameyama e f al. (1985, 1986); Shinagawa et at. (1986)

Klebsiella pnewnoniae

Mg, Ca

Buurman e f at. (1990); Neijssel et at. (1983)

Pseudomonas Juorescens

Mg > Ca

van Schie et at. (1984); Ameyama et 01. (1985); Imanaga (1989)

dehydrogenase (soluble)

Glucose dehydrogenase (membrane)

~~

The presence of a metal ion (Ca2+) has only been demonstrated in the soluble enzyme from Acinetobacter calcoaceticus. The values in parentheses refer to relative activities (%) of activity measured with calcium determined after reconstitution for a fixed length of time: they d o not necessarily indicate the most active form of the enzyme.

PQQ AND QUlNOPROTElNS

23

Figure 4 The equatorial interactions of PQQ and the coordination of Ca” in the active site of methanol dehydrogenase (MDH). This was determined by X-ray crystallography; the structures of alcohol dehydrogenase (type 111) (ADH) and membrane glucose dehydrogenase (GDH) are based on molecular modelling studies (see Section 5 ) . This figure also shows Asp303 which is likely to act as a catalytic base, and Arg33I which may also be involved in the mechanism. Figure 9 shows the axial interactions that are also involved in holding PQQ in place in the active site.

with an abnormal absorption spectrum in the PQQ region (Nunn and Lidstrom, 1986a). The DNA sequences of these genes indicate that MxaA is a periplasmic protein, MxaK is cytoplasmic and MxaL is an integral membrane protein. It is possible that two other proteins, MxaC (cytoplasmic) and MxaD (periplasmic), coded by genes in the same cluster as mxaA, are also involved in Ca2+incorporation (Morris rt a/., 1995). Reconstitution of active enzyme having the normal absorption spectrum was achieved by incubation of the purified enzyme with a high concentration of Ca2+

24

PAT M. GOODWIN AND CHRIS ANTHONY

(Richardson and Anthony, 1992). Reconstitution involved a large conformational change, resulting in active holoenzyme from which Ca2+could not subsequently be removed, and was optimal at high pH and high Ca2+ concentrations (Goodwin et al., 1996). This contrasts with the in vivo situation where the pH in the periplasm is neutral and the concentrations of Ca2+ are likely to be relatively low. Presumably assembly of the holoenzyme in the periplasm is facilitated by MxaA, and possibly also by the membrane protein MxaL. Geiger and Gorisch (1989) had previously shown that, in solution, PQQ forms a complex with Ca2+ or Mg2+; the complex has an absorption maximum of 343 nm, different from that of free PQQ (330 nm) but very similar to that of PQQ in alcohol dehydrogenases. They concluded that either the ion is necessary to induce the proper conformation of the protein so that i t can then bind PQQ at the active site, or that PQQ is anchored as a complex with the calcium. Both of these conclusions have now been confirmed for methanol dehydrogenase by studies of Ca2+ incorporation into the enzyme (Goodwin et al., 1996) and by its X-ray structure. This shows a Ca2+ ion in the active site, coordinated directly to PQQ and to amino acid residues (Fig. 4) (White et al., 1993; Blake et al., 1994; Ghosh et al., 1995; Xia et a f . , 1996) and this has led to the proposal that Ca2+ plays a key catalytic role in this enzyme by facilitating the initial reaction with substrate (Anthony et al., Blake et al., 1994; Anthony, 1996, 1998). Remarkably, by using dehydrogenase prepared from a mxaA mutant, it has been possible to produce a methanol dehydrogenase in which the Ca2+has been replaced with Ba2+,the first enzyme ever described to contain active site Ba2+; although substrate binding is much poorer in the barium enzyme the activation energy is lower and hence the maximum catalytic rate is higher (Goodwin and Anthony, 1996). The first demonstration of a metal ion in an alcohol dehydrogenase other than methanol dehydrogenase was in the (type I) quinoprotein ethanol dehydrogenase of P. aeruginosa, which has one Ca2+ per molecule of PQQ (Mutzel and Gorisch, 1991). This could be removed by treatment with CDTA (not EDTA) and reconstitution achieved by incubation with PQQ and Ca2+ or Sr2+, but not with Mg2+, Mn2+ or Cd2+.Ca2+ (but not Mg2+) is required for formation of active enzyme from the apoenzyme of the type 11 and type I11 alcohol dehydrogenases (the quinohaemoproteins), but the presence of calcium has not yet been demonstrated in the isolated enzymes (Groen ef al., 1986; Shinagawa et al., 1989). However, sequencing and modelling studies of their structures, including the coordination sites for Ca2+ (Fig. 4, Section 5), are sufficiently similar to methanol dehydrogenase for it to be concluded that Ca2+ fulfils a similar role in the type 11 (Stoorvogel et al., 1996) and type I11 (Cozier et al., 1995) alcohol dehydrogenases.

PQQ AND QUINOPROTEINS

25

4.2. The Role of Divalent Metal Ions in Glucose Dehydrogenase

The presence of a metal ion (Ca2+ ) in a glucose dehydrogenase was first demonstrated in the soluble enzyme from A . calcoaceticus (Geiger and Gorisch, 1989). The Ca2+ could be removed by treatment with high salt concentrations, low pH or high temperature, subsequent reconstitution requiring PQQ plus Ca2+, Mn2+ or Cd2+ (Mg2+ was not effective and Sr2+ was not tested). This study has been extended by using a mutant of A . calcoaceticus that is unable to synthesize PQQ and so only produces the apoenzyme (Matsushita et al., 1995a). Active enzyme could be formed by incubation with PQQ and a divalent cation, the most effective ions being Cd2+ and Ca2+, followed by Sr2+ and Mn2+; no reconstitution occurred with Mg2+ (Table 4). It should be noted that the percentage values in Table 4 were taken from experiments in which a standard assay was used and reconstitution was for a fixed length of time. These results do not necessarily show that the catalytic activity of the enzyme is highest with the metal ion giving the highest rate in this type of experiment. The K , value for PQQ was 1.3 nM. A second approach to the study of the metal ion in the soluble glucose dehydrogenase was developed by Olsthoorn and Duine (1996) who expressed the A . calcoaceticus gene in E. coli, which does not produce PQQ. The soluble apoenzyme was isolated in the dimeric form, monomerization occurring during gel filtration in the presence of a chelating agent. From a study of reconstitution using the monomer and dimer, it appears that Ca2+ plays a dual role in this enzyme as it is required for dimerization as well as for incorporation of PQQ in a functional form. The ions that could support reconstitution of the wild-type enzyme were also effective in this system (Table 4). After reconstitution, the holoenzyme resembled reconstituted methanol dehydrogenase (Goodwin et al., 1996) in that Ca2+ could no longer be removed by chelating agents. It is difficult to come to a firm conclusion about the role of the divalent metal ion in membrane glucose dehydrogenases. They are completely different from the soluble enzyme in their sequence, structure and location and the metal ion content of the membrane glucose dehydrogenase has never been determined. The only evidence available is from studies of reconstitution of inactive apoenzyme with PQQ to form the active holoenzyme. For this process Mg2+ often is better than Ca2+ and many other metals can be used instead of Ca2+ (Matsushita et al., 1995a) (Table 4). For example, using the same mutant of A . calcoaceticus as was used in the study of the soluble enzyme (see above), it was shown that Mg2+ and Zn2+ were as effective as Ca2+ for reconstitution of active enzyme; in this case the K , value for PQQ (40 nM) was much higher than for the soluble enzyme (1.3 nM) (Matsushita e f al.. 1995a). The metal content of the reconstituted holoen-

26

PAT M. GOODWIN AND CHRIS ANTHONY

zyme cannot be determined because the process is readily reversible, and removal of the excess Mg2+ required for reconstitution will also remove the ion from the enzyme. If the function of the metal ion during reconstitution is to provide a metal ion at the active site, then this will have implications for our understanding of the mechanism of glucose dehydrogenase, as it is unlikely that Mg2+ could replace Ca2+ for some functions. Modelling studies using the predicted amino acid sequence of glucose dehydrogenase, with the coordinates of methanol dehydrogenase, have shown that some of the residues important in coordination with the Ca2+ are different in the glucose dehydrogenase (Cozier and Anthony, 1995) (Fig. 4).

5. THE STRUCTURE AND MECHANISM OF PQQ-CONTAINING QUINOPROTEINS

The predicted amino acid sequences of all the PQQ-containing quinoproteins so far studied show regions similar to that of the superbarrel structure of the a subunit of methanol dehydrogenase (Anthony, 1992a), although none has sequence similar to that of the subunit. The sequence of the catalytic subunit of the type 111 alcohol dehydrogenase, which contains PQQ, has an N-terminal region of 600 residues with 31% identity to the methanol dehydrogenase sequence; this is followed by a C-terminal extension containing a haem-binding site (Fig. 5). In the membrane glucose dehydrogenase, by contrast, the N-terminal region (residues 1-154) forms a membrane anchor with five transmembrane segments and this region is likely to contain the ubiquinone binding site. The structure of methanol dehydrogenase has been determined (Section 5.1) and, although the overall identity level is not particularly high, i t has been possible to use the sequences of the alcohol dehydrogenase and glucose dehydrogenase, together with the coordinates of the methanol dehydrogenase, to produce reliable model structures of the 'superbarrel regions' (Fig. 5; see also Fig. 10) of these two enzymes. A key feature enabling this to be done is the high level of conservation of the tryptophan-docking motifs which form the basic structure of the propeller superbarrel.

5.1. The Structure and Mechanism of Methanol Dehydrogenase

This is the only PQQ-containing dehydrogenase for which a structure is available (Ghosh et al., 1995; Xia et ul., 1996), the highest resolution structure (1.94 A) being that from Methylobacterium exforyuens. This structure

27

PQQ AND QUINOPROTEINS

Methanol dehydrogenase

Glucose dehydrogenase

Alcohol dehydrogenase

Reqion d superbarrel

5 membrane helices

LlDll ,\Dl1 ,iDII

I

150 amino acids

A b u l GO3

amino arl(h

Haem-binding region 100 amino acids

Superbarrel reglon W8

MDH

w1

w2

w.7

101 IABCDI IAECDIIABCDI

w4

I A B CI D A B IC D

I

J A B C D I I A B C D I~

AEC

1

Figure 5 Amino acid sequence alignment of quinoprotein dehydrogenases. Each ’W’ is a four-stranded p sheet (or propeller blade); the letters ABCD correspond to the four strands of each ‘W’ motif (see Fig. 6). These are the regions showing greatest similarity of sequence between the quinoproteins. There are many loops between, and within, the p sheets which show least similarity. For example, there is a long region with little conservation of sequence (including a large loop) between the end of the D-strand in W5 and the end of the D-strand of W6. The highly conserved region between strand-A in W7 and the end of strand-B in W8 was originally proposed t o be a PQQ-binding domain; this is not the case (see Fig. 12).

and its implications with respect to enzyme mechanisms have been reviewed elsewhere (Anthony et al., 1994; Anthony, 1996, 1998). The basic structure of the ci subunit is a ‘propeller fold’ superbarrel made up of eight p sheet ‘propeller blades’ (‘W’ motifs) which are held together by novel tryptophandocking motifs (Figs. 6 and 7). The p subunit is most unusual as it has no hydrophobic core and forms a very extended structure which wraps around the u subunit (Figs. 6 and 8). In the absence of any other obvious function for this subunit, it has been suggested that it acts to stabilize the folded form of the large chain. The absence of p subunits in most other PQQ-containing quinoproteins, however, indicates that it may have a specific (unknown) function in methanol dehydrogenase. The PQQ is in the centre of the ci subunit, coordinated to a Ca2+ ion (Fig. 4) and is maintained in position by a stacked tryptophan and a novel eight-membered ring structure made up of a disulphide bridge between adjacent cysteine residues (Fig. 9). The methanol oxidation reaction is initiated by abstraction of a proton from the alcohol by a base (Asp303) followed by attack, ,on the electrophilic C-5 of PQQ, of the resulting oxyanion to form a hemiketal intermediate; or attack by a hydride from the methyl group of the methanol (Anthony, 1996, 1998). It has been proposed that the Ca2+ acts as a Lewis acid through coordination to the C-5 carbonyl oxygen, thus facilitating formation of

28

PAT M. GOODWIN AND CHRIS ANTHONY

Figure 6 A drawing of an up unit of MDH looking down the pseudo X-fold axis, simplified to show only the ‘W’ motifs of the u chain, and the long u helix of the p chain, but excluding other limited p structures and short u helices. The PQQ prosthetic group is in skeletal form and the calcium ion is shown as a small sphere. The outer strand of each ’W’ motif is the D strand, the inner strand being the A strand. The ‘W’ motifs are arranged in this view in an anti-clockwise manner. The exceptional motif W8 is made up of strands A< near the C-terminus plus its D strand from near the N-terminus. This figure is based on the structure in Ghosh el a / . (1995).

PQQ AND QUINOPROTEINS

29

Figure 7 The girdle of tryptophan residues involved in docking the p sheets together. The tryptophan residues involved in docking are shown in spacefill mode and the rest of the chain as backbone. The PQQ prosthetic group is in skeletal form and the calcium ion is shown as a small sphere.

the electrophilic C-5 of PQQ. An alternative possibility is that Arg331 plays this role (Anthony, 1996; Xia er al., 1996). A key question in relation to this enzyme, and to the other PQQ-dependent dehydrogenases, is how electrons pass from the quinol form of PQQ (PQQH?) to the outside of the protein and thence to the electron acceptor. The novel disulphide bridge structure is in close contact with the PQQ in the active site and, because reduction of this disulphide bond leads to loss of electron transfer to the cytochrome, it was thought at one time that it must play some direct role in this process; this has subsequently been shown to be unlikely (Avezoux et al., 1995). Electron transfer to the cytochrome must

30

PAT M. GOODWIN AND CHRIS ANTHONY

Figurc 8 The alp2tetrameric structure of methanol dehydrogenase

occur one electron at a time; this leads to formation of the free radical semiquinone form, and the novel disulphide ring structure may play a role in stabilizing this semiquinone form of PQQ (Avezoux er al., 1995; Anthony, 1996).

5.2. The Structures and Mechanisms of Alcohol and Glucose Dehydrogenases

The model alcohol dehydrogenase being considered here is the N-terminal region of subunit I of the type 111 alcohol dehydrogenase of acetic acid

PQQ AND QUINOPROTEINS

31

Figure 9 The novel disulphide ring in the active site of methanol dehydrogenase. The ring is formed by disulphide bond formation between adjacent cysteine residues. The PQQ is 'sandwiched' between this ring and the tryptophan that forms the floor of the active site chamber. The calcium ion is coordinated between the C-9 carboxylate, the N-6 of the PQQ ring and the carbonyl oxygen at C-5.

bacteria (Fig. 10) (Cozier et ul., 1995). In this structure there are considerable differences in the external loops, particularly those involved in the formation of the shallow funnel leading to the active site in methanol dehydrogenase. However, the active site region is highly conserved, including the tryptophan and the disulphide ring on opposite sides of the plane of the PQQ, and most of the equatorial coordinations to the PQQ (Fig. 4). Especially important with respect to the mechanism is the conservation of the active site base (Asp303 in methanol dehydrogenase) and all the coordinations to the calcium ion. This suggests that the mechanism of this alcohol dehydrogenase is essentially similar to that of the methanol dehydrogenase. Comparison of the protein sequence of the soluble quinohaemoprotein ethanol dehydrogenase from Cornamonus testosteroni leads to a similar conclusion for that enzyme (Stoorvogel et a/., 1996). In the model structure of the periplasmic portion of the membrane glucose dehydrogenase of E. coli (Fig. 11) (Cozier and Anthony, 1995), there is a sequence of about 80 amino acids where there is little similarity to the methanol dehydrogenase sequence and so this region cannot be modelled. The novel disulphide ring is replaced by a histidine residue which maintains the position of PQQ in the active site, consistent with the previous demonstration that a histidine residue is essential for binding PQQ (Imanaga, 1989) (Fig. 12). There are fewer equatorial interactions between the protein and PQQ (Fig. 4), perhaps explaining why it is possible to effect the reversible dissociation of PQQ from glucose dehydrogenase but not from methanol

Figure 10 Schematic representation of the backbones of the quinohaemoprotein alcohol dehydrogenase (ADH)and showing their major secondary structure. These model structures are based on that of glucose dehydrogenase (GDH), methanol dehydrogenase wluch was determined by X-ray diffraction (see Fig. 6) (Ghosh er a/., 1995). The model ADH structure is of the N-terminal region of the quinohaemoprotein subunit I of the membrane complex (residues 1-590), omitting the C-terminal haem domain (Cozier et al., 1995). The model GDH structure is of the C-terminal section of the membrane-bound GDH (residues 155-796). omitting the N-terminal membrane region (Cozier and Anthony. 1995). These residues are not present in MDH or ADH and the sequences are too long to model. The prosthetic group is shown as a ball and stick structure. and the Ca’+ as a van der Waal’s sphere. The major loops are in black.

33

PQQ AND QUINOPROTEINS

Trp243

Trp404

Figure 11 Comparison of the stacking interactions of the PQQ in methanol dehydrogenase (MDH) and the model glucose dehydrogenase (GDH). In M D H the PQQ is stacked between the coplanar Trp243 and the disulphide ring system of CyslO3 and CyslO4. In G D H the coplanar tryptophan is retained (Trp404) but the disulphide is not conserved. Instead, His262 may perform a similar role in helping to bind the PQQ into the active site region.

dehydrogenase. One clear difference between these proteins is that there is more ‘space’ in the glucose dehydrogenase active site, perhaps to accommodate the larger substrate. By analogy with the methanol dehydrogenase structure, Asp466 is likely to be involved in base catalysis, initiation of the reaction being by abstraction of a proton from the anomeric hydroxyl of the pyranose ring. As discussed in Section 4, it is possible that the active site of glucose dehydrogenase may sometimes contain big2+ instead of Ca2+, although many of the groups that are involved in coordination to Ca” in methanol dehydrogenase are conserved in the model glucose dehydrogenase. It has been suggested that Arg331 and not Ca2+ may act as a Lewis acid in the mechanism of methanol dehydrogenase; the equivalent residue in glucose dehydrogenase is Lys494 which might facilitate formation of the electrophilic C-5 of PQQ if Mg2+ is present in the active site and unable to fulfil the function proposed for Ca2+. Relatively little is known about the mechanism of glucose dehydrogenase, but some information is available from chemical modification studies of lmanaga (1989), and modelling studies have suggested that many features of its mechanism are likely to be similar to that of methanol dehydrogenase (Cozier and Anthony, 1995; Anthony, 1996, 1998). One key difference is that electron transfer from the reduced PQQ does not occur in two stages to a cytochrome c as in methanol dehydrogenase - electrons

Figure 12 The z2 dimer of MDH showing the highly conserved region. The p subunits are omitted for clarity. The highly conserved region, between strand-A in W7 and the end of strand-B in W8. shown by the dark ribbon, was originally proposed to be a PQQ-binding domain; t b s is clearly not the case. The Ca’+ is shown as a sphere. The PQQ is shown in the active site stacked between the hydrophobic Trp243 and the disulphide ring.

PQQ AND QUINOPROTEINS

35

must pass through the protein to the ubiquinone in the membrane. Although this must also involve transfer of the electrons one at a time, this can be by a rapid direct route between the two redox centres; it is not necessary for a stable semiquinone to be formed, and indeed no semiquinone has ever been observed in glucose dehydrogenase. This is perhaps consistent with the absence of the novel disulphide ring structure present in the alcohol dehydrogenases. There is no suggestion from the model structure or from the primary sequence that there is any hydrophobic region of the protein that could interact with the membrane other than the N-terminal transmembrane segments.

5.3. The Conserved Sequence that was wrongly identified as the PQQ-binding Domain in Quinoproteins

When the primary sequences of these dehydrogenases were first compared it was seen that there was one region of greater identity than any other and it was reasonably concluded that this might represent the one feature known to be common to all PQQ-containing enzymes - their ability to bind PQQ; this region was therefore designated a putative PQQ-binding domain (for review see Anthony, 1992a). Remarkably, this sequence constitutes part of the main propeller structure and is not in any way directly involved in PQQ binding (Fig. 12). Why this region has such a relatively high level of identity is not known but it provides the most obvious region for designing DNA probes for use in the identification of genes coding for PQQ-dependent quinoproteins.

6. QUINOPROTEINS IN ENERGY TRANSDUCTION

The PQQ-dependent dehydrogenases all function in respiration and usually also in energy transduction; in all cases so far investigated, this involves the oxidation of substrates in the periplasm of Gram-negative bacteria (Table 1). This contrasts with oxidation systems involving the membrane flavoproteins which catalyse reactions on the inner face of the cytoplasmic membrane. The reason for the periplasmic location of the quinoprotein dehydrogenases is unclear, but it presumably relates to their nature and/ or to their function. It might be that active enzymes must be assembled outside the cell because of some aspect of PQQ incorporation, or because proper folding of the protein will not occur in a reducing environment. It is probable that all of the quinoproteins contain Ca2+ or Mg2+, and perhaps their periplasmic location avoids the problem of transporting these ions into

36

PAT M. GOODWIN AND CHRIS ANTHONY

cells or of having high concentrations of them within the cells. In the case of methanol dehydrogenase, electrons are passed directly to high potential ctype cytochromes which are only found in the periplasm and so the respiration system is similar to that of bacteria oxidizing inorganic substrates. In this case the product of substrate oxidation must then be transported into the bacteria and this is also necessary with some of the alcohol- and glucoseoxidizing systems. In other cases, however, the product is not used by the bacteria and is released into the medium. Clearly in these systems it is more appropriate to have the substrate oxidized in the periplasm of the bacteria. Whatever the role of the quinoprotein (see Section 7), it must be coupled to an electron transport system, and for ATP production this must be arranged so as to produce a protonmotive force across the inner cytoplasmic membrane to drive ATP synthesis by the membrane ATP synthase (Anthony, 1988, 1993a). The type of electron transport system for the quinoprotein dehydrogenases depends on the type of enzyme and on the type of terminal oxidase present. These fall into two main functional categories: those that oxidize periplasmic cytochrome c (cytochrome aa3 and cytochrome co); and those that oxidize ubiquinol without the mediation of a cytochrome hcl complex (cytochromes ho, hd and ha) (Poole, 1988); most bacteria are able to synthesize only one of these two general types of oxidase, although more than one of the specific type may be produced. The specific oxidase that is in operation at any one time depends upon the growth conditions, the most usual determinant being the oxygen concentration. For example, enteric bacteria only produce quinol oxidases; cytochrome ho has a low affinity for oxygen and is usually produced when oxygen is plentiful whereas cytochrome hd, which has an extraordinarily high affinity for oxygen, is produced when oxygen is scarce or when it must be removed to a very low level in order to protect oxygen-sensitive enzymes. During methanol oxidation in some methylotrophs there is the less usual situation in which the carbon status of the cell determines the oxidase; in carbon excess conditions cytochrome co is synthesized, whereas only the cytochrome 0 0 3 is produced in carbon-limited culture (Cross and Anthony, 1980). Although the number of molecules of ATP produced per pair of electrons passing down the electron transport chain is not as clearly defined as previously thought, the potential yield of ATP will be higher in systems in which ubiquinone is oxidized by way of cytochrome bel complex and periplasmic cytochrome c than when the terminal oxidase is a ubiquinol oxidase. The yield will also depend on whether or not the terminal oxidase has a proton-pumping function. When the electron transport chain bypasses the low redox potential ubiquinone/cytochrome h part of the chain (as in methanol oxidation) no more than one ATP per methanol oxidized is likely to be achieved.

PQQ AND QUINOPROTEINS

37

6.1. Electron Transport Chains involving Soluble Alcohol Dehydrogenases The electron transport systems from methanol dehydrogenase are summarized in Fig. 13 (for reviews see Anthony, 1988, 1992b, 1993a). They have in common the first step, which is electron transfer from the dehydrogenase to the specific cytochrome cL in the periplasm (called cytochrome c551, in Paracoccus). The interaction of these proteins depends on electrostatic interactions and so methanol oxidation is strongly inhibited by high ionic strength. During this first step in the oxidation of methanol, electrons pass from the dehydrogenase by way of cytochromes to the oxidase, and protons are liberated from the reduced PQQ into the periplasm, thus contributing to the protonmotive force. The cytochrome cL is subsequently oxidized by cytochrome C H , which is similar in all respects to the other small c-type cytochromes that mediate electron transfer between cytochrome bc, complexes and oxidases (cytochrome aa3 or cytochrome co). The oxidase consumes protons on the inside face of the membrane and it may also act as a direct proton pump. A critical point with respect to energy transduction is that all these electron transport chains are similar to those operating in the oxidation of inorganic substrates in by-passing the low potential ubiquinone/cytochrome b parts of the chain. The result of this is that the first step in the oxidation of methanol is likely to yield only one molecule of ATP (or less). Hyphomicrobium sp. and Paracoccus denitrijicans are both able to act as denitrifying bacteria during anaerobic growth on methanol with nitrate, and in these conditions the cytochrome c is oxidized by a nitrite reductase. The electron transport systems for periplasmic type I alcohol dehydrogenases (Fig. 14) are likely to be essentially similar to those for methanol oxidation (Fig. 13). The soluble quinoprotein alcohol dehydrogenase from P . aeruginosa reacts rapidly with a small c-type cytochrome called cytochrome cEDH which might be assumed to be related to the cytochrome cL of methylotrophs. It is, however, rather smaller (14.5 kDa compared with about 20 kDa) and there is no similarity between the 19 N-terminal residues of cytochrome cEDH and cytochrome cL (Schrover et al., 1993). Although P . aeruginosa usually also contains a typical small c-type cytochrome which is the substrate for its oxidase, during growth on ethanol by way of the quinoprotein alcohol dehydrogenase there appears to be no cytochrome c able to mediate electron transport between cytochrome cEDH and the oxidase; its electron transport chain (Fig. 14) might therefore be similar to that in the methylotrophic organism 4025 during growth on methanol in media containing a high concentration of copper (Auton and Anthony, 1989a,b; Anthony, 1992b) (Fig. 13). Thus, in P . aeruginosa a blue copper protein like azurin and an ‘azurin oxidase’ may be involved in electron transport

38

PAT M. GOODWIN AND CHRIS ANTHONY

Methylobacterium extorquens

MDH

- - - Cyt. cL

Methylophllus methylotrophus

Cyt.c,

Cyt. aa3

O2

carbon-limited conditions

- - - -

MDH

Cyt. cL

Cyt.c,

Cyt. aa3

O,

Methylophllus methylotmphus carbon-excess conditions

- - - -

MDH

Cyt. cL

Cyt.C,

Cyt. co

0,

Acetobacter methanollcus

MDH

- - - Cyt. cL

Cyt.~,

Cyt. co

0,

Organlsm 4025 low copper MDH

- - - -I - Cyt. cL

Cyt.C,

Cyt. co

0,

kudn

Cyt. co

0,

Organlsm 4025 high copper MDH -Cyt.

cL

Paracoccus denltrMcans

MDH -Cyt.,,,

Cyt. aa3

-

O,

Figure 13 Electron transport chains ofmethylotrophs. The details of these chains are discussed in Anthony (1992b).

39

PQQ AND QUINOPROTEINS

Qulnoproteln alcohol dehydrogenase (type I) (periplasmic) e.g. Pseudomonas ADH

-

-

Cyt.cEoH

Azurln

-

-

Cyt.co

0,

Qulnohaemoprotein alcohol dehydrogenase (type II) (perlplasmic) e.g. Comamonas festosteronl

-

ADH haem c -

Pseudomonas putlda

- + Cyt.c - - + Cyt.c - - +

\ \

\

\r

f

w

l

f

' \ r

'Azurln'

Cyt. aaj cyt.co

-

0 2

/

'Azurln'

Qulnohaemoprotein alcohol dehydrogenase (type Ill) (membranes) Acetic acid bacteria : Acefobacfer and Gluconobacter ADH -haemsc

- UQ

Cyt. ba

\Cyt.bo alternative oxidase

-

0, 0, 0 2

Figure 14 Electron transport chains involved in the oxidation of alcohols. Little is known about the electron transport chain from the quinohaemoprotein type I1 alcohol dehydrogenase. The arrangement with respect to the membrane and ubiquinone (UQ) of the subunits of the quinohaemoprotein type Ill alcohol dehydrogenase is illustrated in Fig. 3. The type of oxidase in the acetic acid bacteria depends on the genus; Gluconohncter produces only cytochrome ho (or the cyanide-insensitive alternative oxidase) whereas Acriohucter produces either cytochrome hn (previously called cytochrome (I,), or cytochrome bo, depending on the growth conditions (see text). Dotted lines indicate alternative routes of electron transfer.

during growth on ethanol. A similar electron transport chain presumably operates in P . putidu during growth on ethanol (Toyama et al., 1995). The electron acceptor for the type I1 alcohol dehydrogenases (quinohaemoproteins) in C. testosteroni and P . putidu is not known but, because

40

PAT M. GOODWIN AND CHRIS ANTHONY

this is a periplasmic enzyme, it is likely to be a specific cytochrome c or blue copper protein (Fig. 14).

6.2. Electron Transport Chains involving Membrane-bound Quinohaemoprotein Alcohol Dehydrogenase (Type 111) in Acetic Acid Bacteria

This type of enzyme has only been described in acetic acid bacteria, which are strict aerobes and have highly active oxidase systems for metabolizing sugars and alcohols. Much of our knowledge of electron transport systems in these organisms (Fig. 14) comes from the work of the Yamaguchi group who have written a comprehensive review of the subject (Matsushita ef al., 1994). The enzyme is a quinohaemoprotein/cytochrome c complex able to react with ubiquinone, and an active respiratory chain has been successfully reconstituted using the enzymes from Acefobacferaceti and G . suboxyduns together with the purified quinol oxidases formed by these bacteria (cytochrome bo or cytochrome ha, previously called cytochrome a , ) (Matsushita ef al., 1992b). Usually reconstitution of membrane enzymes into proteoliposomes requires incubation of all the components together. However, in the case of the type 111 quinohaemoprotein alcohol dehydrogenase it was possible to reconstitute an active system by addition of the enzyme to preformed proteoliposomes containing the oxidases. This is consistent with our previous suggestion that the interaction of the dehydrogenase complex with the membrane must be an unusual one (Section 3.2). There is some evidence that subunit 11 (the tri-haem cytochrome c) may also be able to mediate electron transfer from membrane glucose dehydrogenase (see below) but the physiological significance of this has yet to be determined. Cultures of A . aceti produce either the cytochrome bo or cytochrome ba (Matsushita rt ul., 1994). Cytochrome bo is predominant in static cultures whereas cytochrome ha has a higher affinity for oxygen and is predominant in shaking cultures. The change from one oxidase to the other is not due to a straightforward induction mechanism, however, but involves a poorly understood change from one genetic cell type to another (Matsushita et al., 1994). By contrast with Acefohacferstrains, Gluconohacfer is only able to produce the cytochrome bo but in some conditions it also produces a cyanide-insensitive by-pass which may make use of the cytochrome subunit of alcohol dehydrogenase as part of the electron transport chain for oxidation of ubiquinol (Matsushita et al., 1994, 1995b). Acefobacter mefhanolictrs is an exceptional organism. It is able to grow at low pH on a range of multicarbon compounds such as glycerol and is the only species of acetic acid bacteria able to grow on methanol. During growth on glycerol, ethanol is produced and is oxidized to acetic acid; as

41

PQQ AND QUINOPROTEINS

in other acetic acid bacteria this involves the ubiquinol oxidase cytochrome ho; cytochrome ha is not produced. By contrast, during growth on methanol the soluble periplasmic c-type cytochromes must be synthesized for reaction with methanol dehydrogenase together with an appropriate terminal oxidase (cytochrome co) able to oxidize cytochrome c (Fig. 13) (Elliott and Anthony, 1988; Chan and Anthony, 1991a,b; Matsushita et al., 1992a).

6.3. Electron Transport Chains involving Membrane-bound Glucose Dehydrogenase

Electron transport from glucose dehydrogenase, which interacts directly with membrane ubiquinone, varies from organism to organism, depending on the nature of the oxidase(s) produced and their substrates (Fig. 15).

Pseudomonas

GDH

- UQ

Cyt. bC1

-

cyt.c -cyt.co

-

0,

Aclnetobacter calcoacetlcus and Escherlchla coli

GDH

- UQ

cfl. bo

\ Cyt.bd Gluconobacter

GDH

- UQ

\

Cyt.bo cyt.

of ADH Figure 15

-

02

0,

-

0,

-

[ High oxygen tension ]

[Low oxygen tension]

cyanidesensitive

-

alternative oxldase cyanide insensitive

o2

Electron transport chains involved in the oxidation of glucose.

42

PAT M. GOODWIN AND CHRIS ANTHONY

In P . aeruginosu, the oxidation of ubiquinol is by way of the cytochrome he, complex which is subsequently oxidized by a typical periplasmic cytochrome c, the terminal oxidase being a cytochrome co (Matsushita et ul., 1982b). In all the other bacteria the ubiquinol is oxidized directly by a ubiquinol oxidase; in Acinetohacter, Escherichia and Klebsiella the oxidase is cytochrome ho in oxygen-sufficient conditions but cytochrome bd in oxygen-deficient conditions (van Schie el al., 1985; Beardmore-Gray and Anthony, 1986; Ameyama el ul., 1987; Matsushita et al., 1987; Smith et ul., 1990; Juty et ul., 1997). As is often the case. the situation with the acetic acid bacteria is complex. In Gluconobacter, which only grows in conditions of high aeration, there is usually a typical cytochrome ho (Matsushita et af., 1987). At low pH, however, a second ‘cyanide-insensitive’ and non-energygenerating pathway appears to operate. An intermediate in this pathway may be the cytochrome c subunit (subunit 11) of the membrane alcohol dehydrogenase (Matsushita et ul., 1989c, 1994). Many of the systems for glucose oxidation shown in Fig. 15 operate in reconstituted membrane systems and can generate a protonmotive force; this is consistent with observations that glucose oxidation by way of membrane glucose dehydrogenase leads to increased growth yields (van Schie et a f . , 1985, 1987a,b,c; Mueller and Babel, 1986; Neijssel et al., 1980; Adamowicz el ul., 1991; Neijssel and Demattos, 1994).

7. THE PHYSIOLOGICAL FUNCTIONS OF THE QUINOPROTEIN DEHYDROGENASES I t might be expected that the physiological function of enzymes that oxidize common growth substrates would not merit much discussion, but for the PQQ-containing quinoprotein dehydrogenases this is not the case. To summarize the following discussion in advance, it can be concluded that during growth on methanol and other alcohols as the sole source of carbon and energy the relevant periplasmic dehydrogenases usually play a key role in energy production, catalysing oxidation of the alcohol to the corresponding aldehyde, which is then either further oxidized or assimilated into cell material. In the case of the acetic acid bacteria, however, the PQQ-dependent alcohol and glucose dehydrogenases catalyse the first step in the incomplete oxidation of the energy source and most of the products of these oxidations are released into the growth medium. The roles of the glucose dehydrogenases in other bacteria are varied and often a matter of debate, particularly in the case of the enteric bacteria.

PQQ AND QUINOPROTEINS

43

7.1. The Role of Periplasmic Ouinoproteins that oxidize Alcohols 7.1.1. Methanol dehydrogenase

Methylotrophic bacteria are able to grow on reduced carbon compounds such as methane or methanol. Methane is produced in anaerobic environments by methanogenic bacteria and this provides a substrate for growth of methanotrophs at the aerobic surface. As well as being a product of methane oxidation, methanol arises in nature by hydrolysis of methyl ethers and esters present in pectin and lignin which are structural components of plants; methylotrophs growing on methanol are therefore abundant on the surfaces of leaves and in soil and water (Anthony, 1982). Methanol dehydrogenase is the only enzyme present in methylotrophic bacteria which is able to catalyse the oxidation of methanol to formaldehyde and it is therefore essential for growth on methanol or on methane. Although methanol dehydrogenase can also oxidize formaldehyde to formate this is unlikely to be important in vivo; indeed, some bacteria contain a regulatory modifier protein, the M-protein, which prevents this from happening by decreasing the affinity of the enzyme for formaldehyde (Long and Anthony, 1991). Every molecule of methanol that is used by the bacteria during growth on methane or methanol is oxidized to formaldehyde by methanol dehydrogenase and this step results in ATP production but not in production of NADH. Many methylotrophs are therefore unusual in having growth that is limited by reductant (NADH) rather than by ATP availability, a conclusion of considerable importance when predicting growth yields on methanol and methane (Anthony, 1986). During growth of methylotrophs on ethanol, methanol dehydrogenase may also be responsible for oxidation of this growth substrate to acetaldehyde (Dunstan el al., 1972a; Anthony, 1982).

I . 1.2. T?lpe I and Type 11 Periplasmic Alcohol Dehj)drogenases The role of the type I quinoprotein alcohol dehydrogenase in P . aeruginosa and P . putida is straightforward; it is induced during growth on ethanol or other short-chain alcohols when it is the key enzyme for production of energy and precursors for carbon assimilation (Gorisch and Rupp, 1989; Schrover et al, 1993; Toyama et al., 1995). Although the affinity of this enzyme for methanol is low, in Rhodopseudomonas acidophila it can also function in energy production during anaerobic growth on methanol in the light; in this case it has a role in the production of reducing power by reverse electron transfer (Anthony, 1982).

44

PAT M. GOODWIN AND CHRIS ANTHONY

Comamonas testosteroni grows slowly on ethanol or butanol, oxidizing them by means of an NAD-linked ethanol dehydrogenase. In these conditions the apoenzyme of the type I1 alcohol dehydrogenase (a quinohaemoprotein) is induced; this contains haem but not PQQ, which cannot be synthesized by this organism. However, when PQQ is included in the growth medium the active holoenzyme is produced and the growth rate increases, implying that the PQQ-dependent alcohol dehydrogenase takes over the main role of ethanol oxidation (Groen et a f . , 1986). By contrast with C‘. testosteroni, in P . putida the type I1 dehydrogenases, which are induced during growth on butanol or glycerol, are produced in the fully active form containing PQQ (Toyama el a f . , 1995).

7.2. The Roles of Glucose Dehydrogenase in Acinetobacter, Pseudomonads and Enteric Bacteria

The first step in glucose metabolism by bacteria usually involves uptake across the cytoplasmic membrane. This occurs either by the phosphotransferase system, when transport is coupled with phosphorylation of the glucose to glucose 6-phosphate, or by a glucose transporter, the glucose then being phosphorylated in the cytoplasm. There are three main pathways for subsequent metabolism - glycolysis, the Entner-Doudoroff pathway or the hexose monophosphate pathway (Fig. 16). In some bacteria all three pathways can operate and in these cases glycolysis is the major route for glucose catabolism whereas the Entner-Doudoroff pathway is used mainly for gluconate metabolism. Although the hexose monophosphate pathway can effect the complete oxidation of glucose, its main function is usually to provide Cs sugars and NADPH for biosynthesis. In organisms which contain an active glucose dehydrogenase, there is a fourth variant for glucose metabolism sometimes referred to as the direct (non-phosphorylating) oxidation pathway, glucose being oxidized in the periplasm (Fig. 16). This route is widely distributed among Gram-negative bacteria but a major bioenergetic advantage has not been obvious for this pathway and it was often referred to as a ‘dissimilatory by-pass’, expressing the apparent inefficient use of glucose. Operation of this pathway is evident by excretion of gluconate or 2-ketogluconate into the medium, but whereas the functional significance of this in some bacteria is clear (e.g. acetic acid bacteria, Section 7.3). in other cases it is debatable. Indeed, a mutant of Pscwdomonas cepuciu lacking glucose dehydrogenase grew as well as the wild-type on glucose as the sole carbon source (Lessie et al., 1979). In the natural environment, however, the presence of glucose dehydrogenase may give bacteria an advantage over competitors for a variety of reasons and this is discussed below.

45

PQQ AND QUINOPROTEINS Glucose

Glucose 8 P

-

EmbdenMeyerhof pathway (glycolysls)

Gluconate

6-Phosphogluwnate

2-Ketogluconate

t 2ketogluconate 8-P

Entner-Doudoroff pethway

Hexose monophosphate pathway

Figure 16 Alternative pathways for glucose metabolism. The abbreviation Pts refers to the PEP-linked phosphotransferase system.

1.2.1. Glucose Dehydrogenase in Pseudomonads Members of the genus Pseudatnonas are typically found in soil and water and are characterized by their ability to use a large variety of substrates as carbon and energy sources. They are aerobes, although some can grow in the absence of oxygen, using nitrate as the terminal electron acceptor. Most lack a complete glycolytic pathway and the Entner-Doudoroff pathway is the main route for glucose metabolism. The initial step in glucose metabolism usually involves uptake of glucose into the cell, followed by phosphorylation to glucose 6-phosphate (Fig. 16). Alternatively, glucose can be directly oxidized by glucose dehydrogenase to gluconic acid, which may then be further oxidized to 2-ketogluconic acid. As a result, acid accumulates in the growth medium and this is a diagnostic feature of the pseudomonads. The gluconate and 2-ketogluconate may be subsequently taken up by the bacteria and metabolized by the EntnerDoudoroff pathway.

46

PAT M. GOODWIN AND CHRIS ANTHONY

In P. aeruginosa the affinities of these two systems for glucose have been ~ the glucose uptake system and measured, the apparent K , being 8 p for 1 mM for glucose dehydrogenase; the latter is inducible and only operates when there is excess glucose in the growth medium (Ng and Dawes, 1973; Whiting et al., 1976). In natural environments, when glucose availability is limited, it would clearly be advantageous for P. aeruginosa to take up glucose as rapidly as possible by the high-affinity system and then convert it to glucose 6-phosphate. The significance of glucose oxidation by glucose dehydrogenase under conditions of glucose excess is not clear, although there is evidence that it can generate a protonmotive force in membrane vesicles (van Schie et d., 1985). Furthermore, a mutant of P. aeruginosa defective in glucose 6-phosphate dehydrogenase could grow aerobically using glucose as the sole source of carbon and energy, demonstrating that, if necessary, all the glucose required for growth may be metabolized via glucose dehydrogenase followed by uptake of the oxidation products (Hunt and Phibbs, 1983). The ability of the wild-type to rapidly oxidize glucose to gluconic acid, producing extra energy for growth, could be advantageous when it is growing in the presence of excess glucose. In other pseudomonads glucose dehydrogenase can operate during growth in the absence of glucose. The enzyme has a broad substrate specificity and may enable energy to be produced from the oxidation of sugars which cannot be used as carbon sources. This has been demonstrated in P. putida, which can oxidize xylose, but cannot use it as a carbon source. Thus, when xylose was added to a chemostat culture growing under conditions of limiting carbon source (glucose or lactose), xylonolactone and xylonate formed and the growth yields, Yglucose and YIaclale,increased, indicating that additional energy for growth can be provided by the oxidation of xylose (Hardy e f al., 1993). In the natural environment this versatility could give an advantage over competitors. A role for glucose dehydrogenase in some bacteria, including P. crpuciri and other soil pseudomonads, has recently been convincingly demonstrated by Goldstein, who has shown that such bacteria can make phosphate available from rock phosphate ore (Goldstein, 1995). This is particularly important in the region immediately surrounding the roots of plants (the rhizosphere) in arid and semi-arid soils where calcium phosphates provide a significant source of phosphate. There is evidence that this ‘mineral phosphate-solubilizing’ phenotype is a result of acidification of the periplasmic space and surrounding medium by the direct oxidation of glucose or other aldose sugars (produced by plant roots), the first step of which is catalysed by the membrane glucose dehydrogenase; the key acidic products are gluconic acid and 2-ketogluconic acid, which is particularly important in this context because it has the lowest pK of any organic acid produced by bacteria.

PQQ AND QUINOPROTEINS

47

7.2.2. Glucose Dehydrogenase in Acinetobacter calcoaceticus Acinetohacter species are aerobic chemoheterotrophs and, like the pseudomonads, they are found in soil and water and can use a wide range of substrates as carbon and energy sources. Some species can grow on glucose, metabolizing it by way of the Entner-Doudoroff pathway; others cannot do so, most strains of Acinetohacter calcoaceticus falling into this category. These strains usually synthesize an active glucose dehydrogenase and under certain growth conditions (for example carbon-limited growth on a mixture of glucose and acetate) they oxidize glucose to gluconate, which accumulates in the growth medium. The oxidation of glucose is associated with the formation of a protonmotive force and ATP synthesis and it has therefore been suggested that it functions as an auxiliary energy-generating system (van Schie et al., 1987~).This is consistent with the observation that when glucose was added to a chemostat culture of A . calcoaceticus growing increased (Mueller and Babel, 1986). on acetate, the yield, Yacetate, In addition to the membrane-bound enzyme, A . calcoaceticus contains a soluble periplasmic glucose dehydrogenase, the only known example of such an enzyme (Section 3.3.2). There is no evidence that it is involved in electron transport but it does have a high affinity for PQQ and it has been suggested that this enzyme functions as a PQQ carrier, accumulating PQQ derived either from the external medium or endogenously, and then transferring it to the membrane-bound apoenzyme on the outer surface of the cytoplasmic membrane (Matsushita et al., 1995a).

7.2.3. Glucose Dehydrogenase in Klebsiella pneumoniae Klebsiella pneumoniae is a facultative anaerobe capable of nitrogen fixation. It is found in soil and water and metabolizes glucose mainly via the phosphotransferase system and the glycolytic pathway. However, during aerobic growth in the presence of excess glucose, gluconate and 2-ketogluconate are sometimes excreted into the growth medium (Hommes et al., 1985; Buurman et al., 1994). This occurs when there is a high energy demand on the cell and it has been suggested that the function of glucose dehydrogenase is to provide an additional contribution to the protonmotive force and ATP synthesis (Hommes et al., 1985). However, there is evidence that in some situations (for example, when flux through glyceraldehyde 3-phosphate is limited by low phosphate availability), it provides an alternative source of energy, replacing glycolysis and the TCA cycle (Buurman et al., 1994). It is possible that sometimes oxidation of glucose by way of glucose dehydrogenase could provide extra energy, whereas at other times it could be an alternative energy-producing system; further work is needed to clarify

48

PAT M. GOODWIN AND CHRIS ANTHONY

this situation. It would also be of interest to ascertain if the enzymes of the Entner-Doudoroff pathway are induced when K . pneumoniae is grown under conditions of glucose excess. This would be comparable with growth of E. coli in excess glucose in the presence of PQQ, which results in the formation of active glucose dehydrogenase and induction of the enzymes of the Entner-Doudoroff pathway (Fliege et al., 1992). We speculate that an additional role for the direct oxidation of glucose in enteric bacteria might be to help provide protection against oxygen inactivation during transfer from aerobic growth to anaerobic fermentative growth. Many of the fermentative enzymes (e.g. pyruvateeformate lyase) are oxygen-sensitive, but are induced before conditions become completely anaerobic; they must, therefore, be protected from the remaining oxygen. To achieve this it is essential to have a source of electrons for rapid electron transfer to consume oxygen by way of the high-affinity oxidase (cytochrome hd) which is also induced during these conditions. Glucose dehydrogenase, which supports a high rate of respiration, could provide this. There is some evidence for such a role for glucose dehydrogenase in protecting the oxygensensitive pyruvate metabolism and nitrogenase in conditions supporting microaerobic nitrogen fixation in K . pneumoniue. In these conditions the yield of dinitrogen fixed per mole of glucose consumed is greater than in strictly anaerobic conditions due to the induction of cytochrome bd which has a very high affinity for oxygen. The electron transport chain involving this oxidase fulfils two functions; one is to provide ATP for nitrogenase function and the other is to remove trace amounts of inhibitory oxygen (Smith et ul., 1990; Hill et al., 1990; Juty et al., 1997). In whole cells glucose is an excellent substrate for electron transport by way of the cytochrome hd oxidase during microaerobic respiration and for nitrogen fixation in these conditions (Juty et al., 1997). This respiration is likely to be by way of glucose dehydrogenase and unlikely to involve the glycolytic enzymes, but some rigorous experiments with appropriate mutants are needed to confirm this. It would also be of interest to determine if synthesis of the glucose dehydrogenase in K . pneumoniae is regulated in the same way as that of E. coli, where the glucose dehydrogenase structural gene has two promoters, one regulated negatively by cyclic AMP and the other regulated positively by oxygen (Yamada et ul., 1993a).

7.2.4. Glucose Dehydrogenase in E. coli Escherichiu coli is a facultative anaerobe found in the intestines of mammals and also in water, particularly sewage effluent. It usually metabolizes glucose by way of the phosphotransferase transport system followed by glycolysis and, in aerobic conditions, the TCA cycle. It cannot synthesize

PQQ AND QUINOPROTEINS

49

PQQ but does make the membrane-bound glucose dehydrogenase apoenzyme and in the presence of PQQ this can be converted to the active holoenzyme both in vivo and in vitro. PQQ can act as a chemoattractant for E. coli (de Jonge ef al., 1996) and under conditions of low phosphate availability the PhoE porin, which is thought to be involved in PQQ uptake, is induced. Active enzyme may, therefore, form when E. coli is growing naturally in aquatic environments, alongside bacteria which excrete PQQ (Adamowicz et al., 1991; Nickerson and Aspedon, 1992). PQQ-dependent oxidation of glucose to gluconate by membrane vesicles of E. coli can generate a protonmotive force (van Schie et al., 1985). In this organism the enzymes of the Entner-Doudoroff pathway are not subject to catabolite repression, so if active glucose dehydrogenase is available, glucose could be metabolized by the glycolytic pathway and the Entner-Doudoroff pathway at the same time (Fliege et al., 1992). Metabolism by way of glucose dehydrogenase and the Entner-Doudoroff pathway is sufficient to support growth of E. coli, as shown by the ability of mutants lacking glycolytic enzymes to grow on glucose (Adamowicz et al., 1991; Fliege et al., 1992). Wild-type E. coli does not normally oxidize glucose to gluconate but when grown in continuous culture under conditions of glucose excess and, for example, phosphate limitation, gluconate accumulated if PQQ was supplied in the growth medium (Hommes et al., 1991). The biomass produced was similar to that of cultures grown in the absence of PQQ and the Yglucose decreased because of increased utilization of glucose. In contrast, when the bacteria were grown in the presence of PQQ under glucose limitation there was no accumulation of gluconate; this is because the affinity of glucose for the phosphotransferase system is much greater than that for glucose dehydrogenase. These results indicate that in cells grown in the absence of PQQ, under conditions of glucose excess, the respiratory chain was not working to full capacity; in the cells grown in the presence of PQQ, additional electron flow must have resulted from the oxidation of glucose to gluconate and presumably extra molecules of ATP were synthesized. In the natural situation in low-phosphate conditions where PQQ is available, the direct oxidation system and PhoE porin will be induced, and the Entner-Doudoroff pathway will operate (Fliege ef al., 1992; Yamada et al., 1993a). The resulting extra electron flow may provide a bioenergetic advantage. In addition to these possible roles in energy metabolism in E. coli, the respiratory chain involving glucose dehydrogenase and cytochrome bd might also play some role in respiratory protection during transfer from aerobic growth to anaerobic growth, when induction of oxygensensitive fermentative enzymes occurs, as suggested for K. pneurnoniae (Section 7.2.3).

50

PAT M. GOODWIN AND CHRIS ANTHONY

7.3. The Roles of Alcohol and Glucose Dehydrogenases in the Membranes of Acetic Acid Bacteria

We have found the literature on these bacteria rather confusing and contradictory, and here we attempt to summarize some of the key points of the relevant physiology (for reviews of growth and metabolism of acetic acid bacteria, see Asai, 1968; Swings, 1991; Matsushita el af., 1994). Acetic acid bacteria grow in sugary, alcoholic environments and they are typically found on the surfaces of leaves, fruits and alcoholic beverages. Most of the earlier work has concentrated on their use in vinegar production and as spoilers in beers and wines (Swings, 1991). They have subsequently been exploited for microbial transformations that depend on their characteristic of catalysing incomplete oxidations, many of which are catalysed by quinoproteins (Asai, 1968; Matsushita et al., 1994). Examples of industrial uses include the production of acetic acid from ethanol, gluconic acid from glucose, and sorbose from sorbitol. In the absence of added energy sources acetic acid bacteria are unable to grow on complex media such as peptone, nutrient broth or yeast extract, but they grow well at low to neutral pH when the medium includes an energy source such as ethanol, glucose or glycerol. Some strains are also able to use these substrates as a sole source of carbon and energy. The acetic acid bacteria obtain their energy from the incomplete oxidation of ethanol or glucose. All acetic acid bacteria (Acetobacter and Gluconohacter) have a type 111 alcohol dehydrogenase (quinohaemoprotein) which oxidizes ethanol to acetaldehyde. A membrane aldehyde dehydrogenase (Section 4) then oxidizes the acetaldehyde to acetic acid, which is excreted into the growth medium. Sometimes the acetic acid can then be transported back into the cell and further metabolized. Most Gluconohacter strains can also obtain energy from the incomplete oxidation of glucose to gluconate, catalysed by the membrane-bound glucose dehydrogenase. The gluconic acid is sometimes further oxidized in the periplasm to 2-ketogluconic acid by a membrane-bound flavoprotein dehydrogenase. Gluconohacter do not have a TCA cycle, and metabolize sugars (including gluconate) by way of the pentose phosphate pathway (Fig. 16). A rather unusual situation is found in Acetohacter diazotrophicus, which is remarkable in being able to fix atmospheric dinitrogen while using glucose or ethanol as sole source of carbon and energy (Stephan et al., 1991; Galar and Boiardi, 1995). Nitrogen fixation imposes a heavy demand for energy on the cell and there is evidence that in A . diazotrophicus synthesis of glucose dehydrogenase is three- to four-fold higher when cells are grown in batch culture under nitrogen-fixing conditions than when an alternative nitrogen source is present in excess (Galar and Boiardi, 1995). Gluconate accumu-

51

PQQ AND QUINOPROTEINS

lated in the growth medium during the lag and early exponential phases, demonstrating that glucose dehydrogenase was responsible for oxidizing glucose. It was suggested that this oxidation provides an ancillary energygenerating system during nitrogen fixation and also that it might play a role in protecting nitrogenase from oxygen (Stephan et al., 1991; Galar and Boiardi, 1995), as suggested for K . pneumoniae (Section 7.2.3).

8. SYNTHESIS OF PO0 8.1. Origin of PO0 Backbone

The most important work on the origin of the PQQ backbone has come from analysis, using NMR spectrometry, of the PQQ produced when Hyphoniicrobium X and M . extorquens AM1 were grown on labelled substrates. These results demonstrated that PQQ is derived from glutamate and tyrosine, both amino acids being incorporated intact (Houck et al., 1988, 1989, 1991; van Kleef and Duine, 1988; Unkefer, 1993; Unkefer et al., 1995). Thus, the tyrosyl side chain provides the six carbon atoms of the orthoquinone ring of PQQ and the pyrrole-2 carboxylic acid moiety is derived from internal cyclization of the amino acid backbone of tyrosine; the remaining five carbon atoms are from glutamate (Fig. 17). These observations led to proposals of routes for conversion of tyrosine and glutamate to PQQ (see Unkefer, 1993 for review), but there is no direct chemical or biochemical evidence to support them. In the meantime the genetics of PQQ biosynthesis was under investigation, and a small polypeptide of 23-39 amino acids was shown to be involved in PQQ production in a number of bacteria. These polypeptides have a high identity and contain a conserved motif with glutamate and tyrosine residues separated by three other residues (Fig. 18). Site-directed mutagenesis was used to construct COOH

COOH

I

,H

\ H*

Glutamate

PQQ

Figure 17 The origin of the carbon atoms in PQQ.

Tyrosine

52

PAT M. GOODWIN AND CHRIS ANTHONY MQWTKPmDLRIGFEVTMYFEAR

I

l

-

l

I Ill I

M WKKPAFIDLRLGLEVTLYISNR

I

I

I

I Ill I

MKWAAPIVSE ICVGMEVTSYESAEID’I‘FN

I

l

l

I

Ill I

MYRQHPSHPPQRSNFMTWSKPAYTDLRI GFEVTMYFASH 1 1 I I Ill I MM WTKPEVTEMRFGFE VTMYVC NF7

,4 calcoocrticus PqqlV 24 amino acids K. pneumoniae PqqA 23 amino acids

hf. exiorquens PqqA 2 9 amino acids 1’. ,fluore.vcensPqqA 39 amino acids hf ,flagellrrlurn PqqD 24 ~IIIIIIIOacids

Figure 18 Alignment of the amino acid sequences of the proposed polypeptide precursor of PQQ in different bacteria. Data are taken from Goosen el id. (1989), Meulenberg ei al. (1992). Morris el al. (1994), Schnider el a/. (1995) and Gomelsky ef ul. (1996).

mutants of A . calcoaceticus containing either aspartate in place of the conserved glutamate residue (Glu 16) or phenylalanine in place of the conserved tyrosine residue (Tyr20); in both cases PQQ synthesis was abolished (Goosen et al., 1992). However, replacement of a nearby glutamate (Glu22) with aspartate had no effect on PQQ synthesis. Although this does not rule out the possibility that the small polypeptide has, for example, a regulatory function, the finding that genes with similarity to those coding for peptidases are also essential for PQQ synthesis (Section 8.2) lends support to the hypothesis that this cofactor is derived from a peptide precursor.

8.2. The Genetics of PQQ Biosynthesis

Genes involved in PQQ synthesis have been identified in a number of bacteria (Table 5 ; Fig. 19). In most cases these genes have been isolated and sequenced following complementation analysis of mutants which require PQQ for growth on relevant substrates and their role in PQQ biosynthesis has been confirmed by demonstrating that mutagenesis of the cloned gene abolished its ability to complement the relevant mutation. The most detailed information is available for K . pneumoniae, A . calcoaceticus and M . extorquens. Klebsiella pneumoniae contains a cluster of six pqq genes -pqqABCDEF (Meulenberg et al., 1992). Upstream o f p q q A is the end o f another open reading frame, o r f x , which is not essential for PQQ synthesis. In A . cukoaceticus there are genes equivalent to p q q A (gene IV), pqqB (gene V). pqqC (gene I), pqqD (gene 11) and pqqE (gene 111) (Goosen et ul., 1989), arranged in the same order as in K . pneumoniae. No equivalent of

Table 5 Genes required for PQQ synthesis.

Gene symbol for strain shown (number of amino acids in predicted protein if known) A. K . pneumoniae calcoaceticus

M . extorquens (old

A (23 aa)

IV (24 aa)

B (308 aa)

V (303 aa)

C (251 aa)

I (252 aa)

D (92 aa)

I1 (94 aa)

E (380 aa)

I11 (331 or 384 aa)

M. organophilum

M. jlagellatum P. ,fluorexens

A @qqD) (29 aa)

D

D (24 aa)

A (39 aa)

PQQ precursor

B @qqG, moxO) (299 aa) CID @qqC, mo.uT) (372 aa) CID @qqB, moxV) (372 aa) E @qqA, moxCIP) (384 aa)

G

G (305 aa)

B (303 aa)

Transport

C

C

C

Oxygenase

symbol in brackets)

E. herbicola

Proposed function

B

F (761 aa)

Synthesis of metalcontaining cofactor Peptidase

orfx

Peptidase noncatalytic subunit Dipeptidase

orj R

A

(378 aa)

Data are taken from Goosen et al. (1987), Mazodier et al. (1988). B i d e et al. (1989), Meulenberg e f al. (1990, 1992), Liu et al. (1992), Morns et al. (1994), Schnider er al. (1995), Gomelsky et al. (1996), Springer et al. (1996), Turlin et al. (1996), Toyama e f al. (1997).

54

PAT M. GOODWIN AND CHRIS ANTHONY

A calcoacctrcus

K.pneumoniae h1 rxtorquens new

old M organophrlum

t'.fluorescens hL flagelloturn

Figure 19 Organization of PQQ genes in different bacteria. Data are taken from Goosen el ul. (1987). Mazodier et ul. (1988), Biville et ul. (1989), Turlin et a/. (1996), Meulenberg et ul. (1990; 1992), Morris et a/. (1994), Springer et ul. (1996), Toyama e / a / . (l997), Schnider et a/. (1993, Gomelsky et ul. (1996). The dotted lines indicate genes which are functionally equivalent.

pqqF has been reported, but downstream of gene 111 is an open reading frame, orfR, which is probably not essential for PQQ synthesis. Methylobacterium extorquens also contains genes equivalent to the pqqABCDEF genes of K . pneumoniae, but in this case they are situated in two separate clusters (Morris et al., 1994; Springer et al., 1996; Toyama et al., 1997). The nomenclature of the M . extorquens pqy genes is confusing; when they were first identified they were designated mox (methanol oxidation) genes; later, when their function in PQQ synthesis was demonstrated, they were called pqq genes and the labelling system used for the pqq genes of another methylotroph, Methylobacterium organophilum, was followed. However, recently it has been proposed that they should be renamed, to correspond to their functional equivalents in K . pneumoniae (Toyama et al., 1997). The new and old nomenclature is shown in Table 5 and Fig. 19, but in the text only the new system is used. One cluster of M . extorquens p q y genes is adjacent to the mxh genes which are involved in the regulation of methanol dehydrogenase synthesis. It contains pqqAB and E . Between p q y B and pqqE is a gene encoding a protein which is similar at the N-terminal end to PqqC of K . pneumoniae (a polypeptide of 29.7 kDa) and at the C-terminal

PQQ AND QUINOPROTEINS

55

end to PqqD of K . pneumoniae (a polypeptide of 10.4kDa). Expression of the M . extorquens DNA in E. coli indicated that this region encodes a single protein of 42 kDa and it appears that in M . extorquens the pqqC and D genes have fused to give p q q C / D and the resulting protein can carry out the functions of both PqqC and PqqD. In M . extorquens there are another two pqq genes -pqqF and pqqC, which are linked; an equivalent ofpqqC has not yet been described in K . pneumoniae or A . calcoaceticus. A model for PQQ synthesis is shown in Fig. 20. The putative polypeptide precursor of PQQ is encoded by the pqqA gene of K . pneumoniae and its functional equivalents in other bacteria. Processing of this precursor must involve several steps, including cleavage by a specific protease or proteases, formation of the PQQ backbone from the relevant glutamate and tyrosine residues, and formation of the quinone groups. Analysis of those pqq genes which have been sequenced indicates that PqqF of K . pneumoniae and PqqF and PqqG of M . extorquens have similarity to a family of divalent cationcontaining endopeptidases which are involved in processing small peptides (Meulenberg et al., 1992; Springer et a f . , 1996). The two PqqF proteins seem to be members of different subfamilies - the K . pneumoniae analogue is most closely related to the subfamily which contains pitrilysin, a periplasmic oligopeptidase found in E. coli, while the M . extorquens protein belongs to the subfamily containing mitochondria1 processing peptidases. Members of this subfamily form heterodimers containing two similar subunits, one of which lacks the catalytic site. The predicted PqqG protein shows some identity to the C-terminal half of two members of the mitochondrial processing peptidase family, and it has been suggested that PqqF (which is predicted to contain the catalytic site) and PqqG (which does not appear to contain the catalytic site) may associate to form a heterodimer (Springer et al., 1996). Interestingly, the orfX located upstream of the K . pneunioniae pqq operon and the orfR located downstream of the A . calcoaceticus pqq gene cluster both encode proteins similar to a human dipeptidase. However, mutations in these genes do not abolish the ability to synthesise PQQ, indicating that they are not essential for PQQ biosynthesis (Goosen et al., 1989; Meulenberg et al., 1990, 1992). The predicted PqqE proteins of K . pneumoniae and M . extorquens contain a CxxxCxYC motif similar to that found in MoaA, which is involved in the biosynthesis of the molybdopterin cofactor of E. coli and NifB, which is involved in the biosynthesis of the iron-containing cofactors of nitrogenases (Toyama et al., 1997). These proteins probably function in the donation of metal atoms to the relevant cofactor and the conserved CxxxCxYC sequence may be a metal-binding site (Menendez et al., 1995). This suggests that an unknown metal-binding cofactor may be involved in PQQ synthesis. PqqC probably catalyses the last step in PQQ biosynthesis; pqqC mutants of K . pneumoniae and M . exforquens accumulated an intermediate which

56

PAT M. GOODWIN AND CHRIS ANTHONY

/ -

E-Y-

and formation of PQQbackbone

I

Peptide precursor PqqF (peptldase PqqG (dipeptidase) ')'WE 7PqqD

precursor Formationof

Signal

Figure 20 A model for PQQ synthesis in Merhylobocteriuni exrorquetis. A signal (which could, for example, be formaldehyde) is transmitted by way of MxbD and MxbM, leading to the activation of the pqqA promoter; this is also regulated by MxaB. The polypeptide product of p q q A is processed in the cytoplasm to form PQQ and this probably involves the PqqCDEFG proteins. PQQ is then transported into the periplasm; PqqB, which is predicted to be a cytoplasmic protein, is necessary for this.

could be converted to PQQ in vifro on addition of an extract containing PqqC or PqqC/D respectively (Velterop c'f ul., 1995; Toyama r f a/., 1997). Attempts to purify the intermediate were unsuccessful. Aerobic conditions and NAD(P)H were both essential for its conversion to PQQ, suggesting involvement of an oxygenase in the formation of the quinone groups of PQQ. However, if pqqC does encode an oxygenase it is not similar at the amino acid sequence level to any known monooxygenase or dioxygenase, and i t is not obvious how this step would be catalysed in Purucoccus or Hyphomicrohium growing anaerobically on methanol with nitrate.

PQQ AND QUINOPROTEINS

57

PqqB may be involved in transport of PQQ into the periplasm. This was suggested by Velterop et al. (1995) and Gomelsky et al. (1996). who demonstrated that K . pneumoniae pqqB mutants and mutants of Methylobacillus jlagellaturn defective in the equivalent gene, pqqG, accumulated PQQ intracellularly but did not excrete it into the growth medium. The PqqB protein does not contain any hydrophobic regions so it is unlikely to be a membrane protein directly involved in transport, but it might be required as a PQQbinding protein or for modification of a membrane protein involved in PQQ transport.

8.3. Does Escherichia coli contain pqq Genes?

Escherichia coli is unable to synthesize PQQ, and produces glucose dehydrogenase as an apoenzyme, the holoenzyme only forming on addition of exogenous PQQ. There have been several reports that E. coli strains carrying heterologous pqq genes can synthesize PQQ and can therefore make active glucose dehydrogenase. However, the number of genes needed to obtain PQQ synthesis varies, depending on the source of the heterologous DNA. When K . pneumoniae DNA was used all six pqq genes seemed to be needed for PQQ production by E. coli (Meulenberg et al., 1990, 1992) although later work indicated that small amounts of PQQ could be made in the absence of pqqB (which is possibly involved in PQQ transport) or pqqF (which encodes a peptidase) (Velterop et al., 1995). This is consistent with the report that four A . calcoaceticus pqq genes (IV, I, I1 and 111, equivalent to pqqA, C , D and E of K . pneumoniae) were required for PQQ synthesis by E. coli (Goosen et al., 1989). Escherichia coli does contain a gene which is functionally equivalent to the pqqF genes of K . pneumoniue and M . extorquens. Evidence for this was obtained by the demonstration that a 7.3 kb fragment of E. coli DNA containing an ORF with low identity to the pqqF gene of M . extorquens (Springer et al., 1996) can complement a mutant of Methylobacterium organophilum which is defective in the equivalent gene (Turlin et al., 1991, 1996). This might explain why it was not essential to provide the pqqF gene of K . pneumoniae in order to obtain PQQ synthesis in E. coli carrying the K . pneumoniae pqq genes. The E. coli fragment also contained an ORF with low identity to the pqqG gene of M . extorquens (Springer et al., 1996) and complemented a M . organophilum mutant defective in the equivalent gene. Thus, E. coli contains genes which can function in the same way as the PqqF and PqqG proteins of M . extorquens. There is no evidence from sequence analysis that E. coli contains a gene with similarity to pqqB, but it is possible that another E. coli protein may be able to substitute for it in functional terms.

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PAT M. GOODWIN AND CHRIS ANTHONY

The work described above ihdicates that E . coli can synthesize PQQ, but only if provided with at least four heterologous PQQ genes. It is difficult to reconcile this with reports suggesting that E . coli strains carrying single genes from Erwiniu herhicolu (Liu et ul., 1992) and Pseudomonas crpacia (Babu-khan et al., 1995) produced PQQ. However, in neither case was PQQ measured directly - synthesis was inferred because the E. coli strains carrying the heterologous DNA produced acid from glucose. The gene from Erwiniu hrrbicolu had a high degree of identity with pqqE of K . pneumoniae but the gene from P. cepacia (guhY) was not similar to any previously described pyy genes. It is predicted to encode a 14.2 kDa protein which has some regions similar to parts of the membrane bound components of periplasmic permeases such as HisQ and GlnP. I t was suggested that GabY might catalyse PQQ production by a novel pathway, cause production of a cofactor which could replace PQQ in the apo-glucose dehydrogenase, or activate cryptic pqq genes. Biville and colleagues (1991) also suggested that E. coli contains cryptic pyy genes. They were working with an E. coli strain lacking the phosphotransferase system (PTS), and therefore unable to metabolize glucose via the glycolytic pathway. This mutant could, however, grow on glucose if PQQ was added to the medium - the holoform of glucose dehydrogenase was produced and thus glucose could be oxidized to gluconic acid and catabolized via the Entner-Doudoroff pathway. When these PTS- strains were plated onto glucose minimal medium (lacking PQQ) spontaneous ‘revertants’ arose, which could oxidize glucose to gluconic acid, and PQQ and low glucose dehydrogenase activities were detected in membrane preparations. The ability to grow on glucose could not, therefore, be explained by reversion of the original PTS mutations. The authors concluded that E. coli contains all the genetic information required for PQQ synthesis and that the spontaneous mutation resulted in expression of these genes. However, the data supporting this conclusion have recently been challenged (Matsushita e f al., 1997). Analysis of the E. coli genome indicates that it does not contain any genes with high similarity to pyyBCDE of M . extorquens, suggesting that there is no cryptic pyq operon comparable with those described in the well-characterized PQQ-producing bacteria. However, proteins which do not share similarity at the amino acid level may still share functions; for example, ActA of Listeriu monocytogenes and IcsA of Shigellu jiesneri have virtually no sequence similarity, but carry out similar functions in initiating actin assembly (Strauss and Falkow, 1997). The possibility that E. coli contains genes which have no sequence similarity to the known pqq genes, but can carry out the same functions, needs to be explored, as does the suggestion that there might be a novel pathway for PQQ synthesis in E. coli. This may

PQQ AND QUINOPROTEINS

59

clarify the conflicting results concerning the ability of this organism to produce PQQ.

9. REGULATION OF SYNTHESIS OF PQQ AND QUINOPROTEIN DEHYDROGENASES 9.1. Synthesis of PQQ and Apoenzymes is not Coordinated

Synthesis of PQQ and the apoenzyme dehydrogenases is not necessarily coordinated and sometimes enzyme activity in vitro is several fold higher when PQQ is added to the assay mixture than in its absence (van Schie et al., 1984; Hommes et al., 1989). An investigation of PQQ synthesis and production of the apo- or holoenzymes of glucose dehydrogenase, methanol dehydrogenase and quinoprotein alcohol dehydrogenase in a variety of bacteria indicated that PQQ synthesis is not essential for apoenzyme production (van Kleef and Duine, 1989). By contrast, PQQ is only made when the quinoprotein is also being synthesized.

9.2. Regulation of PQQ Synthesis

It has been suggested, on the basis of DNA sequence analysis, that the pqqA and B genes of K . pneumoniae and M . extorquens and the equivalent genes in A . calcoaceticus are co-transcribed. Prediction of the mRNA structures suggests that a hairpin can form between the sequence for pqqA and pqqB, which might cause transcription termination. Analysis of the mRNA transcripts of M . extorquens confirmed that the major mRNA formed from transcription from the pqqA promoter was approximately 240 bases, and encoded only PqqA; a second, less abundant transcript was also detected, which encoded PqqA and PqqB (Ramamoorthi and Lidstrom, 1995). Furthermore, in K . pneumoniae expression of pqqA is 20-fold higher than that of pqqC or pqqE (Velterop et al., 1995). These observations are consistent with the suggestion that pqqA encodes the precursor of PQQ - it would be needed in larger amounts than the products of other pqq genes, which presumably have catalytic roles in processing the precursor. The promoters of the pqqA genes have not been conclusively defined, but possible -10 and -35 sequences have been identified in K . pneumoniae (CAATAT and TTGATC) and M . extorquens (CGATAT and TTGCAG) (Ramamoorthi and Lidstrom, 1995). The latter differs from the postulated promoter sequence (-10 TAGAA, -35 AAGACA) upstream of the mxaF

60

PAT M. GOODWIN AND CHRIS ANTHONY

operon, which encodes the methanol dehydrogenase and cytochrome CL structural proteins (Barta and Hanson, 1993; Chistoserdova et af., 1994). However, upstream of the putative pqqA promoter, at bases -54 to -48, is the sequence AGAAACG, which is similar to the septanucleotide sequence AGAAATG found upstream of mxaF and other methanol-regulated promoters in Merhylobacrerium organophifum (Xu et a f . , 1993) and it has therefore been suggested that mxaF and pqqA share some common regulators. This is supported by the observation that expression of pqqA is not regulated normally in mutants of M . extorquens defective in three genes essential for expression of the inxaF operon - mxbM and mxbD, which encode a twocomponent regulatory system, and mxaB, which encodes a positive regulator (Section 9.3.3). However, as noted above, synthesis of the apoenzyme and PQQ is not always coordinated and comparison of the level of expression of pqqA and the amounts of PQQ produced in these mutants indicated that regulation of pqqA transcription is not the only step at which synthesis is controlled (Ramamoorthi and Lidstrom, 1995). Whether the other factor(s) which regulate PQQ synthesis act at the transcriptional or post-transcriptional level has still to be determined. However, there is preliminary evidence that there is a weak C , -inducible promoter upstream of pqqF and that expression from this promoter is not under the control of the regulatory genes thought to control methanol dehydrogenase production (Springer et at., 1996). 9.3. Factors Affecting the Synthesis of the Quinoprotein Dehydrogenases

There is evidence from several studies that active methanol dehydrogenase, alcohol dehydrogenase and glucose dehydrogenase can be synthesized to quite high levels in the absence of their substrates, although a number of factors do affect their synthesis; for example, growth rate, the nature of the growth substrate or oxygen availability (Dunstan et af., 1972b; Ng and Dawes, 1973; O’Connor and Hanson, 1977; Roitsch and Stolp, 1985, 1986; de Vries et uf., 1988; van Kleef and Duine, 1989; Frebortova cr ul., 1997). Much of the early work is difficult to interpret because it was done using batch cultures and growth conditions were not well defined. The conclusions of some of the more detailed studies. using chemostat cultures, are summarized below. 9.3.1. Synthesis of’ Glucose Dehydrogenase In K . pneumoniae several factors affect expression of glucose dehydrogenase. During growth in chemostat cultures in the presence of excess glucose under

PQQ AND QUINOPROTEINS

61

potassium or phosphate limitation, glucose dehydrogenase activities were high; however, they were relatively low in glucose-limited cultures and in sulphate- or ammonia-limited cultures, despite the presence of excess glucose, indicating that synthesis was not necessarily regulated in response to the level of glucose in the medium (Hommes et al., 1985). Growth rate also influenced glucose dehydrogenase production, although the effect depended on the growth conditions - activity in potassium-limited cultures increased with increasing dilution rate, whereas in phosphate-limited cultures activity decreased with increasing growth rate. It has been suggested that glucose dehydrogenase plays a role in the generation of reducing power when there is a high energy demand on the cell (Hommes et al., 1985; Section 7.2.3) and it may be synthesized in response to this. However, the situation is clearly complex and further work is required to define precisely how changes in enzyme synthesis relate to the energy demand of the cell under different growth conditions. Most strains of A . calcoaceticus cannot grow on glucose, but they can make glucose dehydrogenase and oxidize glucose to gluconic acid, and this may provide an additional energy source during growth on other substrates (Section 7.2.2). Synthesis appears to be regulated by derepression rather than induction because active enzyme is made during growth on a variety of substrates, independent of the presence of glucose in the medium. However, activity does vary with growth rate, being high at low growth rates, when there is a high requirement for maintenance energy. It has therefore been suggested that, as with K. pneumoniae, glucose dehydrogenase synthesis is regulated in response to the energy status of the cells (van Schie e f al., 1988). By contrast, in P. aeruginosa glucose dehydrogenase is induced by glucose. Gluconate, the product of glucose oxidation, and glycerol are also inducers. However, during growth on a mixture of glucose and citrate, it is also regulated by catabolite repression (Midgley and Dawes, 1973; Ng and Dawes, 1973). Active enzyme is not produced when P. aeruginosa is grown anaerobically on glucose with nitrate as the terminal electron acceptor, but this is because PQQ is not made. The apoenzyme is induced, and active enzyme can be formed on addition of PQQ (van Schie et al., 1984). Escherichia coli only produces apo-glucose dehydrogenase, but there is evidence that it is induced by glucose. Levels are high in glucose-grown cells which are potassium-, phosphate- or sulphate-limited, and low in cultures grown on limiting glucose (Hommes el al., 1991). The glucose dehydrogenase structural gene, gcd, has been sequenced (Cleton-Jansen et al., 1990) and analysis of the upstream region indicates that there are two promoters. Studies using transcription fusions demonstrated that glucose induces transcription from the first promoter, which is also regulated negatively

62

PAT M. GOODWIN AND CHRIS ANTHONY

by cyclic AMP; the second promoter is regulated positively by oxygen (Yamada et al., 1993a).

9.3.2. Synthesis qf’ Methanol Dehydrogenase In Methylophilus methj~lotrophus,which is an obligate methylotroph able to grow only on methanol and methylated amines, methanol dehydrogenase activity is strictly controlled in response to the growth conditions. I t is maximally repressed during growth in oxygen-limited cultures and, when cells are grown at different dilution rates under methanol limitation. the activity decreases as the standing concentration of methanol in the growth medium increases (Greenwood and Jones, 1986; Jones et al., 1987; Southgate and Goodwin, 1989). However, methanol dehydrogenase does not catalyse the rate-limiting step in methanol oxidation, and the activity of the overall methanol oxidase system was high at all dilution rates. It has therefore been suggested that, when there are high levels of methanol in the medium, methanol dehydrogenase is repressed in order to prevent accumulation of formaldehyde, the toxic product of methanol oxidation. The methanol oxidase system is regulated by oxygen and methanol such that the energy demands imposed by the growth rate can be met. Repression of methanol dehydrogenase was also observed under similar conditions in Hyphomicrobiuni X (Duchars and Attwood, 1989). In Paracoccus denitriJicans, an autotrophic methylotroph which assimilates methanol by the ribulose bisphosphate pathway after oxidation to carbon dioxide, there is evidence for two mechanisms of regulation of methanol dehydrogenase synthesis (de Vries et al., 1988). There is a basal level of synthesis during growth on high levels of multicarbon compounds which are good growth substrates, whereas under conditions of carbon limitation or at low growth rates it is derepressed. It is also regulated by induction but the inducer is formaldehyde, the product of methanol oxidation, and not methanol itself. Autotrophic growth on methanol is not energetically favourable compared with growth on multicarbon substrates. The combination of repression/derepression of methanol dehydrogenase in response to the availability of other growth substrates, together with product induction (which requires that the substrate is present at high enough levels for a long enough time period for the concentration of the product to be high enough to initiate induction), would therefore ensure that, in the natural environment, the methanol dehydrogenase system is only synthesized to high levels when there is no preferred substrate available. Similar patterns of regulation, i.e. derepression at low growth rates and induction by methanol, have been observed in another autotrophic methylotroph, a Xanthohacter sp., and in Methylohacterium sp., but

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63

further work is needed to determine if formaldehyde rather than methanol is the inducer (O’Connor and Hanson, 1977; Roitsch and Stolp, 1986; Croes et al., 1991).

9.3.3. Molecular Mechanism of Regulation of Methanol Dehydrogenase Syn thesis More than 25 genes (mox genes) are required for methanol dehydrogenase synthesis (Tables 5 and 6). Seven are pqq genes (Section 8.2), three (mxaFGI) are structural genes and three (mxaAKL) are required for insertion of calcium into methanol dehydrogenase. Of the remainder, seven are thought to be regulatory genes and the rest are of unknown function. The structural genes mxaFGI are located in an operon, and in the Methylobacterium strains msaF and G are separated by mxaJ, which encodes a polypeptide of unknown function (Fig. 21). Using transcriptional fusions it has been shown that the mxaF promoter of the Methylobacterium strains is expressed at a high level during growth on methanol compared with growth on succinate and that it is positively regulated by msaB (Morris and Lidstrom, 1992; Xu et al., 1993). The mxaB gene also regulates p44A (Section 9.2). The promoters of two other mox genes m.xa W and mxcU, both of unknown function - are activated during growth on methanol, but this is not dependent on mxaB. Transcription of the mxaF operon of the Methylobacterium strains is also regulated by two pairs of sensor kinase-response regulator proteins, encoded by m s b D M and mxcEQ (Xu et al., 1993, 1995; Springer et al., 1997). Two-component regulatory systems are common in bacteria and involve sensing of a signal by the membrane protein kinase which is then autophosphorylated. It can then interact with the response regulator, resulting in activation of transcription at a specific promoter(s). Sequence analysis indicates that MxbD and MxcQ are membrane proteins belonging to the histidine kinase superfamily, although their putative periplasmic loops are quite different, suggesting that they respond to different signals. MxbM and MxcE are DNA-binding proteins of the response regulator family. The rnxbDM genes are probably co-transcribed and studies using mxhD transcriptional fusions showed that expression was considerably reduced in mxcE and mxcQ mutants, suggesting a hierarchical regulatory system. Thus, the mxcQE proteins would respond to a specific signal and switch on expression of m s b D M . The MxbD protein then presumably responds to a second signal, resulting in enhanced transcription of mxaF. The mxbD and M genes are also essential for expression of pqqA (Section 9.2), mxa W and mxcCJ; however, mxcQ and mxcE are not, indicating that there must be a ~

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PAT M. GOODWIN AND CHRIS ANTHONY

Tahle 6 Genes involved in methanol oxidation in Methylohacterium and P. denitrificans. Proposed function and location of gene product Structural genes

Organism

M. organopldum

P. dentrificans

mxaF

nixaF

mxaG

mxaC

rnxal

mxal

mscQE m.xhDM

VlSU Y X

nixhDM mxaB

mxciB

mxh N

nisb N

M. extoryuens

a subunit of

mxaF MDH (P) cytochrome CL ( P ) mxaG p subunit of mxal M D H (P)

Regulatory genes sensor kinase response regulator (M/C)

-

(C)

mxcQE

n1xuZ

(MI Insertion of Ca” into M D H

Other

? Ca2+ binding (P) mxaA

(C) (M)

mxaK mxuL

Third subunit of MDH or molecular chaperone (P) Unknown ( C ) Unknown Unknown Unknown (C) Unknown (P) Unknown Unknown Unknown

msaJ

mxnJ

mxa R

ni.xaR

mxuS

mxuS

tnxa W

nisuC nxaD m.w U

nisdR mxdS

(C), cytoplasm; (M), membrane; (P) periplasm; MDH, methanol dehydrogenase. Key references for M . extoryuens: Nunn and Anthony, 1988; Nunn et a/., 1989; Anderson et nl.. 1990; Morris and Lidstrom, 1992; Lidstrom el d , , 1994; Morris et a/., 1995; Amaratunga rt a/., 1997a,b; Springer et al., 1995, 1997. M . organophilum: Machlin and Hanson, 1988; Xu el al., 1993, 1995; P . denitrifcans: Harms el a/., 1987; van Spanning ef a / . , 1991; H a r m et a / . , 1993.

Figure 21 A model for the expression of methanol dehydrogenase in Merhylobacrerium exiorquens; it is probably similar in Paracoccus denirrrfcans and related bacteria. Signal 1 is transmitted by way of MxcQ and MxcE. leading to transcription of mxhDM. These genes encode a second signal transduction system which responds to signal 2, resulting in activation of the msaFJGI operon. The signals have not been identified, but one is likely to be methanol or formaldehyde. The m.uaF promoter is also controlled by MxaB and MxbN. The m.uaFJGI preproteins are transported into the periplasm where they are assembled into the proteins that are specifically involved in methanol oxidation - methanol dehydrogenase and cytochrome c L .The * indicates two possibl: steps at which PQQ may be inserted. At least three proteins (MxaA. MxaK and MxaL) are involved in incorporation of Ca-+ into methanol dehydrogenase and MxaC and MxaD may also be involved in this.

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PAT M. GOODWIN AND CHRIS ANTHONY

basal level of mxhDM expression, independent of mxcQE, which allows transcription of pqqA, mxa W and mxcU. In addition to mxaB, mxbDM and mxcQE, there is thought to be another regulatory gene - mxbN (Xu et al., 1995) and these six genes are all required for growth on methanol and for transcription of mxaF. I t is likely that there are additional regulatory genes, which are not essential for growth on methanol but are responsible for the fine-tuning of methanol dehydrogenase expression. Such genes have not yet been identified in the Methylobacterium strains, but in Paracoccus denitrijicans two putative regulatory genes which are not essential for growth on methanol have been described. These are mxaY and Z , which, with mxaX, are located upstream of the mxaF operon (Harms et al., 1993). The genes mxaY and X have significant similarity to m.ucQE of Methylobacterium orgmophilum (Xu et a / . , 1995) but, although the response regulator encoded by mxuX is essential for growth on methanol, the histidine kinase encoded by mxaY is not, indicating that an alternative sensor can replace MxaY. The mxaZ gene is not essential for growth on methanol either, but mxaZ mutants grow slowly and have reduced expression of the mxaF operon, indicating that this gene, which encodes a novel protein, is involved in the regulation of methanol dehydrogenase (Yang el nl., 1995). To date, genes equivalent to mxuB and m.uhDM of the Melhylohacterium strains have not been identified in P . denit r ifi cuns .

10. CONCLUDING REMARKS

Since the first isolation of PQQ more than 30 years ago considerable progress has been made in understanding the biochemistry of some of the PQQdependent dehydrogenases, particularly methanol dehydrogenase. Less is known about the synthesis of PQQ and its interaction with the apoenzymes to form the holoenzymes. The physiological function of methanol dehydrogenase and other alcohol dehydrogenases is relatively clear but in most bacteria the role of glucose dehydrogenase is not well understood. The description of glucose metabolism via this enzyme as ‘the dissimilatory pathway’ indicates inefficient use of glucose and, although we have speculated on its role in Section 7.2, we finish with a final, perhaps heretical, suggestion; that this enzyme had an important role to play in an ancestor of the presentday bacteria. providing a rapid means of metabolizing glucose, but that in many bacteria it may now be merely an ‘evolutionary relic’ which is occasionally useful.

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Work in the authors’ laboratory has been supported by the BBSRC and The Wellcome Trust.

Adachi, 0..Matsushita, K.. Shinagawa, E. and Ameyama, M. (1988) Enzymatic determination of pyrroloquinoline quinone with a quinoprotein glycerol dehydrogenase. Agric. Biol. Chern. 52, 208 1-2082. Adachi, O., Matsushita, K., Shinagawa, E. and Ameyama, M. (1990a) Calcium in quinoprotein methanol dehydrogenase can be replaced by strontium. Agric. B i d . Chem. 54, 2833--2837. Adachi, 0..Okamoto, K., Matsushita, K., Shinagawa, E. and Ameyama, M. (1990b) An ideal basal medium for assaying growth stimulating activity of pyrroloquinoline quinone with Acetohacter uceti IFO-3284. Agric. Biol. Chem. 54, 2751-2752. Adamowicz. M., Conway. T. and Nickerson, K.W. (1991) Nutritional complementation of oxidative glucose metabolism in Escherichiu coli via pyrroloquinoline quinonedependent glucose dehydrogenase and the Entner-Doudoroff pathway. Appl. Environ. Microhiol. 57. 2012-2015. Amaratungd, K . , Goodwin. P.M., O’Connor, D.C. and Anthony, C. (1997a). The methanol oxidation genes m.xaFJGIR(S)ACKLD in Methylobacterium extoryuens. FEMS Microhiol. Lett. 146, 3 1-38. Amaratunga, K., Goodwin, P.M., O’Connor, D.C. and Anthony, C. (1997b) Erratum to ‘The methanol oxidation genes mxaFJGIR(S)ACKLD in Methylobacterium extorquens’. FEMS Microbiol. Lett. 150, 175-177. Ameyama, M. and Adachi, 0. (1982) Alcohol dehydrogenase from acetic acid bacteria, membrane-bound. Methods Enzynol. 89, 450.-457. Ameyama, M.. Shinagawa, E., Matsushita, K. and Adachi, 0. (1981) o-glucose dehydrogenase from Gluconohucter suboxydans: solubilization, purification and characterization. Agric. B i d . Chern. 45. 851- 861. Ameyama, M., Nonobe, M., Hayashi, M., Shinagawa, E., Matsushita, K. and Adachi 0. (1985) Mode of binding of pyrroloquinoline quinone to apo-glucose dehydrogenase. Agric. Biol. Chcwi. 49. 1227-1231. Ameyama, M., Nonobe, M., Shinagawa, E., Matsushita, K., Takimoto, K . and Adachi, 0. (1986) Purification and characterization of the quinoprotein o-glucose dehydrogenase apoenzyme from Escherichiu coli. Agric. Biol. Chem. 50, 49-57. Ameyama, M., Matsushita. K.. Shinagawa. E. and Adachi, 0. (1987) Sugar-oxidizing respiratory chain of Gluconohucter suho.xyduns. Evidence for a branched respiratory chain and characterization of respiratory chain-linked cytochromes. Agric. Biol. Chem. 51, 2943 2950. Ameyama, M., Matushita, K., Shinagawa, E., Hayashi, M. and Adachi. 0. (1988) Pyrroloquinoline quinone: excretion by methylotrophs and growth stimulation for microorganisms. Biojuctors 1. 5 1-53, Anderson, D.J., Morris, C.J., Nunn, D.N., Anthony, C. and Lidstrom, M.E. (1990) Nucleotide sequence of the Methylobacterium e.utorqurn.s AM 1 nioxF and rno.xJ genes involved in methanol oxidation. Gene 90, 173- 176.

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Molecular Phylogeny as a Basis for the Classification of Transport Proteins from Bacteria, Archaea and Eukarya Milton H. Saier, Jr Department of Biology, University of’ California at San Diego. La Jolla. CA 92093-0116. USA

ABSTRACT Although enzymes catalyzing chemical reactions have long been classified according to the system developed by the Enzyme Commission (EC), no comparable system has been developed or proposed for transport proteins catalyzing transmembrane vectorial reactions. We here propose a comprehensive system, designated the Transport Commission (TC) system, based both on function and phylogeny. The TC system initially categorizes permeases according to mode of transport and energy coupling mechanism, and each category is assigned a one-component TC number (W). The secondary level of classification corresponds to the phylogenetic family (or superfamily) to which a particular permease is assigned, and each family is assigned a two-component TC number (W.X). The third level of classification refers to the phylogenetic cluster within a family (or the family within a superfamily) to which the permease belongs, and each cluster receives a three-component TC number (W.X.Y). Finally, the last level of categorization is based on substrate specificity and polarity of transport, and each entry is assigned a four component T C number (W.X.Y.Z). This system is based on the observation that mode of transport and energy coupling mechanism are fundamental properties of transport systems that very seldom transcend familial lines, but substrate specificity, being readily alterable by point mutations, is a superficial characteristic that often transcends familial lines. The proposed ADVANCES IN MICROBIAL PHYSIOLOGY VOL 40

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system has the potential to include all known permeases for which sequence data are available and has the flexibility to accommodate the multitude of permeases likely to be revealed by future genome sequencing and biochemical analysis. Major conclusions resulting from our classification efforts are described. The classification system, which will be continuously updated, is available on our World Wide Web site (http://www-biology.ucsd.edu/-msaier/ transport/titlepage.html). 1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 83 2. Considerations for the systematic classification of transmembrane solute 84 permeases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. Proposed transport protein classification system . . . . . . . . . . . . . . . . . . . . . . . 86 4. Diverse evolutionary origins of integral membrane transport protein families . 95 5. The major facilitator superfamily (MFS) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 107 6. The ATP-binding cassette (ABC) superfamily. . . . . . . . . . . . . . . . . . . . . . . . . . 109 121 7. Prokaryotic genome sequence analyses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8. Independent evolution of distinct transport modes and energy-coupling 124 mechanisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9. Proposed independent evolution of different channel and carrier families . . . 127 131 10. Conclusions and perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgement . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 131 Note added in proof . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 131 132 References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Abbreviutions. Standurd A, archaea; B, bacteria; CFTR, cystic fibrosis transmembrane conductance regulator; E, eukarya; MDR, multidrug resistance; PEP, phosphoenolpyruvate; pmf, protonmotive force; spanner, transmembrane cl-helical spanner; TC, Transport Commission; TMS, transmembrane u-helical spanner. Family uhhreviutions: AAAP, amino acid/auxin permease; ABC, ATP-binding cassette; ACC, ATP-gated cation channel; APC, amino acid-polyaminecholine; Ars, arsenical efflux; BR, ion-translocating bacteriorhodopsin; CIC, chloride channel; Colicin, channel-forming colicin; DASS, divalent anion:Na+ symporter; ENaC, epithelial Na+ channel; F-ATPase ( Fo. F, ATPase), H+- or Na+-translocating F-type, V-type and A-type ATPase; GntP, gluconate:H+ symporter; Holin, Holin functional superfamily; MC, mitochondria1 carrier; MFS, major facilitator superfamily; [Families of the MFS: 1) SP, sugar porter; 2) DHA14, drug:H+ antiporter, 14 spanners; 3) DHA 12, drug:H+ antiporter, 12 spanners; 4) OPA, organophosphate:P, antiporter; 5) OHS, oligosaccharide:H+ symporter; 6 ) MHS, metabolite: H symporter; 7) FHS, fucose:H+ symporter; 8) NNP, nitrate/nitrite porter; 9) PHS, phosphate:H+ symporter; 10) NHS, nucleoside:H+ symporter; I I ) OFA, +

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0xalate:formate antiporter; 12) SHS, sialate:H+ symporter; 13) MCP, monocarboxylate porter; 14) ACS, anion:cation symporter; 15) AAHS, aromatic acid:H+ symporter; 16) UMF, unknown major facilitator; 17) CP, cyanate permease]; MIP, major intrinsic protein; Mot, H+- or Na+-translocating bacterial flagellar motor; MscL, large conductance mechanosensitive ion channel; Na-NDH, Na+-translocating NADH:quinone dehydrogenase; NaT-MMM, Na+-transporting methy1tetrahydromethanopterin:coenzyme M methyltransferase; PTS, phosphotransferase system; RIR-CaC, ryanodine-inositol 1,4,5-triphosphate receptor Ca2+ channel; RND, resistance-nodulation-cell division; Sit, silicon transporter; SMR, small multidrug resistance; SSS, solute: sodium symporter; TPT triose-phosphate translocator; VIC, voltage-sensitive ion channel. “A theory is the more impressive the greater is the simplicity of its premises, the more different are the kinds of things it relates and the more extended is its range of applicability”. Albert Einstein

1. INTRODUCTION

Over the past decade, our laboratory and others have been concerned with molecular archeological studies aimed at revealing the origins and evolutionary histories of permeases. These studies have revealed that several different families, defined on the basis of sequence similarities, arose independently of each other, at different times in evolutionary history, following different routes. When complete microbial genomes first became available for analysis, we adapted pre-existing software and designed new programs that allowed us quickly to identify probable transmembrane proteins, estimate their topologies and determine the likelihood that they function in transport. This work allowed us to expand previously recognized families and to identify dozens of new families. All of this work then led us to attempt to design a rational but comprehensive classification system that would be applicable to the complete complement of transport systems found in all living organisms on earth. The classification system that we have devised is based primarily on mode of transport and energy coupling mechanism, secondarily on molecular phylogeny, and lastly on the substrate specificities of the individual permeases. In this chapter we shall initially present the system we have developed for permease classification and then discuss and update some of our earlier phylogenetic work. We shall present some of the evidence that establishes that different permease families arose independently of each other and sum-

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marize some of the more interesting conclusions that have resulted from complete microbial genome analyses. Finally, we shall conclude with a discussion of pertinent observations that have resulted from the development of our transporter classification system. It will be argued that molecular phylogeny provides a useful basis for classifying permeases. Most of the information to be presented represents just the tip of the iceberg of our efforts. The detailed results can be found in our recent publications (Pao et al., 1998; Paulsen et al., 1998) and on our World Wide Web sites [http://www-biology.ucsd.edu/-ipaulsen/transport/titlepage.html;http:// www-biology.ucsd .edu/-msaier/transport/titlepage.html]. 2. CONSIDERATIONS FOR THE SYSTEMATIC CLASSIFICATION OF TRANSMEMBRANE SOLUTE PERMEASES

Enzymes have long been classified in accordance with the directives and recommendations of the Enzyme Commission (EC) (Dixon and Webb, 1979). The system of classification utilized by the Commission was developed several decades ago, before protein and nucleic acid sequencing techniques had become refined. Consequently, nothing was then known about the protein families to which the constituent enzymes belong. The EC system of enzyme classification which proved to be of tremendous value was based exclusively on function. In developing the EC system, it was tacitly assumed that proteins of similar catalytic function would be closely related, and that they therefore should be grouped together. However, as increasing numbers of enzyme sequences became available, molecular phylogenists interested in protein evolution developed programs and conducted computational sequence comparisons that revealed the probable evolutionary origins of these proteins. This work showed that while enzymes catalyzing a specific reaction from different organisms were often related in sequence and used the same mechanism, two different enzymes catalyzing exactly the same reaction sometimes: (i) exhibited completely different amino acid sequences; (ii) were of different three-dimensional structures; (iii) were of different subunit compositions; (iv) took advantage of different cofactors to catalyze the reaction; (v) exhibited different reaction kinetics; (vi) went through different reaction intermediates; (vii) showed sensitivities to different amino acyl-specific chemical reagents; (viii) utilized different active site residues to bind and orient the substrate(s); (ix) utilized different active site catalytic residues to lower transition state energy levels; (x) exhibited different active site configurations; (xi) functioned by completely different mechanisms; and (xii) apparently evolved independently of each other, converging only with respect to the final reactions catalyzed.

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In spite of these sequence, structural and catalytic differences, and in spite of the probability that these enzymes are not related to each other in any way, they occupied adjacent positions in the EC classification system. Thus, this system reflected only the reactions catalyzed by and the substrate specificities of the enzymes, but it did not reflect the structures, mechanisms of action or phylogenies of these proteins. As has been extensively documented, molecular phylogeny reflects protein structure and mechanism of action, and it only secondarily reflects the specific reaction catalyzed and the substrate specificity. Since the former characteristics are fundamental traits of a protein while the latter characteristics are more superficial traits, sometimes merely reflective of single amino acyl residue changes in an enzyme, it would be reasonable to suggest that as more and more sequence and phylogenetic data become available, these data should be used to provide the most reliable basis for protein classification. Since single residue substitutions can give rise to different substrate binding properties, these characteristics should be used in the final level of classification rather than a primary level. We therefore suggest that the evolutionary process provides the most reliable indication of structure, mechanism and function. If molecular phylogenetic studies can accurately retrace the evolutionary process, they should be used as the basis for any rational system of protein classification. In defense of the monumental efforts and accomplishments of the Enzyme Commission, it should be pointed out that the EC classification system was based on the best criteria available at the time. In fact, only now that we have entered the era of complete genome sequencing, are sufficient phylogenetic data available to allow the design of novel comprehensive classification systems based on molecular phylogeny. The usefulness of the EC system cannot be overestimated, and the fact that the system these pioneers developed does not reflect the structural, mechanistic or evolutionary traits of these proteins, merely reflects the normal scientific process and does not imply a lack of insight. The best means available at the time were utilized. However, it is now time to move on. The Enzyme Commission recognized and classified only those proteins that catalyze chemical modification reactions, i.e. proteins that are ‘enzymes’ according to the classical definition. Some of the enzymes classified within the EC system are asymmetrically situated within an anisotropic, hydrophobic lipid membrane that separates two aqueous compartments. The resultant asymmetry allows these enzymes to catalyze vectorial as well as chemical modification reactions as clearly enunciated several decades ago by Peter Mitchell (Mitchell and Moyle, 1959; Mitchell, 1961, 1962). In fact, a significant fraction of these integral membrane proteins do catalyze transmembrane transport of small solutes. In spite of the fundamental biological significance of vectorial processes, the vectorial nature of the reac-

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tions catalyzed was not emphasized in the system developed by the Enzyme Commission. Moreover, most currently recognized solute permeases do not catalyze a chemical reaction, are therefore not enzymes, and consequently are not included within the EC classification system. N o coherent and comprehensive system of permease classification has yet been proposed or implemented to the best of my knowledge. The system I wish to present was designed to correct this deficiency. 3. PROPOSED TRANSPORT PROTEIN CLASSIFICATION SYSTEM

According to the proposed classification system, transport systems should be grouped on the basis of four criteria, and each of these criteria corresponds to one of the four numbers within the Transport Commission (TC) number for a particular type of permease. Thus, a TC number has four components as follows: W.X.Y.Z. W corresponds to the transporter type and energy source (if any) used to drive transport; X specifies the permease family (or superfamily); Y represents the subfamily (or family in a superfamily) in which a particular permease is found; and Z delineates the substrate(s) transported. Any two transport proteins in the same subfamily of a permease family that transport the same substrate(s) using the same mechanism are given the same TC number, regardless of whether they are orthologues (i.e. arose in distinct organisms by speciation) or paralogues (i.e. arose within a single organism by gene duplication). Sequenced homologues of unknown function are not normally assigned a TC number, and functionally characterized permeases for which sequence data are not available are also not included. These deficiencies will be eliminated with time as more and more sequenced permeases are characterized biochemically and as sequences become available for the functionally but not molecularly characterized permeases. The primary level of classification is based on permease type and energy source. Twelve primary categories of permease types are currently recognized as follows: 1. Channel-type facilitators. Proteins in this category have transmem-

brane channels which usually consist largely of a-helical spanners. Transport systems of this type catalyze facilitated diffusion (by an energy-independent process) by passage through a transmembrane aqueous pore or channel without evidence for a carrier-mediated mechanism. Outer membrane porin-type channel proteins are excluded from this category and have been put into their own category (category 9).

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\

87

2. Carrier-type facilitators. Transport systems are included in this category if they utilize a carrier-mediated process to catalyze uniport (a single species is transported by facilitated diffusion in a process not coupled to the utilization of a primary source of energy), antiport (two or more species are transported in opposite directions in a tightly coupled process not directly linked to a form of energy other than chemiosmotic energy) and/or symport (two or more species are transported together in the same direction in a tightly coupled process not directly linked to a form of energy other than chemiosmotic energy). 3. ATP-driven active transporters. Transport systems are included in this category if they hydrolyze the terminal pyrophosphate bond in ATP or of another nucleoside triphosphate to drive the active uptake and/or extrusion of a solute or solutes. The transport protein may or may not be transiently phosphorylated, but the substrate is not phosphorylated. 4. PEP-dependent, phosphoryl transfer-driven group translocators. Transport systems of the bacterial phosphoeno1pyruvate:sugar phosphotransferase system are included in this category. The product of the reaction, derived from extracellular sugar, is a cytoplasmic sugarphosphate. 5 . Carboxylic acid-dependent, decarboxylation-driven active transporters. Transport systems that drive solute (e.g. sodium ion) uptake or extrusion by decarboxylation of a cytoplasmic substrate are included in this category. 6. Electron flow-driven active transporters. Transport systems that drive transport of a solute (e.g. an ion) energized by the flow of electrons from a reduced substrate to an oxidized substrate are included in this category. 7. Light-driven active transporters. Transport systems that utilize light energy to drive transport of a solute (e.g. an ion) are included within this category. 8. Mechanically driven active transporters. Transport systems are included within this category if they drive movement of a cell, organelle or other physical structure by allowing the flow of ions (or other solutes) through the membrane down their electrochemical gradients. 9. Outer membrane channel-type facilitators ( p o r k ) . The proteins of this category exhibit transmembrane p strands that form p barrels through which solutes pass. They are found in the outer membranes of Gram-negative bacteria, mitochondria and eukaryotic plastids.

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MILTON H. SAIER, JR

10. Methyltransferase-driven active transporters. A single characterized

protein, the Na+-transporting methy1tetrahydromethanopterin:coenzyme M methyltransferase, currently falls into this category. 98. Auxiliary transport proteins. Proteins that function with, or are complexed to, known transport proteins are included in this category. An example would be a protein that facilitates transport across the two membranes of the Gram-negative bacterial cell envelope in a single step driven by the energy source (e.g. ATP or the protonmotive force, pmf) utilized by a cytoplasmic membrane transporter. Energy coupling and regulatory proteins that d o not actually participate in transport represent other examples. 99. Transporters of unknown classification. Transport protein families of unknown classification are grouped under this number and will be classified elsewhere when the transport process and energy coupling mechanism are characterized. The current index of transport system families is presented in Table 1. Over 150 families are represented. Some of these families are large superfamilies with hundreds of currently sequenced members. Others are small families with only one or two members. Most families, however, are of intermediate sizes, with between five and 100 members. All of these families will undoubtedly expand with time, and new families will be identified. The availability of new protein sequences will occasionally allow two or more currently recognized families to be placed together under a single TC number. In a few cases, two families are already known for which some evidence is available suggesting that these families are related. This evidence is usually based on: (i) limited sequence similarities; (ii) common function; and/or (iii) similar protein size, topology and structure. When ‘missing link’ sequences or three-dimensional structural data become available so that proteins of two families can be unequivocally grouped together within a single family, the lower TC number will be adopted for all of the family members, and the higher TC number will be abandoned. The complete index (Table 1) and representative tables describing some of the families to be discussed in this chapter will be presented below. The complete classification system is available on our WEB site [http://www-biology.ucsd.edu/-msaier/transport/ titlepage.html1. It will be updated continuously as new information becomes available. Anyone noting errors or incomplete listings is encouraged to contact the author providing the missing information and references by email, fax, phone or mail. In almost all cases, members of a transporter family utilize a single energy coupling mechanism, thus justifying the use of transport mode and energy coupling mechanism as the primary bases for classification. However, two exceptions have been noted. First, the arsenite (Ars) permease of Escherichiu

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Table 1 Families of transport proteins. 1.

Channel-type facilitators, a-type 1.1 The Major Intrinsic Protein (MIP) Family 1.2 The Epithelial Na+ Channel (ENaC) Family 1.3 The Large Conductance Mechanosensitive Ion Channel (MscL) Family 1.4. ATP-gated Cation Channel (ACC) Family 1.5 The Voltage-sensitive Ion Channel (VIC) Family 1.6 The Ligand-gated Ion Channel (LIC) Family of Neurotransmitter Receptors 1.7 The Glutamate-gated Ion Channel (GIC) Family of Neurotransmitter Receptors I .8 The Channel-forming Amphipathic Peptide (CAP) Functional Superfamily 1.9 The Ryanodine-Inositol 1,4,5-trisphosphate Receptor Ca2+ Channel (RIRCaC) Family 1.10 The Chloride Channel (CIC) Family I . 1 1 The Holin Functional Superfamily 1.12 The Channel-forming Colicin (Colicin) Family 1.13 The Channel-forming S-Endotoxin Insecticidal Crystal Protein (ICP) Family 1.14 The a-Hemolysin (uHL) Family 1.15 The Aerolysin Channel-forming Toxin (Aerolysin) Family

2. Carrier-type facilitators (uni-, sym- and antiporters) 2. I The Major Facilitator Superfamily (MFS) 2.1.1 Sugar Porter (SP) Family 2.1.2 The Drug:H+ Antiporter (14 Spanner) (DHA14) Drug Efflux Family 2.1.3 The Drug:H+ Antiporter (12 Spanner) (DHA12) Drug Efflux Family 2.1.4 The Organophosphate: Pi Antiporter (OPA) Family 2.1.5 The 0ligosaccharide:H' symporter (OHS) Family 2.1.6 The Metabolite:Ht Symporter (MHS) Family 2.1.7 The Fucose:H+ Symporter (FHS) Family 2.1.8 The Nitrate/Nitrite Porter (NNP) Family 2.1.9 The Phosphate:H+ Symporter (PHS) Family 2.1.10 The Nuc1eoside:H' Symporter (NHS) Family 2. I . I 1 The 0xalate:Formate Antiporter (OFA) Family 2.1.12 The Sialate:H+ Symporter (SHS) Family 2. I , 13 The Monocarboxylate Porter (MCP) Family 2.1.14 The Anion:Cation Symporter (ACS) Family 2.1.15 The Unknown Major Facilitator (UMF) Family 2.1. I6 The Aromatic Acid:H+ Symporter (AAHS) Family 2.1.17 The Cyanate Permease (CP) Family 2.2 The Glycoside-Pentose-Hexuronide(GPH):Cation Symporter Family 2.3 The Amino Acid-Polyamine-Choline (APC) Family 2.4 The Cation Diffusion Facilitator (CDF) Family 2.5 The Zinc (Zn*+)-Iron (Fe2+)Permease (ZIP) Family 2.6 The Resistance-Nodulation-CellDivision (RND) Family 2.7 The Small Multidrug Resistance (SMR) Family 2.8 The Gluconate:H+ Symporter (GntP) Family 2.9 The L-Rhamnose Transporter (RhaT) Family 2.10 The 2-Keto-3-Deoxygluconate Transporter (KDGT) Family

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MILTON H. SAIER, JR

Table 1 continued

2. I I The Citrate-Mg*+:H+ (CitM) - Citrate:H+ (CitH) Symporter (CitMHS) Family 2.12 The ATP:ADP Antiporter (AAA) Family 2.13 The C4-Dicarboxylate Uptake (Dcu) Family 2.14 The Lactate Permease (LctP) Family 2.15 The Betaine/Carnitine/CholineTransporter (BCCT) Family 2,16 The Telurite-resistance/DicarboxylateTransporter (TDT) Family 2.17 The Proton-dependent Oligopeptide Transporter (POT) Family 2.18 The Amino Acid/Auxin Permease (AAAP) Family 2. I9 The Ca2+:CationAntiporter (CaCA) Family 2.20 The Inorganic Phosphate Transporter (Pit) Family 2.21 The So1ute:Sodium Symporter (SSS) Family 2.22 The Neurotransmitter:Sodium Symporter (NSS) Family 2.23 The Dicarboxy1ate:Cation (Na'or H+) Symporter (DCS) Family 2.24 The Citrate:Na+ Symporter (CSS) Family 2.25 The Alanine:Na+ Symporter (ASS) Family 2.26 The Branched Chain Amino Acid:Na+ Symporter (LIVSS) Family 2.27 The Glutamate:Na+ Symporter (ESS) Family 2.28 The Bile Acid:Na+ Symporter (BASS) Family 2.29 The Mitochondrial Carrier (MC) Family 2.30 The Na-K-CI Cotransporter (NKCC) Family 2.31 The Anion Exchanger (AE) Family 2.32 The Silicon Transporter (Sit) Family 2.33 The NhaA Na+:H+ Antiporter (NhaA) Family 2.34 The NhaB Na+:H+ Antiporter (NhaB) Family 2.35 The NhaC Na+:H+ Antiporter (NhaC) Family 2.36 The Monovalent Cation:Proton Antiporter-l (CPA-I) Family 2.37 The Monovalent Cation:Proton Antiporter-2 (CPA-2) Family 2.38 The K + Transporter (Trk) Family 2.39 The Nuc1eobase:Cation Symporter-I (NCSI) Family 2.40 The Nuc1eobase:Cation Symporter-2 (NCS2) Family 2.41 The Nucleoside Uptake Permease (NUP) Family 2.42 The Aromatic Amino Acid Permease (ArAAP) Family 2.43 The Serine/Threonine Porter (STP) Family 2.44 The Formate-Nitrite Porter (FNP) Family 2.45 The Metal Ion Transporter (MIT) Family 2.46 The Benzoate:H+ Symporter (BenE) Family 2.47 The Sulfate/Carboxylate Uptake Permease (SCUP) Family 2.48 The Reduced Folate Carrier (RFC) Family 2.49 The Ammonium Transporter (Amt) Family 2.50 The Triose Phosphate Translocator (TPT) Family 2.5 I The Nucleotide-Sugar Transporter (NST) Family 2.52 The Ni "-Co2' Transporter (NiCoT) Family 2.53 The Sulfate Permease (SulP) Family 2.54 The Mitochondrial Tricarboxylate Carrier (MTC) Family 2.55 The Acetyl-Coenzyme A Transporter (AcCoAT) Family 2.56 The Tripartite ATP-independent Periplasmic Transporter (TRAP-T) Family 2.57 The Equilibrative Nucleoside Transporter (ENT) Family

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TRANSPORT PROTEIN CLASSIFICATION

Table I

continued

3. ATP-driven active transporters 3.1 The ATP-binding Cassette (ABC) Superfamily 3.2 The H+- or Na+-translocating F-type, V-type and A-type ATPase (FATPase) Superfamily 3.3 The Cation-translocating P-type ATPase (P-ATPase) Superfamily 3.4 The Arsenical (Ars) Efflux Family 3.5 The Type I1 (General) Secretory Pathway (IISP) Family 3.6 The Type 111 (Virulence-related) Secretory Pathway (IIISP) Family 3.7 The Type IV (Conjugal DNA-Protein Transfer or VirB) Secretory Pathway (IVSP) Family 4. Phosphotransferase systems 4.1 The PTS Glucose-Glucoside (Glc) Family 4.2 The PTS Fructose-Mannitol (Fru) Family 4.3 The PTS Lactose-Cellobiose (Lac) Family 4.4 The PTS Glucitol (Gut) Family 4.5 The PTS Galactitol (Gat) Family 4.6 The PTS Mannose-Fructose-Sorbose (Man) Family 5. Decarboxylation-driven active transporters 5.1 The Na'-transporting Carboxylic Acid Decarboxylase (NaT-DC) Family 6. Electron flow-driven active transporters 6.1 The Proton-translocating NADH Dehydrogenase (NDH) Family 6.2 The Proton-translocating Transhydrogenase (PTH) Family 6.3 The Proton-translocating Quino1:Cytochrome c Reductase (QCR) Superfamily 6.4 The Proton-translocating Cytochrome Oxidase (COX) Superfamily 6.5 The Na+-translocating NADH:Quinone Dehydrogenase (Na-NDH) Family 7. Light-driven active transporters 7.1 The Ion-translocating Bacteriorhodopsin (BR) Family 7.2 The Proton-translocating Reaction Center (RC) Family 8. Mechanically driven active transporters 8.1 The H+- or Na+-translocating Bacterial Flagellar Motor (Mot) Family 9. Outer 9.1 9.2 9.3 9.4 9.5 9.6 9.7 9.8 9.9 9.10

membrane porins, p-type The General Bacterial Porin (GBP) Family The Chlamydia1 Porin (CP) Family The Sugar Porin (SP) Family The Erucella-Rhizobium Porin (BRP) Family The Pseudomonas OprP Porin (POP) Family The OmpA-OprF Porin (OOP) Family The Rhodobacter PorCa Porin (RPP) Family The Mitochondria1 and Plastid Porin (MPP) Family The FadL Outer Membrane Protein (FadL) Family The Nucleoside-specific Channel-forming Outer Membrane Porin (Tsx) Family 9. I I The Outer Membrane Fimbrial Usher Porin (FUP) Family 9.12 The Autotransporter (AT) Family

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MILTON H. SAIER, JR

Table 1 continued 9.13 The Alginate Export Porin (AEP) Family 9.14 The Outer Membrane Receptor (OMR) Family 10. Methyltransferase-driven active transporters 10.1 The Na+-transporting Methy1tetrahydromethanopterin:CoenzymeM

Methyltransferase (NaT-MMM) Family 98. Auxiliary transport proteins 98.1 The Membrane Fusion Protein (MFP) Family 98.2 The Outer Membrane Factor (OMF) Family 98.3 The Cytoplasmic Membrane-Periplasmic Auxiliary-1 (MPAI) Protein with Cytoplasmic (C) Domain (MPAI-C or MPAI + C) Family 98.4 The Cytoplasmic Membrane-Periplasmic Auxiliary-2 (MPA2) Family 98.5 The Outer Membrane Auxiliary (OMA) Protein Family 98.6 The TonB-ExbB-ExbD / TolA-TolQ-TolR (TonB) Family of Auxiliary Proteins for Energization of Outer Membrane Receptor (0MR)-mediated Active Transport 98.7 The Phosphotransferase System Enzyme I (EI) Family 98.8 The Phosphotransferase System HPr (HPr) Family 98.9 The rBAT Family of Putative Transport Accessory Proteins 98.10 The Slow Voltage-gated K+ Channel Accessory Protein (MinK) Family 99. Transporters of unknown classification 99.1 The Polysaccharide Transporter (PST) Family 99.2 The MerTP Mercuric ion (H 2 + ) Permease (MerTP) Family 99.3 The MerC Mercuric Ion (I-$+) Uptake (MerC) Family 99.4 The Nicotinamide Mononucleotide (NMN) Uptake Permease (PnuC) Family 99.5 The K+ Uptake Permease (Kup) Family 99.6 The L-Lysine Exporter (LysE) Family 99.7 The Chromate-resistance (ChrA) Family 99.8 The Ferrous Iron Uptake (FeoB) Family 99.9 The Low Affinity Fe2+ Transporter (FeT) Family 99.10 The Oxidase-dependent Fez+ Transporter (OFeT) Family 99.11 The Copper Transporter-1 (Ctr-I) Family 99.12 The Copper Transporter-2 (Ctr-2) Family 99.13 The MnZt Uptake Transporter (MnT) Family (also called the Nramp Family)

coli consists of two proteins, ArsA and ArsB (Table 2). ArsB is an integral membrane protein which presumably provides the transport pathway for the extrusion of arsenite and antimonite (Silver et al., 1993). ArsA is an ATPase that energizes ArsB-mediated transport. However, when ArsB alone is present, as in the case of the arsenical resistance pump of Sraphylococcus aureus (Table 2), transport is driven by the pmf (Broer et al., 1993). The presence or absence of the ArsA protein thus determines the mode of energy coupling. Such promiscuous use of energy is exceptionally rare and has to my knowledge been documented in only a very few instances. When such an

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Table 2 The arsenical (Ars) efflux family (TC #3.4)

The arsenical resistance (Ars) efflux pumps of bacteria consist either of two proteins (ArsB, the integral membrane constituent with 12 transmembrane spanners, and ArsA, the ATP-hydrolyzing, transport energizing subunit, as for the chromosomally-encoded E. coli system), or of one protein (the ArsB integral membrane protein of the plasrnidencoded Staphylococcus system (Silver et al., 1993)). The ArsB membrane constituents of these systems possess homologues in animals, while the ArsA constituents have homologues in both plants and animals. ArsA proteins have two ATP binding domains and probably arose by a tandem gene duplication event. ArsB proteins all possess 12 transmembrane spanners and may also have arisen by a tandem gene duplication event. Structurally, the Ars pumps resemble ABC-type emux pumps, but there is insufficient sequence similarity between the Ars and ABC pumps to establish homology. When only ArsB is present, the system operates by a pmf-dependent mechanism. When ArsA is also present, ATP hydrolysis drives efflux (Bruhn et a/., 1996). These pumps actively expel both arsenite and antimonite. The overall reaction catalyzed by ArsB is: Arsenite or Antimonite (in)

2 Arsenite or Antimonite (out)

That by ArsB-ArsA is: Arsenite or Antimonite (in)

+ ATP + Arsenite or antimonite (out) +ADP P,

+

TC #

Name

Source

Example

Arsenical resistance efflux pump (ATP-dependen t) Arsenical resistance efflux pump (pmf-dependent)

Bacteria; eukaryotes

ArsA-ArsB of E. coli (ArsA:spP08690; ArsB: spP373 10) ArsB of Staphylococcus aureus (spP30329)

~~

3.4.1.1 3.4.1.2

Bacteria; eukaryotes

effect is reported, we shall classify the permease in accordance with the more complicated energy coupling mechanism [in this case, as an ATP-driven primary active transporter (Class 3), rather than as a secondary carrier (Class 2)]. The potentially different energy coupling mechanisms will be described in the table characterizing that family (see, for example, Table 2). Examples of permease families in which promiscuous transport modes have been reported include the mitochondria1 carrier (MC) family (Table 3; TC #2.29) and the triose phosphate translocator (TPT) family (Table 4; TC #2.50). Proteins of both these families are apparently restricted to eukaryotic organelles. Members of both families normally catalyze carrier-mediated substrate:substrate antiport, and are therefore classified as secondary carriers. However, treatment of MC family members with chemical reagents, such as N-ethylmaleimide or Ca2+ (Dierks et al., 1990a,b; Brutovetsky and Klingenberg, 1994, 1996; Jezek et al., 1994), or imposition of a large mem-

94

MILTON H. SAIER, JR

Table 3 The Mitochondrial Carrier (MC) family (TC #2.29).

Permease protein subunits of the M C family possess six transmembrane a-helical spanners (Kuan and Saier, 1993). The proteins are of fairly uniform size (about 300 residues). They arose by tandem intragenic triplication events in which a genetic element encoding two spanners gave rise to one encoding six spanners. This event may have occurred less than 1.5 billion years ago when mitochondria first developed their specialized endosymbiotic functions within eukaryotic cells. Members of the family are found exclusively in eukaryotic organelles although they are nuclearly encoded. Most are found in mitochondria, but some are found in peroxisomes of animals and in amyloplasts of plants. Structurally characterized members of the M C family are dimers. Many of them preferentially catalyze the exchange of one solute for another (antiport) (Sullivan et al., 1991; Fiermonte et al., 1992; Fernandez et al., 1994; Ahringer, 1995; Liu and Dunlap, 1996; Tzagoloff et al.. 1996; lndiveri et al., 1997). Fifteen paralogues of the M C family are encoded within the genome of Saccharornyces cerevisiae. The generalized transport reaction for carriers of the M C family is: S, (out) S2(in) + S , (in) S2(out).

+

+

TC #

Name

Source(s)

Example

2.29. I . 1

ATP/ADP antiporter

Animals, plants, fungi

2.29.2. I

Oxoglutarate/malate antiporter

Animals

2.29.3.1

Uncoupling protein

Animals

2.29.4. I

Phosphate carrier

Animals, yeast

2.29.5.1

M R S protein

Yeast

2.29.6.I

Peroxisomal carrier

Yeast

2.29.7.1

Tricarboxylate carrier

Animals

2.29.8.1

Mitochondrial carnitine/ acyl carnitine carrier (CAC) Embryonic differentiation (DIF-1) protein

Mammals

ATP/ADP carrier of Homo sapiens (gbJ02683) Oxoglutarate/malate carrier of Bos ruurus (gbM60662) Uncoupling carrier o f Bos raurus (gbX 14064) Phosphate carrier of Bos taurus (gbX05340) M R S protein of Saccharomyces cerevisiae (spP10566) (transport substrate unknown) Peroxysomal carrier of Candida hoidnii (pirS50283) (transport substrate unknown) Citrate carrier of Ratrus norvegicus (gbL12016) C A C of Rattus norvegicus (gbX9783 1)

2.29.8. 2

Animals

DIF- 1 of Caenorhabdiris elegans (pirS55056) (transport substrate unknown)

95

TRANSPORT PROTEIN CLASSIFICATION

Table 3 continued TC #

Name

Source(s)

Example

2.29.9. I

Mitochondria] basic amino acid carrier (BAAC) Flavin adenine dinucleotide (FAD) carrier (FADC; FLXI) Amyloplast Brittle-1 (BTI) protein

Fungi

BAAC of Neurospora crussu (gbL36378) FLX 1 of Saccharomyces cerevisiae (spP40464)

2.29.10.1

2.29.1 I . I 2.29.12. I 2.29.13.1

Grave’s Disease Carrier (GDC) protein Regulator of acetylCoA-synthetase activity, ACR-1

Yeast

Plants Mammals Yeast

BTI of Zea Mays (spP29518) (transport substrate unknown) GDC of Bos iaurus (spQ01888) (transport substrate unknown) ACR 1 of Saccharomyces cerevisiae (spP33303) (transport substrate unknown)

brane potential (AW)across a membrane into which a TPT member has been incorporated (Wallmeier et ul., 1992; Schultz rt ul., 1993; Schwarz el al., 1994), has been reported to convert these antiport-catalyzing carriers into anion-selective channels capable of functioning by uniport. ‘Tunneling’ of ions and other solutes through carriers with little or no conformational change has been discussed (Frohlich, 1988). Again, the more complicated carrier-type mechanism, which appears to be relevant under most physiological conditions, provides the basis for classifying these proteins (i.e. as Class 2 carriers rather than Class 1 channels).

4. DIVERSE EVOLUTIONARY ORIGINS OF INTEGRAL

MEMBRANE TRANSPORT PROTEIN FAMILIES Early work from several laboratories, including ours, revealed that the integral membrane proteins that comprise or partially comprise many permeases share a common apparent topology (for reviews see Saier, 1994, 1996). This common feature is presented in Table 5. In this table each entry represents a different currently recognized family of transport systems, where the members of a particular family are defined on the basis of their sequence similarities.

96

MILTON H. SAIER, JR

Table 4 The Triose-phosphate Translocator (TPT) family (TC #2.50).

The functionally characterized members of the TPT family are derived from the inner envelope membranes of chloroplasts and non-green plastids of plants (Fliigge and Heldt, 1991; Fliigge, 1992; 1995; Fliigge et al., 1992; Fischer et al., 1997). However, homologues are also present in yeast (Loddenkotter et a/., 1993). Sacchnromyces cerevisiae has three functionally uncharacterized TPT paralogues encoded within its genome. Under normal physiological conditions, chloroplast TPTs mediate a strict antiport of substrates, frequently exchanging an organic three-carbon compound phosphate ester for inorganic phosphate (PI). Normally, a triose-phosphate, 3-phosphoglycerate, or another phosphorylated C3 compound made in the chloroplast during photosynthesis, exits the organelle into the cytoplasm of the plant cell in exchange for PI. However, experiments with reconstituted translocator in artificial membranes indicate that transport can also occur by a channel-like uniport mechanism with about 10-fold higher transport rates. Channel openings may be induced by a membrane potential of large magnitude and/or by high substrate concentrations. Non-green plastid TPT carriers, such as those from maize endosperm and root membranes, mediate transport of C3 compounds phosphorylated at carbon atom 2, particularly phosphoenolpyruvate, in exchange for PI. Glucose 6-P has also been shown to be a substrate of the plastid translocator. The chloroplast and nongreen plastid proteins are divergent in sequence and substrate specificity. Each TPT family protein consists of about 4 0 M 5 0 amino acyl residues with five to eight putative transmembrane a-helical spanners (TMSs). The actual number has been proposed to be six for the plant proteins as for mitochondria1 carriers (TC #2.29). However, proteins of the TPT family do not exhibit significant sequence similarity with the latter proteins, and there is no evidence for an internal repeat sequence. The generalized reaction catalyzed by the proteins of the TPT family is: organic phosphate ester (in) + P I (out) organic phosphate ester (out) P, (in).

+

TC #

Name

Source

Example

2.50.1. 1

Chloroplast TPT

Plants

2.50.2. I

Non-green plastid TPT

Plants

2.50.3. 1

Sly4lp (transport function unknown)

Yeast

TPT of Zea mays (spP49 133) TPT of Brassica oleracea (spP52 178) Sly4lp of Saccharonryces cere visiae (spP222 15)

All of the members of a family thus exhibit a statistically significant degree of sequence similarity, but they lack significant sequence similarity with the members of all other families. The members of a single family are therefore said to be homologous as they are all derived from a single, primordial, ancestral protein, but they cannot be shown to be homologous to the members of any other well-defined protein family. The criteria for establishing homology have been enunciated and evaluated elsewhere (Doolittle, 1986; Saier, 1994). It should be noted, however, that lack of significant sequence similarity between any two proteins, or between the protein mem-

97

TRANSPORT PROTEIN CLASSIFICATION

Table 5 Topological features of representative well-characterized transporter families.

Family

Example(s)

Mitochondria1 Carrier ATP/ADP antiporters (MC) Family (TC #2.29) of mitochondria Major Intrinsic Protein Aquaporins of plants, (MIP) Family (TC #I.]) animals, yeast and bacteria Major Facilitator (MF) Lactose permease of Superfamily (TC #2.1) E. coli Resistance-NodulationMultidrug resistance Cell Division (RND) pumps of Gramnegative bacteria Family (TC #2.6) ATP-binding Cassette Maltose permease of E. coli (ABC) Superfamily (TC #3.1) Voltage-sensitive Ion K+ channels of animals Channel (VIC) Superfamily (TC #IS) Voltage-sensitive ion Na+ and Ca2+ channels Channel (VIC) of animals Superfamily (TC #IS)

No. of TMSs/ unit

No. of Units

No. of polypeptide chains

6

2

2

6

2or4

2or4

6

2

1

6

2

1

6

2

1 or 2

6

4

4

6

4

1

bers of two distinct families, does not by itself establish that these families did not stem from a common ancestor. It is always possible, unless demonstrated otherwise using definitive criteria, that the two families arose from a single ancestor, but that the members of these two groups of proteins diverged in sequence from each other to such a degree that homology between the two groups could not be established on the basis of sequence comparisons alone, or even that statistically significant sequence similarity could not be detected. The families presented in Table 5 are each presumed to have evolved independently of each other, and in the cases of the mitochondrial carrier (MC) family (Table 3), the major intrinsic protein (MIP) family (Table 6 ) , the major facilitator (MF) superfamily (Table 7) and the resistance-nodulation-cell division (RND) family (Table 8), this presumption can be established. Thus, for example, as illustrated schematically in Fig. 1, the mitochondrial carrier (MC) family (TC #2.29; Table 3) consists of over 100 currently sequenced proteins, all of which are found in mitochondria and a few other eukaryotic organelles (peroxisomes of animals and amyloplasts of plants). They are homodimeric permeases with a subunit size

98

MILTON H. SAIER, JR

Table h The Major Intrinsic Protein (MIP) family (TC #1.1). The MIP family is large and diverse, possessing members that form transmembrane channels and function in water, small molecule (e.g. glycerol; urea) and possibly ion transport by an energy independent mechanism (Reizer et al., 1993; Park and Saier, 1996). They are found ubiquitously in bacteria and eukaryotes. Phylogenetic clustering of the proteins is largely according to phylum of the organisms of origin. They are believed to form aqueous pores that selectively allow passive transport of their solute(s) across the membrane without apparent recognition. Aquaporins selectively transport water (but not glycerol) while glycerol facilitators selectively transport glycerol and other small organic molecules, but not water. Glycerol facilitators function as solute nonspecific channels, and physiologically may transport glycerol, dihydroxyacetone, urea and/or propanediol. One member of the family, the yeast FPS protein (1.1.5.1 below) may transport both. Several reports of MIP family proteins transporting ions may or may not be physiologically significant. However, demonstration of the involvement of the cyanobacterial channel protein ( I . I .4.1 below) in copper homeostasis suggests that it may transport Cu”. The physiological functions of many of these proteins are unknown. They probably consist of homodimers (GlpF of E. coli; I . I . I . 1) or tetramers (MIP of Bos raurus; 1.1.8.1). Each subunit spans the membrane six times as putative ahelices and arose from a three-spanner-encoding genetic element by a tandem. intragenic duplication event. The transport reaction for channel proteins of the MIP family is: H 2 0 ( o u t ) H 2 0 ( i n ) (e.g. aquaporins) or solute (out) * solute (in) ( e g glycerol facilitators).

*

TC #

Name

Source(s)

Example

I.1.1. 1

Glycerol facilitator

Gram (-) bacteria

1.1.2. I

Glycerol facilitator

1.1.3. I

Aquaporin Z

Gram ( + ) bacteria and Haemophilus injluenzae Enteric bacteria

GlpF of E. coli. (spP22144) GlpF of Bacillus suhtilis (spP18 156)

1.1.4. I

Channel protein

Cyanobacteria

1.1.5. 1

FPS glycerol facilitator

Yeast

1.1.6. I

P9h77.5 gene product

Yeast

1.1.7. 1

70.5 KDa protein

Yeast

AqpZ of E. coli (gbU38664) Copper homeostasis protein (SmpX) of Synechococcus sp. (gbD43774) FPSl protein of Succharomyces cere visiae (spP23900) P9677.5 gene product of Saccharomyes cerevisiae (spP53386) 70.5 KDa protein o f Sacchurotnvces cerevisiae (spP43549)

99

TRANSPORT PROTEIN CLASSIFICATION

Table 6 continued

TC #

Name

Source(s)

Example

1.1.8. 1

Aquaporin 1

Animals

1.1.9. 1

Aquaporin 3

An i rnaI s

1.1.10. I

Tonoplast intrinsic protein Tonoplast intrinsic protein (wTIP) Nodulin-26

Plants

Aquaporin 1 of Homo sapiens (spP29972); Major intrinsic protein (MIP) of Bos taurus (spPO6624); Big Brain (BIB) of Drosophila melanogaster (spP23645) Aquaporin 3 of Rurtus norvegicus (gbL35 108) TIP of Arabidopsis thaliuna (spP26587) oTIP of Pisum sativum (spP25794) Nodulin-26 of Glycine Max (spPO8995)

1. I . I 1 . l

1.1.12 . I

Plants Plants

minimally of about 300 amino acyl residues. Each subunit possesses a topology with six transmembrane a-helical spanners. These permeases transport a variety of solutes (nucleotides, organic acids, protons or hydroxyl ions, phosphate, carnitine and its acylated derivative, amino acids, enzyme cofactors such as flavin adenine dinucleotide (FAD) and other compounds), most frequently by antiport mechanisms (Table 3). The accession numbers provided in Table 3 allow easy access to the sequences of MC family proteins described, and these can be used to identify and retrieve the sequences of all sequenced members of the family (present and future) using, for example, the BLAST program to screen the current databases (Altschul et al., 1990). Based in part on the degree of sequence similarity observed for the three repeat units in each MC protein (Fig. I ) , we estimate that this triplication event occurred less than 1.5 billion years ago, after the invasion of the developing eukaryotic cell by aerobic bacteria to give rise to the prokaryotic precursor of mitochondria. Indeed, members of the MC family are found only in mitochondria and other eukaryotic organelles as noted above (Table 3). Members of the MIP family of channel proteins (Table 6) arose by an intragenic duplication event in which a genetic element encoding three transmembrane a-helical spanning regions (TMS) gave rise to a sixTMS-encoding element. This event was easily recognized by conducting sequence analyses and is believed to have occurred over 2 billion years ago

100

MILTON H. SAIER, JR

Table 7 The Major Facilitator Superfamily (MFS) (TC #2.1).

The MFS is a very old, large and diverse superfamily that includes several hundred sequenced members (Griffith et al., 1992; Marger and Saier, 1993; Pao er ul., 1998). They catalyze uniport, so1ute:cation (H+ or Na') symport and/or so1ute:solute antiport (Baldwin, 1993; Schuldiner et al., 1995; Goffeau et al., 1997). Most are 40MOO amino acyl residues in length and possess either 12 or 14 putative transmembrane a-helical spanners (Paulsen et a/., I996a.b). They exhibit specificity for sugars, polyols, drugs, neurotransmitters, Krebs cycle metabolites, phosphorylated glycolytic intermediates, amino acids, peptides, osmolytes, nucleosides, organic anions, inorganic anions, etc. They are found ubiquitously in all three domains of living organisms. The generalized transport reactions catalyzed by MFS permeases are: (I)

Uniport :

(2) Symport : (3) Antiport :

S (out)

* S (in) + +

+

S (out) [H' or Naf](out) + S(in) [H' or Na'](in) SI(out) S2(in) SI (in) S2 (out) (S, may be H+ or Na')

TC#

Name

2.1.1

Sugar Porter (SP) family

+

Source

Example

GalP of E. coli (spP37021) AraE of E. eoli (spP09830) XylE of E. coli (spP09098) Glf of Zyinomonas mobilis (spP2 1906) HxtO of Saccharomyces cerevisiue (spP4358 1) Gal2 of Saccharomyces cerevisiue (spP13181) Qay of Neurospora crassa (spP11636) ITR 1 of Saccharomyces cerevisiae (spP30605) LacP of Kluyveroniyces 1ucti.s (spPO7921) MAL6 of Sacchuromyces cerevisiae (spPI 5685) uGlc permease of Saccharomyces cerrvisiui~ (pirS59368) Gtr3 of Ratrus norvegicirs (spQ07647) Ftr of Homo sapiens (gbUI 1843) Hup 1 of Chlorella kesslui (spPI 5686) SYV2 of R a m s norvrgicus (spQO2563) Oca of Ruttus norvegicrrs (gbX78855)

2.1.1.1 2.1.1.2 2.1 .l.3 2. I . 1.4

Ga1actose:H' symporter Arabinose:H' symporter Xylose:H+ symporter Glucose uniporter

Bacteria Bacteria Bacteria Bacteria

2. I. 1.5

Hexose uniporter

Yeast

2.1.1.6

Galactose:H+ symporter

Yeast

2.1.1.7

Quinate:H' symporter

Fungi

2.1.1.8

Myoinosito1:H' symporter

Yeast

2. I . 1.9

Lactose:H symporter

Yeast

2.1.1.10 Maltose:H+ symporter

Yeast

2.1.1. I I a-glucoside:H' symporter

Yeast

2.1. I .I2 Glucose uniporter

Animals

2.1.1.13 Fructose uniporter

Animals

2.1. I .I4 Hexose:H' symporter

Plants

2.1.1 . I 5 Synaptic vesicle neurotransmitter transporter 2. I . I . I6 Organic cation transporter

Animals Animals

101

TRANSPORT PROTEIN CLASSIFICATION

Table 7 continued

TC#

Name

2.1.1.17 Glucose transporter 2.1.2 2.1.2.1 2.1.2.2 2. I .2.3 2. I .2.4 2.1.2.5 2. I .2.6 2.1.2.7 2. I .2.8

2.1.2.9 2.1.2.10 2.1.2.1 I 2.1.2.12 2.1.2. I3 .2.14 .2.15 .2.16 .2.17

Source

Examp1e

Protists

Th2A of Trypanosoma brucei (spQO6222)

The Drug:H+ antiporter (14 Spanner) (DHA14) drug efflux family Actinorhordin:H+ Gram-positive ActVa of Streptomyces coelicolor (gbX5 8833) bacteria antiporter Cephamycin:H+antiporter Gram-posi tive CmcT of Nocardia lactamdurans (spQ04133) bacteria Lincomycin:H+ antiporter Gram-posi tive LmrA of Streptomyces lincolnensis (gbX59926) bacteria Methylenomycin:H+ Gram-posi tive MmrB of Bacillus subtilis (spQOO538) bacteria antiporter Puromycin:H+ antiporter Gram-positive Pur8 of Streptomyces lipmanii (gbX76855) bacteria Tetrdcenomycin:H+ Gram-positive TemA of Streptomyces antiporter glaucescens (gbM80674) bacteria (Bicyclomycin, Gram-negative Bcr of E. coli (pirJN0659) sulfathiazole, etc.):H+ bacteria antiporter Gram-positive Blt of Bacillus subtilis (Fluoroquinolones, (gbL32599) acriflavin, bacteria chloramphenicol, ethidium bromide, etc.):Hi antiporter (Hydrophobic uncouplers Gram-nega tive EmrD of E. coli (spP31442) e.g., CCCP):H+ bacteria antiporter Gram-positive LmrP of Lactococcus lactis (Daunomycin, ethidium (gbX89779) bromide, etc.):H+ bacteria antiporter (Benomyl, cycloheximide, Yeast CaMDR1 of Candida albicans (spP28873) methotrexate, etc.):H+ antiporter VMAT1 of Rattus norvegicus (Doxorubicin, ethidium Mammals (gbM97380) bromide, rhodamine6-G):H' antiporter Car 1 of Schizosaccharomyces Amiloride:H+ antiporter Yeast cerevisiae (spP33532) CyhR of Candida maltosa Cyc1oheximide:H' Yeast (spP32071) antiporter CmlA of Pseudomonas Chloramphenicol:H+ Bacteria aeruginosa (spP32482) antiporter Tetracyc1ine:H'antiporter TetA of E. coli (gbX00006) Bacteria Acetylcholine:H+ Uncl7 of Caenorhabditis Animals elegans (spP34711) antiporter

102

MILTON H. SAIER, JR

Tuhle 7 continued

TC#

Name

2. I .3 2. I .3. I

The Drug:H+ antiporter (12 Spanner) (DHAl2) drug efflux family (Aminotriazole, 4Yeast Atrl of Sacchuromyces nitroquinoline-N-oxide, cerevisiae (gbZ49210) etc.):H+ antiporter (CCCP, nalidixic acid, Gram-negative EmrB of E. coli (spP27304) organomercurials, bacteria etc.):H+ antiporter (Acriflavin, ethidium Gram-positive LfrA of Mycohacteriuni smegtnatis (gbU40487) bromide, fluoroquinobacteria lone, etc.):H+ antiporter (Mono- and divalent Gram-positive QacA of Staphylococcus organo-cation):H' bacteria uureus (gbX56628) antiporter Gram-positive Ptr of Streptomyces (Pristinamycin 1 and 11, bacteria prist inuespiralis rifamycin):H+ antiporter (gbX84072) Tetracyc1ine:H' antiporter Bacteria TetK of Staphylococcus aurcws (gbM 162 17)

2.1.3.2 2.1.3.3 2. I .3.4 2.1.3.5 2.1.3.6

Source

Example

The 0rganophosphate:Pi antiporter (OPA) family Hexose-P:P, antiporter Bacteria UhpT of E. coli (spPI3408) P-glycerate:P, antiporter Bacteria PgtP of Sahnonellu typhitnurium (spP1268 I ) Bacteria GlpT of E. coli (spP08 194) 2. I .4.3 Glycerol-P:P, antiporter

2.1.4 2.1.4. I 2. I .4.2

2.1.5 2.1 S.1 2.1.5.2 2.1.5.3

The 0ligosaccharide:H' symporter (OHS) family Lactose:H' symporter Bacteria Lacy of E. coli (spP02920) Raffinose:H+ symporter Bacteria RafB of E. coli (spP16552) Sucrose:H+ symporter Bacteria CscB of E. coli (spP30000)

The Metabolite:H+ symporter (MHS) family 2.1.6 Bacteria 2. I .6.I Citrdte: H' symporter Bacteria 2. I .6.2 a-ketog1utarate:H' symporter Bacteria 2. I .6.3 Dicarboxy1ate:H' symporter 2.1.6.4 (Proline/betaine):(H+/Na+) Bacteria symporter Bacteria 2.1.6.5 4-Methyl-o-phthalate:H+ symporter 2.1.7 2.1.7.1 2. I .7.2

2.1.7.3

The Fucose:H+ symporter (FHS) family L-fucose:H+ symporter Bacteria Glucose/galactose Bacteria permease Glucose/mannose:H+ Bacteria symporter

Cit of E. coli (spP07860) KgtP of E. coli (spP17448) PcaT of Pseudomonas putida (gbU48776) Prop of E. coli (spP30848) MopB of Burkholderia cepacia (gbU 29532) FucP of E. coli (spP11551) Ggp of Brucella cibortus (gbU43785) GlcP of Bacillus suhtilis (gbAFOO2191 )

103

TRANSPORT PROTEIN CLASSIFICATION

Table 7 continued

TC#

Name

Source

2.1.8 2.1.8.1 2.1.8.2

The Nitrate/Nitrite porter (NNP) family Nitrite extrusion permease Bacteria Bacteria Nitrate uptake permease

2.1.9 2.1.9. I

The Phosphate:H+ symporter (PHS) family Pi uptake permease Yeast

Example ~~~

2.1.9.2

Pi uptake permease

Fungi

2.1.9.3

Pi uptake permease

Plants

The Nucleoside:H+ symporter (NHS) family 2.1.10 Bacteria 2.1.10. I Nucleoside permease Bacteria 2.1.10.2 Xanthosine permease

~

~

~

~

~~~~

~~

NarK of E. coli (spP10903) NasA of Bacillus subtilis (spP42432) Ph84 of Saccharornyces cerevisiae (spP25297) Pho-5 of Neurospora crassa (gbL36127) PTI of Solanum tuberosum (gbX98890) NupG of E. coli (spPO9452) XapB of E. coli (spP45562)

The 0xalate:Formate antiporter (OFA) family 2.1.1 1 Bacteria OxlT of E. coli (gbU40075) 2. I . 1 I , 1 The 0xalate:formate antiporter The Sialate:H+ symporter (SHS) family 2.1.12 2.1.12.1 The sialic acid permease Bacteria

NanT of E. coli (spP41036)

The Monocarboxylate antiporter (MCP) family 2.1.13 2. I . 13.1 The monocarboxylate Animals, yeast, Mot-I of Homo sapiens (lactate, pyruvate, fungi (pirA55568) mevalonate) uptake/ efflux permease The Anion:Cation symporter (ACS) family 2.1.14 2. I . 14.1 Glucarate permease Bacteria 2.1.14.2 Hexuronate permease 2.1.14.3 Putative tartrate permease 2. I. 14.4 Allantoate permease

Bacteria Bacteria Yeast

2.1.14.5 Phthalate permease

Bacteria

2.1.14.6 Na:Pi symporter

Animals

Aromatic Acid:H+ symporter (AAS) family 2.1.15 2. I . 15. I 4-Hydroxybenzoate Bacteria (Protocatachuate) permease 2. I . 15.2 3-Hydroxyphenyl Bacteria propionate permease 2.1.15.3 2.4-DichlorophenoxyBacteria acetate permease

GudT of Bacillus subtilis (spP42237) ExuT of E. coli (spP42609) TtuB of Agrobacterium vitis (gbU32375) Da15 of Saccharotnyces cerevisiae (spP15365) Phtl of Pseudomonas putida (spQO518 1) Nptl of Mus musculus (gbX7724 I ) PcaK of Pseudomonas putida (gbU10895) MhpT of E. coli (gbX97543) TfdK of Ralstonia eutropha (gbU16782)

104

MILTON H. SAIER, JR

Table 7 continued

TC#

Name

Source

Unknown Major Facilitator (UMF) family 2.1.16.1 Permease of unknown Yeast specificity

Example

2.1.16

The Cyanate Permease (CP) family 2.1.17.1 Cyanate transport Bacteria system

Ykrl06w of Saccharomyces cerevisiae (spP36173)

2.1.17

CynX of E . coli (spP17583)

Tuhk 8 The Resistance-Nodulation -Cell Division (RND) family (TC #2.6).

Members of the R N D family all probably catalyze substrate efflux via an H+ antiport mechanism (Paulsen et ul., 1996b). These proteins are found only in Gram-negative bacteria and fall into three phylogenetic clusters (Saier el a / . , 1994). Clustering pattern correlates with substrate specificity with cluster I catalyzing export of heavy metals, cluster 2 catalyzing export of multiple drugs, and cluster 3 probably catalyzing export of lipooligosaccharides concerned with plant nodulation for the purpose of symbiotic nitrogen fixation. These transport systems are large (950-1 100 amino acyl residues) possessing a single transmembrane spanner (TMS) at their N-termini followed by a large periplasmic domain, then six additional TMSs, a second large periplasmic domain, and five final C-terminal TMSs. In the case of one system (NolGHI) the system consists of three distinct polypeptide chains, but all others consist of a single polypeptide chain. The first halves of these systems are homologous to the second halves, and they therefore probably arose as a result of an intragenic tandem duplication event that occurred in the primordial system prior to divergence of the family members. These proteins function in conjunction with a 'membrane fusion protein' (MFP; T C #98.1) and an 'outer membrane factor' (OMF; TC #98.2) to effect efflux across both membranes of the Gram-negative bacterial cell envelope (Dinh, el ul., 1994). The generalized transport reaction catalyzed by R N D proteins is:

+

+

Substrates (in) nH'(out) + Substrate (out) nH+(in). Substrates: (a) heavy metals, (e.g. Co2+,ZnZf, Cd2+ and Ni"); (b) multiple drugs (e.g., tetracycline, chloramphenicol, fluoroquinolones, p-lactams, etc.); or (c) lipooligosaccharides (nodulation factors). TC #

Name

2.6.1.1

Heavy metal efflux Pump

Gram-negative bacteria

2.6.2.1

Multidrug resistance pump Putative lipooligosaccharide nodulation factor exporter

Gram-negative bacteria Gram-negative bacteria

2.6.3. I

Source

Example CzcA of Alcaligenes eutrophus (gbM91650) AcrE (EnvC) of E. coli (gbX57948) NolGHl of Rhizobium meliloti (gbX58632)

105

TRANSPORT PROTEIN CLASSIFICATION hmlly:

MC

<

MIP

<

MF

Figure 1 Reconstructed histories of three transport protein families. The three families represented are the mitochondria1 carrier (MC) family (left; Table 3), the major intrinsic protein (MIP) family (middle; Table 6) and the major facilitator (MF) superfamily (right; Table 7).

in prokaryotes. Indeed, members of the MIP family are found in both prokaryotes and eukaryotes. The major facilitator superfamily (MFS; Table 7) also arose by an intragenic duplication event, but in this case the primordial unit encoded a protein of about 225 amino acyl residues with six TMSs, and the product of the duplication event was a gene encoding a protein minimally of about 450 residues with 12 TMSs. All members of the MFS have 12 (or more; see below) TMSs, and these proteins occur ubiquitously throughout the living world (Table 7). The MFS is one of the two largest superfamilies of permeases found on earth, and it is probably also one of the oldest (see below). Finally, the RND family (Table 8) arose by a duplication event that clearly differed from those observed for the MIP family or the MFS. In this case, the genetic element that was duplicated encoded a large primordial protein of about 500 residues with six TMSs and a large, extracytoplasmic, hydrophilic domain. Present-day members of the RND family all possess 12 putative TMSs as well as two large, repeated, hydrophilic domains (Table 8). These proteins have been found only in Gram-negative bacteria. The degree of sequence similarity observed for the two halves of the RND family proteins clearly suggests that they arose relatively recently in bacteria, after subdivision of the living world into bacteria, archaea and eukaryotes over 2 billion years ago (Doolittle et al., 1996). Members of the RND family have not yet been found in Gram-positive bacteria, archaea or eukarya, suggesting an absence, or at least a minimal degree of horizontal gene transfer between the major prokaryotic and eukaryotic kingdoms of life.

106

MILTON H. SAIER, JR

When the established or putative topologies of the subunits of channel protein families (Class 1 families) are compared with those of carrier families (Class 2 families) (see Table I), a startling observation can be made (Fig. 2 ) . Channel protein subunits usually possess two to six TMSs (Fig. 2A) while carrier protein subunits usually possess 10-14 TMSs (Fig. 2B). Further, the channel proteins usually consist of oligomers while the carriers have often been found to be monomers. This interesting distinction (of which exceptions exist for both classes of proteins) suggests that these two classes of

20 0

t

-mu C

-

A

1 5 10-

c

0

5 -

0

-

. . . . 6

2 0 -

0

5 t

.u

1 5 -

10-

L.

a

0

5 -

... ... :..

. .

0

I/

~

0

5

i

C

201 5 -

1 0 -

.E C

=

5 -

0

0

. . I

I

. . . 1

I

I

I

I

2

4

6

8

10

12

14

**I/

I

24

No. TYSslpolypeptIde chain

Figure 2 Proposed topologies [numbers of putative transmembrane a-helical spanners (TMSs)] for channel (Class I; A) versus carrier (Class 2; B) protein families. The putative topologies of permease families functioning by unknown mechanisms (Class 99; C) are also represented. Each dot represents one of the families listed in Table I under the appropriate category). Only families with reasonably well-established topologies are represented in the figure.

TRANSPORT PROTEIN CLASSIFICATION

107

proteins have different structural requirements which allow them to be distinguished on the basis of topological features alone. Thus, one can predict with reasonable confidence what transport mode may be catalyzed by a recognized permease that functions by an unknown mechanism (Fig. 2C).

5. THE MAJOR FACILITATOR SUPERFAMILY (MFS)

When accounting for the phenomenon of solute transport, two superfamilies of permeases always come to mind, the major facilitator superfamily (MFS), and the ATP-binding cassette (ABC) superfamily. Of the many dozens of transporter families that are currently recognized, these two families are by far the largest and functionally most diverse. Together, they account for a major fraction (nearly one-half) of the recognized permeases found in microorganisms with completely sequenced genomes so far examined (Paulsen et al., 1998). Properties of the MFS are summarized in Table 9. Within the MFS are 17 currently recognized families, each generally specific for a different class of compounds (see also Table 7). MFS permeases transport almost any type of small solute including sugars, drugs, all kinds of metabolites and a variety of organic and inorganic cations and anions. These proteins can catalyze solute uniport, so1ute:cation symport and/or so1ute:solute or so1ute:cation antiport, a fact that has led Goswitz and Brooker (1995) to suggest the alternative designation for the MFS, the USA family, much to the chagrin of many scientists who live outside the United States. While some MFS members catalyze only one of these processes under normal physiological conditions, many catalyze two or even all three of these processes, depending on conditions. MFS proteins are usually of about 450 amino acyl residues in length although some are longer due to N-terminal or C-terminal hydrophilic extensions that are particularly prevalent in eukaryotes (Pa0 et al., 1998). They possess either 12 or 14 a-helical TMSs depending on family (Paulsen et

Table 9 Properties of the Major Facilitator Superfamily (MFS).

I. 2. 3. 4. 5. 6.

17 families, specific for sugars, drugs, metabolites, anions, etc. Consists of symporters, antiporters and uniporters. Size, -400 residues or larger; 12 or 14 spanners. Found in bacteria, archaea and eukaryotes. Nearly 500 sequenced members. Well-characterized members: Lacy, Glut1 and TetB.

108

MILTON H. SAIER, JR

af., 1996a). Thus, 14 of the 17 MFS families have members with 12 TMSs while the remaining three families have members with 14 TMSs (Pa0 rf af., 1998). The three 14 TMS families are believed to have arisen independently of each other from different 12 TMS precursors long after the intragenic duplication event that gave rise to the 12 TMS topology that is characteristic of most MFS permeases. At least in one case, that of the drug:H+ antiporter family with 14 TMSs (DHA14; TC #2.1.2; see Table 7), the two extra TMSs are found between the two six-TMS units of these permeases. MFS permeases are found ubiquitously in all organisms for which they have been sought, including members of the three major kingdoms of life the Bacteria, Archaea and Eukarya. Nearly 400 members of the superfamily are currently available for analysis (Tables 7 and 9; Pao ef al., 1998). Wellcharacterized members include the lactose permease of E. coli (Lacy; TC #2.1.5.1; Kaback, 1986; Varela and Wilson, 1996) and the hexose facilitators of mammals (TC #2.1.1.12 and #2.1.1.13) (Baldwin, 1993). Figure 3 illustrates one of the mechanisms of energy coupling catalyzed by MFS permeases. While some permeases catalyze the free facilitation of their molecular substrates across the membrane in an energy uncoupled process (uniport), others catalyze the active uptake of nutrients using a nutrierkcation symport mechanism. In such a process, energy is provided for nutrient accumulation when the cation (either H+ or Na') flows down its electrochemical gradient into the cell. Alternatively, a substrate:cation antiport mechanism can be used. In such a process, energy is provided for the active extrusion of a toxic substance or an end product of metabolism ~

Environment

ti'

Figure 3 lllustration of solute:H+ antiport (MFS). The figure illustrates the process of drug:H+ antiport catalyzed by the QacA protein of Stciphylococcus mmws (TC #2.1.3.4; Table 7). In this process, drug efflux is driven by the f o w of protons. down their electrochemical gradient, into the cell.

109

TRANSPORT PROTEIN CLASSIFICATION

because expulsion is coupled to uptake of the cation, again, down its electrochemical gradient. This last mentioned process is illustrated in Figure 3. Many uniporters, so1ute:H + symporters and so1ute:H antiporters also catalyze so1ute:solute antiport. Sometimes this process is of greater physiological significance than uniport or the cation coupled process. So1ute:solute antiport is particularly important when the nutrient taken up is converted to a structurally related metabolic end product that must be extruded from the cell. The extrusion of formate in exchange for oxalate, taken up into E. coli cells via the OxlT permease (TC #2.1.11.1; Table 7) is a prime example. Others include the 0rganophosphate:Pi antiporter (OPA) family members (TC #2.1.4; see Table 7). The two permease topologies found within the MFS (12 and 14 TMSs) are shown in Fig. 4A and B, respectively, and an unrooted phylogenetic tree for these permeases is shown at the bottom of Fig. 4. Representative members of most of the 17 recognized MFS families (see Table 7) are included (Pa0 ef al., 1998). Examination of the tree reveals that some families are apparently more closely related to each other than they are to other families. For example, the DHA14 and DHA12 families (Table 7), both of which catalyze drug:H+ antiport, diverged from each other later than they did from any other MFS families (Fig. 4, bottom). Other examples of related MFS families can be identified by examination of Fig. 4 (see Pao et al., 1998 for a more detailed discussion of the MFS evolutionary process). +

6. THE ATP-BINDING CASSElTE (ABC) SUPERFAMILY

Only one permease superfamily is larger and more diverse in function than is the MFS, and that is the ABC superfamily (Tables 10 and 11). This immense superfamily consists of at least 40 currently recognized clusters or families of permease systems. The individual permeases exhibit specificity for virtually any kind of small molecule as do the permeases of the MFS, but in addition they can transport macromolecules such as proteins, complex carbohydrates and lipids (Table 10). Many of these permeases are capable of transporting multiple substrates. They fall into three major phylogenetic and functional categories: prokaryotic uptake systems, prokaryotic efflux systems and eukaryotic efflux systems (Table 11). ABC-type uptake systems are apparently lacking in eukaryotes. Uptake or efflux of substrates of ABC permeases is almost always driven by ATP hydrolysis. These multidomain, multicomponent systems exhibit a total size of over 1000 amino acyl residues each and usually (but not always) possess a 12 ( 6 + 6 ) TMS topology. The efflux systems are found ubiquitously. About 400 sequenced members of the ABC superfamily are available

110

MILTON H. SAIER, JR

A MFS-12TMS

B

MFS-I4TMS

,#%

FigIJre 4 The two topological types of permeases found within the MFS (top; A and B, respectively) and the phylogenetic tree for the MFS (bottom). Fourteen of the 17 MFS families exhibit the topology shown in A with 12 putative or established transmembrane a-helical spanners (TMSs) while the remaining three families exhibit the 14 TMS topology. Most of the 17 families included within the MFS (Table 7) are to be found within the unrooted tree shown (see Table 7 and Pao rt a/., 1998 for abbreviations of the individual permeases represented).

TRANSPORT PROTEIN CLASSIFICATION

111

Table 10 Properties of the ATP-binding Cassette (ABC) superfamily ~

1.

40 families, specific for sugars, amino acids, ions, drugs, antibiotics, vitamins, iron complexes, peptides, proteins, complex carbohydrates, etc. Driven by ATP hydrolysis. Multicomponent, multidomain systems; total size, > 1000 residues, usually 12 (6 + 6) spanners. Found in bacteria, archaea and eukaryotes. About 500 sequenced members. Well-characterized members: MalEFGK, MDR and CFTR.

2. 3. 4. 5. 6.

for analysis. Well-characterized members include the maltose uptake permease of E. coli (TC #3.1.1.1), the multidrug resistance pump (MDR; TC #3.1.61.1) and the cystic fibrosis transmembrane conductance regulator (CFTR; TC #3.1.62.1). Both of these mammalian efflux permeases may be able to catalyze efflux of a variety of substances including anions, peptides and hydrophobic drugs. The bacterial uptake permeases, which consist of two integral membrane units and two water-soluble ATP-hydrolyzing units (both either homo- or heterodimeric or fused), function in conjunction with an extracytoplasmic solute-binding receptor. The three-dimensional structures of many of these receptors have been determined by X-ray crystallographic analyses, and although their sequences are very divergent, they all exhibit similar structures, suggesting a common phylogenetic origin (Quiocho and Ledvina, 1996). In conducting phylogenetic analyses of ABC permease constituents, we were interested in answering three unrelated questions: 1.

Did all or most ABC permeases evolve from a single primordial system without appreciable shuffling of their protein constituents between systems, or did these systems exchange constituents during their evolution? 2. Did the three essential constituents (extracytoplasmic receptors, transmembrane channel-forming proteins and cytoplasmic energycoupling, ATP-hydrolyzing constituents) evolve at similar or dissimilar rates? 3. What was the primary driving force governing the relative rates of sequence divergence upon which phylogenetic tree construction is based: the evolutionary process or restrictions imposed upon the process of sequence divergence due to functional constraints? To answer these questions, phylogenetic trees were constructed. The receptors analyzed fell into several distinct families as revealed, for example,

112

MILTON H. SAIER, JR Table 1 I

The ATP-binding Cassette (ABC) Superfamily (TC #3.1).

The ABC superfamily contains both uptake and efflux transport systems, and the members of these two permease groups generally cluster together with just a few exceptions. ATP hydrolysis without protein phosphorylation energizes transport. There are dozens of families within the ABC superfamily and each family generally transports a single class of compounds. Thus, family classification correlates with substrate specificity (Tam and Saier, 1993; Saurin and Dassa, 1994; Kuan ef al., 1995). However, there are exceptions (Fath and Kolter, 1993; Saurin and Dassa, 1994; Paulsen el al., 1998). The permeases of the ABC superfamily consist of two integral membrane domains/ proteins and two cytoplasmic domains/proteins. The uptake systems (but not the efflux systems) additionally possess extracytoplasmic solute-binding receptors (one or more per system) which in Gram-negative bacteria is found in the periplasm, and in Gram-positive bacteria is present either as a lipoprotein, tethered to the external surface of the cytoplasmic membrane, or as a cell surface-associated protein, bound to the external membrane surface via electrostatic interactions. Both the integral membrane channel constituent(s) and the cytoplasmic ATP-hydrolyzing constituent(s) may be present as homodimers or heterodimers. In many of these permeases, the various domains are fused in a variety of combinations. Uptake permeases generally have their constituents as distinct polypeptide chains, while efflux systems usually have them fused. ABC-type uptake systems have not been identified in eukaryotes, but ABC-type efflux systems abound in both prokaryotes and eukaryotes. The eukaryotic efflux systems often have the four domains (two cytoplasmic domains and two integral membrane domains) fused into either one or two polypeptide chains. The integral membrane permease domains each usually possesses five (uptake) or six (efflux) transmembrane spanners, but exceptions exist. The three structurally dissimilar constituents of the ABC permeases have generally arisen from a common ancestral permease system with minimal shuffling of constituents between systems. Thus, phylogenetic clustering of the three protein/domain constituents is almost always the same. However the rates of sequence divergence differ drastically with the extracytoplasmic solute-binding receptors diverging most rapidly, the integral-membrane channel-forming constituents diverging at an intermediate rate, and the cytoplasmic ATP-hydrolyzing constituents diverging most slowly. Thus, all ATP-hydrolyzing constituents are demonstrably homologous, but this is not true for the integral membrane constituents or the receptors. Nevertheless, clustering patterns are generally the same for all three types of proteins, and three-dimensional structural data suggest that, in spite of their extensive sequence divergence, the extracytoplasmic solute-binding receptors are homologous to each other. The generalized transport reaction for ABC-type uptake systems is: Solute (out) ATP + Solute (in) ADP P,. The generalized transport reaction for ABC-type efflux systems is: Substrate (in) + ATP + Substrate (out) ADP + P,.

+

+

+

+

ABC-type uptake permeases (all from bacteria) ___~

TC#

~

~

~~

Name

Cluster 1: carbohydrates# I 3.1. I . I Maltooligosaccharide permease

~

Example MalEFGK of E. coli MalE (receptor (R)): spPO2928 MalF (membrane (M)): spPO2916 MalG (membrane (M)): spPO2622 MalK (cytoplasmic (C)): spP029 I4

113

TRANSPORT PROTEIN CLASSIFICATION

Table 11 continued

ABC-type uptake permeases (all from bacteria) TC#

Name

3. I. 1.2

Multiple sugar (melibiose; raffinose, MsmEFGK of Streptococcus mutans MsmE (R): spQOO749 etc) permease MsmF (M): spQ00750 MsmG (M): spQOO751 MsmK (C): spQOO752 UgpABCE of E. coli Glycerol-phosphate permease UgpB (R): spP10904 UgpA (M): spP10905 UgpE (M): spP10906 u g p c (C): spP10907 LacEFGK of Agrobacterium Lactose permease radiobacter LacE (R): spP29822 LacF (M): spP29823 LacG (M): spP29824 LacK (C): spP29825

3.1.1.3

3. I . 1.4

Cluster 2: carbohydrates #2 3.1.2.1 Ribose permease

3.1.2.2

Arabinose permease

3.1.2.3

Galactose/glucose (methyl galactoside) permease

Example

RbsABCD of E. coli RbsB (R): spPO2925 RbsC (M): spPO4984 RbsD (M): spPO4982 RbsA (C): spPO4983 AraFGH of E. coli AraF (R): spPO2924 AraG (C): spPO8531 AraH (M): spPO8532 MglABC of E. coli MglA (C): spP23199 MglB (R): spPO2927 MglC (M): spP23200

Cluster 3 polar amino acids and derivatives 3.1.3.1 Histidine; arginine/lysine/ornithine HisJ (histidine receptor)-ArgJ (arg/lys/orn receptor)-HisMPQ of permease Salmonella typhimurium HisJ (R): spP02910 ArgJ (R): spPO2911 HisM (M): spPO2912 HisQ (M): spPO2913 HisP (C): spPO2915 GlnHPQ of E. coli 3.1.3.2 Glutamine permease GlnH (R): spP10344 GlnP (M): spP10345 GlnQ (C): spP10346

114

MILTON H. SAIER, JR

Table I 1

continued

ABC-type uptake permeases (aU from bacteria) TC#

Name

Example

3.1.3.3

Arginine permease

3.1.3.4

Glutamate/aspartate permease

3. I .3.5

Octopine permease

3.1.3.6

Nopaline permease

ArtJ (arginine receptor)/ArtI (receptor of unknown specificity)-ArtMQP of E. coli ArtP(C): spP30858 ArtQ(M): spP30861 ArtM(M): spP30862 ArtJ( R): spP30860 Artl(R): spP30859 GltJKLX of E. coli GltJ (M): spP41074 GltK (M): spP41075 GltL (C): spP41076 GltX (R): not available OccQMPT of Agrohacterium tumefaciens OccT (R): gbM77784 OccQ (M): gbM77784 OccM (M): gbM77784 OccP (C): gbM77784 NocQMPT of Agrobacterium tumefacicws NocT(R): gbM77785 NocQ (M): gbM77785 NocM (M): gbM77785 NocP (C): gbM77785

Cluster 4 hydrophobic amino acids 3. I .4.1 Leucine; leucine/isoleucine/valine permease

Cluster 5: peptides and nickel 3.1.5.1 Oligopeptide permease

3.1.5.2

Dipeptide permease

LivK (leucine-specific receptor)-LivJ (Leu/Ile/Val receptor)-LivHMGF LivJ (R): spPO2917 LivK (R): spPO4816 LivH (M): spP08340 LivM (M): spP22729 LivG (C): spP22730 LivF (C): spP2273 1 OppABCDF of Salmonella typhimurium OppA (R): spP06202 OppB (M): spP08005 OppC (M): spP08006 OppD (C): spPO4285 OppF (C): spP08007 DppABCDE of Bacillus subtilis DppA (C): spP26902 DppB (M): spP26903

115

TRANSPORT PROTEIN CLASSIFICATION

Table 11 continued

ABC-type uptake permeases (all from bacteria) TC#

3.1.5.3

Name

Nickel permease

Cluster 6 sulfate and nitrate 3.1.6.1 Sulfate/thiosulfate permease

3.1.6.2

Nitrate permease

Cluster 7: phosphate 3.1.7.1 Phosphate permease

Cluster 8: molybdate 3.1.8.1 Molybdate permease

Cluster 9 phosphonates 3. I .9.1 Phosphonate/organophosphate ester permease

Example DppC (M): spP26904 DppD (C): spP26905 DppE (R): spP26906 NikABCDE of E. coli NikA (R): gbX73143 NikB (M): gbX73143 NikC (M): gbX73143 NikD (C): gbX73143 NikE (C): gbX73143 Sbp (sulfate receptor)-CysP (thiosulfate receptor)-CysTWA Sbp (R): spPO6997 CysP (R): gbM32101 CysT (M): gbM32101 CysW (M): gbM32101 CysA (C): gbM32101 NrfABCDX of Synechococcus sp (PCC7942) NtrA (M): spP38043 NtrB (M): spP38044 NtrC (C): spP38045 NtrD (C): spP38046 NtrX (R): not available PhoS (phosphate receptor)-PstABC of E. coli PhoS (R): gbK01992 PstA (M): gbK01992 PstC (C): gbK01992 PstB (C): gbK01992 ModABC of E. coli ModA (R): gbL34009 ModB (M): gbL34009 ModC (C): gbL34009 PhnCDE of E. coli PhnC (C): spP16677 PhnD (R): spP16682 PhnE (M): spP16683

MILTON H. SAIER, JR

116 Table 11 continued

ABC-type uptake permeases (all from bacteria) TC#

Name

Cluster 1 0 iron 3. I . 10.I Iron permease

Cluster 11: polyamines 3. I . 1 1. 1 polyamine (putrescine/spermidine) permease

Cluster 12: glycine/betaine Glycine/betaine permease 3. I . 12. I

Cluster 13: vitamin BI2 3.1.13.1 Vitamin B12permease

Cluster 14: iron chelates 3.1.14.1 Iron-enterobactin permease

3.1.14.2

Iron-dicitrate permease

3.1.14.3

Iron-hydroxamate permease

Cluster 1 5 manganese ions 3.1.15.I Manganese permease

Example SfuABC of Serratia marcescens SfuA (R): spP21408 SfuB (M): spP21409 SfUC (C): spP21410 PotABCD of E. coli PotA (C): gbM64519 PotB (M): gbM64519 PotC (M): gbM64519 PotD (R): gbM64519 ProVWX of E. coli Prow (M): gbK01992 ProX (R): gbK01992 ProV (C): gbK01992 BtuECD of E. coli BtuC (M): gbM14031 BtuD (C): gbM1403I BtuE (R): gbM14031 FecBCDE of E. coli FecB (R): gbM26397 FecC (M): gbM26397 FecD (M): gbM26397 FecE (C): gbM26397 FepBCDG of E. coli FepB (R): spP14609 FepC (C): spP23878 FepD (M): spP23876 FepG (M): spP23877 FhuBCD of E. coli FhuB (M): spPO6972 FhuC (C): spPO7821 FhuD (R): spPO7822 MntABC of Synechocystis 6803 MntA (C): gbL34630 MntB (M): gbL34630 MntC (R): gbL34630

117

TRANSPORT PROTEIN CLASSIFICATION

Table 1 1 continued

ABC-type efflux permeases (bacterial) TC#

Name

Source

3.1.31.1

Capsular polysaccharide exporter

Gram-negative bacteria

3.1.32.1 3.1.33.1

3.1.34.1 3.1.35.1

3.1.35.2

3.1.35.3 3.1.35.4 3.1.35.5 3.1.36.1 3.1.37.I

3. I .38.1 3.1.39.1 3.1.39.2 3.1.40.1 3.1.40.2 3.1.41.1

Example

KpsMT of E. coli KpsM (M): spP24584 KpsT (C): spP24586 Lipo-oligosaccharide Gram-negative NodIJ of Rhizobium galegae exporter bacteria NodJ (M): gpX87578 Nod1 (C): gpX87578 Lipopol ysaccharide Gram-negative RfbAB of Klebsiella exporter pneumoniae bacteria RfbA (M): gbL41518 RfbB (C): gpL41518 Teichoic acid exporter Gram-positive TagGH of Bacillus subtilis bacteria TagG (M): gpU13832 TagH (C): spP42954 Daunorubicin; doxorubicin Gram-positive DrrAB of Streptomyces (drug resistance) peucetius bacteria exporter DrrA (C): spP32010 DrrB (M): gpM73758 Oleandomycin (drug Gram-positive OleC4-01eC5 of resistance) exporter bacteria Strep tomyces an I ibio ticus OleC4 (C): pirS32904 OleC5 (M): pirS32909 Macrolide (drug resistance) Gram-positive SrmB of Streptomyces exporter bacteria ambofaciens (gbX63451) Erythromycin (drug Gram-positive MsrA of Stnphylococcus resistance) exporter epdermidis (gbX 5208 5) bacteria Tylosin (drug resistance) Gram-positive TlrC of Streptomyces fradiae exporter bacteria (gbM57437) Microcin B17 exporter Enteric bacteria McbEF of E. coli McbE (M): spPO5528 McbF (C): spPO5529 Heme exporter Gram-negative CycVWX of Bradyrhizobium japonicum bacteria CycV (C): spP30963 CycW (M): spP30964 CycX (M): spP30959 P-glucan exporter Gram-negative NdvA of Rhizobium meliloti (spP18767) bacteria cr-hemolysin exporter Gram-negative HlyB of E. coli (spPO8716) bacteria Cyclolysin exporter Gram-negative CyaB of Bordetella pertussis (spP 18770) bacteria Protease exporter Gram-negative PrtD of Erwinia chrysanthemi (spP23956) bacteria Enteric bacteria CvaB of E. coli (spP22520) Colicin V exporter Hemolysin/bacteriocin Gram-positive CylB of Enterococcus faecalis (gbM38052) exporter bacteria

118

MILTON H. SAIER, JR

Tuhle I I

continued

ABC-type efflux permeases (bacterisl) TC#

Name

Source

Example

3.1.41.2

Subtilin (toxic peptide) exporter Competence factor exporter Pediocin PA- 1 exporter

Gram-posi t ive bacteria Gram-posi tive bacteria Gram-positive bacteria Gram-negative bacteria Gram-nega tive bacteria Gram-positive bacteria

SpaB of Bucillus suhtilis (spP33116) ComA of Streptococcus pneumoniue (spQ03727) PedD of Pediococcus ucidiluctici (spP364Y7) SyrD of Pseudomonus s.vringue (spP3395 1 ) HasADE of Serratiu murce.wen.s (gbX8 1 195) NatAB of Bucillus suhtilis (gbU38073)

3.1.42. I 3. I .42.2 3. I .43.1 3. I 4 . 1

3.1.45.1

Siderophore exporter (drug exporter) Fe uptake transporter ( Lantibiotic exporter) Na' efflux pump NatAB

ABC-type efflux permeases (mostly eukaryotic) TC#

Name

Source

3.1.61.1

Multidrug resistance (MDR) efflux pump (peptide efflux pump; phospholipid flippase) Cystic fibrosis transmembrane conductance regulator (CFTR); cyclic AMPdependent chloride channel Peroxysomal transporter associated with Zellweger syndrome Eye pigment precursor transporter Sporidesmin toxicity suppressor (STSI) (MDR) a-Factor sex pheromone exporter (STE6) Metal resistance protein (yeast cadmium factor YCFI)

Animals, fungi, MDRl of Homo sapiens bacteria (spP08 183)

3. I .62.1

3.1.63.1 3 . I .M.I

3. I .65. I 3.1.66.1 3.1.67. I

Example

Animals

CFTR of Homo siipiens (spP1356Y)

Animals

PMP7 of Homo sapiens (spP28288)

Animals

Whit of Drosophilu melanogaster (spP 10090) STS 1 of Succhurotnyces cerevisiuc (gbX74113)

Yeast Yeast Yeast

STE6 of Sacchuromyces cerevisiue (gbX15428) YCFI of Sacchuromyces cercvisiue (gbL35237)

119

TRANSPORT PROTEIN CLASSIFICATION

Table I I

continued

ABC-type efflux permeases (mostly eukaryotic)

TC#

Name

Source

Example

3.1.68.1

Multi-drug resistanceassociated protein, MRP. (Leukotriene; glutathione conjugates; drug exporter) MHC peptide exporter (TAP) Pleiotropic drug resistance (PDR) exporter; steroid exporter

Animals

MRP of Rattus norvegicus (gbX90642)

Animals

TAP2 of Homo sapiens (gbZ22935) Pdr5 of Saccharomyces cerevisiue (gbL19922)

3.1.69. I 3. I .70.1

Yeast

in Fig. 5A-D (Tam and Saier, 1993). Figure 5A includes receptors for one group of sugars (carbohydrates #1, T C #3.1.1 in Table 1 1 ) as well as for iron (TC #3.1.10), and Fig. 5B includes receptors for a second group of sugars (carbohydrates #2, T C #3.1.2 in Table 1 1). Figure 5C shows representative receptors specific for polar amino acids and their derivatives (TC #3.1.3 in Table 1 I), and Fig. D presents the phylogenetic tree for receptors specific for peptides and nickel (TC #3.1.5 in Table 1 I). When a phylogenetic tree was constructed for the ATP-hydrolyzing, energy-coupling constituents of corresponding permeases (Fig. 6), clustering patterns were found to be similar to those of the receptors shown in Fig. 5. Thus, sugar # I permease energizers all clustered together (top of the tree shown in Fig. 6); sugar #2 permease energizers clustered together (lower right side of the tree shown in Fig. 6 ) , and the polar amino acid energizers (lower left) and peptide energizers (middle left) each clustered into a coherent group. The integral membrane constituents of the ABC-type uptake permeases exhibited similar clustering patterns (Saurin and Dassa, 1994). These observations suggested that the permease constituents of all of these systems had evolved with minimal shuffling of constituents between systems during their evolution. The fact that the energy coupling proteins could all be included within a single phylogenetic tree, while those of the integral membrane constituents and receptors could not, argued that the rates of sequence divergence for the receptors was substantially greater than those of the energizers, and that the integral membrane constituents of these systems diverged at an intermediate rate (Saurin and Dassa, 1994; Kuan et al., 1995). The fact that clustering patterns were the same for the three permease constituents provided strong evidence for the conclusion that the evolutionary process rather than restrictions imposed upon the proteins due to substrate

A

Glp G o Chv ALU

Siu Smu

AnGo

: AIu

Figure 5 Phylogenetic families of extracytoplasmic receptors that function in conjunction with ABC permeases. The abbreviations of the proteins and the original analyses are described in Tam and Saier, (1993). Most of the proteins represented are presented in Table 1 1 (Reproduced from Saier. 1994. with permission.) See text for explanation.

121

TRANSPORT PROTEIN CLASSIFICATION MsmK Smu ‘

NikD Eco

\

PotA Eco 8

I

[I4 ., / 1’

UgpC E cob ,I MalK Eco\2g321 NikE Eco

PotG Eco

CysA Ssp-

CysA ECO MbpX Mpo

SfuC Sma

ModC Rca ModC Eco

FepC E c o y

FecE Eco GlnQ E c o ’ y GlnQ B s t L p

0,c P AtU NocF LivF EciD

BraG Pae

AraG Eco

Figure 6 Phylogenetic tree for the cytoplasmic energy coupling, ATP-hydrolyzing constituents of bacterial ABC uptake permeases. The protein abbreviations and phylogenetic tree construction were as described in Kuan et al. (1995). Most of the proteins represented are presented in Table 11 which also provides the abbreviations used. (Reproduced from Kuan er al., 1995, with permission.)

recognition was the primary force driving sequence divergence (Kuan et al., 1995). On the other hand, functional constraints presumably gave rise to the different rates of evolutionary divergence observed for the three different constituents of these systems (Saier, 1994, 1996).

7. PROKARYOTIC GENOME SEQUENCE ANALYSES

Recently we have analyzed the completely sequenced genomes of six prokaryotes: two Gram-negative bacteria (E. coli and Haemophilus infZuenzae); two Gram-positive bacteria (Mycoplasma genitalium and M . pneumoniae); one cyanobacterium (Synechocystis PCC 6803); and one archaeon (Methanococcus jannaschii) for their complements of cytoplasmic membrane permeases. In addition, we analyzed the 50% of the Bacillus subtilis genome that was available at the time of our analyses (Paulsen et al., 1998; see this

122

MILTON H. SAIER, JR

reference for primary references describing the various genome sequencing efforts). We have similarly analyzed the complete genome of the eukaryotic organism, Sacchuromyces cerevisiue, but these results will not be discussed here. Within the seven prokaryotic genomes analyzed, 62 cytoplasmic membrane solute permease families were identified, and the occurrence of permease family members in each organism examined was tabulated (Paulsen ef ul., 1998). All of these families are included within groups 1,2, 3 , 4 and 99 in Table 1. Of these 62 families, only four families were represented in all seven of the organisms analyzed. These four families were the MF and ABC superfamilies discussed above, the so-called amino acid-polyamine-choline (APC) family (TC #2.3), in which individual permeases are specific for the three classes of compounds mentioned, and the H+- or Na+-translocating F, Fo-ATPases (F-ATPase superfamily; TC #3.2). While many members of each of the first three of these families are encoded within the genomes of most of these organisms, only one F-ATPase family member is encoded within the genome of each of these prokaryotes. Most, but not necessarily all, of these F-ATPases are probably orthologous. Interestingly, 58 of the 62 families represented were identified in E. coli, showing that this organism provides an excellent model organism for understanding prokaryotic transport. We estimate that about 80% of all cytoplasmic membrane transporters encoded within the six fully sequenced genomes analyzed have been identified. Some of the major conclusions resulting from these analyses are summarized below.

I . The numbers of solute transporters encoded within eubacterial genomes are approximately proportional to genome size (10% of all genes), but are two-fold lower for cyanobacteria and archaea. The first observation, that the number of solute transporters encoded within a genome is approximately proportional to genome size, was particularly surprising in view of the fact that E. coli can biosynthesize essentially all of its biosynthetic precursors and vitamins while M . genitalium, with only one-tenth the amount of DNA, can make very few of them. The latter organism must obtain these essential nutrients from exogenous sources, presumably as a result of the activities of nutrient uptake permeases. One must hypothesize that M . gmituliurn permeases exhibit broad specificity in contrast to those characterized in E. coli which usually exhibit a high degree of specificity for just one or a few compounds. The fact that the archaeon, M . jannuschii. and the cyanobacterium, Synechocystis PCC 6803, exhibit a two-fold lower percentage of transport genes correlates with finding 2 (below).

TRANSPORT PROTEIN CLASSIFICATION

123

2 . M . jannaschii and Synechocystis exhibit a 2- to 3-fold greater percentage of transporters for inorganic ions and a concomitant decrease in transporters for organic compounds. The fact that permeases for organic compounds greatly predominate over those for inorganic compounds provides a partial explanation for the lower numbers of permeases encoded (finding 1 above). The differences in specificity noted presumably reflect the distinctive life styles (i.e. metabolic activities) of these bacteria. 3. About half of the 62 families represented within the prokaryotic genomes analyzed have representation in eukaryotes. Some 15% are restricted to Gram-negative bacteria, but only one family is restricted to Gram-positive bacteria. None is found exclusively in archaea or cyanobacteria. To what extent this last observation reflects the limited amount of sequence and biochemical data available for the archaea and cyanobacteria cannot be established at this time. 4. Proteins of one to three TMSs have far fewer homologues than proteins of zero or more than three TMSs, implying either more rapid evolutionary divergence or independent histories. We suggest that this surprising observation has an explanation in the types of functions most frequently performed by integral membrane proteins of one to three TMSs. Perhaps many of these proteins serve structural rather than catalytic roles, allowing more rapid sequence divergence from their primordial proteins. Rapid sequence divergence would be expected to mask the common ancestry of many of these proteins and hence limit the numbers of identifiable homologues in the databases. 5. Two superfamilies (ABC and MFS) account for nearly 50% of all transporters in each of the six bacteria examined. This observation is even more surprising when viewed in terms of point 6. 6. The ratio of ABC to MFS permeases varies over a 10-fold range, depending on organism and energy availability. Thus, the E. coli genome encodes 63 ABC permeases and 64 MFS permeases, but M . genitalium, with a total of 22 identified permeases, has 11 ABC permeases and only one MFS permease. This skewed distribution of permease types correlates with point 7. 7. Bioenergetics of transport frequently correlates with the primary source of energy generated via available metabolic pathways. Thus, E. coli possesses both substrate-level phosphorylation for the synthesis of ATP and electron flow for the primary generation of a proton electrochemical gradient (pmf) while M . gmitalium lacks an electron transport chain and therefore generates energy only by substrate-level phosphorylation. The primary availability of ATP to M . genitalium correlates with a preponderance of ATP-

124

MILTON H. SAIER, JR

dependent transporters. This explanation cannot, however, explain the relative distribution of ABC- versus MFS-type permeases in Synechocystis PCC 6803 which also has a ratio of ABC-type to MFS-type permeases of about 10:1. Synechocystis catalyzes both substrate-level phosphorylation and electron flow. However, this organism normally lives in freshwater ponds where nutrient concentrations are, in general, very low. ATP-driven permeases can accumulate their substrates against much greater concentration gradients than can pmf-driven permeases, and the former systems usually do so with higher affinities for substrate. In this case, the ecological niches in which' these organisms find themselves may explain the observed distribution of permease types. 8. Finally, &13% of all solute transporters are drug efflux pumps with comparable percentages in pathogens and non-pathogens. This fact suggests that the active extrusion of end products of metabolism and toxic substances is probably important to all prokaryotic organisms. Furthermore, the use of antibiotics and other drugs in medicine did not appreciably enhance the distribution of these efflux permeases encoded within the genomes of pathogens.

8. INDEPENDENT EVOLUTION OF DISTINCT TRANSPORT

MODES AND ENERGY-COUPLING MECHANISMS As noted above, we have classified transport systems on the basis of four criteria: 1. Permeases were first grouped according to transporter type and mode of energy coupling. 2. Each permease type was subdivided into recognizable families. 3. Each family was subdivided into phylogenetic clusters. 4. Each cluster was subdivided according to substrate specificities of the individual permeases. Table 12 identifies the ten different permease types listed in greater detail in Table 1 and summarizes the distributions in the various families of each type in the three major kingdoms of life, the Bacteria (B), the Archaea (A) and the Eukaryotes (E). The ten types include: (i) channels (except porins); (ii) secondary carriers; (iii) ATP-driven primary carriers; (iv) phosphoenolpyruvate (PEP)-driven, sugar-transporting group translocators; (v) organic acid decarboxylation-driven Na' pumps; (vi) electron flow-driven H' or Na' pumps; (vii) light-driven ion pumps; (viii) mechanically driven ion pumps; (ix) methyl transfer-driven Na' pumps; and (x)

125

TRANSPORT PROTEIN CLASSIFICATION

Table 12 Distribution of transporter types in the three major domains of living organisms.'

Transporter type Channels Secondary carriers ATP-driven carriers PEP-driven carriers Decarboxylation-driven carriers Electron flow-driven carriers Light-driven carriers Mechanically driven carriers Methyl transferase-driven carriers Porins

B 3 1

2 6 1 1 0 1 0

8

A

E

BA

BE

AE

BAE

0

6

0

0

7 0 0 0 0 0 0 0 1

2 10 0 0 0 3 0 0 0 0

0 0 0 0 0 0 0 0 0 0

1 8

0 0 0 0 1 0 1

1 0

1 3 0

0 0

0 0 0 0 0

5 0

0 1 0 0 0 0

The number indicated in each category (B, bacteria; A, archaea and E, eukarya) for each entry represents the number of families found in this category at the time this chapter was prepared. BA, found in both bacteria and archaea, but not eukarya. BE, found in both bacteria and eukarya, but not archaea. BAE, found in all three domains. The categories exhibiting a majority or a large number of family entries are indicated in bold print.

porins, outer membrane channels having exclusively P-structure. Four of these transporter types include families that are found in at least two, and probably all three, of the primary domains of life. These four transporter types include channels (Class l), secondary carriers (Class 2), ATP-driven primary carriers (Class 3) and electron flow-driven proton pumps (Class 6). Each of these four categories includes transporter families that are represented only in bacteria or eukaryotes, but several of the families included within each of these four categories are represented in both bacteria and eukaryotes, or even in bacteria, archaea and eukaryotes. On the other hand, all other energy-coupled transporter types are restricted to just one of the major domains of life. These unique types of energy-coupling mechanisms are presented in more detail in Table 13. The light-driven, Ht-and Cl--transporting bacteriorhodopsin (BR) family members are found in just one subgroup of the archaea, the halotolerant archaea, while the Na+-transporting methy1tetrahydromethanopterin:coenzyme M methyl transferase (NaT-MMM) family members are found in another archaeal subgroup, the methanotrophs (Table 13). All other energycoupled transport processes listed in Table 13 are restricted to bacteria. These include the PEP:sugar phosphotransferase systems (PTS) which modify their sugar substrates during transport; the unique Na+-transporting NADH dehydrogenase (NaNDH) family which is not homologous to

126

MILTON H. SAIER, JR

Tfihle 13 Novel energy-coupling mechanisms found in only one Kingdom

Family

TC #

PTS

4.1 6

NaT DC

5.1

Na NDH BR

6.5

Mot

8. I

Porins

9.1-1.14

NaTMMM

11.1

7.1

Energy-coupling mechanism Kingdom Phosphory I transfer Decarboxylation Electron flow Light absorption Flagellar rotation None Methyl transfer

No. of sequenced members

No. of substrate classes

Bacteria

30

I

Bacteria

10

1(3)

Bacteria Archaea

1 10

I 2

Bacteria

10

I

Bacteria; 100 eukaryotic organelles Archaea 2

10

I

members of the Ht-transporting NADH dehydrogenase family; the flagellar motor (Mot) family of cation (H’ or Na’) transporters, and the P-type porins found in the outer membranes of Gram-negative bacteria and eukaryotic organelles. It is interesting to note that the protein complexes of the F-ATPase family may in fact couple proton flux through the Fo channel of the complex to the ‘mechanical’ rotation of the FI stalk of the complex which catalyzes ATP synthesis or hydrolysis (Noji et al., 1997). Thus, this one family may actually use a mechanical device to couple ATP synthesis/hydrolysis to proton transport. This unique family includes distantly related archaeal and vacuolar ATPases as well as the better studied F-type ATPases. It is presumably an ancient family (Blair et al., 1996). Based on these observations, we suggest that: (i) channels. secondary carriers, ATP-driven primary carriers and electron-flow-driven H+ pumps are ubiquitous. They undoubtedly arose before divergence of the three kingdoms of life; and (ii) all other energy-coupling mechanisms are kingdomspecific, none being found in eukaryotes. They may have arisen after divergence of the three kingdoms. Thus, novel mechanisms of energy coupling are kingdom-specific and occur only in bacteria and archaea. Eukaryotes may have been the least inventive in designing new modes of energy coupling to transport although they cleverly adapted old mechanisms to unique physiological situations.

127

TRANSPORT PROTEIN CLASSIFICATION

9. PROPOSED INDEPENDENT EVOLUTION OF DIFFERENT CHANNEL AND CARRIER FAMILIES In the previous section, we observed that some transport modes and energycoupling mechanisms appear to occur ubiquitously in all of the three domains of living organisms while others are restricted to one domain or subdomain. We proposed that these facts most commonly reflect the time in evolutionary history in which they arose. Most of those that proved to be ubiquitous may have arisen early, while those that are restricted in their organismal distribution may have arisen later. We shall now attempt to extend this same argument to specific transporter families. Table 14 presents a list of representative transporter families that are believed to occur ubiquitously, and therefore are believed to have arisen early, before the divergence of eukarya from archaea and bacteria. The first three families listed consist of channel proteins; the next six include only secondary carriers; and the last two families have members that are all ATP-driven primary carriers. All of these families have members identified in both bacteria and eukaryotes, and all but two also have known archaeal members. The large major intrinsic protein (MIP) family consists of proteins that transport water and small neutral molecules such as glycerol, urea and ammonia (Table 6; Park and Saier, 1996). The even larger voltage-sensitive ion channel (VIC) family includes members that transport K', Na' or Ca2+

Table 14 Representative ubiquitous transporter families with variable substrate ranges. No. of sequenced members

No. of substrate classes

Family

TC #

No. of Kingdoms

MIP VIC CIC

1.1 I .5 1.10

2 3 3

> 100 > 100 30

2 3 1

sss

MFS

2.1 2.21

3 2

> 300 20

9 6

FNP DASS

2.44 2.47

3 3

10 20

2 3

CaCA Amt

2.19 2.49

3 3

50 20

1 I

A BC ARS

3.1 3.4

3 3

> 300

13 I

20

128

MILTON H. SAIER, JR

with a fairly high degree of cation specificity (Hille, 1992). Finally, characterized members of the smaller but still ubiquitous chloride channel (CIC) family are apparently highly specific for a single anionic species, chloride (Huang et al., 1994). With respect to carriers, both the MFS (Tables 7 and 9) and the solute: sodium symporter family (Table 15) transport a variety of different compounds. However, each permease transports its substrate(s) with a high degree of specificity. Interestingly, the MFS is an exceptionally large family while the SSS is a much smaller family. These two families can be distinguished functionally in that MFS permeases catalyze uniport, symport and/ or antiport and exhibit either inwardly directed or outwardly directed polarity, while the SSS permeases apparently catalyze only Na' symport with inwardly directed polarity. Thus, while the MFS and SSS transporters are both promiscuous with respect to substrate specificity, only the MFS is promiscuous with respect to cation coupling and polarity. If these two

Tabk 15 The So1ute:Sodium Symporter (SSS family (TC #2.21).

Members of the SSS family catalyze solute:Na+ symport (Reizer e/ a/.,1994). The solutes transported may be sugars, amino acids, nucleosides, vitamins, anions or inositols, depending on the system. Members of the SSS family have been identified in bacteria and in animals, and all catalyze solute uptake. They vary in size from about 400 residues to about 700 residues and possess 12-14 putative transmembrane helical spanners (Sarker el ul., 1997). The generalized transport reaction catalyzed by the members of this family is: solute (out)

+ Na'(out)

TC#

Name

2.21.1.1

Pantothenate:Na+ symporter Pro1ine:Na' symporter G1ucose:Na' symporter

2.21.2.1 2.21.3.1

-+ solute (in) + Na+(in).

Source

Example

Bacteria

PanF of E. coli (spP16256)

Bacteria Animals

PutP of E. coli (spPO7 I 17) SGLT of Homo sapiens (spP13866) SglS of Vibrio purahaemolyticus (gbD78137) SNST of Oryctologus cuniculus (spP26430) SMIT of Cunis~furniluris (gbM85068) Na'I- symporter of Homo sapiens (gpU66088)

2.21.3.2

Glucose or galactose:Na+ symporter

Bacteria

2.21.3.3

Nuc1eoside:Na' symporter

Animals

2.2 1.4.1

Myoinosito1:Na' symporter Animals

2.21.5.1

Sodium iodide symporter

Animals

TRANSPORT PROTEIN CLASSIFICATION

129

families are both ancient, the functional diversity of the MFS may explain why it is so much larger than the SSS. Other presumably ancient families include members, all of which are highly specific for one or a few substrates. Thus, the formatenitrite porter (FNP) family members appear to transport only these two anionic species while the divalent anion:sodium symporter (DASS) family members are capable of transporting a variety of organic anions (e.g. succinate, fumarate, oxaloacetate, etc.) as well as inorganic anions (phosphate and sulfate), each with fairly high specificity (Saier et al., 1998). Moreover, proteins of the fairly large Ca2+:cation antiporter (CaCA) family catalyze transport only of Ca2+in exchange for H' or Na', while the ammonium transporter (Amt) family only transports one species, ammonium (Saier et af., 1998). Finally, in comparing ABC superfamily transporters (Table 11) with the arsenical (Ars) family transporters (Table 2), the former transport virtually every solute of biological importance with either inwardly and outwardly directed polarity, but characterized Ars family permeases only catalyze extrusion of arsenite and antimonite (Silver et al., 1993). In general, then, we conclude that the largest families are ubiquitous and most diverse in function. However, some small ubiquitous families are functionally diverse while some larger families are functionally restricted. It is suggested that permease architecture in part determines the potential for functional diversification. Functional diversity undoubtedly provided a major driving force for family expansion during evolutionary history. Table 16 lists selected channel and carrier families identified only in bacteria or in eukaryotes. Among the channel protein families, the mechanosensitive channels with large conductance (MscL) comprise a small family of bacterial-specific ion channels that may function in response to osmotic pressure (Sukharev et al., 1994, 1996). The large and diverse holin functional superfamily apparently evolved for the export of autolysin proteins (Young and Blasi, 1995). In contrast, channel-forming colicins are used in bacterial warfare (Gouaux, 1997). Turning to channels identified only in animals, the epithelial Na' channels (ENaC) appear to be specific for Na' and other monovalent cations (Le and Saier, 1996), the ATP-gated cation channels (ACC) are apparently specific for similar molecular species (North, 1996) and the ryanodine-inositol 1,4,5-triphosphate receptor Ca2+ channels (RIR-CaC) are specific just for Ca2+ (Lee, 1996). Turning to the selected carrier families listed in Table 16, the currently sequenced resistance-nodulation-cell division (RND) family members are restricted to Gram-negative bacteria (Table 8), but they transport a wide variety of structurally unrelated drugs, heavy metals and lipo-oligosaccharides, all with outwardly directed polarity (Saier et al., 1994; Paulsen et al.,

130

MILTON H. SAIER, JR

Tuhle 16 Representative channel and carrier families found in only one Kingdom.

Family'

TC #

Kingdom

ENaC ACC RIR-CaC

I .2 I .4 1.9

Animals Animals Animals

MscL Holin Colicin

I .3 1.11 1.12

Bacteria Bacteria Bacteria

RND

2.6

SMR GntP

2.7 2.8

Gram-negative bacteria Bacteria Bacteria

AAAP MC Sit

2.18 2.29 2.32

Eukaryotes Eukaryotes Diatoms

No. of sequenced permeases

No. of substrate classes

24

1

10 10

I

6 > 20

I 3 1

20

3

II 12

I

30

2 9 I

> 30

> 100

6

1

1

*The full family designations together with the abbreviations for these Families are provided in the text.

1996b). The recently discovered small multidrug resistance (SMR) family includes two groups of permeases, one that catalyzes drug efflux and another of unknown specificity and function (Paulsen et al., 1996b,c). The functionally characterized members of the GntP family all take up gluconate (Peekhaus et a / . , 1997). The three eukaryotic-specific carrier families listed in Table I6 include: (i) the amino acid-auxin permeases (AAAP) of plants and fungi, all catalyzing the uptake of amino acids and their analogues (Bennett et d,, 1996); (ii) the mitochondria1 carrier family (MCF; Table 3), all found in eukaryotic organelles but diverse in their transport specificities; and (iii) the silicon transporters (Sit), all specific for silicate and so far found only in diatoms (Hildebrand, et al., 1997). The occurrence and characteristics of these kingdom-specific permeases lead us to suggest that many families of channels and carriers have arisen independently in specific eukaryotic kingdoms or in specific bacterial kingdoms. These families presumably arose late in the evolutionary process, and little inter-kingdom transfer has occurred. Our observations therefore lead us to suggest that horizontal transmission and fixation of genetic material across kingdom lines has been exceedingly rare.

TRANSPORT PROTEIN CLASSIFICATION

131

10. CONCLUSIONS AND PERSPECTIVES

In conclusion, we have seen that permease families arose repeatedly and independently, at different times in evolutionary history, following different routes. In spite of similar apparent topological features, we believe that several permease types must exhibit distinctive architectural features that confer differing capacities for functional diversification. Genome analyses have helped to reveal the numbers of permease families and the breadth of their functionalities. These analyses have led us to devise a novel permease classification system based on both functionality and phylogeny. We believe that a functional-phylogenetic basis for permease classification provides the most rational approach to protein classification in general. It also provides the maximal yield of information concerning the evolution, structures and functions of any class of proteins.

ACKNOWLEDGEMENT This Chapter is dedicated to the memory of my father, Milton H. Saier, Sr.

NOTE ADDED IN PROOF Since this manuscript was submitted for publication, the transport protein classification system described here has been expanded and updated. The continuously updated version can be found on our world wide web site [http://www-biology.ucsd.edu/-msaier/transport/titlepage.html]. Major changes in the primary categories of classification (see Table 1) are as follows: category 1 now includes 21 families of channel proteins while category 2 now includes 64 families of carriers. Category 3, ATPdriven active transporters, is now referred to as pyrophosphate bond hydrolysis-driven transporters and includes 11 families, one of which is an H+- translocating vacuolar pyrophosphatase family. Category 9 now includes 19 porin families. Category 99 (transporters of unknown classification) has been expanded to include 23 families. A new category, 100, includes putative transporter families. Additional information regarding this classification system has been prepared for publication and will appear as follows: Saier, M.H., Jr. (1998). Classification of Transmembrane Transport Systems in Living Organisms. In: Biomemhrane Transport (L. Vanwinkle, ed.). Academic Press, San Diego, in press. Descriptions of the individual families discussed here, par-

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MILTON H. SAIER, JR

ticularly the ABC and M F superfamilies, have been substantially expanded. This updated information is available on our web site. Milton H. Saier, Jr. msaier@ ucsd .edu Professor of Biology, Department of Biology UCSD La Jolla, CA 92093-0116 Phone: +619-534-4084; Fax: +619 534 7108

Ahringer, J. (1995) Embryonic tissue differentiation in Caenorhabditis ekgans requires dif-I, a gene homologous to mitochondrial solute carriers. EMBO J. 14, 2307-2316. Altschul, S.F., Gish, W., Miller, W., Myers, E.W. and Lipman D.J. (1990) Basic local alignment search tool. J. Mol. B i d . 215, 403410. Baldwin, S.A. ( I 993) Mammalian passive glucose transporters: members of an ubiquitous family of active and passive transport proteins. Biochim. Biophys. Acta 1154, 17-49, Bennett, M.J., Marchant, A.. Green, H.G., May, S.T., Ward, S.P., Millner, P.A., Walker, A.R., Schulz, B. and Feldmann, K.A. (1996) Arabidopsis AUXl gene: a permease-like regulator of root gravitropism. Science 273, 948-950. Blair, A,, Ngo, L., Park, J., Paulsen, I.T. and Saier, M.H., Jr (1996) Phylogenetic analyses of the homologous transmembrane channel-forming proteins of the FoFI-ATPasesof bacteria, chloroplasts and mitochondria. Microbiology 142, 17-32. Bruhn, D.F., Li, J., Silver, S., Roberto, F. and Rosen, B.P. (1996) The arsenical resistance operon of lncN plasmid R46. FEMS Microhiol. Lett. 139, 149-153. Broer, S., Ji, G., Broer. A. and Silver, S. (1993) Arsenic efflux governed by the arsenic resistance determinant of Siaphylococcus aurercs plasmid pI258. J . Bacteriol. 175, 348G3485. Brutovetsky, N. and Klingenberg, M. (1994) The reconstituted ADP/ATP carrier can mediate H+ transport by free fatty acids, which is further stimulated by mersalyl. J. Biol. Cheni. 269, 27 329-21 336. Brutovetsky, N. and Klingenberg M . (1996) Mitochondria1 ADP/ATP carrier can be reversibly converted into a large channel by Ca2+. Biochemistry 35, 8483-8488. Dierks, T., Salentin, A,, Heberger, C. and Krimer, R. (1990a) The mitochondrial aspartate/glutamate and ADP/ATP carrier switch from obligate counterexchange to unidirectional transport after modification by SH-reagents. Biochim. Biophys. Aciu 1028. 268-280. Dierks, T., Salentin. A.. Heberger, C. and Kramer, R. (1990b) Pore-like and carrier-like properties of the mitochondria1 aspartate/glutamate carrier after modification by SHreagents: evidence for a preformed channel as a structural requirement of cnrriermediated transport. Biochim. Biophys. A m 1028, 281-288. Dinh, D., Paulsen, I.T. and Saier. M.H., Jr (1994) A family of extracytoplasmic proteins that allow transport of large molecules across the outer membranes of Gram-negative bacteria. J. Bucteriol. 176, 3825-3831. Dixon, M. and Webb, E.C. (1979). Bizymes, 3rd ed. Academic Press, Inc., New Y o r k .

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Jezek, P., Orosz, D.E., Modriansky. M. and Garlid, K.D. (1994) Transport of anions and protons by the mitochondrial uncoupling protein and its regulation by nucleotides and fatty acids. J . Biol. Chem. 269, 26 184-26 190. Kaback, H.R. (1986) Active transport in Escherichia coli: passage to permease. Anriu. Rev. Biophys. Biophys. Chem. 15, 279-3 19. Kuan. G., Dassa, E., Saurin, W., Hofnung, M. and Saier, M.H., Jr (1995) Phylogenetic analyses of the ATP-binding constituents of bacterial extracytoplasmic receptordependent ABC-type nutrient uptake permeases. Rex Microhiol. 146, 271 278. Kuan, J. and Saier. M.H., Jr. (1993) The mitochondrial carrier family of transport proteins: structural, functional and evolutionary relationships. Crif. Rev. Biochm. Mol. B i d . 28, 209%233. Le, T. and Saier, M.H., Jr (1996) Phylogenetic characterization of the epithelial Na' channel (ENaC) family. Mol. Memhr. Biol. 13, 149-157. Lee, A.G. ( 1996) The ryanodine receptor. In: Biomembrune, Vol. 6 . Trunsmc,mhrane Receptors und Channels (A.G. Lee, ed.), pp. 291-326. JAI Press, Denver, CO. Liu, Q. and Dunlap, J.C. (1996) Isolation and analysis of the arg-13 gene of Nc~urosporfr crassa. Generics 143, 1 163- I 174. Loddenkotter. B., Kammerer, B., Fischer, K. and Flugge, U.-I. (1993) Expression of the functional mature chloroplast triose phosphate translocator in yeast internal membranes and purification of the histidine-tagged proteins by a single metal-affinity chromatography step. Proc. Null. Acad. Sci. USA. 90,2155-2159. Marger, M.D. and Saier, M.H., Jr (1993) A major superfamily of transmembrane Facilitators that catalyse uniport, symport and antiport. Trends Biocheni. Sci. 18, 13 20. Mitchell, P. ( I96 I ) Coupling of phosphorylation to electron and hydrogen transfer by ;I chemi-osmotic type of mechanism. Nnfure 191, 144-148. Mitchell, P. (1962) Metabolism, transport, and morphogenesis: which drives which? J . Cen. Microhiol. 29. 25- 37. Mitchell, P. and Moyle, J. (1959) Coupling of metabolism and transport by enzymic translocation of substrates through membranes. Proc. R . Sot. Edinh. [Phys.] 28, 19-27. Noji, H., Yasuda, R.. Yoshida, M. and Kinosito, K., Jr (1997) Direct observation of the rotation of FI-ATPase. Nuture 386, 299-302. North, R.A. (1996) Families of ion channels with two hydrophobic segments. Cur. Opin. Cell Biol. 8, 474-483. Pao, S.S., Paulsen. I.T. and Saier, M.H., Jr (1997) Major facilitator superfamily. Microhiol. Mol. Biol. Rev. 62, 1-32. Park, J.H. and Saier, M.H., Jr (1996) Phylogenetic characterization of the MIP family of transmembrane channel proteins. J . Mrmhr. Biol. 153, 171-1 80. Paulsen, I.T. and Skurray, R.A. (1993) Topology, structure and evolution of two families of proteins involved in antibiotic and antiseptic resistance in eukaryotes and prokaryotes - an analysis. Gene 124, 1-1 1. Paulsen, I.T., Brown, M.H., Littlejohn, T.G., Mitchell, B.A. and Skurray, R.A. (1996a). Multidrug resistance proteins QacA and QacB from Stuphylococcits uureus: membrane topology and identification of residues involved in substrate specificity. Proc. Nufl. Acrid. Sci. U S A 93, 3630-3635. Paulsen, I.T., Brown, M.H. and Skurray R.A. (l996b) Proton-dependent multidrug efflux systems. Microhiol. Rev. 60,575608. Paulsen, I.T., Skurray, R.A., Tam, R., Saier, M.H., Jr, Turner, R.J., Weiner, J.H., Goldberg, E.B. and Grinius, L.L. (19964 The SMR family: a novel family of niultidrug efflux proteins involved with the eftlux of lipophilic drugs. Mol. Microbiol. 19, 1167--1175.

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Paulsen, I.T., Beness. A.M. and Saier, Jr. (1997) Computer-based analyses of the protein constituents of transport systems catalysing export of complex carbohydrates in bacteria. Microbiology 143, 2685- 2699. Paulsen, 1. T.. Sliwinski M.K. and Saier, M.H.. Jr (1998) Microbial genome analyses: global comparisons of transport capabilities based on phylogenies, bioenergetics and substrate specificities. 1.Md. Biol. 277, 573-592. Peekhaus. N., Tong, S.. Reizer. J., Saier. M.H., Jr, Murray, E. and Conway, T. (1997) Characterization of a novel transporter family that includes multiple Esclic~richiacoli gluconate transporters and their homologues. FEMS Microhid. Leu. 147, 233-238. Quiocho. F.A. and Ledvina. P.S. (1996) Atomic structure and specificity of bacterial periplasmic receptors for active transport and chemotaxis: variation of common themes. Mol. Microbiol. 20, I7--25. Reizer. J.. Reizer, A . and Saier, M.H. Jr (1993) The MIP family of integral membrane channel proteins: sequence comparisons, evolutionary relationships, reconstructed pathway of evolution and proposed functional differentiation of the two repeated halves of the proteins. Crir. Rev. Biochem. Mol. Biol. 28, 235-257. Reizer. J.. Reizer, A. and Saier, M.H., Jr (1994) A functional superfamily of sodium/ solute symporters. Biochirn. Bi0phy.s. Aria 1197, 133-166. Saier, M.H., Jr (1994) Computer-aided analyses of transport protein sequences: gleaning evidence concerning function, structure, biogenesis, and evolution. Microhiol. Rev. 58. 71-93. Saier. M.H., Jr (1996) Phylogenetic approaches to the identification and characterization of protein families and superfamilies. Microbial Comp. Genomics 1, 129-1 50. Saier, M.J., Jr, Tam, R., Reizer, A. and Reizer, J. (1994) Two novel families of bacterial membrane proteins concerned with nodulation, cell division and transport. Mol. Microhiol. 11, 841-847. Saier, M.H., Jr., Eng, B.H., Fard, S., Garg, J.. Haggerty. D.A., Hutchinson, W.J., Jack, D.L., Lai, E.C., Liu, H.J.. Nesinew. D.P.. Omar, A.M., Pao, S.S., Paulsen, I.T., Quan J.A.. Sliwinski. M.. Tseng, T.-T., Wachi, S. and Young, G.B. (1998) Phylogenetic characterization of novel transport protein families revealed by genome analysis. Biochim. Biophys. Acra (in press). Sarker, R.I., Ogawa, W., Shimamoto, T., Shimamoto, T. and Tsuchiya, T. (1997) Primary structure and properties of the Na+/glucose symporter (SglS) of Vihrio parahueniol~vticus. J . Bacteriol. 179, 1805-1808. Saurin, W. and Dassa, E. (1994) Sequence relationships between integral inner membrane proteins of binding protein-dependent transport systems: evolution by recurrent gene duplications. Prof. Sci. 3, 325-344. Schuldiner, S., Shirvan, A. and Linial, M . (1995) Vesicular neurotransmitter transporters: from bacteria to humans. Physiol. Rev. 75, 369-392. Schulz, B.. Frommer, W.B., Flugge. U.-I., Hummel, S., Fischer K . and Willmitzer L. (1993) Expression of the triose phosphate translocator gene from potato is light dependent and restricted to green tissues. Mol. Gen. Gene,. 238, 357-361. Schwarz, M . . Gross, A,, Steinkamp, T., Flugge, U.-I. and Wagner R. (1994) Ion channel properties of the reconstituted chloroplast triose phosphate/phosphate translocator. J . Biol. Chern. 269, 29481-29489. Silver. S., Ji, G.. Broer, S., Dey, S.. Dou D. and Rosen B.P. (1993) Orphan enzyme or patriarch of a new tribe: the arsenic resistance ATPase of bacterial plasmids. Mol. Microhiol. 8, 637-642. Sukharev, S.I., Blount, P.. Martinac. B., Blattner, F.R. and Kung, C . (1994) A largeconductance mechanosensitive channel in E. coli encoded by mscL alone. Narure 368, 265- 268.

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Sukharev, S.I., Blount, P., Martinac, B., Guy. H.R. and Kung, C. (1996) MscL: A Mechanosensitive Channel in Escherichia coli. In: Organellar Ion Channels and Transporters, pp. 133-141. The Rockefeller University Press, New York. Sullivan, T.D., Strelow, L.I., Illingworth, C.A., Phillips, R.L. and Nelson, O.E., Jr (1991) Analysis of maize brittle- 1 alleles and a defective suppressor-mutator-induced mutable allele. Plant Cell 3, 1337- 1348. Tam, R. and Saier, M.H., Jr (1993) Structural, functional, and evolutionary relationships among extracellular solute-binding receptors of bacteria. Microbiol. Rev. 57, 320-346. Tzagoloff, A,, Jang, J., Glerum, D.M. and Wu, M. (1996) FLXI codes for a carrier protein involved in maintaining a proper balance of flavin nucleotides in yeast mitochondria. J. Biol. Chem. 271, 7392-7397. Varela, M.F. and Wilson, T.H. (1996) Molecular biology of the lactose carrier of Escherichia coli. Biochim. Biophys. Acta. 1276, 21-34. Wallmeier, H., Weber, A,, Gross, A,, and Fliigge, U.-I. (1992). Insights into the Structure of the Chloroplast Phosphate Translocator Protein. In Transporf and Receptor: Proteins of Plant Membranes (D.T. Cooke and D.T. Clarkson, eds), pp. 77-89. Plenum Press, New York. Young, R. and Blasi, U. (1995) Holins: form and function in bacteriophage lysis. FEMS Microbiol. Rev. 17, 191-205.

The Physiology and Metabolism of the Human Gastric Pathogen Helicobacter py/ori David J. Kelly Department of Molecular Biology and Biotechnology. University of Shefjeld. Western Bank, Shefield, SlO 2TN, U K

ABSTRACT Helicobucter pylori is a spiral Gram-negative microaerophilic bacterium that causes one of the most common infections in humans; approximately 3&50% of individuals in Western Europe are infected and the figure is nearly 100% in the developing world. It is recognized as the major aetiological factor in chronic active type B gastritis, and gastric and duodenal ulceration and as a risk factor for gastric cancer. H . pylori normally inhabits the mucus-lined surface of the antrum of the human stomach where it induces a mild inflammation, but its presence is otherwise usually asymptomatic. A variety of virulence factors appear to play a role in pathogenesis. These include the vacuolating cytotoxin VacA, cytotoxin-associated proteins, urease and motility. All are under intense study in an attempt to understand how the bacterium colonizes and persists in the gastric mucosa, and how H . pylori infections lead to the disease state. Although an explosion of research on H . pylori has occured within the past 15 years, most efforts have been directed at aspects of the bacterium and disease process which are of direct clinical relevance. Consequently, our knowledge of many aspects of the physiology and metabolism of H . pylori is relatively poor. This should change rapidly now that the complete genome sequence of a pathogenic strain has been determined. This review focuses attention on these more fundamental areas of Helicobucter biology. Analysis of the genome sequence and some detailed metabolic studies have ADVANCES IN MICRORIAL PHYSIOLOGY VOL 40 ISBN 0- 12-027740-9

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revealed solute transport systems, an incomplete citric acid cycle and several incomplete biosynthetic pathways, which largely explain the complex nutritional requirements of H . pylori. The microaerophilic nature of the bacterium is of particular interest and may be due in part to the involvement of oxygen-sensitive enzymes in central metabolic pathways. However, the biochemical basis for the requirement for C 0 2 has not been completely explained and a major surprise is the apparent lack of anaplerotic carboxylation enzymes. Although genes for glycolytic enzymes are present, physiological studies indicate that the Entner-Doudoroff and pentose phosphate pathways are more active. The respiratory chain is remarkably simple, apparently with a single terminal oxidase and fumarate reductase as the only reductase for anaerobic respiration. NADPH appears to be the preferred electron donor in vivo, rather than NADH as in most other bacteria. H . pylori is not an acidophile, and must possess mechanisms to survive stomach acid. Many studies have been caried out on the role of the urease in acid tolerance but mechanisms to maintain the protonmotive force at low external pH values may also be important, although poorly understood at present. In terms of the regulation of gene expression, there are few regulatory and DNA binding proteins in H . pylori, especially the two-component ‘sensor-regulator’ systems, which indicates a minimal degree of environmentally responsive gene expression. 1. Introduction.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 139 140 2. Helicobecter pylori as a gastric pathogen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 140 2.1. Epidemiology and transmission of H. pylori . . . . . . . . . . . . . . . . . . . . . . . 2.2. Virulence factors and the cag pathogenicity island . . . . . . . . . . . . . . . . . . 142 2.3. H. pylori-associated disease. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 142 144 3. Characteristics of Helicobacter pylori . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 144 3.1. Taxonomy and evolutionary relationships. ........................ 144 3.2. Cellular features and growth requirements . . . . . . . . . . . . . . . . . . . . . . . . 3.3. The spiral to coccoid cell transition. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 145 147 3.4. Motility and chemotaxis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4. Solute transport, ion movements and acid tolerance in H. pylori. . . . . . . . . . . 149 4.1. Major types of transport systems present. . . . . . . . . . . . . . . . . . . . . . . . . 149 149 4.2. Mechanisms of iron acquisition.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3. Ion homeostasis and its relationship to acid tolerance . . . . . . . . . . . . . . . 151 5. The microaerophilic nature of H. pylori. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 152 5.1. Physiology of microaerophilic growth . . . . . . . . . . 5.2. Mechanisms to combat oxidative stress in H. pylo 6. Current knowledge of H. pyloricarbon metabolism and substrate utilization . . 155 6.1. Early studies of metabolism . . . . . . . . . . . . . . . . . . . . . . . . 155 6.2. Glucose metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 156 159 6.3. Pyruvate metabolism.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

PHYSIOLOGY AND METABOLISM OF HELlCOBACTER PYLORl 6.4. The importance and physiological role of POR and OOR in H. pylori provision of NADPH and an explanation for the microaerophilic growth phenotype? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.5. Alcohol dehydrogenase. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.6. Fumarate metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.7. The nature of the citric acid cycle in H. pylori. . . . . . . . . . . . . . . . . . 6.8. The C02 requirement of H. pylori . . . . . ................... 6.9. Anabolic pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7. The respiratory chain of H. pylon'. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.1. Substrate oxidation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.2. Composition of the respiratory chain. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.3. The terminal oxidasek). . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.4. The genome sequence indicates a simple respiratory chain organization . 8. Nitrogen metabolism in H. pylori. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.1. Assimilation and management of nitrogen . . . . . . . . . . . . . . . . . . . . . . . . .......................... ..... 8.2. The urease of H. pylori 9. Concluding remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Note added in proof. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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169 169 171 174 175 176 176 176 179 180 180 180

1. INTRODUCTION

Until the early 1980s, gastrointestinal ulceration was largely attributed to elevated levels of stomach acid and was treated with powerful antisecretory drugs, particularly the H2-receptor antagonists. Through development, these drugs became more effective at easing the symptoms of disease, but they did little to prevent the recurrence of ulcers. Martin et al. (1981) were surprised at their discovery that duodenal ulcers remained healed for considerably longer after treatment with a bismuth salt than after H2 antagonist treatment. A reason for this became clear when, in the same year, an Australian pathologist and doctor began to connect the pathologist's earlier identification of microorganisms in the biopsies of patients with active chronic gastritis, to the occurrence of this disease (Marshall, 1988). These biopsies contained large numbers of curved and spiral bacteria which were closely associated with the surface of the gastric epithelium, both within and between the gastric pits (Warren, 1983). By light microscopy, the organisms resembled Campylobacter ,jejuni and they proved difficult to culture from antral biopsy specimens. However, after a prolonged incubation period, the bacteria grew in a microaerophilic environment at 37°C on moist chocolate agar (Marshall, 1983). As this bacterium appeared to have several similar features to the genus Campylobacter, it was originally named Campylobacter pyloridis (Marshall et al., 1984). The specific epithet was grammatically incorrect and the name was changed to C. pylori (Marshall and Goodwin, 1987). Further studies revealed that certain features of C. pylori were very

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different from those of all other campylobacters; therefore a new genus was established and C . pylori was transferred to it. This new genus was named Helicobacter and the bacterium became Helicobacter pylori (Goodwin et a/., 1989). The association of H. pylori with gastritis and gastrointestinal ulceration has revolutionized the treatment of such disease; it is now possible to completely heal gastric and duodenal ulcers, instead of merely suppressing the symptoms. A large number of investigations have shown that the complete eradication of the bacterium from the stomach can heal ulcers and prevent recurrence of disease (e.g. Hosking et al., 1994; Sung et al., 1995). It should be appreciated that the scale of human infection with this bacterium is enormous, even though most individuals are essentially asymptomatic; up to 50% of the population of the Western world and up to 90% or more of people in the developing world may be infected, making H. pylori one of the most common infections in humans. Because of this, and the world-wide occurrence of gastrointestinal ulceration, there has been an explosion of research into H. pylori, particularly within the past decade. The reader is referred to recent reviews in Calam (1995) and the series edited by Malfertheiner et a/. (1 995,1996) for access to the general and specialized literature on the diagnosis, epidemiology, transmission, pathology, pathogenesis and eradication of H. pylori. It is clear that the vast majority of publications concerning H . pylori are not concerned with fundamental aspects of the organism’s biology. The aim of this review is therefore to focus attention on aspects of the microbial physiology of H . pylori, with an emphasis on those metabolic processes which constitute the core of central metabolism. Not all areas can be covered, however, but the selection of topics hopefully reflects the areas where the most significant work has been carried out to date. A major advance in the understanding of the biology of H. pylori has occurred recently, with the publication of the complete genome sequence of a pathogenic strain (Tomb et al., 1997). Throughout this review, reference will be made to this important piece of work and attention drawn to those features which are supported by or which conflict with experimental evidence.

2. Helicobacter py/ori AS A GASTRIC PATHOGEN 2.1. Epidemiology and Transmission of H. py/ori

A feature of H. pylori-associated disease is that infection induces a signifcant host immune response and antibodies are present in the serum which are directed against the bacterium. Many epidemiological studies have

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involved the use of serum antibody tests for H. pylori. These have proved to be a reliable indirect method of detecting the presence of H. pylori infection, as endoscopy to collect samples for histology and culture is a cumbersome method of obtaining data to identify world-wide epidemiological trends. It is well established that the seroprevalance of H. pylori infection increases with age (Veldhuyzen van Zanten et al., 1994; Malaty et al., 1996). It has also been concluded that the pattern of clinical H. pylori disease found within a population is determined by the age of acquisition of the infection. Infection in childhood leads to a predominance of gastric ulcer and gastric cancer, whereas infection as an adult generally leads to duodenal ulcer and gastric cancer is rarer. There is little relationship between seropositivity for anti-H. pylori antibodies and alcohol intake, smoking or gender. Evidence indicates that infection rates are higher in developing countries and are inversely related to socioeconomic class (Graham ef ul., 1992; Malaty el ul., 1996). In developing countries, where poverty prevails, there is overcrowding and poor childhood health. Many individuals of these populations become infected at an early age and atrophic gastritis is common in young adults, yet there is a low incidence of duodenal ulceration in these populations (Graham, 1991). Using Japan as an example, the change in epidemiology of H . pylori as a result of Westernization was studied. By testing serum banks for anti-H. pylori antibodies, it was concluded that in 1940 infection was most likely to occur before adulthood. A similar investigation in 1990, i.e. after an improvement in sanitation and healthcare, indicated that individuals were more likely to become infected between the ages of 20 and 40 (Graham et al., 1992). The precise mechanism of transmission of H. pylori is unknown, but any method that introduces the organism into the stomach of a susceptible person may lead to infection. The oral-oral route is favoured, and although the human stomach is relatively resistant to oral inoculation, a reduction in acid secretion may make it easier for the organism to become established (Marshall et al., 1985). H . pylori has also been successfully cultured from human faeces, which raises the possibility of a faecal-oral route of transmission (Kelly et a]., 1994). Although the natural niche for H. pylori is the human stomach, for widespread infection the organism may need to survive in the external environment. Evidence has been obtained from studies in Peru suggesting that H . pylori is waterborne (Klein ef al., 1991), but there is no conclusive evidence that H. pylori is a widespread environmental contaminant. The bacterium can survive for several days in water and chilled foods, but it has never been isolated from these sources (West et al., 1992). Domestic cats (Handt ef al., 1995) have been reported to carry viable H. pylori and may act as potential animal reservoirs for the pathogen.

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2.2. Virulence Factors and the ceg Pathogenicity Island A number of general and specific virulence factors have been implicated in the pathogenicity of different strains of H. pylori. These include motility, ureasc production, lipopolysaccharide and adhesins. One of the most important specific virulence factors produced is a 94 kDa vacuolating cytotoxin protein encoded by the vucA gene (Cover and Blaser, 1992), which induces the formation of acidic vacuoles in gastric epithelial cells. Another ’cytotoxin-associated’ protein, encoded by the cagA gene, is consistently produced by virulent isolates (Tummuru et d.,1993). The cugA gene is part of a pathogenicity island in H. pylori; this is a large contiguous set of genes that encode proteins with poorly defined functions, but at least some of which appear to be necessary to elicit production of interleukin-8 (IL-8) a/., 1995; Censini et ul., 1996). by gastric epithelial cells (Tummuru Some of the gene products have homologies with proteins in other bacteria known to be involved in the transfer of DNA between cells (e.g. VirBDG, TraBO) and toxin export (Ptl).

2.3. H. pylori-associated Disease

2.3.1. Gastrilis Infection with H . p j h r i may result in an acute (often poorly characterized) or chronic gastritis. It is now widely accepted that chronic diffuse antral gastritis is a result of H . pylori infection. This is found predominantly in the antrum, is often associated with hypochlorhydria and peptic ulceration, does not involve an autoimmune response and is classified as ‘type B’ gastritis (Wyatt and Dixon, 1988). It is characterized by epithelial degeneration, neutrophil infiltration, lymphocyte and plasma cell influx, glandular atrophy and intestinal metaplasia. Prior to the association with H. py/ori, the aetiology of this category of antral inflammation was unknown, although irritants such as bile reflux, therapeutic drugs, hot drinks and salted or spicy foods were all suggested. However, not all investigators are convinced that all nonautoimmune gastritis is H . pylori-associated. This is because there are varying patterns of inflammation accompanying different disease states. Many infected individuals exhibit symptoms of gastritis which may lead to gastric ulcer and may even develop into carcinoma, while the majority are asymptomatic (Dixon, 1994). The strongest evidence for H . pylori being a causal agent of acute gastritis was produced by non-infected, healthy human volunteers ingesting a culture of the bacterium and developing symptomatic gastritis several days later (Marshall et d.,1985). Subsequent eradication of the bacteria resulted in remission of the symptoms of gastritis.

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Duodenal and Gastric Ulceration

The initial problem with the proposal that H . pylori was associated with duodenal ulceration was that the organism had only been shown to colonize gastric-type epithelium (Moss and Calam, 1992). However, patches of gastric-type epithelium (metaplasia) can be present in the duodenal bulb (Wyatt et al., 1987), and colonization of this region can lead to duodenitis which probably predisposes to duodenal ulceration. Gastric metaplasia was only present in 5-30% of people who were H . pylori-positive but had no symptoms of duodenal ulceration (Wyatt et al., 1987; Fitzgibbons r f al., 1988). I t is thought that H. pylori, once established within the duodenum, may cause ulcers by inducing inflammation and by releasing an ulcerogenic toxin (VacA; see above). It has been suggested that H . pjllori strains which are specifically associated with duodenal ulceration are genetically different to strains associated with other gastric diseases (Moss and Calam, 1992). Further evidence for specific duodenal ulcer causing H . p-vlori has been obtained. Antibodies to the vacuolating cytotoxin were present in 100% of duodenal ulcer patients with H . pylori compared with 61 YOof patients with H . pylori but no ulcer (Cover et al., 1990). Another important piece of evidence which links H . pylori to duodenal ulcers is that eradication of infection prevents ulcer relapse (O’Connor, 1994). Therefore, to prevent recurrence, a therapy should involve both the treatment of the ulcer and the removal of the infection. A successful eradication therapy may also accelerate duodenal ulcer healing. Most duodenal ulcer patients have H . pylori infection, whereas about 30% of gastric ulcer patients are H . pylori-negative (O’Connor, 1994). This suggests that in addition to H . pylori, other causative agents are involved in gastric ulceration. Non-steroidal anti-inflammatory drugs (NSAIDs) are known to be exogenous causes of ulcer disease. These promote mucosal inflammation and the development of lesions and haemorrhages which lead to ulcer formation in uninfected people (Sobala ef al., 1990). In one study, about 60% of cases of gastric ulcers in NSAID users were H . pylorinegative (Taha et al., 1992).

2.3.3. Gastric Cancer As H . pylori infection was determined as a causative agent of chronic gastritis, it was proposed that when the gastritis became atrophic and mucosal alterations occurred, the bacterial infection could be involved in the development of gastric cancer (Forman et al., 1991). This hypothesis was supported by the discovery that population groups at high risk of gastric cancer had a high prevalence of H . pylori infection at an early age (Fox el al., 1989).

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In Finland, about 70% of gastric cancer cases are associated with H . pyloripositive chronic gastritis (Sipponen, 1994). H . pylori is not known to produce carcinogens that directly damage the DNA of epithelial cells. Therefore, H . pylori infections must possess features which have an indirect role in carcinogenesis. It has been proposed that ammonia, or ammonium-containing substances, produced as a result of H . pylori urease activity, may act as cancer promoters by enhancing rates of cell division (Tsujii et al., 1992). Bacterial phospholipases may damage the phospholipid bilayer of the epithelial cell membrane and degrade the protective mucus layer (Goggin et al., 1991; Marshall, 1991). H . pylori strains that produce the vacuolating cytotoxin VacA, have been reported in populations at high risk of developing gastric cancer (Fox et al., 1992). Finally, cytokines and reactive oxygen metabolites, which are produced as part of the host immune response to H . pylori-induced gastritis, may stimulate proliferation of epithelial cells and cause damage to the DNA of these cells (O'Connor, 1992).

3. CHARACTERISTICS OF Helicobecter py/ori

3.1. Taxonomy and Evolutionary Relationships

After the initial isolation of H . pylori (Marshall, 1983; Warren, 1983), the bacterium was classified as a new species in the genus Campylohacter. However, initial 16s rRNA sequence data and additional taxonomic features, such as the presence of sheathed flagella and a distinctive fatty acid and SDS-PAGE protein profile, led to the establishment of the new genus Helicobacter (Goodwin et al., 1989). There are now over 13 members of the genus Helicohucter and several of these species have known pathogenesis. More recent analysis of 16s rRNA sequences has shown that the genus Helicobacter has relationships to Campylobacter, Wolinellu and Arcohacter. H . pylori was most closely related to H . acinonyx and H . fX.s (Stanley et al.. 1993). Thus, H . pylori is a member of the &subdivision of the Proteobacteria, which also includes genera such as Desulfovihrio and My.xoc0ccus. 3.2. Cellular Features and Growth Requirements

Helicobacter pylori is a non-spore-forming Gram-negative, spiral or curved rod (0.54.9 pm wide by 2-4 pm long). The cells are normally motile and typically possess five to six sheathed, polar flagella. Flagella are about 30 nm in diameter with a filament of 12-15 nm. They have terminal bulbs, but no definite function has been determined for these (Jones et a f . , 1985). The

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microaerophilic nature of H . pylori necessitates that it is cultured in an atmosphere containing 5-10% (v/v) oxygen and 5-10% (v/v) carbon dioxide. H . pylori is a fastidious organism and it is routinely cultured in complex media with additional growth supplements. For the initial culture of H . pylori, blood derivatives were added as supplement, but growth can also be achieved with charcoal, starch, bovine serum albumin (BSA), catalase and b-cyclodextrin (Buck and Smith, 1987; Hazell et al., 1989; Olivieri et al., 1993). The function of these supplements may be to adsorb potentially toxic long-chain fatty acids. The amino acid requirements of H . pylori were determined through the development of defined media. The details of two such media have been published and the approaches of the investigators were quite different. Reynolds and Penn (1994) added components to a defined tissue culture medium and initial studies identified that omission of lipoic acid, FeS04, BSA and non-essential amino acids was detrimental to growth of H . pylori. Glucose was also shown to be an important component of this liquid medium to support growth which was comparable with growth of the bacterium in complex media. BSA was added as a growth supplement. Nedenskov (1994) used solid media in his studies, with charcoal as a growth supplement. This medium was similar to that of Reynolds and Penn (1994) in that it was essentially a buffered amino acid mixture, but the growth of H . pylori showed no requirement for glucose, indicating the utilization of amino acids as carbon and energy sources. This has since been confirmed in metabolic studies (Mendz and Hazell, 1995; Stark et al. 1997). Nedenskov (1994) briefly described the use of liquid defined media; surprisingly, no long-chain fatty acid-adsorbing growth supplement was necessary for growth. The use of both these defined media identified similar amino acids that were necessary for growth. All tested strains in the two separate investigations required arginine, histidine, isoleucine, leucine, methionine, phenylalanine and valine. Reynolds and Penn (1994) concluded that, as these seven essential amino acids are generally produced at the end of biosynthetic pathways, mutations in or deletions of genes which encode enzymes of these pathways may have occurred in the H . pylori genome. This has been largely confirmed by the results of genome sequencing, which indicate the apparent absence of the genes encoding some of the key enzymes in these pathways (Tomb et al., 1997).

3.3. The Spiral to Coccoid Cell Transition

After several days of growth in vitro, H . pylori batch cultures undergo a morphological change from spiral-bacillary to coccoid forms (Catrenich and Makin, 1991). This morphological conversion is characterized by the loss of

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culturability and an increased pH of broth cultures. Brucella broth was used in this investigation, but the use of a defined medium for H . pylori growth reduced the eventual conversion to the coccoid form from nearly 100% to 50% (Reynolds and Penn, 1994). This indicated that the change to the coccoid form may be regulated by the organism’s nutritional requirements. It has been proposed that even though the coccoid form of H . pjdori is not culturable, it may still be viable (Bode et al., 1993; Shahamat c’t d., 1993). Therefore, this form may represent a starvation-survival mechanism, by which the bacterium can adapt temporarily to a hostile environment, and may thus function in transmission or relapse of infection following what was initially thought to be a successful eradication (Bode et al., 1993). However, there is no evidence that the coccoid form of H . pylori can revert back to the spiral form, and there is considerable controversy surrounding the whole phenomenon of ‘viable but non-culturable’ (VBNC) cells, which have been reported for a number of bacterial genera (Oliver, 1993). This arises over the criteria used to define ‘viability’. Shahamat et ul. (1993) used autoradiography after incubation of starved cells of H . pylori with [3H]thymidine to show that thymidine uptake continued to occur long after the cells became nonculturable (i.e. unable to form colonies on plates). In many studies, the demonstration of a variety of such metabolic activities in VBNC cells has been taken as one measure of their potential viability, but only in a very few cases has the recovery of normal vegetative cells from VBNC cells been achieved, thus demonstrating their eventual culturability. Gribbon and Barer (1995) studied growing and non-culturable cells of H . pjdori using substrate-enhanced tetrazolium reduction combined with digital image processing to observe the retention of oxidative metabolism. Cells starved for carbon or nitrogen in air lost their tetrazolium reducing capacity within 24 h when stored at 37°C but virtually all of the cells starved at 4°C retained succinate-, 2-oxoglutarate- or aspartate-dependent activity for up to 250 days. No attempts were made to convert the starved (largely coccoid) cells back to a spiral, culturable form. It has thus not been established that the retention of metabolic activities in the coccoid cells is actually a measure of viability. In the absence of clear evidence that coccoid cells can ever undergo cell division or return to a spiral form which is able to divide, the role of these cells in the environment and in the transmission of disease remains doubtful. Indeed, in a recent detailed study, Kusters et al. (1997) showed that inhibition of protein or RNA synthesis did not affect conversion to the coccoid form and that coccoid cells did not exhibit a measurable membrane potential. This strongly indicates that these cells are dead, and are the end result of a passive conversion from the spiral form. Despite this, it is clear from the genome sequence (Tomb et al., 1997) that H . pylori possesses homologues of proteins which may be important in carbon starvation

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conditions and in stationary phase survival, and this will be an important area for future research.

3.4. Motility and Chemotaxis

The motility of H . pylori is one of its most characteristic and prominent features. The spiral shape of the cells may give them an advantage in viscous environments and early experiments with solutions of methyl cellulose showed that H . pylori is capable of efficiently migrating in solutions of high viscosity (Hazell et al., 1986). Examination of human biopsy specimens suggested that the spiral morphology is most noticeable when the bacteria are motile and is less apparent once they have adhered to the gastric epithelium (Hazell et al., 1986). The importance of motility in H . pylori colonization was first confirmed when a gnotobiotic piglet model was only poorly colonized by a non-motile variant of a clinical isolate in comparison with the parent strain which colonized well and induced chronic gastritis (Eaton et al., 1992). The major H . pylori flagellin gene WaA) has been sequenced, and encodes a protein with a predicted molecular mass of 53.2 kDa. The amino acid sequence had good homology in the N-terminal and C-terminal regions to other bacterial flagellins, including species of the genus Campylohacter (Leying et a/., 1992). A second flagellin gene, JlaB has been identified which is unlinked to,flaA and encodes a protein only distantly related to it (Leying et al., 1992; Suerbaum et al., 1993). FlaB is located at the hookproximal end of the flagella (Kostrzynska et al., 1991). Both of these genes are necessary for full motility and mutation of either one severely reduced the colonization efficiency of H . pylori in gnotobiotic piglets (Eaton rt ul., 1996). A j a A B double mutant completely failed to colonize this animal model. However, in addition to motility per se, the chemotactic responses of pathogenic bacteria are likely to be very important in contributing to virulence, but although this is now receiving increasing attention in some important pathogens (e.g. Campylohucter jejuni; Yao et a/., 1997). there is a real lack of molecular data in this area. The best understood chemotaxis systems remain those of the enteric bacteria, where it is known that chemoeffectors (attractants/repellents) interact with membrane-bound methyl-accepting chemotaxis proteins (MCPs). A signalling complex is formed between the MCPs, a linker protein (Chew) and CheA, a histidine protein-kinase and a member of the ‘two-component’ signal transduction protein family. Autophosphorylated CheA then transfers its phosphate group to CheY, which interacts with the ‘switch protein’ FliM in the flagellar motor and alters the direction of flagellar rotation. CheA also phosphorylates CheB, which acts as a methylesterase and thus brings about adaptation to the

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stimulus by restoring the original (lower) level of MCP methylation catalysed by the methyltransferase, CheR. CheY has autophosphatase activity, but interaction with an additional protein, CheZ, enhances this considerably and effectively terminates the signal by competing with FliM for CheY-P. Although the enteric system has acted as the paradigm for chemotaxis in bacteria, there is now no doubt that the majority of other bacteria have more complex systems, usually with completely novel components or with multiple MCP, CheA and CheY proteins or domains. The pathways and mechanisms of signal transduction in these systems are likely to be quite distinct from the enteric model. Very little is currently known about the types of molecules important for chemotaxis in H . pylori; there is evidence for positive chemotaxis towards components of gastric mucin (Turner et al., 1997) which may have a role in directing the bacteria to the gastric mucosa. In addition, both urea and sodium bicarbonate reportedly have a positive chemotactic effect (Mizote et al., 1997). Jackson et al. (1995) first reported the identification and cloning of chemotaxis genes from H . pylori, encoding MCP, CheA and CheY homologues. The recently released genome sequence of H . pylori (Tomb et al., 1997) indicates that there are three MCP genes, with highest homology to tlpA/B/C of B. subtilis. There is also an additional gene which encodes another TlpC like MCP. It is not yet known if the MCPs are membranebound or cytoplasmic. All of these homologues have the conserved MCP signalling domain containing appropriately located glutamate residues which may be expected to be subject to carboxymethylation as part of the adaptation response to ligand binding. However, there is no evidence from the genome sequence that H . pylori contains CheR (methyltransferase) or CheB (methylesterase) homologues, or proteins similar to CheC/D of B. subtilis, which are also involved in methylation reactions. The basis for adaptation in the H . pylori MCPs is thus unclear. There are two additional unlinked chemotaxis gene clusters, one containing an isolated che Y gene, which appears to be part of a larger operon containing apparently chemotaxis-unrelated stress induced genes (Beier et al., 1997), and the other containing an operon encoding the following homologues: (i) a CheV protein (31% identity to B. subtilis CheV), which has an N-terminal Chew domain linked to a C-terminal CheY domain; (ii) a bi-functional protein (Pittman et al., 1997), with homology to the myxobacterial FrzE protein, containing both CheA and CheY domains; and (iii) a Chew homologue. There is no cheZ gene. Surprisingly, the genome sequence also reveals the presence of two additional cheV homologues (proteins 27% and 25% identical to B. subtilis CheV), unlinked to each other and to the other cheV gene. This combination of chemotaxis genes is novel and it is interesting to note the presence of no less than four Chew domains and five CheY domains, all on separate proteins. In B. subtilis, Chew and CheV are partially functionally

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redundant (Frederick and Helman, 1994), but in H . pylori these multiple domains could have very specific roles in what appears to be a complex system. How these proteins interact to transduce information to the flagellar motor will require substantial future biochemical investigations, but it is worth noting that there are very few sensor-regulator pairs of the ‘twocomponent’ families in H. pylori (Tomb et al., 1997) and, indeed, few regulatory proteins in general. The complexity of the chemotaxis signal transduction system suggests that chemotaxis is an important feature of H . pylori physiology.

4. SOLUTE TRANSPORT, ION MOVEMENTS AND ACID TOLERANCE IN H. py/ori 4.1. Major Types of Transport Systems Present

The genome of H . pylori is relatively small at 1.7 Mb, and the fastidious nature of the bacterium and its limited capacity for biosynthesis suggests that there will be a plethora of transport systems for the acquisition of essential amino acids, other nutrients and ions from the external environment. Some detailed work has been done on specific transport systems related to glucose and iron utilization and nickel transport for the urease enzyme, and these will be dealt with below. The genome sequence of strain 26695 reveals a large number of genes encoding transport functions (Tomb et al., 1997). Primary transport systems dependent on ATP hydrolysis for energy coupling are well represented, and include periplasmic binding protein-dependent systems for oligo- and dipeptides, amino acids, osmoprotectants, molybdate and ferric iron, as well as P-type ATPases like CopA, CopP and CadA which are involved in transport of essential ions like nickel and resistance to heavy metals. Ion-linked secondary transporters for the uptake of organic solutes include those for amino acids, carboxylic acids (lactate, 2-oxoglutarate, fumarate, malate) and glucose. A recently identified group of binding protein-dependent secondary transporters (TRAP transporters) widely distributed in bacteria (Forward et al., 1997) is, however, not present in H . pylori. Two aspects of solute transport will be considered in more detail; iron acquisition and the maintenance of a protonmotive force under acid conditions. 4.2. Mechanisms of Iron Acquisition

In common with all pathogenic bacteria, a major problem facing H . pylori in establishing itself in the gastric mucosa is obtaining sufficient iron for

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essential metabolic roles, such as the biosynthesis of cytochromes and other electron transport components. Thus, the ability of the bacterium to satisfy its iron requirement can be considered to be an important virulence determinant. Although there is a significant amount of iron present in mammalian body fluids, the amount of free iron available to invading pathogens is extremely small. Most iron is held inside cells as haem or in ferritin, and that which is extracellular in plasma or other body fluids is bound to the high-affinity iron-binding glycoproteins transferrin and lactoferrin. Pathogens like H . pylori which can successfully establish extracellular infections must possess high-affinity uptake systems which allow them to compete effectively with host iron-binding proteins for essential iron, or to acquire it from traces of haemoglobin or haem which may be liberated from cells. Relatively little is known regarding the sources of iron used by H. pylori in the gastric mucosa or the iron uptake mechanisms it employs it1 v i w . As for other bacteria, iron-limited growth of H . pylori leads to the induction of a number of outer-membrane proteins, some or all of which could serve as receptors for the uptake of iron complexes (Husson et a/., 1993; Worst c’t ul., 1995). Three such proteins (77, 50 and 48 kDa) were isolated by heminagarose affinity chromatography in the study of Worst et ul. (1995) and thus may be involved in the uptake of hemin as an iron source. The 77 kDa protein was expressed in vivo and appeared to be strongly immunogenic (Worst et a/., 1996). The study by Husson et al. (1993) concluded that N.pylori does not secrete siderophores and is unable to utilize the siderophores enterobactin and pyochelin. In contrast, another study suggests that H. pylori does produce siderophores (Illingworth et al., 1993). There are no obvious candidate siderophore-related genes in the genome sequence of strain 26695. There is strong evidence that H . pylori is able to use human lactoferrin as the sole iron source, but not human transferrin, ovotransferrin or bovine lactoferrin (Husson ct ul., 1993). These findings are significant in view of the presence of lactoferrin in human gastric tissues, and there is evidence that these levels are increased in patients infected with H . pylori (Nakao et al., 1997). Dhaenens el al. (1997), using an affinity chromatography method, identified a 70 kDa lactoferrin binding-protein (Lbp) from the outer membrane of H . pylori. This protein was only present when the bacteria were cultured under iron-depleted conditions and competitive binding experiments with lactoferrin and transferrins from different sources showed that the Lbp was highly specific for human lactoferrin. With the publication of the genome sequence of strain 26695, some interesting insights can be gained into the iron acquisition mechanisms which H . pylori possesses (Tomb et al., 1997). There are genes encoding a Fec system, homologous to that in E. coli, which could mediate the uptake of ferric iron,

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although the associated regulatory genes CfecR and fecl) are absent. An unusual feature is that in H . pylori, there are three copies of thefecA transport protein gene, none of which is linked to the fecDE genes encoding the membrane protein and ATPase subunits respectively, of this ABC-type transporter. There are also two copies of a gene encoding a putative periplasmic ferric iron-binding protein CfeuA), closely linked to each other but not to the other jec genes. It is also apparent that H . pylori possesses a ferrous iron uptake system (FeoB protein) which may be important in the assimilation of iron under microaerobic conditions. This system has been characterized by Velayudhan ef al. (l997), including the construction of a jeoB mutant. Interestingly, the f e o S gene is adjacent to one of the f2cA genes. There is, however, nofeoA gene, as is found in E. coli. The H . pylori genome also encodes three homologues offrpB, which may possibly encode haem or lactoferrin binding-proteins, consistent with the biochemical data described above. Another intriguing feature is the presence of no less than three sets of exhBD genes, one of which is linked to a single copy of tonB. The TonB protein is an important energy transduction protein for several iron-uptake systems, and ExbBD may act as assembly factors or chaperonins essential for TonB operation. In many bacteria, the expression of ironuptake systems under iron-limiting conditions is controlled by the global regulator, Fur, and this is also present in H . pylori (Bereswill et al., 1998). 4.3. Ion Homeostasis and its Relationship to Acid Tolerance

Despite being able to colonize the stomach, where the lumenal pH is around 1.0-2.0, H . pylori is not an acidiphilic bacterium. On entering the stomach, the organism must therefore rapidly gain access to the mucous layer overlying the gastric epithelium where the pH is near neutral. Much work has focused on the role of the active urease of H. pylori in neutralizing acid in the immediate microenvironment of the bacterium, but ammonia production in this way could not alter the bulk phase lumenal pH significantly and thus other mechanisms of acid tolerance must exist which allow. the bacterium to survive the transient initial pH decrease. Such mechanisms must ensure the maintenance of an internal pH near neutrality, which necessarily will involve the active exclusion of protons from the cytoplasm, implying the existence of a large ApH across the cytoplasmic membrane. Measurements have been made of the magnitude and composition of the protonmotive force in H . pylori at acidic and neutral external pH values (Matin et al. 1996; Meyer-Rosberg et a/., 1996). In the study of Matin ef al. (1996), ''C-labelled salicylate was concentrated by the cells more than 800-fold at an external pH of 3.0, corresponding to an internal pH of 6.2 and thus a ApH of 3.2 (-193 mV). Addition of a

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protonophore released the accumulated salicylate. The uptake of thiocynate (S14CN-) at this low external pH indicated a positive inside membrane potential, with a magnitude of + 81 mV. Thus the overall Ap at an external pH of 3.0 was -1 12 mV. At an external pH of 7.0, ApH was zero, and a membrane potential, inside negative, of -132 mV was measured by tetraphenylphosphonium (3H-TPPt) uptake. From these data, it appears that H . pylori is able to reverse the polarity of its membrane potential and thereby maintain a significant Ap at low pH, comparable with that measured in acidiphilic bacteria, but unlike E. coli where Ap = 0 at pHo 3.0 (Matin et al., 1996). In contrast, Meyer-Rosberg el al. (1996), using fluorescent dyes as indicators of Ap, found that although the membrane potential increased over the external pH range 4.0 to 7.0, in the absence of added urea the membrane potential collapsed to zero outside of these pH limits. The nature of the ion-transport mechanisms responsible for pH homeostasis in H . pylori is not yet clear. To generate a positive inside membrane potential either electrogenic anion efflux or active proton extrusion is required, and Tomb et a/. (1997) have concluded that there is little evidence for the former mechanism from the number and types of anion transporters present in the genome sequence, although they acknowledge that many candidate systems are at present unidentified. Active proton extrusion by P-type ATPases has been proposed (Melchers et al., 1996) but none of the systems characterized thus far has been shown to be proton-translocating. Whatever the mechanism(s) involved, it is clear that H . pylori is unable to grow at low external pH values, and so it may only be able to sustain the necessary ion gradients transiently.

5. THE MICROAEROPHILIC NATURE OF H. py/ori 5.1. Physiology of Microaerophilic Growth

The microaerophilic nature of H . pylori suggests that O2 is required for growth. However, O2 can present problems to most organisms owing to the toxic effects of the products of the stepwise one-electron reduction of 02: 0 2 -+

0; -+ HzO2 -+ HO' + H20

The superoxide radical (OF)and hydrogen peroxide (H202)are relatively poorly reactive in aqueous solution and can be eliminated by enzymatic activity. However, the interaction of these two molecules causes damage to living cells owing to the formation of the highly reactive hydroxyl radical (HO'), which can attack and destroy all known biomolecules (Halliwell and Gutteridge, 1986). In vivo, the formation of the hydroxyl radical is

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commonly catalysed by iron. The superoxide radical reduces Fe(II1) to Fe(II), and the reduced Fe(I1) can react with hydrogen peroxide in the Fenton reaction. The overall reaction is known as the Haber-Weiss reaction and this is one of the main ways in which oxidative stress can affect biological systems:

+ Fe3+ O2 + Fe2' Fez+ + H 2 0 2+ Hf --+ Fe3+ + HO' + H 2 0 0; + H 2 0 2+ HO' + OH- + O2 0,

Fenton reaction: Haber-Weiss reaction:

--+

5.2. Mechanisms to Combat Oxidative Stress in H. py/ori

Most aerobic, and some anaerobic, organisms possess major defence mechanisms against such oxidative stress. These defence mechanisms include such iron-dependent proteins as superoxide dismutase, catalase and ferritin and the presence and function of these proteins in H. pyfori has been investigated. Superoxide dismutases (SODs), are a family of three types of metalloenzymes containing manganese (MnSOD or SodA) or iron (FeSOD or SodB) or both copper and zinc (CuZnSOD or SodC) cofactors which catalyse the breakdown of superoxide anion to hydrogen peroxide and dioxygen (Fee, 1991):

The gastritis and peptic ulcer disease associated with chronic H. pyfori infection are characterized by the concentration of polymorphonuclear granulocytes (PMNs) in the infected area (Sobala et uf., 1991). PMNs have been shown to have a reduced microbicidal effect against H. pyfori (Andersen et a f . ,1993); however, activation of the oxidative burst does take place (Nielsen and Andersen, 1992). The oxidative burst is one of the major killing mechanisms of PMNs so the lack of killing of H. pyfori by PMNs suggests that the organism possesses a defence mechanism against superoxide, such as SOD. The gene encoding a H. pyfori FeSOD (so&) has been cloned and sequenced (Spiegelhalder et a f . , 1993; Pesci and Pickett, 1994). The enzyme consists of two identical subunits and has a native molecular mass of 50 000 Da. By analysis of primary structure and inhibition studies it was revealed that the H. pyfori enzyme was typical of prokaryotic iron-containing SODs. Sequence analysis determined that the H. pyfori SOD had greatest homology to that of facultative intracellular pathogens such as Listeriu ivanovii,

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Listeria monocytogenes and Legionella pneumophila, which may be evidence for the enzyme’s role in resistance to phagocytic attack. Immunolocalization studies of SOD using gold immunostaining techniques has suggested that the H . pylori SOD could be located on the cell surface (Spiegelhalder et al., 1993), although there is no evidence of a signal sequence in the protein. In addition to an FeSOD, H . pylori has been found to possess an active catalase (Hazell et al., 1991). This is an important enzyme in bacteria as it catalyses the breakdown of hydrogen peroxide to water and oxygen, thus protecting the bacterium from the damaging effects of hydrogen peroxide. The H . pylori catalase is a tetramer with a subunit molecular mass of 50 000 Da on SDS-PAGE. This soluble enzyme was found to be reversibly and non-competitively inhibited by sodium azide which provides evidence for the presence of an iron-porphyrin prosthetic group. The enzyme was heat stable and had a broad pH activity, similar to the catalases of Klehsiella pneumoniae and E. c d i . The H . pylori catalase activity was affected by the presence of blood, serum or erythrocytes in the growth medium. The greatest activity was detected when serum was added to the medium and the activity decreased as the relative concentration of erythrocytes in the medium increased (Hazell et al., 1991). This suggested that either a serum factor(s) stimulates catalase activity and/or the presence of haem-containing proteins released by erythrocytes has an inhibitory effect. H . pylori catalase activity may act alongside the SOD activity in protecting the bacterium against the in vivo microbicidal activity of PMNs which can release H 2 0 2 both intra- and extracellularly. Further molecular details of the H . pylori catalase came from the serendipitous cloning of the katA gene by Odenbreit et al. (1996). The published N-terminal sequence of a putative 63 kDa lipid-binding adhesin, purified by Lingwood et al. (1993), was used in a search for the corresponding adhesin structural gene by hybridization of a degenerate oligonucleotide probe with an H . pylori gene library. However, when the positive clones were sequenced, it became clear that the product of the gene encoding the supposed adhesin had strong homology with both prokaryotic and eukaryotic catalases, particularly with KatA from Bordetella pertussis (64.9% amino acid identity). This gene functionally complemented an E. coli catalase mutant and an insertion mutation in the chromosomal copy of the gene resulted in the complete abolition of catalase activity in H . pylori (Odenbreit et al., 1996). Thus the katA gene encodes the only functional catalase in H . pylori, although there is an additional ‘catalase-like protein’ encoded in the genome of strain 26695 (Tomb et al., 1997). The katA mutant displayed no differences in adhesion to epithelial cells compared with its isogenic parent, suggesting that the N-terminal sequence reported by Lingwood et a/. (1993) originated from catalase which contaminated the adhesin preparation. As yet, no detailed analysis of the physiological

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consequences of the lack of catalase activity in the katA mutant has been reported. Ferritin is an iron-storage protein which enables iron to be stored in a soluble, non-toxic and bioavailable form (see the review by Andrews, this volume). A 19.6 kDa H . pylori protein, which binds to laminin and is capable of causing weak hemagglutination of erythrocytes, has been characterized and has been described as a ferritin-like molecule owing to the relatively high concentrations of iron which were associated with it (Doig et al., 1993). The gene encoding this protein was cloned and the sequence has homology with the ferritin-like protein produced by E. coli K12 (Frazier et al., 1993). Sequence analysis has shown that all amino acids involved in chelation of inorganic iron by ferritins from other species are conserved in the H . pylori protein. Using immunoelectron microscopy techniques, it has been demonstrated that the H . pylori gene product (termed Pfr) is located in the cytoplasm where it forms paracrystalline inclusions. The studies of Doig et al. (1993) and Frazier et al. (1993) suggest that the H . pylori ferritin stores only non-haem iron, and it may have the function of protecting against intracelMar iron-mediated toxicity. The genome sequence also indicates the presence of a protein with some similarities to bacterioferritin (NapA; Tomb et al., 1997), but an analysis of the sequence of this protein suggests that it is not homologous to other bacterioferritins (Andrews, this volume). Helicobacter pylori has thus been shown to possess several defence mechanisms against oxidative stress, and there is ample evidence that the bacterium is quite capable of survival and growth in atmospheres containing oxygen. One interesting feature is the absence of genes in this bacterium encoding global regulators like OxyR and SoxRS, which control the expression of important oxidative stress-related genes in enteric bacteria. Yet the microaerophilic nature of H . pylori indicates an oxygen sensitivity. which must have a physiological basis. It has long been recognized that the possession of oxygen-sensitive enzymes or proteins may be an important component of a microaerophilic phenotype (Kreig and Hoffman, 1986). This possibility is highlighted below in considering studies on the carbon metabolism of H . pylori.

6. CURRENT KNOWLEDGE OF H. p y h r i CARBON METABOLISM AND SUBSTRATE UTILIZATION 6.1. Early Studies of Metabolism

Using preformed-enzyme tests, a basic enzyme profile of H . pylori was determined (Megraud et a f . , 1985; McNulty and Dent, 1987). Clinical iso-

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lates produced oxidase, catalase, urease, alkaline phosphatase, y-glutamyl aminopeptidase, leucine aminopeptidase and DNAase. These tests were useful for the rapid identification of H . pylori and showed that H . pylori strains are a homogeneous group with respect to enzyme profile. The studies of McNulty and Dent (1987) and Megraud et al. (1985) used rapid enzyme identification procedures to characterize new H . pylori strains and not as an investigation into the organism's metabolic pathways. No evidence was obtained on specific metabolic mechanisms as these rapid enzyme identification techniques did not detect any enzymes which utilize carbohydrates either fermentatively or oxidatively. Evidence of how H . pylori utilizes organic substrates has been accumulated more recently using nuclear magnetic resonance (NMR) spectroscopy, which has proven to be a powerful tool (Chalk et al., 1997). By analysing the growth medium by proton ('H) NMR, before and after growth of H . pylori, it was reported that the bacterium metabolizes lactate and alanine (Dick and Gamcsik, 1989). The major products of metabolism were shown to be acetate, succinate, glycine and citrate. A similar study with multinuclear NMR showed that formate, 2oxoglutarate and lysine were completely metabolized, but no products were detected. These studies by Dick and Gamcsik (1989) provided some evidence for citric acid cycle activity as malate was metabolized to fumarate and an excess of fumarate was metabolized to malate. However, more recent NMR and enzyme studies have called into question most of the results of Dick and Gamcsik (1989) and have provided more conclusive results about carbohydrate metabolism. 6.2. Glucose Metabolism

Mendz and Hazell (1991) produced evidence for the presence of enzymes of the pentose phosphate pathway in H . pylori using 3'P NMR spectroscopy. When glucose 6-phosphate and NADP+ were incubated with bacterial lysates, 6-phosphogluconate and NADPH were produced, indicating glucose 6-phosphate dehydrogenase and phosphogluconolactonase activities. These oxidative activities are initial steps in both the Entner-Doudoroff and pentose phosphate pathways. To distinguish between the two possibilities, bacterial lysates were incubated with 6-phosphogluconate and NADP+. The products were ribulose 5-phosphate and NADPH, which indicated the activity of 6-phosphogluconate dehydrogenase and, therefore, under the conditions used, the presence of the oxidative phase of the pentose phosphate pathway (Mendz and Hazell, 1991). By incubating bacterial lysates with substrates of the other enzymes of the pentose phosphate pathway and identifying the products of the reactions by NMR, activities were indicated for phosphopentose epimerase, transketolase and transaldolase

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(Mendz and Hazell, 1991). Therefore, H . pylori has all the enzymes necessary for a complete pentose phosphate pathway, and its function in this bacterium is likely to be as a source of ribose 5-phosphate for nucleotide synthesis. The discovery that H . pylori possesses enzymes of the pentose phosphate pathway was taken as strong evidence that the bacterium could in fact utilize carbohydrates. Therefore, saccharide kinase activities were investigated as phosphorylation is an important step in the uptake and initial metabolism of saccharides by bacteria. Mendz and Hazell (1993a) used I3C and 31P NMR spectroscopy to monitor the appearance of phosphorylated products when bacterial lysates were incubated with ATP and a selection of mono- or disaccharides. Under the experimental conditions, D-glucose was the only sugar phosphorylated of the 22 investigated. The enzyme involved had a high substrate specificity, was found to have a relatively high K,, and was not inhibited by excess substrate, which suggested that the enzyme was a glucokinase rather than a hexokinase. The evidence that H . pylori did possess mechanisms for utilizing carbohydrates led to investigations being carried out into the transport and incorporation of D-glucose into the bacterium (Mendz et al., 1993, 1995a). Using ~-[U-'~C]glucose in radioactive tracer analysis experiments, a rate for transport and incorporation was determined which indicated directly that H . pylori was able to utilize the monosaccharide. The characteristics of glucose transport into intact cells were determined by Mendz et al. (1995a) using the 1,2,3-3H]glucose. The measured K,,, non-metabolizable analogue 2-deoxy-~-[ value for this substrate was 4.8 mM and significant competition for uptake was obtained with D-galactose and L-arabinose. Sodium ions appeared to strongly stimulate uptake and low concentrations of the sodium transport inhibitor amiloride and the sodium ionophore monensin were inhibitory. However, the classical glucose transport inhibitors cytochalsin B, phloretin and phloridzin did not inhibit uptake, suggesting that the glucose transporter(s) in H . pylori may be novel. In order to determine how glucose was utilized, ~-["C]glucose was incubated with bacterial cells and the loss of label analysed by NMR spectroscopy at regular time intervals (Mendz et al., 1993). The disappearance of the ['3C]glucose label was shown to have biphasic characteristics, with an initial slow period of glucose metabolism, followed by a second phase where the decline of label occurred at a rate at least an order of magnitude faster than the initial rate. There was little product accumulation during the initial phase, but resonances arising from catabolic products were observed during the second phase. In this investigation, several [ '3C]glucose molecules were used, each with the label in a different position. The enrichment levels declined almost twice as fast with ~ - [ l - ~ ~ C ] g l u c othan s e with ~-[6-"C]glucose. Mendz et al. (1993) concluded that the label in position

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1 of glucose was being lost, possibly as COz, in the oxidative phase of the pentose phosphate pathway. This suggested that this pathway may account for the metabolism of a significant amount of the glucose that was available to H . pylori. However, catabolic products were also observed when [ I-'3C]glucose was incubated with H . pylori cells. To identify these products, Mendz et al. (1993) used [13C]glucoseuniversally labelled in all positions. Under their experimental conditions, lactate was identified as the major catabolic product that accumulated in the second phase of glucose metabolism. This led to the conclusion that, in addition to the pentose phosphate pathway, which may be used as a source of carbon for the biosynthesis of macromolecules, there could be at least one other pathway involved in glucose metabolism by H . pylori. These other pathways may have a more significant role in energy conservation in the bacterium. Helicobacter pylori glucose metabolism has also been investigated by Chalk et al. (1994) using 13C NMR spectroscopy. The major experimental difference between this study and that of Mendz et al. (1993) is that Chalk et al. (1994) attempted to determine the importance of oxygen in the atmosphere to H . pylori metabolism. The work of Chalk et al. (1994) confirmed that glucose metabolism is relatively slow, which suggested that glucose may not be a major energy source in vivo. When [I-"C]glucose was incubated anaerobically with a dense cell suspension, two products were accumulated. These were produced immediately on incubation with substrate and were present in low, yet significant, concentrations. The products were identified, using gas chromatography and mass spectrometry, as sorbitol and gluconate. Further anaerobic incubations with [ I-I3C], [2-13C], [3-13C] and [5-'3C]glucose also identified sorbitol and gluconate as products. It was concluded that, as a reduction product of glucose, sorbitol may be involved in the maintenance of the cell redox state during anaerobiosis. Gluconate may have been produced by the breakdown of 6-phosphogluconate, a key intermediate of the pentose phosphate and Entner-Doudoroff pathways (Chalk et al., 1994). Sorbitol and gluconate were also products when cells were incubated aerobically with [2-13C],[3-"C] and [5-'3C]glucose; however, an additional oxidation product was produced which was identified as acetate. The catabolism of glucose to acetate may occur by three routes: glycolysis. a phosphoketolase, or the Entner-Doudoroff pathway. Chalk et al. (1994) investigated the fate of glucose carbons labelled at different positions because, depending on the pathway in use, acetate is generated from different glucose carbons. They showed that [2-13C]and [5-"C]glucose gave rise to acetate labelled on the carboxylic acid carbon (CI). The label of [3-'3C]glucose, was incorporated into the methyl carbon (C2) of acetate and when [l-'3C]glucose was used, no labelled products were observed which could be explained as the loss of gaseous "COZ. This labelling pattern suggested that glucose oxidation may follow the Entner-Doudoroff path-

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way. Under their experimental conditions, Mendz et al. (1993) found no evidence for this pathway, but they did suggest that in addition to a pentose phosphate pathway, which possibly only has a biosynthetic role, there may be other pathways of glucose metabolism. Chalk et al. (1994) found no evidence that lactate was a product of H . pylori glucose metabolism, under either aerobic or anaerobic conditions, which may be due to the experimental conditions of the two investigations being quite different. Results from the genome sequencing of strain 26695 (Tomb et al., 1997) confirms the presence of the pentose phosphate pathway and the EntnerDoudoroff enzymes 2-keto-3-deoxy-6-phosphogluconatealdolase and 6phosphogluconate dehydratase. Interestingly, however, H . pylori also contains a complete set of genes encoding classical glycolytic enzymes, although it is clear from the above data that the actual metabolic evidence for this as the major route of glucose degradation is weak.

6.3. Pyruvate Metabolism

Pyruvate is a major metabolic junction at which several metabolic pathways originate which may lead to either energy conservation or biosynthesis of important products. Chalk et al. (1994) showed that the products of [2-"C]pyruvate incubated anaerobically with H . pylori cells suspended in growth medium were lactate, acetate, alanine and ethanol. There was a more rapid utilization of pyruvate than there was with glucose as 80% of the added pyruvate was metabolized within 1 hour. The production of alanine was found to be dependent on the presence of a nitrogen source in the incubation medium as little alanine was produced when cells were resuspended in phosphate-buffered saline (PBS). This led to experiments in which cells resuspended in PBS were incubated with [2-13C]pyruvateand urea to determine whether the potent H . pylori urease activity was sufficient for urea to act as a nitrogen source. Under these conditions, it was shown that 40% of [2-"C]pyruvate was converted to alanine and there was a decrease in the amount of lactate produced (Chalk et al., 1994). It had previously been suggested that the urease activity had a role in creating an alkaline microenvironment which could aid bacterial survival in the acidic in vivo environment (Ferrero and Lee, 1991; Ferrero and Labigne, 1993). However, the results of Chalk et al. (1994) suggest that urease may also be involved in providing a nitrogen source. It was also concluded from this study that alanine production from pyruvate may be due to transaminase and glutamate dehydrogenase activities, the latter of which has been reported in H . pylori (Mendz and Hazell, 1991). The investigation of Chalk et al. (1994) into the anaerobic metabolism of pyruvate gave no evidence for the accumulation of succinate, which suggested that H . pylori does not possess the

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enzymatic activity to utilize pyruvate as a precursor for the reductive (C4) side of the citric acid cycle. An alternative route for anaerobic pyruvate metabolism, as found in certain facultative anaerobes, is the production of acetate via a pyruvate-formate lyase activity. The accumulation of acetate was shown by H . pylori but experiments with [I-"Clpyruvate showed that this was probably not due to pyruvate-formate lyase activity, as no formate was produced. Aerobic metabolism of [2-I3C]pyruvate by cells resuspended in PBS was also rapid with several products being detected within 20 min (Chalk et al., 1994). The major product was acetate, with smaller amounts of lactate, ethanol and alanine, and in addition there were several unidentified products. However, with the exception of acetate, these were all shown to be further metabolized and were not detected after 80 min. From these results, Chalk et ul. (1994) concluded that there was no evidence for the operation of the citric acid cycle as [2-13C]pyruvate would have been expected to be partially metabolized to "CO2 which would have been detected as a significant decrease in ['klacetate. However, it should be noted that the catabolite concentrations in this investigation were not quantified in absolute terms, but were reported as relative integrals. Mendz et al. (19944 have also studied pyruvate metabolism by H . pylori using 'H and I3C NMR spectroscopy. This study showed that pyruvate was the major product when bacterial lysates were incubated with either L-serine or phosphoenolpyruvate which indicated the presence of serine dehydratase, phosphatase and/or pyruvate kinase activities respectively. These activities and the presence of the Entner-Doudoroff pathway (Chalk et al., 1994) confirm that pyruvate is an important intermediate in the physiology of the bacterium. Under the conditions used, Mendz et ul. (1994a) provided evidence that, in addition to acetate, alanine and lactate, formate and succinate were products of anaerobic pyruvate metabolism. In agreement with the data of Chalk et al. (1994), H . pylori cells resuspended in media containing a nitrogen source were shown to produce alanine from pyruvate. The additional detection of formate in the study of Mendz et al. (1994a) suggested that H . pylori possesses a mixed-acid fermentation pathway involving pyruvate-formate lyase. However, it is now clear from the genome sequence that there is no gene present which encodes pyruvate-formate lyase, so this is unlikely to be the case (at least in strain 26695). Mendz et al. (1994a) also detected the accumulation of small amounts of succinate during the latter stages of pyruvate metabolism. This is evidence for the presence of a citric acid cycle operating in the reductive direction. As with the results of H . pylori glucose metabolism, it can be concluded that different experimental conditions - particularly with regard to the oxygen and C 0 2 concentrations the cells experienced - are the probable reason for the differing products of pyruvate metabolism detected.

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Although these NMR investigations provided evidence for the presence of various routes for pyruvate metabolism, only the activities of certain reactions were detected and no specific enzymes were identified which may be involved in pyruvate metabolism. However, during a study to identify H . pyfori carboxylation enzymes, Hughes et af. (1995) detected a pyruvatedependent, ATP-independent, and avidin-insensitive H14C03- fixation activity. This was shown to be due to an isotope exchange reaction between the carboxyl group of pyruvate and the radiolabelled bicarbonate. This exchange reaction is known to be catalysed by pyruvate:acceptor oxidoreductases (Raeburn and Rabinowitz, 1971). A pyruvate:acceptor oxidoreductase (POR) activity was confirmed in H . pyfori by the ability of cell extracts to catalyse the pyruvate and CoA-dependent reduction of methyl viologen. Methyl viologen reduction was also observed when pyruvate was replaced by 2-oxoglutarate, indicating the presence of a 2-oxog1utarate:acceptor oxidoreductase (OOR) activity, but this enzyme had a lower specific activity. Using the methyl viologen reduction assay, H . pyfori POR activity in cell extracts was found to have an optimum temperature for activity of 30°C and optimum pH of 8.0. Hughes et a f . (1 995) have purified the H . pyfori POR and it was found to be composed of four subunits of 47, 36,24 and 14 kDa. The purified enzyme was specific for pyruvate. Therefore, it was concluded that the 2-oxoglutardte-dependent methyl viologen reduction observed in cell extracts was due to a separate oxidoreductase which possibly has a role in citric acid cycle reactions of H . pyfori. N-terminal sequence analysis of the four H . pyfori POR subunits identified regions of conserved amino acid residues observed in four-subunit pyruvate:ferredoxin oxidoreductases from the hyperthermophilic archaebacteria Archaeoglobus,fufgidus(Kunow et af., 1995) and Pyrococcusfuriosus (Blarney and Adams, 1993) and the hyperthermophilic bacterium Thermotoga maritima (Blarney and Adams, 1994).

6.4. The Importance and Physiological Role of POR and OOR in H. pylork Provision of NADPH and an Explanation for the Microaerophilic Growth Phenotype?

Pyruvate and 2-oxoglutarate:acceptor oxidoreductases are enzymes generally associated with anaerobic metabolism, and are employed by a large number of obligate anaerobes. H . pyfori possesses both activities, but is a microaerophile which does not appear to grow under anaerobic conditions. Hughes et al. (1995) found that the H . pyfori POR enzyme was highly oxygen-sensitive, and the presence of oxygen and the omission of dithiothreitol from purification buffers resulted in its rapid inactivation. It is highly likely that this property is a major contributor to the microaerophilic growth phenotype, but it was suggested that there may be some form of

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protection to prevent inactivation of POR by oxygen in vivo. Several artificial electron acceptors were found to couple with POR; however, in vivo, a low redox-potential electron acceptor such as ferredoxin or flavodoxin would be likely to be involved. A pigmented MonoQ fraction was shown to contain flavodoxin by virtue of its absorption spectrum. A mixture of partially purified H . pylori POR, pyruvate and CoA was shown to reduce this flavodoxin (FldA), suggesting that this was indeed the in vivo electron acceptor. The OOR (also very oxygen-labile) has been purified and the genes encoding the four subunits of both POR and OOR have been cloned and sequenced (Hughes et al., 1998). Although both enzymes are very similar biochemically, their subunits are not highly similar in terms of amino acid sequence identity, and they may have evolved independently. Attempts to produce mutants in either por or oor genes by allelic exchange mutagenesis were unsuccessful, indicating that both POR and OOR are essential enzymes in H . pylori (Hughes et al., 1998). Interestingly, FldA appears unable to act as an in vitro electron acceptor for OOR (Hughes et al., 1998). The most likely alternative electron acceptor is a ferredoxin, and there are candidate genes encoding ferredoxin-like proteins in strain 26695 (Tomb et al., 1997). In many anaerobes, the electrons from reduced flavodoxin or ferredoxin are used in a variety of reductive processes which regenerate the oxidized acceptor and which may or may not be linked to energy conservation. The simplest route is the formation of molecular hydrogen via a hydrogenase. Other reactions include nitrogen fixation and sulphate reduction. In H . pylori, pyruvate- and 2-oxoglutarate-dependentreduction of NADP but not NAD has been observed in cell-free extracts incubated anaerobically, implying the presence of a flavodoxin and/or ferredoxin NADP reductase activity (Hughes et al., 1998; Fig. I). This may be important in energy conservation in H . pylori, as NADPH appears to be the preferred respiratory electron donor rather than NADH (Chang et al., 1995; Hughes et al., 1998) and there is no evidence for a transhydrogenase in the genome sequence. Thus, POR and OOR are crucial enzymes in H . pylori not only for the generation of acetyl-CoA and succinyl-CoA for biosynthetic purposes, but also in the provision of electrons for respiration (Fig.1).

6.5. Alcohol Dehydrogenase

Helicohacter pylori contains an alcohol dehydrogenase (ADH) which has been characterized in several studies (Roine et al.. 1992; Salmela ef al., 1993, 1994), and a gene encoding a short-chain alcohol dehydrogenase is present in strain 26695 (Tomb et al., 1997). In the presence of NAD, cell-free extracts oxidized ethanol to acetaldehyde in a concentration-dependent

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flavodoxirl, Acetate Acetyl- oA

NADP

Oxaloacetate

J

lo L-Malate

Fumarate

Succinate

itrate

l6

Isocitrate

NADP

2-oxoglutarate C O A X F4x SuccinyCCoA

Fd,

x

NADPH NADP

Figure I Pyruvate metabolism in H. pylori and its relationship to reactions of the citric acid cycle. Enzymes are denoted by numbers. I ; Pyruvate:flavodoxin oxidoreductase. 2; flavodoxin: NADP oxidoreductase, 3; Acetate kinase, 4; phosphotransacetylase, 5 ; citrate synthase; 6, aconitase, 7; isocitrate dehydrogenase, 8; 2-oxoglutarate: ferredoxin oxidoreductase, 9; ferredoxin:NADP oxidoreductase, 10; malate dehydrogenase, 11; fumarase, 12; fumarate reductase. The cycle is incomplete and the exact mechanism for oxaloacetate generation is unknown. Enzymes 2 and 9 are putative at present.

manner with a rather high apparent K , of 103 mM ethanol (Roine et al., 1992), although significant acetaldehyde production was formed from a low ethanol concentration in the range which might be expected in vivo (Salmela et al., 1993). The activity was competitively inhibited by 4-methylpyrazole. Acetaldehyde was also produced by intact cells in the absence of nictotinamide nucleotides (Salmela et al., 1994), and it was suggested that the formation of toxic acetaldehyde could contribute to pathogenesis in vivo. However, Graham et al. (1994) pointed out that the cytoplasmic location of the enzyme and the high reactivity of acetaldehyde would argue against this, and considered the finding of doubtful clinical significance. Considering that in several of the NMR studies discussed above, ethanol was detected, it is more likely that, in vivo, the function of ADH is to produce ethanol as a fermentation product from acetaldehyde. This implies the presence of a pyruvate decarboxylase activity to form acetaldehyde from pyruvate, which has yet to be demonstrated.

6.6. Fumarate Metabolism

The investigations into H . pylori pyruvate metabolism have provided evidence that this 2-oxoacid is metabolized by enzymes which are normally

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associated with anaerobic processes (Mendz et al., 1993; Hughes el a/., 1995). Although the optimal in vitro environment for H . pylori growth is microaerobic, these anaerobic features led to studies into the fumarate metabolism of the bacterium. The reduction of fumarate is catalysed by the enzyme fumarate reductase. which is normally a terminal step of a proton-translocating electron transport chain. This can be an important source of ATP for anaerobic bacteria (Kroger et al., 1992). Mendz and Hazell (1993b) identified the products of fumarate catabolism of H . pylori cells or lysates using one- and two-dimensional NMR techniques. Under their experimental conditions, the primary product of fumarate metabolism was malate and this was subsequently converted to pyruvate after approximately 10 h. After this time interval there was still a considerable amount of fumarate present. Further incubation showed the disappearance of pyruvate and, over a 40 h incubation period, the final products were identified as succinate, acetate, lactate, alanine and formate. By studying the pattern of metabolite evolution over time, Mendz and Hazell (1993b), showed that the decline of fumarate levels had biphasic characteristics. The initial period of fumarate utilization was due to a burst of malate production and was comparatively short, but had a fast rate. I t was suggested that this reaction was catalysed by fumarase and the data indicated that an equilibrium was established between the concentrations of both metabolites. At a similar time to this equilibrium being achieved, succinate was produced which was indicative of a second phase of fumarate metabolism. This reaction indicated a fumarate reductase activity which led to the exhaustion of fumarate substrate. It was concluded from this investigation that H . pylori may have a reductive C4 electron sink pathway from oxaloacetate to succinate which could have an important role in maintaining a redox balance for a proton-translocating electron transport chain. This provided some evidence that H . pylori may generate ATP via anaerobic respiration. Using "C-NMR, and much shorter incubation times, Chalk et at. (1997) clearly showed that the major end product of fumarate metabolism under anaerobic conditions was succinate, again indicating an active fumarate reductase, and they also noted the transient accumulation of malate. The fumarate reductase from H . pylori has been purified by Birkholz tit ul. (1994). They identified an immunogenic protein with a molecular mass of 80 kDa which was recognized by 55% of serum samples from patients infected with H . pylori, using Western blots of butanol extracts of H . p j h r i membranes. The N-terminal sequence of this protein showed 80% identity with the N-terminal sequence of subunit A of the fumarate reductase of Wolinellu succinogenrs (Lauterbach et a/., 1990). The W . succinogmes enzyme consists of three subunits with apparent molecular masses of 70 (FrdA), 31 (FrdB), and 25 (FrdC) kDa. To confirm the identification of a H . pylori fumarate reductase subunit A and to try to identify a subunit B,

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Birkholz et al. (1994), carried out Western blots by reacting rabbit antisera raised against both these subunits of the W . succinogenes enzyme with a partially purified H . pylori fumarate reductase preparation. This showed reaction of each antiserum with the fumarate reductase preparation and identified 80 and 31 kDa H . pylori polypeptides, which was consistent with the sizes of subunits A and B of W. succinogenes fumarate reductase respectively. Native gel electrophoresis and SDS-PAGE of purified H . pylori fumarate reductase indicated that these two polypeptides were part of the same protein complex. There was also evidence for further subunits of the protein complex as polypeptides with molecular masses of 25, 23.5 and 22 kDa were present in the same sample. These were found not to be breakdown products of the 80 kDa protein as they were not recognized by the antiserum against subunit A of W . succinogenes fumarate reductase. However, it was not clear whether these polypeptides co-purified with the fumarate reductase complex. The 25kDa polypeptide may be identified as subunit C as this was the same size as subunit C of W . succinogenes fumarate reductase, which is a dihaem cytochrome b (Kortner et al., 1990), and the active fraction obtained from ion-exchange chromatography had a reddishbrown colour that is characteristic of cytochromes. H . pylori fumarate reductase activity was demonstrated in purified fractions of cell membranes by either measuring the succinate oxidation to fumarate by methylene blue or by fumarate reduction to succinate by reduced benzyl viologen. Enzyme activity was exclusive to membrane preparations, suggesting that the fumarate reductase was membrane-bound like the fumarate reductase of W. succinogenes. The 80 kDa protein was found to be ubiquitous as sonicates of all H . pylori strains tested were shown to react with rabbit antiserum against subunit A of W . succinogenes fumarate reductase complex. This confirmation of a H . pylori fumarate reductase activity provides further evidence that this microaerophilic bacterium may be capable of anaerobic respiration, yet it has thus far proved impossible to grow it anaerobically with fumarate as the electron acceptor (A.A. Davison, P.A. Chalk and D.J. Kelly, unpublished results; Hoffman et al., 1996). The cloning and sequencing of fumarate reductase structural genes was briefly reported by Davison et al. (1994a). A cluster of genes encoding close homologues of the W . succinogenes frdCAB genes, and in the same order, was identified and frdCA were completely sequenced. The sizes of the deduced products (FrdA, 80. I kDa and FrdC, 28.8 kDa) are in good agreement with the sizes of the polypeptides observed by Birkholz et al. (1994). In facultative anaerobes, fumarate reductase is normally induced under anaerobic conditions and repressed aerobically, but in H . pylori cells grown under a range of oxygen concentrations, the specific activity of the enzyme in cell-free extracts did not change markedly, indicating that it is constitutive

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(Davison et al., 1993). Genome sequencing confirms the presence ofJrdCAB 1997). genes in H . pylori (Tomb et d., Fumarate reductase is not present in humans, therefore the H . pylori enzyme could be a potential chemotherapeutic target. Mendz et (11. (1995b) employed NMR spectroscopy and growth culture techniques to investigate the cytotoxicity of morantel, oxantel and thiabendazole, which are known to inhibit fumarate reductase in parasitic worms. This study showed that the three compounds had an inhibitory effect on the H . yylori enzyme, The strength of the inhibitory effect was found to be oxantel > thiabendazole > morantel. When a dense cell suspension was incubated with 40 mM fumarate in the presence of 0.9 mM oxantel, the rate of succinate production was reduced to 36 f 5 nmol/min/mg protein from 207 f 25 nmol/min/mg protein when no inhibitor was present. By examining the Michaelis constant and maximal velocity it was concluded that oxantel was a competitive inhibitor of fumarate reductase; the Michaelis constant increased and maximal velocity remained constant. It was discovered that adding excess fumarate to actively growing H . p.ylori cultures led to a decreased cell growth and viability. Mendz el ul. (1995b) concluded that excess substrate has an inhibitory effect on the fumarate reductase enzyme and a bactericidal effect on the cells. In addition, they showed a correlation between the inhibitory potency of the three compounds and their bactericidal effects. This suggested that inhibition of fumarate reductase was the basic cause of cell death, but to ensure that the inhibitors had no other effects, the inhibition by oxantel of enzymes of the citric acid cycle was tested. Oxantel did not inhibit fumarase, malate dehydrogenase or cisaconitase activities, which suggested some degree of specificity of this compound. However, all three compounds had a relatively high minimum inhibitory concentration (MIC) against H . pylori and the in vitro effects of oxantel were shown to be reversible when the compound was removed from the growth medium. This investigation also provided evidence that fumarate reductase is an important enzyme in the metabolism of the bacterium but none of the tested compounds would be likely to be effective in the eradication of H . pylori from its natural niche. 6.7. The Nature of the Citric Acid Cycle in H. py/ori

It should be apparent from the foregoing sections that there has been considerable uncertainty over the activity and role of the citric acid cycle in H . pylori. The accumulation of acetate from pyruvate under aerobic incubation conditions in a number of the NMR investigations reviewed above strongly suggests a major diversion of the acetyl-CoA resulting from the POR reaction towards acetate production rather than oxidation through the citric

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acid cycle (Fig. I). This could occur via phosphotransacetylase and acetate kinase, genes for which have been identified in strain 26695 (Tomb et al., 1997), and would yield IATP per mole of acetyl-CoA. Nevertheless, it is equally clear that H . pvlori does show oxidative activity. In a recent report, in vitro enzyme assays have demonstrated the presence of a number of the enzymes required for the operation of a citric acid cycle (Hoffman et al., 1996), with the apparent exception of succinyl-CoA synthetase. It cannot be concluded from this, however, that there is a complete cycle operating solely in the oxidative direction, because of the clear evidence for an apparently constitutive fumarate reductase activity in H . pylori. Indeed, it is likely that the succinate dehydrogenase activity reported by Hoffman et al. (1996) is in fact due to fumarate reductase. An additional point of interest is the very low activity (< 10 nmol/min/mg protein) of NADH-linked malate dehydrogenase activity in H . pylori (Hoffman et a/., 1996). In bacteria with complete oxidative citric acid cycles, this enzyme is invariably present at high specific activity. Davison et al. (1993) reported the presence in H . pylori of a dyelinked (NADH-independent) malate dehydrogenase activity which was loosely membrane bound and which was present at higher specific activity than the NADH-linked activity. This is probably a flavoprotein MDH of the same type as found in some other bacteria (Ohshima and Tanaka, 1993) and may have a separate function in donating electrons from malate directly to the respiratory chain, as L-malate-dependent cytochrome reduction has been observed in virro (Davison et al., 1993). Interestingly, Tomb et al. (1997) have concluded from the genome sequence of strain 26695 that the citric acid cycle is incomplete, there being no genes for succinate dehydrogenase, succinyl-CoA synthetase or NAD-linked malate dehydrogenase. There is also no evidence of genes encoding glyoxylate shunt enzymes (isocitrate lyase and malate synthase). although again the activities of these enzymes have been reported in some strains (Hoffman rr al., 1996). Genes encoding homologues of citrate synthase, aconitase, isocitrate dehydrogenase, fumate reductase and fumarase are present, and although there is no 2-oxoglutarate dehydrogenase complex, it has been shown that the conversion of 2-oxoglutarate to succinyl-CoA is effected by OOR (Hughes et al., 1995, 1998; Hoffman et al., 1996). Considering all of the available data, it seems most likely that both an oxidative (C6) branch from acetyl-CoA to succinyl-CoA and a reductive (C4) branch from oxaloacetate to succinate is operating in H . pylori, according to the scheme of Fig. 1. The main function of the oxidative branch would be in the provision of succinyl-CoA and, to a lesser extent, NADPH for biosynthesis and respiration. The reductive branch may function in fumarate respiration. Both branches require oxaloacetate as their starting point and it is a paradox that H . pylori 26695 contains no

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genes for carboxylation enzymes that could produce this intermediate (see Section 6.8, below). 6.8. The COz Requirement of H. pylori

In addition to a lowered oxygen concentration, H . pylori requires a much higher COz level for growth (5-10%, v/v) than is present in the normal atmosphere. It is possible to grow H . pylori in an atmosphere containing 21% (v/v) O2 as long as the C 0 2 level is increased to 10% (v/v) (P.A. Chalk, personal communication). Microorganisms which have an elevated dependence for C 0 2 are classified as capneic or capnophilic and the best known examples of pathogenic capnophilic organisms are Neisseria spp. (Cox and Baugh, 1977). This high requirement for C 0 2 relates to the reported nature of the niche occupied by the organism. It has been calculated that gastric exudates contain bicarbonate concentrations of 10 mM in dogs and 25 mM in humans (Dittmer, 1961). These values can be converted into the partial CO1 pressure in gastric exudates and the mean of these two values calculates to 7 1.8 mmHg. This partial pressure corresponds to a potential concentration of aqueous C 0 2 of 9.45% (v/v). This concentration may be further increased in the natural niche of H . pylori as a result of the high levels of urease activity exhibited by the bacterium. The apparent dependence on high COz levels should have a bearing on the organism's metabolism and was initially investigated by Hughes ef al. (1995). Acetyl-CoA carboxylase (involved in fatty acid biosynthesis) and a low malic enzyme activity were detected, but no activities were identified for other important C 0 2 fixation enzymes such as pyruvate carboxylase (PC), phosphenolpyruvate (PEP) carboxylase and PEP carboxykinase. An apparent pyruvate-dependent C 0 2 fixation activity was detected, but was in fact found to result from the isotope exchange reaction of POR, discussed above. In contrast, Hoffman rf ul. (1996) reported the detection of PC, PEP carboxylase and PEP carboxykinase in some strains of H . pylori, but the specific activities were very low and their physiological significance is uncertain, particularly in view of the fact that there are no candidate genes encoding these enzymes (or indeed malic enzyme) in the genome sequence of strain 26995 (Tomb et ul., 1997). In fact. in view of the obligate C 0 2 requirement, it is astonishing that the genomic data indicate there to be no anaplerotic COz fixation mechanisms in this bacterium, and thus no obvious way to explain the production of oxaloacetate, as noted above. Biotin carboxylases employ a catalytic mechanism in which the bicarbonate ion (HC03-), rather than C 0 2 itself, is transferred to the covalently bound biotin prosthetic group. A study of the acetyl-CoA carboxylase activity in H . pylori by Burns et al. (1995) revealed that the apparent K ,

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value of this enzyme for bicarbonate was 16.6 mM. This high value was proposed as a major contributor to the cellular C 0 2 requirement. This is an interesting suggestion, but the observation that the addition of bicarbonate alone cannot apparently replace the requirement for the addition of gaseous C02 for the growth of H. pylori in liquid media (P.A. Chalk, personal communication; Tuckwell and Chalk, 1993) may indicate a more complex role for C02. It is also of interest that there are two genes in H. pylori that may encode carbonic anhydrases (Tomb et al., 1997). These enzymes could be involved in some mechanism of C 0 2 concentration in the bacterium which may be important in explaining the C 0 2 requirement. 6.9. Anabolic Pathways

The studies of Nedenskov (1994) and Reynolds and Penn (1994) indicated that H. pylori may have defective anabolic pathways which result in complex nutritional requirements and, in particular, the apparent inability to synthesize a number of amino acids. There have been few physiological studies of amino acid biosynthesis in H . pyiori, and the absence of key enzymes which might explain this nutritional phenotype has not been established biochemically, although as noted above, the genome sequence gives some insights into the capability of the bacterium for de novo amino acid biosynthesis. Some studies have been carried out into the synthesis of pyrimidine and purine nucleotide precursors in H. pylori (Mendz et al., 1994b,c). Cells incorporated the nucleobases adenine, guanine and hypoxanthine in addition to the nucleosides adenosine, guanosine and deoxyadenosine, but not deoxyguanosine. Evidence was obtained from both radiotracer and N M R assays for the presence of salvage pathways for purine synthesis (Mendz et al., 1994~).Aspartate, bicarbonate and orotate carbon was incorporated into genomic DNA, providing evidence that a de novo route to pyrimidine nucleotide synthesis was present. Uracil and uridine were also incorporated but to a much lesser extent. These data indicated that a salvage pathway for pyrimidine nucleotide synthesis was present, but that the de novo pathway is likely lo be more important.

7. THE RESPIRATORY CHAIN OF H. py/ori 7.1. Substrate Oxidation

An early investigation into the respiration of H . pylori showed that whole cells were capable of oxidizing D-glucose, formate, DL-lactate, succinate and

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pyruvate (Baer et al., 1993). The rates of respiration were low for D-glucose and formate, but DL-lactate, succinate and pyruvate were much more readily oxidized. Baer et al. (1993) concluded from these observations that glucose and formate did not significantly influence the flow of electrons along an electron transport chain, which supported the notion that H . pylori did not metabolize common saccharides (Megraud et al., 1985; McNulty and Dent, 1987). However, NMR spectroscopy suggested that H . pylori is capable of metabolizing glucose, as activities of enzymes of the pentose phosphate pathway have been identified (Mendz and Hazell, 1991) and there is evidence for the presence of an Entner-Doudoroff pathway (Chalk c’t a/., 1994). Therefore, the identification of a low rate of respiration of glucose by intact cells may have been more significant than originally thought by Baer et ul. (1993). A more comprehensive study into the kinetics of substrate oxidation by whole cells and cell membranes of H . pylori was carried out by Chang et nl. (1995). Under their experimental conditions, cell preparations from stirred broth cultures showed an endogenous metabolism which consumed oxygen in the absence of substrate for several hours. By adding metabolizable substrates, variable rates of oxygen uptake were observed; 25 p ~ ~ l a c t a stite mulated oxygen uptake, but more frequently endogenous metabolism was not increased even when high concentrations of substrates were added. Cells harvested from chocolate agar plates also showed variable rates of oxygen uptake in the absence of added substrates. It was concluded that this endogenous metabolism from cells grown in stirred broth cultures may have been due to the leakage of cell metabolites from stressed or autolysed cells. Cells from broth cultures which were incubated without agitation were shown to have no or only a low endogenous oxygen uptake. These cells oxidized ethanol, fumarate, glucose, D-lactate, pyruvate and succinate. Low concentrations (25 p ~ of) pyruvate, D-lactate and succinate were rapidly oxidized and the respiration rates were relatively high. Further substrate addition when oxygen uptake had ceased resulted in a similar rate of respiration. The K , values for pyruvate, D-lactate and succinate were low, which indicated a high affinity for these substrates. Chang et al. (1995) concluded that this was evidence that H . pylori cells may be adapted to utilizing these substrates in vivo. In addition, a lower affinity for ethanol and fumarate was detected which suggested that these substrates may not be oxidized at significant rates in vivo. From this study it was calculated that the total oxygen taken up during lactate and pyruvate oxidation was insufficient for complete oxidation to C02 and H 2 0 via the citric acid cycle. This did not rule out the possibility of the presence of some H. pylori citric acid cycle activity as the amount of oxygen taken up was consistently greater than that required for the oxidation to acetate and C02. This study, in comparison with that of Baer et al. (1993), definitely showed that glucose was oxidized

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by H . pylori cells. The rate of glucose oxidation was shown to be variable and the rate declined quite rapidly. The initial rate was similar to that when the same concentration of succinate was added; however, this rate declined to an almost undetectable level within 30 minutes. This observation was not due to substrate exhaustion as further additions of glucose did not result in the stimulation of oxygen uptake. Chang et al. (1995) were unable to detect oxygen uptake when acetate, glycerol, L-lactate, oxaloacetate, 2-oxobutyrate and several amino acids including aspartate and glutamate were added to H . pylori cells. Experiments with H . pylori cell membranes showed the oxidation of NADH, NADPH, D-lactate and succinate. However, there was a much higher rate of NADPH oxidation compared with that for NADH, which suggested that the former was likely to be a more significant physiological electron donor. There is also evidence for a separate, soluble NADPH dehydrogenase in H . pylori (Clayton et al., 1994). The mechanism of succinate respiration is of some interest because acetate has been identified as the major product by ‘H-NMR analysis (Chalk et al., 1997). In one experiment, over a 180 min incubation period with a starting concentration of 2 mM succinate, 1.5 mM succinate was consumed and 1.0 mM acetate was produced. From a comparison of the rates of succinate consumption and oxygen uptake, a stoichiometry of 1.5 moles O2 per mole of succinate was obtained (A.A. Davison, P.J. White and D.J. Kelly, unpublished results). The overall reaction is thus: (CH2COOH)Z + 1.502 = CH3COOH

+ 2C02 + H20

Bearing in mind the data reviewed above concerning the known citric acid cycle reactions in H . pylori, a plausible pathway for succinate oxidation is shown in Fig. 2, which involves a reversal of the C4 branch reactions to malate or oxaloacetate, the decarboxylation of malate or oxaloacetate to pyruvate and the oxidative decarboxylation of pyruvate to acetate (via POR and acetate kinase/phosphotransacetylase). In this pathway, two moles of C 0 2 are produced per mole of succinate oxidized and the three electron pairs produced would require 1.5 moles of O2 for complete oxidation via the respiratory chain, consistent with the above overall equation.

7.2. Composition of the Respiratory Chain

Until recently, very little was known about the nature of the respiratory apparatus in H . pylori, apart from the early observations that menaquinone was the sole isoprenoid quinone present (Collins et al., 1984) and that the bacterium is ‘oxidase positive’. Baer e f al. (1993) investigated the effects of inhibitors on the respiratory activity of H . pylori; as would be expected,

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5 mM cyanide was an efficient inhibitor of the oxidation of lactate, with the respiration rate being decreased by almost 90%. Antimycin A and myxothiazol also inhibited lactate respiration by 54% and 33% respectively, which suggested inhibition of a cytochrome bcl complex. In parallel to this investigation into the effect of inhibitors on respiration, intracellular ATP levels were measured and cyanide caused the greatest decrease of intracellular ATP levels, presumably as a consequence of the inhibition of respiration. This study also produced evidence that certain bismuth salts were partial inhibitors of H. pylori respiration. Bismuth gallate and bismuth subsalicylate showed a significant lowering of the rate of respiration of lactate, which suggested one mechanism for the reported antibacterial effects of bismuth salts in vitro (Vogt et al., 1989). Davison et ul. (1994b, 1995) reported the cloning and sequencing of the structural genes for a hydrogenase from H. pylori, the deduced amino acid sequence of which was very similar to the hydrogenase of the related rumen bacterium Wolinella succinogenes. Gilbert el al. ( 1995) also cloned a hypB homologue, which in Bradyrhizobium juponicum encodes a nickel-binding protein involved in hydrogenase biosynthesis. It has since been found that H . pylori is capable of hydrogen oxidation (Maier et al., 1996), and the presence of a membrane-bound hydrogenase activity which could couple with a variety of artificial and physiological electron acceptors with a positive redox potential was described. From Western blotting experiments with antisera raised

Succinate

Fumarate L M a l a t e -!-w (Oxaloacetate)

Acetate Fixt4re 2 Possible mechanism for succinate respiration that accounts Tor acetate a s the observed end product and the ratio of 1.S moles of oxygen consumed per mole of succinate oxidized. The pathway involves a reversal of the fumarate reductase and fumarase reactions and requires the presence of malic enzyme for the oxidative decarboxylation of malate to pyruvate and CO,. This activity has been detected in H . pylori (Hughes r t ul., 199S), although no candidate gene for it has been identified. Another mechanism of malate decarboxylation (e.g. via oxaloacetate, dotted lines) is not ruled out. Pyruvate is converted to acetate via POR, phosphotransacetylase and acetate kinase.

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against the Bradyrhizobium japonicum uptake hydrogenase, Maier et al. (1996) identified H . pylori hydrogenase polypeptides of about 65 and 26 kDa. Taken together, the available data indicate that H . pylori possesses a NiFe type of H2-uptake hydrogenase which could act as an electron donor to the respiratory chain, and thus contribute to energy conservation if molecular hydrogen is present (Fig. 3). The quinone and cytochrome complement of H . pylori, and how this varies with different oxygen concentrations during growth, has only recently been investigated (Maier et a!., 1996; Marcelli et al., 1996; Nagata et al., 1996). In agreement with Collins et al. (1984), Marcelli et al. (1996) identified the major isoprenoid quinone as menaquinone-6 (MK-6), with traces of MK-4, but no ubiquinone. Moss et al. (1990) reported an unusual type of MK-6 in H . pylori, but this was not seen in the study of Marcelli et al. (1 996), who also found no changes in the amounts or types of menaquinone in cells grown at different oxygen concentrations (over the range of 2-15%, v/v). Their spectroscopic analysis of cells and membranes revealed the presence of h- and c-type cytochromes but there was no evidence for terminal oxidases of the a- or d-types (but see below). Membrane-free ‘cytoplasmic’ extracts contained cytochrome(s) c (undoubtedly localized in the periplasm in intact cells), which 0 d u m and Andersen (1995) had previously identified as being responsible for ascorbate oxidation in cell extracts. In CO difference spectra, a peak at 416 nm was attributed to the CO-ligated form of cytochrome o and a trough at 428 nm was thought to originate from a CObinding cytochrome b. One novel feature noted in the CO difference spectra

Hydrogen

D-Lactate + MQ

-bc,

c~~~

cS5,- peroxidase # + cb-oxidase

G-3-P Fumarate Figure 3 Organization of the respiratory chain in H . pylori based on available biochemical and genomic data. A linear electron transport pathway to the cb-type terminal oxidase is shown, although there is some evidence for an alternative oxidase (see text). The source of electrons for the peroxidase has yet to be established, but it is likely that cytochrome cSs3 is the immediate electron donor.

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of both membranes and intact cells by Marcelli st al. (1996) was a peak at 628 nm. Although similar in wavelength to that expected for the CO complex of cytochrome d, the corresponding band of the ferrous form (at 626630 nm) was not present in the reduced minus oxidized spectra. I t was suggested that this may be due to the rapid turnover of a cytochrome (1, except when the ferrous form is trapped as the CO-adduct, rendering it spectrally visible.

7.3. The Terminal Oxidase(s)

The question of the nature of the terminal oxidase(s) present in H . py1or.i is of considerable interest, because of the possible relationship to microaerophily. There is in fact both genetic and biochemical evidence that H. pylori contains a ch-type haem-copper cytochrome c-oxidase (Clayton ct d., 1995; Alderson et ul., 1996; Nagata et al., 1996). This type of oxidase was first characterized in symbiotic rhizobia, where it is encoded by the ,fixNOQP operon (Preisig et ul., 1993). It has a very high affinity for oxygen which enables efficient respiration to be carried out within the microaerobic environment of the bacteroids (reviewed in Garcia-Horsman L J ~ul., 1995). Helicohucter pylori contains homologues of the rhizobial ji.uNOQP genes 1996, 1997; Tomb et ul., 1997) located (Clayton et ul., 1995; Alderson et d., in an operon as in other bacteria, but surprisingly there is no evidence of associated regulatory genes or a closely IinkedjxCHIS like-operon which in other bacteria is invariably associated with the structural genes. The oxidase activity has been characterized by Nagata rt ul. ( 1 996) who reported a K , value of 0.4 ~ L Mfor oxygen and a high sensitivity to KCN (ICSOof 4 p ~ ) . Some of the polypeptides of the oxidase were tentatively identified from haem-stained gels and a spectral analysis of cytochromes in the membrane fraction was in broad agreement with the findings of Marcelli et ul. ( 1 996). However, this may not be the sole terminal oxidase in H. pylori, at least from biochemical studies (but see below). Alderson et al. (1996, 1997) reported that high concentrations (20 p ~ of ) the specific cytochrome hc, complex inhibitors antimycin A and niyxothiazol did not completely inhibit lactate respiration in intact cells, indicating that the respiratory chain is branched at the level of the menaquinone pool. Furthermore, although cytochrome c oxidase activity was very sensitive to cyanide (lCso of 5 ~ L M in ) , agreement with Nagata et al. (1996), lactate respiration was much less sensitive. These data are consistent with the presence of both a quinol oxidase of low cyanide sensitivity and a cytochrome c oxidase of high cyanide sensitivity (the ch-oxidase). Cytochrome hd oxidase is generally cyanide-insensitive and thus could be a candidate for the quinol oxidase.

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Maier et al. (1996) reported the characteristic spectral signal at 595 nm for the high-spin haem b of such an oxidase (in addition to haem d diagnostic signals) in H2-reduced minus Oroxidized membranes of a clinical isolate of H . pylori grown in the presence of hydrogen, but this was not observed by Marcelli et al. (1996) using strain NCTC 11637, and there is equivocal spectroscopic data concerning the presence of haem d, as noted above.

7.4. The Genome Sequence indicates a Simple Respiratory Chain Organization

The genome sequence of strain 26695 has yielded some surprising insights into the respiratory chain of H . pylori, not all of which agree with the above biochemical data. The main feature is the apparent simplicity of the electron transport pathways inferred, with the cb-type cytochrome c-oxidase being the sole terminal oxidase identified. Apart from fumarate reductase, discussed above, there is no evidence that H . pylori possesses any other type of reductase for alternative electron acceptors such as nitrate, nitrite, dimethyl sulphoxide (DMSO) etc. In addition to the hydrogenase, genes for only three types of primary dehydrogenases have been identified as encoding potential donors to the menaquinone pool; glycerol 3-phosphate dehydrogenase, D-lactate dehydrogenase and a multi-subunit NAD(P)Hquinone oxidoreductase (Tomb et al., 1997), although there is biochemical evidence for an NAD-independent L-malate dehydrogenase, as noted above. Intermediate electron carriers between the MQ-pool and the oxidase are a cytochrome bc, complex @c gene products) and a cytochrome c553which is the most likely immediate donor to the cb-type oxidase. There is no obvious candidate for an alternative (quinol) oxidase in the genome sequence. A cytochrome c551 peroxidase gene is present, which may encode a periplasmic type of hydroperoxidase for detoxification purposes. By analogy with electron transport in a related microaerophile, Campylobacrer mucosalis (Goodhew et al., 1988), the cSs3is also the most likely electron donor to this peroxidase. Thus, the electron transport chain is astonishingly simple, and H . pylori seems unusual in not having the type of branched respiratory chain common in the majority of bacteria studied to date. Another unusual feature is the presence of only two obvious homologues of proteins required for the biogenesis of c-type cytochromes; Gram-negative bacteria normally contain a complex system for haem transport, insertion and assembly into the apoprotein in the periplasm. Figure 3 summarizes knowledge about membrane-associated electron transport in H . pylori.

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8. NITROGEN METABOLISM IN H. py/ori

8.1. Assimilation and Management of Nitrogen

Helicohacter pylori is capable of rapid growth in complex media where the main available carbon, nitrogen and energy sources are amino acids. In addition to the direct utilization and incorporation of alanine, arginine, aspartate, glutamate, proline and serine into cell biomass (Mendz and Hazell, 1995; Stark et al., 1997), it has been demonstrated that amino acids such as glutamine and asparagine can be converted to central intermediary metabolites by deamination reactions which occur at fast rates (Mendz and Hazell, 1995; Stark et a/., 1997). Such reactions would also generate ammonium ions which could act as a nitrogen source for biosynthesis. Ammonia is also one of the products of the urease reaction (see below). The assimilation of ammonia in H . pylori appears to be carried o u t by glutamate dehydrogenase and/or glutamine synthetase, genes for which have been identified (Tomb et. al., 1997). Interestingly, however, there is no candidate gene encoding a glutamate synthase. Many bacteria employ a two-component sensor-regulator system (NtrB/C) along with a special sigma factor (NtrA or RpoN) to regulate the synthesis of nitrogenmetabolizing enzymes in response to environmental nitrogen availability. Although H . pylori has an RpoN homologue, there are no other obvious components of an Ntr-like system in this bacterium (Tomb et al., 1997). There is biochemical evidence that H . pylori possesses a urea cycle of the type normally found in eukaryotes, and a few other prokaryotes. Mendz and Hazell (1996) used one- and two-dimensional NMR spectroscopy and radioisotopic labelling to demonstrate the formation of ornithine and ammonium from L-arginine in bacterial lysates. The ornithine was converted to citrulline by an ornithine transcarbamoylase activity, and both arginosuccinate synthetase and arginosuccinase activites were also demonstrated. It was suggested that this cycle may be involved in maintaining nitrogen balance in the cells, perhaps disposing of excess nitrogen generated by the rapid catabolism of amino acids in the form of urea, which would be subsequently hydrolysed by urease.

8.2. The Urease of H. pylori

H . pylori urease is an extremely active enzyme which catalyses the hydrolysis of urea to carbon dioxide and ammonia. The native enzyme consists of two polypeptides with molecular masses of 29.5 and 61 kDa, which combine to produce complexes of between 300 and 650 kDa (Turbett et al., 1992). The

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importance of urease in colonization was demonstrated when a ureasenegative variant of H. pylori failed to colonize gnotobiotic pigs (Eaton et al., 1991). The activity of urease may help the bacterium to survive in acidic environments as the production of ammonia may neutralize acid in the immediate region surrounding the organism. Therefore, survival in the stomach for a finite time may be possible before penetration of the less acidic mucus (Marshall et al., 1990). If H. pylori is supplied with physiological amounts of urea in vitro, it can survive for 120 min at pH 2.0 (Goodwin et al., 1986). The reason for the survival of H. pylori in these urea-supplemented acidic conditions was explained when pH optima experiments showed that although the major peak of activity was at pH 7.0, there was a second peak of activity at pH 4.0 (Ferrero and Lee, 1991). To support the findings that urease helps the survival of H. pylori in acidic conditions, urease-negative mutants were shown to be quickly killed when exposed to low pH (Segal et af., 1992). However, it is clear that urease is more than simply a defence against acid, and it can be regarded as a virulence factor with several distinct functions. The activity of urease may cause some of the tissue damage associated with gastritis, in several ways. It has been hypothesized that the accumulation of ammonia from urea hydrolysis might significantly increase the pH of the gastric mucosa which could be responsible for the back diffusion of hydrogen ions. This would increase the acidity at tissue level and may cause tissue injury (Hazel1 and Lee, 1986). In addition, the presence of ammonia itself could directly act as a cytotoxic agent (Smoot et al., 1990). As the H. pylori urease activity was one of the most prominent features of the bacterium, the genes for this enzyme were amongst the first to be cloned and sequenced. The first reported cloning of H. pylori urease genes was by Clayton et al. (1989a). This group detected 66 and 31 kDa antigens with antiserum raised against the purified H. pylori urease. The cloned sequence encoded two polypeptides, UreA and UreB, which had molecular masses of 26.7 and 60.5 kDa respectively (Clayton et al., 1989b). Labigne et al. (1991) cloned urease genes by a different strategy. They identified a 44 kb portion of the H. pylori genome by cosmid cloning, which, when transferred to C. jejuni, enabled temporary synthesis of urease. Subcloning localized the H. pylori urease gene cluster to a 4.2 kb region of DNA which consisted of four open reading frames. These were in the order ureCDAB and the translated products had predicted molecular masses of 49.2, 15.0, 26.5 and 61.6 kDa respectively. The UreA and UreB polypeptides correspond to the two structural subunits of the urease enzyme and, based on this, the H. pylori urease may be more closely related to jack bean urease than to the three subunit ureases normally found in other bacteria. The functions of UreC and UreD were originally unclear, but as UreD has typical transmembrane features it was originally suggested to transport or anchor the enzyme (Labigne et al.,

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1991). However, in their extensive review of microbial ureases, Mobley rt al. (1995a) noted that no other bacterial urease gene clusters contain homologues of ureC or ureD. Recently, it has been established that UreC encodes a phosphoglucosamine mutase, involved in peptidoglycan synthesis (deReuse el al., 1997), and the gene has been renamed glmM. Expression of H . pylori urease activity in E. coli was only possible after the discovery of a contiguous (but probably separately transcribed) 3.3 kb DNA region just downstream of ureAB, which contained five open reading frames (urelEFGH) that encoded additional accessory proteins (Cussac e / al., 1992). Of those genes identified, ureA, ureB, ureF, ureG and urrH were shown to be required for expression of urease activity in E. coli, as mutations in each of these genes led to urease-negative phenotypes. These accessory genes are required for functions related to the assembly of the nickel metallocentre within the enzyme and for its activation. An important requirement for urease activity in vivo is clearly an adequate supply of nickel ions for incorporation into the apoenzyme. In the gastric mucosa, nickel and other divalent cations are likely to be tightly bound by a variety of proteins, and the question arises as to how H . pylori manages to obtain sufficient Ni2+ to sustain the level of urease activity which it exhibits. Mobley et ul. (1995b) identified an H . pylori gene (nixA) which significantly enhanced urease activity in E. coli clones carrying the H . pylori urease gene cluster. This gene encoded a 34 kDa protein with the characteristics of an integral membrane transport protein, and was shown to confer upon E. coli a high-affinity nickel transport activity (KT of 1 1 nM NiCI2). NixA is homologous to the HoxN nickel transporter from Alcaligenes eutrophus and its predicted topology suggests seven transmembrane helices. Insertional inactivation of nixA in H . pylori by allelic exchange mutagenesis resulted in a 42% reduction in urease activity and a significant reduction in nickel transport activity (Bauerfiend et al., 1996), indicating the presence of an additional transporter(s). There is evidence that a P-type ATPase is involved in urease activity, which could have nickel transport activity (Melchers et al., 1996). Recently, a binding protein-dependent ABC-transporter has also been implicated in urease activity (Hendricks and Mobley, 1997) but mutations in this system, although dramatically reducing urease activity (especially in combination with a nixA mutation), did not result in a measurable reduction in nickel transport, and the substrate for this system is currently unknown. Mechanisms for acquiring and processing nickel in H . pylori are clearly complex and well-developed. A heat shock protein (HSP) is closely associated with H . pylori urease, both in crude preparations and after gel filtration (Evans e l al., 1992). This 62 kDa protein (HspB) can be separated from the urease by ion-exchange chromatography and was found to belong to the Hsp60 family of stress

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proteins, with similarity to GroEL. H . pylori also synthesizes a GroES homologue (HspA), which has an interesting structure and relationship to urease activity (Kansau et al., 1996). In addition to a conventional N-terminal domain homologous with other GroES proteins, HspA also contains an additional 27-residue C-terminal domain which is rich in histidines and which was shown to be capable of binding nickel ions with high affinity and specificity. Co-expression of HspA in E. coli with the urease gene cluster led to a 4-fold increase in urease activity, indicating a specific function in urease assembly (Kansau et al., 1996).

9. CONCLUDING REMARKS

From a microbial physiologist’s viewpoint, Helicobacter pylori is a fascinating organism. It has a mixture of aerobic and anaerobic physiologies which combine to produce a microaerophilic phenotype, but the molecular basis for this is now only beginning to be understood. There are still a large number of unanswered questions regarding the metabolism of this bacterium, despite the publication of the genome sequence of strain 26695. Indeed, in many ways this has highlighted some important paradoxes and conflicts with experimental data which suggest future directions for research. For example, the biochemical basis for the requirement for COz has not been completely explained and a major surprise is the apparent lack of carboxylation enzymes. Although genes for glycolytic enzymes are present, a number of physiological studies indicate that the Entner-Doudoroff and pentose phosphate pathways are more active physiologically. The respiratory chain is remarkably simple and it is of interest that NADPH appears to be the preferred electron donor, rather than NADH as in most other bacteria. The fastidious nature of H . pylori can be understood to a large extent in terms of the absence of some key biosynthetic enzymes and the incomplete citric acid cycle and this is compensated for by the significant number of specific transport systems which enable the bacterium to satisfy its nutritional requirements from the host. Another remarkable feature is the relatively small number of regulatory and DNA binding proteins in H . pylori, especially the two-component ‘sensor-regulator’ systems, which indicates a minimal degree of environmentally responsive gene expression. Perhaps many of these features are related to the close relationship between H . pylori and its host and its apparent inability to grow in the environment. Future work on the physiology and metabolism of H . pylori will doubtless lead to deeper insights into this relationship.

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ACKNOWLEDGEMENTS

I would like to acknowledge the help of Alistair Davison in writing this review and Peter Chalk, Peter White, Chris Clayton, Nicky Hughes, Colin Jackson, Jess Alderson, Marc Pittman and Jyoti Velayudhan for useful discussions. I also acknowledge the financial support of Glaxo-Wellcome, The UK Biotechnology and Biological Sciences Research Council and the Wellcome Trust.

NOTE ADDED IN PROOF Since this review was written, many papers relevant to the physiology of H . pylori have been published. The fumarate reductase operon has been characterised in detail, and anfLdA mutant was found to be viable but exhibited a prolonged lag phase (Ge, Z., Jiang, Q., Kalisiak, M.S. and Taylor, D.E. ( 1 997)). Cloning and functional characterisation of Helicobacterpylori fumarate reductase operon comprising three structural genes coding for subunits C, A and B (Gene 204, 227-234). A de novo purine synthesis pathway has been demonstrated (Mendz, G.L., Shepley, A.J., Hazell, S.L. and Smith M.A. (1997) Purine metabolism and microaerophily of Helicobacter pylori. Arch. Microbiol. 168, 448456). The expression of ferric iron reductase activity and its dependence on riboflavin synthesis have been established as important factors in iron acquisition (Worst, D.J., Gerrits, M.M., Vandenbrouke-Grauls, C.M.J.E. and Kusters, J.G. (1998) Helicobacter pylori ribBA-mediated riboflavin production is involved in iron acquisition. J . Bacteriol. 180, 1473-1479).

REFERENCES Alderson, J., Clayton, C.L. and Kelly, D.J. (1996) Respiration in Helicohacter pylori is carried out by at least two-terminal oxidases, one of which is a novel haem-copper oxidase of the ch-type. Gut 39 (S2), A68. Alderson, J . , Clayton, C.L. and Kelly, D.J. (1997) Investigations into the aerobic respiratory chain of Helicohucter pylori. Guf 41 (SI) A7. Andersen, L.P., Blom, J . and Nielsen, H. (1993) Survival and ultrastructural changes of Helicohacrer pylori after phagocytosis by human polymorphonuclear leukocytes and monocytes. Acta puihologicu, Microhiologica e f Immunologica Scandinavica 101,6 1-72. Baer, W., Koopman, H. and Wagner, S. (1993) Effects of substances inhibiting or uncoupling respiratory-chain phosphorylation of Helicohacter pylori. Zbl. Bakt. 280, 253-258.

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Genes Involved in the Formation and Assembly of Rhizobial Cytochromes and their Role in Symbiotic Nitrogen Fixation Maria J. Delgadol, Eulogio J. Bedmar' and J. Allan Downie'

' Departamento de Microhiologia del Suelo y Sistemas Simbioricos, Estacidn Experimental del Zaidin, CSIC, P.O. Box 419, 18080-Granada, Spain 'Department of Genetics. John Innes Centre, Norwich Research Park, Colney, Norwich, NR4 7 U H , U K

ABSTRACT

Rhizobia fix nitrogen in a symbiotic association with leguminous plants and this occurs in nodules. A low-oxygen environment is needed for nitrogen fixation, which paradoxically has a requirement for rapid respiration to produce ATP. These conflicting demands are met by control of oxygen flux and production of leghaemoglobin (an oxygen carrier) by the plant, coupled with the expression of a high-affinity oxidase by the nodule bacteria (bacteroids). Many of the bacterial genes encoding cytochrome synthesis and assembly have been identified in a variety of rhizobial strains. Nitrogen-fixing bacteroids use a cytochrome cbb3-type oxidase encoded by the fixNOQP operon; electron transfer to this high-affinity oxidase is via the cytochrome bcl complex. During free-living growth, electron transport from the cytochrome bcl complex to cytochrome aa3 occurs via a transmembrane cytochrome c (CycM). In some rhizobia (such as Bradyrhizobium japonicum) there is a second cytochrome oxidase that also requires electron transport via the cytochrome bc, complex. In parallel with these cytochrome c oxidases there are quinol oxidases that are expressed during free-living growth. A cytochrome bb3 quinol oxidase is thought to be present in B. japonicum; in Rhizobium leguminosarum, Rhizobium etli and Azorhizobium caulinodans cytochrome d-type oxidases have been ADVANCES IN MICROBIAL PHYSIOLOGY VOL 40 ISBN 0-12-027740-9

Copyright (01998 Academic Press All rights of reproduction in any form reserved

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identified . Spectroscopic data suggest the presence of a cytochrome o-type oxidase in several rhizobia. although the absence of haem 0 in B . japonicurn may indicate that the absorption attributed to cytochrome o could be due to a high-spin cytochrome b in a cytochrome bh,. type oxidase. In some rhizobia. mutation of genes involved in cytochrome c assembly does not strongly affect growth. presumably because the bacteria utilize the cytochrome cindependent quinol oxidases. In this review. we outline the work on various rhizobial mutants affected in different components of the electron transport pathways. and the effects of these mutations on symbiotic nitrogen fixation and free-living growth .

1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 193 195 2 Haem-copper respiratory oxidases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 197 2.1. The haem-CuB bimetallic centre . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 198 2.2. The CuA redox centre . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 198 3 . Respiratory chains of free-living rhizobia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 199 3.1. The cytochrome bcl-aa3 respiratory chain ......................... 3.2. Identification and characterization of the fbcFH genes encoding the B . japonicum cytochrome bel complex ........................... 199 3.3. R . leguminosarum biovar viciae cytochrome bc, complex . . . . . . . . . . . . . 201 201 3.4. B . japonicum cytochrome CycM ................................. 3.5. R . leguminosarum biovar viciae cytochrome CycM . . . . . . . . . . . . . . . . . . . 202 3.6. Identification and characterization of coxA encoding the B . japonicum aa3-type cytochrome c oxidase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 203 204 3.7. R . tropicicoxA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 204 3.8. Regulation of coxA expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 205 4 . Other terminal oxidases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 205 4.1. B . japonicum CoxMNOP oxidase ................................ 206 4.2. B . japonicum CoxWXYZ oxidase ................................ 4.3. Non-cytochrome-containing branch of the respiratory chain . . . . . . . . . . . 207 4.4. Cytochrome d its involvement in free-living microaerobic respiration . . . 208 4.5. A . caulinodans multiple oxidases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 208 5. Symbiosis-specific cytochromes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 209 5.1. B . japonicum fixNOQP genes encoding a cytochrome cbb3-type oxidase involved in symbiotic nitrogen fixation ..................... 210 5.2. Biochemical characterization of the B. japonicum cytochrome cbb3-type oxidase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 212 5.3. Topological model of the cytochrome ebb3 subunit I. . . . . . . . . . . . . . . . . 213 6. Other fixNOQP genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 214 6.1. A . caulinodans fixNOQP genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 214 215 6.2. A . tumefaciens fixNOQP genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.3. R . leguminosarum biovar viciae symbiosis-specific oxidase . . . . . . . . . . . . 215 216 6.4. The fixNOQP-related fixGHIS operon . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 217 6.5. Regulation of the fixNOQP and fixGHlS operons . . . 218 7 . Genes involved in cytochrome c biogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . .

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8. Rhizobial mutants with altered oxidase activity and improved symbiotic nitrogen fixation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 221 Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 222 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 222

1. INTRODUCTION

The genera Rhizobium, Bra&,rhizobium and Azor..izobium, collectively referred to as rhizobia, are members of the bacterial family Rhizobiaceae of the a subdivision of the Proteobacteria (Elkan and Bunn, 1992; Young, 1992). They are Gram-negative soil bacteria with the unique ability to establish a nitrogen-fixing symbiosis on legume roots, and on the stems of some aquatic legumes. As free-living cells, rhizobia are aerobic microorganisms that utilize O2 as the final electron acceptor of the respiratory chain for the generation of ATP. Electrons are fed into the quinone/quinol pool via various dehydrogenases and passed down an electron transport chain to oxidases that reduce oxygen to water as the terminal step in aerobic respiration. Whereas the oxygen concentration in air-saturated water is about 250 p ~ , in nodules it is extremely low, ranging from 3 to 22 nM (Witty and Minchin, 1990; Hunt and Layzell, 1993). Under these microaerobic conditions, the endosymbiotic rhizobia, the so-called bacteroids, depend on respiratory ATP for their maintenance and for the energy-demanding process of atmospheric dinitrogen (Nz) fixation. At least 16 molecules of ATP are required to reduce one molecule of NZ. Microaerobiosis is essential for the function of nitrogenase, which is irreversibly inactivated by oxygen. Thus, bacteroids must be maintained in a low-O2 environment to prevent nitrogenase inactivation, but supplied with a high 0 2 flux to generate metabolic energy. The combined effects of specialized plant cells acting as an oxygen diffusion barrier and an abundant nodule protein, leghaemoglobin, which reversibly binds oxygen, result in a very low concentration of free oxygen within the infected nodule tissue (Layzell et a f . ,1993). Bacteroids develop a specialized branch of the respiratory system terminated by a high-affinity oxidase that guarantees optimal utilization of the extremely low oxygen concentration within the infected plant cells. In contrast to the single respiratory chain present in most eukaryotic mitochondria, rhizobia - like all aerobic bacterial species examined (Poole, 1983; Anraku, 1988) - have branched electron transport chains terminating with oxidases that have different affinities for oxygen. The different respiratory oxidases allow the cells to customize their respiratory systems to meet the demands of the (often rapidly changing) oxygen

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5

P

3 ‘ n

cytc

1

1

AEROBIC RESPIRATORY CHAINS

6

CycM

1 1

SY MBIOSISSPECIFIC,

MICROAEROBIC RESPIRATORY CHAINS

Figure 1 Model of branched respiratory chains showing the known or proposed terminal oxidases in rhizobia. During free-living growth, the aa3-type cytochrome c oxidase ( I ) has been genetically characterized in B. ,juponicum (Bott e f al., 1990; Gabel and Maier, 1990). R. tropici (Gabel et al., 1994). A. caulinodans (Mandon et al., 1994). and detected spectroscopically in R. leguminosarum biovar viciae (Kretovich e f ul., 1973; Delgado e f ul., 1995) and R. frifolii (De Hollander and Stouthamer, 1980). The cytochromes CoxMNOP (2) and CoxWXYZ (3) have been genetically identified in B. japonicum and thought to be a cytochrome c oxidase (Bott el ul., 1992) and a quinol oxidase of the hh3 type (Surpin er al., 1994, 1996),respectively. The presence ofcytochrome o has been observed spectroscopically in B. juponicum (Appleby, 1969b). A. cuulinodans (Stam et al., 1984). R. efli (Soberon e f al., 1989), R. leguminosarum biovar viciae (Kretovich e f al., 1973; Delgado er al., 1995) and many other rhizobial species (Chakrabarti rt d., 1987)although in view of the work of Surpin ef ul. (1996) it is evident that, in B. juponicum, the signal attributed to cytochrome o is actually caused by a bb3 quinol oxidase. This might also be the case in other rhizobia. A non-cytochrome-containingbranch of the respiratory

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concentration in the environment. The biogenesis of respiratory cytochromes in bacteria has recently been reviewed by Thony-Meyer (1997). In this review, we summarize the present knowledge of the biochemistry and molecular genetics of the respiratory pathways present in both freeliving and symbiotic rhizobia (Fig. 1). The characterization of mutants (Table 1) affected in components of different electron transport chains has helped elucidate the sequential steps of electron transfer in the respiratory chains. In view of the recent review of haem biosynthesis in the Rhizobiumlegume symbiosis (O’Brian, 1996), we have not included this aspect of cytochrome biosynthesis in this review [see also Chauhan et af. (1997), Chauhan and O’Brian (1 997), King and O’Brian (1 997) and references cited therein for recent work on haem biosynthesis and the roles of metals and oxygen in its regulation].

2. HAEM-COPPER RESPIRATORY OXIDASES

Respiratory oxidases, including the bacterial oxidases, contribute about 90% of the total reduction of molecular oxygen in the biosphere (Slater et af.,1965). Two kinds of homologous respiratory oxidases have been found in bacteria (Poole, 1988). The quinol oxidases take electrons from the quinol pool, whereas the cytochrome c oxidases receive electrons from cytochrome c that had been reduced by ubiquinokytochrome c oxidoreductase, also called the cytochrome bc, complex (Trumpower, 1990). During the reduction of molecular oxygen to water, the free energy available from the process is used to generate a transmembrane proton eletrochemical gradient, or protonmotive force, which, in turn, is used for ATP synthesis. The cytochrome c oxidases are more efficient, i.e. they generate a larger protonmotive force per electron equivalent transported compared with the quinol oxidases. pathway (5) has been described in B. japonicum (Frustaci et a/., 1991) and R. tropic; (Barquera et a/., 1991b). Cytochrome d (6) has been implicated as terminal oxidase during free-living, microaerobic growth of R. leguminosarum biovar viciae (Kretovich et a/., 1973; Wu ef a/., 1996), R. trifolii (De Hollander and Stouthamer, 1980), A . caulinodans (Stam et a/., 1984; Kitts and Ludwig, 1994) and R. etli (Barquera er a/., 1991a). The symbiosisspecific chh3-type haem-copper cytochrome c oxidase (7) has been characterized genetically and biochemically in B. japonicum (Preisig et a/., 1993, 1996a,b). and thefixNOQP genes, encoding the cbb3-type oxidase, have been identified in R. me/iloti (Batut et a / . , 1989). A . caulinodans (Mandon el al., 1994), A . tumefaciens (Schliiter et al., 1995), and R. leguminosarum biovar viciae (Schluter ct a/., 1997); both the cbb3-type oxidase (6) and the quinol oxidase d (7) are present in bacteroids of A . caulinodans (Kaminski el a/., 1996).

Table 1 Rhizobial genes and their functions in free-living respiration. Genes or operons

Known or proposed function of gene products

cycA cjcB cycc cydAB cycM

Cytochrome c550 Cytochrome c552 Cytochrome cSs5 Cytochrome d Membrane-bound 20 kDa Membrane-bound 23 kDa cytochrome c Subunit I of aa3-type cytochrome c oxidase

coxA

coxMNOP cox WXYZ

fbcFH fbcBC

Subunits 11, I, IIIa and IIIb of haem/copper cytochrome c oxidase Ubiquinol oxidase of the bb3-type Ubiquinol-cytochrome c oxidoreductase (cytochrome bc, complex) Cytochromes b and c ,

Species

B. japonicum B. japonicum B. japonicum A . caulinodans B. japonicum R. leguminosanun biovar viciae B. japonicum

Symbiotic phenotype

References

Fix' Fix+ Fix+ Fix+/Fix+ Fix+

Bott et al. (1995) Rossbach et al. (1991) Tully et a/. (1991) Kitts and Ludwig (1994) Bott et al. (1991) Wu et al. (1996)

Fix+

A . caulinodans R. tropici B. japonicum

Fix+ Fix+

Gabel and Maier ( 1990) Bott et al. (1990) Kitts and Ludwig (1994) Gabel and Maier (1994) Bott et al. (1992)

B. japonicum B. japonicum

Fix+ Fix-

Surpin et al. (1994, 1996) Thony-Meyer et al. (1989)

R. leguminosarum

Fix-

Wu et al. (1996)

biovar viciae

Fix+

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Most bacterial oxidases contain homologues of the three subunits I, I1 and I11 of mitochondrial cytochrome aa3. Some exceptions are the ba3-type cytochrome c oxidase from Thermus thermophilus (Zimmermann et al., 1988) and the aa3-type quinol oxidase from Sulfolobus acidocaldarius (Lubben et al., 1992), which lack subunit 111, and the ebb3-type oxidases (Garcia-Horsman et al., 1994a; Gray et al., 1994), which lack both subunits I1 and 111. Homologues of the 10 other subunits of the eukaryotic oxidases have not been found in bacteria, but some of the bacterial oxidases contain an additional subunit IV which is unrelated to any eukaryotic gene product (Ishizuka et ul., 1990). Despite differences in their reductants, oxygen affinities, haem types and metal compositions, most bacterial oxidases are now recognized to be related members of a single family called the haemxopper oxidase superfamily (for reviews see Garcia-Horsman e f al., 1994b; Gray et al., 1994; van der Oost et al., 1994).

2.1. The Haem-CuBBimetallic Centre

A common structural feature of the oxidase superfamily is the active site where oxygen is reduced. It is a bimetallic centre, also called the binuclear centre, formed by the ifon of a pentacoordinated haem and a copper called CuBwhich is within 5 A of the haem (Babcock and Wikstrom, 1992). These two metals are bound to subunit I via four conserved histidines. The haem of the binuclear centre is high-spin; it is coordinated to a single histidine and has an available coordination position providing the oxygen-binding site. Subunit I also binds another low-spin haem that is responsible for most of the absorption in the visible region of the spectrum and whose function is to facilitate the transfer of electrons to the binuclear centre; this haem is axially coordinated to two histidine residues (Brown et al., 1993; Hosler et al., 1993). Subunit I is the one common feature in the superfamily of haemxopper respiratory oxidases. Since the two haems associated with subunit I may be the same or different, several combinations may occur in the two subunit I haems; hence, the great variation observed in the haemxopper oxidases. The use of the subscript ‘3’ for the haem A associated with subunit I is maintained in the oxidase nomenclature because of historical reasons; it served to differentiate the ligand-binding (high-spin) haem A in the binuclear centre of the mitochondrial oxidase from the low-spin haem A. Oxidase nomenclature by haem types, however, does not indicate the oxidase affinity for its substrate nor the class (ubiquinol or cytochrome c oxidase) to which the oxidase belongs. By analogy with cytochrome aa3 many groups refer to the oxidase containing B haems as cytochrome bb3 (or ebb3) to denote the presence of high- and low-spin haem B components. As pointed out by

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Poole and Chance (1995). this nomenclature is problematic and they suggest the use of the term cytochrome hh' instead. However, since most of the groups working with rhizobial cytochromes refer to cytochrome hh3, we follow that nomenclature to avoid confusion. 2.2. The CuA Redox Centre

With the exception of the chh3-type oxidases, all of the cytochrome oxidases contain a subunit I1 where a second copper-containing redox centre, called CuA, is located. Residues in subunit I1 that are conserved in all these cytochrome oxidases have been implicated in either the binding of cytochrome c or the ligation of CuA (Capaldi, 1990; Saraste, 1990). CuA is the primary acceptor of electrons coming from cytochrome c (Taha and FergusonMiller, 1992; Hill, 1993). is located in the membrane-exposed part of subunit I1 and has two copper atoms in a mixed valence configuration (van der Oost et a/., 1992; Kelly et a / . , 1993; Lappalainen et al., 1993; Malmstrom and Aasa, 1993; von Wachenfeldt rt a/., 1994). The amino acid sequences of subunit I in cytochrome c oxidases share significant similarities with those of subunit I from ubiquinol oxidases (Chepuri et a / . , 1990; Saraste et a / . , 1991). In the latter class. a clear distinction is the absence of residues implicated in binding either cytochrome c or CUA(Chepuri r ~ ul., t 1990; Saraste et a/., 1991; Santana et al., 1992); subunits I1 of all quinol oxidases studied so far lack the CuA redox centre (Lauraeus et al., 1991; Minghetti et a/., 1992; Fukaya et a/., 1993). In most cases analysed, subunit I1 has two transmembrane-spanning helices and a large hydrophilic domain that faces the bacterial periplasm (Capaldi, 1990; Saraste, 1990). Besides providing the substrate-binding site either for quinol or cytochrome L', the CuA centre in subunit I1 transfers electrons directly to the haem A of subunit I, and then to the binuclear haem-CuB centre, where O2 is reduced (Ramirez et al., 1995).

3. RESPIRATORY CHAINS OF FREE-LIVING RHlZOBlA

Appleby and Bergersen (1958) first showed that the haemoprotein pattern of aerobically cultured Bradyrhizobium juponicum cells differs from that of root-nodule bacteroids. Photochemical action spectra indicated that oxidases of the u q - and o-types were expressed in aerobically grown cells but were absent in bacteroids (Appleby, 1969a). Recently, Surpin et al. (1996) analysed the haems present in B. japonicuni and concluded that haem 0 was not present. The spectroscopic properties of cytochrome o cannot readily be

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distinguished from those of a high-spin h-type cytochrome (Poole, 1994). Therefore, although there are many reports of cytochrome o being present in rhizobia (on the basis of spectroscopic analyses), there is a possibility that the absorption peaks could in fact correspond to a high-spin b-type cytochrome. possibly as part of a bb3 complex (see Surpin et af., 1996). In the absence of data on the presence or absence of haem 0 in other rhizobia, in this review we will continue to refer to ‘cytochrome 0 ’ as described by others, while recognizing that it could be a high-spin cytochrome h. Cytochromes aa3, h and c were seen in spectra from Rhizohium leguminosarum biovars viciae and rrifolii (Kretovich et ul., 1973; De Hollander and Stouthamer, 1980; Vargas er ul., 1994) and Delgado er ul. (1995) noted the presence of a CO-binding component with the characteristics of cytochrome o or a high-spin cytochrome h. Azorhizobium caulinoduns actively fixing N2 in continuous culture has a spectrum typical of cytochromes m3,d and o (Stam er d., 1984). Cytochrome ua3 generally terminates a low-oxygenaffinity branch of the aerobic electron transport chains, while cytochrome o usually terminates a high-oxygen-affinity branch (O’Brian and Maier, 1985). Cytochromes b, c, aa3 and o were also seen in difference spectra of crude cell-free extracts in each of the 22 aerobically grown fast- and slow-growing rhizobia examined by Chakrabarti er uI. (1987). In cells of Rhizohium erli (formerly Rhizohium leguminosurum biovar phaseoli type I) growing under well-aerated conditions, oxidases o and au3, in addition to b-type and c-type cytochromes, were identified by photodissociation spectra and oxygen binding (Soberon et al., 1989).

3.1. The Cytochrome bc,-aa3 Respiratory Chain

The presence of an au3-type oxidase in aerobically grown cells of different rhizobia suggested the existence of an electron transport pathway similar to that occurring in mitochondria and many aerobic bacteria:

2[H] -+ Q -+ Fe-S/hcl

.+

Cytc + aa3 .+ O2

This was demonstrated after identification, isolation and mutagenesis of genes encoding each of the components of the chain.

3.2. Identification and Characterization of the fbcFH Genes Encoding the B. japonicum Cytochrome bc, Complex A B. japonicum mutant (Regensburger et al., 1986) unable to fix nitrogen was found to be unable to oxidize ‘Nadi’ reagent in a standard cytochrome

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oxidase assay on plates. This test is dependent on oxidation of tetra- or dimethyl-phenyl-diamine (TMPD or DMPD) via cytochromes cl, c and aa+ Cloning and sequencing of the wild-type DNA fragment corresponding to the Tn5-containing DNA in the mutant identified two adjacent genes, fbcF and fbcH, arranged in one operon (Thony-Meyer el al., 1989). fbcF encodes a Rieske iron-sulphur protein of 176 amino acids. Four conserved cysteines in the C-terminal part of the protein are believed to serve as the ligands for the catalytically active 2Fe-2S centre of the protein, although two histidine residues could also act as potential Fe ligands. A stretch of hydrophobic amino acids in the N-terminus of the protein was predicted to form a transmembrane helix, that might act as a membrane anchor. ThefbcH gene product was predicted to be 687 amino acids long. The Nterminal domain resembles cytochrome h from different organisms, and the C-terminal domain of about 250 amino acids, is similar to cytochrome c. In the N-terminal domain of FbcH there are four histidines, that are predicted to be haem-iron-binding ligands for the two non-covalently bound B-type haems in the active centre of the protein. Other residues predicted to be part of the active site are three proline residues and eight glycine residues; two of the prolines and six of the glycines are located in potential membranespanning domains. The C-terminal half of the protein contains the sequence Cys-Ala-Ser-Cys-His, which is characteristic of a c-type haem binding site (Cys-X-X-Cys-His); the cysteines form the covalent thioether bonds to one C-type haem and the histidine provides the fifth ligand for the haem iron. A methionine, thought to be the sixth ligand of the haem iron, is also present. Of the 11 transmembrane domains predicted in FbcH, domain XI is located in the C-terminus and is believed to function as a membrane anchor for the hydrophilic cytochrome c. In contrast to the three genes, fbcF, fbcB and fbcC that encode the hcl complex found in mitochondria and many aerobic and photosynthetic bacteria, ,fhcF and fbcH alone encode the entire bcl complex of B. juponicum (Thony-Meyer et al., 1989). ThefbcH gene encodes a precursor that is posttranslationally processed into the two individual cytochromes. A protease, recognizing the site Ala-Arg-Ala in a hydrophobic domain of about 30 amino acids connecting the N-terminal and C-terminal sequences of the FbcH protein, could be responsible for the cleavage of the large FbcH precursor (Thony-Meyer et al., 1991). A C-haem membrane protein corresponding to uncleaved FbcH was not detected in the wild-type strain; two C haem-staining bands with apparent M , of 28 000 and 20 000 were found. Both were missing in membrane preparations from the fhcH mutant. (The M , 20 000 component was subsequently shown to be the membrane-bound cytochrome c, CycM, that was not correctly assembled in the absence of cytochrome c1, see below). Since the haem-stained 28 kDa protein cross-

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20 1

reacted with cytochrome c1 antiserum it was concluded that this corresponds to cytochrome cI processed from a larger FbcH precursor (ThonyMeyer et ul., 1989). This example of a single gene coding for two or more functional proteins after post-translational cleavage is unusual in prokaryotes. B. juponicum fbcF and fbcH mutants are unable to fix nitrogen in freeliving cultures and induce Fix- nodules in soybeans, demonstrating that the cytochrome bc, complex is essential for a N2-fixing symbiosis (Thony-Meyer et ul., 1989). 3.3. R. leguminosarum biovar wiciae Cytochrome bcl Complex

Membranes from R. leguminosarum biovar viciae contain two C-haem proteins of M , 3 1 000 and 23 000 (Vargas et al., 1994; Delgado et al., 1995). Two Tn5-induced mutants were identified (Wu et al., 1996), which express lowered levels of c-type cytochromes and cytochrome au3, but increased levels of cytochrome d. Membrane fractions from both mutants lacked haemstaining proteins, but the soluble fraction contained, in addition to the expected protein of M , 14 000, a haem-stained band at M , 23 000 that was not present in the wild-type strain. Molecular analysis of mutants revealed that one contained Tn5 in a cytochrome c I gene, whereas the other had Tn5 in a gene encoding the cytochrome b component of the cytochrome bcl complex. Two ORFs encoding cytochromes b and c1 were found separated by an intergenic region of 28 nucleotides. The predicted sequence of the cytochrome c1 gene product has a typical N-terminal transit peptide, which suggests that the protein is secreted to the periplasm. Thus, in contrast to the situation with B. japonicum, there are two genes coding for cytochrome b and cytochrome c1 in R. leguminosarum biovar viciae. Because of the similarities with the Rhodopseudomonas sphaeroides fbcB and fhcC genes, the genes identified in R. leguminosarum biovar viciae were called fbcB and fbcC (Wu et al., 1996). 3.4. B. japonicum Cytochrome CycM

The isolation and characterization of Tn5-induced mutants unable to oxidize TMPD provided evidence for the involvement of a membrane-bound cytochrome c in respiration of aerobically grown B. japonicum. Analysis of c-type cytochromes by haem-staining in membrane preparations from one mutant revealed the absence of a 20- kDa component, while the 28 kDa cytochrome c1 and all the soluble c-type cytochromes (~550,~ 5 5 2and ~ 5 5 5 ,

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see below) were still present (Bott e f al., 1990).The gene affected was named cycM, and has been shown to be the structural gene for the membraneanchored 20 kDa c-type cytochrome homologue of the soluble mitochondrial cytochrome c (Bott et al., 1991). The apoprotein was predicted to be 184 amino acids long. A hydrophobic domain at the N-terminal end forms a transmembrane helix that acts as a translocation signal and a membraneanchor. The haem-binding site motif Cys-Gly-Ala-Cys-His identified the protein as a monohaem c-type cytochrome that is most probably involved in electron transfer from the cytochrome bel complex to the aa3-type terminal oxidase (Bott et al., 1991).This cytochrome c (CycM) may interact with two acidic domains in cytochrome c I analogous to soluble cytochrome c binding domains found in beef heart cytochrome el (Broger et al., 1983; Stonehuerner et al., 1985), and conserved in the FbcH protein of B. japonicum (Thony-Meyer et al., 1989). The cycM mutant lacks cytochrome aa3 in addition to the CycM protein. Since CycM is also absent from fbcH mutants, it was suggested that the cytochrome bel complex, the CycM protein, and cytochrome aa3 form a complex whose assembly in the membrane occurs only in a strictly unidirectional order: bcl-CycM-aa3.The incorporation of cytochrome aa3 into the complex is proposed to be dependent on the incorporation of CycM, which in turn requires assembly of the cytochrome bcl complex in the membrane (Bott et al., 1991). This differs somewhat from observations made with R . leguminosarum biovar viciae; although a cycM mutant lacked cytochrome aa3, cytochrome bcl mutants were found to have spectroscopically detectable cytochrome aa3 (Wu et al., 1996).

3.5. R. legurnhoserum biovar wiciae Cytochrorne CycM

A R . leguminosarum biovar viciae mutant was identified that was specifically blocked in the formation of a membrane bound c-type cytochrome of M , 23 000, but retained both the membrane-bound M , 31 000 cytochrome cI and the soluble M , 14 000 haem-C proteins. DNA sequencing from the end of the Tn5 that caused the mutation revealed it to be in a gene that shows 65% identity with the B. juponicum cycM gene encoding the 20 kDa membrane-bound c-type cytochrome; this suggested that the gene encodes a homologue of CycM (Wu et al., 1996). The R . leguminosarum biovar viciae cycM mutant had a normal symbiotic phenotype whereas thefhcB and.fhcC mutants were blocked for symbiotic nitrogen fixation. This correlates with the observation in B. japonicum that the cytochrome hcl complex is necessary for symbiotic nitrogen fixation, but CycM is not (Thony-Meyer et al., 1989; Bott et al., 1991).

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3.6. Identification and Characterization of coxA Encoding the B. jeponicum 883-wpe Cytochrome c Oxidase

Mutants of B. japonicum defective in cytochrome aa3 have been generated by chemical and Tn5 mutagenesis (El Mokadem and Keister, 1982; O’Brian el al., 1987; Nautiyal et al., 1989; Bott et al., 1990). DNA sequencing of complementing plasmids revealed a predicted hydrophobic protein of 541 residues with extensive similarities to mitochondria1 and prokaryotic cytochrome aa3 sequences, including 74% similarity with Saccharomyces cerevisiae cytochrome c oxidase subunit I (O’Brian and Maier, 1987; Bott et al., 1990; Gabel and Maier, 1990; Maier et al., 1990). The gene was named coxA in view of its similarity to subunit I of cytochrome aa3. The most important feature in the amino acid sequence of CoxA is the conservation of six histidines, two of which are predicted to act as ligands for the haem A, one for the high-spin haem A(a3), and possibly three for CUB.A seventh conserved histidine residue is substituted for a glutamine in the B. japonicum enzyme. The transcription initiation site of the identified coxA gene is located 72 bases upstream of the proposed translation initiation site, and shows no homology with either other known B. japonicum promoters or other coxA promoters in other organisms (Gabel and Maier, 1993). Analysis of proteins with covalently bound haem in a coxA mutant showed that the soluble and membrane-bound cytochromes c are all present (Bott et al., 1990). In plant infection tests with soybeans, cycM and coxA mutants form normal nodules that are able to fix nitrogen at similar or higher levels than the wild-type strain (Bott et al., 1990). Thus, B. japonicum cells grown aerobically have a cytochrome c pathway that is not essential for symbiosis. In this mitochondria-like electron transport pathway, electrons are transferred from the quinol pool via the Rieske Fe-S protein/cytochrome hc, complex, then to a membrane-bound cytochrome c (CycM protein), and then passed to the cytochrome aa3-type terminal oxidase, the site of O2 reduction:

2[H] -+ Q -+ Fe-Slhc,

-+

CycM -+ aa,

-+

O2

A similar respiratory chain has been proposed in aerobically grown cells of R . leguminosarum biovar viciae (Wu et al., 1996) although a coxA-like gene has not yet been described in this species. An additional gene required for the assembly of a-type cytochromes is tlpA, which encodes a thioredoxin-like protein (Loferer et al., 1993). Mutation of tlpA blocked maturation of holocytochrome aa3, although the subunit I apoprotein was incorporated into the membrane. TlpA is anchored in the cytoplasmic membrane, leaving the bulk of the protein exposed to the periplasm, and has disulphide reductase activity (Loferer et

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MARIA J. DELGADO, EULOGIO J. BEDMAR AND J. ALLAN DOWNIE

al., 1993; Loferer and Hennecke, 1994). It is thought to reduce cysteine residues in a protein required for the synthesis and/or incorporation of haem A. Interestingly, mutation of tlpA blocked development of the nitrogen fixing symbiosis. Since cytochrome a03 is not required for nitrogen fixation, Loferer et ul. (1993) concluded that TlpA is involved in an independent process required for symbiotic nitrogen fixation.

3.7. R. tropici coxA The R. tropici (formerly R . leguminosarum biovar phaseoli type 11) coxA gene has also been identified and characterized (Gabel and Maier, 1990; Gabel et al., 1994). The sequenced gene encodes a polypeptide of 538 amino acids about 90% identical to the B. japonicum CoxA. The putative promoter is 5 1 bases upstream of the proposed translation initiation codon, and shows no homology with other described rhizobial promoters, including the coxA promoter of B. japonicum (Gabel et ul., 1994). 3.8. Regulation of coxA Expression

Expression of the B. juponicum coxA gene is affected by O2 (Gabel and Maier, 1993). RNA from cells grown at various 02 levels probed with the coxA gene (Gabel and Maier, 1990), showed that there is a 6-fold reduction in coxA transcription by cells grown with 1% 02,compared with fully aerobically grown cells. Although no spectrophotometrically discernible cytochrome au3 was found in bacteroids, RNA isolated from bacteroids had a coxA message which was about 19% of the level of that found with cells grown with 1% 02.It was speculated that there is a basal level of transcription in bacteroids, or that the message detected is actually due to a message still being synthesized by undifferentiated bacteria (Gabel and Maier, 1993). In contrast to B. japonicum, R. tropici shows no significant 02-mediated reduction in the level of either coxA transcription or cytochrome au3 level, even in cells incubated at 1% 02.Bacteroids isolated from bean nodules contained 65% of the fully aerobic free-living levels of the coxA transcript, indicating that R. tropici cytochrome au3 expression is regulated differently from that of B. juponicum (Gabel et al., 1994). The effects of decreased copper concentrations on the levels of this cytochrome were investigated in B. juponicum and R . tropici (Gabel et al., 1994). Both strains had the same doubling times in copper-free and copper-containing medium, and coxA transcript levels were the same for cells grown

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with or without copper. This indicates that expression of coxA is not transcriptionally regulated by copper. The fact, however, that membranes from cells of B. japonicum grown without copper lacked spectroscopically detectable cytochrome aa3 (whereas those from R . tropici retained about 50% of normal cytochrome aa3 levels) suggests a post-translational effect of copper on cytochrome aa3 expression (Gabel et a f . , 1994). In those experiments, the spectral contribution of cytochromes b and c remained unchanged between samples with and without copper, implying that copper depletion does not affect levels of these cytochromes. Whether B. japonicum does not synthesize functional or spectrally detectable cytochrome aa3, or copper is required for proper structural stability of the enzyme, is not known.

4. OTHER TERMINAL OXIDASES 4.1. B. japonicum CoxMNOP Oxidase

During identification and cloning of coxA, a B. japonicum DNA region was found that hybridized with a fragment of the subunit I gene (ctaDI) from Paracoccus denitrlficans cytochrome c oxidase. A cluster of at least four genes organized in the coxMNOP operon was identified. The predicted gene products are homologous to subunit I (CoxN), subunit I1 (CoxM) and subunit I11 (CoxO, CoxP) of haem
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MARIA J. DELGADO, EULOGIO J. BEDMAR AND J. ALLAN DOWNIE

Aerobically grown cells of a coxN mutant had TMPD terminal oxidase activity and nitrogen fixation in soybean nodules was like that of wild-type (Bott et al., 1992). It was speculated that the coxMNOP gene products may form a CuA-containing cytochrome o-type oxidase (Bott et al., 1992), but this is unlikely in the light of the more recent results of Surpin el al. (1996) who demonstrated an absence of haem 0 in B. japonicum. The prediction that coxMNOP encode a cytochrome oxidase implies that a cytochrome c is required as the reducing substrate. Three soluble c-type cytochromes, first identified in soybean bacteroids (Appleby, 1969b). termed cytochromes csSO,('552 and cSs5(formerly cSs4) because of their spectroscopic properties (Daniel and Appleby, 1972; Appleby. 1984; Appleby et al., 1991) and later isolated, purified and biochemically characterized (Appleby et d., 1991; Tully et al., 1991), were suggested as candidates for electron transfer from the cytochrome hel complex to the cytochrome CoxMNOP oxidase (Bott et al., 1992). These cytochromes are dispensable for symbiosis, because mutations in their respective genes, cycA (css0) (Bott ef al., 1995), cycB (cs5?) (Rossbach et al., 1991) and c,ycC (cS55) (Tully et al., 1991) produce strains with a Fix' phenotype in symbioses with soybeans. The following sequence of reactions for the respiratory chain containing the alternative CoxM NOP oxidase has been proposed (Bott et a/., 1992): 2[H+]

-+

U Q -+ Fe-S/hc, -+ c -+ CoxMNOP -+ O2

4.2. B. japonkurn CoxWXYZ Oxidase

Since cytochrome hel mutants are still capable of growing aerobically (Bott et al., 1990), it was speculated that an alternative. cytochrome hcl-independent respiratory pathway would be functional in aerobically grown B. .japonicum cells. In addition to coxA and coxMNOP, another gene, named coxX, was identified in B. japonicum (Surpin et al., 1994). The coxX gene was found after hybridization of PCR-amplified genomic DNA from a B. juponicum strain using probes from degenerate primers designed from the known sequences of the Escherichia coli cytochrome o subunit I and the au3-type cytochrome oxidase subunits I of PS3, P . denitriJjcans and beef heart mitochondria. coxX encodes a 666-amino acid protein with 59% identity to that of E. c d i cytochrome o subunit 1 (cyoB gene product), and a lesser degree of similarity with CoxA, CoxN and FixN ( f i x N O Q P operon, see below). c~i.uX is located within the coxWXYZ operon that is proposed to encode a hh3type ubiquinol oxidase (Surpin et al., 1996). CoxW is homologous to subunits I1 of ubiquinol oxidases including CyoA of the E. coli ho3 quinol oxidase and has a conserved ubiquinol-binding domain; CoxY is homolo-

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gous to subunits 111 of terminal oxidases including CyoC from E. coli; CoxZ is similar to subunits IV such as E. coli CyoD. A B. japonicum COXX mutant grew normally in 1% and 20% 02,but when grown in 1 YOO2 had markedly different cyanide inhibition of terminal oxidase activity compared with equivalent wild-type cells. Surpin et al. (1996) failed to detect haem 0 in wild-type cells and this, taken together with the complete absence of cytochrome aa3 in coxA mutants, led them to conclude that the co.uWXY2 operon encodes a bh3-type oxidase that probably oxidizes quinols in a pathway:

It is likely that this hh3 oxidase accounts for the cytochrome d i k e spectra that have been detected in B. ,japonicum, these actually corresponding to high-spin h cytochrome absorption spectra.

4.3. Non-cytochrome-containing Branch of the Respiratory Chain

In addition to cytochromes, flavoproteins are also involved in bacterial electron transport chains, functioning as dehydrogenases and, in some cases, as oxidases (Massey et a/., 1988). Some flavoproteins form complexes with cytochrome c, but others react with oxygen without participation of cytochromes (Appleby, 1978; Malmstrom, 1982; O'Brian and Maier, 1983). The existence of a NADH-dependent flavoprotein oxidase has been reported in free-living R. etli. This is characterized by its low affinity towards oxygen and NADH, its low sensitivity to quinacrine inhibition, and its selectivity towards ferricyanide over 2,6-dichlorophenol-indophenol (DCPIP) as artificial electron acceptors. The oxidase activity is expressed in aerobically and semi-aerobically cultured cells, but not in anaerobically cultured cells or in bacteroids (Barquera et al., 1991b). A B. japonicum mutant (Guerinot and Chelm, 1986) defective in 6-aminolevulinic (ALA) synthase, the first committed step in haem biosynthesis, could be induced to grow well in an 02-dependent manner if yeast extract was added to the growth medium. Although no haem could be detected in cell extracts as measured by absorption spectra or the peroxidase activity of the haem moiety (Frustaci et al., 1991), the mutant had rates of endogenous respiration identical to those found in the parental strain. These unusual results indicate the existence 01' an alternative, aerobic pathway composed exclusively of non-haem proteins.

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4.4. Cytochrome d: its Involvement in Free-living Microaerobic Respiration

During late stationary phase of aerobic cultures and in 02-restricted conditions, cytochrome d has been described in spectra of R . leguminosarum biovars viciae and trifolii (Kretovich et al., 1973; Vargas et al., 1994; Wu et al., 1996), R . trifolii (De Hollander and Stouthamer, 1980), A . caulinoduns (Stam et al., 1984; Kitts and Ludwig, 1994) and R. etli (Barquera et al., 1991a). In contrast, B. japonicum cells cultured under 02-restricted conditions do not synthesize cytochrome d . Difference spectra also revealed a high-spin 6-type cytochrome (hSg5)in aged, R . leguminosarum biovar viciae (Wu et al., 1996) and in microaerobically grown R . etli (Barquera et al., 1991a). There is a large increase in the amount of cytochromes d and 6595 in mutants lacking either the cytochrome bcl complex (Wu et al., 1996) or all c-type cytochromes (Vargas et al., 1994). This could result from the cytochrome d branch of the respiratory chain compensating for the loss of another branch, clearly indicating that R . leguminosarum biovar viciae can use cytochrome d as a terminal oxidase. The high affinity of cytochrome d towards O2 (Poole, 1983; Au et al., 1985) has led to the proposal that its biological role would be to enable bacteria to continue growth and respiration in low-O2 environments. With the exception of A . caulinodans, where cytochrome d appears to be active in nodules during symbiotic N2 fixation (Kaminski et al., 1996), expression of this cytochrome seems to be restricted to free-living rhizobia. 4.5. A. ceulinodenr Multiple Oxidases

Nitrogen-fixing cells of A . caulinodans contain spectroscopically-detectable cytochromes aa3, d and o (Stam et al., 1984). A second, a-type cytochrome was detected by difference spectroscopy of mutants of A . caulinodans lacking cytochrome 4a3 (Kitts and Ludwig, 1994). This alternative a-type cytochrome is also present in the wild-type, where it is probably masked by cytochrome aa3. In late exponential growth, the cytochrome aa3 mutants induced a new, membrane-bound, CO-binding cyt550, which might serve as a cytochrome c oxidase. Therefore, at least five terminal oxidases, in addition to the symbiosis-specific terminal oxidase, have been proposed to mediate respiratory electron transport to 02. Using B. japonicum coxA (O’Brian and Maier, 1987), and Azotohacter vinelandii cydAB gene probes (encoding cytochrome d oxidase; Moshiri et al., 1991), Azorhizobium coxA and cydAB gene regions have been cloned (Kitts and Ludwig, 1994). Cytochrome aa3 mutants, a cytochrome d mutant and a cytochrome aa3-d double mutant were constructed. Pleiotropic mutant strains lacking all cytochrome c activity were also isolated by screen-

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ing for strains unable to oxidize TMPD (Kitts and Ludwig, 1994). Cytochrome 4a3 mutants show normal growth rates when cultured in both rich and defined media, fix N2 at essentially wild-type rates, and are fully able to oxidize general respiratory electron donors. Cytochrome d mutants have 40% lower N2 fixation rates in culture and in planta, but aerobic growth rates are wild-type. A cytochrome aa3-d double mutant had growth rates similar to that of the wild type, but showed 70% lower N2 fixation rates in planta. Pleiotropic cytochrome c mutants excrete coproporphyrin IX to the growth medium, and are unable to utilize glutamate as carbon source. They grow normally in aerobic minimal medium, but show poor growth in rich medium and are impaired in N2 fixation, both in culture and in planta (Kitts and Ludwig, 1994). A . caulinodans JixNO-deleted mutant strains (the JixNOQP operon coding for the cbb3-type bacteroidspecific terminal oxidase, see below) show nitrogenase activity only slightly lower than that of the wild-type under free-living conditions, and still retain 50% of the activity in the symbiotic state (Mandon et al., 1994). Mutants defective in cytochrome d or cbb3 oxidases remain able to fix N2 both in culture and during symbiosis. Cytochrome d-cbb3 double mutants, while able to grow and fix N2 in aerobic cultures, are the only cytochrome oxidase-deficient mutants of A . caulinodans reported to be completely unable to fix N2 in symbiosis (Kaminski et al., 1996). This indicates that both cytochromes d and cbb3 seem to contribute as respiratory terminal oxidases in the symbiotic state. However, in O,-limited continuous cultures, cytochrome d oxidase maintains dissolved O2 at 3.6 mM in the steady state, whereas cytochrome cbb3 oxidase activity depleted O2 to submicromolar levels (Kaminski et al., 1996). Since the dissolved 0 2 level in nodules is estimated to be in the 1&20 nM range (Bergersen et al., 1986), the reason why cytochrome d is active during symbiosis is not known. Since Azorhizobium terminal oxidase mutants are able to grow under all physiological conditions tested, no single terminal oxidase seems essential for any particular Azorhizobium growth regime, including ability to fix N2 both in pure cultures and symbiosis with the host legume Sesbania rostrata. It has been suggested that this broad repertoire of terminal oxidases, although somewhat redundant as to the physiological role of each individual oxidase, might allow A . caulinodans to grow in wide-ranging oxygen environments (Kitts and Ludwig, 1994). 5. SYMBIOSIS-SPECIFIC CVTOCHROMES

A striking feature of the respiratory system of many strains of rhizobial bacteroids is the apparent absence of cytochromes aa3, ‘cytochrome 0’ and cytochrome d (Appleby, 1969b, 1984; Vargas et al., 1996). Although

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MARIA J. DELGADO, EULOGIO J. BEDMAR AND J. ALLAN DOWNIE

this situation is the general rule, there are species and specific strains that retain cytochrome a43 under symbiotic conditions (Keister and Marsh, 1985, 1990; Maier et al., 1990). Since cytochrome aa3 has an apparent 0 2 affinity of 4 7 mM (O’Brian and Maier, 1989), and is not essential for symbiosis (Bott et al., 1990), its presence under microaerobic conditions has been the subject of speculation (O’Brian and Maier, 1989), and attributed to a lack of tight 02-dependent regulation in those strains (Gabel and Maier, 1993; Gabel et al., 1994). In bacteroids of B. japonicum, soluble and membrane-bound haem proteins c550. ~552,c~~~~P-420, P-428 and P-450, and one putative flavincontaining protein, are present, and these (except the flavin protein) bind CO (for reviews, see O’Brian and Maier, 1989; Hennecke et al., 1993). A photolysable pigment, with an absorption maximum in the reduced form at about 445 nm has also been shown in B. japonicum, R . leguminosarum and R . lupini bacteroids, and suggested as a terminal oxidase in those species (Williams et al., 1990). P-420, P-428 (haemoprotein bsg0,showing peroxidase and catalase activities) and ~ 5 5 2have been shown not to be involved in electron transfer to O2 via a CO-sensitive terminal oxidase (Appleby and Poole, 1991). A B. japonicum mutant whose bacteroids contained no detectable cytochromes P-450 formed effective nodules on soybeans, suggesting that these cytochromes are not involved in an essential symbiotic function (Tully and Keister, 1993). The three soluble c-type cytochromes, ~ 5 5 0 c552 , and ~ 5 5 5 were , also shown to be dispensable for symbiosis because mutations in the respective genes produced strains which have no observable defects in symbiotic nitrogen fixation (Rossbach el al., 1991; Tully et al., 1991; Bott et al., 1995). Furthermore, a cycB-cycC double mutant, and a cycA-cycB-cycC triple mutant elicited N2-fixing nodules on soybeans. Hence, none of these cytochromes c is essential for respiration supporting symbiotic N2 fixation (Bott et al., 1995).

5.1. B. jeponicum fixN0OP Genes Encoding a Cytochrome cbk-type Oxidase Involved in Symbiotic Nitrogen Fixation

A DNA region located upstream of the previously reported genes for the oxygen-responsive two-component regulatory FixLJ system of B. japonicum (Anthamatten and Hennecke, 1991, see below) was cloned and shown by interspecies hybridization to have genes related to the so-called ‘JixN region’ of R. meliloti (Preisig et al., 1993). The R. meliloti ‘jixN region’ consists of four genes organized in the operonJixNOQP (Batut et al., 1989), and was first described as a duplicated Jix region linked to the regulatory genes.fixLJ

Table 2 Rhizobial genes involved in symbiosis-specific oxygen respiration. ~

Genes or operons

Known or proposed function of gene products

Species

Symbiotic phenotype

References ~

fixN fix0 fixP 3XQ

Subunit I, of cytochrome &-type cytochrome c oxidase monohaem cytochrome c, dihaem cytochrome c membrane-bound protein of unknown function

All rhizobia examined

FixFixFix Fix+

Batut et al. (1989) Preisig et a!. (1993) Mandon et al. (1 994) Schliiter er al. (1997)

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MARIA J. DELGADO, EULOGIO J. BEDMAR AND J. ALLAN DOWNIE

andfixK in R . meliloti (Batut et al., 1985; David et al., 1987; Renalier et al., 1987). Four open reading frames (ORFs) were identified in cloned DNA from B. japonicum showing homology to the jixNOQP region (Preisig et al., 1993). Based on the similarity with R . meliloti, four of the ORFs were named$xN,.fixO,$xP andjixQ. Analysis of the predicted protein sequences suggested that FixN (549 amino acids) is a subunit I of the haemxopper oxidases and FixN also shows about 20% identity with B. japonicum CoxA and CoxN. There is conservation of 12 membrane-spanning helices, and five of the six histidines thought to be the CuB ligands, the high-spin haem B ligand, and the low-spin haem B ligand. Fix0 (244 amino acids) and FixP (290 amino acids) are predicted to be membrane-bound mono- and dihaem c-type cytochromes, respectively, which are anchored in the cytoplasmic membrane by their hydrophobic N-termini. FixQ (54 amino acids) is thought to be membrane-bound by its hydrophobic N-terminal half. Analysis of membrane proteins from B. japonicum wild-type and mutant cells revealed that two c-type cytochromes, the likely products of the fix0 and jixP, are synthesized only under oxygen-limited growth, and that j x N andJixNO mutants exhibit a strong decrease in whole cell oxidase activity as compared with the wild-type (Preisig et al., 1993). Because of the presence of only c- and h-type cytochromes and the absence of subunit I1 carrying the CuA centre, this novel oxidase was called a chh3-type cytochrome oxidase, which is now recognized as a new subfamily of the haem-copper oxidase superfamily (Castresana et al., 1994; Garcia-Horsman et al., 1994b; van der Oost e f al., 1994). It appears to be present in all rhizobia examined (Table 2).

5.2. Biochemical Characterization of the B. jeponicum Cytochrome cbk-type Oxidase A cytochrome-c oxidase supercomplex consisting of seven to eight subunits and possessing a molecular mass of 358-425 kDa was purified from membranes of B. japonicum bacteroids (Keefe and Maier, 1993). The purified complex oxidizes exogenously added cytochrome c , and contains both hand e-type haem proteins that were shown spectrophotometrically to form complexes with CO. Two of the components of the complex have been identified as cytochromes b and e l , as they cross-react with antibodies previously raised against these two proteins from B. japonicum (Thony-Meyer et a[., 1991). Furthermore, the complex efficiently functions at free oxygen concentrations below l m ~ a;t that time, however, it was not possible to determine specific affinity of the oxidase for 02.

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The cbb3-type oxidase encoded by JixNOQP was purified 27-fold from B. japonicum cells grown anaerobically with nitrate as the final electron acceptor (Preisig et al., 1996a). The purified oxidase had TMPD oxidase activity as well as cytochrome c oxidase activity. Haem B but not haem A was detected in the haem extracts from the purified oxidase, and FixO and FixP were visualized as C-type haem proteins in polyacrylamide gels after haem staining. Difference spectra of the cbh3-type purified oxidase revealed the absence of a-type cytochromes and confirmed the presence of both b- and c-type cytochromes. Spectral characteristics of both the purified cbh3-type oxidase and the oxidase complex purified by Keefe and Maier (1993) are similar. N-terminal amino acid sequences of the three major constituents (42, 31 and 29 kDa, respectively) of the purified ebb3-type oxidase are identical to the N-termini sequences predicted from the JixN, Ji.0 and JixP gene products. No indication for the presence of the small JixQ gene product has been obtained thus far in SDS-polyacrylamide gel electrophoresis (Preisig et al., 1993, 1996a; Thony-Meyer et a/., 1995). The O2 affinities of the purified cbb3-type oxidase and membranes from anaerobically grown cells of B. japonicum wild-type, JixNOQP, and coxA mutants, were measured (Preisig et al., 1996a) using a leghaemoglobinbuffered spectrophotometric assay (Appleby and Bergersen, 1980; d’Mello et al., 1994). A coxA mutant had monophasic kinetics, with a K , value of 7 nM corresponding to the ebb3-type oxidase. In thefixNOQP mutant, a K , for O2 of 56 nM, corresponding to the aa3-type oxidase, was found. Biphasic kinetics were observed in the wild-type, with K , values of 19 and 4 nM 02, suggesting the existence of at least two oxidases with different affinities for oxygen in anaerobically grown B. japonicum cells. Taken together, the above findings clearly indicate that the chb3-type oxidase encoded by the JixNOQP operon is suited to support microaerobic respiration in endosymbiotic bacteroids (Preisig et al., 1993, 1996a; ThonyMeyer et al., 1995, Zufferey et al., 1996a) in the pathway: 2[H] +. Q +. Fe-S/bc, -+ebb3 +. O2

5.3. Topological Model of the Cytochrome c b k Subunit I

Analysis of symbiotic nitrogen fixation, TMPD- and cytochrome oxidase activities, difference spectroscopy, haem stains and Western blots of nonpolar in-frame deletion mutations in JixN, JixO and fixQ, and an insertion mutant ofJixP have shown that for assembly of the cbb,-type oxidase, FixN and FixO are first required to form a core complex, which assembles independently of FixP. FixP is then incorporated to the complex, its persistence

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MARIA J. DELGADO, EULOGIO J. BEDMAR AND J. ALLAN DOWNIE

in the membranes depending on formation of that core complex. FixQ is not essential for oxidase assembly (Thony-Meyer et al., 1995; Zufferey et a]., 1996a) and does not appear to play an essential role because a,fixQ deletion mutant has a Fix’ phenotype (Zufferey et al., 1996a). During genetic characterization of subunit I (FixN) of the chh3-type oxidase, only five (instead of the normal six) canonical histidine residues could be assigned as the putative haem or copper ligands (Preisig et al., 1993). The ambiguity concerned the position of the sixth strictly conserved histidine because two histidines at positions 43 and I31 as the presumptive N-terminal ligands of the low-spin haem h might fulfil this role. Replacement by site-directed mutagenesis of His 43 to alanine produced a mutant strain in which the three Fix proteins FixN, FixO and FixP are present in similar amounts as in the wild-type strain. By contrast, the FixN protein is absent in the mutant where His I3 I was changed to alanine, and FixO and FixP are only weakly detectable. This indicated that the His 131 to alanine replacement causes a defect in assembly or stability of the oxidase subunits, and suggests that His 131 serves as the N-terminal lowspin haem B ligand (Zufferey et al., 1996b). Identification of all six functionally important histidine residues and their location on the periplasmic face of the membrane have led to a topological model of the fixN subunit of the cytochrome chh3-type haemxopper oxidase containing 12 transmembrane helices organized in a 3-fold symmetry (Zufferey et al., 1996b). This organization resembles the situation in classical subunits I of haem-copper oxidases (Iwata et al., 1995; Tsukihara et al., 1996).

0. OTHER fixNOQP GENES

0.1. A. caulinodans fixNOQP Genes

TheJixNOQP genes were localized in the A . caulinodans genome after hybridization with the R . melilotifixN gene as a probe (Mandon et al., 1993). They encode polypeptides whose amino acid sequences are 76, 71, 43 and 58% similar to B. japonicum FixN, FixO, FixQ and FixP, respectively, and 64,66 and 46% with R . meliloti FixN, FixO and FixP, respectively (Mandon et al., 1994). Five conserved histidines typical of subunit I terminal oxidases are present in FixN, while FixO and FixP contain consensus haem-binding sites. Membrane protein analysis of wild-type and mutant strains revealed the presence of two covalently bound haem polypeptides of M , 32 and 34 kDa, which were assigned to FixO and FixP, respectively. Spectral analysis of whole cells showed a large decrease in the c-type cytochrome content of a$xNO deletion mutant (Mandon et al., 1994). By

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215

contrast to the Fix- phenotype of B. japonicum and R . meliloti j'i.uN mutants, the A . cmlinodans jxNO-deleted mutant strain retains 50% of the nitrogenase activity of the wild-type in the symbiosis, and under freeliving conditions the nitrogenase activity is not significantly different from that of the wild-type (Kaminski et al., 1996).

6.2. A. tumefaciens fixNOQP Genes

Hybridization analysis using the R . meliloti.fi.xN as a probe also revealed the presence of an homologous DNA region in the phytopathogenic bacterium A . tumclfbciens (Schluter et al., 1995). Cloning and sequencing of the hybridizing fragment predicted the presence of two contiguous ORFs, whose nucleotide-deduced amino acid sequences are 86% and 82% identical to the R . melilofi FixN and FixO proteins, respectively. A strain of A . tumefaciens mutated in thefixN-like gene showed decreased TMPD-specific oxidase activity under microaerobiosis, indicating that thefixN gene codes for proteins involved in respiration under reduced oxygen availability. DNA of R. leguminosarum biovar phoseoli, Rhodobacter capsirlatus and E. coli also show hybridizing bands when R . melilotiJixN is used as a probe (Schluter et al., 1995). These findings, together with the fact that genes homologous to those of the ,fi.uNOQP operon are expressed in non-fixing bacteria such as A . tumefaciens, have led to the speculation that the fixNOQP genes in the Rhizobiaceae are not directly involved in nitrogen fixation, but are required for bacteroid respiration under the microaerobic conditions in the nodule environment. Similarly, expression of these genes in Agrohucterium might be necessary for colonization of microaerobic niches in the soil and in the plant tumour tissue (Schluter e f al., 1995).

6.3. R. leguminosarum biovar wiciae Symbiosis-Specific Oxidase

Analysis of covalently bound haem proteins revealed that the total amount of haem-stained proteins in membrane fractions of bacteroids isolated from nodules of pea plants inoculated with R . leguminosarum biovar viciae is considerably higher than that in membranes from free-living cells (Vargas et a/., 1996). The increased level of spectroscopically detectable cytochrome c is correlated with the appearance of two haemproteins of calculated M , of 30000 and 28000, close to those of the FixO and FixP proteins of B. japonicum (Preisig et al., 1993) and A . caulinodans (Mandon et ul., 1994). CO difference spectra indicated that cytochromes aa3 and d are absent in bacteroid membranes, and that at least two CO-binding components are

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MARIA J. DELGADO, EULOGIO J. BEDMAR AND J. ALLAN DOWNIE

present, which could not be found in free-living cells. Photodissociation spectra provided further evidence for the presence of an unusual ligandbinding haemprotein that appears to react with traces of oxygen at low temperature, a behaviour consistent with the postulated high oxygen affinity of the bacteroid oxidase. These results have led to the suggestion that the M , 30 000 and 28 000 haemproteins correspond to the FixO and FixP proteins of the cytochrome oxidase in R . leguminosarum biovar viciae bacteroids (Vargas et al.. 1996). The JixNOQP genes were identified by DNA hybridization and sequencing (Schluter et al., 1993, 1997). Two copies of t h e j x N O Q P operon were identifed, one on the plasmid carrying the nodulation genes and the other adjacent to theJixK-jxL region (Patschkowski et al., 1996) on a different plasmid. Insertional mutagenesis showed that both copies ofJixNOQP were functional and analysis of the expression ofJixN-gusA reporter gene fusions revealed that both operons were induced by microaerobiosis and in the symbiotic zone of nodules. Mutation of fnrN and .fixL strongly reduced expression of both operons although mutation of JixK had only a slight effect on their expression (Schliiter et al., 1997). 6.4. The fixNOQPrelated fixGHlS Operon

The ,fixCHIS operon is formed by four tightly linked genes first identified, cloned and sequenced in R . meliloti (Kahn et al., 1989), and subsequently found in B. japonicum (Preisig et al., 1993, 1996b; Thony-Meyer et al., 1995) and A . caulinodans (Mandon et al., 1994). In R . meliloti, on the basis of the deduced amino acid sequences, all four genes products are predicted to be transmembrane proteins. FixG is probably involved in a redox process because it contains two cysteine clusters typical of iron-sulphur centres present in bacterial ferredoxins. FixI shows homology to the catalytic subunit of bacterial and eukaryotic ATPases involved in cation pumping. It has been speculated that FixI is a symbiosis-specific cation pump whose function is coupled to a redox reaction catalysed by the FixG subunit (Kahn ri al., 1989). The B. japonicum FixGHIS proteins share 27-58'10 amino acid sequence identity with theJixGHIS gene products of R . melilori. Fix1 has been predicted to be a P-type ATPase involved in copper transport, and FixG could be a membrane-bound oxidoreductase enzyme catalysing the oxidation of Cu(1) to the biologically useful Cu(I1) after transport. FixH and FixS could be additional subunits associated with this transport (Thony-Meyer c'r a/., 1995; Preisig et al., 1996b). The FixN, FixO and FixP proteins are absent (or present in barely detectable amounts) in aJixGHI B. japonicum mutant. This finding, together

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with the coordinated expression with (see below), and the conserved localization downstream of, thefixNOQP operon have lent support to the suggestion that the FixGHIS proteins form a membrane protein complex required for maturation of the cbb3-type oxidase. It is suggested that the specific function of the complex would be the uptake and metabolism of copper required for the cytochrome cbb3-type haem-copper oxidase (Thony-Meyer et al., 1995; Preisig et al., 1996b). ThefixGHI B. japonicum mutant had the same phenotypes as those known from the fixNOQP mutants, such as defective symbiotic nitrogen fixation and decreased cytochrome oxidase activity in cells grown under oxygen deprivation (Preisig et al., 1996b)

6.5. Regulation of the fixNOQP and fixGHlS Operons The jixNOQP operon is located between the regulatory genes JixLJIJixK CfiXK2 in B. japonicum) and the fixGHIS operon. About 5 kb of DNA separate $xLJ and fixG both in R . meliloti (Batut et al., 1989) and in B. japonicum (Preisig et al., 1993). JixLJ/K(K2), JixNOQP and JixGHIS are contained within the so-called fix cluster I1 of R . meliloti and fix cluster 111 of B. japonicum, respectively (Fisher, 1994; Thony-Meyer et al., 1995). ThejixNOQP operon of A . caulinodans is adjacent tofixGHIS (Mandon et al., 1994) withinfix cluster V, but is not linked to thefixLJ/K genes, which are located infix cluster IV (Fisher, 1994). FixLJ is a two-component regulatory system, of which the haem moiety bound to the central cytoplasmic FixL domain senses the cellular oxygen levels and, in turn, inversely modulates the kinase and phosphatase activities of the C-terminal domain. Low oxygen conditions favour the kinase activity, resulting in an increased amount of phosphorylated FixJ protein, which then activates transcription of the fixK/fixK2 genes. In R . leguminosarum biovar viciae, an unusualfixL gene was found that encodes a protein that is analogous to a FixLJ hybrid in which the receiver domain characteristic of FixJ is fused near the FixL C-terminus (Patschkowski et al., 1996). The FixK proteins of R . meliloti (Batut et al., 1989), A . caulinoduns (Kaminski et al., 1991), R . leguminosarum biovar viciae (Patschkowski et al., 1996) and the FixK2 protein of B. japonicum (Anthamatten et al., 1992) are homologous to the E. coli Fnr protein, a transcriptional regulator involved mainly in anaerobic respiratory processes (Spiro and Guest, 1990; Unden and Tragesser, 1991; Spiro, 1994). Strains carrying mutations in j.xL or fixJ of R. meliloti, B. japonicum and A . caulinodans form symbiotically inefficient nodules, and nitrogenase activity in free-living A . caulinodans mutant cells is drastically diminished. However, a R. leguminosarum biovar viciae fixL mutant was only slightly

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MARIA J. DELGADO, EULOGIO J. BEDMAR AND J. ALLAN DOWNIE

affected (40% reduction) in symbiotic nitrogen fixation. Null mutations in the fixK genes of R. meliloti (JixK JixK’ double mutants) and A . caulinodans, as well as inJixK2 of B. japonicum, abolish nitrogen fixation by these bacteria (for a review, see Fisher, 1994). Mutation ofJixK in R. leguniinosarum biovar viciae only reduced (by about 25%) symbiotic nitrogen fixation due to the presence of a second JixK-like gene, f n r N . Mutants lacking both fnrN andJixK are Fix- (Patschkowski et al., 1996). Under oxygen-limited conditions, the FixLJ/FixK(K2) regulatory cascade activates JixNOQP expression in R . meliloti (Batut et al., 1989), B. japonicum (Fisher, 1994; Thony-Meyer et al., 1995) and A . caulinnduns (Mandon at ul., 1994). The A . tumefaciens JixN-like gene is also preferentially expressed under microaerobiosis and its transcriptional activation is mediated by an Fnr-like protein (Schluter et al., 1995). Motifs highly similar to the core of the binding site for E. coli Fnr (5’-AAA-TTGAT-ATCAATTT-3’, Fnr box or anderobox; Eiglmeier el ul., 1989) have been found in the promoter regions ofJixNOQP genes or operons of R . meliloti (Batut et al., 1989), B. japonicum (Preisig et al., 1993), A . caulinodans (Mandon et al., 1994), R. leguminosarum biovar viciae (Schliiter et al., 1997) and A . tumefaciens (Schluter et al., 1995). The conserved motif 5’-TTGA-C-GATCAA-G3’ is considered to be the consensus sequence for putative FixK-binding sites in the Rhizohiuceae (Fisher, 1994). The FixK proteins of R. meliloti and the FixK2 of B. japonicum also control expression of the JixCHlS operon, so that concomitant expression of this operon occurs upon activation of JixNOQP by FixK or FixK2, respectively. This is supported by the presence in both microorganisms of a potentialJixK/K2-bindingsite upstream ofJixG (Kahn et al., 1989; Preisig et al., 1993). Activation of transcription ofJixC in A . caulinodans is independent of FixK, and a motif resembling an anaerobox has not been identified upstream of theJixG coding sequence (Mandon et al., 1993). Whether a JixGHIS operon is conserved in A . tumefaciens is not known. The critical role of oxygen in expression of genes involved in bacteroid respiration and energy metabolism in relationship to nitrogen fixation has been reviewed (Hennecke et al., 1990; Fisher et al., 1993; Hennecke, 1993; Batut and Boistard, 1994).

7. GENES INVOLVED IN CYTOCHROME c BIOGENESIS

Cytochromes c have a haem group attached covalently at the motif C-X-XC-H; the two cysteines form thioether linkages with the protohaem vinyl groups. The mature holoproteins, when soluble, are located, in the periplasmic space or, when membrane-anchored, the haem-binding domain is

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located on the periplasmic side of the bacterial cytoplasmic membrane. In many cases, cytochromes c associate in a super-complex with other redox proteins. Given the participation of c-type cytochromes in both aerobic and symbiosis-specific (el, CycM, FixO, FixP) respiratory chains of rhizobia, the proteins involved in biogenesis of c-type cytochromes are necessary for respiration in nodules. Cytochrome c biogenesis involves a post-translational pathway for the conversion of pre-apocytochrome c into the mature holocytochrome c, including translocation of the precursor polypeptide and haem into the periplasm and the covalent linkage between these two molecules (for a review, see Thony-Meyer et al., 1994). Analyses of cytochrome c-deficient mutants of rhizobia led to the identification of several genes involved in the formation of functional c-type cytochromes (Table 3). The cyc V and cyc W genes from B. japonicum encode proteins homologous to the ATP-binding subunit of an ABC-membrane translocator, and to the channel-forming subunit of such translocators, respectively. Thus, the CycV and CycW proteins are thought to be involved in transport of a specific molecule (possibly haem) required for the synthesis of mature c-type cytochromes (Ramseier et al., 1989, 1991). Downstream of the cycVW genes, another gene, cycY, has been identified which encodes a periplasmic thioredoxin-like protein that may be involved in the reduction of the apoprotein cysteine residues before haem attachment (Ramseier et a/., 1991; Fabianek et a f . , 1997). A cycY homologue was identified in R . leguminosarum biovar viciae; mutation of this gene blocked the formation of all c-type cytochromes and symbiotic nitrogen fixation (Vargas et a f . , 1994). In R . etli, the ccmA and ccmB genes (homologues of cyc V and cyc W) were also shown to be required for the assembly of c-type cytochromes and symbiotic nitrogen fixation (Aguilar and Soberon, 1996). A four-gene cluster cycHJKL has been identified in B. japonicum (Ritz et al., 1995), R . meliloti (Kereszt et al., 1995) and R . leguminosarum biovar viciae (Delgado et al., 1995). It has been suggested that these genes might encode a haem lyase complex involved in the attachment of haem to apocytochrome c. Mutations in any of these genes result in the loss of all c-type cytochromes, and block symbiotic nitrogen fixation. Notably, the stage of nodule development reached by cytochrome c-deficient mutants varies among different species. In the pea nodules induced by R . leguminosarum biovar viciae mutants, cells were fully infected with many bacteroid forms surrounded by peribacteroid membranes (Delgado et al., 1995). In contrast, soybean nodules induced by ~aycHmutants of B. japonicum contained very few bacteroids (Ritz et al., 1993), and nodules induced on Phaseolus beans by a cytochrome c-deficient mutant of R . phaseoli lacked bacteroids (Soberon et a / . , 1993). Nevertheless, several of these mutants can grow

Table 3 Rhizobial genes involved in cytochrome c assembly.

Genes or operons

Known or proposed function of gene products

cycvwx

Components of an ATP-dependent transport system for haem

ccmAB (cycVW) cyc Y

Membrane-bound thioredoxinlike protein

cycHJKL

Attachment of haem to apocytochrome c? Influence iron uptake

Species

Symbiotic phenotype

References

B. japonicum

Fix-

Ramseier et al. (1991)

R . etli

Fix-

B. japonicum R. legwninosarum biovar viciae B. japonicum R. melilori R. leguminosarum biovar viciae

Fix-

Aguilar and Soberon ( 1996) Ramseier et al. (1991) Fabianek er al. (1997) Vargas er al. (1994) Ritz et al. (1993, 1995) Kereszt et al. (1995) Delgado et al. (1995) Yeoman et al. (1997)

FixFixFix-

GENES FOR RHlZOBlAL CYTOCHROMES

22 1

aerobically, presumably by relying on the cytochrome d and/or cytochrome hb3 (o?)-terminated branches of the electron transport pathway, which do not involve c-type cytochromes. Analysis of strains mutated in the cycHJKL operon and hence defective in cytochrome c assembly revealed an unexpected role for c-type cytochromes in iron uptake (Yeoman et af., 1997). A mutant that had been thought to accumulate haem precursors due to a defect in iron uptake (Nadler et a f . , 1990) was in fact found to carry a mutation in cycK (Yeoman et af., 1997). Other mutations affecting cycH and cycK induced accumulation of protoporphyrin IX and abolished the production of siderophores. This absence of siderophores caused a defect in high-affinity iron acquisition, although the precise reason for this is not yet known (Yeoman et u f . , 1997). Such pleiotropic effects of mutations in the cycHJKL may account for the different levels of nodule infection by various rhizobia in different legumes.

8. RHlZOBlAL MUTANTS WITH ALTERED OXIDASE ACTIVITY AND IMPROVED SYMBIOTIC NITROGEN FIXATION

Since bacterial respiration is closely associated to supply of ATP, it has been considered that isolation of mutants with increased respiratory efficiency could result in strains with improved symbiotic nitrogen fixation. A R . etfi mutant strain (CFN4205), had reduced levels of cytochrome o and 4-fold higher cytochrome au3 levels in culture. Nodules formed by this strain fixed 33% more N2 that those formed by the wild-type (Soberon et af., 1989). Two Tn.5-induced mutants of R . erli, strains CFN037 (Soberon et al., 1990) and CFN030 (Miranda rt af., 1996), that were selected because of their increased ability to oxidize TMPD and azide-resistant phenotypes, respectively, express cytochrome aa3 and have increased respiratory activities when cultured microaerobically. These mutants had improved symbiotic nitrogen fixation capacity as judged by nitrogenase activity and measurements of the total nitrogen content of nodulated plants. Both strains, besides expressing higher levels of h- and c-type cytochromes, contain an additional CO-reacting cytochrome which is not present in the wild-type strain, and has spectral signals similar to those of the cbb3-type oxidase (Miranda et a/., 1996). Therefore, it has been suggested that rhizobial strains with derepressed expression of cytochrome chh3-type oxidase may provide improved strains for inoculation of legume crops.

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We thank R.K. Poole and S. Marroqui for comments on the manuscript. E.J.B. and M.J.D. acknowledge support by DGICYT (grant PB94-0117) and by EU-TMR (Return grant 950206), and J.A.D. is supported by the BBSRC (UK) and the EU (ERBCIl*CT94-002).

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Preisig, O., Zufferey, R., Thony-Meyer, L., Appleby, C.A. and Hennecke, H. (1996a) A high-affinity &,-type cytochrome oxidase terminates the symbiosis-specific respiratory chain of Bradyrhizobium japonicum. J. Bacreriol. 178, 1532-1 538. Preisig, O., Zufferey, R. and Hennecke, H. (l996b) The Bradyrhizobium japonicurn fixCHIS genes are required for the formation of the high-affinity chb,-type cytochrome oxidase. Arch. Microbiol. 165, 297-305. Ramirez, B.E., Malmstrom, B.G., Winkler, J.R. and Gray, H.B. (1995) The currents of life: the terminal electron transfer complex of respiration. Proc. Natl. Acad. Sci. USA 92, 1 1949-1 1951. Ramseier, T.M., Kaluza, B., Studer, D., Gloudemans, T., Bisseling, T., Jordan, M.P., Jones, R.M.. Zuber, M. and Hennecke, H. (1989) Cloning of a DNA region from Bradyrhizohium japonicum encoding pleiotropic functions in heme metabolism and respiration. Arch. Microbiol. 151, 203-212. Ramseier, T.M., Winter, H.W. and Hennecke, H. (1991) Discovery and sequence analysis of hacterial genes involved in the biogenesis of c-type cytochromes. J . B i d . Chem. 266, 1193-1803. Regensburger, B., Meyer, L.,Filser, M., Weber, J., Studer, D., Lamb, J.W., Fisher, H.M., Hahn, M. and Hennecke, H. (1986) Bradyrhizobium japonicum mutants defective in root-nodule bacteroid development and nitrogen fixation. Arch. Microhiol. 144, 355366. Renalier, M.H., Batut, J., Ghai, J., Terzaghi, B., Gherardi, M., David, M., Garnerone, A.M., Vasse, J., Truchet, G., Huguet, T. and Boistard, P. (1981) A new symbiotic cluster on the pSym megaplasmid of Rhizobium rnelilori 201 1 carries a functional ,fix gene repeat and a nod locus. J. Bacteriol. 169, 2231-2238. Ritz, D., Bott, M. and Hennecke, H. (1993) Formation of several bacterial c-type cytochromes requires a novel membrane-anchored protein that faces the periplasm. Mol. Microbiol. 9. 129-140. Ritz, D., Thony-Meyer, L. and Hennecke, H. (1995) The cycHJKL gene cluster plays an essential role in the biogenesis of c cytochromes in Bradyrhizohiuin japonicum. Mol. Gen. Gene!. 247, 21-38. Rossbach, S., Loferer, H., Acuiia, G., Appleby, C.A. and Hennecke. H. (1991) Cloning, sequencing and mutational analysis of the cytochrome cSs2 gene (cycB) from Bradyrhizobiuin ,japonicum strain I 10. FEMS Microbiol. Lerr. 83, 145-1 52. Santana, M., Kunst, F., Hullo, M.F., Rapoport, G., Danchin, A. and Glaser, P. (1992) Molecular cloning, sequencing and physiological characterization of the qox operon from Bacillus suhtilis encoding the aa,-600 quinol oxidase. J. B i d . Cliem. 267, 10 225 10231. Saraste, M. (1990) Structural features of cytochrome oxidase. Q . Rev. Biophys. 23, 331 336. Saraste, M., Holm, L., Lemieux, L., Liibben, M. and van der Oost, J. (1991) The happy family of cytochrome oxidases. Biochein. Soc. Trans. 19, 608412. Schluter, A., Patschkowski, T., Weidner, S., Unden, G., Hynes, M.F. and Priefer, U. ( 1993) Functional and regulatory characteristics of FnrN, an oxygen-responsive transcriptional activator in Rhizobium legumino.sarum biovar viciae. In: New Horizons in Nirrogen Fi.ucrlion (R. Palacios, J. Mora and W.E. Newton, eds), p. 493. Kluwer Academic Publishers, Dordrecht, The Netherlands. Schliiter, A., Ruberg, S., Kriimer, M., Weidner, S. and Priefer, U. (1995) A homolog of the Rhizohiuni nielilori nitrogen fixation genefixN is involved in the production of a microaerobically induced oxidase activity in the phytopathogenic bacterium Agrohucteriuin ruinejuciens. Mol. Gen. Gent.!. 247. 206-2 15. ~

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Schliiter, A,, Patschkowski, T., Quandt, J., Selinger, L.B., Weidner, S., Kramer, M., Zhou, L.M., Hynes, M.F. and Priefer, U.B. (1997) Functional and regulatory analysis of the two copies of theJixNOQP operon of Rhizobium leguminosarum strain VF39. Mol. Plant. Microbe Interact. 10, 605-616. Slater, E.C., van Gelder, B.F. and Minnaert, K. (1965) Cytochrome c oxidase. In: Oxidases and Related Redox Systems (T.E. King, H.S. Mason and M. Morrison, eds), pp. 667-706. John Wiley & Sons, New York. Soberon. M., Williams, H.D., Poole, R.K. and Escamilla, E. (1989) Isolation of a Rhizobium phaseoli cytochrome mutant with enhanced respiration and symbiotic nitrogen fixation. J. Bacteriol. 171, 465472. Soberon, M., Membrillo-Hernindez, J., Aguilar, G.R. and Sanchez, F. (1990) Isolation o f Rhizobium phaseoli Tn.5-induced mutants with altered expression of cytochrome terminal oxidases o and aa3. J. Bacrcriol. 172, 167&1680. Soberon, M., Aguilar, G.R. and Sinchez, F. (1993) Rhizohiurn phaseoli cytochrome cdeficient mutant induces empty nodules on Phaseolus vulgaris L. Mol. Microhiol. 8. 159- 166. Spiro, S . (1994) The F N R family of transcriptional regulators. Anfonie van Leeuwenhoek 66, 23-36. Spiro, S. and Guest, J.R. (1990) Fnr and its role in oxygen-regulated gene expression in Escherichia coli. FEMS Microbiol. Rev. 75, 399428. Stam, H. Van Verseveld, W.H., de Vries, W. and Stouthamer, A.H. (1984) Hydrogen oxidation and efficiency of nitrogen fixation in succinate-limited cultures of Rhizobiurn ORS571. Arch. Microbiol. 139, 53-60. Stonehuerner, J., O’Brien, P., Geren, L., Millet, F., Steidl, J., Yu, L. and Yu, C. (1985) Identification of the binding site on cytochrome C I for cytochrome c. J. Biol. Chem. 260, 5392-5398. Surpin, M.A., Moshiri, F., Murphy, A.M. and Maier, R.J. (1994) Genetic evidence for a fourth terminal oxidase in Bradyrhizobium ,japonicum. Gene 143, 73-77. Surpin, M.A., Lubben, M. and Maier, J. (1996) The Bradwhizobiurn japonicum cox W X Y Z gene cluster encodes a &type ubiquinol oxidase. Gene 183, 201-206. Taha, S.M. and Ferguson-Miller, S. (1992) Interaction of cytochrome c with cytochrome c oxidase studied by monoclonal antibodies and a protein modifying reagent. Biochemistry 31, 9090-9097. Thony-Meyer, L. (1997) Biogenesis of respiratory cytochromes in bacteria. Microbiol. Mol. Biol. Rev.. 61, 337-376. Thony-Meyer, L., Stax, D. and Hennecke, H. (1989) An unusual gene cluster for the and its requirement for effective cytochrome he, complex in Bra~i~rhizohium,japonicum root nodule symbiosis. Cell 57, 683497. Thony-Meyer, L.,James, P. and Hennecke, H.(1991) From one gene to two proteins: The biogenesis of cytochromes h and c, in Bradyrhixhium japonicum. Proc. Null. Acad. Sci. USA 88, 5001-5005. Thony-Meyer, L..Ritz, D. and Hennecke, H. (1994) Cytochrome c biogenesis in bacteria: a possible pathway begins to emerge. Mol. Microbiol. 12, 1-9. Thony-Meyer, L., Preisig, O., Zufferey, R. and Hennecke, H. (1995) The role of a microaerobically induced ch-type cytochrome oxidase in symbiotic nitrogen fixation. In: Nirrogen Fixarion: Fundamentals and Applications (I.A. Tikhonovich, N.A. Provorov, V.I. Romanov and W.E. Newton, eds), pp. 383-388. Kluwer Academic Publishers, Dordrecht, The Netherlands. Trumpower, B.L. ( 1 990) Cytochrorne bc, complexes of microorganisms. Microhiol. Rev. 54, 101-129.

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Tsukihara, T.. Aoyama. H., Yamashita, E., Tomizaki, T., Yamaguchi, H., ShinzawaItoh. K., Nakashima, R., Yaono, R. and Yoshikawa, S. (1996) The whole structure of the 13-subunit oxidized cytochrome c oxidase at 2.8 A. Science 272, 1136- 1144. Tully, R.E. and Keister, D.L. (1993) Cloning and mutagenesis of a cytochrome P-450 locus from Eradyrhizobium japonicum that is expressed anaerobically and symbiotically. Appl. Environ. Microhiol. 59, 41 3-142. Tully, R.E., Sadowsky, M.J. and Keister, D.L. (1991) Characterization of cytochrome c550 and c555from Bradyrhizobiurn japonicum: cloning, mutagenesis. and sequencing of the c555 gene (cycC). J. Bucteriol. 173, 7887-7895. Unden. G. and Tragesser, M . (1991) Oxygen regulated gene expression in Escherichio coli: control of anaerobic respiration by the FNR protein. Antonie van Leeuwenhoek 59, 65-76. van der Oost, J., Lappalainem, P., Musacchio, A,, Warne, A,, Lemieux, L., Rumbley, J., Gennis, R.B., Aasa, R., Pascher, T., Malmstrom, B.G. and Saraste, M. (1992) Restoration of a lost metal-binding site: construction of two different copper sites into a subunit of the Escherichio coli cytochrome o quinol oxidase complex. EMEO J. I f . 3209 3217. van der Oost, J., de Boer, A.P.N., de Crier, J.W., Zumft, W.G., Stouthamer, A.H. and van Spanning, R.J.M. (1994) The haemcopper oxidase family consists of three distinct types of terminal oxidases and is related to nitric oxide reductase. FEMS Microhiol. Lett. 121, 1-10, Vargas. C., Wu, G., Davies, A.E. and Downie. J.A. (1994) Identification of a gene encoding a thioredoxin-like product necessary for cytochrome c biosynthesis and symbiotic nitrogen fixation in Rhizobium leguminosarum. J. Bucteriol. 176,4117-4123. Vargas. C . , Wu, G., Delgado, M.J., Poole, R.K. and Downie. J.A. (1996) Identification of symbiosis-specific c type cytochronies and a putative oxidase in bacteroids of Rhizohium legutnitiosarum biovar viciae. Microbiology 142, 41 4 6 . von Wachenfeldt, C., de Vries, S. and van der Oost, J. (1994) The CuA- site of the cou3type oxidase of Eui~iI1u.ssuhrilis is a mixed-valence binuclear copper center. FEES Lett. 340, 109-113. Williams, H.D., Appleby, C.A. and Poole, R.K. (1990) The unusual behaviour of the puta live terminal oxidases of Eradyrhizohiuni juponicurn bacteroids revealed by lowtemperature photodissociation studies. Eiochim. Biophys. Acta 1019, 225-232. Witty, J.F. and Minchin, F.R. (1990) Oxygen diffusion in the legume root nodule. In: Nitrogen Fixufion: Achievements and Objectives (P.M. Gresshoff, L.E. Roth, G . Stacey and W.E. Newton, eds), pp. 285-292. Chapman & Hall, New York. Wu, G., Delgado, M.J., Vargas, C., Davies, A.E., Poole, R.K. and Downie, J.A. (1996) The cytochrome bcl complex but not CycM is essential for symbiotic nitrogen fixation by Khizohiutn Ii~gwninosarum.Microhiology 142, 338 1-3388. Yeoman, K.H., Delgado, M.J., Wexler, M., Downie, J.A. and Johnston, A.W.B. (1997) High affinity iron acquisition in Rhizohium leguminosarum requires the cycHJKL operon and theJeuPQ gene products, which belong to the family of two-component transcriptional regulators. Microbiology 143, 127-1 34. Young, J.P.W. (I992) Phylogenetic classification of nitrogen-fixing microorganisms. In: Biolrigicul Nitrogen Fixation (G. Stacey, R.H. Burris and H.J. Evans, eds), pp. 43 -86. Chapman & Hall, New York. Zimmermann, B.H., Nitsche, C.I., Fee, J.A., Rusnak, F. and Munck, E. (1988) Properties of a copper-containing cytochrome ha3 a second terminal oxidase from the extreme thermophile Thernius thermophilus. Proc. Nail. Acud. Sci. USA 85, 5779 5783.

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Zufferey. R.. Preisig, O., Hennecke. H. and Thony-Meyer, L. (1996a) Assembly and function of the cytochrome chh3 oxidase subunits in Brudyrhizohiutn juponicum. J . Biol. Chem. 271. 91 14-91 19. Zufferey, R., Thony-Meyer. L. and Hennecke, H. (1996b) Histidine 131, not histidine 43. of the Brad-vrhizohiumjrrponicuni FixN protein is exposed towards the periplasm and essential for the functioning of the chh3-type oxidase. FEBS Let/. 394. 349-352.

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The Starvation-Stress Response (SSR) of Salmonella Michael P. Spector Lkpartment of' Biomedical Sciences, University of South Alabama, Mobile, Alabama 36688. USA

ABSTRACT Salmonella serovars are common etiologic agents of intestinal-based disease of animals and humans. As a result of their lifestyle, salmonellae occupy and survive in a wide range of niches where they can encounter an even broader range of environmental stresses. One of the most common stresses is starvation for an essential nutrient such as a carbon/energy (C)-source. The genetic and physiologic changes that the bacterium undergoes in response to starvationstress are referred to as the starvation-stress response or SSR. The genetic loci whose expression increases in response to the starvationstress compose the SSR stimulon. Several loci of the SSR stimulon have been identified in Salmonella typhimurium and grouped, based on putative or known functions or products, into transport systems, C-compound catabolic enzymes, known protective enzymes, respiratory enzyme systems, regulatory proteins, virulence loci and unclassified products. The majority of loci identified are under positive control by the rpoS-encoded sigma factor, 0'. However, a few are under (indirect) negative control by os, but only during starvation-induced stationary phase. Most of the loci identified are also under either positive or negative control by the cAMP:CRP complex. For many, additional regulatory proteins (e.g. FadR, OxyR, and RelA and others) play a role in their regulation as well. Furthermore, most of the SSR loci identified are induced during other stresses or environmental conditions. For example, some are induced during P- or N-starvation, in addition to C-starvation; some ADVANCES IN MICROBIAL PHYSIOLOGY VOL 40

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are induced by extremes in pH or osmolarity; and some are induced in the intracellular environment of epithelial cells, and/or macrophages, and/or medium designed to mimic the intracellular milieu of mammalian cells (ISM). Several SSR loci are required for long-term starvation-survival (core SSR loci), e.g. narZ, dad.4, sfiC and rpoS. In addition, a few of the core SSR loci are also required for stress-specific-inducible and/or C-starvation-inducible resistancc to HzOz (e.g. sfiC), thermal (e.g. stiC), and/or acid pH (e.g. narZ). challenge. Interestingly, C-starved cells are resistant to challenge with the antimicrobial peptide, polymyxin B. However, this resistance mechanism(s) is different from the resistance mechanisms for H 2 0 2 and other environmental stresses. Furthermore, a link between the SSR and Salmoriella virulence can be hypothesized since the two major regulators of the SSR, as and cAMP:CRP, are required for full virulence of SalmonelIa. Moreover, the spv (Salmonella plasmid-associated virulence) genes, required for Salmonellu to cause systemic disease, are C (and P- and N-)starvation-inducible. However, a direct link between starvationstress and virulence has not been established conclusively. 1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 235 237 2. The starvation-stress response (SSR) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Physiologic changes during the SSR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 238 2.2. The SSR stirnulon . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 239 2.3. Genetic, environmental and physiologic regulation of SSR loci . . . . . . . . 252 3. The SSR and long-term starvation survival. . . . . . . . . . . . . . . . . . . . . . . . . . . . 264 3.1. Core SSR loci . . . . . . . . . . . . . . . 4. The SSR and resistance to other envir . . . . . . . . . . 266 4.1. H202resistance. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 268 4.2. Therrnotolerance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 269 . . . . . . . . . . . . . . . . . . 269 4.3. Osrnotolerance 4.4. Acid tolerance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 270 4.5. Polyrnyxin resistance. . . . . . . . . . . . .................. 5. The SSR and Sa/mone//a virulence . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6. Concluding remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 272 273 Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 273

Abbreviations: CRP, CAMP receptor protein; product of the crp gene; cAMP:CRP, CAMP-bound CRP complex; C, carbon/energy source; P, phosphate source; N, nitrogen source; SSR, starvation-stress response; MDCK cells, Madin-Darby canine kidney (epithelial) cells; ISM, intracellular salts medium; AIDS, acquired immunodeficiency disease syndrome; PTS,

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phosphoeno1pyruvate:carbohydrate phosphotransferase system; ABC transporters, transport proteins possessing a consensus ATP-binding region; GABA, y-aminobutyric acid; rrg, generic designation for an rpoS-regulated gene; LB medium, Luria-Bertani liquid or solid growth medium; NR, nitrate reductase; WET in vivo expression technology; CSI. C-starvation-inducible.

1. INTRODUCTION

Nutrient limitation and other environmental stresses are routine occurrences for enteric bacteria such as Salmonella serovars. Situations of true feast or non-stress conditions are few and far between for bacteria outside the nutrient-rich media of the research laboratory. As a result, bacteria are most frequently found in a state of starvation- or stress-induced slow or non-growth (Koch, 1971; Harder and Dijkhuizen, 1983; Tempest el al., 1983; Roszak and Colwell, 1987). This is supported intuitively by the fact that, given bacterial growth rates under non-limiting conditions in the laboratory, if growth were non-limiting in natural and host environments it would not be hard to imagine the consequences. The study of how nonsporulating bacteria respond to, thrive in and survive periods of nutrient starvation and other environmental stresses has seen a resurgence in recent years developing into an exciting area of modern fundamental and applied biology. This question is particularly intriguing for pathogenic bacteria, especially those that can occupy niches in natural microcosms as well as those of animal and human hosts. These bacteria not only must respond to and survive potentially lethal limitations and extremes of natural environments but must also tolerate the deluge of antimicrobial weapons launched by the infected host. In fact, one may postulate that it is the inability of specific bacteria to handle either one or more of these milieus that condemns them to a life as obligate intracellular parasites or non-pathogenic saprophytes. However, few bacteria cope with the trials and tribulations of both natural and host environments as well ;is the many serovars of Salmonella enterica (Cabello et al., 1993; reviewed in Foster and Spector, 1995). Salmonella entericu serovars, e.g. Salmonella enterica serovar Typhimurium or Salmonella typhimurium, are common etiologic agents of gastrointestinal-based disease in humans and account for tens of thousands of reported cases (only a small percentage of the estimated three million or so cases going unreported), hundreds of deaths, and significant financial costs each year in the United States alone (Chalker and Blaser, 1988; Pavia and Tauxe, 1991). S. typhimurium is most commonly acquired from

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contaminated food or water, especially poultry products. Typically, this manifests as a self-limiting ‘food poisoning’ or enteritis in otherwise healthy individuals. However, in compromised individuals, e.g. AIDS patients, S . typhimurium can cause more serious potentially life-threatening systemic disease, e.g. bacteremia (Cohen et al., 1987; Finlay and Falkow, 1988, 1989; Wilkens and Roberts, 1988). If one examines a typical life cycle of Salmonella it is easy to see how the ability to respond to, thrive in and survive environmental stress may influence the epidemiology and virulence of these pathogens. Upon release from the host in feces this relatively nutrient-rich condition does not last long since nutrients can quickly be diluted in aquatic environments. The organism is then met by temperature downshifts, extremes in osmolarity and pH, ongoing limitation of one or more essential nutrients and the stresses associated with predation by protozoa (Barker and Brown, 1994). Ingestion of the organism in contaminated food or water begins a whole new journey through numerous hostile microcosms, i.e. pH extremes, anaerobiosis, bile salts, various weak acids, hyperosmolarity, cationic antimicrobial peptides and nutrient limitation resulting from competition with commensals and the host itself for nutrients in the lumen of the intestinal tract. Those organisms surviving these assaults face further attacks in various host microenvironments as they traverse the intestinal epithelium and are engulfed by macrophages where they can survive within phagosomes and/or phagolysosomes (Finlay and Falkow, 1989). Although technically not virulence factors per se, many of the functions needed for survival during prolonged starvation and various specific or multiple environmental stresses both within and outside the host may be important factors influencing the epidemiology and virulence of salmonellae. Further links between nutrient starvation/ environmental stress and Salmonella virulence have been demonstrated. Nutrient limitation, osmolarity. acid pH and anaerobiosis have all been shown to control the expression of Salmonella virulence factors (Miller et al., 1989; Ernst et al., 1990; Foster and Hall, 1990; Galan and Curtis, 1990; Lee and Falkow, 1990; Fang et al., 1991; A. Turk, P. Gulig and M. Spector, unpublished results). Thus, there appears to be a clear empirical relationship between the expression of survival factors and survival both in nature and in host organisms. Although the picture is becoming clearer, still relatively little is known about how non-sporulating bacteria, especially enteric bacteria, respond to and survive environmental stresses such as nutrient starvation. This review attempts to summarize what is known about the starvation-stress response in the enteropathogen Salmonella, but where appropriate will also draw upon what is known for its commensal cousin, Escherichia coli.

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2. THE STARVATION-STRESS RESPONSE (SSR)

The starvation-stress response (SSR) refers to the genetic and physiologic changes the bacteria undergo upon starvation for an essential nutrient. Typically, when we refer to the SSR, the essential nutrient for which we are starving the bacteria is a carbon/energy (C) source. However, phosphate (P) and nitrogen (N) starvations also elicit a SSR (Spector et al., 1986, 1988; reviewed in Magasanik, 1996; Wanner, 1996). The SSR elicited by starvation for each of these three nutrients involves both unique and overlapping sets of genes/proteins (Foster and Spector, 1986; Spector et al., 1986, 1988; Matin ef a/., 1989; Spector, 1990). It should also be noted that a distinction must be made between starved cells and stationary-phase cells. The distinction made here is that stationary-phase cells are those that populate cultures that have stopped growing following log-growth in a rich, or non-limiting minimal, media as opposed to starved cells which populate cultures that have stopped growing as a result of the exhaustion of one or more defined nutrients. For ‘stationary-phase’ cultures, the condition limiting growth is not necessarily defined nor is it typically limited to a single stress. For ‘starved’ cultures, the limitation or stress that restricts cell growth is defined. So, for example, if additional amounts of the limiting nutrient are added to the culture the populating cells will once again begin to grow logarithmically. In addition, stationary-phase cultures normally achieve a much higher cell density compared with starved cultures which can have a significant effect on overall cellular responses and long-term survival. Furthermore, the genes/proteins expressed in stationary-phase cells may or may not overlap with those expressed in cells undergoing a specific defined starvation (Foster and Spector, 1986; Spector et al., 1986; 1988; Matin et al., 1989; Spector, 1990. It should also be noted that a distinction must be made between starved cells and stationary-phase cells. The distinction made here is that stationary-phase cells are those that populate cultures that have stopped growing following log-growth in a rich, or non-limiting minimal, media as opposed to starved cells which populate cultures that have stopped growing as a result of the exhaustion of one or more defined nutrients. For ‘stationary-phase’ cultures, the condition limiting growth is not necessarily defined nor is it typically limited to a single stress. For ‘starved’ cultures, the limitation or stress that restricts cell growth is defined. So, for example, if additional amounts of the limiting nutrient are added to the culture the populating cells will once again begin to grow logarithmically. In addition, stationary-phase cultures normally achieve a much higher cell density compared with starved cultures which can have a significant effect on

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overall cellular responses and long-term survival. Furthermore, the genes/ proteins expressed in stationary-phase cells may or may not overlap with those expressed in cells undergoing a specific defined starvation (Foster and Spector, 1986; Spector et id., 1986; 1988; Matin t’t ul., 1989; Spector and Foster, 1993; Huisman et al., 1996). The SSR refers specifically to the response of starved cells, typically C-starved cells, unless otherwise defined. However, some loci initially characterized as stationary-phase-inducible have also been found to be starvation-inducible and will be included as part of the SSR stimulon.

2.1. Physiologic Changes during the SSR

Morphologically and physiologically starved cells are dramatically different from logarithmically growing (log-phase) cells. The major morphologic and physiologic characteristics of starved and stationary-phase (enteric) bacteria have recently been reviewed in detail (Huisman et uf.,1996). An overview of some of the major changes will be presented for the purposes of later discussions. The cell’s first response to C-starvation is to try and avoid the adverse effects of the stress as well as to gear cell growth to nutrient availability. Persistence of the starvation condition eventually results in a cell that is smaller and much more hardy and efficient than growing cells. This is, at least partially, mediated by the accumulation of two cellular nucleotides, cyclic 3’3’-adenosine monophosphate (CAMP) (reviewed in Botsford and Drexler, 1978; Saier rt ul., 1996) and guanosine 3’3’-bis(diphosphate) (ppGpp) (Gallant et uf., 1976; reviewed in Cashel r t id., 1996). and an alternative sigma transcription factor encoded by the rpoS gene, as or 03’ (reviewed in Hengge-Aronis, 1996). The net effects are to redirect the transcription and overall expression of genes in the cell (discussed later). This ultimately leads to the expression of new or higher-affinity nutrient utilization systems to scavenge the environment for carbon/energy sources as well as other nutrients, degradation of cellular RNA and proteins, reduction in the number of ribosomes, alteration of the components of the inner (including the amounts and types of lipid components) and outer membranes (including the lipopolysaccharide or LPS components), alterations in the peptidoglycan structure and its association with the outer membrane, redirecting of metabolism to make it even more efficient, and the condensation of the chromosomal DNA in order to protect it from damage (El-Khani and Stretton, 1981; Wensink et ul., 1982; Druilhet and Sobek, 1984; Reeve (’1 uf., 1984; lvanov and Fomchenkov, 1989; Leduc rt ul., 1989; Almiron et d., 1992; Dougherty and Pucci, 1994; for reviews see Cashel et ul., 1996; Huisman et ul., 1996; Miller, 1996; Saier et af., 1996).

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As will be discussed later, in salmonellae these changes typically result in smaller highly efficient cells that can withstand the detrimental effects of long-term starvation (starvation-survival) and a variety of other environmental stresses (starvation-inducible general stress or cross-resistance). The morphologically and physiologically discrete starved cell is a direct consequence of the turning on, and off, of specific genes. The evidence for a starvation-stress response stimulon comes from the findings that cells, to which protein synthesis inhibitors (e.g. chloramphenicol) have been added in late log-phase through early starvation-induced stationary phase, do not survive the long-term effects of starvation-stress or other environmental stresses. This is supported by biochemical analysis of specific protein activities, whole cell two-dimensional gel electrophoretic (2-D PAGE) analysis, and operon/gene fusion analysis (Spector et al., 1986, 1988; Matin et al., 1989; Spector and Cubitt, 1992; Spector and Foster, 1993; O’Neal et al., 1994; Seymour et al., 1996). 2.2. The SSR Stimulon

The sets of genes/proteins induced during the (carbon-) starvation-stress response are referred to as the SSR stimulon. The response of S . typhimurium to various starvation-stresses (C-, P-, N- and iron-starvation) as well as other environmental stresses has been studied extensively via 2-D PAGE electrophoresis (Foster and Spector, 1986; Spector et a[., 1986; Foster and Hall, 1990; Foster, 1991; Spector and Foster, 1993). In addition to 2-D PAGE analysis, our laboratory and others have used operon/gene fusion techniques to identify genetic loci of Salmonella induced during various starvation conditions (Foster, 1983; Foster and Spector, 1986; Spector et a[., 1988, 1998a; Fang et al., 1991, 1992, 1996; M. Spector, unpublished results). When examining the results from these various studies, several points become clear. Firstly, initial 2-D PAGE analysis of the SSR clearly missed a number of proteins whose expression is increased during various stress (e.g. C-starvation) conditions (Spector et al.. 1986). This has become apparent from our operon fusion studies. We have accumulated over 70 independently isolated Mud-directed operon fusions to promoters induced during C-starvation conditions. Of these 70-plus fusions, admittedly only about 18 have been studied in any detail. Of these 18, some 25% have represented separate insertions into the same locus. If this holds true for the remaining C-starvation-inducible fusions, yet to be identified, then our collection of fusions should represent around 4&50 different genetic loci. This estimation is much higher than our original estimation of around 15 C-starvationinduced proteins based on 2-D PAGE analysis but is close to the 30-40

2 40

MICHAEL P. SPECTOR

C-starvation-induced proteins estimated in E. coli (Spector et al., 1986; Matin et ul., 1989). Secondly, results from 2-D PAGE analysis from both E. coli and Sulmonellu have shown that induction of the SSR stimulon occurs in sequential programmed phases (Matin ef al., 1989; M. Spector and J . Foster, unpublished results). These findings are supported by results from kinetic studies of C-starvation-inducible lac operon fusions. Inductionkinetics of more than 30 starvation-inducible fusions indicate that there are at least four major phases of gene expression designated phases 0 to 3 (Spector and Cubitt, 1992; Spector and Foster, 1993; M. Spector, unpublished results). Phase 0 genes are induced in late log-phase just prior to entry into C-starvation-induced stationary phase; however, they are not induced in late log-phase in LB or non-limiting minimal medium. A subgroup of phase 0 genes appears to be turned off or exhibit a reduction in expression as the cell enters starvation-induced stationary phase. Another subgroup of phase 0 genes continues to increase, eventually peaking and then reaching a steady-state level during the first 24 h of starvation. Phase 1 genes are induced during the transition from log-phase to starvation-induced stationary phase typically reaching a peak by 4 h of starvation. Two subgroups of phase 1 genes can be described, one subgroup is turned off over the next 24 h and the other subgroup of phase 1 genes reaches a (lower) steady-state level of expression over the next 24 h of starvation. Interestingly, the vast majority of loci identified, thus far, by operon fusion analysis are phase 1 loci. Moreover, only phase 1 loci have been shown to affect long-term starvationsurvival of the bacteria. Phase 2 genes are induced after 1 h of starvation and phase 3 genes are induced after about 4-5 h of starvation. Unfortunately, the latter two groups of genes are the least characterized because each is represented by only two and one fusion, respectively (Spector and Cubitt, 1992; M. Spector, unpublished results). Furthermore, it is evident that no stress-response niodulon (e.g. C - , P- or N-starvation, heat-shock, anaerobiosis, acid-shock) is completely distinct from the others. In other words, the SSR and other stress responses possess both unique and overlapping sets of genes and proteins (Spector and Foster, 1993). These findings and others presented below support the idea that there is a good deal of redundancy and overlap in the bacterium's response to environmental stress. Lastly, the interconnection of the various bacterial stress responses suggests a complex regulatory network of control at both the global and stressspecific levels. This last point has clearly been demonstrated over the past several years for both Sulmonellu and E. coli (see reviews, see Foster and Spector, 1995; Hengge-Aronis, 1996; Huisman ef a/., 1996; this review). The identification and characterization of the remaining fusions in our collection and in those of other researchers should provide a much clearer

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look at the SSR stimulon of Salmonella in the next several years. Furthermore, the knowledge gained from similar studies in Salmonella, E. coli and other bacteria should create a much better understanding of the stress responses of non-differentiating bacteria, in particular enteric bacteria. However, we have begun to develop a picture of the SSR of salmonellae by identifying and characterizing a number of C-starvation-inducible loci identified via Mud-lac fusions in S . typhimurium. Our preliminary analysis has allowed us to group these loci into several categories (Table 1).

2.2.1.

Transport Systems

As mentioned previously, one of the physiologic changes that occurs in starved cells is the induction of new or higher-affinity transport systems presumably to scavenge the environment for needed nutrients. As a recent example, two C-starvation-inducible genes have been identified as members of putative transport systems (M. Spector, M . Pallen and G . Dougan, unpublished results). The first of these is the csiA locus. The csiA locus is a phase 1 C-starvation-inducible locus ( M . Spector, unpublished results). Partial sequencing analysis across the Mud-lac insertion site has revealed that CsiA represents an Enzyme 11-like component of a phosphoenolpyruvate:carbohydrate phosphotransferase system (PTS; reviewed in Postma et al., 1996). Comparison with known DNA and amino acid sequences in various databases indicate homology to the consensus sequence of Enzyme IIB domains of known PTS Enzyme I1 components. Interestingly, the three identified E. coli sequences obtained from the search did not exhibit a high enough degree of similarity to be considered orthologous to CsiA (M. Pallen, G. Dougan and M. Spector, unpublished results). Thus, CsiA appears to represent a previously unidentified Enzyme 11-like component involved in the phosphorylation-coupled transport of some unknown compound. The second locus encoding a putative transport system is csiC. The csiG locus is a phase 0 C-starvation-inducible locus. It is induced about 1 h prior to starvation and reaches a peak upon entry into starvation-induced stationary phase; its expression then slowly declines to a lower steady-state level over the next few hours of starvation ( M . Spector, unpublished results). Sequence analysis of DNA adjacent to the Mud-lac insertion site has divulged homology with a variety of periplasmic binding protein-dependent ABC transporters from a wide range of bacteria (Boos and Lucht, 1996). However, none of these transporters appeared orthologous to CsiG ( M . Pallen, G. Dougan and M . Spector, unpublished results), suggesting that

Table 1 Known carbon-starvation-inducible SSR loci of Salmonella.

Genetic locus (genetic map location)* Transport systems csiA (nm) csiC (nm) C-Compound catabolic enzymes aldB (80.2) fudF (6.5)

gubD (61.1) dadA (41.1) rrg-5: (nm)

Known Protective enzymes katE (29.6) ofsA (nm)

Product/functiont PTS Enzyme I1 component homolog - Phosphyorylation-coupled transport of unknown compound ABC transporter homolog (paralogous family offtsE, .fepC and phnC) - influx or efRux system for unknown compound(s) Aldehyde dehydrogenase - converts glycolaldehyde and lactaldehyde to glycolate and lactate, respectively; possibly others; may function in alternative C-source utilization Medium/long-chain fatty acid acyl-CoA dehydrogenase - poxidation of medium/long-chain fatty acids (e.g. oleic acid) for use as C-source Succinate-semialdehyde dehydrogenase - enzyme in pathway for utilization of y-aminobutyric acid (GABA) as N-source (Salmonellu unable to use it as C-source) D-Amino acid dehydrogenase - works with dadB-encoded alanine racemase for utilization of Palanine (other D-amino acids) as C-sources Putative oxidoreductase - substrate unknown Catalase HPII

-

detoxification of HzOz

Trehalose-6-phosphate synthase - in operon with trehalose-phosphate phosphatase (orsB); functions in trehalose synthesis; osmotolerance and thennotolerance

Respiratory Enzyme Systems narZ (35.8)

Regulatory Proteins rpoS (63.6) crp (75.0)

(cryptic) Nitrate reductase 11, a subunit operon; physiologic function unknown

-

part of nitrate reductase system encoded by narZYWV

Second major o subunit of starved, stationary-phase, or stressed cells - os or 03',synonymous with KatF; reprograms RNA polymerase molecules to genes needed during inducing condition cAMP receptor protein (CRP) - binds with cAMP to function as either a transcriptional activator of C-source utilization systems upon exhaustion of glucose or a transcriptional repressor of several starvation-inducible genes

Table 1

Virulence loci rpoS CrP spv R/spvABCD@SLT)

Unclassified stic (47.3) csiH (92.5)

continued

See above See above SpvR is a positive regulator of spvABCD transcription; required for ability of Salmonella to cause systemic disease; exact functions of these proteins are unknown but increase growth rate of the bacteria in mice Homolog of hypothetical 22 kDa protein of E. coli - no DNA or deduced amino acid sequence homology to any known sequences; potential inner membrane association; putative C-starvationinducible general stress protein Homolog of hypothetical 12 kDa protein of E. coli - potential inner membrane association; potential function in ion-coupled transport of an unknown solute

csiC (nm) csiM (nm) stiG (88.2)

? ?

stiH (57.4) rrg-I+ (nm)

?

rrg-2 (nm) rrg-3 (nm)

No DNA sequence homology to any known E. coli or Salmonella sequences Homolog of a hypothetical E. coli protein 0186 - function unknown

rrg-4 (nm)

No DNA sequence homology to any known E. coli or Salmonella sequences

9

No DNA sequence homology to any known E. coli or Salmonella sequences

*Genetic map locations are in centisomes (Cs) on the Salmonella genetic map (Sanderson et al., 1996). Map locations given are from published or unpublished mapping results or are estimated based on location of E. coli homologue on E. coli genetic map (Berlyn et al., 1996) with respect to shared local genetic markers between E. coli and Salmonella. (nm) = not mapped. ?Product or function deduced from partial DNA sequence analysis, homology searches in GenBank and other databases, and phenotypic confirmation when possible. ? = no sequence data available. See text for references and further explanation. $The designation rrg is for rpoS-regulated gene; rrg-1 through rrg-5 refers to the rpoS-regulated MudJ insertions in strains XF259 through XF326 in Fang er al. (1996), respectively.

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CsiG represents an as yet unidentified ABC transporter of a still unknown substrate.

2.2.2. Carbon (C)-Compound Catabolic Enzymes Another important aspect of starved cell physiology is a switch from a predominantly anabolic metabolism in growing cells to a more diverse and efficient catabolic metabolism. This has been reviewed recently by Huisman et ul. (1996). Thus, one feature of starved cells is the increased or new expression of a variety of catabolic enzyme systems that allow maximum utilization of available carbon/energy sources and other nutrients. This is supported by the recent findings that several C-starvation-inducible lac operon fusions from our collection represent a diverse mix of carboncompound catabolic enzymes. Two, designated csi-5 and csi-10, were both found to be phase 1 C-starvation-inducible lac fusions since both are induced at the onset of C-starvation (M. Spector, unpublished results). Sequence analysis of DNA adjacent to the Mud-luc insertion sites indicate that both are in the aldB gene (Xu and Johnson, 1995; M. Pallen, G. Dougan and M. Spector, unpublished results). The aldB gene encodes an aldehyde dehydrogenase whose substrates include glycolaldehyde and lactaldehyde and possibly others. The regulation of this enzyme activity by growth on several different sugars and certain amino acids yielding 2-0x0glutarate indicates that this enzyme plays an important role in the use of alternative C-sources and central energy metabolism (Quintilla et al., 1991). Three other fusions, designated csi-11, csi-20 and csi-101, were also found to be phase 1 C-starvation-inducible lac fusions (M. Spector unpublished results). Analysis of the DNA sequence adjacent to each insertion site indicates that they are all located within thefadFgene which encodes a medium/ long-chain fatty acid acyl-CoA dehydrogenase. This enzyme is key to the utilization of medium- or long-chain fatty acids (e.g. oleic acid) as C-sources via P-oxidation (reviewed in Clark and Cronan, 1996). The identity of this locus as ,fadF is supported by two findings: (i) strains carrying these insertions are defective in the ability to utilize oleic acid as a C-source; and (ii) these strains are also defective in P-oxidation. In addition to the C-starvation-inducibility of thefadFgene, several findings indicate that the ability to utilize medium/long-chain fatty acids is a key event in the SSR. First, the cellular levels of monounsaturated fatty acids such as oleic acid decrease significantly during starvation and, second, fadF mutants exhibit altered Cstarvation-survival (M. Spector, C.C. DiRusso, M. Pallen and G. Dougan, unpublished results). Another phase 1 C-starvation-inducible lac fusion, designated csiL, proved to be in the gahD gene (M. Pallen, G. Dougan and M. Spector,

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unpublished results). The gabD locus encodes a succinate-semialdehyde dehydrogenase activity which is part of an operon including at least gahT and gahP which encode a glutamic acid-succinic semialdehyde aminotransferase activity and a y-aminobutyrate (GABA) permease, respectively. These activities are found in a number of bacteria where they are required for the utilization of GABA as a carbon and/or nitrogen source (Dover and Halpern, 1972a,b; McFall and Newman, 1996; Reitzer, 1996). It is interesting that several S. typhimurium laboratory and virulent strains cannot utilize GABA as a C-source but can utilize it as a N-source (M. Spector, unpublished results). Further evidence that the csiL locus encodes the gabDTP operon is that strains carrying the csil-lac insertion are defective in the ability to utilize GABA as a N-source (M. Spector, unpublished results). Although S. typhimurium can use GABA as a N-source but not a C-source we have yet to demonstrate guhD(TP) induction during N-starvation even though it is clearly induced during C-starvation (M. Spector, unpublished results). The importance or significance of this is not clear at this time. It should be noted, however, that E. coli apparently possesses a functional GABA utilization pathway but cannot utilize GABA as a C- or N-source under normal circumstances. In contrast, Klebsiellu aerogenes can use GABA as both a sole C- and N-source (McFall and Newman, 1996). It appears that GABA utilization capability may be associated with specific niches of specific bacteria. Why Salmonella appears to be able to utilize GABA as a N-source but not a C-source is not known, nor is why it appears to be under C-starvation control but apparently not N-starvation control. An additional lac fusion shown to be a phase 1 C-starvation-inducible locus is sfiB. The regulation and phenotypic characterization of this locus has been carried out in some detail and will be discussed in later sections (Spector et ul., 1988; Spector and Cubitt, 1992; Seymour e f al., 1996). Sequence analysis of DNA adjacent to the stiB-lac insertion site revealed that the stiB locus encodes the dadAB operon (S. Bearson, M . Pallen, G. Dougan, J. Foster and M. Spector, unpublished results). The insertion itself is in the dadA gene which encodes the smaller subunit of D-amino acid dehydrogenase; dads encodes an alanine racemase. Together they are required for the utilization of L-alanine and certain D-amino acids (e.g. Disomers of phenylalanine, methionine and asparagine) as carbon/energy sources (McFall and Newman, 1996). The identity of the stiB locus as daclAB is supported by the finding that a sfiB mutant cannot utilize L-alanine as a sole C-source (M. Spector, unpublished results). The finding that stiB encodes an L-alanine/D-amino acid utilization system coupled with previous findings that stiB is required for starvation-survival indicate that the ability to utilize L-alanine and/or D-amino acids is an important part of the Salmonella SSR. The source of these substrates is not clear; likely possibilities include the environmental milieu of the bacterium or the turnover of

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bacterial cell wall components or proteins. Another possible scenario is that the dudA insertion affects dudB expression. DadB is one of two alanine racemases that are found in Sulmonrllu. The other is encoded by the ulr locus. The latter is thought to be expressed at a constitutively low level, adequate for supplying D-alanine for peptidoglycan synthesis. This has led to the proposal that DadB is the ‘catabolic’ alanine racemase and Alr is the ‘biosynthetic’ alanine racemase (Wasserman et ul.. 1983; McFall and Newman, 1996). However, this may change during starvation/stationary phase where the addition of the DadB racemase may be needed to supply D-alanine residues for the thicker and more cross-linked peptidoglycan of starved or stationary-phase cells (Huisman et al., 1996). Although induction of the ciudAB operon during C-starvation is more likely due to its catabolic function, a biosynthetic function during these specific conditions cannot be ruled out. Another possible SSR locus involved in C-compound catabolism is the rrg-5 locus. According to Fang et ul. (1996) sequence analysis indicates that it encodes a putative oxidoreductase activity. Unfortunately, its substrate and physiologic role are unknown.

2.2.3. K n o w Protective Enzyines One characteristic of C-starved cells is that they exhibit increased resistance or tolerance to the detrimental effects of a number of environmental stresses in addition to long-term starvation (Lange and Hengge-Aronis, 1991; Matin. 1991; Spector and Cubitt, 1992; O’Neal et a/., 1994; Foster and Spector, 1995; McLeod and Spector, 1996). I t is not surprising therefore that the expression of proteins/enzymes known to protect the cell against environmental stresses are increased during starvation or stationary phase. Two of these known protective systems are encoded for by the kutE and orsA(B) loci. The KatE protein is a second catalase activity known as catalase HPII (Loewen et al., 1985). Although kutE has been found in E. coli to be induced during stationary phase in rich medium (Mulvey et ul., 1990), we have recently shown that it is also induced significantly during Cstarvation (M. Spector, unpublished results). Catalase HPII functions in protecting the cell from oxidative damage by degrading any H 2 0 2 that might be present or generated (Loewen et al., 1985). The otsA locus of Sulnionellu was found by Fang el a/. (1996) to be stationary-phase-induced and later we showed that it is also C-starvation-inducible (M. Spector and F. Fang, unpublished results). Along with OtsB, OtsA is involved in the synthesis of (periplasmic) trehalose which functions in osmoprotection and thermotolerance (Fang et ul., 1996).

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2 47

Other known protective activities that may be part of the SSR based on findings in E. coli (see review by Hengge-Aronis, 1996) include the glycinebetaine transport systems encoded by the pro VWX operon and p r o p , both of which function in osmotolerance, xthA which encodes exonuclease 111 and plays a role in repairing DNA damage caused by H202, and uspA (Farewell, et a/., 1996) which encodes what is hypothesized to be a universal stress protein required for resistance to a variety of environmental stresses. In addition, other Salmonella SSR loci described can also play a role in resistance to certain environmental stresses. Specific roles of these loci in stress-specific or starvation-induced general resistance to various stresses will be discussed later.

2.2.4. Respiratory Enzyme Systems As noted so eloquently in Nystrom et a f . (1996), aerobically C-starved cells and anaerobically shifted cells are remarkably similar physiologically. These workers showed that a strain defective in the global regulator ArcA is unable to reduce the expression of several TCA cycle enzymes during aerobic C-starvation, resulting in increased respiration and metabolic activity during starvation. As a result arcA mutants are severely defective in Cstarvation-survival. These findings allowed these authors to propose that the ArcA-dependent reduction in electron donor production and aerobic respiratory enzyme levels and activity that occurs during growth arrest is key to starvation-survival because it reduces the generation of reactive oxygen intermediates (thus limiting potential damage to macromolecules) and controls the rate of energy source utilization. It is well known that, during anaerobiosis, E. cofi and Salmoneffu can induce alternative respiratory enzyme systems (e.g. fumarate reductase, nitrate reductase and nitrite reductase) for the utilization of electron acceptors such as fumarate, nitrate and nitrite, respectively, in order to efficiently generate utilizable energy. However, in the case of the anaerobic induction of the nitrate and nitrite reductases, the electron acceptor must be present. Taken together, the reduction in oxygen-dependent respiratory chain activity during starvation, the production of alternative respiratory enzyme systems during anaerobiosis, and the similarity of starved and anaerobically shifted cells, suggest that non-oxygen-dependent alternative respiratory chains may be important in starved cells. This hypothesis is supported by findings with two of our C-starvationinducible lac fusions designated stiA1 and sti-99. Both are phase 1 Cstarvation-inducible fusions. The regulation and phenotypic characterization of stiA has been described in some detail and will be discussed further in later sections (Spector et ul., 1988; Spector and Cubitt, 1992; Seymour et

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al., 1996). Sequence analysis of DNA flanking the insertion sites of these two fusions indicated that they are both in the narZ gene but at different locations (M. Spector, S.M.D. Bearson, M. Pallen, G. Dougan and J. Foster, unpublished results). The narZ gene is the first gene of an operon (narZYWV) encoding a second (cryptic) nitrate reductase activity, NR-I1 or NR-Z. NR-Z in both E. coli and Salmonella exhibits a high degree of both DNA and amino acid sequence homology to the major anaerobic nitrate reductase activity, NR-I or NR-A, encoded by the narCHJl operon. Nitrate reductase-Z activity was thought to be constitutive since regulation by nitrate or anaerobiosis could not be demonstrated (reviewed in Gennis and Stewart, 1996). Regulation of the stiA-lac and sti-99-lac fusions both confirm that narZ is not nitrate or anaerobically-induced (M. Spector, unpublished results). One model proposed that NR-Z provided a basal level of NR activity that could be utilized by the cell upon an anaerobic shift in the presence of nitrate providing enough energy to allow for the production of the major nitrate and anaerobiosis-inducible NR-A activity (Barrett and Riggs, 1982; Gennis and Stewart, 1996). However, results from our laboratory indicate that narZ( Y W V ) (or stiA) is induced in N-starved and P-starved as well as C-starved cells, suggesting a key role in starved cell physiology. Further-more, this locus exhibits regulation that is similar to other starvation/stationary-phase loci, e.g. rpoS-dependence and CAMPCRP repression. Moreover, this locus is needed for starvation-survival and adaptive resistance to H 2 0 2(Spector and Cubitt, 1992; O’Neal et al., 1994; Seymour e f al., 1996). Thus, the NR-Z activity may be important in the defense of Salmonella against so-called aging as proposed by Nystrom et al. (1996). However, the exact physiologic function of the NR-Z activity in starved cells and/or H202-adapted cells is not known. Other possible respiratory enzymes that may be part of the SSR based on findings in E. coli include the hydrogenase-I complex encoded by the hyaABCDEF operon and CyxAB (a cytochrome bd homolog) encoded by the cyxAB operon. Both these have been shown to be stationary-phaseinducible but direct C-starvation-induction has not been demonstrated in E. coli or Salmonella (Hengge-Aronis, 1996).

2.2.5. Regulafory Proteins Regulation of the SSR stimulon of Salmonella and E. coli occurs at several levels and is highly interconnected. Many SSR genes are controlled by multiple repressor and activator proteins which fine tune their expression under different environmental conditions. The two major regulatory proteins of the SSR, os and the cyclic AMP (CAMP) receptor protein, both exhibit increased expression in C-starved cells.

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The rpoS-encoded alternative sigma transcription factor, us or c3*or KatF (Mulvey and Loewen, 1989), exhibits multilevel regulation (discussed further below). The net result is increased expression of the us protein to such a level that it becomes a second major sigma factor for RNA polymerase (E) in starved and stressed cells (Hengge-Aronis, 1996).Accumulation of os during these conditions reprograms RNA polymerase molecules to recognize, bind to, and initiate transcription from a new group of promoters. Those promoters recognized by EoS belong to genes and operons of the rpoS regulon. Since not all SSR loci are under os control, the rpoS regulon represents a subset of genes within the SSR stimulon. However, the rpoS regulon is a very important component of the SSR since rpoS mutants are defective in starvation-survival and the generation of stress-specific and Cstarvation-induced cross-resistance (Fang et al., 1992, 1996; O’Neal et al., 1994). The major regulatory protein of the SSR is the cAMP receptor protein (CRP). CRP acts as both a repressor and an activator protein of SSR gene/ operon expression (Spector, 1990; Spector and Cubitt, 1992; Fang et al., 1996). CRP along with its co-activator cAMP have long been associated with the regulation of carbon utilization metabolism (reviewed in Saier et al., 1996). Both cAMP and the CRP protein increase as a result of C-source starvation. The increase in cAMP is due primarily to an activation of adenylate cyclase (product of the cya gene) activity rather than increased expression of the cya gene. In contrast, increases in C R P levels are due to increased transcription of the u p gene. cAMP:CRP is best characterized as an activator of numerous catabolite repressed systems for carbohydrate utilization but is becoming increasingly recognized as a repressor of gene expression (Saier et al. 1996). Therefore, it is not surprising that cAMP:CRP is a central player in the regulation of C-starvation-inducible gene expression. i.e. the SSR. 2.2.6.

Virulence Functions

Although a specific link between starvation and pathogenicity of Salmonella has not been found, many virulence factors have been shown to be starvation-inducible (reviewed in Mahan et al., 1996). Thus, they are part of the SSR of Salmonella. It is intriguing, however, to ponder why virulence factor expression has maintained the starvation-regulatory circuit over the evolution of Salmonella as a pathogen if starvation is not an important signal within the host. Two SSR loci that have been shown to play a role in virulence, based on the fact that when mutated they significantly affect the virulence of Salmonella, are the two major regulators of the SSR, rpoS and crp. Both

2 50

MICHAEL P. SPECTOR

rpoS and crp mutants exhibit reduced virulence, by several logs, in the mouse virulence model for Salmonella. The most logical explanation for their roles in virulence is the regulation of virulence factor expression, e.g. spv gene expression. However, this explanation is not straightforward, particularly in the case of crp, since cAMP:CRP can positively and negatively regulate different virulence-associated functions (Fang et al., 1992; Mahan et al., 1996). Several Salmonella serovars possess a relatively large (5C90 kb) plasmid (virulence plasmid) that is essential for the bacteria to cause systemic disease in their hosts. Although different serovars can possess different sized plasmids, they all appear to contain a conserved, approximately 8 kb, region that if cloned and placed into virulence plasmid-cured strains can by itself restore systemic virulence to the strain. This region contains five genes designated spvR and spvABCD. The spvR gene product is a positive regulator of spvABCD operon expression. The exact functions of the SpvA, SpvB, SpvC and SpvD in virulence are not known. There is evidence that SpvA is a negative regulator of spvR, thus providing a negative feedback control loop (Gulig et al., 1993). The spvABCD operon is Cstarvation-inducible (Fang et al., 1992; A. Turk, P. Gulig, and M. Spector, unpublished results). However, since spvABCD induction requires SpvR, C-starvation probably induces SpvR expression which in turn induces spvABCD. Other virulence factors which may be part of the SSR of Salmonella include the phoPQ regulon and the slyA regulon (Groisman e f al., 1989; Miller et al., 1989; Buchmeier et al., 1997). The phoP locus was originally identified as a positive regulator of a non-specific acid phosphatase activity (phoN gene product) during P-starvation (Kier et al., 1979). Later it was found to be required for survival within macrophages and for Salmonellu virulence (Miller et al., 1989). Behlau and Miller (1993) showed that some of the pag (PhoP-activated gene) and prg (PhoP-repressed gene) loci are oppositely regulated during aerobic starvation, with pug loci being induced and prg loci being repressed. This would suggest that the PhoPQ sensor-regulator system is responding to a starvation signal and in turn regulating the expression of pag and prg genes. Recent preliminary experiments in our laboratory, however, found that a phoP-lac fusion carried on a plasmid with a wild-type phoPQ on the chromosome is not induced during the first few hours of C-starvation; in fact, phoP-lac expression was found to decrease by almost 50% during this time (M. Spector, unpublished results). However, Behlau and Miller (1993) measured expression of pag and prg genes after 48 h of what appeared to be multiple nutrient starvation. Thus, it is possible that phoPQ expression may be turned on later during C-starvation or the reduced level of PhoPQ expression observed early in Cstarvation may be sufficient to regulate pag and prg expression under these

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conditions. Interestingly, we have found that some pug genes are induced during the first few hours of C-starvation even though phoPQ expression appeared to be decreasing (M. Spector, unpublished results). Although we have not determined if this C-starvation induction is PhoP-dependent, this suggests, at least, two possibilities: (i) that some pug loci may be under the direct control of additional regulatory systems besides PhoPQ; or (ii) that the PhoPQ-regulon may represent another subset of genes/proteins composing the SSR stimulon. The slyA locus was originally identified as a gene required for Salmonella virulence and macrophage survival. It was later identified as a transcriptional regulator needed for the expression of an E. coli cryptic hemolysin but a similar scenario has not yet been demonstrated for Salmonella. It has recently been demonstrated that SIyA itself is induced and regulates the induction of a number of proteins during stationary phase and upon macrophage infection. Furthermore, the stationary-phase induction of the slyAregulon is os-independent (Buchmeier et al., 1997). Unfortunately, the parameters of the stationary-phase conditions were unclear so it is not known to what signal sIyA is responding. One possibility is C-starvation, and thus it may also represent another subset of genes/proteins composing the SSR stimulon.

2.2.7.

UnclassiJed

There are several C-starvation-inducible loci identified by lac fusion techniques that have eluded classification into any of the above groupings. For several, there are sequence data available but because the genes do not result in any known auxotrophy or classifying defect and exhibit little or no homology to any known sequences in the various databases it is difficult to propose a function for the gene product. For others there are no sequence data available and no observable phenotype associated with the Mud-lac insertion. Examples of the former group of unclassified loci include stiC, csiH and rrg-l to rrg-4 (rrg-lto rrg-5 refer to the ypoS-regulated gene fusions in strains XF259 to XF326 in Fang et al. (1996). respectively). The stiC gene appears to encode a homolog of a hypothetical 22 kDa protein of E. coli; amino acid sequence characteristics suggest that it may be an inner membrane protein. Phenotypic characterization of strains carrying the sriC-lac insertion mutation suggest that it may represent a major C-starvation-induced general stress protein (V. Makam Nataraj, A. Mahmud and M. Spector, unpublished results). The csiH locus appears to encode a homolog of another hypothetical E. coli protein (12 kDa) that may be associated with the inner membrane. Amino acid sequence

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homology seems to suggest a potential function of this protein in ioncoupled transport of an unknown solute (M. Spector, M . Pallen and G. Dougan, unpublished results). Unfortunately, no clear conclusions could be drawn from the sequence data for the rrg-1 through rrg-4 loci with respect to the potential function(s) of their gene products (Fang et al., 1996). The other group of unclassified loci, for which no sequence data or classifying phenotype is available, includes csiC, csiM, stiE, stiC and sriH. The unifying characteristic of all the unclassified loci, however, is that they are all at least C-starvation-inducible; thus, they are members of the SSR of Salmonella (Spector et al., 1988; M . Spector, unpublished results). 2.3. Genetic, Environmental and Physiologic Regulation of SSR Loci

As more is learned about the regulation of SSR gene/operon expression in Salmonella and E. coli as well as other bacteria it becomes increasingly clear that regulation of the SSR stimulon is very complex and multileveled. Individual genes or sets of genes within the SSR stimulon share a level of common genetic regulation as well as sometimes many levels of regulation specific to that gene. The same can be said for the physiologic and/or environmental regulation of the genes and operons composing the SSR stimulon. They are all by definition C-starvation-inducible but most are also induced by other conditions. Thus, it is difficult to predict under what conditions various SSR loci will be induced or what regulators will be involved in their regulation. The complex genetic, environmental and physiologic regulation of C-starvation-inducible SSR loci of Salmonella is summarized in Table 2. 2.3.1. Genetic Regulation of SSR Loci Several global regulatory systems have been implicated in the regulation of SSR gene expression. The two most important regulators in terms of the number of loci under their control are cAMP:CRP and 0’. 2.3.1.1. cAMP:CRP Most C-starvation-inducible loci identified in Salmonella are under direct or indirect control by the cAMP:CRP complex. This control can be either positive or negative; in some cases cAMP:CRP can be both a positive and negative regulator under different conditions. Studies in E. coli have estimated that as many as two-thirds of C-starvationinducible proteins are positively regulated by cAMP:CRP with the remain-

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ing one-third being either negatively regulated or not regulated at all by cAMP:CRP (Matin et al., 1989). The cAMP:CRP complex is best characterized as an activator protein for certain sugar utilization operons, e.g. the lac operon of E. coli (reviewed in Botsford and Drexler, 1978; Saier et al., 1996). Several C-starvationinducible SSR loci are under positive control by cAMP:CRP (Table 2). These include the two putative transport proteins encoded by csiA and csiG, several C-compound catabolic enzymes encoded by aldB, fadF, gabD( T P ) , and two unclassified SSR loci designated by csiH and csiM. It has been hypothesized that the positively regulated C-starvation-inducible proteins are involved in starvation-stress avoidance. In this model, their primary function would be to scavenge and utilize all available nutrients in the hope that, teleologically speaking, nutrients will become available again and the cell will not have to enter into the programmed non-growing state (Matin et al., 1989). Thus, having two putative transport systems and three C-compound utilization enzymes under positive cAMP:CRP control fits this hypothesis. In contrast, C-starvation-inducible proteins that are not dependent on cAMP:CRP for their induction are proposed to be required for long-term starvation-survival (Matin ef a/. 1989). This is supported by the fact that 12 SSR loci are negatively regulated by cAMP:CRP and, of these, five are required for starvation-survival. For most of these loci, knock-out mutations in cya and/or crp result in complete derepression of the locus, to induced levels, under non-inducing conditions. These include the dadA ( B ) operon, narZ( Y V W ) operon, rrg-1 to rrg-5 loci, rpoS and spvRlspvABCD. For three other loci, stiC, aldB and csiC, a cya or crp mutation results in partial derepression, to a level intermediate between fully induced and fully repressed levels, under non-inducing conditions. This indicates that cAMP:CRP is required, along with one or more other negative regulators, for their full repression. Thus, the aldB appears to require cAMP:CRP for its full induction during C-starvation and full repression under non-limiting conditions in minimal media (Spector and Cubitt, 1992; Gulig et al., 1993; Fang et al., 1996; Mahan e/ al., 1996; M. Spector, M. Pallen and G. Dougan, unpublished results). An interesting aspect of the negative regulation of narZ and stiC by cAMP:CRP, for example, is that the addition of exogenous cAMP or growth on CAMP-increasing C-sources (e.g. mannitol) does not prevent the induction of these loci under inducing conditions (Spector and Cubitt, 1992). Thus, cAMP:CRP-mediated repression does not appear to respond to increasing levels of cAMP and CRP, indicating that inducing conditions lead to derepression independent of cAMP:CRP levels. This is convenient since C-starvation-induced gene expression must occur in the presence of high cAMP:CRP levels. Curiously, in the case of dadA(B), repression by CRP is independent of cAMP in minimal medium

Table 2

Regulation of C-starvation-inducible SSR loci of salmonella.

Regulators* Other stresses/conditio s Known to induce locus

9

Genetic locus

cr’

cAMP:CRP

Others

csiA csiG aldB fadF gabD dadA kafE otsA narZ

P n-stat P 0 P n-stat P P P

P P p-fulljn-full P P n

PPGPP (PI ? Fis (n) PPGPP @); FadR (n)

?

?

?

rpoS spvR

0

P

n n

spvABCD

P

n

stiC

P

n

csiH csiC csiM stiG sfiH

0 0 n-stat

P n-full P

? ?

?

0

n

? ?

?

Epithelial Epithelial: P-starvation

P-starvation; HzOz; LB-log; r-ala Weak acids; Alkaline pH High salt Lrp (PI; H-NS (n) P- and N-starvation; HzOz; Epithelial: PPGPP(P); OxyR (n-full) ISM P- and N-starvation: Acid pH; High salt PPGPP(P): M v i ~ (RssB) (n) H-NS (n); Spv A(n); SpvR (p) PhoP (p) P- and N-starvation; Iron-starvation; Acid pH; Macrophage; Epithelial; ISM H-NS (n); SpvR @) PhoP (p) P- and N-starvation; Iron-starvation: Acid pH; Macrophage; Epithelial: ISM P- and N-starvation; Epithelial; ISM; PPGPP (PI Alkaline pH Epithelial ? 9 Epithelial PPGPP (P) ?

? ?

? ? ?

Table 2 . continued rrg- 1 rrg-2 rrg-3 rrg-4 rrg-5

P P P P P

n

9

n

? ? ?

n

n n

9

? ?

P-starvation

*Regulation (direct or indirect) during C-starvation and/or log-phase in minimal media. p = positive regulator required for induction; n = negative regulator required for repression under non-inducing conditions; p-full = required only for the full induction of the gene; n-full = required only for the full repression of the gene under non-inducing conditions; n-stat = negative regulation only under stressinduced stationary-phase; 0 = no role; ? = unknown. See text for references and further explanation. ?Epithelial = induced within MDCK cells: Macrophage = induced within macrophages; ISM = induced in intracellular salts medium; P-starvation = induced during phosphate starvation; N-starvation = induced during nitrogen starvation; H 2 0 2 = induced by 60 PM HzOz in log-phase; High salt = induced under hyperosmotic conditions; Acid or Alkaline pH = induced under acid or alkaline pH conditions; LB-log = induced during log-phase in LB medium but not minimal medium; 1.-ala = induced by presence of 1-alanine; ? = unknown. See text for references.

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MICHAEL P. SPECTOR

(Spector and Cubitt, 1992). The C-starvation-inducible and osmoticallyregulated otsA(B) operon does not appear to be regulated by cAMP:CRP (Fang et al., 1996). The finding that cAMP:CRP is a negative regulator of a number of starvation- /stationary-phase-expressed genes brings up an interesting side point, Those who have worked with cya and crp mutants know that they grow much slower than wild-type cells and form much smaller colonies on agar plates. The reason for this slow growth was unclear but one explanation was that they were unable to express various cAMP:CRP-dependent metabolic enzyme and/or transport systems. In light of the fact that cAMP:CRP is a negative regulator, during log-phase growth, of genes normally expressed in high amounts during starvation or stationary phase, another explanation for the slow growth of cya or crp mutants may be the inappropriate expression of these proteins under non-limiting growth conditions. For example, the effects of the inappropriate expression of csdependent cAMP:CRP repressed SSR genes. 2.3.1.2. RpoS or us An important subset of C-starvation-inducible loci comprises members of the rpoS-regulon. The importance of this regulon to the overall SSR is that rpoS itself and individual loci regulated by rpoS are critical to the development of long-term starvation-survival mechanisms (core SSR loci) and cross-resistant to other stresses (discussed in more detail later). Many C-starvation-inducible SSR loci identified in Salmonella are under the direct or indirect control by the alternative sigma transcription factor, us or 038 (or RpoS or KatF; Mulvey and Loewen, 1989; O’Neal et al., 1994; Fang et al., 1996; M. Spector, unpublished results; Table 2). Most os-dependent SSR loci require us for their induction during C-starvation. These include csiA, aldB, gabD(TP), katE, otsA(B). narZ( YWV), spvRl spvABCD, stiC and rrg-1 through rrg-5. The otsA and the rrg loci were originally identified as rpoS-regulated genes (Fang et al., 1996) and later shown to be C-starvation-inducible (M. Spector, unpublished results). The others were selected for specific phenotypes, e.g. C-starvation-inducible or attenuated virulence, and later shown to be us-dependent. It is not surprising then that several of these loci are needed for maximal starvation-survival and/or cross-resistance to other stresses. It is hypothesized that these loci are transcribed under C-starvation conditions by EoS holoenzyme complexes and thus are under direct us control. However, without data showing direct interaction with promoter regions and/or the identification of a us promoter consensus sequence, an indirect effect cannot be ruled out. In fact, there is evidence which suggests that the rpoS-dependent induction of spvA BCD occurs indirectly through the rpoS-dependent induction of spvR (a positive regulator of spvABCD) (Kowarz et al., 1994). Similar scenarios may exist for other rpoS-dependent C-starvation-inducible loci.

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os has also been found to play a negative role in the regulation of certain C-starvation-inducible genes. This is the case for three SSR loci identified in Salmonella, the csiC, dadA(B) and csiM loci (O’Neal et al., 1994; M. Spector, unpublished results). For each of these loci their induced levels of expression during C-starvation are about 2-, 5- and 2.5-fold higher in a rpoS background than in a wild-type rpoS background. Interestingly, their log-phase expression is not significantly different in these two genetic backgrounds. The most logical explanation is that os controls the expression of one or more negative regulators that reduce or limit the expression of these loci during C-starvation. These putative os-dependent repressor(s) seem to play a role in reducing the expression of two phase 0 SSR loci, csiC and csiM, as cells move from log-phase growth to starvation-elicited non-growth. In the case of dadA (stiB), which also exhibits 2-fold higher levels of induction during P-starvation in a rpoS background, the putative repressor appears to play a role in limiting the expression of dadA during C (or P) -starvation. It is not known why the cell would need to limit the expression of this locus. Perhaps excessive levels of the dadA gene product, D-amino acid dehydrogenase, or dadB gene product, alanine racemase, may be detrimental to the cell. Although a number of C-starvation-inducible loci of Salmonella are controlled by os, many others are not. These include rpoS itself, fadF, and two unclassified SSR loci, csiH and csiC. Hoyever, as mentioned above, all four are controlled by cAMP:CRP. Both rpoS and csiC are negatively regulated by cAMP:CRP while fadF and csiH are positively regulated by cAMP:CRP. 2.3.1.3. Other Regulatorji Systems Results from E. coli and Salmonella have implicated a number of additional regulatory networks associated with starvation- and stationary-phase-inducible gene expression (Spector and Cubitt, 1992; Gulig et al., 1993; Altuvia et al., 1994; O’Byrne and Dorman 1994b; O’Neal et al., 1994; Foster and Spector, 1995; Xu and Johnson, 1995; Hengge-Aronis, 1996; Seymour et al., 1996; M. Spector, C.C. DiRusso, M. Pallen and G. Dougan, unpublished results). One regulatory network common to many C-starvation-inducible SSR gene regulation is the stringent response (reviewed in Cashel et al., 1996). This is mediated by the accumulation of an unusual cellular nucleotide guanosine (penta or) tetraphosphate or (p)ppGpp (Gallant et al., 1976). The classical stringent response was described for cells starved for an amino acid. The lack of a particular amino acid leads to accumulation of its corresponding uncharged tRNA. As a result, translating ribosomes will stall when encountering a codon for the starved amino acid. Ribosome stalling in this manner is then thought to induce the RelA protein which converts GTP and ATP to pppGpp plus AMP. pppGpp is typically dephosphorylated by the product of the gpp gene. (p)ppGpp is degraded by the product of the SPOTgene to

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MICHAEL P. SPECTOR

GTP or GDP depending on the substrate. Interestingly, SpoT can also synthesize (p)ppGpp under certain circumstances. The accumulation of (p)ppGpp ultimately leads to a reduction in both ribosomal protein and RNA synthesis and a general increase in amino acid anabolism, both hallmarks of the stringent response. A variety of other complex changes in cell metabolism were also described (reviewed in Cashel et al., 1996). Thus, there is both a negative and positive arm of the stringent response. In addition to amino acid starvation, C- and P-starvation all directly or indirectly trigger (p)ppGpp accumulation and a stringent-like response. Several SSR loci exhibit positive control by (p)ppGpp in a relA-dependent manner during C-, and in some cases N-starvation. These include csiA,fUdF, dadA(B), narZ(YWV). stiC, as well as rpoS (Spector and Cubitt, 1992; Gentry et al., 1993; O’Neal et al., 1994; M. Spector, C.C. DiRusso, M. Pallen and G. Dougan, unpublished results). In the case of csiA, narZ and stiC, this may be an indirect effect since each is rpoS-dependent and rpoS itself is positively regulated by (p)ppGpp (Gentry et a/., 1993). For dadA (B), (p)ppGpp is believed to act directly to regulate dadAB expression. This interpretation is based on the facts that: (i) as is involved in the negative regulation of dadA ( B ) ; (ii) (p)ppGpp is a positive regulator; and (iii) a rpoS null mutation can suppress the effects of a relA null mutation on dadA expression (O’Neal et al., 1994). In the proposed model, asis needed for the expression of a putative dadAB repressor during C-starvation, and (p)ppGpp is needed to relieve this repression at the dadAB promoter. Thus, rpoS mutants and rpoS relA double mutants exhibit increased levels of dadA-lac expression because they do not synthesize sufficient levels of the putative repressor. Furthermore, relA null mutants show severely reduced CladA-lac induction during C-starvation because they cannot relieve the repression by the putative dadAB repressor. In the case of rpoS, studies with E. coli have shown that (p)ppGpp exerts its positive regulatory effect at the level of transcription-translation coupling, since the inability to make (p)ppGpp results in decreased levels of full-length transcript and increased transcriptional termination resulting from transcription/translation uncoupling (Hengge-Aronis, 1996). A similar mechanism is likely to exist in Salmonella. Additional regulatory circuits known to be involved in Salmonella SSR gene regulation include Fis, FadR, Lrp, H-NS, OxyR, MviA, SpvR and SpvA. The Fis protein is a member of a family of nucleoid-associated proteins that can bend DNA upon binding to certain sites. Fis levels are high in growing cells, dropping as the cell passes through late log-phase and into starvation-/stationary phase. Not surprisingly, based on its expression pattern, it is a negative regulator during log-phase growth, in both E. rol; and Salmonella, of aldB (Xu and Johnson, 1995; M . Spector, unpublished results). Another member of a family of nucleoid-associated proteins,

THE STARVATION-STRESS RESPONSE (SSR) OF SALMONELLA

259

H-NS (encoded for by the hns gene) is also found to negatively regulate several SSR loci in both E. coli and Salmonella during exponential growth. Unlike Fis, however, H-NS protein levels (which are fairly high in log-phase cells) continue to increase as cells enter stationary phase, even though H-NS seems to negatively autoregulate itself (Dersch et al., 1993). H-NS seems to function as a negative regulator of starvation-/stationary-phase-induced gene expression only in exponentially growing cells since such genes become derepressed postexponentially, suggesting another function for H-NS in stationary-phase or starved cells. SSR loci negatively regulated by H-NS in Salmonella include the otsA(B) and spvRlspvABCD (O’Byrne and Dorman, 1994b; Hengge-Aronis, 1996). Intriguingly, studies in E. coli show that H-NS appears to negatively regulate the expression of rpoS but at some point following transcription of the rpoS gene (Barth et al., 1995). Therefore, the negative regulation of otsA and spvR/spvABCD by H-NS may be indirect, resulting from its negative effect on as expression (as being a positive regulator of both). The role of Fis and H-NS as well as another nucleoid-associated protein, IHF (encoded for by the himA and himD genes), in the regulation of other Salmonella SSR loci is yet to be determined. Like H-NS, I H F is found to regulate starvation-/stationaryphase-inducible gene expression and to increase in expression in stationary phase in E. coli (Hengge-Aronis, 1996). The FadR protein is a negative regulator of fadF and P-oxidation in Salmonella (M. Spector, C . C . DiRusso, M. Pallen and G. Dougan, unpublished results). Recently, studies in E. coli have also shown that FadR is also a negative regulator of the uspA gene which encodes universal stress protein A, a serine/threonine phosphorylated cytoplasmic protein. UspA is also positively regulated by (p)ppGpp (Farewell et al., 1996). A uspA homolog of Salmonella has not yet been identified. Otherfad genes have also been shown to be induced during stationary phase in LB and to be repressed by the FadR protein in E. coli (Farewell et al., 1996). The coordinate regulation of fatty acid degradation and a protein required for general stress resistance is very interesting. It suggests that fatty acid/phospholipid metabolism may have a broader role in the SSR than the mere utilization of fatty acids as carbon-/energy-sources. In addition, the leucine-responsive regulatory protein Lrp has been implicated as a positive regulator of otsA (Hengge-Aronis, 1996). Lrp is a regulatory protein that appears to respond to nutritional downshifts by turning-on a number of amino acid biosynthetic operons, degradative operons and other metabolic functions (reviewed in Newman e f al., 1996). A role for Lrp in the regulation of other Salmonella SSR loci has yet to be established. As mentioned previously, expression of the spv virulence plasmid locus of Salmonella possesses its own specific regulatory elements. The SpvR protein

260

MICHAEL P. SPECTOR

is a positive regulator of its own transcription and the spvABCD operon. In addition, SpvA is found to function as a negative regulator of spvR; providing negative feedback control under certain circumstances (Gulig er al., 1993; Wilson et al., 1997; P. Gulig, personal communication). It has been postulated that SpvA negative regulation of SpvR and in turn spvABCD expression has evolved to limit the expression of these genes under inducing conditions either outside the host or at sites within the host where their expression is not needed (P. Gulig, personal communication). Another transcriptional regulator involved in SSR gene expression is the oxyR gene product. OxyR acts as both a sensor and a regulator protein during oxidative stress resulting from H202 exposure. It is best characterized as a transcriptional activator of a subset of HzO2-inducible proteins, the OxyR regulon (Storz and Altuvia, 1994). The OxyR regulon is essential to the development of H202-inducible adaptive H202 resistance (discussed further below). The OxyR protein can be found in two forms in the cell, a reduced form and an oxidized form. Both forms bind to DNA but at different sites. In the presence of H202,OxyR becomes oxidized at key cysteine residues leading to a conformational change allowing it to act as a transcriptional activator of a number of genes in response to H202-mediated oxidative stress (Storz and Altuvia, 1994), including the dps gene in E. coli (Altuvia et al., 1994). OxyR, in both reduced and oxidized forms, also negatively regulates its own transcription and a Mu phage gene mom; however, the oxidized form is much more efficient at repressing transcription of these genes than is the reduced form (Storz and Altuvia, 1994). Recently, OxyR was also shown to regulate the induction by H 2 0 2 of narZ ( s t i A ) (Seymour et a/., 1996). OxyR regulation of narZ is different than that described for clps and other genes where the oxidized form is found to act as a transcriptional activator or repressor protein. In the case of narZ it is the reduced form (i.e. the form predominantly present in the absence of exogenous H202)that acts as a repressor. In log-phase cells, OxyR represses narZ transcription (along with cAMP:CRP) in the absence of exogenous H202. In the presence of exogenous H202, OxyR becomes oxidized and either falls off its narZ binding site or moves to a different site resulting in the partial derepression of the narZ gene. Induction of narZ-lac fusions by H202 is only about one-third of that achieved by C-starvation, indicating that cAMP:CRP still negatively regulates narZ expression under these conditions (Seymour et al., 1996). Thus, nnrZ expression undergoes derepression rather than transcriptional activation, as is the case for d’ps and other Oxy R-dependent genes, in response to H202. An interesting level of oS regulation is control of its stability. This involves a putative two-component response regulator system. However, only the regulator component of this system has been identified and is referred to as MviA in Sulmonella and RssB in E. coli. The mviA gene

THE STARVATION-STRESS RESPONSE (SSR) OF SALMONELLA

261

was originally identified as a regulator of S. typhimurium virulence in Ztys mice (reviewed in Foster and Spector, 1995). Recent evidence suggests that MviA, at least partially, regulates virulence by controlling as stability. One hypothesis is that MviA inhibits a protease (ClpXP) involved in as degradation during starvation and other inducing stresses (e.g. acid pH) allowing as to accumulate and turn-on as-dependent gene expression. This hypothesis is supported by findings that, in mviA null mutants, as accumulates in log-phase non-stressed cells. However, the exact mechanism of MviA regulation of os stability is still unclear (Bearson et al., 1996). Clearly, the fact that the C-starvation-inducible loci characterized thus far do not show a general pattern of regulation indicates that individual SSR loci exhibit complex overlapping and unique multileveled regulation. For example, they are not all rpoS-dependent or -independent or under positive or negative control by cAMP:CRP, etc. Thus, it is difficult to look at the regulation of one gene and use it as a paradigm for the regulation of genes under these conditions.

2.3.2. Environmental and Physiologic Regulation of SSR Loci In addition to C-starvation, most SSR loci are induced in response to one or more environmental and physiologic stresses or conditions. These include starvation for other essential nutrients such as phosphate, nitrogen or iron, extremes in pH or osmolarity, and intracellular phagosomes of epithelial cells or macrophages. 2.3.2.1. Other Defined StresseslConditions Several of the SSR loci identified in Salmonella are induced under other starvation conditions. The fadF, dadA(B), narZ( Y W V ) , stiC, rrg-5, spvRlspvABCD and the rpoS genes all show increased expression during P-starvation. For dadA( B), narZ( Y W V ) , stiC, rrg-5 and spvRlspvABCD, P-starvation induction is as-dependent. The narZ( Y W V ) , stiC, spvRlspvABCD and rpoS loci are also N-starvation-inducible and, except for rpoS, this N-starvation induction is again os-dependent (O’Neal et al., 1994; M. Spector and P. Gulig, unpublished results; M. Spector and F. Fang, unpublished results). Furthermore, the induction of dadA, narZ and stiC during P-starvation, and N-starvation for narZ and stiC, also requires (p)ppGpp. However, only the N (and C)-starvation induction of these loci is relA-dependent (Spector and Cubitt, 1992; O’Neal et al., 1994). Their P-starvation induction appears to require the spoT-encoded (p)ppGpp synthetase activity (M. Spector, unpublished results; Cashel et al., 1996). In addition to P- and N-starvation, the spvRlspvABCD virulence locus is also induced during iron starvation (Gulig et al., 1993). Iron regulation of the other loci has not yet been shown.

262

MICHAEL P. SPECTOR

A number of SSR loci have also been found to respond to extremes in pH. For example, the spv genes and rpoS are all induced at low pH (Foster and Spector, 1995; Mahan et ul., 1996). It is not known if spv gene induction under this condition is rpoS-dependent; thus, their induction at acid pH may be an indirect effect. I t is interesting to note however, that even though as levels increase under acid pH conditions, not all as-dependent genes are acid pH-inducible. In contrast, at least two SSR loci have been shown to be alkaline pH-inducible, kutE and stiC (I.-S. Lee and M. Spector, unpublished results). It is not known if this induction is as-dependent. Two SSR loci have also been shown to be inducible by H202, cludA(B) and narZ( Y W V ). Both are also needed for H202-inducible adaptive H202 resistance (Seymour er ul., 1996). They are both induced during log-phase growth in the presence of 60 V M H202.H 2 0 2is found to induce both loci to about one-third the level observed for their C-starvation induction. However, only nur2-luc expression is regulated by OxyR (via derepression resulting from oxidation of the OxyR protein; discussed previously). Therefore, dudA(B) represents an oxyR-independent H202-inducible locus (Seymour er ul., 1996). The mechanism of the induction by H 2 0 2 of dudA is at present unknown. Lastly, as and the as-dependent gene otsA are induced under conditions of high osmolarity (Hengge-Aronis, 1996). Not surprisingly, both are needed for development of osmotolerance in both E. coli and Sulmonellu (Hengge-Aronis, 1996; Fang el ul., 1996; M. Spector, unpublished results). 2.3.2.2. Intracellular Environments The fact that Sulmonella is an intracellular pathogen allows us to examine the role of SSR during growth within cultured cells in vitro and/or host environments in vivo. Several of the Sulrnonellu SSR loci are induced within cultured epithelial cells and macrophages in vitro as well as within host cells in vivo. In addition, a defined medium believed to mimic the intracellular milieu of mammalian cells, called intracellular salts medium or ISM, has also been employed to study the expression of some of the SSR loci of Sulmonellu. A number of C-starvation-inducible SSR loci with diverse functions are induced within cultured epithelial cells. These include a putative ABC transporter protein (csiC), a fatty acid degradation enzyme vide,a cryptic nitrate reductase activity (narZYWI/), a putative inner membrane-associated starvation-inducible general stress resistance protein (stiC), a hypothetical inner membrane-associated protein (csiH), an unknown gene product (csic), and several Salmonella plasmid-associated virulence functions ( . T ~ V R / S ~ V A B(Wilson C D ) et al., 1997; F. Garcia-dC1 Portillo, M. Spector and B. Finlay, unpublished results). Of these, the spv genes have also been shown to be induced within macrophages (Wilson et ul., 1997). However, it should be noted that the other loci have not yet been tested for macrophage induction. The only known common feature of all these

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THE STARVATION-STRESS RESPONSE (SSR) OF SALMONELLA

genes is their C-starvation-inducibility; unfortunately, not all C-starvationinducible loci are induced under these conditions. Thus, it is difficult to make concrete assessments of the intracellular environment. We can hypothesize that the intracellular environment may generate similar signals to C-starvation to which some, but not all, C-starvation-inducible loci may respond. The studies of Gulig and colleagues have provided some interesting insights into the intracellular environment and the expression of certain SSR loci (Wilson el ul., 1997; M . Spector, J. Wilson, and P. Gulig, unpublished results; P.Gulig, personal communication). These studies have examined the expression of the spv genes, nurZ (sfiA)and stiC in the intracellular salts medium, ISM. All three loci are induced during log-phase in ISM compared with log-phase growth in LB medium. However, only sfiC was induced further upon entry into stationary phase in ISM. For spvA(BCD), the expression during exponential growth in ISM is both SpvR- and osdependent. The expression of .stiA and sfiC under these conditions is also likely to be os-dependent, although this has yet to be confirmed (O’Neal et ul., 1994; Wilson et ul., 1997; M . Spector, J . Wilson and P. Gulig, unpublished results). These findings agree with induction patterns for these three loci within cultured epithelium cells. Thus, further dissection of ISM to find the inducing signal(s) may give insight into the inducing signal(s) present in the intracellular environment. Mahan et ul. (1993, 1995) developed a creative technique for identifying Sulmonrllu genes induced during, and/or required for, growth in vivo. This technique, referred to as in vivo expression technology or WET, couples the use of a purine auxotroph of S . typhimurium which grows poorly in host mice and a library of suicide plasmid vectors containing cloned fragments of S. typhimurium chromosomal DNA in front of promoterless purA and lucZY+ genes. Other genes, thyA and cut, have also been employed to vary the in vivo selectable marker. Thus, cloned fragments possessing promoters of genes expressed in vivo restore virulence to the strain due to complementation of the purine auxotrophy. Bacteria recovered from the infected mouse are then tested in vitro on lac indicator plates such as MacConkey agar plates to identify fusions exclusively expressed in vivo. Any relationship with loci identified by this method and the SSR remains to be explored. However, one interesting finding is that one of the in vivo induced (ivi) fusions identified was in fudB, a gene required for fatty acid degradation. As mentioned above, the .fudF gene is a SSR locus and is induced within cultured epithelial cells. The fudB and fudF genes exhibit very similar regulation (reviewed in Clark and Cronan, 1996). Although, the f u d B gene has not been identified as a SSR locus, its similar regulation to fudF provides support for a potential link between certain aspects of the SSR and Sulmonella pathogenesis. +

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3. THE SSR AND LONG-TERM STARVATION SURVIVAL

Results from 2-D PAGE analysis, operon fusion analysis, and other studies discussed below all indicated that long-term starvation survival of Salmonellu requires new or increased expression of genes and proteins, i.e. induction of the SSR stimulon. Studies with the protein synthesis inhibitor chloramphenicol (Cam) showed that Cam addition during the transition from log-phase to starvation-induced stationary-phase or after 24 h of starvation all significantly decreased starvation-survival after 10 days of starvation. The earlier the Cam was added, the more dramatic was the effect on starvation-survival (Spector and Foster, 1993). From these experiments it was concluded that the synthesis of new or increased production of existing proteins during the first 4 6 h of starvation was most critical for long-term starvation-survival. However, the continued synthesis of these or expression of new proteins even after 24 h of starvation is also important to achieve wild-type starvation-survival levels. This is very interesting because it indicates that proteins induced within the first 4-6 h of starvation are essential for survival of the bacteria after days or even months of starvation. This agrees with similar results described for E. coli (Matin et al., 1989).

3.1. Core SSR Loci

When one examines the starvation-survival curve for typical laboratory or virulent strains of Salmonella, several general features can be seen (Spector and Cubitt, 1992). First, cell density achieved at the onset of starvation, under the conditions tested, is approximately 3-5 x 10’ colony forming units (cfu)/ml. Over the next 48 h of starvation there is little loss of viability of the culture, as indicated by the fact that cfu/ml. values remain fairly constant. However, between three and 12 days of starvation, culture viability begins to decline precipitously to a level of around 3-8% of the maximum viability observed for the culture. This level of viability (or survival) has been maintained for up to six months of starvation (the longest period of starvation tested). We know from chloramphenicol studies that this ‘typical’ starvation-survival curve requires protein synthesis. Thus, one of the goals of the SSR is to allow Salnionella to survive long-term starvation and maintain some level of bacterial cell viability over prolonged periods of starvation. Therefore, we have coined the term core SSR locus to describe a genetic locus whose product(s) is required for starvation-survival. At least five or six core SSR loci have been described in Salmonella; narZ ( Y W V ) , dadA ( B ) , .stiC, rrg-l and rpoS (Spector and Cubitt, 1992; O’Neal ef al., 1994; Fang t’t d . ,

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1996). In addition, afadF‘mutation was found to significantly reduce starvation-survival over the first 10 or so days of starvation but did not effect the ultimate steady-state level of viability achieved by the culture (M. Spector, C.C. DiRusso, M. Pallen and G . Dougan, unpublished results). In the case of narZ (stiA), dadA ( s t i l l ) , sriC and rpoS, individually each mutation reduces the level of starvation-survival by about 75 (f25)-fold after about 20 days of starvation. Interestingly, when the sriA, B or C mutations were combined in the same background, starvation-survival was reduced some 500- to 2000-fold depending on which mutations were combined (Spector and Cubitt, 1992). However, when the rpoS null mutation was combined with the three s t i mutations either individually or in combination there was no additional effect on starvation-survival (O’Neal et a/., 1994). This was surprising since stiA and stiC are rpoS-dependent and os is thought to regulate additional loci that are required for starvationsurvival. Why then does deleting the required positive regulator function for a number of these putative core SSR loci not (i) reduce starvation-survival, at least, to the same level as deleting the functions of two core SSR loci together or (ii) have any additional effect when combined with mutations in other core SSR loci? The answers to these questions are still unclear. One possible explanation is that basal levels or os-independent levels of expression of these core SSR loci can contribute to starvation-survival. Another possibility is that the loss of os-regulation may trigger cryptic or unidentified mechanism(s) of control or redundant functions that can partially compensate for the loss of oS function. However, if two (or more) core SSR loci are specifically knocked out then starvation-survival is more drastically affected. Recently we have found that fadF mutants, compared with wild-type strains, lose viability more quickly over the first two weeks of starvation and exhibit about a 7- to I0-fold lower level of survival than wild-type cells after this period. However, after about 20 days of starvation there was no significant difference between the wild-type parent strain and the fudF mutant strains (M. Spector, C.C. DiRusso, M. Pallen and G. Dougan, unpublished results). This suggests that the ability to degrade medium/ long-chain fatty acids is critical to starvation-survival during the first few days of starvation but does not appear to be important to establishing the low level population of viable cells observed in cultures starved for 20 days or more. Fang er al. (1996) showed that the rrg-1 locus appears to be involved in starvation-survival. In that paper, survival of strains carrying insertions in the various rpoS-regulated genes and a rpoS null mutation were compared with the wild-type strain after 14 days in stationary-phase following logphase growth in rich (LB) medium. Results from these experiments demonstrated that, although all five of the rrg insertion mutations seemed to reduce

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MICHAEL P. SPECTOR 100

Ill

2 'E

z 1 Y

8b

&I

0.1

0.01

TbCllllal

H202

Hyperosmotic

Acid pH

Polymyxin

Challenge Stress Figure 1 Carbon-starvation-inducible cross-resistance to thermal, oxidative, hyperosmotic, acid pH and polymyxin challenge. In all cases, Sulnione//a mterico, serovar Typhimurium strain SL1344 was grown in minimal media with non-limiting glucose to generate log-phase cells or limiting glucose to generate 5 h and 24 h C-starved cells. Logphase, 5 h C-starved, and 24 h C-starved cells were then challenged with one of the following stresses: 55'C for 20 min (Thermal), 15 mM Hz02 for 40 min (HzOz), 2.5 M NaCl for 48 h (Hyperosmotic), pH 2.7 for 45 min (Acid pH), and 100 pg of polymyxin B for 60 min (Polymyxin). Percent survival is the number of viable cells at termination of challenge divided by the number of viable cells at the beginning of the challenge multiplied by 100.

survival under these conditions, only the rrg-1 insertion reduced survival to the same extent as the rpoS mutation. This suggests that the r g - 1 locus may represent another core SSR locus. However, this will need to be confirmed by determining its survival profile during more defined C-starvation conditions.

4. THE SSR AND RESISTANCE TO OTHER ENVIRONMENTAL

STRESSES

For Salmonella, E. coli and other bacteria, C-starvation not only induces functions needed for long-term starvation-survival. core SSR genes, but it

Table 3

Differential requirements for SSR loci in stress-specific and/or C-starvation-inducible resistance to environmental stresses in Salmonella. ~~

~

Challenge stress* Genetic locus dadA narZ sric fadF rrg- I rrg-5

karE otsA rpoS

HI02

Starvation ( 1 0 1 days) (15 mM)

ss ss ss ss ss Nr Nd Nr

ss

Thermal (55 “C)

ss ss

CSI CSI

CSI Nd Nd CSI CSI Nd

Nd Nd Nd Nd CSI

ss; CSI

ss; CSI

ss; CSI

H ypersomotic

( 2 . 5 v NaCI) Nr Nr CSI Nd Nd Nd Nd CSI

ss; CSI

Acid (pH 2.7) Nr

CSI (early) Nr Nd Nd Nd Nd Nd

ss; CSI

*SS = required for stress-specificadaptive resistance to indicated stress; CSI = required for C-starvation-inducible cross-resistance to indicated stress; Nr = Not required; Nd = Not done. See text for further explanation and references.

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P. SPECTOR

can also induce cross-resistance to other environmental stresses. This is in addition to the stress-specific adaptive resistance mechanisms that have been characterized for some stress conditions (Foster and Hall, 1990; reviewed in Neidhardt and VanBogelen, 1987; Matin, 1991; Storz and Altuvia, 1994; Foster and Spector, 1995; Gross, 1996). C-starvation-inducible cross-resistance of Salmonella to various other defined stresses is presented in Fig. 1. A discussion of the roles of various SSR loci in both stress-specific and C-starvation-inducible cross-resistance to five different defined stresses is presented below and summarized in Table 3.

In Salmonella, resistance to H 2 0 2can be induced by both exposure to sublethal levels of H202(stress-specific; SS) or C-starvation (C-starvation-inducible; CSI). Analysis of the roles of various SSR loci in both these mechanisms has revealed some interesting findings (Seymour et u/., 1996; M. Spector, S.M.D. Bearson, M. Pallen, G. Dougan and J. Foster, unpublished results; V. Makam Nataraj, A. Mahmud and M. Spector, unpublished results). As alluded to previously, the rpoS, narZ (stiA) and dudA ( s t i l l ) loci were all found to be required for the development of H202-inducible adaptive H 2 0 2resistance. Both a rpoS and narZ mutant are hypersensitive to H 2 0 2 in log phase and neither develops resistance to a 40 min challenge with H202 following a 60 min incubation with 60 ~ L MH202 (adapting conditions). The dadA mutant was not hypersensitive to H202 in log phase compared with a wild-type strain but it did exhibit a significantly reduced level of survival after a 40 min challenge with H 2 0 2following incubation under adapting conditions, although the effect of the mutation was delayed. Neither narZ nor dadA appeared to be required for C-starvation-inducible cross-resistance to H 2 0 2 , suggesting that other factors can compensate for their loss during starvation (Seymour c)t al., 1996). It is unknown why the functions encoded by narZ or dadA are required for resistance to H202. One possible explanation is that they are involved in detoxifying some by-products of H202-mediated oxidative damage, but that has yet to be shown. The rpoS, katE, stiC and rrg-5 genes are all needed for the development of CSI H202 cross-resistance. Strains carrying mutations in any of these loci fail to induce cross-resistance to a 40 min H 2 0 2 challenge following either 5 h or 24 h of C-starvation. Interestingly, development of CSI H 2 0 2 resistance is independent of the o.uyR gene product, which is essential for the development of the adaptive response (Storz and Altuvia 1994; Seymour et a/., 1996; M. Spector, unpublished results). Thus, the H202-inducible adaptive resistance and CSI cross-resistance mechanisms exhibit both unique and

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269

overlapping features. The function of katE in H202 resistance is to degrade the H 2 0 2before it can produce oxidative damage, but clearly more than the production of a catalase activity is required to establish resistance to H 2 0 2 challenge. At present, the functions of stiC and rrg-5 are not known.

4.2. Thermotolerance

As for H2O2 resistance, Salmonella possesses both stress-specific and Cstarvation-inducible resistance, or thermotolerance, to heat challenge (55 "C). Strains carrying null mutations in either rpoS, dadA, narZ or stiC were all hypersensitive to a 20 min challenge at 55 "C in log phase and all failed to induce a C-starvation-inducible thermotolerance response. In addition, the rpoS and stiC mutant failed to mount a heat-shock (42 "C)inducible thermotolerance in log-phase cells (M. Spector, S.M.D. Bearson, M. Pallen, G. Dougan and J. Foster, unpublished results; V. Makam Nataraj, A. Mahmud and M. Spector, unpublished results; for reviews, see Neidhardt and VanBogelen, 1987; Matin, 1991; Gross, 1996). Thus, there is a clear link between the SSR and thermotolerance. The roles these loci play in thermotolerance is unclear at this time. As in H 2 0 2 resistance, they may play roles in detoxifying by-products of a 55 "C challenge, e.g. damage to membrane components or accumulation of D-amino acids resulting from isomerization or protein degradation. Furthermore, Fang et a/. (1996) have suggested that otsA plays a minimal role in C-starvation-inducible thermotolerance, indicating that the synthesis of periplasmic trehalose may be important for thermotolerance as well as for osmotolerance.

4.3. Osmotolerance As with resistance to the above stresses, salmonellae possess both stressspecific and C-starvation-inducible mechanisms of tolerance to hyperosmotic stress. At least two SSR loci were found to be important for the development of C-starvation-inducible osmotolerance, rpoS and otsA . Strains carrying mutations in these loci do not develop C-starvationinducible osmotolerance. The rpoS and otsA loci also play a role in the stress-specific (0.35 M NaC1)-inducible osmotolerance mechanism. The role of orsA(B) in osmotolerance is in the synthesis of periplasmic trehalose, which serves as an osmoprotectant for the cell (Fang et al., 1996; V. Makam Nataraj, R. Ellison and M. Spector, unpublished results). The function of RpoS is, as for other stresses, one of the regulator of both osmotically- and C-starvation-inducible genes.

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4.4. Acid Tolerance

Again, Salmonellu has both a stress-specific and C-starvation-inducible acid tolerance response. The stress-specific response known as the adaptive acid tolerance response (ATR) is complex and is partially dependent on rpoS (Foster and Hall, 1990; Foster, 1991; Foster and Spector, 1995; Bearson et a/., 1996). No other SSR loci have been shown to play a role in the adaptive ATR; however, only a few have been tested under these conditions. The rpoS gene is also required for the C-starvation-inducible ATR, indicating again that, as with the other stress resistance mechanisms, rpoS-regulated genes are critical to the generation of C-starvation-inducible cross-resistance to acid pH. For example, strains carrying the m r Z null mutation failed to induce maximal tolerance to a 45 min challenge with pH 2.7 following 5 h of C-starvation but was not as critical for acid tolerance devcloped in 24 h Cstarved cells. An interesting side note is that 24 h C-starved wild-type cells are about 40-fold less acid-tolerant than 5 hr C-starved wild-type cells, which may explain why we did not detect a significant effect of the izurZ mutation in 24 h C-starved cells (M. Magut and M. Spector, unpublished results). Unfortunately, again, the function of nczrZ in acid tolerance is not known at this time.

4.5. Polymyxin Resistance

Recently we have also demonstrated that C-starvation, as well as P- and Nstarvation and LB-stationary phase, produces cells that are resistant to the membrane-permeabilizing effects of the antimicrobial peptide polymyxin B (McLeod and Spector, 1996). More interestingly, this resistance is os-independent. Its os-independence and the fact that P-starvation induces equivalent levels of polymyxin resistance indicate that this resistance is different from the cross-resistance described for other stresses (described previously), since they are os-dependent, and P-starvation typically induces only minimal resistance to these other stresses. The polymyxin resistance we identified does involve both phoP-dependent (Groisman et ul., 1989; Miller pi a/., 1989) and phoP-independent pathways. The phoP-dependent mechanisms play a role in log-phase cells and during the first few hours of C-starvation but decline in importance as cells continue to starve until they have no role in 24-h C-starved cells. The greatest effect of a phoP null mutation is early in LB stationary phase in Luria--Bertani (LB) medium. The phoP-independent mechanisms are essential for polymyxin resistance upon entry into C-starvation-induced stationary phase through the first 24 h of C-starvation (similar results are seen in P-starved and LB stationary-phase cells). Unfortunately, we have not examined cells that have been starved for longer

THE STARVATION-STRESS RESPONSE (SSR) OF SALMONELLA

27 1

than 24 h for polymyxin resistance to determine what happens beyond the 24 h of starvation point. Characterization of the phoP-independent mechanisms is currently under investigation.

5. THE SSR AND Salmonella VIRULENCE

Starvation-stress has yet to be directly linked to the virulence of Salmonellu and the expression of virulence factors within the host organism. We know from studies in vitro, however, that several loci involved in virulence are induced during C-starvation or similar conditions and can play a central role in the SSR of Salmonclla. These include the rpoS, crp and the spv genes. As described earlier, several SSR loci are also induced by other environmental stresses, such as extremes in pH or osmolarity and the presence of H 2 0 2 , which suggests that virulence factor expression may be geared to multiple stresses occurring in various niches within the host (Fang et al., 1992; Gulig et a1.,1993; Mahan et a/., 1993, 1995, 1996; Bearson rt ul., 1996; Seymour et a/., 1996; Wilson et a/., 1997). Moreover, one might hypothesize that having multiple environmental stresses/signals being capable of regulating virulence factor expression may help to ensure the appropriate expression of these functions at the desired times within the host. The roles that rpoS and crp have in Salmonella virulence appear to be to regulate virulence factor expression, e.g. the spv genes. For the spv genes, both regulators are thought to act primarily through SpvR to regulate spvABCD expression (Gulig et a/., 1993; Kowarz et a/., 1994; O’Byrne and Dorman, 1994a). The spvABCD operon gene products in turn are required for the systemic pathogenesis of non-typhoidal Salmonella serovars (Gulig et al., 1993). Evidence from our laboratory and others (A. Turk, P. Gulig and M. Spector, unpublished data; Gulig et a/., 1993; O’Byrne and Dorman 1994b) have shown that s p v R l ~ p v A B C Dare induced during C - , P- and Nstarvation as well as stationary phase in LB. Recent evidence has provided some new insights into spvR gene regulation and in turn spvABCD expression. Gulig and colleagues (Wilson et al., 1997; J . Wilson and P. Gulig, personal communication) have shown that the spvR gene and spvABCD operon are also induced during exponential growth in ISM medium as well as within epithelial cells and macrophages. In addition, SpvA protein which was previously proposed to be a negative regulator of spvR expression (Gulig et al., 1993) was recently found by Gulig and co-workers to negatively regulate spvR expression only under certain conditions, e.g. stationary phase in LB. Interestingly, deletion of the spvA gene alone did not affect spvR expression during growth in ISM medium. These findings suggest that

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MICHAEL P. SPECTOR

spv gene expression is controlled by different regulatory factors under different growth conditions. This hypothesis is supported by new data from this laboratory (Wilson el al., 1997; P. Gulig, personal communication) showing that spv gene induction in ISM is PhoP-dependent but induction during stationary phase in LB is PhoP-independent. However, spv gene induction under both these conditions is rpoS-dependent. One might hypothesize then that, within host cells, slow or non-growth is not an important signal for spv gene expression and that spv gene expression under slow or non-growth conditions is blocked by the negative feedback action of the SpvA protein. Add to this the role of PhoPQ, as and cAMP:CRP in spv gene expression under different conditions and one thing becomes clear - the regulation of just this one virulence locus is extremely intricate and complex. Abshire and Neidhardt (1993), based on 2-D PAGE analysis of S . typhimurium proteins expressed intracellularly within macrophages, suggest that there are two populations of bacteria within infected macrophages. One population is actively growing and the other is static. They estimated that the growing bacteria have a generation time of about 40 min but represent only a small percentage of the intracellular bacteria. Therefore, most of the intracellular salmonellae are not actively growing, but are viable. The individual importance of these two populations of cells to the virulence of Salmonella is not evident. One might expect, however, that in the context of the findings of Gulig and co-workers (Wilson ci al., 1997; J . Wilson and P. Gulig, personal communication) the growing population would be expressing the spv genes at high levels and thus would be more likely to spread and lead to systemic disease. In contrast, the non-growing population would be expressing low levels of the spv genes but presumably several of the starvation-/stationary-phase-/stressinducible proteins, making these cells highly resistant to a number of environmental stresses and allowing them to persist locally, triggering inflammatory responses in the host tissue and so forth. Thus, Salmonella pathogenesis may represent a tale of two cell populations; if or where the SSR fits in to this tale remains to be determined.

6. Concluding Remarks

The purpose of this review was to summarize and place into perspective what is known concerning the response of Salmonella to starvation-stress. As a result, some information obtained in E. coli was not presented or only briefly referred to in the text. However, what should be clear from this review is that the cumulative knowledge of studies in both Salmonella and E. coli, as well as other bacteria, has allowed huge leaps to be made in the understanding of starvation and stationary-phase physiology over the past

THE STARVATION-STRESS RESPONSE (SSR)OF SALMONELLA

273

few years. Fortunately, the apparent increasing complexity and interrelationship of the SSR and other stress responses and the ever-increasing number of SSR and stress-associated regulatory networks, should provide the fodder for study for many years to come. Continued understanding of starvation/stationary-phase physiology and genetics is invaluable since bacteria spend the vast majority of their existence under conditions that promote the expression of SSR and stress-inducible genes. Knowledge obtained from these endeavors may contribute to the development of new vaccines, new disease therapies, and new antimicrobial agents, as well as advances in biotechnology. Furthermore, the many research accomplishments already seen - and those yet to come - should continue to provide a greater understanding of host-pathogen relationships, and the fundamental biology of bacterial and mammalian cells. Thus, the study of the starvation-stress responses will continue to be an exciting area of basic and applied biomedical research.

ACKNOWLEDGEMENTS The author would like to thank his many colleagues and students who have contributed ideas and results over the years on whose hard work this review was based. I would also like to thank those colleagues who provided results and other information prior to publication as well as useful discussions. Some of these individuals in particular include Drs John Foster, Mark Pallen, Ferric Fang, Stephen Libby, Paul Gulig, Francisco Garcia-del Portillo, Brett Finlay and Concetta DiRusso. I would particularly like to thank Drs Mark Pallen, Gordon Dougan, John Foster and Shawn Bearson for their recent help in determining the identity of some of the genes discussed in this review. Also, due to space considerations, recent review articles or chapters were used to reference certain information. This was not meant to disregard or slight the importance of the work performed in the original publications. Those interested are encouraged to go to the cited reviews to obtain the citations for the original papers. Portions of the work performed in the author’s laboratory and cited in this review were supported by grant awards from the National Institutes of Health (GM47628 and AI/OD41170-01) and University of South Alabama Research Committee (3-61366, 3-61375 and 3-61407).

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Almiron, M., Link, A,, Furlong, D. and Kolter, R. (1992) A novel DNA binding protein with regulatory and protective roles in starved Escherichia coli. Genes Dev. 6, 2646-2654. Altuvia, S., Almiron, M., Huisman, G., Kolter, R. and Storz, G. (1994) The (Ips promoter is activated by OxyR during growth and by I H F and 0’ in stationary phase. Mol. Microhiol. 13, 265-272. Barker. J. and Brown, M.R.W. (1994) Trojan horses of the microbial world: protozoa and survival of bacterial pathogens in the environment. Microbiology 140, 1253-1259. Barrett, E.L. and Riggs, D.L. (1982) Evidence for a second nitrate reductase activity that is distinct from the respiratory enzyme in Salmonella typhimurium. J. Bacteriol. 150, 563--571. Barth, M., Marschall, C., Muffler, A,, Fischer, D. and Hengge-Aronis, R. (1995) Role for the histone-like protein H-NS in growth phase-dependent and osmotic regulation of 0’ and as-dependent genes in Escherichia coli. J. Bacteriol. 177. 3455-3464. Bearson, S.M.D., Benjamin, W.H., Jr, Swords, W.E., and Foster, J.W. ( I 996). Acid shock induction of RpoS is mediated by the mouse virulence gene mviA of Salmonella typhimurium. J. Bucteriol. 178, 2512-2519 Behlau, I. and Miller, S.I. (1993) A PhoP-repressed gene promotes Salmonella typhimuriurn invasion of epithelial cells. J. Bacteriol. 175, 4475 4484. Berlyn, M.K.B., Low, K.B., Rudd, K.E. and Singer, M. (1996) Linkage map of Escherichia coli K-12, Edition 9. In: Escherichia coli und Salmonella Cellulur and Molecular Biology (F.C. Neidhardt, R. Curtiss, 111, J.L. Ingraham, E.C.C. Lin, K.B. Low, B. Magasanik, W.S. Reznikoff, M. Riley, M. Schaechter and H.E. Umbarger, eds), pp. 1715-1902. ASM Press, Washington, DC. Boos, W. and Lucht, J.M. (1996) Periplasmic binding protein-dependent ABC transporters. In: Escherichia coli and Salmonella Cellular and Molecular Biology (F.C. Neidhardt, R . Curtiss, 111, J.L. Ingraham, E.C.C. Lin, K.B. Low, B. Magasanik, W.S. Reznikoff, M. Riley, M. Schaechter and H.E. Umbarger, eds), pp. 1175- 1209. ASM Press, Washington, DC. Botsford, J.L. and Drexler, M. (1918) The cyclic 3’, 5’-adenosine monophosphate receptor protein and regulation of cyclic 3’, 5’-adenosine monophosphate synthesis in Escherichiu coli. Mot. Gen. Genet. 165, 47-56. Buchmeier, N., Bossie, S., Chen, C.-Y., Fang, F.C., Guiney, D.G. and Libby, S.J. (1997) SlyA, a transcriptional regulator of Salmonella typhimurium, is required for resistance to oxidative stress and is expressed in the intracellular environment of macrophages. Infect. Immun. 65, 3125 -3130. Cabello, F., Hormaeche, C., Mastroeni. P. and Bonina, L. (eds) (1993) Biology o / Sirlmoni4a, Plenum, New York. Cashel, M., Gentry, D.R., Hernandez, V.J. and Vinella, D. (1996) The stringent response. In: Escherichia coli and Salmonella Cellulur and Molecular Biology (F.C. Neidhardt, R. Curtiss, I l l , J.L. Ingraham, E.C.C. Lin, K.B. Low, B. Magasanik, W.S. Reznikoff, M . Riley, M. Schaechter and H.E. Umbarger, eds), pp. 1458- 1496. ASM Press, Washington, DC. Chalker, R.B. and Blaser, M.J. (1988) A review of human salmonellosis. 111. Magnitude of Sa~monelluinfection in the United States. Rev. Infect. Dis. 10, I I 1 ~124. Clark, D.P. and Cronan, J.E., Jr. (1996) Two-carbon compounds and fatty acids as carbon sources. In: Escherichia coli and Salmonella Cellulur and Molecular Biology (F.C. Neidhardt, R. Curtiss, 111, J.L. Ingraham. E.C.C. Lin, K.B. Low, B. Magasanik, W.S. Reznikoff, M. Riley, M . Schaechter and H.E. Umharger. eds), pp. 343- 357. ASM Press, Washington, DC.

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Iron Storage in Bacteria Simon C. Andrews School of Animal and Microbial Sciences, The University of Reading, Whiteknights, PO Box 228, Reading RG6 6 A H , U K

ABSTRACT

Iron is an essential nutrient for nearly all organisms but presents problems of toxicity, poor solubility and low availability. These problems are alleviated through the use of iron-storage proteins. Bacteria possess two types of iron-storage protein, the haemcontaining bacterioferritins and the haem-free ferritins. These proteins are widespread in bacteria, with at least 39 examples known so far in eubacteria and archaebacteria. The bacterioferritins and ferritins are distantly related but retain similar structural and functional properties. Both are composed of 24 identical or similar subunits (-19 kDa) that form a roughly spherical protein (-450 kDa, -120 diameter) containing a large hollow centre (-80 diameter). The hollow centre acts as an iron-storage cavity with the capacity to accommodate at least 2000 iron atoms in the form of a ferric-hydroxyphosphate core. Each subunit contains a four-helix bundle which carries the active site or ferroxidase centre of the protein. The ferroxidase centres endow ferrous-iron-oxidizing activity and are able to form a di-iron species that is an intermediate in the iron uptake, oxidation and core formation process. Bacterioferritins contain up to 12 protoporphyrin IX haem groups located at the two-fold interfaces between pairs of two-fold related subunits. The role of the haem is unknown, although it may be involved in mediating iron-core reduction and iron release. Some bacterioferritins are composed of two subunit types, one conferring haem-binding ability (a) and the other (p) bestowing ferroxidase activity. Bacterioferritin genes are often adjacent to genes encoding a small [2Fe-2S]-ferredoxin (bacterioferritin-associated ferredoxin or Bfd). Bfd may directly interact with bacterioferritin and could be Copyright @I 1998 Academic Press ADVANCES IN MICROBIAL PHYSIOLOGY VOL 40

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involved in releasing iron from (or delivering iron to) bacterioferritin or other iron complexes . Some bacteria contain two bacterioferritin subunits. or two ferritin subunits. that in most cases co.assemble . Others possess both a bacterioferritin and a ferritin. while some appear to lack any type of iron-storage protein . The reason for these differences is not understood . Studies on ferritin mutants have shown that ferritin enhances growth during iron starvation and is also involved in iron accumulation in the stationary phase of growth . The ferritin of Cumpylobacter jejuni is involved in redox stress resistance. although this does not appear to be the case for Escherichiu coli ferritin (FtnA). No phenotype has been determined for E . coli bacterioferritin mutants and the precise role of bacterioferritin in E . coli remains uncertain . 283 1. Biologically relevant features of iron . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 283 1.1. Beneficial properties of iron . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 283 1.2. Detrimental properties of iron . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.3. Countering the problems of iron dependence . . . . . . . . . . . . . . . . . . . 2 . Ferritins, rubrerythrins and bacterioferritins . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Eukaryotic ferritins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 288 291 2.2. Rubrerythrins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 293 2.3. Bacterioferritins . . . . . . . . . . . 302 2.4. Bacterial ferritins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 . Primary structures and evolution of iron-storage proteins . . . . . . . . . . . . . . . . 305 3.1. The ferritin-bacterioferritin-rubrerythrin superfamily . . . . . . . . . . . . . . . . . 305 3.2. Bacterioferritin heteropolymers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 311 3.3. Evolution of the ferritin-bacterioferritin-rubrerythrin family from a simple two-helix protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 313 3.4. The ubiquity of bacterioferritins and ferritins ....................... 314 3.5. Multiple iron-storage proteins within bacteria . . . . . . . . . . . . . . . . . . . 3.6. A relationship between the ferritin-bacterioferritin-rubrerythrin superfamily and the Dps family? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 315 316 4 . Structures of bacterioferritin and bacterial ferritin . . . . . . . . . . . . . . . . . . . . . . . 4.1. E . coli bacterioferritin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 316 4.2. E . coli ferritin (FtnA). . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 320 321 4.3. Comparison of ferroxidase centres . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 . Core formation and the iron core . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .323 323 5.1. Iron uptake . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ........................ 326 5.2. The iron core .................... 329 6 Bacterioferritin-associated ferredoxin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 333 7 . lntracellular iron metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 333 7.1. Regulation and physiological roles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 337 7.2. The low-molecular weight iron pool . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 339 7.3. lntracellular ferric reductases. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8 . Perspective . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 339 Note added in proof . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 340 Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 340 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 341

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1. BIOLOGICALLY RELEVANT FEATURES OF IRON 1.1. Beneficial Properties of Iron

Iron is a major trace element for all bacteria, other than the lactobacilli (Archibald, 1983). It normally represents about 0.02% of bacterial dry weight but can be as high as 1.8% (Sphaerotilus natans; Rouf, 1964). The utility of iron arises from its abundance in the Earth’s crust (it is the fourth most common element; Zajic, 1969) and its useful chemical properties. Iron possesses two stable oxidation states (11) and (111) (ferrous and ferric) that are readily interchangable, allowing iron to participate in redox reactions. The redox potential of the Fe(lI):Fe(III) couple is strongly dependent on the coordinating ligands and can range from +330 (Chromatium HiPiP) to -490 mV (Chromatium ferredoxin). This flexibility makes iron an extremely useful redox mediator in biology. Furthermore, iron is able to undergo rearrangments of its d orbital electrons into different spin states. This enables it to mediate reactions involving molecular oxygen. Thus, during oxygenase and oxidase reactions, changes in the spin state of iron are used to compensate for otherwise energetically unfavourable spin alterations in oxygen (Wrigglesworth and Baum, 1980). Iron can be specifically incorporated into proteins in several different forms: haem (A, B, C, D, E, sirohaem); iron-sulphur ([Cys4-Fe], [2Fe-2S], [3Fe-4S], [4Fe-4S], [8Fe-7/8S], [7Fe-9S-Mo-homocitrate]); iron-nickel (Ni-Fe-CO[CNI2); di-iron (Fe-OFe); and mononuclear iron. Processes in which iron-containing proteins participate include: respiration (cytochromes, ferredoxins and other ironsulphur proteins); activation of oxygen (cytochrome oxidase); degradation of H202 and 0; (haem-catalase, iron-superoxide dismutase, peroxidase); amino acid and pyrimidine biosynthesis (glutamate synthase, dihydro-orotate dehydrogenase); the citric acid cycle (fumarase, aconitase, succinate dehydrogenase); DNA synthesis (ribonucleotide reductase); nitrogen fixation (nitrogenase); carbon fixation metabolism (methane monooxygenase); photosynthesis (light-harvesting complexes, reaction centre, ferredoxin); oxygen binding (globins); degradation of aromatic compounds (toluate 1,2-dioxygenase); and gene regulation (FNR, SoxR, CooA). It appears that there are few aspects of metabolism that iron does not touch and its first association with life on Earth is likely to have been an early event in evolution (Hartman, 1975; Williams, 1990). 1.2. Detrimental Properties of Iron

Although iron is extremely beneficial to cellular organisms, it also possesses properties that are problematic. The solubilities of Fe(I1) and Fe(II1)

284

SIMON C. ANDREWS

at pH 7 are -10-1 and 1 0 - l 8 ~ ,respectively, indicating that Fe(II1) is unstable in solution under physiological conditions (Biedermann and Schindler, 1957). Fe(II1) tends to hydrolyse and polymerize in aqueous environments, lowering the effective solubility of iron ( 1 0 - 3 6 ~still ) further. Since the oxidized, Fe(III), form of iron predominates under aerobic conditions, poor iron availability is often a major difficulty for iron-dependent organisms. Indeed, lack of iron is one of the most common forms of malnutrition and is particularly acute in certain environments such as some oceans (e.g. the Southern Ocean and tropical Pacific) where growth of phytoplankton is strictly limited by iron availablity. A further problem posed by iron is its tendency to potentiate oxygen toxicity by interacting with oxygen and oxygen-reduction products to generate highly reactive and toxic free radicals (Halliwell and Gutteridge, 1984). Reactive oxygen species such as 0; and H 2 0 2 are the by-products of normal aerobic metabolism and can be generated by environmental agents. The damage inflicted during redox stress is to a large extent dependent on the collaboration of intracellular iron (Repine et al., 1981; Flitter et al., 1983; Floyd, 1983; Mello Filho and Meneghini, 1984, Mello Filho ef al., 1984; Sutton and Winterbourn, 1984; Zigler et al., 1985; Byler et al., 1994). 0, and H 2 0 2 are only mildly reactive in aqueous solutions but, in the presence of iron, highly damaging and reactive hydroxyl radicals (HO') can be formed via the Fenton reaction (equation 2) (Halliwell, 1978). 0; is involved in redox cycling of Fe(II1) to Fe(I1) (equation 1). Combination of equations 1 and 2 gives the Haber-Weiss reaction (equation 3) in which iron catalyses the production of the highly reactive OH' (hydroxyl) radical from H202 and 0; 0;

+ Fe3+ -Fe2+

+0 2 + OH- + HO'

Fe2+ + H 2 0 2-Fe3+ Net : 0,

+H202

(1)

Fe calalyst

HO'

+ OH- + O2

(2) (3)

It is the OH' radical that is the major culprit in inflicting damage during redox stress.

1.3. Countering the Problems of Iron Dependence

Iron therefore presents a dual paradox for aerobic iron-requiring organisms. It is at once useful and hazardous, and is abundant but poorly available.

IRON STORAGE IN BACTERIA

285

Bacteria (and other cellular organisms) utilize a variety of mechanisms to counter the problems presented by their dependence on iron. Iron toxicity is alleviated through anti-oxidants (e.g. glutathione) and enzymes (e.g. superoxide dismutases, catalases, peroxidases) that degrade reactive oxygen species and by repair systems (e.g. endonucleases) that repair the damage inflicted during redox stress. The problem of low Fe(II1) solubility is overcome by highly efficient iron-transport systems employing high-affinity extracellular Fe( 111) chelators (called siderophores) (Briat, 1992; Cuerinot, 1994). Bacteria are able to use siderophores synthesized and secreted by themselves or by other organisms. Ferri-siderophore complexes are internalized by specific membrane-associated transport systems. There are often multiple siderophore-mediated iron-transport systems in any given organism (up to eight in Escherichia coli; Earhart, 1996), and so far over 200 siderophores have been identified from microbial sources (Neilands. 1981). The expression of these systems is generally tightly regulated by iron availability. Bacteria can also transport Fe(I1) directly and, like fungi and plants, some bacteria can reduce Fe(II1) prior to transport as Fe(I1) (Guerinot, 1994). Invasive pathogenic bacteria circumvent the withdrawal of free iron by their hosts through the acquisition of iron directly from host sources, such as by the binding and uptake of iron from host iron-binding proteins (Fe-lactoferrin, Fe-transferrin, haem-haemopexin, haemoglobin-haptoglobin, haemoglobin, haem) and by the haemolysin-mediated lysis of erythrocytes followed by the use of released haemoglobin as an iron source, as well as by siderophore-mediated transport (Litwin and Calderwood, 1993). Bacteria can also adapt to iron deficiency by reducing their requirement for iron (e.g. by expressing flavodoxin in place of ferredoxin; discussed in Bovy et al., 1993). Another mechanism for dealing with the problems posed by iron involves the use of iron-storage proteins or ‘ferritins’. Ferritins are found in eubacteria, archaebacteria and eukaryotes (Table 1). The universality of ferritins implies a fundamental role in iron metabolism. The physiological purpose of ferritins is thought to be to sequester iron in a soluble and readily available form, providing reserves of iron that can be drawn upon when required. Ferritins are also thought to maintain iron in a form that is less likely to participate in harmful, free-radical-generating reactions (Theil, 1987; Andrews et al., 1992). Therefore, ferritins assist in dealing with both of the problems that iron imposes, namely poor solubility/bioavailability and toxicity. Since iron-storage proteins are viewed as the major mediators of the iron-storage process, the focus of this review will be the structurefunction, physiological role and regulation of synthesis of the ferritins and bacterioferritins of bacteria.

Table 1 Ferritins. bacterioferritins. rubrerythnns and bacterioferritin-associated ferredoxins in bacteria (and one organelle and one fungus).

Species (Absidia spinosa, a fungus) Alcalagenes eurrophus Archaeoglobus fulgidus Azotobacter chroococcwn Azotobacter vinelandii Bacillus subtillis Bacteroides fragilis Brucella melitensis Campylobacter jejuni Clostridium acetobutylicum Clostridium perfringens (Cyanophora paradoxa cyanelle) Deinococcus radiodurans Desulphovibrio vulgaris Escherichia coli Helicobacter pylori Haemophilus influenzae Magnetobacterium magnerotacticum Methanobacterium thermoautotrophicum Methanococcus jannaschii Mycobacterium avium Mycobacterium leprae Mycobacterium paratuberculosis M~~cobacterium tuberculosis Mycoplasma genitaliuni

Genome size (MB)

2.20

c

Ftn

Bfr

1

Rer

Bfd

Reference

4

0

a a b

0

1 0

C

4.2 c

0 1 1 1

4.1 P

1

1

1

C 3.00 P

0 0

0 0

0 0

4.60 C 1.66 C 1.83 C

2 1 2

1 0

1.75 P 1.66 C 4.70 P 4.40 P

1 0

4.40 P 0.58 C

1 0

1 0 0 2 0 0 I 1 1 1 0

0 2 1

? 0

1

0 0

1 0

a h a a a d a a b a a b b a d b a a a b b

Table 1 continued

Genome size Species

(MB)

Ftn

Bfr

Rer

Mycoplasma pneumoniae Neisseria gonorrhoeae Neisseria meningitidis Niirobacier winogradskii Pseudomonas aeruginosa Pseudomonas put ida Rhodobacter capsulatus Rhodopseudomonas sphaeroides Rhodospirillurn rubrum Streptococcus pyogenes Synechocystis PCC6803 Thermatoga maritima Treponema pallidum Vibrio cholerae

0.81 C 2.20 P 2.30 P

0 0 0

0 0

0

e

0

0

0

f b a

5.9 P

0

0 2 2 1 2

0

1

g

Totals (bacteria only): 37

1.98 P 3.57 c 1.80 P 1.05 P 2.50 P

Bfd

Reference

1 1

a a

1 1

C

0

C

0 0 1 0 1

2 0 0 1

0 0 1 0 0

13

26

12

f a b b b

Numbers of ferritin (Ftn), bacterioferritin (Bfr), rubrerythrin (Rer) and Bfd proteins in each species are given, where known. Genome size is given for genomes fully (C) or partly (P) sequenced. Proteins whose sequences are absent from Figs 6 and 10, but are present in this table, are either not available as amino acid sequences or are unavailable as single amino acid sequences. References for the data above are as follows: (c) seeTable 2; (d) http://www.cric.com/htdocs/sequences; (a) see Figs 6 and 10; (b) http://www.ncbi.nlm.nih.gov/cgi-bin:BLAST/nph-tigrbl; (e) Himmelreich er al. (1996); (f) http://dna 1 .chem.uoknor.edu; (g), http://www.ncbi.nlm.nih.gov/cgi-bin~BLAST/nph-pseudoabl; and (h), http://www.pasteur.fr/Bio/SubtiList.html.

288

SIMON C. ANDREWS

2. FERRITINS, RUBRERYTHRINS AND BACTERIOFERRITINS

2.1. Eukaryotic Ferritins The structure, function and regulation of expression of eukaryotic ferritins have been extensively characterized, particularly those in mammals, e.g. human, rat, mouse and horse (Ford et al., 1984; Theil, 1987; Harrison and Arosio, 1996). The X-ray structures of several eukaryotic ferritins are known (Banyard et al., 1978; Lawson et al., 1991; Trikha et a/., 1994, 1995; Gallois rt al., 1997; Hempstead et al., 1997) revealing that ferritins have a unique molecular architecture. They are composed of 24 structurally identical subunits (approx. 19 kDa) that assemble to form a roughly spherical protein shell (approx. 500 kDa, 120 A diameter) surrounding a central storage cavity (approx. 70 A diameter) with the capacity to accommodate up to 4500 iron atoms as a ferric-oxy-hydroxyphosphate core (Figs 1 and 2). The surrounding protein coat maintains the otherwise insoluble iron core in a soluble state. Each subunit is composed of a four ct-helix bundle (helices

1

I

Figure I Schematic representation of the structure of horse-spleen apoferritin. ( 1) The 4-3-2 fold symmetrical structure of the 24 equivalent subunits of an apoferritin holomer, centred at the four-fold axis. Each subunit is represented by a capsule-shaped unit and the orientation of each subunit is indicated : N, N-terminal end; E, E-helix end. (2) Ribbon representation of the alpha-carbon backbone of a single subunit. The five helices (A-E) and the long loop (L) are indicated. as is the N-terminus. Reprinted with permission from Ford et al. (1984).

IRON STORAGE IN BACTERIA

289

Figure 2 A schematic representation of iron core formation in ferritin. (a) Ferrous iron passes through channels in the protein shell and is oxidized at ferroxidase centres. (b) An iron-core nucleation centre is formed. (c,d) The ferrihydrite-microcrystal grows as ferrous iron is oxidized on the surface of the growing core. The rate of core formation first increases as the available surface area for iron deposition on the growing iron core (indicated by the thickened line) increases, and then the rate declines as the molecule fills and the exposed surface of the core is reduced. Arrows indicate entry of ferrous iron. Reprinted with permission from Harrison ef al. (1980).

A-D) with helices B and C connected by a long loop (L) that traverses the length of the bundle, and a short helix (E) at the C-terminal end (see Fig. I). Most ferritin subunits possess an active site at the centre of their fourhelix bundles, known as the ‘ferroxidase centre’, which is responsible for the oxidation of ferrous to ferric iron that occurs during the uptake and storage reaction catalysed by ferritins (Fig. 3). This ferroxidase activity endows ferritins with enzymatic properties, although ferritins do not ‘turn-over’ in the way that enzymes do. Once the storage cavity is full, the iron uptake reaction cannot continue. Furthermore, it appears that only the first few ferrous ions (-50) are oxidized by the protein and that subsequent oxidation occurs on the surface of the growing iron core (Fig. 2). The function of the ferroxidase centre may be simply to oxidize the first few iron atoms to provide a nucleus for core growth which, once initiated, grows autocataly-

290

SIMON C. ANDREWS

Human H-chainferritin

~ h14 1

Glu-27

Horse '2-chain"ferritin

~ l , , - 1 41

Tyr-27

Figurc 3 Schematic diagrams of the dinuclear metal site of mammalian ferritin Hsubunits and the equivalent site in mammalian ferritin L-subunits. The metal sites I and 2 in human H-chain ferritin are indicated. The equivalent region of the horse ferritin Lsubunit lacks the dinuclear metal species, and instead there is a salt bridge involving Lys62 (residue numbering is as for human H-chain ferritin). Adapted from Harrison and Arosio ( 1996).

tically. An intermediate p-0x0-bridged di-iron species is formed at the ferroxidase centre during the oxidation reaction and the amino acid residues acting as ligands to the di-iron species are highly conserved among ferritins and are crucial for full ferroxidase activity (see later, Fig. 6 for conserved ferroxidase centre residues). The oxidant used physiologically by ferritins is thought to be molecular oxygen, which is converted to hydrogen peroxide in the initial protein-catalysed step, or water in the subsequent iron-corecatalysed step. Kinetic, spectroscopic and site-directed engineering studies

IRON STORAGE IN BACTERIA

29 1

indicate that the reaction pathway involves formation of a di-ferric species at the ferroxidase centre, followed by decomposition of the dimer to a ferric monomer and then build-up of iron clusters or cores. X-ray and electron diffraction experiments have shown that the iron core is normally ordered or crystalline, and is like the mineral ferrihydrite in its structure (Towe and Bradley, 1967; Fischbach et al., 1971). The structure of the core is clearly dictated by the protein shell and the environmental conditions under which the core is formed. The physiological mechanism of iron release from eukaryotic ferritins is unknown, but could involve lysosomal degradation. Ferritins often possess more than one type of subunit (e.g. H and L subunits in mammalian ferritins, 55% identical) and these have similar sequences, are structurally similar and are believed to be mutually interchangeable in the protein coat allowing the formation of a series of ‘heteropolymers’ or ‘isoferritins’ with subunit compositions ranging from HOL24to H24Lo. For such heteropolymeric ferritins, one type of subunit may lack a functional ferroxidase centre (e.g. the L subunit of mammalian ferritins; Fig. 3). L-subunit-rich ferritins are found in iron-rich tissues and have a lower ferroxidase activity than H-rich ferritins. They are more stable and better at forming iron cores. It is probable that the purpose of L-rich ferritins is to provide a ‘long-term’ iron store whereas H-rich ferritins are likely to be more important in short-term iron storage or intracellular ‘iron-flux’. Ferritins tend to be extremely resilient to physical and chemical stresses (e.g. 80°C or 8~ urea for horse-spleen ferritin), a feature often exploited during their purification. The durability of ferritins is likely to reflect a requirement for a stable iron-storage system that will not release sequestered iron in an uncontrolled manner. Appropriately, apoferritin synthesis is induced by iron.

2.2. Rubrerythrins

It has recently been shown that ferritins are structurally related to rubrerythrins (deMar6 et al., 1996), implying an evolutionary relationship which is supported by amino acid sequence comparisons (Harrison et a]., 1998). The function of rubrerythrins is unknown, although it has been suggested that they act as superoxide dismutases (Lehmann et al., 1996), and it has been shown that they have ferroxidase activity similar in magnitude to that of ferritins (Bonomi et al., 1996). These findings suggest roles in redox-stress resistance or iron metabolism. Rubrerythrins have so far been found only in ‘anaerobic’ bacteria (Archaeoglobus fulgidus, Clostridium acetobutylicum, Clostridium perjiringens, Desulphovibrio vulgaris, Methanococcus jannaschii, Methanobacterium thermoautotrophicum and Therrnotoga maritima; see Table 2 ) . Rubrerythrins are homodimers of 22 kDa subunits. Each subunit

Table 2 Comparison of the properties of bacterioferritins from different species

Size ( m a ) Species

Subunit

24-mer

E. coli P . aeruginosa A . vinelandii N . winogradrkyi R . sphaeroides R. rubrum A . chroococcum Synechocystis A . spinoso R. capsulatus

18.5 18 and 18.5 18 19.5 16 23 17 19 20 and 20 18.2

452 430 443 260 > 100 450 -410 400

-480 437

Haems/ holomer

Fe atoms/ holomer

ct peak

12 3-9 12 12 10 Yes Yes 6 Yes 6

-980 7W840, 60&2400

557 557 557.5 559 558 557.5 557.5 559 557 557

-100

2300 750 600-950

(nm)

Haem Em (mW

Fe:phosphate (mo1e:mole)

Subunits (no of types)

-340t, -20*

2.2:l 1.7:1 1.4:1

1 2

-474, -2251

>o

-204 1.5:1 1.61.9:l

1 probably 1

1 1 1 1 (or 2) 2 1

Data from 'Fujita et al. (1963), + D e b and Hager (1964), Yariv et al. (1981), Andrews et 02. (1989a,b), Moore et al. (1986; 1994), Stiefel and Watt (1979), Mann et al. (1987), Watt et 01. (1986), Kurokawa et 01. (1989), Meyer and Cusanovich (1985), Bartsch et 01. (1971), Chen and Crichton (1982), Laulhere er al. (1992), Carrano et al. (1996), Penfold et of. (1996) and Ringeling et 01. (1994). Value determined for apobacterioferritin (nonhaem-iron free). Value determined for iron-core-containing bacterioferritin.

*

IRON STORAGE IN BACTERIA

293

possesses two domains: an N-terminal four-helix bundle, similar to those of ferritins, containing a ferritin-like di-iron cluster; and a C-terminal rubredoxin-like domain containing an Fe-Cys4 cluster (in place of the E-helix of ferritins).

2.3. Bacterioferritins 2.3.1. Azotobacter vinelandii Ferritin was first discovered in mammals -y Schmiedeberg in 1894 and called ‘ferratin’ (Harrison et a / . , 1974). The protein was first purified by Laufberger (1937) from horse spleen, later discovered in plants by Hyde et al. (1963) (and initially termed ‘phytoferritin’), and then in fungi by David and Easterbrook (1971). However, the first report of an iron-storage protein from a prokaryote was not until 1979 when Stiefel and Watt described the isolation of a ‘bacterioferritin’ from the nitrogen-fixing bacterium, Azotobacter vinelandii (Stiefel and Watt, 1979). Moreover, as recognized by Stiefel and Watt (1979), this protein had been isolated earlier by Bulen et a/. (1973) and described as a b-type cytochrome containing large amounts of non-haem iron and no labile sulphide. The A . vinelandii bacterioferritin was, like eukaryotic ferritins, heat stable (at 60°C) and contained a subunit of 17 kDa, 1100-1800 iron atoms and, unlike ferritins, contained 12 haems per molecule (assuming a 24-mer). A later SDS-PAGE analysis by Harker and Wullstein (1985) suggested the presence of two types of subunit (21 and 23 kDa), but this finding has not been substantiated. The protein was defined as a ‘bacterioferritin’ by Stiefel and Watt (1979) largely on the basis of its appearance under the electron microscope as an electron-dense core of about 55 diameter (the iron core) surrounded by a roughly spherical electron-transparent shell of about 105 A in outer diameter (the protein coat). This morphology, together with the dimensions, are characteristic of ferritin molecules. Mossbauer and magnetic susceptibility measurements on the iron core revealed superparamagnetic behaviour (as for horse ferritin) and a magnetic moment of 3.7 Bohr magnetons per Fe atom (similar to that for horse ferritin). The haem group was shown to be protoporphyrin IX (Bulen rt al., 1973) and the oxidized and reduced spectra had a Soret band at 417 nm and a,p and Soret bands at 557.5, 527 and 425 nm, respectively (see later, Fig. 5). The haem was shown to be intimately associated with the protein and redox measurements gave a reversible Nernst plot (n = 1) with an extemely low mid-point potential of -416 mV at pH 7 (later revised, see Section 2.3.4). In reduction experiments, one electron was taken up for each iron atom in the protein, resulting in the reduction of both haem and non-haem iron. The reduction potential of the haem was shown to be about 30 mV more nega-

A

294

SIMON C. ANDREWS

tive than the iron core (although this finding was later modified; Watt et ul., 1986), and therefore the reduction potental of the iron core is much lower than that of horse-spleen ferritin (which is -190 mV at pH 7; Watt et d., 1985). Apart from the presence of haem and the low redox potential of the iron core, the other difference between bacterioferritin and horse-spleen ferritin was thought to be the greater ability of bacterioferritin to retain iron once reduced. However, a similar capacity to retain reduced iron core was later established for horse-spleen ferritin (Watt e l a/., 1985). Stiefel and Watt (1979) speculated that bacterioferritin serves as an ironstorage protein, in the same way as eukaryotic ferritins do, and that the associated haem acts to facilitate reduction of Fe(II1) to the soluble and more labile Fe(I1) during iron release. They suggested that the low redox potential for Fe(lI1) reduction may ensure that iron is only released when the redox potential of the cell is low, as required for nitrogenase activity (- -430 mV). Another suggested role was one of electron storage, given the huge capacity of the protein to act as an electron sink at low redox potentials. The A . vinrlandii bacterioferritin was also partially characterized by another group (Jiudi r t ul., 1980, 1985; Baoguang et ul., 1984) whose results were consistent with those of Stiefel and Watt (1979). 2.3.2. Escherichia coli At approximately the same time as the A . vinelundii studies, others were investigating the properties of an iron-storage protein from E. coli (Bauminger et ul., 1976, 1979, 1980). Initial whole-cell Mossbauer studies at low temperatures showed that E. coli contains high-spin Fe(II1) aggregates that become magnetically ordered at very low temperature ( 3 K ) (Bauminger et u/., 1976, 1979), suggesting the presence of iron-cores similar to those of ferritin. Subsequent Mossbauer studies on 57Fe-labelled E. coli and bacterioferritin purified from the same source revealed that most of the iron in ‘iron-rich’ E. coli is in aggregates similar to those in isolated bacterioferritin, although only 1 YOof cellular iron could be attributed to bacterioferritin itself (Bauminger et ul., 1980). However, this estimation is likely to be erroneous since, as E. coli was grown aerobically with 36 V M 57Fe in the absence of any chelator, it is likely that much of the 57Fe preciptated and was harvested with the bacteria giving an excessively high apparent content of cellular iron. Furthermore, the amount of iron measured in the harvested bacteria was equivalent to 0.27% of dry weight, which is 10-fold that normally found. Similar unexpectedly high bacterial iron contents arising from iron precipitation have been reported previously (Hartmann and Braun, 1981). The real proportion of cellular iron associated with bacterioferritin during the logarithmic phase is therefore likely to be approximately lo%, rather than 1%.

295

IRON STORAGE IN BACTERIA

Two forms of bacterioferritin were isolated from aerobically grown E. coli by Yariv et al. (1981), one containing a polynuclear iron core (holobacterioferritin) and another form (apo-bacterioferritin) lacking an iron core. The proteins contained 12 haems (protoporphyrin IX) per 24 subunits (J. Yariv unpublished work, in Smith et al., 1988b) and had subunits of 15 kDa as estimated by SDS-PAGE. Electron microscopy revealed that the molecule is a sphere of 95 diameter (later measured as -124 A; Smith et al., 1989) with an inner electron-dense core of 60 diameter (Fig. 4). The holo form was purified using high-speed centrifugation which exploited its high density, and by heating (60°C, 10 min) and CsCl crystallization. The apo form was purified by immunoprecipitation (using antibodies raised against the holo form) followed by gel filtration in 8 M acid-urea. A mixture of apo and holobacterioferritin was obtained following growth with 10 mg/l FeS04 .7H20,but only the apo-form was present following growth on 5 mg/l FeS04.7H20 and casamino acids. This indicated that the iron content of bacterioferritin is subject to iron availability. The CsC1-bacterioferritin crystals had a 1432 space group, suggesting 24 identical subunits and a quaternary structure resembling those of eukaryotic ferritins. It later became apparent (Yariv, 1983; Smith et al., 1988b) that, as for the A . vinelandii bacterioferritin, the E. coli bacterioferritin had also been described and isolated previously, and identified as cytochome bl (Keilin, 1934; Fujita et al., 1963; Deeb and Hager, 1964; Hager and Deeb, 1967; Hagihara et al., 1975). The cytochrome b, had been shown to possess a haem group with a redox potential of -340 mV or -20 mV and to have a molecular mass of 500 kDa or 700 kDa (Fujita et al., 1963; Deeb and Hager, 1964). The first bacterioferritin gene (bfr) to be cloned was from E. coli (Andrews et al., 1989a,b). The gene was isolated by screening a lambda library using an anti-bacterioferritin immunostaining procedure. The cloned segment of DNA contained two open-reading frames, one (gen64 or /~fi/,) encoding a putative 64-residue protein (P64 or the bacterioferritin-associated ferredoxin, Bfd) and another (bfr) encoding a 1%-residue polypeptide (18.5 kDa) with an N-terminal sequence closely matching that of purified bacterioferritin. The two genes are located at 74.6 min, have the same polarity, are separated by 71 bp and each have well-predicted promoters and ribosome-binding sites (Andrews et al., 1989a; Berlyn ef d., 1996). The gene- and protein-derived amino acid sequences of bacterioferritin reported by Andrews et al. (1989a,b) differed markedly from the protein-derived sequence reported previously by Tsugita and Yariv (1989, indicating that much of the earlier analysis was incorrect. Furthermore, reports of an E. coli gene at 53 min encoding a protein corresponding to bacterioferritin (according to two-dimensional PAGE and peptide mapping analyses; Neidhardt et al., 1983) proved to be mislead-

A

A

296

SIMON C. ANDREWS

Figure 4 Electron micrographs of E. coli bacterioferritin. (a), (c) and (d) are negatively stained with uranyl acetate, (b) is unstained. Magnification is x 132 000 (a) or x255 000 (bd).Arrows indicate electron-dense cores (b), ordered arrays (c), and noncontiguous iron cores. Reprinted with permission from Yariv e / al. (1981).

IRON STORAGE IN BACTERIA

297

ing (Andrews et al., 1991~).The initial sequence analysis indicated that the gene-derived bacterioferritin amino acid sequence was unlike any other in the databases, including ferritin sequences, although secondary structure prediction suggested a high a-helical content consistent with a four-helix bundle conformation, as found in the ferritins (Andrews et al., 1989b). However, further analyses later revealed a convincing sequence similarity between bacterioferritins and ferritins indicative of shared structural and functional properties (see Section 3.1). The E. coli bacterioferritin and several variants have been overproduced in E. coli, purified and characterized (Andrews et al., 1993, 1995; Le Brun et al., 1995). Two assembly forms of overproduced bacterioferritin were identified, the usual 24-subunit form and a novel haem-containing subunit-dimer form. The proportions of these two assembly forms varied in different preparations from 0 to 65% dimer. The reason for this variability was not discovered (Andrews et al., 1993). During native-PAGE the 24-mer partially disassembled into two assembly forms, a subunit dimer and a subunit monomer. These findings indicate that bacterioferritin may be prone to disassembly under some conditions. Whether this has any physiological significance is unclear, as is the possibility that the subunit dimer has a physiological role which differs from that of the 24-mer. Disassembly of the bacterioferritins of A . vinelandii, Pseudomonas aeruginosa and E. coli at low pH during isoelectric focusing has also been reported (Andrews et al., 1991a). The overproduced bacterioferritin contained 3.5-10.5 haems/24-mer and 25-75 nonhaem iron atoms/24-mer (Andrews et al., 1993). Despite its low initial iron content, the protein was able to accumulate up to 1800 iron atoms in vitro by the addition of ferrous iron to aerobic solutions (pH 6.5-7.0) containing apobacterioferritin. Thus, E. coli bacterioferritin is able to catalyse the oxidation of iron (and so has ‘ferroxidase activity’) and the formation of an iron core, although the rate of iron uptake was 10-fold slower than that of human H-chain ferritin. The iron core had no effect on the electrophoretic or crystallographic properties of the protein, demonstrating that the core was within the hollow central storage cavity. The low iron content of overproduced bacterioferritin is probably due to the increased storage capacity of the overproducing strain. The low haem content may reflect an inability of the overproducing strain to synthesize sufficient haem to meet the extra demand placed upon it by the overproduction of bacterioferritin. 2.3.3. Other bacteria In addition to A . vinelandii and E. coli, bacterioferritins have now been purified from Azotobacter chroococcum (Chen and Crichton, 1982), Nitrobacter winograd.skyi (Chaudhry et al., 1980; Kurokawa et al., 1989),

298

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P . aeruginosa (Moore et al.. 1986), Synechocystis PCC 6803 (Laulhtre et al., 1992), Rhodopseudomonas sphaeroides (Meyer and Cusanovich, 1989, Rhodobacter rubrum (Bartsch et al., 1971; Okada and Okunuki, 1973), Rhodobacter capsulatus (Ringeling et al., 1994) and Mycobacterium paratuherculosis (Brooks et al., 1991) (Table 1). In pathogenic bacteria, bacterioferritins have been recognized following their initial identification as antigens (Brooks el a/., 1991; Denoel et al., 1995). The properties of bacterioferritins characterized so far are very similar although there is variability in haem content (3-12 haems per 24 subunits), subunit composition (either one or two types) and iron content (Table 2). Bacterioferritins appear to be widespread in bacteria (Table 1) having been found (either as the protein or the encoding gene) in the high- and low-G + C Gram-positive bacteria, cyanobacteria and in protobacteria, and possibly fungi (Carrano rt al., 1996). 2.3.4.

The huem group

All bacterioferritins characterized so far contain the protoporphyrin IX haem group and therefore haem appears to have an important role in the function of the protein. A typical bacterioferritin spectrum is shown in Fig. 5 and the spectral properties of the haem groups of several bacterioferritins are summarized in Table 3. The presence of haem distinguishes the bacterioferritins from the ferritins, although it remains unclear why ferritins do not contain haem whereas bacterioferritins do. Clearly, an understanding of the precise function of the haem group in bacterioferritin is crucial if a proper appreciation of the role of bacterioferritin in iron metabolism is to be obtained. Perhaps the most likely role of the haem is in release of iron from the iron core by mediating electron transfer through the protein shell. There is some evidence in support of this notion (see below) although little is known regarding either the source of electrons used for haem reduction or the mechanism governing control of the release and uptake reactions. The axial haem ligands of bacterioferritins from A . vinelandii, E. coli and P . aeruginosa were ini tally identified by near infrared (NIR) magnetic circular dichroism (MCD) and electron paramagnetic resonance (EPR) as the thioether side chains of a pair of methionine residues (Cheesman et al., 1990, 1992, 1993; McKnight et al., 1991). A haem ligation scheme of this nature is not known to exist in any other type of haemoprotein apart from cytochrome hShz variants (Barker and Freund, 1996). An extended X-ray absorption fine structure (EXAFS) analysis on A . vinelandii bacterioferritin supported the NIR-MCD and EPR studies by showing that the haem iron group has two axial sulphur ligands at a distance of 2.35 and four nitrogen ligands at 1.97 A (George et a/., 1993). The identity of the

A,

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Figure 5 Ultraviolet absorption spectra of overproduced E. coli bacterioferritin (BFR) and a BFR variant (BFR-1) with a 14-residue C-terminal extension. BFR-h was fully assembled but contained only one to two haems per holomer and could not take up iron. The C-terminal extensions project into the central cavity to occupy the space normally used for iron storage, and they appear to perturb the haem-binding pocket (hence the low haem content and altered spectrum in the 725-737 nrn region). (A) BFR (400 pg/ ml), pH 7. The oxidized spectrum was obtained by using an aerobic solution and the reduced spectrum was obtained by addition of a few grains of Na2S204 to the aerobic solution. (B) Comparison between the oxidized spectra of BFR-1 (10 mg/ml) and BFR (1.5 mg/ml) at 62CMOO nm, revealing a slight perturbation in the BFR-h haem-derived spectrum. Reprinted with permission from Andrews ef al. (1993).

coaxial haem ligands as a pair of methionines was finally confirmed by the X-ray crystallographic determination of the E. coli bacterioferritin structure (Frolow et al., 1993). This work showed, as predicted (Grossman et al., 1992; Cheesman et al., 1993), that the haem-iron-coordinating residues are a pair of methionines (Met52 and Met52') from two-fold related subunits (see later, Section 4.1 and Fig. 8). Thus, the haems were located at the interfaces between two-fold related subunits, towards the inner surface of the protein shell. This provided an explanation for the general observation that there are 12 haems per bacterioferritin holomer, i.e. there are only 12 haem-binding sites, one at each of the two-fold axes. EPR spectroscopy showed that bacterioferritin contains mainly low-spin haem Fe3+, although some high-spin Fe3+ haem (g-values at 6.55 and 5.38 for P. aeruginosa bacterioferritin) was also present at a low proportion ( < 5 % ) of total haem (Cheesman et al., 1992). A broad, low-intensity band at 690-800 nm observed in the absorbance spectra of P. aeruginosa,

Table 3 Spectral parameters of the haem groups of bacterioferritins.

UV-visible (nm) MCD (nm) wavelength max of CT band

Species

Oxidized

Reduced

EPR (g-values) Oxidized

E. coli

418, -530, 650, 737

426, 528, 558

2.88, 2.31, 1.46

2270

P . aeruginosa

417, -530, 69@800

422, 526, 557

2.86, 2.32, 1.40

2240

A . vinelandii

417

425, 527, 558

2.88, 2.31, 1.46

2270

N. winogradskyi

418

426, 528, 559

Synechocystis

-420

425, 527, 559

A. spinosa

417, 530

424, 526, 557

R. sphaeroides

411. -540, 726

-425, 526, 557

2.86. 2.33, 1.46

Data are from Andrews el a/. (1993), Cheesman er a/. (1990). Moore er al. (1986), Stiefel and Watt (1979), Kurokawa el al. (1989), Laulhere et al. (1992) and Carrano et a/. (1996).

IRON STORAGE IN BACTERIA

30 1

A . vinelandii and E. coli holo- and apobacterioferritins (Fig. 5 and Table 3) was attributed to the bis-methionine-ligated haem group (Cheesman et al., 1992). This assignment was later confirmed by analysis of haem-free sitedirected variants (Andrews et d., 1995). A minor peak observed in the spectrum of E. coli bacterioferritin, at 650 nm, was also absent from the haem-free variants, indicating that this peak is likewise haem-derived (Andrews et al., 1993, 1995). Studies on the bacterioferritin from A . vinelandii (Watt et al., 1986) have shown that the haem redox potential is markedly affected by the presence (-475 mV) or absence (-225 mV) of an iron core, indicating that the core affects the electronic environment of the haem group. This may be related to changes in charge distribution around the haem group, resulting in a more negative local environment in the presence of core, or could result from a change in the net charge of the protein (Cheesman et al., 1992). Small changes in charge have previously been shown to have major effects on redox potentials of metalloproteins (e.g. Leitch et al., 1984). Low-temperature MCD and EPR spectra were not affected by the presence or absence of an iron core, indicating that the iron core does not influence the electronic state and stereochemistry of the haem group. This suggested that the haem is not directly influenced by short-range bonding interactions with the core (Cheesman et al., 1992) and, in turn, falsely indicated that the haem must be located towards the outer edge of the protein shell. Furthermore, experiments by Kadir and Moore (1990a) indicated that fully assembled bacterioferritin can incorporate externally added haem, to give 24 haems per holomer, in a state indistinguishable from that found in the untreated protein. This also incorrectly suggested that the haem-binding pocket must be oriented towards the outer surface of the protein shell and further suggested 24 haem-binding sites. Later studies showed that incorporation of haem by bacterioferritin in vitro is sample-dependent, much haem is incorporated in the high-spin state and that the earlier studies had been performed with ‘damaged’ protein (Moore et al., 1994). In an attempt to identify the haem-iron ligand and to determine the effect of the absence of haem on bacterioferritin, three conserved methionine residues (Met31, Met52 and Met86) in E. coli bacterioferritin were substituted for leucines and/or histidines (Andrews et al., 1995). The M52H and M52L variants were devoid of haem, whereas the other variants possessed full haem complements. This result confirmed the identity of the haem ligand determined by X-ray crystallography (Frolow et a/., 1993, 1994). The haem-free proteins were fully assembled, heat-stable proteins that could take up iron to form iron cores at rates similar to that of the wildtype protein. However, as isolated from the overproducing strains, the haem-free Met52 variants contained at least 4-fold more non-haem iron than the other overproduced variant or wild-type bacterioferritins. The

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C. ANDREWS

differences in iron contents were reflected in differences in the iron contents of the corresponding overproducing strains (Andrews et al., 1995). These studies clearly show that haem is unnecessary for iron uptake by bacterioferritin in vivo or in vitro. The higher iron contents of the haem-free bacterioferritins suggested that haem may either regulate the iron uptake and release steps in vivo or, more likely, that it could be involved in iron release through mediating delivery of electrons to the iron core. Such an effect has been demonstrated with haem incorporated into mammalian ferritins (Kadir and Moore, 1990b; Moore et ul., 1992). However, the removal of haem from A . vinelundii holobacterioferritin had no apparent effect on the rate of iron core reduction, indicating that haem does not facilitate iron core reduction, at least under the conditions employed (Watt et al., 1988). In contrast, a kinetic study of iron reduction and release from bacterioferritin has shown that, initially, the haem group is reduced, and then the iron core is reduced by the reduced haem (Richards et nl.. 1996) which suggests a role for haem in electron transfer. A two-phase iron release reaction was described in which the iron is removed rapidly in the first stage but about the last 145 iron atoms/holomer are removed more slowly presumably because they are bound to the protein or haem (Richards et al., 1996).

2.4. Bacterial Ferritins

The first evidence for haem-free ferritin-like iron storage proteins in bacteria was the discovery of an open-reading frame (gen-165,.f t n o r f m A ) in E. c d i encoding a predicted 165-residue polypeptide reported to possess 25% sequence identity to human H-chain ferritin (Izuhara et al., 1991). The ftnA gene was cloned as part of an attempt to isolate genes regulated by RNase 111 (involved in processing stable RNA species). The 550-nucleotide mono-cistronicffnA-gene transcript was found to be > 50% less abundant in RNase 111-deficient cells. The gene was located at 42 min on the E. coli chromosome, and potential transcriptional start and ribosome-binding sites were identified. The gene was overexpressed generating a 19 kDa polypeptide corresponding to the .finA product (FTN or FtnA). The ferroxidasecentre residues of human H-chain ferritin were absolutely conserved in E. coli FtnA, indicating the protein would be an active ferritin with the ability to catalyse the oxidation of iron (Andrews et al., 1991b). The first prokaryotic ferritin (or Ftn) to be purified was from the obligate anaerobe, Bacteroides,frugilis (Rocha ef ul., 1992). The protein was purified by heat denaturation and chromatography, and found to be a non-haemiron-containing protein with a native molecular mass of about 400 kDa. The subunits were 16.7 kDa in size and the N-terminal amino acid sequence was very similar to those of eukaryotic and prokaryotic ferritins (43% identity to

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human ferritin, 37% identity to E. coli FtnA), but was much less similar to the bacterioferritins (-20% identity). The lack of haem and the sequence similarity data clearly placed the B. fragilis Ftn among the ferritins, rather than the bacterioferritins. Although the protein stained positively for nonhaem iron and incorporated ”Fe in vivo, the iron content of the purified protein was only three iron atoms per molecule, suggesting that it may not have an important iron-storage role but could play a part in iron detoxification, especially during transient exposure to oxygen (Rocha et al., 1992). Another prokaryotic ferritin-like protein (designated Pfr) was isolated from the microaerophilic Helicobacter pylori and the corresponding gene was cloned and sequenced (Doig et al., 1993; Frazier et al., 1993). The sizes of the Pfr subunit and Pfr 24-mer were measured as 19.3 and 350 kDa, respectively. The Pfr amino acid sequence was found to be 42% identical to E. coli FtnA, but only 17% identical to E. coli bacterioferritin, and the seven highly conserved ferroxidase centre residues of human H-chain ferritin were, as for the E. coli ferritin, conserved. Inclusion bodies observed in H . pylori P466 were labelled by anti-Pfr antibodies suggesting that the inclusion bodies were ‘enriched’ with Pfr. No inclusion bodies were seen in H . p-vlori strain MO 19, although there was cytosolic labelling with Pfr-antibodies. This study indicated that Pfr is a highly abundant protein in H . pylori, although the absolute amount of Pfr was not reported nor was the proportion of total cell iron incorporated into Pfr. Pfr was overproduced in E. coli to 49% of total protein, resulting in the formation of inclusion bodies and a 10-fold increase in the amount of cellular iron, indicating that the overproduced protein contains iron (Frazier et al., 1993). Doig et al. (1993) purified Pfr from H . pylori as a 19.6-kDa polypeptide which apparently had binding affinity for human erythrocytes, epithelial cells and laminin. Upon isolation, the protein was shown to be a non-specific binding protein containing up to 1700 iron atoms per molecule, having a high phosphate content (1.4: 1 molar ratio of Fe to phosphate) tnd appearing under the electron microscope as a ‘ringlike’ protein of 91 A diameter with an electron-dense central core. Its lack of haem identified it as a ‘ferritin’ rather than a ‘bacterioferritin’ and its Nterminal amino acid sequence matched that of the Pfr protein described by Frazier et al. (1993). Subcellular fractionation revealed that most (5680%) of the protein was cytosolic. The.ftnA gene of E. coli was later overexpressed by Hudson et al. (1993) simply by providing the gene on a multicopy plasmid and relying upon expression from the natural promoter and translational initiation region. The overproduced FtnA, representing up to 14% of total protein, was purified and found to lack haem and possess properties similar to those of eukaryotic ferritins. Overproduction of FtnA resulted in a 2.5-fold increase in cellular iron, although FtnA contained only five to 20 iron atoms per molecule upon purification. Low iron contents have been reported for other

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overproduced ferritins and bacterioferritin ( E . coli bacterioferritin, 25-75 Fe/molecule; human H-chain ferritin, 100-1 30 Fe/molecule; human Lchain ferritin, 5-1 5 Fe/molecule; H . pylori Pfr, 24 Fe/molecule; see Hudson et al., 1993), suggesting that iron-storage proteins have a relatively passive role in iron homeostasis since their iron contents appear to reflect intracellular iron availability. The protein was able to accumulate up to 2000 iron atoms per molecule at rates similar to those of human H-chain ferritin. The presence of an iron core had no discernible effect on the surface properties of FtnA, showing that the core was completely encompassed by the protein shell. Electron microscopy revealed the characteristic ferritin morphology for holo-FtnA (a spherical shell of 120 diameter and inner core of 79 diameter). The core generated in vitro was shown to be ferrihydrite-like when it had been formed in the absence of phosphate, but in the presence of phosphate an amorphous non-crystalline core was produced. These properties reflect those of mammalian ferritins (see Section 5.2). During purification, the protein was prone to proteolytic cleavage at sites in the region between the predicted D and E helices, and between the E helix and the C-terminus. This proteolysis was prevented by chelators, free radical scavengers, reductants or by removal of iron from the protein. This suggested that FTN cleavage was caused by free radicals generated by an ironmediated Fenton reaction. Whether or not free-radical-induced proteolytic degradation of FTN is physiologically relevant (i.e. in iron release, as proposed for pea-seed plastid ferritin; Laulhere et al., 1989) is uncertain. In addition to those from B. fragilis, E. coli and H . pylori, ferritins have been isolated and/or a ferritin-encoding gene cloned from the following bacteria: Campylobacter jejuni (Wai et al., 1995, 1996), Haemophilus influenzae (Fleischmann et al., 1995), Methanobacterium thermoautotrophicum, Clostridium acetobutylicum, Thermatoga maritima, A rchaeoglobus julgidus, Mycobacterium tuberculosis and Vibrio cholerae (Table I ). Thus, like bacterioferritins, ferritins would appear to be relatively widespread within eubacteria, being present in proteobacteria, bacteroides, high- and low-G + C Gram positive bacteria and thermotogales. In contrast to the situation for bacterioferritins, ferritins are also present in archaebacteria ( M . thermoautotrophicum and A . fulgidus contain genes encoding ferritin-like proteins). Therefore, there is firm evidence for the presence of ferritins in all three domains of life, whereas current evidence indicates that bacterioferritins are restricted to the eubacterial domain. However, the current lack of evidence for bacterioferritins in archaebacteria (or other groups), should not be mistaken for conclusive proof of their absence from such organisms. It is likely that as more microbial genome sequences become available this appraisal will have to be revised. Interestingly, the ferritin gene of M . thermoautotrophicum appears to be in an operon consisting of two rubredoxin genes, followed by a flavoprotein

A

A

IRON STORAGE IN BACTERIA

305

A gene, then the ferritin, alkylhydroperoxidase, and Mn/Fe-superoxide disIt is mutase genes (http://www.cric.com/htdocs/sequences/methanobacter). possible that the products of these genes are required under redox-stress conditions, suggesting a role for the ferritin of this bacterium in redox-stress resistance, as is the case for the ferritin of C. jejuni (Section 7.1). Alternatively, the putative operon may be induced by high iron availability (e.g. as for the sodB and ftnA genes of E. coli; Niederhoffer et al., 1990; Section 7.1).

3. PRIMARY STRUCTURES AND EVOLUTION OF IRONSTORAGE PROTEINS 3.1. The Ferritin-Bacterioferritin-Rubrerythrin Superfamily

As described above, two types of iron-storage protein are known, the haemcontaining bacterioferritins and the haem-free ferritins. The bacterioferritins and ferritins are structurally, functionally and evolutionarily related to each other and to the rubrerythrins (Andrews et al., 1991b; Grossman et al., 1992; Harrison et al., 1998; Figs. 6 and 7). These proteins therefore constitute a superfamily, designated the ferritin-bacterioferritin-rubrerythrin (F-B-R) superfamily. A multiple amino acid sequence alignment of the bacterioferritins, bacterial ferritins and rubrerythrins (together with four eukaryotic ferritins) is shown in Fig. 6. The alignment was constructed using the ‘Pileup’ program and was optimized by reference to conserved secondarystructure elements. Although the bacterioferritins and ferritins are likely to be related along the entire lengths of their polypeptide chains, the rubrerythrins are only related to the iron-storage proteins in the four-helix bundle region, not in the C-terminal rubredoxin/E-helix region. The E-helix region is the least conserved part of the iron-storage proteins. Indeed, this region can be deleted from human H-chain ferritin without any major structural or functional consequences (Levi et al., 1988). This indicates that it does not have an important function, which would explain its poor conservation. The sequence similarity is fairly evenly distributed within the four-helix bundle region of the alignment. Some residues are very well conserved in all three subfamilies. The best examples are Leu 1 1, Glu 18, Tyr25, Gly36, Glu5 1, His54, Ala55, Glu85, (311,194,Glu127, Leu138 and Ile141 ( E . coli bacterioferritin numbering). These 12 residues include five di-iron ligand residues and an absolutely conserved Tyr residue which is near the di-iron site and has been shown to be vital for full ferroxidase activity in mammalian ferritins (Harrison and Arosio, 1996). There is no region that is exceptionally well conserved in comparison with other regions. This is probably because such

H H H

E E E

E

E

C

IRON STORAGE IN BACTERIA

309

regions normally represent active sites and the active site of the F-B-R proteins are composed of residues from non-contiguous parts of the polypeptide. Therefore, although there are six highly conserved ferroxidase centre residues within the F-B-R family members, these residues are widely distributed along the polypeptide chain and so do not form a clear pattern or motif. Very few padding characters were required to optimize the alignment. Of the 22 padding characters inserted in the E. coli Bfr sequence, only one lies in the A helix, none is in helix B, one is in C helix, and two are in helix D. This supports the validity of the alignment since large numbers of

Figure 6 Optimum multiple-amino-acid-sequence alignment of bacterial ferritins, animal ferritins, bacterioferritins and rubrerythrins. The ‘gcg/egcg’ programs (with default settings) ‘Pileup’, ‘Elineup’ and ‘Prettyplot’ (Devereux, 1989) were used to produce and display the optimal alignment of eight bacterial ferritins (one incomplete), four representative animal ferritins, twenty bacterioferritins (six incomplete) and seven rubrerythrins (one erythrin) amino acid sequences. The regions corresponding to the a-helices of E. coli bacterioferritin (A-E), and key residues of E. coli FtnA (bacterial ferritin), human H-chain ferritin (animal ferritin), E. coli bacterioferritin (bacterioferritin) and D . vulgaris rubrerythrin (rubrerythrin) are shown below the alignment: E, H and Q , dinuclear metal ligands; Y, conserved tyrosine implicated in the ferroxidase activity of ferritins; C, FeS4 ligand for rubrerythrins; M, coaxial haem-iron ligand for bacterioferritins; E, glutamate residue of unknown function that changes position in bacterioferritin upon occupation of the dinuclear metal site; and E, ligands unique for metal site C of bacterial ferritins. Residue numbers for each sequence are shown on the left. The database accession numbers or sources of the sequences used (and corresponding organisms) are below. Bacterial ferritins: eco-ftnb/ecoliftn (E. coli), D90832/P23887; hiftnl/2 (Haenrophilus influenzae), P43707/P43708; hpylorif (Helicobacter pylori), P52093; cjejuni (Campylohacter jejuni), D64082; mth-ftn (Methanobacterium thermoautotrophicum), www.cric.com/htdocs/ sequences; and bfragftn (Eacteroides jragilis), Rocha et al. (1992). Animal ferritins: sch-man2 (Schistosoma mansoni), P25320; humanh (Homo sapiens), P02794; and human1 ( H . sapiens). Bacterioferritins: alceubfr (Alcalagenes eutrophus), M69036; syn-bfr2 (Synechocystis PCC6803), D90905; ngo-bfrb (Neisseria gonorrhoeae), U76634; mmbfr I (Magneiospirillum magnetotacticurn), L.E. Bertani (personal communication); absibfra (Absidia spinosa), Carrano et al. (1996); ecobfr (E. coli), PI 1056; pae-bfra (Pseudomonas aeruginosa), Moore et al. (1994); avibfr (Azotobacter vinelandii), P22759; brucella (Brucella melitensis), P49944; rcap-bfr (Rhodobacter capsulatus), 254247; nwino-bfr (Nitrobacter winogradskyi), Kurokawa et a / . (1989); mavbfr (Mycobacterium avium), P433 14; mpara-bfr (Mycobacterium parutuherculosis), Brooks el al. (1991); mlebfr (Mycobucterium Ieprae), P433 15; mmbfr2 (M. magnetotacticum), L.E. Bertani (personal communication); syn-bfrl (Synechocystis PCC6803). D909 16; ngo-bfra ( N . gonorrhoeae), U76633; absibfrb ( A . spinosa), Carrano et al. (1996); putida-bfr (Pseudomonas putida), U667 17; and pae-bfrb (Pseudomonas aeruginosa), Moore et a/. (1994). Rubrerythrins: cyanopho (Cyanophora paradoxa cyanelle), P48379; mth-rerb (M. thermoautotrophicum), see mth-ftn; desvh-nige (Desulphovibrio vulgaris strain Hildenborough), U71215; ruby-des (nigerythrin from D . vulgaris), P2493 1; ruby-cpr (Clostridium perfringens), P5159 I ; ruby-met (Methanococcus jannaschii), U67520; and mth-rera (M. thermoautofrophicum), see mth-ftn.

310

SIMON C. ANDREWS

Rubrerythrins

SCALE

1

M Innnor hrr M IhrrmooiIlolnJptinrm

Ncr

C perfringrnr

D

vul#arilrrs

Figure 7 Unrooted phylogenetic tree showing the evolutionary relationship between members of the ferritin-bacterioferritin-rubrerythrin superfamily. The programs ‘Protdist’ (with the Dayhoff PAM matrix), ‘Fitch’ and ‘Drawtree’ from the ‘Phylip’ package were used to create the phylogenetic tree (Felsenstein, 1989, 1993). The input file for Protdist consisted of an abbreviated form of the alignment shown in Fig. 6 in which the C-terminal rubredoxin-like regions of the rubrerythrin proteins were omitted, as were incomplete sequences.

padding characters in the major structure element regions would be inconsistent with common structures and evolutionary relatedness. Although the bacterioferritins and ferritins have many features in common, their sequence similarity is limited, indicating that they are very distantly related in evolution (Figs 6 and 7; Andrews et al., 1991b; Grossman et al., 1992). This is illustrated by comparing the sequence identities between E. coli bacterioferritin and the ferritins (12-21 YO)or other bacterioferritins (41-67%) in Fig. 6, and between E. coli FtnA and the other ferritins (19.57%) and the bacterioferritins (1 1-1 8 %). Likewise, the sequence similarities between the rubrerythrins and the bacterioferritins or ferritins is relatively weak: the amino acid sequence identity of E. coli bacterioferritin or E. coli FtnA and the rubrerythrins in Fig. 6 is 13-18% or 1&15%; and the identities between D. vulgaris rubrerythrin and the ferritins, bacterioferritins or the other rubrerythrins in Fig. 6 is 8-17%, 11-19% and 25-54%, respectively. This limited sequence identity between the three groups of the F-B-R

IRON STORAGE IN BACTERIA

31 1

superfamily indicates that the bacterioferritins, ferritins and rubrerythrins form distinct families within the F-B-R superfamily. This notion is supported by the phylogenetic analysis shown in Fig. 7: the bacterioferritins, ferritins and rubrerythrins clearly constitute evolutionarily distinct domains within the F-B-R superfamily. One of the biggest surprises in recent studies on bacterioferritins was the discovery of a bacterioferritin in a fungus, Absidia spinosa (Carrano er a/., 1996)- as all other known bacterioferritins have been found in eubacteria. The A . spinosa bacterioferritin contains two subunits (aand p) and is thus a heteropolymer. In this respect it is like the heteropolymeric bacterioferritin of P. aeruginosa (Moore et al., 1994). Furthermore, there are close sequence similarities between the CY subunits (65% identity) of these two organisms, and also between the subunits (78% identity). Also, phylogenetic analysis indicates that the two CY subunits share a very recent common ancestor, as do the two 0 subunits (data not shown). This suggests that either a very recent horizontal gene transfer (bacteria to fungi) has occurred or that the A . spinosa bacterioferritin is from a bacterium, not a fungus.

3.2. Bacterioferritin Heteropolymers

Interestingly, there are now five examples of organisms containing two types of bacterioferritin subunit. These are Neisseria gonorrhoeae, Synechocystis, P. aeruginosa, A . spinosa and Magnetospirillurn magnetotacticurn (Fig. 6; Table 1). It is likely that in each case the two different subunits coassemble to form heteropolymers (see below). It also appears that in each case, one subunit confers haem-binding ability upon the heteropolymer whereas the other confers ferroxidase activity. This is somewhat reminiscent of the situation for mammalian ferritins where only one of the two subunits possesses a ferroxidase centre; Section 2.1). One of the two bacterioferritin subunits of N. gonorrhoeae, M . magnetotacticurn and Synechocystis (subunits A, 2 and 1, respectively) contains a full complement of ferroxidase centre residues but no coaxial haem-ligating Met (instead they contain Thr or Leu which are not thought to function as haem ligands), whereas the other subunit (B, 1 and 2, respectively) would appear to lack a functional active site (Gln, Leu or His in place of Glu 18; Ala in place of Glu5 1 for Synechocystis Bfr2 only; Gln or Ala in place of His54; Asn or deletion in place of Glu127) but does contain the coaxial haem-ligating Met residue. The partial sequences of the P. aeruginosa and A . spinosa bacterioferritin subunits are insufficient to reveal for certain whether a similar arrangement exists for these bacterioferritins. However, the N-terminal sequences of P. aeruginosa subunits extend far enough to show that for BfrB the haem-ligating Met is replaced by a Thr residue and that the first four ferroxidase centre residues are

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maintained, whereas BfrA contains the Met52 residue but also the first three ferroxidase centre residues. For the A . spinosu subunits, BfrA contains one out of the first two ferroxidase centre residues whereas BfrB contains both of the first two. Therefore, the partial sequence data are mostly consistent with the BfrB and BfrA subunits of A . spinosa and P . ueruginosa acting as haem-binding and ferroxidase active subunits, respectively, although i t is possible that both subunits of P . aeruginosu bacterioferritin contain ferroxidase centres. The phylogenetic tree (Fig. 7) shows that the 'haem-free' subunits are clustered into a subfamily (which includes the BfrB subunits of A . spinosu and P . aeruginosn; data not shown). This subfamily also includes a Psmdomonus putida bacterioferritin subunit which also lacks the Met52 residue but contains a full complement of ferroxidase centre residues. The corresponding 'ferroxidase-inactive haem-containing' subunits are also clustered into a subfamily (which includes the BfrA subunits of A . spinosa and P . pseudomonas. data not shown). It is likely that P. putida contains a second as yet undiscovered bacterioferritin subunit that falls into the 'haem-containing' subfamily. In the cases of the bacterioferritins of P . nerirginosn and A . spinosa, it appears that the BfrA and BfrB subunits form heteropolymers since the purified proteins each contain the corresponding subunits, distinguished on the basis of differing sizes and N-terminal sequences (Moore et a/., 1994; Carrano et d.,1996). For the bacterioferritin subunits of Sjnechocystis, N . gonorrhoear and M . magnetotacticuni, only nucleotide sequences are available but it is highly likely that they also co-assemble to form heteropolymers. However, the S.vnechoc:g.sti.s bacterioferritin has been purified and only a single N-terminal sequence was obtained which corresponded to subunit 1 that lacks Met52 (Laulhere ci d., 1992). Nevertheless, it is still possible that a second minor subunit was present that was not revealed by the PAGE or amino acid sequencing analyses. The two bacterioferritin genes found in both N . gonorrhoear and M . magnrtotacticurn are adjacent on the chromosome and are co-polar, indicating that they could be co-transcribed. The two bacterioferritin genes of Svnei~hoc~vtis are not co-located on the chromosome and therefore could be independently regulated. The physiological rationale for the presence of two types of subunits, which appear to confer different functional properties, is unknown. One possibility is that the subunit composition varies according to environmental conditions (e.g. iron availability) resulting in appropriate changes in the properties of the heteropolymeric bacterioferritin. The subunit composition of P . uerzrginosn bacterioferritin has been shown to vary from preparation to preparation, although the conditions influencing this variability and the significance have not been reported (Moore et ul., 1994). Also, the haem content of P . ncwrginosa bacterioferritin is highly variable. This could reflect the subunit composition: presumably, high levels of the a subunit would

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result in relatively high haem contents, whereas high levels of the /3 subunit would cause low haem contents. The reverse trend might be expected for ferroxidase activity. However, no functional consequence of variable subunit composition has been reported and such studies are eagerly awaited.

3.3. Evolution of the Ferritin-Bacterioferritin-Rubrerythrin Family from a Simple Two-helix Protein

The N-terminal A-B helices and the central C-D helices of rubrerythrin display sequence similarity and thus rubrerythrins possess internal homology within their four-helix bundles (Kurtz and Prickril, 1991). This strongly suggests that the rubrerythrins (and therefore the related ferritins and bacterioferritins) have arisen from an ancient and simple two-helix-containing protein, through duplication and fusion of the encoding gene, followed by divergence. Intra-polypeptide homology is particularly apparent for the rubrerythrin-like predicted protein (designated ‘erythrin’) from the cyanelle organelle of Cyunophora paradoxu (a flagellated, freshwater protozoan containing an endosymbiotic cyanobacterium called the cyanelle). This predicted protein lacks the C-terminal rubredoxin domain (and so is not a true rubrerythrin); comparison of the central segments of its A-B (residues 9-49) and C-D (residues 126-166) helix regions shows that they are highly similar (39% identical) in amino acid sequence. The function of erythrin is unknown, although it possesses all of the di-iron cluster ligands found in the other rubrerythrins (Fig. 6) and is therefore likely to form a di-iron centre. The C. puruci‘oloxa protein is the most distantly related member of the rubrerythrin group (possessing an average of only 22-28% sequence identity when compared with the other six members in Fig. 6) and is more closely related to the ferritins and bacterioferritins than are the other rubrerythrin family members (Fig. 7). The common ancestor of the F-B-R superfamily is likely to have been a simple protein displaying strong sequence identity between its N- and C termini, similar to the C . paradoxu protein, probably more rubrerythrin-like than ferritin-like in terms of subunit composition (i.e. monomeric or dimeric). It is therefore possible that the C . paradoxu protein closely resembles the original common ancestor of the F-B-R superfamily. For this reason. the branch leading to the F-B-R ancestor has been placed within the rubrerythrin branch, close to the C . paradoxu branch. There is little sequence similarity between the A-B and C-D helix regions of the ferritins and bacterioferritins, indicating that in contrast to the rubrerythrins, the Nand C-terminal halves of these proteins have diverged markedly through the course of evolution.

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3.4. The Ubiquity of Bacteriofertitins and Ferritins

The ability of living organisms to utilize and sequester iron safely is likely to be a fundamental requirement. The increasing number of completely or almost completely sequenced microbial genomes allows us to begin to determine whether ferritins/bacterioferritins are indeed ubiquitous in cellular organisms, and therefore to reveal how fundamental the need for an ironstorage capability is. Of the 19 bacterial genome sequences that have been completed (or nearly completed) and released to the public, ferritins or bacterioferritins are found in 14 (Table 1). No ferritins/bacterioferritins were found in the two Mycoplasma species, in Streptococcus pyogenes, Treponema pallidum or in the archaebacterium Methanococcus jannuschii. Ferritins are also apparently absent in Saccharomyces cerevisiue (http://genome-www.stanford.edu; but note that a ferritin from this organism has been isolated and characterized; Raguzzi et al., 1988). It is possible that the latter organisms possess ferritins or bacterioferritins having very low sequence similarity to other family members and are therefore not recognizable. Alternatively, these organisms may not utilize iron-storage proteins. This is likely to be the case for the Mycoplasma species which are obligate intracellular parasites having extremely small genome sizes. (Mycoplasma genitallizim possesses only 470 coding regions and lacks many metabolic pathways considered essential for free-living organisms). The finding that four free-living microorganisms do not appear to possess iron-storage proteins suggests that such proteins are not ubiquitous. 3.5. Multiple Iron-storage Proteins within Bacteria

Ferritins and bacterioferritins are sometimes found together within a single bacterium (e.g. in E. roli, Vihrio cholerae, Mycohacterium tuberculosis; Table 1). As already seen, some bacteria possess two bacterioferritin subunits which appear to co-assemble. However, other bacteria possess two ferritin subunits (e.g. Haemophilus influenzae, E. coli; Table 1). The two H . influenzae ferritin subunits are 64% identical and are encoded by genes that are adjacent on the chromosome and are co-polar, suggesting co-transcription. The ferroxidase centre residues are conserved, indicating that both subunits would confer ferroxidase activity. It is not therefore clear why H . injiuenzue possesses two such similar subunits, but given their high sequence identity it is probable that they co-assemble to form heteropolymers. The two ferritins of E. coli are also encoded by adjacent, co-polar genes, although there is an -1.4 kb mostly non-coding, intergenic region that separates them. Although the presence of the ftnA gene (previously calledftn) has been known for some time, the second gene (ftnl? or yecl) was revealed only recently

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through genome sequencing studies. Two complete nucleotide sequences are available for the,finB gene (accession no. P52091 and D90832), which differ slightly. The P52091 sequence would appear to contain several errors affecting the predicted translational-initiation codon, the amino acid sequence at the C-terminal region of the translation product and the codon-reading frame. The predicted amino acid sequence of the,ftnB-encoded FtnB subunit (accession no. D90832) is only 33% identical to that of FtnA, and possesses several ferroxidase centre substitutions which suggests that it will not confer ferroxidase activity, although it may have the capacity to bind metal at the di-iron site (see later, Fig. 9). The inter-subunit contacts observed in the FtnA X-ray crystal structure are not conserved in FtnB indicating that it does not coassemble with FtnA (P.M. Harrison, personal communication).

3.0. A Relationship between the Ferritin-BacterioferritinRubrerythrin Superfamily and the Dps Family?

Recently, it has been proposed that there is an evolutionary linkage between the bacterioferritin and Dps families (Peiia and Bullerjahn, 1995) and this relationship has been extended by Evans et a/. (1995) to include the ferritins. Any such relationship would be expected to extend to the rubrerythrins also. The Dps proteins are DNA-binding, dodecameric, hexagonal-ring-forming proteins (possibly haem-containing) from bacteria that are induced by stress. They are believed to provide resistance to oxidative stress and starvation by protecting DNA. A similarity of 55% in amino acid sequence was identified in the C-terminal regions of A . vinelandii bacterioferritin and Synechococcus DpsA, and a weaker similarity between the N-terminal regions. Other proteins in the Dps family did not have high sequence similarity to the bacterioferritins (Peiia and Bullerjahn, 1995). A multiple alignment (Peiia and Bullerjahn, 1995) comprising a ‘bacterioferritin consensus sequence’ and the Dps proteins was interpreted as revealing ‘shared residues across both groups’, suggesting a common origin. However, virtually none of the highly conserved residues in the bacterioferritin group was well conserved in the Dps group (none of the seven ferroxidase centre residues of bacterioferritins is highly conserved in Dps proteins, only one of the 12 highly conserved F-B-R residues described in Section 3.1 are conserved in Dps proteins). The multiple alignment reported by Evans et d.(1995) included ferritins as well as bacterioferritins and Dps proteins, and the relationship between sequences was very different to that reported by Peiia and Bullerjahn ( 1 995). The reported multiple alignment incorporated many padding characters required for alignment optimization and these were in positions that would be expected to lead to gross structural perturbations (22 padding characters in the E. coli bacterioferritin sequence, of

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which 19 lie in the A-, B-, C - or D-helix regions). Furthermore, highly conserved ferroxidase centre residues of the F-B-R family were not maintained in the Dps group, and the alignment of the bacterioferritins and ferritins differed from that of Fig. 6 (e.g. Glu127 of E. coli bacterioferritin was not aligned with the structurally equivalent Gln127 of E. coli FtnA). ‘Profile Analysis’ searches of the databases using a profile of aligned F-B-R sequences failed to detect sequence similarity between the bacterioferritin and Dps proteins, as did sequence comparisons using BLAST or FASTA (S.C. Andrews, unpublished). Also, there were very few residues that were well conserved in both the Dps proteins and the F-B-R proteins. It must be concluded, therefore, that the validity of the proposed relationship between bacterioferritins (and ferritins) and the Dps family remains doubtful, as has been suggested previously (Penfold et al.. 1996). The four prokaryotic proteins described as new members of the bacterioferritin family (NapA of H . pvlori, the mrgC product of Bacillus subtilis, two proteins encoded by unidentified genes from Anabaena vuriabilis and Treponema pullidurn; Evans et al., 1995) are likely to be Dps proteins, not bacterioferritins. Very recently, a novel non-haem-iron binding protein, related to the Dps proteins, was isolated from Listeria innocua (Bozzi et al., 1997). The protein was isolated on the basis of its iron content (5-10 Fe atoms per holomer) and heat stability, and was found to be composed of 18 kDa subunits and to have a native molecular mass of 240 kDa. It was able to catalyse iron oxidation at rates equivalent to those of ferritin to form an iron core of 500 ferric atoms. The cores were located in the centre of the doughnutshaped protein. Like the ferritins, this Dps-like ferritin is predicted to possess four long a-helices. Some limited sequence similarity was observed between mammalian ferritin L-subunits and the Dps-like ferritin, but no convincing similarity could be detected with other ferritins or bacterioferritins. However, this was regarded as proof of an evolutionary relationship between ferritins/bacterioferritins and Dps proteins. The work of Bozzi et al. (1997) strongly suggests an iron-related role for the L. innocua Dsp protein, but any relationship with ferritins remains to be made convincingly and the possibility that other Dps proteins have a role in iron metabolism has yet to be tested. 4. STRUCTURES OF BACTERIOFERRITINAND BACTERIAL

FERRlTlN 4.1. E. coli Bacterioferritin

Initial X-ray crystallographic studies on four crystal forms of E. coli bacterioferritin showed that the molecule has a diameter of 119-128 and 24

A

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subunits arranged in 4-3-2 symmetry (Smith et al., 1988a, 1989). Attempts to solve the structure by isomorphous replacement were unsuccessful (Smith et al., 1990) and so model structures of E. coli bacterioferritin were generated (Smith, 1991; Cheesman et a/., 1993). The X-ray crystal structure of the bacterioferritin of E. coli was eventually solved at 2.9 A resolution and shown to closely resemble those determined for animal ferritins (Section 2.1). The 24 subunits of bacterioferritin form a roughly spherical -120 protein shell surrounding a central -80 A cavity (Frolow et NI., 1994). The subunits are arranged in a manner approximating a rhombic dodecahedron (i.e. a polygon with 12 rhombic faces). Each subunit folds into a four-helix bundle (helices A-D) with a short C-terminal helix (helix E) lying on top of the bundle at an angle of 60“ to that of the bundle-central axis. The 12 haem h groups are located at the two-fold axes, in a groove between twofold-related subunits, towards the inner surface of the protein shell (see Fig. 8). As suggested by EPR and NIR-MCD spectroscopy, and protein engineering studies (Cheesman er al., 1990, 1992; McKnight et al., 1991; George et a/., 1993; Andrews et al., 1995), the haem-iron is coaxially coordinated by the thio-ether side chains of two methionine residues, Met52 and Met52’, one from each of the two-fold related subunits (Frolow et al., 1993). The distance between the haem iron and sulphur of Met52 is 2.4 which compares well to the value of 2.35 determined by EXAFS (George et al., 1993). The interaction between the ligating methionine residues and the haem group is extensive. The two methionine side chains adopt an extended conformation running along the opposite faces of the haem group contributing over 58 out of the 108 van der Waals contacts between protein and haem, and thus acting to clamp the haem in place. A pair of two-fold related Phe26 residues wedge the haem in place from opposite sides. The two haem-propionyl groups project inwards into the central cavity. There is no obvious access to the haem from the outer surface. As for most types of ferritin, a dinuclear metal-binding site (ferroxidase centre) is located in the middle of each of the four-helix bundles (Frolow er a/., 1994; Le Brun et al., 1995). The metals are approximately 4 A apart and are 12.5 A from the haem iron. I t is likely that the two metal atoms located at this site in the bacterioferritin structure are Mn(II), since the crystals were generated in the presence of this metal and Mn(I1) is reported to bind to the ferroxidase centre (Thomson er al.. 1997). Four glutamate and two histidine residues act as ligands to the dinuclear metal species; Glu51 and Glul27 act as bidentate-bridging ligands between metal atoms A and B, Glu 18 and His54 act as monodentate ligands to ‘metal A’, and Glu94 and His130 are monodentate ligands to ‘metal B’. This arrangement of metalbinding ligands differs from those of human H-chain ferritin and E. coli FtnA (see Section 4.3).

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B

Figure 8 Ribbon diagrams of the alpha-carbon backbones of an E. coli bacterioferritin subunit dimer showing the haem and dinuclear metal sites. (A) View along the twofold interface between a pair of subunits. The outer surface of the bacterioferritin molecule is towards the top of the page, the inner surface towards the bottom. The haem is represented by a stick model, the haem iron by a large black sphere, and the dinuclear metal atoms by small black spheres. The haem propionates are seen to project into the central cavity. (B) View of the two-fold interface, looking outwards from the inside of the molecule. Derived from Frolow ei al. (1994).

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Since the haem group is sandwiched between two ferroxidase centres (one from each of the two-fold-related subunits) at a distance ofjust 12.5 from each of the corresponding dinuclear centres, it is possible that haem mediates electronic interaction between the dinuclear centres. Also, as the haem group projects into the central iron-storage cavity, it is possible that the haem enables electron interaction with the iron core. Studies have shown that the redox status of the haem group does not alter during the ironuptake process; however, initial iron-binding does result in a small, transient and very rapid blue/red shift in the position of the haem Soret band, followed by a slower red/blue shift (Le Brun, 1993; Le Brun et d.,1993b). This may be related to the close juxtapositions of the haem and dinuclear ironbinding residues (Met52 and Glu51). The homology between the bacterioferritins and ferritins had earlier allowed the three-dimensional structure of E. coli bacterioferritin to be modelled upon that of human H-chain ferritin (Cheesman et al., 1993). This model turned out to be relatively accurate. Cheesman et al. (1993) identified two potential haem-binding sites containing methionine residues. These were an intra-subunit site employing Met31 and Met86, and the intersubunit site employing Met52 which was subsequently shown to be correct. The inter-subunit site was also correctly predicted by Grossman el a/. (1992). The dinuclear metal site ligands were correctly anticipated by Cheesman et al. (1993), including the use of His130 rather than Asp5O. Also, a potential role for Glu47 in metal binding at the dinuclear site was highlighted. More recently, an X-ray crystallographic analysis of E. coli bacterioferritin site-directed variants carrying either M52H or E18A substitutions has been carried out on crystals of the apoproteins (iron-free) generated in the absence of added metal (Barynin et al., 1997). This study has shown that there are significant differences in the positions of three side chains (Glu47, Tyr25 and Hisl30) located at or near the ferroxidase centre, depending on whether the di-iron site is occupied or unoccupied. The absence of metal at the di-iron site allows the His130 side-chain to disengage from its position as a terminal ligand at Fe site B, and instead it is swung by approximately 90" and rotated so that it is oriented towards the inner surface where it interacts with Glu47. Consequently, the Glu47 side chain is also drawn from a position where it is directed towards the inner surface, to a position where it is close to Fe site B. These side-chain movements result in an adjustment in the position of the highly conserved Tyr25 residue. Although the mechanistic significance of these movements was not reported, it is possible that the Glu47 residue replaces His130 as a terminal ligand at Fe site B during the initial stages of the iron-binding/oxidizing reaction, and that its subsequent repositioning towards the inner surface provides a third iron-binding site for core nucleation and/or migration of iron from the di-iron site.

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The structural consequences of the M52H mutation and lack of haem are closer only slight: the two-fold-related subunits are approximately 0.5 together, and the His52, Phe26 and Tyr45 residues move into the space left by the absent haem group.

A

4.2. E. coli Ferritin (FtnA)

The ferritin (FtnA) of E. co/i has been crystallized in the absence of metal ions and its three-dimensional structure solved by X-ray diffraction (Hempstead et al., 1994). A full description of the structure has not yet been given, although it is reported to be very similar to that of other ferritins. The absence of metal in the crystallization medium allowed iron (in the ferrous form) to be soaked into the crystals. As a result, electron-density maps of both the iron-free (2.4 resolution) and iron-containing (3.0 A resolution) forms of FtnA were obtained. Three iron-binding sites (originally presumed to contain ferric iron, now thought to contain ferrous iron; Harrison et d., 1998) were observed in iron-containing FtnA. A pair of iron were present at the atoms (Fe site A and Fe site B), separated by 3.8 ferroxidase centre. These were ligated by one histidine and four glutamate residues: Glu50 acts as bidentate-bridging ligand between Fe site A and Fe site B; Glu17 (bidentate) and His53 (monodentate) are ligands to Fe site A; and Glu94 (bidentate) and Glu130 (monodentate) act as ligands to Fe site B (see Fig. 9). A sixth potential ligand, Gln127, may be linked to Fe site A or Fe site B via water. FtnA crystals soaked with TbCI3 had a pair of Tb(II1) atoms positioned and ligated in the same way as Fe site A and Fe site B. TbCI3-soaked human H-chain ferritin crystals also had a pair of Tb(lI1) atoms, 3.1 A apart, in very similar positions to Fe site A and Fe site B of FtnA (Lawson et al., 1991). However, the coordination pattern of the FtnA di-metal site differs slightly from those of human H-chain ferritin and E. c d i bacterioferritin (see Section 4.3). The third iron site (Fe site C) is unlike any metal site yet seen in the eukaryotic ferritins or in the bacterioferritins. It possesses four ligands, all glutamates: Glu49, -126. -129 and -130. Glu130 acts as a ligand for both Fe site C and Fe site B, and it is suggested that it has two alternative positions depending on which of the two iron sites it binds. All three iron sites are reported to have high occupancy, and therefore are probably jointly occupied. It is suggested that Fe site C could represent the monomeric Fe(II1) species observed by Mossbauer spectroscopy (Bauminger et af., 1994), and since it is on the inner surface it could be the core nucleation site.

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4.3. Comparison of Ferroxidase Centres

The dinuclear iron sites of the F-B-R family members resemble those of ribonucleotide reductase (R2 subunit), methane monooxygenase (hydroxylase subunit) and haemerythrin (Fig. 9; Norlund et al., 1990; Holmes and Stenkamp, 1991; Atta et al., 1992; Rosenzweig et al., 1993). Like the F-B-R proteins, these proteins contain their dinuclear iron sites at the centres of four-helix bundles. However, no evolutionary linkage appears to exist between these three proteins, nor between them and the F-B-R proteins. It is probable that the di-Mn’+-centre structure of bacterioferritin is very similar to that of the diferrous centre. It has been speculated that oxidation of Fe(I1) at the ferroxidase centre of bacterioferritin is accompanied by breakage of the Glu127 bridging bond, as anticipated for ribonucleotide reductase (Harrison and Arosio, 1996). The ferroxidase centre residues of the F-B-R proteins are very well conserved (see Fig. 6), and where this is not the case it is presumed that corresponding proteins lack ferroxidase activity (as for mammalian ferritin L subunits). However, although the ferroxidase centres of the F-B-R group are similar, they are not identical (Fig. 9). The residues equivalent to Glu27, Glu62 and Glu107 in human H-chain ferritin are also used by bacterioferritin, FtnA and and rubrerythrin (Fig. 9). However, other dinuclear metal site ligands are not consistently used. The H-chain residue Gin141 (possibly liganded to iron through water) is conserved in FtnA, but is replaced by the bidentate bridging residue, glutamate, in bacterioferritin and rubrerythrin. The iron site A ligand, His65, of H-chain ferritin is maintained in the other ferritins but in rubrerythrin it is not used although it is conserved. Instead, Glu97 from helix C (rather than helix B) is used. The residues equivalent to Glu97 in the other proteins are hydrophobic and therefore cannot act as iron ligands. The iron site B ligand, Glu61, of H-chain ferritin is not used by any of the other proteins. In place of this B-helix residue, FtnA, bacterioferritin and rubrerythrin use residues from equivalent positions in the Dhelix: Glu130, His130 or Hisl31, respectively. In H-chain ferritin, the equivalent residue in the D helix is Ala144 which is clearly unable to act as an iron ligdnd. In FtnA, bacterioferritin and rubrerythrin, the residue equivalent to Glu61 of H-chain ferritin is a glutamate, an aspartate and a glutamate, respectively - all potential iron ligands. However, in the rubrerythrins this residue is poorly conserved (Fig. 6) and in FtnA it acts as an iron site C ligand (Section 4.2). Tyr34 of H-chain ferritin is conserved in all the F-B-R proteins, where it is hydrogen bonded to Glu94 (H-chain numbering). This residue has been implicated in the formation of a transient tyrosylradical (or ferric-tyrosinate) species during the iron oxidation/storage reaction of human H-chain ferritin (Harrison and Arosio, 1996). The differences in the dinuclear sites of the F-B-R proteins are likely to result in functional

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r 0

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contrasts, but the way in which these differences influence ferroxidase activity and core formation are not understood as yet. The model of the dinuclear metal site region of N . gonorrhoeae BfrB is shown as a representative of the ‘a-subunit group’ of bacterioferritins (Fig. 7). N . gonorrhoeae BfrB shows three differences when compared with the same region of E. cofi bacterioferritin (Fig. 9). These differences are likely to result in a nonfunctional ferroxidase centre, although it may bind metal. The model of the FtnB dinuclear metal site region reveals five differences when compared with the equivalent region of FtnA. These differences indicate that FtnB is unlikely to possess ferroxidase activity, but the residues at the ferroxidase-centre region are all potential iron ligands indicating that it may bind metal.

5. CORE FORMATION AND THE IRON CORE 5.1. Iron Uptake 5.1.1.

Bacterioferritin

Ferritins and bacterioferritins acquire iron in the reduced, ferrous form and store it in the oxidized, ferric form. A . vinefandii and P . aeruginosa bacterioferritins have been shown to exhibit sigmoidal iron-uptake kinetics with progress rates dependent on initial iron contents, as displayed by mammalian ferritins (Mann et al., 1987). The rates of bacterioferritin iron uptake were similar to those of horse-spleen ferritin (Mann et a f . , 1987). For E. cofi bacterioferritin. iron uptake consists of at least three kinetically distinguishable phases corresponding to an initial rapid binding (phase 1; ti -50 ms) and oxidation (phase 2; ti -5 sec) of two ferrous ions at each ferroxidase

Figure 9 Schematic diagrams of the dinuclear metal sites of rubrerythrin, methane monooxygenase, haemerythrin, bacterioferritin, FtnA and human H-chain ferritin, derived from their crystal stuctures (DeMare el al., 1996; Norlund er al., 1990; Norlund and Ecklund, 1993; Rosenzweig ef al., 1993; Frolow el a/., 1994; Hempstead et a/., 1994; Holmes and Stenkamp, 1991; Lawson et a/., 1991). Also shown are the equivalent regions of N . gonorrhoeae BfrB ( N . g o BfrB, model), E. coli FtnB (model) and the L-subunit of horse-spleen ferritin (Lawson, 1990). The defined metal sites A and B are indicated as shaded spheres. The model of N. gonorrhoeae BfrB is used to represent bacterioferritin group. FtnB and N. go BfrB may the ‘ferroxidase-centre-free/a-subunit’ not possess functional dinuclear-metal binding sites or ferroxidase centres (the corresponding positions of the dinuclear metal species observed in FtnA or E. coli bacterioferritin are indicated as open spheres). Adapted from Harrison and Arosio (1996). and from Harrison ei a/. (1998).

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centre followed by slower oxidation and iron core formation (phase 3; ti -6 min) (Le Brun et al., 1993b; 1995). The first 48 iron atoms added per apobacterioferritin 24-mer are oxidized rapidly at the ferroxidase centre (phases 1 and 2), but any additional ferrous iron is slowly oxidized (phase 3). The first phase was monitored by measuring a transient and very rapid blue shift in the haem spectrum which occurs upon binding of ferrous iron at the ferroxidase centre (which is adjacent to the haem site). Phase 1 was saturated by the addition of 50 or more ferrous ions per holomer. Phases 2 and 3 were monitored by measuring the increase in absorbance at 340 nm arising as the result of iron oxidation, and phase 3 was seen only when more than 50 ferrous iron atoms were added per holomer. Other metals can bind at the ferroxidase centre (Co”, Mn2+, Zn2+; binding affinities are 10-4-10-7) and act as iron uptake inhibitors (Frolow et al., 1994; Le Brun et al., 1995; Keech et ul., 1997; unpublished work in Thomson et ul., 1997). EPR studies indicate that a short-lived di-iron species is formed (probably at the ferroxidase centre) during iron uptake that is rapidly converted to a mononuclear, magnetically isolated high-spin species which may be at a site other than the ferroxidase centre (Le Brun et al., 1993a,b, 1995). A similar monomeric high-spin Fe” species has been seen in the bacterioferritins of A . vinelutzdii, P . arrzrginosa and E. coli following their purification (Cheesman et al., 1992), indicating that this species is stable and common among the bacterioferritins. A stable iron species absorbing maximally at 475 nm (and very weakly at 425 nm) has been observed by visible spectroscopy in E. coli bacterioferritin (Andrews et ul., 1995). Whether this species corresponds to any of those seen by EPR is uncertain. Substitution of residues acting as metal ligands at the dinuclear site resulted in loss of both ferrous oxidation and iron core formation capability (Le Brun et al., 1995). confirming the identity of this site as the ‘ferroxidase centre’. The binding of metal ions at the ferroxidase centre is accompanied by changes in protonation that compensate for alterations in charge (Le Brun et al., 1996). Furthermore, cobalt binding studies indicate that Co2+ binds cooperatively at the ferroxidase centre (Keech e / al., 1997), i.e. once one metal binding site is occupied at the ferroxidase centre, the remaining site is more readily occupied. Between 25 (pH 5.5) and 150 (pH 9.0) ferrous ions bind anaerobically to A . vinelutrdii bacterioferritin, and the number of protons released per ferrous ion bound rises with increasing pH (two protons released per ferrous ion at pH 9). A relatively high amount of ferrous iron binding to holobacterioferritin was attributed to its high phosphate content, and a role for the ironcore-phosphate in mediating electron transfer at the core surface was suggested (Watt et al., 1992). Mossbauer spectroscopic analysis of the initial oxidation of iron by E. coli bacterioferritin has shown that the reaction pathway involves the rapid conversion of Fe(I1) into Fe(II1) dimers, then the dissociation of dimers into

IRON STORAGE IN BACTERIA

325

monomers, followed by the formation of iron clusters (Hawkins et a/., 1996). This reaction pathway is consistent with that determined by EPR spectroscopic studies described above. The Mossbauer parameters of the ferric dimer species differed from those of the dimer associated with human ferritin, which is consistent with the differences in amino acid residues at the respective ferroxidase centres. A new component was observed, different from any species so far identified in ferritin, that probably represents a small cluster of Fe(II1) atoms. Oxidants other than molecular oxygen have been shown to allow iron core formation in bacterioferritin and mammalian ferritin (Watt et a/., 1988). The alternative oxidants were Fe(CN);- and proteins (cytochrome c, stellacyanin and plastocyanin) too large to penetrate the protein shell. In horse-spleen ferritin, the initial protein-catalysed Fe” oxidation step is specific for 0 2 whereas the subsequent oxidation on the core surface is non-specific for oxidant (Treffry el ul., 1979). It has also been shown that the iron core of bacterioferritin and mammalian ferritin can be reduced by large proteins (flavoprotein and ferredoxins). These findings indicate that electrons can pass through the protein shells of iron-storage proteins via long-range electron transfer processes (Watt ct a/., 1988).

5.1.2.

Ferririn

A Mossbauer analysis of iron core formation in E. coli ferritin revealed the presence of three types of small iron cluster: magnetic ferrihydrite clusters; non-magnetic ferrihydrite clusters; and novel non-magnetic non-ferrihydrite clusters (Bauminger et a/., 1994). Studies on E. coli ferritin and corresponding site-directed variants using a sequential stopped-flow assay employing 1,lO-phenanthroline to bind free Fe” indicates that the first 48 Fez+ atoms to enter the molecule are bound and oxidized at the ferroxidase centres. The rate of Fe” oxidation was maximal when 48 Fe2+ atoms were added per molecule, consistent with a two-electron oxidation of a pair of Fe” atoms in the ferroxidase centre. Substitution of iron-site A ligands was found to eliminate fast Fe” binding and oxidation, whereas substitution of ironsite B ligands resulted in retention of fast Fe” binding but loss of rapid oxidation. It was suggested that the differences in the effects of the site A and B modifications could mean that dioxygen is initially bound to the Fe” at site B (Treffry PI d., 1997). A comparison of rates of ferrous iron oxidation and core formation revealed that FtnA is about 4-fold more active than bacterioferritin (Hudson et al., 1993). This presumably relates to differences at and around the respective ferroxidase centres. The reaction mechanisms of the two proteins appear similar, both involving the rapid binding and oxidation of two

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ferrous irons per ferroxidase centre generating a di-iron species that converts to mononuclear ferric species. This step is followed by a slower oxidation and core formation reaction. Unfortunately, a detailed direct comparison of the reaction pathways of bacterioferritin and bacterial ferritin has not been reported so it is not possible to determine how the proteins differ in this respect. The route taken by ferrous iron entering the ferroxidase centre is unknown. In human H-chain ferritin, a ‘one-fold channel’ is present that could allow iron to pass from the outside of the molecule through a cavity between the A and C helices (Lawson et al., 1991). A similar ‘cleft’ has been observed in the Desulphovibrio gigas rubrerythrin structure (deMare P I d . , 1996). Also, channels at the three- and four-fold axes have been described for eukaryotic ferritins (Harrison and Arosio, 1996). In E. coli bacterioferritin and FtnA, the residues surrounding the channel regions are different from those in mammalian ferritins and this possibly reflects differences in channel function (Harrison and Arosio, 1996; Harrison et d . , 1998). Sitedirected alteration (D118A) of a potential iron-binding residue in the threefold channel region of E. coli bacterioferritin had little effect on the iron core formation process, suggesting that in bacterioferritin the ‘three-fold channel’ has no important role in iron uptake and oxidation (Le Brun et al., 1995). Currently, the route(s) taken by iron entering and exiting the bacterial ferritins and bacterioferritins are uncertain. 5.2. The Iron Core

Animal ferritins tend to contain ordered cores with crystalline domains having diffraction properties like those of ferrihydrite, which has the composition 5Fe203.9H20(Towe and Bradley, 1967; Fischbach et al., 1971). Although animal ferritins contain variable amounts of inorganic phosphate, bacterioferritin and bacterial ferritin cores are far richer in phosphate with Fe:Pi ratios of between 1 and 2 (Moore et al.. 1986; Watt et al., 1986; Frazier et al., 1993; Table 4). The phosphate is apparently distributed throughout the iron cores of bacterioferritns and animal ferritins (Rohrer et al., 1990). The high phosphate contents of bacterioferritin cores result in structural differences between the cores of mammalian ferritins and bacterioferritins (Stiefel and Watt, 1979; Bauminger et al., 1980; Ford et al., 1984; Mansour et al., 1985; Mann et al., 1986, 1987; Moore et al., 1986; St Pierre et al., 1986, 1989). However, these differences in phosphate content are likely to arise mainly from differences in the cytosolic environments in which the cores were formed, rather than any fundamental protein structure-function contrasts. High-resolution electron microscopy and electron diffraction reveal that the phosphate-rich cores of bacterioferritin are disordered, whereas those of mammalian ferritins are ordered (Mann et al.,

Table 4 Bacterioferritin native core properties.

Species A . vinelandii P . aeruginosa E. coli Horse spleen (ferritin)

Ordering temperature ( T , )

Blocking temperature ( T , )

> 20

-18 23

3 -3 3

ND m

&SO

Crystallinity

Phosphate:iron

Iron atomsiholomer

Amorphous Amorphous ND Good

1: 1.5-1.9 1:1.41.7 1:2.2 1:8

1000 900

ND, not determined. Data are from Bauminger ef al. (1980). and Treffry e f al. (1987).

-980 1 100 or 2000

328

SIMON C. ANDREWS

1986). However, bacterioferritin cores reconstituted in the absence of phosphate are ordered and ferrihydrite-like in structure (Mann et d., 1987). The magnetic ordering of the cores of bacterioferritins differs from those of animal ferritins, and indeed the core of A . vineluridii bacterioferritin differs from those of P . Lierugiiiosci and E. coli bacterioferritin (Bauminger et d . , 1980; Watt et al., 1986; St Pierre et al., 1986). The iron cores of the P . uerzcginosu and E . coli bacterioferritins generate magnetically split sextet Mossbauer spectra a t < 1.3 K , but at higher temperature the magnetic splitting collapses to a quadrupole split doublet. This behaviour is interpreted as characteristic of magnetically ordered cores at 1.3 K, that undergo a transition to a paramagnetic state at higher temperature (St Pierre et d., 1986), and therefore, in contrast to mammalian ferritin cores, the cores are not superparamagnetic. The lack of superparamagnetism for these bacterioferritins is thought to be due to the disorder of their phosphate-rich iron cores. In contrast, the iron cores of A . vincdmilii bacterioferritin, despite being phosphate-rich and apparently disordered, are superparamagnetic (Watt c't ul., 1986; Mann et d.,1987). The reason for this discrepancy is unclear. Reconstitution of the iron core of bacterioferritins in the absence of inorganic phosphate results in the formation of superparamagnetic, crystaline ferrihydrite-like cores similar to those of mammalian ferritins (Mann et al., 1987; Andrews rt ul., 1993). This demonstrates that differences in native core structures are mainly due to the in vivo environmental conditions under which the cores are formed. Presumably, bacteria contain higher levels of free inorganic phosphate in their cytosols than d o mammalian cells. The blocking temperature of reconstituted A . vinelunclii bacterioferritin (22.2 K ) cores is lower than those of reconstituted E. coli bacterioferritin (39 K ) and horse-spleen ferritin (38 K ) (Bell et d . ,1984; Mann et ul., 1987; Andrews ('I id., 1993), probably because the reconstituted A . vinelunriii bacterioferritin cores are smaller than the native cores (Mann et al., 1987). The EPR spectra of horse-spleen ferritin contain a wide signal at g z 4.0 between 5 and 100 K, which converts to a narrower g = 2.0 signal at higher temperature (Weir rt d . , 1984). No core signal is seen at very low temperatures since the core becomes EPR silent. The EPR spectra of bacterioferritin cores are different. At very low temperatures (-3 K ) no signal is seen, as for horse-spleen ferritin, probably because the core becomes ordered. At higher temperatures a broad signal is generated that narrows with increasing temperature but remains centred at g x 2.0. not 4.0. This suggests a lower magnetic anisotropy for the bacterioferritin cores than for those of horse-spleen ferritin, as indicated by Mossbauer spectroscopy (Le Brun, 1993). The redox potential of the iron core of A . vinelundii bacterioferritin is -420 mV (pH 7-9), and rises to -390 mV at pH 6 (Watt et ul., 1986), whereas that of horse-spleen ferritin varies from -190 to -416 mV over

IRON STORAGE IN BACTERIA

329

the pH range 7-9 (Watt et al., 1985). These differences in redox potential and responses to pH are likely to reflect differences in core composition (crystallinity and phosphate content), although this has not been tested using reconstituted iron cores. Whole-cell Mossbauer spectroscopic studies on P . aeruginosa grown anaerobically in rich broth with high or low amounts of nitrate revealed that the predominant form of cellular iron differs according to the growth condition employed (Reid et al., 1990). Surprisingly, despite the lack of oxygen, no ferrous iron was observed. Under conditions of high nitrate a single quadrupole-split doublet iron species was seen, resembling that observed in purified P. aeruginosa bacterioferritin. Under conditions of low nitrate an additional iron species was observed yielding a sextet spectrum like that of A . vinelandii bacterioferritin. These differences are probably due to differences in the nature of the iron cores formed under the different growth conditions. Bacterioferritin isolated from anaerobically grown P. aeruginosa contains an iron core of -670 iron atoms (Moore et al., 1986). The possibility that the P.ueruginosa bacterioferritin iron core is predominantly ferrous iron in vivo and is oxidized during purification was tested by Kadir et al. (1991). Their studies suggested that the iron core of bacterioferritin is not altered on isolation, is typically 85% oxidized within the cell and may contain small iron clusters during the early post-exponential phase. The nature of the oxidant used anaerobically in the formation of ferric-iron cores is unknown.

6. BACTERIOFERRITIN-ASSOCIATED FERREDOXIN

The open-reading frame (originally called gen-64, now redesignated b f i Garg et al., 1996) located 72 bp upstream of the bfr gene was originally of unknown function (Andrews et al., 1989a). However, the gene is now known to encode a protein having 64 amino acid residues (Bfd, bacterioferritin-associated ferredoxin) related to the -60-residue [2Fe-2S] cluster-containing domains of the NifU proteins (required for nitrogenase metallocluster assembly), the nitrite reductases, and the nitrate reductase of Klebsic4la pneurnoniae (Figs 10 and 1 1 ; Fu et al., 1994; Ouzounis et al., 1994). The members of this family all contain the following motif, -C-X(C/H)-X31-34-C-Xl-C-,which is thought to provide a binding site for the [2Fe-2S] centre (Fu et al., 1994). Mvcohucterium leprae (Pessolani et ul., 1994), P . aeruginosa (Fig. 10) and A . vinelandii (Garg et al., 1996) possess homologues of the hfd gene which, as for E. coli, are adjacent to hfr genes. This finding strongly suggests that hfd and bjr are functionally related (the bjd homologue of Svnechocystis is not near either of the corresponding hfr

B E5C

CO

MYC

le

Bfd NWC1CNG.I S5KKIRQAVR CFSPASFQQL K...KF:?VG NQCGKCVRAA R E W D E L M Q L %WCLCAG.A TNQTVCDW. ARGATTSKEI A...AACGAG SDCGRCRRTS RAIIAASNPT V 1 MYICVCRG.1 SDKQIEAAA. QWITSLEEL S...ESMGVG AZWGVCQGHA CQVLEAIARR K 1 1

SYIleCO

NIFT And

5p

And

Vd

A20 ch A20

vi

Kle pn P l e bo N o s co

He1 p y dzo br Unknowr.

142 141 134 134 134 52 53 158 136 140

ALVC3CFG.V SENKVRRIVI ALVCRCFG-I SESKVRRVIR QL1CKSFA.I DEVMVRDTIR AL1CKCFA.V DEVMVRDTIR KL1CKCFG.V DEGHIRRAVQ ALICSCFG.1 SEPKIRRWI ALVCSCFG.1 SESKIRRVIL 1IVCECAR.V SLGTIKEVIK ELVCKCPG.1 DAAMIEMVT AL1CKCFG.V TDARiRRVII

ENDLTDAEQV ENNLTTAEQV ANKLSTVEDV ANKLSTVEDV NNGLTTW ENGLTTVEQV ENHLTDAEQV LNDLKSVEEI VNSLTTLEEV EKJLTTAEQV

T...NYIKAG GGCGSCIAKi T...SYIKAG GGCGSCLADI T...KHTKRG GGCSACHEGI T...NYTKAG GGCSACHFAI I...NYTKAG GGCTSCHEKI T...SWKAG GGCGSCLADI T N?M(AG GGCGSCLANI ?...NYT:KAG AFCKSCVRPG T...HYTKAG GSCQTCHEKI T...NYVKAG GGCSSCLSDI

DDIIWVKEN DDLISAVIKE ERVLSEELAP ERVLTEELAA ELALAEILAQ EDIITAVIDE DDII3SVOOE GHEKRDYYLV EEVLEAVLAK DDILADITQE

K S

V R

Q K

Y D T K

Nitrate reduct.sc A

PSe fl

805 RI1CSCFS.V GERAIGEAIA GGCRSPGGRL G...GKLKCG TNCGSCITEL K A L W A Q A 187 KILCNCiKN.V SESAVCAGIG RGL3LDGLKQ P.....LDCG TQCGSCVPEI KRL;ASTSLP I

Bra j a

488 RN1CVCNR.V DLGTZEDAIS VHGLRSVAAV R...EHTNAA GGC..CQGRI EDWLYSEPSD M

K l e pn

NifE pro& Ifan

p0

BdC

SU

ESC

Co

K l e pn Erne nl N e u cr

Bac

BU

E 5 c CO K l e pn

ni N e u cr COI.mnmU.

Nitrite reductaru TQICSCHN.1 TKGKLVEAV. KNGCSSLADV K...KCTKAG TACGGCEPTV EI1CGCNG.V SKGAIIQAIQ EKGCSSTDEI K...ACTGAS RSCGGCKPSV AQICSCFD.V TKGDLIAAI. NKACHTVAAL K...AETKAG TGCGGCIPLV AQ1CSC:XX.V SKGDICQAV. SGGAGDMAAI K...SRTKAA TGCGGCSALV TQ1CSCHN.V TKGDVVESVK SGTCKTIADV K...SCTKAG TGCGGCMPLV TQ1CSCHN.V TKADLVAPLK SGECTSLGDL K...SCTKAG TGCGGCMPLV EA1CGCTT.L SRDEVVEEIK AKGLSHTXEV MN.VLGWKTP EGCSKCRPAL NNLCEHFA.Y SRQELFHLIR VEGIKTPEEL L...AKHGKG YGCEVCKPTV B KD1CEHFP.W SRQEIYHLVR VNHIRTPEQL I...ARYGQG HGCEVCKPLV B 559 NNLCVE1P.Y SRADLYNVIA IRQLRTFDDV MKSAGKCPDS LGCEICKPAI B 555 "LCPHFPEY SRADLYNIIS VKRLRTLPDV MREAGADADS LGCEACKPAI

A A A A A B B

458 415 422 420 497 493 479 483 481

:

cc

KIIFEKELKK EEILQHTLGS TQVLNAELAK KQVHEYQLAE QSIFNKTMLD TSIFNRTMSA NYYbZMINPT GSLLASCWNE ASVLASCWNE ASILSSLFNP ASIFASLWND

L D Q

Q M L

K Y

Y H H

c c

Figure 10 The [2Fe-2S] domain of the NifU family. (A) Schematic representation of the domain organization of members of the NifU family. (B) Multiple amino-acid-sequence alignment of the [2Fe-2S] domains of members of the NifU family. The conserved Cys (or His) residues, thought to act as iron-sulphur centre ligands, are in bold. Residue numbering is on the left. Sequences and their corresponding database accession numbers are as follows. Bfd: Esc co (Escherichia coli). P13655; Myc le (Mycobacterium leprae), L01095; and Syneco (Synechocystis PCC6803), P73484. NifU: Ana sp (Anabaena sp. PCC 7120), P20628; Ana va (Anabaena variabilis), Q44483; Azo ch (Azotobacter chroococcum), P23 121 ; Azo vi ( A . vinelandii), Q44540; Kle pn (Klebsiella pneumoniae), P05343; Ple bo (Plectoneme boryanum), 400241; Nos co (Nostor commune), P26247; Helpy (Helicobacter pylori). G23133 12; Azo br (A:ospirillum brasilense). 443895; and 'Unknown', 009257. Nitrate reductase A: Kle pn (K. pneumoniae). 406457; and Pse f l (Pseudomonasfluorescens nitrate reductase like protein). 45171 5. N i E : Bra ja (Bradvrhizobium japonicum). P26506. Nitrite reductase A or B domain: Han PO (Hansenulapolymorpha). 400943; Bar su (Bacillus subtillis), P42435; Esc co (E. coli), PO8201; Kle pn (K. pneumoniae), 406458; Eme ni (Emericella nidulans). P22944; and Neu cr (Neurospora crassa), P38681. Adapted from Ouzounis et al. (1994).

332

SIMON

C. ANDREWS

NIFU

Bfd

Bra j a

I

I

/

Ilelpy

\

Psejl

NR -A

~

BacsuR

1 Erne ni

SCALE - = Distance of 0.1

Figure I I Unrooted phylogenetic tree showing the evolutionary relationship between members of the Bfd-NifLJ-Nitrite reductase family. Details are as for Fig. 7 except the sequences in Fig. 10 were used to produce the tree. Four subfamilies are indicated: Bfd, NifU, Nitrite Reductase [2Fe-2S]-domains A or B.

genes; Cyanobase reference ~ ~ 1 2 2 5 0The ) . Bfd protein of E. c d i has been overproduced, purified, and characterized and found to be a positively charged monomer containing two iron atoms and two labile sulphides. The results show that Bfd is a ferredoxin-like protein possessing a [2Fe2S] cluster with properties that are remarkably similar to those of the iron-sulphur-containing domain of A . vinelandii NifU. Spectroscopic and redox studies revealed the presence of a [2Fe-2S]2+3Scentre (E! = -254 mV) closely resembling that of the NifU protein (Garg c't (11.. 1996; Quail r / ( I / . .

IRON STORAGE IN BACTERIA

333

1996). Bfd was shown to associate specifically with bacterioferritin, suggesting that the two proteins may interact within the cell. Regulatory studies have shown that hfd is repressed by the Fe'+-Fur complex and preferentially expressed post-exponentially under iron-starvation conditions (see 1996). Recently, hfil-hfr and hfd null mutants have been proQuail et d.. 1997). It duced and their phenotypes are under investigation (Tehrani et d., is speculated that Bfd is a 2Fe-ferredoxin participating either in release/ delivery of iron from/to bacterioferritin (or other iron complexes such as ferri-siderophores or iron-sulphur clusters).

7. INTRACELLULAR IRON METABOLISM 7.1. Regulation and Physiological Roles

7.1.1. Canipylobacter jejuni The ferritin gene ( c : / i ) of the microaerophilic Cirinpjdohucicr ,jejiini encodes a typical bacterial ferritin highly similar in sequence to the haem-free ferritins of other bacteria. The function of the ferritin of C. ,jejiini has been investigated by constructing a cfi mutant and studying its phenotype (Wai et NI., 1996). These studies revealed that growth of the ferritin mutant is inhibited under conditions of iron deprivation. Also, the mutant was more sensitive to the redox-stress inducing agents paraquat and H102. Only a slight growth defect was observed under standard growth conditions. The iron-starvation and redox-stress growth defects of the cfi mutant were mostly or fully complemented with a cft-encoding plasmid. This study strongly suggests that the C. jejritii ferritin participates in enhancing growth under iron-starvation conditions and in defending against redox stress. and it is likely that these effects are achieved by the sequestration of iron by the ferritin molecule.

7.1.2. Escherichia coli The genes (finA and h j i ) , encoding the two characterized iron-storage proteins of E. coli, FtnA and bacterioferritin, have been inactivated on the chromosome (Andrews el ul., 1997; S.C. Andrews. A.J. Hudson, A.R. Timms. C. Hawkins. J . M . Williams, P.M. Harrison and J.R. Guest, unpublished) generating isogenic single and double mutants. After growth to stationary phase in rich medium, the iron contents of the / i n A and f i n A hfi. mutants were half those of the hfr mutant and parental strain

334

SIMON C. ANDREWS

(W31 lo). The iron deficiencies of the , f h A and j f n A hfi mutants were confirmed by Mossbauer spectroscopy, which further showed that the low iron contents are due to a lack of magnetically ordered ferric iron clusters likely to correspond to FtnA iron cores. N o differences in iron contents were observed following growth under iron-deficient conditions. Growth tests in rich and minimal media, under aerobic and anaerobic conditions, under redox-stress conditions and in the presence of high and low concentrations of iron, failed to reveal any growth defects associated with the mutations. However, growth of the ,ftnA and JtnA hfi. mutants was found to be impaired in iron-deficient minimal medium. This phenotype was only observed when precultures were grown under iron-sufficient conditions indicating that during growth under iron deficient conditions, E . c d i can utilize pre-deposited FtnA-contained iron stores to enhance growth. No redox-stress sensitivity was observed for the iron-storage mutants (Keyer and Inilay, 1996; Andrews et a/., 1997), which contrasts with the findings made with ferritin mutants of C. ,jKjuni (see Section 7.2.1). In summary, it appears that FtnA enables E. c d i to double its iron content in stationary phase during growth in rich medium and provides an intracellular source of iron that can be drawn upon during growth under iron-restricted conditions. The role of Bfr remains unclear. I t has recently been suggested that it participates in Lac permease activity but there is no direct evidence for this (Yariv, 1996). Although the ,ftnA gene appears to have no redox-protective role under normal circumstances, Touati et a/. (1995) have shown that a multicopy plasmid carrying the E . colijinA gene results in the protection offur mutants against ironinduced redox stress. However, multicopy bfr had no protective effect. This result was interpreted as indicating that overproduction of FtnA (but not bacterioferritin) can reduce intracellular free-iron, presumably through its capacity to sequester iron. I t is not known why the hfr gene could not function in a similar way. Overexpression of the hfi and ,finA (and the human H-chain ferritin and the H . pylori pfr genes) genes in E. coli was found to result in the induction of the Fur-regulated JiuF gene, presumably due to sequestration of intracellular iron (Stojiljkovic r t ( I / . , 1994). This finding supports the proposal that FtnA may exert a redoxstress protective effect in j r r mutants through its iron-sequestering activity but fails to explain the inability of bacterioferritin to function similarly. Factors influencing expression of the bl; andJinA genes have been investigated using /rcZ fusions and by Western blotting with appropriate antisera (Andrews et uf., 1997). Results showed that bfr andJtnA are induced by the Fe2+-Fur complex and by post-exponential growth. A stationary-phase induction of bfr expression was suggested by previous studies on the abundance of overproduced bacterioferritin in a transformant carrying a multicopy hfr plasmid (Andrews et a / . , 1993). Post-exponential expression of the

IRON STORAGE IN BACTERIA

335

bfr gene was dependent on the alternative sigma factor, os (Andrews et al., 1997), which is responsible for the stress-induced expression of at least 30 E. coli genes. In contrast, ftnA expression was os-independent and was strongly induced by RNase 111, as previously shown (Izuhara et al., 1991). Neither the bfr nor the ftnA genes were found to possess sequences resembling Fur-binding sites, and the hfr gene was found not to interact with the Fe-Fur complex in a Fur-titration assay (Stojiljkovic et al., 1994). Therefore, the putative Fur-binding site identified in the upstream region of the hfr gene (Andrews et al., 1989a) is unlikely to be functional.

7.1.3. Synechocystis PCC 6803 The abundance of the bacterioferritin of Synechocystis PCC 6803 was not affected by the iron concentration of the medium (between 3 and 300 p~ iron citrate), suggesting that expression of the corresponding gene is not influenced by iron in the concentration range employed (Laulhere et al., 1992). However, cellular contents of soluble iron were found to increase by 250-fold when cells were grown with 300 PM iron citrate, rather than 3 PM. Autoradiography of native polyacrylamide gels containing 59Fe-labelled soluble-cell extracts revealed four major iron-containing components: 'cellular debris'; bacterioferritin; low-molecular mass iron proteins; and 'low molecular mass molecules' ( < 10 kDa). The bacterioferritin and low molecular mass molecules (LMMM) were the major iron components. Preculturing S y n e c k y s t i s in iron-poor medium (3 PM iron citrate), followed by transfer to fresh medium with 1.7 p~ 59Fe citrate, resulted in high "Fe incorporation into bacterioferritin and the LMMM. In contrast, preculturing in iron-rich (300 LLM iron citrate) medium resulted in virtually no subsequent incorporation of 59Feinto bacterioferritin (- 10-fold less than when precultured in 300 p~ iron citrate), and an -2-fold reduced incorporation into LMMM. These experiments suggest that bacterioferritin incorporates iron when iron concentrations in the medium are low, but not when iron levels are high. A redistribution of "Fe from bacterioferritin to the LMMM was observed under low iron growth conditions, indicating that bacterioferritin can release its stored iron to other cellular components. The nature of the LMMM is uncertain (see Section 7.2) but they appear to be important contributors to intracellular iron homeostasis.

7.1.4. Other bacteria The bacterioferritin of R. capsulatus is produced during aerobic and photosynthetic growth, and is located mainly in the cytoplasmic fraction

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(Ringeling el ul., 1994). The levels of bacterioferritin were relatively constant with respect to the growth phase during chemoheterotrophic growth in batch culture. However, levels of bacterioferritin were reduced following growth in iron-deficient medium, suggesting regulation of bacterioferritin synthesis in response to iron availability, as for bacterioferritin and FtnA of E. coli (Section 7.1.2). The low-temperature Mossbauer spectrum of R. cupsulutiis bacterioferritin indicated the presence of an amorphous core. Mossbauer spectroscopy on 57Fe-labelled whole cells grown aerobically or photosynthetically (anaerobically) revealed the presence of ferric iron under both conditions (Ringeling r t d., 1994). Aerobically, three forms of iron were distinguished: ferritin-like Fe(II1) (85%); non-magnetically ordered Fe(II1) (10Y0); and Fe(I1) ( 5 % ) . The predominant 'ferritin-like' iron comprised two species, one (60% of the detected iron) with parameters resembling those of R. cupsulritus bacterioferritin and the other (25% iron) resembling A . vinelandii bacterioferritin and mollusc ferritins. In photosynthetically grown bacteria, bacterioferritin-like iron was abundant (4On/o of iron). This correlated with the observation of bacterioferritin protein in photosynthetically grown R. c.upsulutu.s. Of the remaining iron, 30% was non-magnetically ordered Fe(II1) and 30% was Fe(I1). The observation of bacterioferritin-like cores within anaerobically grown cells suggests that core formation, irr viiw, may not require exogenous molecular oxygen (as for P. creruginoscr; Section 5 . 2 ) . I t was suggested that the photosynthetic reaction centre of photosynthetically grown R . cup.sulatu.s may be responsible for oxidizing ferrous iron during the iron core formation (Ringeling c t ul., 1994). It should be stressed however, that the identity of the 'bacterioferritin-like iron' in photosynthetically grown bacteria was not determined and so its location (in bacterioferritin, ferritin or elsewhere) remains to be established. The intracellular human pathogen, Lrgionellrr pneuniophilo, has a relatively high requirement for iron and because of this the iron-containing proteins of the bacterium were investigated (Mengaud and Horwitz, 1993). Iron-containing soluble proteins were detected in "Fe-labelled whole-cell extracts by autoradiography and native PAGE. Seven iron proteins were detected. One was identified as Fe-superoxide dismutase and the major iron-containing protein was identified as aconitase. Two weakly-labelled, low-mobility iron-proteins (probably corresponding to ferritin and/or bacterioferritin) were observed having mobilities similar to that of a major Feprotein (probably bacterioferritin) from E. coli. Radioactive iron-labelling studies have also revealed a potential bacterioferritiniferritin in Y~~rsiniu peslis having an estimated native molecular mass of -620 kDa (Perry lit ul., 1993).

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7.2. The Low-molecular Weight Iron Pool

Little is known regarding the precise distribution of cellular iron. The subcellular distribution of iron in E. coli was determined by Bohnke and Matzanke ( 1 995) following growth in iron-restricted minimal medium containing Fe-ferricrocin: total iron was -0.006% of cell dry weight; the membrane fraction was 0.0013% iron; the cytoplasm was 0.0043% iron; and the periplasm was 0.00034% iron. Iron entering the cell may become incorporated into iron-containing enzymes and proteins, or into iron-storage proteins (ferritins and bacterioferritins). However, 'free' iron is also present within the cell forming a mobile, low-molecular weight (LMW) iron pool (Tangeras et al.. 1980; Crichton and Charloteaux-Wauters, 1987; Weaver and Pollack, 1989; Fontecave and Pierre, 1991; Keyer and Imlay, 1996). The chemical nature of this iron pool is controversial. In animals it has been suggested that pyrophosphate- or ATP- and GTP-bound iron are major contributors to the LMW-iron pool (Mansour et al., 1985; Nilsen and Romslo, 1985; Weaver and Pollack, 1989). Other candidates for LMWiron binding components include polypeptides, amino acids and some sugars (Gessa et al., 1983; Tonkovic et al., 1983; Bakkeren et al., 1985; Anderson and Porath, 1986). Williams (1982) predicted that the levels of free iron in the cell are likely to be in the range to IO-'M, and that free iron would be predominantly ferrous. Studies on the Fur protein (of E. coli and other Gram-negatives) indicate that it senses ferrous iron, suggesting that this form of iron must be freely available in the cell and that its concentration reflects cellular iron status. The affinity of Fur for ferrous iron is l O P 5 (Bagg ~ and Neilands, 1987), suggesting that free ferrous iron should be approximately at (or above) this concentration. Indeed, levels of free iron in E. coli have been estimated by EPR measurements of desferrioxaminebound cellular iron and found to be lo-' M (Keyer and Imlay, 1996). The ferrous form of iron is generally considered to be the metabolically active form of iron. This is because it is soluble and is the form utilized preferentially by a number of proteins (e.g. ribonucleotide reductase, ferrochelatase, ferritins, Fur). The nature of the LMW iron pool in E. coli has been probed using wholecell Mossbauer spectroscopy. This approach has shown the presence of two major components, a ferrous and a ferric species (Bauminger et al., 1979; Matzanke er al., 1989, 1992; Hudson et al., 1993), also seen in other microorganisms (see Bohnke and Matzanke, 1995). The proportions of the ferrous and ferric components depends on the whole-cell iron content; the proportion of ferrous iron increases as the cellular iron content decreases (Bauminger et al., 1979; Matzanke er al., 1989; Hudson et af.,1993). Ferrous iron levels of between 8 and 50% have been reported, depending on cellular iron levels (Matzanke et al., 1989, 1992; Hudson et al., 1993). It would thus

-

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appear that the absolute amount of cellular ferrous iron remains relatively constant but the ferric content fluctuates greatly in response to cellular iron levels. Hudson rt al. ( 1 993) detected two ferrous species (species B and C ) a s well as the ferric species (A). A temperature-dependent Mossbauer absorption study showed that ferrous species B is weakly bound (and could therefore be 'free' iron) whereas ferrous species C is strongly bound (and so could be protein associated) (Hudson el al., 1993). The levels of species C were low (3-5% of total iron) but were increased in an FtnA overproducing strain (9 25%), resulting in an increase in the total ferrous iron content from 8 -23% of total iron in the control strain, to 18-35% in the overproducer. These findings indicated that ferritin can influence the speciation of intracellular iron and that ferrous species C could be associated with FtnA. During sample storage at -20°C there was a redistribution of the ferrous iron from species C to species B. This was interpreted as evidence for the release of iron from FtnA to give free iron. Whole-cell Mossbauer studies by Matzanke cf al. (1989) revealed that the predominant iron species in E. coli had Mossbauer parameters unlike those of purified holobacterioferritin. The absence of a bacterioferritin-like iron species was presumably because the bacteria were grown under low iron conditions ( 2 4 P M "Fe ferricrocin), conditions that would not be expected to favour the deposition of iron stores. However, a bacterioferritin-like iron species was seen when very high concentrations of iron were used in the growth medium. This observation reflects the absence of iron in bacterioferritin isolated from non-iron supplemented medium (Yariv et al., 198 1 ) supporting the notion that bacterioferritin accumulates iron only under iron-sufficient conditions. It should be noted that in the above study, and other earlier studies, the presence of a ferritin in E. coli was unknown, and so the 'bacterioferritin-like' species could correspond to ferritin as well as, or instead of, bacterioferritin. A s5Fe-labelled component representing 70% of cellular "Fe was isolated from both the cytoplasmic and membrane fractions of "Fe-labelled E. coli grown aerobically in minimal medium under iron deficiency (Matzanke rt d., 1989). The major 55Fe-component eluted as a 155 kDa macromolecule upon gel filtration and was separated into I5 and 17 kDa 5SFe-labelled subunits during SDS-PAGE. The iron content of the 155 kDa protein was 13 atoms per subunit. These features are somewhat reminiscent of the Dps-like ferritin of L. innocurr (Section 3.6). Further studies by the same group resulted in the isolation of a LMW-ferrous species (Bohnke and Matzanke, 1995) of molecular mass 2.2 kDa and a PI of 1.05. The ferrous iron-containing compound had Mossbauer parameters like those of ferrous species observed in whole cells. It was composed mainly of phosphorylated sugar derivatives (pentose and/or uronic acid) binding -40% of cytoplasmic iron and it is speculated that it corresponds to the major component of the

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LMW iron pool. Whether or not the phosphorylated sugar compound has any important role in bacterial iron metabolism has yet to be determined, as has its relationship with the 155 kDa macromolecule.

7.3. lntracellular Ferric Reductases

Although it is fairly well established that bacteria contain ‘free’ ferrous iron in their cytosols, the mechanism employed to maintain iron in the reduced state, particularly under aerobic conditions, is not known. Also, it is unclear how ferric iron is reduced upon entry into the cell or during release from iron-storage proteins. Three cytosolic proteins possessing ferric reductase activity have been identified in E. coli (sulphite reductase, the haemoglobin-like protein and flavin reductase enzyme) and ferric reductases have been reported in other bacteria (Fontecave et al., 1994; Earhart, 1996). However, there is no good evidence that these enzymes have a direct role in iron metabolism and all would appear to have other cellular roles (see Fontecave et ul., 1994 for a recent review). It is possible that, since the cytosol is relatively reduced, the ferrous form is the preferred state for iron. The Fes protein of E. coli is the only protein known to function specifically in the reduction-mediated release of iron from an ironsiderophore complex (ferric-enterochelin) (Earhart, 1996). How iron is released from other ferri-siderophore complexes is unknown, although this is likely to involve reduction of the complexed iron.

8. PERSPECTIVE

There is still a great deal to be learnt regarding the processes of iron storage in bacteria. It is clear that bacteria have adopted different mechanisms to cope with their need for an iron store. Some bacteria contain one or two ferritins, others contain both a ferritin and a bacterioferritin, and some contain two bacterioferritins. In other cases, no iron-storage protein would appear to be present. The reason for these differences is not understood, but it is clear that a comprehension of the iron-storage systems of one bacterium will not provide an understanding of all. Much progress has been made in the 18 years since the existence of bacterial iron-storage proteins was first revealed by Stiefel and Watt (1 979). The structure-function properties of bacterial ferritins and bacterioferritins have been studied in some detail, although a complete understanding of their respective mechanisms and their differences has not yet been achieved. Physiological roles for the ferritins of E. coli and C . jejutii have been determined, although the details

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regarding the manner in which these proteins enhance growth, increase cellular iron content or protect against redox stress have yet to be fully addressed. Bacterioferritins remain more of an enigma. No phenotype has been discovered for E. cofi bacterioferritin-mutants. The role of the haem group and the bacterioferritin-associated ferredoxin have not been proven. The in vivo function of the a/&type bacterioferritins will be of great interest. So far, all those bacteria that appear to possess an cr/(3-bacterioferritin d o not apparently contain a ferritin. This implies that in these bacteria the role of ferritin in iron storage (growth enhancement, redox-stress resistance and increasing cellular iron content) is provided by the bacterioferritin. Whether this proves to be the case, and if so, how this is achieved, awaits further study. The possibility that other components (such as Dsp proteins, Bfd and phosphorylated sugars) have roles to play in intracellular iron homeostasis has been raised. Clearly, the functions of these components and any other as yet unrecognized components in iron metabolism must be fully elucidated in order to obtain a complete picture of iron storage/homeostasis processes in bacteria.

NOTE ADDED IN PROOF The recently determined I .6 A X-ray crystal structure of the Dps protein of E. coli reveals that it is structurally and evolutionarily related to F-B-R Family members. The Dps subunit has the same 4-a-helix motif as F-B-R proteins. The Dps holomer comprises 12 subunits formoinga spherical shell (90 A outer diameter) surrounding a hollow centre (45A diameter) likely to function as an iron-storage cavity. The ferroxidase centre residues are not conserved (Grant, R.A., Filman, D.J., Finkel, S.E.. Kolter, R. and Hogle, J.M. (1998) The crystal structure of Dps, a ferritin homolog that binds and protects DNA. N(iturr Structurul Biologji 5, 294-303).

ACKNOWLEDGEMENTS I thank the BBSRC for providing an Advanced Fellowship and the UKHuman Genome Mapping Project Resource Centre and Seqnet for provision of computing facilities. I would also like to thank Drs P. Artymiuk and V. Barynin for assisting with the production of molecular graphics images, and Professor P.M. Harrison for useful discussion.

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Tonkovic, M., Hadzija, 0. and Nagy-Czako, 1. (1983) Preparation and properties of Fe(1ltsugar complexes. Inorg. Chim. Acto 80, 25 1-254. Touati, D., Jacques, M., Tardat, B., Bouchard, L. and Despied. S. (1995) Lethal oxidative damage and mutagenesis are generated by iron in Deltafur mutants of Escherichiu coli: protective role of superoxide dismutase. J . Bacterial. 177, 2305-23 14. Towe, K.M. and Bradley, W.F. (1967) Mineralogical constitution of colloidal 'hydrous ferric oxides'. J . Colloid Intrrfuce Sci. 24, 384-392. Treffry, A,, Sowerby, J.M. and Harrison, P.M. (1919) Oxidant specificity in ferritin formation. FEBS Lett. 100, 33-36. Treffry, A,, Harrison, P.M., Cleton, M.I., De Bruijn, W.C. and Mann, S. (1987) A note on the composition and properties of ferritin iron cores. J . Inorg. Biochrm. 31, 1L6. Treffry. A,, Zhao, Z., Quail, M.A., Guest, J.R. and Harrison, P.M. (1997) Dinuclear center of ferritin: studies of iron binding and oxidation show differences in the two iron sites. Biochemistry 36, 4 3 2 4 1 . Trikha, J., Waldo, G.S., Lewandowski, F.A., Ha, Y., Theil, E.C., Weber, P.C. and Allewell, N.M. (1994) Crystallization and structural analysis of bullfrog red cell Lsubunit ferritins. Proteins 18, 107-1 18. Trikha, J., Theil, E.C. and Allewell, N.M. (1995) High resolution crystal structures of amphibian red-cell L ferritin: potential roles for structural plasticity and solvation in function. J . Mol. B i d . 248, 949-967. Tsugita, A. and Yariv, J. (1985) Preliminary results for the primary structure of bacterioferritin of Escherichia coli. Biochem. J . 231, 209-212. Wai. S.N., Takata, T., Takade, A,, Hamasaki, N. and Amako, K. (1995) Purification and characterization of ferritin from Campylohactrr jejuni. Arch. Microhiol. 164, I 6. Wai, S.N., Nakayama, K., Umene, K., Moriya, T. and Amako, K . (1996) Construction of a ferritin-deficient mutant of Campylohacter jejuni: contribution of ferritin to iron storage and protection against oxidative stress. Mol. Microhiol. 20, 1127-1 134. Watt, G.D., Frankel, R.B. and Papaefthymiou, G.C. (1985) Reduction of mammalian ferritin. Proc. Nut1 Acud. Sci. USA 82, 3640-3643. Watt, G.D., Frankel, R.B., Papaefthymiou, G.C., Spartalian, K. and Stiefel, E.I. (1986) Redox properties and Mossbauer spectroscopy of Azotohacter vinelandii bacterioferritn. Biochmistry 25, 4330-4336. Watt, G.D., Jacobs, D. and Frankel, R.B. (1988) Redox reactivity of bacterial and mammalian ferritin: Is reductant entry into the ferritin interior a necessary step for iron release? Proc. Nut1 Acad. Sci. USA 85. 1451--746I . Watt, G.D., Frankel, R.B., Jacobs, D., Huang, H. and Papaefthymiou, G.C. (1992) Fez+ and phosphate interactions in bacterial ferritin from Azotohacter vineltmdii. Bioc~hemistry31, 5612- 5619. Weaver. J. and Pollack, S. (1989) Low-M, iron isolated from guinea pig reticulocytes as AMP-Fe and ATP-Fe complexes. Biochem. J . 261, 787 -792. Weir, M.P., Gibson, J.F. and Peters, T.J. (1984) Biochemical studies on the isolation and characterization of human spleen haemosiderin. Biocheni. J . 223, 3 1-38. Williams, R.J.P. (1982) Free manganese(I1) and iron(I1) cations can act as intracellular cell controls. FEBS Left. 140, 3-10. Williams, R.J.P. (1990) Iron and the origin of life. Nature 343, 213-214. Wrigglesworth, J.M. and Baum, H. (1980) The biochemical functions of iron. In: Iron in Biochemistry and Medicine. I1 (A. Jacobs and M. Worwood, eds), pp. 29-86. Academic Press, London, New York. Yariv, J . (1983) The identity of bacterioferritin and cytochrome h,. Biochem. J . 211, 527. Yariv, J. (1996) Circumstantial evidence for cytochrome h l involvement in the functioning of lac-permease in respiring Escherichia coli. J . Theor. Biol. 182, 459462. ~

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How Did Bacteria Come to Be? Arthur L. Koch Department of Biology, Indiana University, Bloomington, Indiana, 47405-6801, USA

ABSTRACT

Bacteria in the modern taxonomic sense are one of the three Domains. They must have split from the other two after the bulk of the development of biochemistry and cell biology had taken place. Up to the time of the Last Universal Ancestor (LUA) the world had been monophyletic with little stable diversity. This is to say that as advances took place the older forms were eliminated and diversity was only temporary. Two kinds of events could, in principle, permit stable diversity to arise. One kind occurs when two nearly simultaneous, different advances occur, both of which overcome the same problem. While the previous type would be supplanted, if the new types did not compete with each other, new niches and habitats could lead to stable diversity. The second kind is a saltation or macroevolutionary event that greatly expands the biota and reduces previous constraints and thereby drastically reduces competition; this generally leads to a ‘species radiation’ and results in the development of a spectrum of biological types some of which persist and do not compete with each other. It is proposed that the two splits to yield the three Domains of Bacteria, Archaea, and Eukarya, resulted from one of each of these two processes leading to diversity. One arose from the consequences of cells accumulating substances from the environment, thus increasing their internal osmotic pressure. This resulted in two nearly simultaneous biological solutions: one (Bacteria) was the development of the external sacculus, i.e. the formation of a stress-bearing exoskeleton. The other (Eukarya) was the development of cytoskeletons and mechanoenzymes, i.e. formation of an endoskeleton. The other event ADVANCES IN MICROBIAL PHYSIOLOGY VOL 40 ISBN 0- 12-027740-9

Copyright 0 1998 Academic Press All rights of reproduction in any form reserved

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causing diversity was the invention of an effective way to tap a new energy source and allow the biomass to increase extensively permitting a radiation of many different types of organisms . I suggest that this seminal advance was the development of methanogenesis . This caused a short-lived expansion and radiation before oxygen-producing photosynthesis allowed a still more major expansion and decreased the number of methanogens . Some details of these processes are elaborated . In particular. the evolutionary process that permitted the development of a sacculus. interpreted in light of the bacterial physiology of today’s organisms is presented . It is argued that many great advances arise by developing a number of totally different processes for other purposes that can then each be modified to combine for yet another purpose . 1 . Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Evolution of Domains - a scenario . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. The monophyletic period . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. The development of diversity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3. Advances in energy transduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4. Two splits to yield three domains . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5. Osmotic challenges and the exoskeleton versus the endoskeletons as

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solutions to this problem . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 363 2.6. Thermophilic or methanogenic origin of Archaea . . . . . . . . . . . . . . . . . . . 364 2.7. The splits into the Domains: Bacteria, Archaea, and Eukarya . . . . . . . . . . 365 Bacterial wall formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 367 3.1. Mechanisms brought together for wall biosynthesis . . . . . . . . . . . . . . 367 . . . . . . . . . . . . . 368 3.2. Biochemistry to make disaccharide pentapeptides . . 3.3. Biophysics of extrusion through the lipid membrane with the help of bactoprenol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 370 372 3.4. Extrusion of proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.5. Energy for the synthesis of the wall fabric . . . . . . . . . . . . . . . . . . . . . . . . . 373 373 3.6. Coupling of autolysis and synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.7. The wall was an innovation at the time of the split of early life into domains . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 374 The function of the bacterial wall - non-growth aspects . . . . . . . . . . . . . . . . . . 374 374 4.1. Hexagonal nature of the wall fabric . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2. Structure of wall fabric in the plane of stress . . . . . . . . . . . . . . . . . . . 4.3. Porosity of the wall . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 380 382 4.4. Elasticity of the wall . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The function of the bacterial wall - growth aspects . . . . . . . . . . . . . . . . . . . . . 382 382 5.1. Rod elongation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2. Inside-to-outside growth for Gram-positive organisms . . . . . . . . . . . . . . . 384 5.3. Insertion into the stress-bearing wall of Gram-negative organisms . . . . . . 384 5.4. Poles of rod-shaped bacteria are metabolically inert . . . . . . . . . . . . . . . . . 386 5.5. Cell division and middle finding strategies . . . . . . . . The wall of the first bacterium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 388 392 6.1 The first Gram-negative bacterium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusions. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 393

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7.1. The model for evolutionary creative breakthroughs . . . . . . . . . . . . . . . . . 393 7.2. The scenario for the formation of the three Domains . . . . . . . . . . . . . . . . 394 7.3. The bacterial way of life . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 394 395 Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 395

1. INTRODUCTION

The evolution and development of the bacterial wall is emphasized here. Like most creative acts, it must have been accomplished by bringing together processes that had been developed for other, quite different, purposes. The bacterial wall is special in that its development involved the interplay of biochemistry, biophysics and membrane biology. The Grampositive and Gram-negative wall is a strong flexible fabric synthesized outside the bacterial cell with peptide and glycoside chains that are cross-linked in the gigantic fabric that surrounds the cytoplasmic membrane. The goal of this speculative review is to deduce from what we know of the biology of current organisms and to present a description of the interaction of physical forces, biochemistry and biophysics, that led during evolution to the Domain of Bacteria. In particular, I will try to bring together the different cellular functions needed to build the sacculus and implement its function in the wall physiology of the bacterium. There is another goal of this paper: this is to consider a major way that truly creative and original developments arise. One way (not considered more here) is that previous developments create an environment that is there only by chance, and which have no function as a result of other developments. This presents an opportunity for novel advancement. This idea has been promulgated by Gould and Lewontin (1979) and reanalyzed recently by Queller (1995). But there is a more profound mechanism generating novelty; this is the development of a new process, an evolutionary breakthrough, a scientific advancement, a new art form, etc. For this mechanism, it is not enough for only one process to be developed, but for a number of quite diverse advances for quite diverse purposes to arise. Then there is an opportunity to combine such advances and modify them for an entirely new purpose. This can be the saltation, the macroevolutionary step, the grand synthesis for the opening of a new realm of opportunities. In the rest of this paper, there are a number of examples of this type of saltation taking place in the biology of early life. In the Conclusion section I will take an example from the development of molecular biology to illustrate the same mechanism for the development of great advances.

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As an article of faith, I believe that life originated on this (or a similar) planet and only became capable of Darwinian evolution when it became cellular and was able to self-improve (Koch, 1984a, 1985a, 1994, 1995a,b). This hypothesis has now been subscribed to by Orgel (1987), Fleischaker (1990) and Morowitz (1992). Darwinian evolution became possible when a cell was able to grow and to undergo mutation and still grow, albeit in the mutated form. Darwinian evolution not only requires a system that can replicate and mutate and propagate the mutant, but it also requires that the living entities are favored or are selected against by their ability to cope successfully with their environment. Once started, life evolved many new aspects and these cellular processes were adapted and selected to do their jobs more effectively.

2. EVOLUTION OF DOMAINS

- A SCENARIO

The development of life on this planet depended very much on its creation, the evolution of its geology and what kind of abiotic synthesis could take place and accretion of organic molecules from space (Haldane, 1929; Oparin, 1936; Miller and Orgel, 1973; Chang et al., 1983; Deamer, 1997). Then followed an extensive period in which much important evolution took place. Almost by definition, before the split into the three Domains, life was monophyletic (Fig. I). That is to say that before the time of the Last Universal Ancestor (LUA), life had been evolving and had created many of the advances that we teach in cell biology courses. This is why the trunk of Fig. I is depicted as being broad and tall. At the time of the LUA, life had proceeded very far, but it had done so without the development of stable diversity. The single origin of successful life on this planet is attested to by the similarity of life processes and the sequence homology between proteins of different organisms. Figure 2 is in the more conventional representation, but emphasizes the points to be developed here. 2.1. The Monophyletic Period

But why was stable diversity not evolved early on? Why, conversely, did diversity develop later? I think that the answers lie in the Competitive Exclusion Principle of Gause (1934). His experiments were done and his theory developed in the 1920s and 1930s with studies on various protozoa and bacteria. His conclusion (expressed in more modern terms) was that an ecosystem can have only as many species as it has different resources or different accessible habitats. How would that apply to the early earth? Once

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Pre-cellular 'life' Figure 1 Early evolutionary tree. This is a different version of evolution compared with the more usual versions, as shown in Fig. 2. The emphasis here is that many things had, and must have, happened before the time of the First Cell. They must also have happened before the time of the Last Universal Ancestor. It also emphasizes the magnitude of the evolutionary changes before the splits that yielded the three Domains.

cellular life was present, the conditions for growth still would have been quite limiting. (Note that at the bottom of Fig. 1 the only energy input is from abiotic processes and only in the crown of the tree are there additional sources.) In Table 1, the energy sources that have been thought to power early life are listed, more or less in the order that they have been proposed in the literature. But most would have required extensive evolution before they could be used. Glycolysis as we know it, would have to await both a major supply of carbohydrates (at least monosaccharides) and the coordinate synthesis of a dozen enzymes whose functions dovetailed into each other. Of course, as a process in intermediary metabolism it occurred earlier, but as a source of energy it must have occurred later than the time of the LUA.

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SURVIVE HIGH TURGOR

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PRESSURE BECAUSEOF

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and photosynthetic

Abiotic generation Biomass small

No EXOSKELETON OR ENWSKELETONFirst

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Cell

Figure 2 Evolution from the point of view of turgor pressure and the availability of energy. The evolutionary tree is presented in contemporary form but adds several additional features. It indicates the increase during evolution of internal osmotic pressure, sources of energy, and the typical lifestyle of members of the three domains.

Anoxygenic photosynthesis would have to await the development of a membrane structure that is very complicated from the point of view of both biophysics and biochemistry. It would not generate biomass unless there were organic compounds at the right oxidation state. Oxygenic photosynthesis would have to wait until two types of photosynthesis from different organisms were linked together. Respiration would have had to wait until there was a complicated arrangement of proteins and prosthetic groups within a membrane system and until there was an abundant supply of reduced material and simultaneously an abundant available supply of a suitable oxidant; i.e. 02.Yes, there are other oxidants, such as nitrate and fumarate, but they would not have become abundant until oxygenic photosynthesis had been functioning for some time. Methanogenesis, it is argued below, came first as a major step in increasing the world biomass. It still had to wait until a series of membrane chemiosmotic systems were developed and coupled together to allow reduction of C 0 2 to methane with the utilization of electrons from low redox materials like H2 and other substances that would have been present before oxygenic photosynthesis. However, abiotic pyrophosphate utilization and a special restricted form of chemiosmosis probably functioned from the time of the First Cell.

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Proposed early sources of metabolic energy.

Process

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Fermentation

GIycol ysis

High-energy phosphate bond formation Respiration

Abiotic pyrophosphate formation Electron chain

Paucity of continuing supply of substrate Uptake into the cell

Anoxygenic photosynthesis Generates a protonmotive force Oxygenic photosynthesis Oxygen and reduced carbon Chemiosmosis Methanogenesis

Protonniotive force (PMF) generation Four chemiosmotic steps; many factors

Had to await oxygenproducing photosynthesis Does not cause a redox reaction but uses one Synthesis of two pathways for anoxygenic photosynthesis Needs available oxidant and reductant Most likely to have arisen the earliest

2.2. The Development of Diversity

Before the development of the major sources of bioenergy (photosynthesis and methanogenesis, and then respiration) there could have been only a small quantity of life on the entire globe. After photosynthesis and methanogenesis, the total biomass of the planet became much larger. The point to be made is that these energy-trapping processes developed just a little before or a little after the time of the LUA and the creation of the three Domains. Before that, when none of these energy-trapping systems was present, life was restricted to small inputs of utilizable resources produced abiotically by meteorological or geological processes or from input from extraterrestrial sources (see Deamer, 1997). Thus, the evolutionary game was the same for most of the period designated by the tree trunk in Fig. I ; it was life on a shoe-string but it was when cellular life and cell biology was developing. Contrast this with what happened when one of the energetic processes developed, such as oxygen-producing photosynthesis or methanogenesis. Then, the total population of the world must have grown explosively. This would have generated a biological radiation, such as occurred when terrestrial air-breathing animals arose and many groups of animals quickly evolved from that start. During the earlier radiations around the time of the LUA many mutational changes might have been established by the founder effect and, at least, persisted temporarily because competition was temporarily decreased by the new wealth of energy availability. For the purposes of this essay it is necessary to emphasize that until the energy-trapping improvements occurred, the number of individual

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organisms would remain small. So in the energy-restricted era, developments to fitter individuals depended on progress in the adaptation of the individual organism to develop better intermediary metabolism, better ability for macromolecular synthesis, etc. The ‘etc.’ includes the six remaining classes of processes listed in Table 2. When an improvement occurred, the individual’s descendants would out-produce and out-grow the others, but within the confines of a fairly constant biomass. This is a situation in which the competitive exclusion principle should operate. It would be this situation that may have blocked the early development of diversity. What if two different advances occurred in different organisms at the same time? While each would displace the earlier parental organism, they finally would compete with each other and one would be the eventual survivor. What special conditions would be necessary for stable diversity to develop? The competitive exclusion principle leads to the following concept. If neither new organism did not change ‘the rules of the game’ then only one would survive. If, however, the ‘rules’ were changed, then they both might survive. In the present context, this means that at least one would enter a new habitat or the new mode of life, but this specialization would eliminate it from the old. The advances that permit stable diversity must have occurred close to each other in time, or at a time when a radiation was taking place. Each of the advances at the beginning of organismal diversity must have opened up new and different opportunities. Then it must have been possible for further changes to become fixed before the two new lines would come into direct competition, and possibly by then secondary changes in each line could have been established that were stabilizing. 2.3. Advances in Energy Transduction

Now let us reconsider the major advances in ability to transduce energy, to funnel it into the biosphere. These include photosynthesis and methanoTuhle 2 Processes developed during the monophyletic period.*

I . Exploitation of chemical energy sources 2. Uptake and extrusion of small molecules 3. Biosynthesis pathways for intermediates 4. Reliable macromolecular synthesis 5. Regulation of macromolecular synthesis 6. Synchrony of DNA synthesis and cell division 7. Shape determination and cell division 8. Responses to environmental challenges *Retained today with relatively minor variations.

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genesis. In what sequence might they have been perfected and why were they critical to the original establishment of a stable diversity? I will argue that it had to be that the earliest advance that produced new supplies of substrates and biologically utilizable energy to the point that it gave rise to a large increase in the population and biomass of the world was methanogenesis. Of the set of processes listed in Table 1, methanogenesis (listed last) must have occurred first, for several reasons. First, it would work in the primitive world because both reducing power and COz would be readily available. Secondly, it involves only substrate metabolism and chemiosmosis - processes that probably were present from the time of the first cell (Koch, 1985a; Koch and Schmidt, 1991). Anoxygenic photosynthesis of the type in Rhodospirillum rubrum yields metabolic energy but does not produce any primary carbon compound at the oxidation/reduction level needed for the general components of cytoplasm. (In air and light, R. rubrum can grow chemoautotrophically with hydrogen and carbon dioxide, but it cannot grow anaerobically in the light without a suitable carbon source at an appropriate oxidation level.) The role of purple sulfur bacteria, which grow autotrophically under anaerobic conditions in the light but require reduced sulfur compounds, could have been of some use, but only appeared to have arisen after the LUA. Presumably, the purple bacterial type of photosynthesis (both sulfur and non-sulfur) developed not only after the Bacteria split from future Archaea/Eukarya, but after the Bacteria had split into several subdivisions, including ‘purple bacteria’. This is a reasonable assumption because only this division of Bacteria has members with this type of photosynthesis, although there are many with similar 16s rRNA molecules that are not capable of photosynthesis. Oxygenic photosynthesis was perfected by the cyanobacteria, and this did happen after the Bacteria had split off and then evolved into several subgroups with various types of anaerobic photosynthesis. Still later of course the cyanobacteria contributed chloroplasts to higher plants. Oxygen-producing photosynthesis is now responsible essentially for the creation of all the biomass of the planet. This recycling or trickle-down of carbon by way of food chains and energy can now support a world that can effectively use glycolysis and respiration to increase further the diversity and lead to the modern complex food chains and food webs.

2.4. Two Splits to Yield Three Domains

As summarized in Fig. 2, the weight of current 16s rRNA evidence is that the LUA gave rise to Bacteria and Archaea/Eukarya, and the latter group split somewhat later. From the Cause principle there had to be two splits each yielding a different non-competing population to form the three

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Domains. The argument of this paper is that one split had to do with the way cells could cope with osmotic problems. The sacculus of the Bacteria (and the quite similar sacculi of some Archaea) was one solution to the problem and the other solution was the development of the cytoskeleton and the mechanoenzymes. These two solutions both acted to keep the cell from expanding due to osmotic pressure differences between the inside and outside of the cell. The latter development occurred at some time in the Archaea/Eukarya lineage. While these were the two independent strategies that were the basis of one major split according to the proposed hypothesis, the second split was due to a different type of evolutionary advance - the development of an energy-exploiting process, methanogenesis. This allowed the radiation of the Archaea because with abundant energy a variety of different forms could grow even if individually a species was not the most efficient competitor. This radiation yielded groups such as those with strong cell walls (pseudomurein, S-layer, etc.), and thermophiles. It is important that the exo- and endoskeletons of Bacteria and Eukarya, respectively (Fig. 3), developed close enough in time to lead to stable diversity. Simply developing the sacculus would have given the Bacteria an advantage. From the Gause principle, if developing a cytoskeleton and mechanoenzymes had occurred much earlier, this would have prevented the Bacteria from arising, or the other way around. To stress the same

Stong material resistant to tensile forces in

Two Dimensions

One Dimension

\

Cytoplasmic membrane

Exoskeleton or Sacculus

Endoskeleton or Cytoskeleton

Figure 3 Two responses to osmotic stress caused by cells that have acquired the ability to concentrate medium constituents to create their own osmotic problem. On the left is the typical bacterial structure; on the right is the eukaryotic form.

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point, simply developing methanogenesis and abundant energy and not other mechanisms would have started a radiation that eventually would become simplified to a monophyletic biosphere without Bacteria and Eukarya to create and open niches for specific species. However, with all of these processes evolving nearly simultaneously it could be understood how the three Domains were stably and successfully initiated. I do not have a reason to choose whether the beginnings of development of the cytoskeleton or sacculus or methanogenesis happened a little before or after the splits of Bacteria, Archaea, and Eukarya from each other, but the timing had to be close. The subsequent radiations in all Domains included very numerous, stable and diverse organisms because new habitats and new resources were being developed by the increasing diversity of life.

2.5. Osmotic Challenges and the Exoskeleton Versus the Endoskeletons as Solutions to this Problem As the ancestors of the LUA progressed and developed and polished intermediary metabolism (see Tables 1 and 2) they had need to improve and expand their uptake systems. In particular, they must have evolved more avid systems for uptake of small molecules and these needed to be coupled to metabolic energy in order to do the thermodynamic work of taking up and concentrating the solutes. A consequence of the improvement in active transport is that the osmolarity of cytoplasm of the cells must have increased. That means the tendency for water to rush into the cell must have concomitantly increased. So, progressively, cellular life had to face the possibility of self-destruction as a result of swelling up and bursting due to its own success in improving metabolism. The avoidance of selfdestruction apparently was solved in two independent ways. There are several ways of counteracting osmotic forces. There are only four strategies used by modern organisms: 1. Having a strong cell wall 2. Having a cytoskeleton 3. Exporting water continuously as it flows in 4. Becoming permeable to salts (for halophiles)

The last two strategies require very specialized organization or require restriction to a very special habitat and we will not consider them further here. The first two are critical to the development of the hypothesis developed in this paper and we will deal with them below.

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2.6. Thermophilic or Methanogenic Origin of Archaea

While the emphasis of this paper is that the need for resistance to osmotic pressure was the raison d’etre of Bacteria, it will be necessary to consider the other Domains because the formation of all depended on the special characters of each. I have suggested (Koch, 1995a,b and here) that another split was due to the perfection of methanogenesis which permitted the expansion of what was to become the Archaea. However, contrary to this suggestion it has been proposed that thermophilia (Woese, 1979, 1987, Yamagata et al., 1991) was the initial basis of the separation of the Archaea from the Eukarya and not the ability to carry out methanogenesis. The extant organisms that are deepest in the Archaea lineage are thermophilic and this was the basis for the suggestion that thermophilia might have been the cause of diversity. If the splitting of the Archaea from the Archaea/Eukarya depended on the evolution of ability to survive at high temperature, then today’s mesophilic methanogens should show signs in their genome of previous high-temperature adaptation. However, I was unable to find data supporting this suggestion. On the other hand, the development of ability to grow methanogenically would have been very successful in a world without oxygen at any temperature. The geological record is quite clear that the first major fossil form of biocarbon has the isotope signature of being produced by methanogenesis (Hayes, 1983). Not surprisingly, this first flush of success in increased biomass production was halted with the development of oxygenic photosynthesis and this decreased the habitat where methanogenesis could be successfully practiced; this decrease in methanogenesis is also recorded in the record of carbon isotopes in ancient rocks. Today, an increasing number of methanogens are continuing to be discovered; however, the total collective habitat and current biomass of these organisms is small. If thermophilia started from mesophiles, it would have required revamping the structure of their proteins to have more hydrogen bonds, more salt bonds, more covalent cross-bridges, and more apolar bonds. It also would have required adaptation of the cytoplasmic membranes and changes in the ionic properties of the cytoplasm. Developing mesophilia starting from the thermophilic state would require the reverse changes. So, if mesophilia had arisen from thermophiles i t might be thought at first that some extra stability and rigidity would not interfere with life at a lower temperature. However, if the protein structures were too stable they would be too inflexible and rigid to be effective in catalysis. It is known that some organisms which can grow in a broader range of temperatures than most organisms succeed by having equivalent enzymes for key catalytic function, but the two enzymes differ in their temperature optima. Such organisms express different genes when grown at different temperatures. The implications are that

365

HOW DID BACTERIA COME TO BE?

non-thermophilic methanogens should have some remembrance in their genomes in the form of now inactive pseudogenes whose estimated protein structures should have features of high-temperature forms. They could be like the evolved P-galactosidase genes that are present and only reappear under strong selection.

2.7. The Splits into the Domains: Bacteria, Archaea, and Eukarya 2.7.1.

The splits

With these issues as a background, the proposed scenario of splits to form domains is as follows: in one line of descent, existing processes (Table 3) that served other purposes were combined to allow the formation of an external sacculus to resist the internal osmotic pressure. In another line, a number of existing membrane transport chemiosmotic systems for different specific purposes combined together to support methanogenesis (Table 4). In the third line, Eukarya developed with cytoskeletons and mechanoenzymes. Table 3 Originally independent processes needed to create an exoskeleton. ~

Class

Process

(i)

Simple modifications of the then existing enzymatic steps The export of polysaccharides with the help of bactoprenol, a lipidsoluble aliphatic chain Extrusion of proteins, such as PBPs The secretion of enzymes with the help of hydrophobic leaders and and autolysins signal peptidases for saprophytic growth Steps of intermediate metabolism Formation of the external links driving endergonic processes in through trans-peptidation and PP the form of disposable splitting. Because the wall is outside pyrophosphate and peptide bonds the cell proper, exportable energy has to be provided Allosteric or carefully regulated Carefully controlling the essential holoenzyme of intermediary action of autolysins in order to metabolism grow safely

(ii)

(iii)

(iv)

(v)

Variation on

Metabolic transformations to make disaccharide pentapeptides Extrusion of the oligopeptidoglycan through the lipid membrane

~

~~

PBP, penicillin binding protein; PP, pentapeptide.

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ARTHUR L. KOCH

Tuhle 4 Chemiosmotic steps of methanogenesis from H2 and C 0 2 . Steps Hz + COz+ M F CHO-MF + H4MPT CHO-H4MPT + H i CHz-HdMPT' + Hz CH2-HdMPT + Hz CH3-H4MPT + HS-COM CH3-SCOM + Hz 4H2 + COz

KJ/CH4 Comment +CHO-MF + HzO +CHO-HdMPT + M F +CHz-HaMPT+ + HzO +CHz-HdMPT + H'THMP +CH3-HdM PT +CH?-SCOM + H4MPT +CH4 + HS-COM + CH4 +H20

+ 16 -5 -2 -5 -20 -29 -85 - I30

Consume P M F Create P M F Create P M F Create P M F 4H Extruded +

PMF, protonmotive force. Modified from Blaut ef ul. (1990).

2.7.2. The Archura The steps of methanogenesis from COz and H2 are shown in Table 4.The individual steps would have been of use when the medium supplied certain particular substances, but it was only when all the steps became linked together to form a metabolic pathway that a great step was accomplished because of the general availability of input substances. Once this happened, the subdivisions of the Archaea could have arisen rapidly because of the expansion in growth opportunities. But as mentioned, the global advantage to methanogenesis was short-lived because of emergence of the much superior process of oxygenic photosynthesis. In part, the latter was effectively superior because the oxygen inhibited the methanogenic process. It was only later that combinations of Bacteria and Archaea could function as consortia such that one produced hydrogen and the other consumed it as in the wellknown cases of syntrophy. The original example was an isolate that was thought to be a pure culture and was called Methanobacillus omelianski, (Bryant et a/., 1967). However, this apparently pure culture was a two-member consortium consisting of a true archaebacterium, Methunohucterium hrvanrii, and a second organism that converts ethanol to acetate and molecular hydrogen. Methanogenesis would create an energy supply almost anywhere on the primitive earth. It would lead to great expansion in strategies and niches for methanogenic organisms. It would also lead to a great diversity of biotypes most of whom could survive because competition was at least temporarily ineffective during the population expansion. The methanogens diverged in sundry ways. For example, one path led to the development of pseudo-

HOW DID BACTERIA COME TO BE?

367

murein, similar to the murein of Bacteria to overcome the osmotic problem, and another was the development of the S-layer that may give some support for the cytoplasmic membrane in resisting the osmotic pressure differential from inside to outside. It may be that thermophilia was one of the other emergent strategies resulting from the radiation and one that occurred very early in the development of Archae.

2.7.3. The Eukarya A third line evolved to produce cytoskeletons and mechanoproteins. These could have arisen before or possibly after the development of methanogenesis. In their own right, both advances had important functions but together they could, to a limited degree, protect the cell from osmotic destruction. This path left the cell membrane flexible and allowed the cell to become bigger and to engage in endocytosis and exocytosis. This branch, the forebears of the Eukarya, subsequently gained entire Bacteria as symbionts, leading to mitochondria and chloroplasts. It was, however, the cytoskeleton and mechanoenzymes that permitted this branch to d o four unique things: (i) to actively extrude water; (ii) to grow large and still not be diffusionlimited; (iii) to control the dimensions of the cell by constructing essential cables from one point on the cytoplasmic membrane to another; and (iv) to be capable of taking up symbionts. I suggest that only the first on this list was an additional part of overcoming the osmotic problem.

3. BACTERIAL WALL FORMATION

The main purpose of this article is to consider how the saccular way of life developed. The bag-shaped super gigantic molecule of peptidoglycan bequeaths very many special properties and presents the opportunity for Bacteria to have a larger collective metabolic repertoire than that of the other Domains. 3.1. Mechanisms Brought Together for Wall Biosynthesis

A number of processes listed in Table 3 must have been developed independently from other functions and combined and modified to implement the production of the sacculus. Although the classes of processes must have evolved for reasons other than for the then novel process of sacculus formation, at some point they were combined to do something essential for

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saccular formation. Tables 5 and 6 list steps for wall synthesis and degradation. In the next section we consider them separately.

3.2. Biochemistry to Make Disaccharide Pentapeptides

Steps shown in the first part of Table 5 are simply modifications of what can be presumed to have been variations on existing intermediate metabolism. The steps are shown in a different format in Table 6 (Heijenoort, 1994, 1996; Labischinski and Maidhof, 1994; Koch, 1995b). Intermediary metabolism involved in amino acid and carbohydrate interconversions no doubt evolved, as Horowitz (1945) suggested, as a necessary consequence of a deficiency of abiotically produced small molecular weight substances needed for cytoplasm formation. After this development, the construction in the cytoplasm of the pentapeptide disaccharide from monosaccharides and amino acids is similar to the numerous processes of intermediary metabolism known today and only exceptional in two, almost trivial, ways. The first is that they involve the linkage to form N-acetylmuramic acid from N-acetylglucosamine

Table 5

Processes for exoskeleton (sacculus) function and turnover.

(i) Intermediary metabolism Form a sugar with a functional carboxyl group e.g. MurNAc from GlcNAc and o-Lactic acid Link this sugar to specific amino acids Of unconventional optical activity: D instead of L Of unconventional structure: Diaminopimelic acid, ornithine and E-lysine With unconventional non-alpha bonds: y-Glutamic acid, meso-Diaminopimelic acid Form a unit with both saccharides and peptides: e.g. disaccharide pentapeptide Pre-energize the head-to-head peptide bond Incorporate a ‘throw-away’ amino acid: e.g. terminal o-alanine Form linkage to isoprenoid bactoprenol carrier: Link unit to bactoprenol carrier with a pyrophosphate linkage (ii) Move the pentapeptide disaccharide to the outer face (iii) Secretion of proteins for external modification (iv) Secrete cross-linking enzymes (v) Secrete ‘smart’ autolysins: i.e. link degradation and synthesis together

HOW DID BACTERIA COME TO

BE?

369

Table 6 Wall biosynthesis in true bacteria such as E. coli and B. subtilis.

--------

Intracellular metabolism Fructose 6-phosphate 4 steps UDP-N-acetylglucosamine (UDP-GlcNAc) 2 steps UDP-N-acetylmuramic acid (U DP-MurNac) 3 steps

UDP-MurNac-L-ala-D-glu-meso-DAP

I step UDp-M urNac-L-ala-D-glu-meso-DAP-DAP-D-ala-D-ala 1 step

Linkage to the bactoprenol in the cytoplasmic membrane Prenyl-PP-MurNac-pentapeptide (lipid I) I step Prenyl-PP-MurNac-GlcNAc-pentapeptide (lipid 11) 2 steps Movement to the external side of the lipid bilayer

-

Linkage in both the glycan and peptide directions Transglycosylase expenditure of PP bond energy Transpeptidase expenditure of D-ala-D-ala bond energy

--

Autolysis of stress-bearing wall Gram-positive Surface action of amidase Surface action of glycosaminidase Gram-negative Synthesis and degradation linked by the 3-for-] model Endopeptidase Lytic transglycosylase Uptake and further degradation Recycling

----

phosphoenol pyruvate and NADPH. The second is that they use uncommon forms of amino acids. This includes D-amino acids, diamino acids, such as diaminopimelic acid, and the linkage of glutamic acid into the structure by virtue of the second (gamma) carboxyl group instead of the usual alpha carboxyl group. Of course, this ‘mix and match’ of enzyme types that had evolved primarily for degradation then became linked for a new purpose of synthesis of a new type of structure.

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3.3. Biophysics of Extrusion Through the Lipid Membrane with the Help of Bactoprenol

When cellular life started, it is likely that the fatty acid chains were not of uniform length and that the number and positioning of the double bonds was not optimal for membrane function to be a permeability barrier. Originally phospholipids, or their equivalent, had to result from abiotic synthesis and not be uniform in physical and chemical properties. This unevenness would have resulted in membranes that were more permeable to small molecules than those in current organisms. Membranes of modern organisms are permeable to water and are quite permeable to protons (Paula et ul., 1996). This means that chemiosmosis in the era of the first cell was quite inefficient (i.e. the pumped protons would return to the cytoplasm without bringing in a solute and were thus short-circuited). This would be of more importance when the processes that return the external protons to the cytoplasm are coupled to the uptake of solutes from the environment that are slow. One could expect that, by the time of the LUA, the phospholipid membranes and the chemiosmotic processes would have been optimized in order to minimize wasted proton flow. Together, mechanisms in the membrane would import predominantly hydrophilic materials and would be able to take up a large class of relatively small polar molecules. Transport systems would have become specialized and specific. Thus, the import of exogenous resources and intermediary metabolism of the parts to make disaccharide pentapeptide is first. But then the problem is reversed for the extrusion of the wall precursor through the cytoplasmic membrane. Once the disaccharide pentapeptide is made. its exportation through the cytoplasmic membrane involves a different type of mechanism than considered above. Its use for the export of oligosaccharides is quite common. Its application to wall metabolism is presented by Matsuhashi (1994) and Heijenoort (1996). This mechanism functions in all three Domains and involves attaching oligosaccharides of a range of compositions to a long chain molecule with 55 carbon atoms that is non-polar except at one end. In animal systems, this molecule is called dolichol and the slight modification used in bacteria is called bactopreno1 or undecaprenol. The long non-polar portion can stretch back and forth across the bilayer (Fig. 4). The polar end has an hydroxyl group that is esterified to a phosphate residue for murein synthesis. This polar end would prefer to extend from the membrane on either the inside or outside face than be inside the hydophobic membrane, but because of the hydrophobicity of the rest of the chain it would be occasionally possible for the polar end to move across from one side to the other of the membrane. Moreover, if the phosphate were coupled to a relatively

HOW DID BACTERIA COME TO BE?

371

Figure 4 Transport and glycan chain elongation. The sequences of events starting from the disaccharide pentapeptide (PP, on the inside of the cytoplasmic membrane) being linked to the bactoprenol. transported, linked to another unit now on the outside of the cell membrane to make a finished tetrasaccharide still attached to a bactoprenol.

small polar molecule it, too, could pass through the membrane. So the necessary step for the export of the disaccharide pentapeptide precursor on completion of its synthesis is that it is attached to the bactoprenol via a pyrophosphate linkage, one of the phosphates being contributed by the bactoprenol phosphate and the other from the disaccharide pentapeptide UDP precursor molecule. Another quite important constraint is that the peptide has to achieve a conformation that exposes a non-polar external surface such that the charged and hydrophilic groups are folded on the inside of a compact structure (Koch, 1990a,c, 1995b). Then it would be possible for synthesis on the inside to drive or push the export of the

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ARTHUR L. KOCH

oligopeptidoglycan to the outside concomitantly with being ‘pulled’ by its coupling to the growing oligopeptidoglycan chain on the outside. It is emphasized that this type of mechanism is a process that must have developed for export of polysaccharide for a variety of purposes in cells before the LUA. For example, one purpose would be to coat the outer surface of the primitive cell with polysaccharide in order to adjust the chemical nature of the surface to support the cell’s adhesive properties. Later in evolution such processes could cause binding of cells to other cells in multicellular organisms.

3.4. Extru8ion of Protein8

The first organisms were not even saprophytes; they could consume only abiotically produced very small molecules. Those organisms that died would represent a sink of non-utilizable biomass - no longer a resource. Eventually, ways to harvest that resource developed and for this purpose enzymes had to be synthesized and exported to the outside to degrade dead organisms (and larger available resource molecules). Naturally, saprophytism would involve a number of new evolutionary directions. For the saprophytic way of life it was necessary to be able to secrete proteins through the cytoplasmic membrane (Table 3, Point iii). This would require the paraphernalia of today’s bacterial cells but probably not the shuffling trafficking mechanisms from organelle to organelle and the controlled exocytosis that require the cytoskeleton and mechanoproteins as used in eukaryotes. But the mechanism for Bacteria is sophisticated enough, requiring docking proteins, signal sequences, signal peptidases, chaperones, and possibly still other players to have required extensive evolution. The development of such methods necessarily had to precede the development of a strong external wall. The seminal process of protein export is that the signal peptide, i.e. the initial part of the protein to be exported, is hydrophobic and can insinuate itself into the phospholipid membrane. Once through, it can relatively easily be forced or threaded further because the thermodynamics is easier as peptide chains are moved because a polar amino acid may enter as another leaves. Wall formation requires a series of PBP (penicillin binding proteins) (Matsuhashi, 1994; Heijenoort, 1996). These are integral membrane proteins and are bound in such a way as to expose their catalytic groups to the external face. Equally essential are the secreted autolysins that cleave the wall fabric in conjunction with enlargement (Doyle and Koch, 1987; Shockman and Holtje, 1994).

HOW DID BACTERIA COME TO BE?

373

3.5. Energy for the Synthesis of the Wall Fabric

An essential and well-established feature of peptidoglycan formation is that two forms of energy transduction are built into the structure of the exported peptidoglycan disaccharide so that the energy can be expended externally to the cell membrane to couple the disaccharides to each other and to couple the peptide portions of neighboring glycan chains to each other in a tail-totail linkage. This subject has been well discussed (Tipper and Wright, 1979; Koch, 1995b) and these stages are listed in Tables 5 and 6 and Figs 4 and 5 .

3.6. Coupling of Autolysis and Synthesis

The strategy of forming an exoskeleton depends on being able to open the structure and insert new material. This must be done safely or else the Bactoprenyl

Bacto-

.D r a w-l

Apposition

Bactoprrnyl

u

Stretching under stress

-34

Figure 5 Transpeptidation and stretching (shown as a view of the cytoplasmic membrane surface). The figures show diagrammatically the sequence of events in the transpeptidation that covalently links two chains. Once the membrane comes to bear stress, it stretches.

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ARTHUR L. KOCH

structure would rupture. Consequently, the cell must have arranged matters so that it adds and links new material before it cleaves old. This argument has been made by myself in many places (Koch et ul. 1981a; Koch, 1982a, 1983, 1984b, 1985b, 1988a, 1990a,b,c, 1993; Koch and Doyle, 1985, 1986) and further review is not necessary here, although aspects will become important in later sections.

3.7. The Wall was an Innovation at the Time of the Split of Early Life into Domains

When all five of the classes of processes listed in Tables 3 and 5 became workable and enmeshed, a synthesis to form a saccular structure would have become possible. This would have allowed bacteria to grow (and grow faster) under conditions to which other metabolically successful bacteria would succumb, due to the self-created osmotic problem.

4. THE FUNCTION OF THE BACTERIAL WALL ASPECTS

- NON-GROWTH

4.1. Hexagonal Nature of the Wall Fabric

To serve its function of covering the bacterium, the fabric must be strong and must fully cover the bacteria. In the growing cell, tensile forces would be applied in all dimensions on the surface of the fabric. Bacteria are small and it is for this reason that the walls are thin; bacteria do not use the stratagem of higher plants of constructing thick functional walls from linear material like cellulose chains. While such chains are very strong, they have a high tensile strength in only one dimension. In the plant this can adequately serve as the pressure-containing covering over the cell because the wall is sufficiently thick. Cellulose serves because the glucose moieties of the chains do form hydrogen bonds with similar parallel chains to produce helical bundles that surround the cell. Although the cellulose chains are very strong in their length dimension, and less in the side-to-side directions or from their ends, it is because of the numerous hydrogen bonds that they successfully serve their purpose. The bacterial sacculus (Weidel and Pelzer, 1964) is quite different from plant walls because it is covalently linked throughout the entire structure, giving two-dimensional tensile strength. Several views of the saccular unit structures are given in Figs 6, 7 and 8. Figure 6 shows five disaccharide pentapeptides linked in a chain which is 10 saccharide units comprising

HOW DID BACTERIA COME TO BE?

375

Figure 6 A portion of the oligopeptidoglycan chain with the disaccharide units rotated though 90”. The linked group of five pentapeptide disaccharides is shown to indicate the rotation of the peptide moieties. The figure also gives the peptide structure of the wall of both E. coli and B. suhrilis.

alternate N-acetylglucosamine and N-acetylmuramic acid. The peptide chains are depicted in an extended configuration and are not to scale. The example of the peptide structure chosen in the figure would apply to either Escherichiu coli or Bacillus sutitilis. The minor variations among other bacteria are discussed elsewhere. For a key to this literature see Koch (1995b). The vital points of the peptidoglycan fabric structure for all bacteria are: 1. A carboxyl group extending from the sugar chain. This is supplied by the carboxyl end of the muramic acid structure. 2. A diamino acid to supply a second amino group for the tail-to-tail linkage of peptide moieties extending from different carbohydrate chains. 3. A terminal D-alanyl-D-alanine peptide bond that is expended in the transpeptidation. This peptide bond supplies the energy that permits formation of the bond holding the two peptides from different peptidoglycan strands together. 4. A sequence of special amino acids such that the pentapeptide can achieve a compact structure as it is moved through the non-polar

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ARTHUR L. KOCH

NAG (N-acetylglucosamine)

NAM (N-acetylmuramic acid)

Octa-peplide cross-bridge

Downpeplide

UPpeplide

Figure 7 The strongest unit of wall fabric: the smallest tessera or unit of structure of the wall that surrounds a space in the surface. The term is used both for the pore and for the material at its edges even though it contains both peptides and saccharide and is made up of two oligopeptidoglycan chains. See key on figure.

cytoplasmic membrane, but can extend subsequently in an aqueous environment. 5 . A sugar chain structure that has disaccharides rotated at 90” relative to the adjacent ones. It is for this reason that the alternate peptide chains are not written out but simply indicated as ‘up-peptide’ or ‘down-peptide’ in Figs 6-10. (The role of the up- and down-peptides in the stress-bearing parts of the saccular structure is not clear, and it may be that they have no role.) For the same reason, the third peptide is shown pointing in a different direction than that of the zeroth and fourth, still within the plane. The chemical reason for the rotation is simply that the d a c t y l group that was added to N-acetylglucosamine to make N-acetylmuramic acid is sufficiently bulky to lead to steric hindrance so that there is no free rotation in the glycoside bond. In formation of the wall, the disaccharide pentapeptides are extruded, but must remain connected with the membrane because the bactoprenol

377

HOW DID BACTERIA COME TO BE? Peptlde in d i n e

D-7-Glul

L-Ala NAH

Glyca chain

NAG

\

NAG

-

NAG

NAG NAU

NAU Down-

-peptide Up NAG

peptide

D-Ah

L.DAP

L-DAP

D-Als D-l-Glut

D-l-Glut L-AlaNAH

/

L DAP

f^’ 2

D-Ah

Glycm chain

NAG

D.l-Glul

D-,-Glut

v

NAM

1 Ala-

L-Ah-

r

Glycan

/ chain

NAH

NAG

L-Ala-

-peptlde

NAW

NAM -UP

NAG NAM -

NAG HAM NAG L-Ala-

Downpept1L

NAG

\Glycan chain

D-r-Gkrt

Peptide in plane

Figure 8 The structural unit of the wall fabric: the tessera. A different view of a tessera is shown than in Fig. 7. Again, the hexagonal nature is clear. Also indicated are the six places where stresses from the rest of the wall act on the tessera in an intact fabric. NAG, N-acetylglucosamine; NAM. N-acetylmuramic acid.

attachment portion cannot move into the aqueous environment on the external face. Instead it must await a transglycosylation event cleaving the saccharide from the pyrophosphate group and linking it to the growing chain of the oligosaccharide. This catalysis is done by membrane bound PBPs. These are the PBP1, PBP2 or PBP3 and each of these are both transglycosylases and transpeptidases (Matsuhashi, 1994). The transpeptidase functions to link two adjacent polysaccharide chains through their pentapeptides. This forms a tail-to-tail bond from the penultimate D-alanine of the pentapeptide to the amino group of the free end of the diaminopimelic acid groups and frees the terminal o-alanine from a partner. There are constraints as to which peptides can engage in cross-linking in the plane of the surface. The up- and down-peptides cannot connect with

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ARTHUR L. KOCH

-

0N-Acetylglucosamine

N-Acetylmuramic acid

0

D-lactyl-L-Ala-D-isoGlut-A2pm-D-Ala-D-A

Donor

Figure 9 Two fused tesserae. In the fabric of the wall the glycan chains are linked to each other. Here just two chains are depicted cross-linked to each other by pairs of donor acceptor tail-to-tail peptides. Other peptides extend outward in the plane and may form tesserae with other adjacent strands. The short lines depict other peptides that extend upward or downward and may not have an active role in saccular structure.

HOW DID BACTERIA COME TO BE?

379

Figure 10 A typical portion of the wall fdbric. This is an extension of Fig. 9. It was drawn to have a similar distribution of glycan chain lengths to that determined analytically.

others in the plane and only every fourth peptide can link together two sugar strands within the same surface plane. Figure 7 depicts the smallest unit that in many copies can cover a surface of the cells with a covalent structure that can support stresses in the plane of the surface. The amino acids are not shown but the octapeptide composed of two tetra peptides in tail-to-tail linkage are shown by the three thick lines symbolizing the two peptides and the cross-link. The box indicates the hexagonal unit of structure, or ‘tessera’ (Koch, 1990a, 1995b). This term was adopted from that used for the pieces of colored stones that are affixed together to make a mosaic. There are six attachment points radiating from any peptidoglycan tessera. In a portion of wall in an intact growing cell there are stresses pulling the structure taut. These would be exerted at all of the six bonds leading from the tessera. It is these stresses that create the turgor pressure inside the living cell that just counterbalances the osmotic pressure differential from the inside to the outside of the cell. Just how the stresses would balance with each other in different directions would depend on their magnitudes and directions and would vary in particular parts of the cell wall such as the side wall or the poles. Also there could be an influence of stresses transmitted by the up- and down-peptides if these peptide are connected to other layers or membranes. They no doubt are attached in Gram-positive organisms to adjacent layers and the influence of these layers is no doubt important.

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4.2. Structure of Wall Fabric in the Plane of Stress

In all cases the fundamental shape of the tesserae is that of hexagons. A fabric composed of them would have the same structure as does the familiar chicken-wire of the hen house. Unfortunately, even though this concept is fundamental to basic microbiology, it has not been picked up and included in reviews on cell walls, and certainly has not found its way into miorobiology textbooks. Hopefully it will soon. Figure 8 has the same information as Fig. 7 and includes more detail about the peptide linkages. However, do not get lost in the details of the chemical structure, because here the problem is to extend the unit of structure to make a covering fabric. A start is shown in Fig. 9 of two fused tesserae. Now there are more internal stresses and there are eight points of stress pulling from the outside. Figure 10 shows a still larger fragment of wall structure. This was drawn to simulate a piece of fabric with a glycan chain length distribution not unlike the experimentally determined values from the laboratory of Holtje and Schwarz (Glauner et al., 1988; Glauner and Holtje, 1990; Kraft, 1997) for E. coli. Thousands of such structures would be necessary to cover the surface of E. coli. If the available analytical composition values are right, the problem arises that there is no stress borne by these sugar chains that project from the fabric fragment because they are free and unconnected. As depicted, the three peptide chains shown as if free might, or might not, connect to other fragments. In the latter case, the magnitude of the stresses in these peptide bonds would be much larger than in those bonds that are part of the intact fabric. These highly stressed bonds would be sensitive trouble spots and very little autolysin activity should lead to their rupture and then, possibly, to the failure of the entire cell. 4.3. Porosity of the Wall

Although the job of the sacculus is to constrain and retain the cytoplasm from undue swelling, it has another function which is to permit some molecules to move in from the outside of the cell membrane and others to move in the reverse direction. This requires that the sacculus be a sieve with sufficiently large pores and at the same time be of a sufficiently fine mesh to be sufficiently strong. Paul Demchick in my laboratory devised a very ingenious method to measure the porosity (Demchick and Koch, 1996). He prepared clean sacculi of both E. coli and B. subtilis. He also prepared commercial dextran fractions and labeled them with fluorescein. He then separated them into very narrow size fractions. His experiments were to view with a fluorescence

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HOW DID BACTERIA COME TO BE?

microscope the sacculi in home-built very shallow depressions formed in microscope slides. The depth of the depression was only 4 pm. If the dextran fraction’s size was too large to penetrate the pores of the sacculi, then the sacculi could be visualized because they stained negatively in the fluorescence microscope. If the molecular weight of fractions were too small then the fluorescein entered rapidly and the sacculi were almost invisible. Figure 1 1 shows diagrammatically the time course for filling the sacculi with dextrans of four sizes that pass through the pores at different rates; the righthand side shows the appearance in the fluorescence microscope at the same point in time. By observing the kinetics of the penetration he was able to ascertain that the pores were approximately 2.2 p m in diameter. This

............... 3

4

5

Log (time)

8

6

(sec)

Figure I 1 Time course of filling the SdCCUlUS with fluorescein-labeled dextran. This illustrates the experiment of Demchick and Koch (1996) in which sacculi of E. coli and B. suhrilis were exposed to fluorescent-labeled dextrans of different sizes. The size of pores (i.e. the mean area of the tesserae) could be estimated from the kinetics of this type of experiment.

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suggested that compact polymers or globular proteins of 55 kDa were just able to pass. Of course, unfolded proteins of larger size could pass if they should penetrate by a threading process, but this would be quite slow. The porosities of Gram-negative and Gram-positive organisms were about the same in spite of the difference in wall thickness. Presumably, bacterial walls, even of species with variant chemistries of their murein would have about the same porosity because the tesserae would be of nearly the same size. The size of the pores determined in this way suggests that the tesserae of Figs 710 are limiting the diffusion process generally. It does not seem that there are significant imperfections in the wall of exponentially growing bacteria to allow larger probes in. 4.4. Elasticity of the Wall

An equally important property that the sacculus of bacteria should have is that it should be flexible and extensible. Another student in our laboratory, Steven Woeste, attacked this aspect (Koch and Woeste, 1992) by using lowangle laser light scattering to measure saccular area under various conditions. He found that the area of the sacculi of E. coli could be made to expand up to 4-fold larger than the size of the isoelectric form by either increasing or decreasing the pH, or by acetylating or succinylating the free amino groups in the sacculi (Fig. 12). These results are consistent with the idea that the compact structure of the peptide containing both plus and minus charges becomes extended with only a little repulsive force generated simply by changing the distribution of positive and negative molecular charges so that they all are of the same sign. This result suggests that the oligopeptides, when they are unlinked to other peptides, will be in a more compact structure than when they are linked tail-to-tail and come to bear the stress due to turgor pressure.

5. THE FUNCTION OF THE BACTERIAL WALL ASPECTS

- GROWTH

5.1. Rod Elongation

Many bacteria grow in elongated forms. Possibly this is advantageous to the bacterium due to the resulting increase in the surface to volume ratio. However, it requires special mechanisms and utilization of special areas of physics for rod-shaped cell growth to occur. How this could occur in the absence of mechanoenzymes might appear impossible. I believe, however,

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1

383 I

Figure 12 Expansion and contraction of the Gram-negative sacculus. The results relate mean succulus surface area as a function of pH. m, in low ionic strength; 10.1 M urea; 0, from top to bottom, they signify succinylated sacculi, acetylated sacculi, untreated sacculi at same pH, and untreated sacculi in 1 M KCI; 0, theoretical net charge of the average charge per million sacculi calculated from the composition of sacculi and the pK values of the ionizable groups. (Results of Koch and Woeste, 1992).

+,

that the basis for this comes from the physics of membranes originally studied by LaPlace (1806) two centuries ago. As a minor part of his studies, he calculated that a bubble with closed ends and of a fixed diameter could grow as a cylinder if the pressure and surface tension were compatible with the radius of the cylinder. I used his physics to explain rod-shaped growth (Koch, 1982a; Koch et al., 1982a). The idea is that insertion into such a membrane, if the physical properties remained constant, would result in elongation and not bulging as the cell grows. The argument was simply that the cylinder extension is thermodynamically a zero free energy process whereas bulging would require expenditure of energy. Thwaites and Mendelson (1991) argued that this mechanism could not function without a force extending the cylinder. The essential point of confusion was in distinguishing the properties of a static system of a pressure-filled vessel and a dynamic growing system equivalent to the extensible membrane of an enlarging soap bubble (see Koch, 1995b). My further analysis of the problem (Koch, 1993) showed that the critical feature was that the new wall had to be added in a way that did not depend on the current static stresses in the cylinder, but once it became a part of the stress-bearing wall it

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must then respond to the stresses to the degree that its elasticity and its elastic limit would permit. In any case, no mechanoenzymatic system (similar to these in eukaryotes) has been found that could function to cause rodshaped growth in some more mechanical way. Thus, because very many bacterial cells do grow as rods, there is a paradox. 5.2. Inside-to-outside Growth for Gram-positive Organisms

The mechanism of growth of B. subtilis is now quite clear (Koch and Doyle, 1986; Koch, 1995b) and can be simply stated. From inside the cytoplasmic membrane cylinder surface new wall precursors are secreted and crosslinked in a two-dimensional sheet (they are probably also linked to some degree in the third dimension to the overlying wall) immediately outside the cytoplasmic membrane. During this polymerization process, the wall is not exposed to stretching by any tensile force. As new layers are added the older ones are displaced outward and become stressed in the longitudinal direction as the cell becomes longer. At some point the elastic limit of the murein is reached and tension builds up in the wall. Consequently, the autolysin cleaves more rapidly and relieves the stress within that layer. At this point the layer is no longer capable of bulging because its ability to stretch has been used up already in the elongation process. The physics of the situation is that the autolysin action results in helical groves that remain as further growth takes place, even though the outer layer of wall is being continuously removed. This causes the cell cylinder to twist as it grows (one end rotating relative to the other) (see Koch, 1989). 5.3. Insertion into the Stress-bearing Wall of Gram-negative Organisms

Gram-negative cells have only one layer of murein (Labischinski el ul., 1991), and so they cannot add a new layer as they autolyze an older one. Effectively, however, they do the same thing of inserting and turning over, but do it all within a layer. They add a new wall in a special way so that the new wall is installed before cleavages take place. At this point the new wall is covalently linked but bears no stress. Then, the old wall is cleaved. I had proposed several models (Koch and Burdett, 1984; Koch, 1988a, 1990a, 1995b) as to how this can happen and allow safe enlargement of the wall. Joachim-Volker Holtje (1993, 1996) modified and improved these ideas very significantly to include his ideas based on his data about the structure of the sacculus of E . coli. The very important point he added was to suggest the role for the 'trimers' during safe enlargement. These structures

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(Glauner et al., 1988; Glauner and Holtje, 1990) (Fig. 13) comprise only a very small fraction of the products of muramidase digestion of the wall ( < l%), but appear to be highly significant for the growth strategy. The trimers have three peptidoglycan units connected together through their peptides. This is chemically possible because the cross-bridged tail-to-tail structure still has one amino group free in one of two diaminopimelic acids of the bridge and it can, therefore, form a peptide with another penultimate D-alanine of a third pentapeptide. The basic idea is that the trimer is formed and then cleaved, but cleaved in a different way so that an old glycan chain is separated from the other two (Fig. 13). The ‘three-for-one’ model of Holtje is shown in Fig. 14. [For a more complete discussion and justification of its function see Holtje (1993, 1996) and Koch (1995b, 1998)l. When other constraints are considered, this model will require a number of enzymes and they probably must function as a holoenzyme and in such a way that the synthesis is orchestrated and completed before the hydrolysis phase removes the old chain. Thus, when an old chain is removed, it is replaced with a structure that contains three chains. This replacement of one oligoglycan chain by a group of three

Incoming Pentapeptide NAG

I

NAM-L-Ala-D-7-GI

c - - - - - - - - -

I I

------I

I I I I I I I

I

Figure 13 Trimer formation. Trimers form when a third pentapeptide is bound to the remaining amino group of diaminopimelic acid on the tail-to-tail bond of an existing cross-bridge.

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Endopeptidase resolution of the trimers

strand

I. II.

111. IV.

V.

Alignment t o template strand Addition of disaccharides t o form an oligopeptidoglycan chain Raft formation by tail-to-tail peptide Joining and extension of the two oligopeptidoglycan side chains Linkage t o form trimers of the two side chains with the stress-bearing bonds of the template strand Cleavage of the stress-bearing bonds to cut out the template and pull the raft into the stress-bearing plane

Figure 14 The ’three-for-one’ model. The model of Holtje (1993) for the safe elongation of stress-bearing wall is shown. It shows a triplet raft of three oligopeptidoglycans docked under a stress-bearing chain of the wall and linked to it with trimers. In the next stage past the point of this diagram the template strand will be excised by autolysins and the raft will be pulled into the plane of the sacculus and then come to bear stress.

is the reason for the ‘three-for-one’ name of the model; this substitution doubles the amount of peptidoglycan by replacing the ‘template’ glycan strand in the stress-bearing wall with the nascent three-chained raft of glycan strands. 5.4. Poles of Rod-Shaped Bacteria are Metabolically Inert

The strategy for the growth of rod-shaped organisms is simply that wall growth has to be partitioned into three morphological phases. First, the formation of new poles or septa must use the cylindrical region as a framework for the inward growing wall. Second, the cylinder must maintain itself at a constant diameter while it is elongating. Third, the extant poles must be metabolically inert and structurally rigid to support the cylinder part of the cell. An early attempt to measure the metabolic stability of the pole (Koch, 1982b; Koch et al.. 1982b) was not very successful. In part this was due to the diffuse nature of the tritium beta rays emanating from pulse-labeled

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walls of E. coli to form silver grains over a wider area than the distribution of the radioactive atoms with the high-resolution technique then available. With the better data of Woldringh e f a/. (1987), a higher resolution analysis could be carried out (Koch and Woldringh, 1995). We concluded that the poles were metabolically inert. Strong and convincing support has recently come from the work of de Pedro et al. (1997). These workers used a way to visualize D-cystine-labeled walls (the D-cystine replaces the D-alanine and with antibodies such walls can be detected by immunogold or fluorescent labeling technique). Their studies with the electron microscope or with the fluorescence microscope demonstrated that the old poles do not turn over. Early studies from Doyle’s laboratory (Mobley et al., 1984) for B. subtilis similarly provided evidence that the poles were inert. Merad el al. (1989) shifted organisms from conditions where teichoic acid-containing walls were formed to conditions where teichuronic acid-containing walls were made, and then stained at various times for the phosphate in the wall. These experiments are totally convincing in showing that the poles are essentially inert, that turnover occurs very slowly, and then only near the cylindrical regions. In addition, with streptococci, both the very old experiments (Cole and Hahn, 1962) and more recent data (Higgins and Shockman, 1976) with Enterococcus hirae show that the old poles are inert. Generalizing from the three organisms, E. coli, B. subtilis and E. hirae, it may be that only the cylindrical regions of Gram-positive and Gram-negative bacteria engage in systematic addition and turnover, though the Gramnegative and positive organisms do so in quite different ways. Otherwise the only way that wall growth takes place is by the formation of wall made of only new material for the new pole. This would seem to be the case for the constriction site of Gram-negative bacteria, the septum of B. suhtilis, and new poles of streptococci.

5.5. Cell Division and Middle Finding Strategies

Some bacteria are able to divide very close to their middle to form almost identical daughter cells (Burdett and Higgins, 1978; Trueba, 1981). In addition to the model for the cell finding its middle because the origin DNA copies are attached to the tips of the poles (discussed above), a second possibility has been recently proposed (Koch and Holtje, 1995). This model applies directly to rod-shaped organisms only. It depends on the fact that the existing poles are metabolically inert (as discussed in Section 5.4 for both Gram-positive and Gram-negative organisms). It assumes that the cytoplasmic membrane grows over the cell’s entire surface. Analysis of the forces that would be generated by this situation shows that if the murein and the cytoplasmic membrane have some affinity for each other, then the stress in the

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wall - which is constant throughout the cylindrical part - will be variably partitioned between the cytoplasmic membrane and the overlying murein. This will happen in such a way that tension in the cytoplasmic membrane part will be maximum at the very center of the cell and will increase during the cell cycle. Both this and our other model discussed below (Koch et al., 1981b), in which the symmetrical replicating DNA structure locates the cell center, may act together, particularly if the oriC DNA is not always bound to the membrane as seems to be the case (Ogden et al., 1988).

6. THE WALL OF THE FIRST BACTERIUM

The discussion above in Section 5 of the way that modern bacteria form and enlarge their walls is used in this section to extrapolate to the way primitive bacteria formed their walls. In Section 3 the formation of external wall stopped with the idea that, subsequent to wall addition, the older wall was cleaved to enlarge the cell. The same questions discussed above can be asked concerning the original bacterium. Where and how was new wall added? How was the wall enlarged? How did that cell divide? Two global strategies for cell enlargement have been studied in some detail: those appropriate for some Gram-negative and Gram-positive bacteria have been described in Sections 5.2 and 5.3.Consideration of plausible strategies and mechanisms for bacterial growth has been my interest for the past 20 years, and the point has been reached by producing a more global theory for modern organisms, so that it can now be applied to suggest how saccular growth originated. First we ask: could either the Gram-positive or -negative case for today’s microorganisms be that which functioned for a cell that had just perfected methods to form the bag-shaped molecule outside its cytoplasmic membrane? When the monolayer is finished, what then? A whole new set of mechanisms must have come into play to deploy as a saccular growth mechanism. While these early mechanisms could not be as sophisticated as those used in modern bacteria, at the start of the Domain of Bacteria they had to be sufficient to enlarge the wall safely. The mechanisms had to be simple and, especially, they had to be present at least in a related form beforehand. On this basis, I suggest that something like a Gram-positive coccus would have been the first bacterium. It would only need to be capable of forming a septum, probably with a mechanism similar to that in cross-wall formation of B. subtilis or E. hirae. This cross-wall is much thicker than a monolayer of murein, but as it is formed, the cytoplasmic membrane invaginates and new coatings of murein are added to the margin of the new layer. As growth takes place,

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the annulus becomes narrowed and eventually finishes closing, at which point the disk of the septum is completed. This septum-closing process requires that the cell must locate where the septum is to be formed and permit only new murein precursors to be added. Within this restricted region, glycan chain formation, chain extension and cross-linking of peptide chains would have occurred at random, as I believe they do in present-day organisms (Koch, 1988a). This is because no evidence exists in any organisms that there is any order in the orientation of the glycan chains, nor has any cogent reason been presented why order is needed (see Koch, 1988a). Most fundamentally, an ordering mechanism would have to be very sophisticated and therefore not likely to have been developed at the dawn of the Domain of Bacteria. For systematic localization of the addition site, a quite complex mechanism would be required for which there is no precedent and no known purpose. Consequently, it would have had to evolve especially for this new purpose. For the simplest process imaginable all that is really needed is the localization of a number of protein factors and precursors at the ingrowing margin of the forming septum, but they would not have to be arranged in a precise order or move systematically, since the wall-enlargement process requires a ternary interaction of donor, acceptor and enzyme. If there were to be a more regimented mechanism that arranges and specifies the location of the murein additions, it would need to control the movements of the bactoprenol molecules, the appropriate penicillin binding proteins (PBP), and also the existing recipient (acceptor) oligopeptidoglycan chain to be cross-linked for the formation of a doubly thick septal fabric. Ternary complexes are rare and therefore such processes usually occur instead in two stages. The rules for solution chemistry do not apply in the same way to membrane-bound molecules. Instead, they require the two-dimensional diffusion of the components. Some PBPs have their catalytic site on a flexible domain of the protein (Spratt et ul., 1988). That no doubt greatly increases the reaction rate, but is probably not an essential part of the first process for the formation of a wall of the earliest bacterium. Such a mechanism, in applying to only the future septal part of the cell wall, generates a circular region at the maximum diameter of the nearly round cell; the second necessary part of the process is the bisecting of the septum. This means that an autolysin with some selectivity would be needed. It must not attack the pole wall, but should attack the external part of the septum and its catalytic activity should respond to the biochemical production of cytoplasm and the consequent need of the cell to enlarge. I have argued previously (Koch, 1982a, 1983, 1995b) that enzymes that attack macromolecules would have increased rate of reaction if the substrate were under tension. While this would not be at all an original idea for chemists and is in fact a thermodynamic and kinetic truism, it has not

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been pointed out in a biological context except in my papers. I have also argued that a change in conformation might eliminate enzymatic activity, even though the stress was greater if it were the case that the bond conformation was not appropriate. It can be suggested that the original type of hydrolytic action in the primordial bacterium depended on autolysin being excreted as are many other proteins in modern organisms, but because autolysins have affinity for wall they will not diffuse away (Koch, 1988~). In this case, the nascent wall would not be under tension and therefore would not serve as substrate for the autolysin. The wall of existing poles would not serve as substrate because the assumption is that the bonds are in the wrong configuration. Only the exposed septum is stressed but is still in the right configuration (Koch, 1983; 1995b). As autolysis occurs the septal wall starts to split and becomes stressed. The physical forces, as tension due to growth occurs, lead to maximum tension in the middle of the thick septum (Koch er UI. 1985). Due to the distribution of stresses, continuing activity then splits the septum evenly down the middle until cell separation occurs. The next part of growth and cell division depends on the physical properties of the murein. The peptide portion is elastic and the glycan chain is not (Labischinski et ul., 1979). Once the tension in the plane of the wall becomes applied to both half portions of the now partially split septum by autolysin, further enzyme action, separation of the two daughters starts and stretching takes place to make the new poles from the septal material. We have found that the septum of B. suhtilis stretches 50% on being stressed (Koch and Burdett, 1986a,b). A less perfect fabric would stress more. So, repetition of autolysin action and the peptide stretching leads to growth of cell size with something approaching a coccal range of morphologies. But how does the septum become properly located? What type of machinery could find the middle of the cell? Consideration of the possibilities many years ago led to the hypothesis that attachment of the chromosome to the ends of the cell envelope had to be the key (Koch et al., 1981b; Koch, 1988b). Of the various cell components, the chromosomal DNA is the only molecule in the replicating cell that has the right dimensions to be a measuring stick to allow the cell to be bisected. The suggestion was proposed (Fig. 15) that the origin of the circular DNA would be attached to the tip of the pole (Koch, 1988b). Then, on initiation of chromosome replication (Donachie 1968; see Cooper, 1991, for an extensive discussion of chromosomal replication and cell division), one of the two sister chromatids would be separated from the pole of origin and would diffuse through the cell, possibly attached loosely to the membrane. At some point it finds the terminus DNA that then is attached to a binding site at the other pole. Then, in our proposed mechanism, a special (but not unique) kind of action would be needed. The receptor localized at the tip of the pole that is then bound to the

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Two newly created Ori/Ter binding sites

Oldest Ori/Ter binding site

Second Oldest Ori/Ter binding site

Figure 15 Model of DNA replication leading to formation of equal-sized cells. A cell near the end of the chromosome replication is shown. The oriC locus of DNA of the newborn cell is attached to the pole of the cell and the ter locus of DNA at the other pole. At the start of the replication of the chromosome, before the phase shown here, one sister remained in place and the other oriC has replaced i e r . The DNA in turn became centered in the cell and determined the site of the cell division and the synthesis of duplicate new OriiTer sites.

terminus DNA exchanges that DNA for the origin DNA. This would happen if it had a higher affinity for the origin DNA. The ejected terminus DNA could not find a free binding site and would be jockeyed through the cell, again probably loosely associated with the cytoplasmic membrane. This jockeying would end up locating the terminus close to the center of the cell because the replicating structure of DNA is of a theta shape (as shown in Fig. 15). This is a symmetrical structure in the cell because its two oriC regions are attached at each of the two poles. At some point the terminus complex would generate and fix the site of formation of two new binding sites for the two termini that result from the completion of chromosome replication. Initially, this model was invented to explain the known fact that E. coli and B. suhtilis divide almost precisely in half. Seemingly there is no biological reason for such a degree of precision and Doyle and I argued that this mechanism itself provided the incidentally high, but unnecessary precision, for cell division almost precisely bisecting the cell. So this would be a default mechanism that only would have required the development of a mechanism so that the terminus DNA as the last stage of chromosome replication would code and locate the paired receptors for origin/terminus DNA at that site. For the consideration of a primitive bacterium the only

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addition is that this must trigger as well the formation of the septum and eventually the oriC DNA must be bound at the tip of the pole.

6.1. The First Gram-negative Bacterium

The crux of the matter is that the very first organism that surrounded itself with an exoskeleton or sacculus was able to enlarge further. From the very first, a means of autolyzing the old wall selectively and safely had to be in place. Perhaps we can relax the requirement and substitute 'nearly always safely' for 'safely'. So far there are only two ways for safe enlargement that have been suggested: an allosteric autolysin that only acts hydrolytically if the integrity of the wall will not be compromised; and a holoenzyme that forms bonds simultaneously with the destruction of other peptide bonds (Koch, 1990a,b). The three-for-one model demands a great deal more than just that. Re-examination of Fig. 9 indicates that the donor peptides are all associated with one glycan strand and the receptor peptides with the adjacent strands. This has been shown to be in accord with the Holtje threefor-one model. This regimentation is required by the way the trimers are formed and resolved in the three-for-one growth mechanism. If the glycan chain in the template was very long then the growth by insertion of the raft of three strands and the removal of a strand from the existing wall (Goodell and Schwarz, 1983) would have to be very well regulated and substantially perfect. While the three-for-one model may, or may not, function in modern Gram-negative bacteria, it is difficult to imagine that it was the original mechanism for the first Gram-negative bacterium. So what are the possibilities for the first much simpler organism? By the time that the Domain of Bacteria split to originate Gram-negative organisms, both allosteric enzymes and aggregates of particular proteins to form holoenzymes must have been well developed and common in the intermediary metabolism of the cell. As distinct from their roles in Gram-positive cells, the linking enzymes and the cleavage enzymes must act in concert and act locally. Growth by addition of disaccharide pentapeptides to oligopeptidoglycan chains and then cross-linking the peptides on the inside of the wall and autolysin (Koch, 1988d) on the outside would not work satisfactorily in the Gram-negative case. What would be the difficulties encountered if besides addition of disaccharide units to the ends of available chains and cross-linking of peptides, another process also took place'? This would be the action of a holoenzyme consisting of a transpeptidase that could form trimers and an autolysin that resolved the trimer by cleaving the original cross-bridging bond. This would cleave some oligoglycan chains and create others probably not aligned with

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the chain with which it was now connected. The other end would be capable of growing and its peptide capable of linking with other chains. If this were sufficiently common then the wall would quickly become covalently intact but have been enlarged. I hope that in a few years we will know whether this is the way it happens in present-day organisms.

7. CONCLUSIONS 7.1. The Model for Evolutionary Creative Breakthroughs

The major mechanism for most steps of evolution is to start with a gene that carries out a useful function. A better variant then arises in time and is selected for; this process repeated many times generates the ‘best’ possible function given historical as well as physical, chemical and biological limitations. I have pointed out that there is a faster process which can work if conditions change so that an enzyme is temporarily not required. The gene can then become silenced but when conditions change back, the mutated gene may later be reactivated in a much improved form (Koch, 1972). A second type of process occurs when the gene is duplicated and one copy proceeds as above and the other is modified to serve a new function. This requires two kinds of change. First, duplication, and then the random change which is able to fill a vacant job opportunity that is useful and will be selected for. This is the well-established dogma of biology. In this review, I have considered processes beyond these two kinds and emphasized the basis of truly major steps forward, whether they be called macromutations or saltations. Quite often these advances depend on combining a number of existing, but quite different, functional systems and utilizing them for a new purpose by meshing them together. Developments of this above-mentioned kind are the creation of the First Cell, the development of the (external) sacculus, the development of methanogenesis, the development of oxygen-producing photosynthesis, and the production of cytoskeletons and mechanoenzymes to allow the eukaryote strategy to function. These processes were all generated by the same kind of fitting together of elements that already existed to form a new system which exhibits qualitatively different abilities and serves entirely different purposes than the individual component processes. As a pertinent example of this creative process, I will cite not an aspect of biological evolution, but the solution to the structure of DNA. No question about it, Watson and Crick (1953) carried out the creative synthesis, but needed intellectual resources. Thus, Crick had learned about cryptograms during the war, while Watson had been part of the phage group and was

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imbued with the idea (or hope) that DNA was the genetic material. Crick’s previous studies on X-ray crystallography had generated a theorem that identified patterns due to helical structures. Wilkins and Franklin had done the work. Donahue knew the Caltech scene and the X-ray work and quantum mechanical work going on there. His key information made it obvious that the stable tautomer of guanine was the keto and not the enol form (as one might have expected from the organic chemistry of the time). Add to this list Chargaff’s analytical measures that showed that the amount of adenine equaled that of thymine and the amount of guanine equaled that of cytosine. It was the sum of all these that generated the structure of the double-stranded helix of antiparallel chains. None of the items mentioned in the above list was aimed at understanding coding of proteins nor the replication of genetic information, but they were all components of the new breakthrough. 7.2. The Scenario for the Formation of the Three Domains

The scenario suggested here has many threads from bacteriology, contemporary biology, ecology and evolutionary theory. It is speculative and its one virtue is that it tries to fit processes as much as possible that do not require special (divine) intervention for their development. To explain the monophyletic portion as well as the generation of diversity I have tried to hold to ecological principles, especially Gause’s exclusion principle. I have argued that the paucity of metabolic energy was an important factor in evolution up to the time of the Last Universal Ancestor and that diversity resulted when the cell became successful and when new energy-trapping mechanisms arose. The result of the major advances considered here was an increase in life, both in numbers and in forms. This diversity, once stable, had a life of its own, and diversity of life created new niches and new habitats for exploitation and the development of still new diversity. 7.3. The Bacterial Way of Life

This review has the purpose of illustrating key principles to the evolution of life and illustrates it with the emphasis on the question of the development of the bacterial sacculus. So many issues had to be dealt with; each is complex. They had to be treated consequently from deductions of our knowledge of the nature of modern prokaryotes. The original bacterium was successful because its sacculus could resist the stresses of high internal osmotic pressure. In a world that was becoming diversified, it is not surprising that the diversity of Bacteria would expand

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tremendously with the development of new niches and habitats. The generation of the photosynthetic process that may have led to the branch of organisms called the purple bacteria now contains many non-photosynthetic forms. Is this a misnomer? Or did the ancestor of this branch develop because a synthesis of the anoxygenic photosynthetic process opened new doors for a radiation and now many of the branches have lost all signs of the facility to carry out photosynthesis? Maybe this apparently complete loss was feasible because all the genes involved in photosynthesis had become linked together and so a region could easily be deleted. This might happen in conjunction with the circumstances that other energy sources (particularly heterotrophic respiration) became more reliable and were less of a liability than having a working anaerobic photosynthesis system when it is not in the light. Similarly, the ability to carry out methanogenesis may have ceased to be an advantage when reducing substrates became more rare and destructive oxygen more generally available.

ACKNOWLEDGEMENTS Ian Burdett, Stephen Cooper. Paul Demchick, Ron Doyle, Frank Harold, George Hegeman, Michael Higgins, Joachim-Volker Holtje, Aaron Novick, Miguel de Pedro, Alan Wheals and Steve Woeste were all essential for this synthesis of disparate ideas.

Blaut, M., Muller, V. and Gottschalk, G. (1990) Energetics of methanogens. In: Bacterial Energetics (T.A. Krulwich, ed.) pp. 405-537. Volume XI1 of The Bacteria (J.R. Sokatch and L.N. Ornston, eds), Academic Press, San Diego. Bryant. M.P., Wolin, E.A., Wolin. M.J. and Wolfe, R.S. (1967) Methanobacillus omeliunrki, a symbiotic association of two species of bacteria. Arch. Microhiol. 59, 20-32. Burdett, I.D.K. and Higgins. M.L. (1978) Studies of the pole assembly in Buci1lu.r subtilis as seen in central, longitudinal, thin sections of cells. J . Bacteriol. 133, 959-971. Chang, S., Mack, R., Miller, S.L. and Stratheran, S.L. (1983) Prebiotic organic syntheses and the origin of life. In: Earth’s earliest Biosphere: Its origin and evolution (J.W. Schopf et al., eds), pp. 53-92. Princeton University Press. Princeton. Cole, R.M. and Hahn. J.J. (1962) Cell wall replication in Streptococcus pyrogcncs: immunofluorescent methods applied during growth show that new wall is formed equatorially. Science 125, 722-724. Cooper. S. (1991) Bactc,rial Growth und Division. Academic Press, Inc., San Diego, CA. Deamer, D.W. (1997) The first living system: a bioenergetic perspective. Microhiol. Mol. Biol. Rev. 61, 239-261.

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Koch, A.L. (1984a) Evolution vs. the number of gene copies per primitive cell. J. Mol. Ev01. 20, 71-76. Koch, A.L. (1984b) How bacteria get their shapes, the surface stress theory. Com. Mol. Cell Biophys. 2, 179-196. Koch, A.L. (l985a) Primeval cells; possible energy-generating and cell-division mechanisms. J. Mol. Evol. 21, 27k277. Koch, A.L. (1985b) How bacteria grow and divide in spite of internal hydrostatic pressure. Cun. J. Microbiol. 31, 1071-1084. Koch, A.L. (1988a) Biophysics of bacterial wall viewed as a stress-bearing fabric. Microbiol. Rev. 52, 337-353. Koch. A.L. (1988b) Speculations on the growth strategy of prosthecate bacteria. Can. J. Microhiol. 24, 39C394. Koch, A.L. (1988~)Partition of autolysins between the medium, the internal part of the wall, and the surface of the wall of Gram-positive rods. J. Theor. Biol. 134, 463472. Koch, A.L. (1989) The origin of the rotation of one end of a cell relative to the other end during the growth of Gram-positive rods. J. Theor. Biol. 141, 391402. Koch A.L. (1990a) Recent extensions of the surface stress theory. In: Microbial Growth Dynamics (R.K. Poole, M.J. Bazin and C.W. Keevil, eds). pp. 39-64. Oxford University Press, Oxford. Koch, A.L. (1990b) Additional arguments for the key role of ‘smart’ autolysins in the enlargement of the wall of Gram-negative bacteria. Res. Microhiol. 141, 529-541, Koch, A.L. (1990~)Growth and form of the bacterial cell wall. Amer. Sci.78. 327-341. Koch, A.L. (1993) Current status of the surface stress theory. In: Bacterial Growth and Lysis. Metuholistn and Stritciure ofthe Bacterial Succulus (M.A. de Pedro, J.-V. Holtje and W. Loffelhardt, eds) pp. 4 2 7 4 3 . Plenum Press, New York. Koch. A.L. (1994) Development and diversification of the Last Universal Ancestor. J. Theor. Biol. 168, 269-280. Koch, A.L. (1995a) Origin of intracellular and intercellular pathogens. Q. Rev. Biol. 70, 423 4 3 7 . Koch, A.L. (1995b) Bucreriul Growth and Form. Chapman & Hall, New York. Koch, A.L. and Burdett, I.D.J. (1984) The Variable-T model for Gram-negative morphology. J. Gen. Microbiol. 130, 2325-2338. Koch, A.L. and Burdett, I.D.J. (1986a) Normal pole formation during total inhibition of wall synthesis. J. G m . Microbiol. 132, 344-3449. Koch, A.L. and Burdett, I.D.J. (1986b) Biophysics of pole formation of Gram-positive rods. J. Gen. Microbiol. 132, 3451-3457. Koch, A.L. and Doyle, R.J. (1985) Mechanism of inside-to-outside growth and turnover of the wall of Gram-positive rod. J. Theor. Biol. 117, 137-157. Koch, A.L. and Doyle, R.J. (1986) The growth strategy of the Gram-positive rod. FEMS Microbiol. Rev. 32, 247-254. Koch. A.L. and Holtje, J.-V. (1995) A physical model for the precise location of the division site during growth of rod-shaped bacterial cells. Microbiology 141, 317 1-3 180. Koch, A.L. and Schmidt, T. (1991) The first cellular bioenergetic process; primitive generation of a protonmotive force. J. Mol. Evol. 33, 297-305. Koch, A.L. and Woeste. S.W. (1992) The elasticity of the sacculus of Escherichiu coli. J. Bucteriol. 174, 48 I 1 4 8 19. Koch, A.L. and Woldringh, C.L. (1995) The metabolic inertness of the poles of a Gramnegative rod. J. Theor. Biol. 171, 415425. Koch. A.L., Higgins. M.L. and Doyle, R.J. (1981a) Surface tension-like forces determine bacterial shapes, Strepptococcus,faecium. J. Gen. Microbiol. 123, 151-161.

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Koch, A.L.. Mobley, H.L.T., Doyle. R.J and Streips, U.N. (1981b) The coupling of wall growth and chromosome replication in Gram-positive rods. FEMS Microhiol. Li,rr. 12.201-208. Koch, A.L., Higgins, M.L. and Doyle, R.J. (1982a) The role of surface stress in the morphology of microbes. J . Gen. Microhiol. 128,927-945. Koch, A.L., Venver. R.W.H. and Nanninga, N. (1982b) Incorporation of diaminopinielic acid into the old poles of Escherichiu coli. J . Gen. Microhiol. 128. 2893 -2898. Koch, A.L., Kirchner. G., Doyle, R.J. and Burdett, I.D.J. (1985) How does a BucYl1u.r split its septum right down the middle? Ann. Inst. Pu.sreur/Microhii~l.136A. 91 98. Elsevier, Amsterdam. Kraft, A. ( 1997) Zuckerspaltenende Enzyme im Murenstoffwechsel von Eschi~richiowli. PhD dissertation, University of Tubingen, Germany. Labischinski, H. and Maidhof. H. (1994) Bacterial peptidoglycan: overview and evolving concepts. In: Bucteriul Cell Wul1.s (J.-M. Ghuysen and R. Hakenbeck. eds). pp. 23 38. Elsevier, Amsterdam. Labischinski, H., Barnickel, G., Bradaczek, H. and Giesbrecht, P. (1979) On the secondary and tertiary structure of murein. Eur. J . Biochrrn. 95, 147 155. Labischinski, H., Goodell. E.W., Goodell, A. and Hochberg, M.L. (1991) Direct proof of' a 'more-than-single-layered' peptidoglycan architecture of Eschivkhiu coli W7, a neutron small-angle scattering study. J . Bucreriol. 173, 751-756. LaPlace, P.S. ( 1 806) Mecanique Celeste. lmprimeur Imperiale, Paris. Matsuhashi, M. (1994) Utilization of lipid-linked precursors and the formation of peptidoglycan in the process of cell growth and division: membrane enzymes involved in the final steps of peptidoglycan synthesis and the mechanism of their regulation. In: Buereriirl Cell Wall (J.-M. Ghuysen and R. Hakenbeck, eds). pp. 5 5 ~ ~ 7 1Elsevier, . Amsterdam. Merad, T., Archibald, A.R., Hancock, I.C., Harwood, C.R. and Hobot, J.A. (1989) Cell wall assembly in Bucillus suhrilis, visualisation of old and new material by electron microscopic examination of samples selectively stained for teichoic acid and teichuronic acid. J . G m . Microhiol. 135,645S655. Miller, S.L. and Orgel, L.E. (1973) The Origins o f l i f e on Eurrh. Prentice-Hall, New York. Mobley, H.L.T., Koch. A.L., Doyle, R.J. and Streips, U.N. (1984) Insertion and fate of cell wall in Bucillus suhtilis. J . Bacteriol. 158, 169--179. Morowitz, H. ( 1992) Beginnings of Cellulur Life: Metuholism Recupirulutes Biogetiesis. Yale University Press, New Haven. Ogden, G.B., Pratt, M.J. and Schaechter, M. (1988) The replicative origins of the E. coli chromosome binds to cell membranes only when hemimethylated. Cell 154, 127- 135. Oparin, A.I. (1936) The Origin o f L ( f t . Dover, New York. Orgel, L.E. (1987) Evolution of the genetic apparatus: a review. Cold Spring tfurhor S.ytnp. Quunr. Biol. 52, 9-16. Paula, S., Vilkov. A.G., Van Hock, A.N., Haines, T.H. and Denmer, D.W. (1996) Permeation of protons, potassium ions and small polar molecules through phospholipid hilayers and as function of membrane thickness. Biophphys. J . 70. 339-348. Queller, D.C. (1995) The spaniels a t St. Marks and the panglossinn paradox: a critique of a rhetorical programme. Q. Rev. B i d . 70, 485 489. Shockman, G.D. and Holtje, J.-V. (1994) Microbial peptidoglycan (murein) hydrolases. In: Buereriul Cell Wall (J.-M. Ghuysen and R. Hakenbeck, eds). pp. 131 166. Elsevier, Amsterdam. Spratt. B.G., Bowler, L.D., Edelman. A. and Broorne-Smith, J.K. (1988) Membrane topology of penicillin-binding proteins I h and 3 of Escherichia coli and the production of water-soluble forms of high-molecular weight penicillin-binding proteins. In:

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Energetics of Alkaliphilic Bacillus Species: Physiology and Molecules Terry Ann Krulwich', Masahiro Ito', Raymond Gilmour', David B. Hicks' and Arthur A. Guffanti' 'Department of' Biochemistry. Mount Sinai School of' Medicine of CUN Y . New York, New York. USA 'Department of Life Sciences, Tokyo University, Gumna 374-01, Japan

ABSTRACT

The challenge of maintaining a cytoplasmic pH that is much lower than the external pH is central to the adaptation of extremely alkaliphilic Bacillus species to growth at pH values above 10. The success with which this challenge is met may set the upper limit of pH for growth in these bacteria, all of which also exhibit a low content of basic amino acids in proteins or protein segments that are exposed to the outside bulk phase liquid. The requirement for an active Na+-dependent cycle and possible roles of acidic cell wall components in alkaliphile pH homeostasis are reviewed. The gene loci that encode Na+/H+ antiporters that function in the active cycle are described and compared with the less Na+-specific homologues thus far found in non-alkaliphilic Gram-positive prokaryotes. Alkaliphilic Bacillus species carry out oxidative phosphorylation using an exclusively H+-coupled ATPase (synthase). Nonetheless, ATP synthesis is more rapid and reaches a higher phosphorylation potential at highly alkaline pH than at near-neutral pH even though the bulk electrochemical proton gradient across the coupling membrane is lower at highly alkaline pH. It is possible that some of the protons extruded by the respiratory chain are conveyed to the ATP synthase without first equilibrating with the external bulk phase. Mechanisms that might apply to oxidative phosphorylation in this type of extensively studied alkaliphile are reviewed, and note is made of the possibility of different kinds of solutions to the ADVANCES I N MICROBIAL PHYSIOLOGY VOL 40 ISBN 0- 12-027740-9

Copyright (c) 1998 Academic Press All rights of reproduction in any form reserved

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problem that may be found in new alkaliphilic bacteria that are yet to be isolated or characterized. 1. Introduction.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Energetics of pH homeostasis. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. The magnitude of the challenge . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Evidence that this process is directly related to the upper pH limit for

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growth . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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2.3. Overview of the active cycle necessary for pH homeostasis and

any other features that may support this function. . . . . . . . . . . . . . . . . . . 407 2.4. The antiporters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 410 2.5. Considerations and hypotheses with respect to Na+ re-entry . . . . . . . . . . 417 2.6. Glucose- and malate-grown cells have different extents of dependence upon a Na+-dependent active cycle. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 418 2.7. Outer cell wall layer involvement in glucose-grown cells . . . . . . . . 2.8. Some other possible mechanisms of defence against profoun change . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Energetics of oxidative phosphorylation . . . . . . . . . . . . . . . . . . . . . 3.1. Overview of the problem. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 421 3.2. The respiratory chain. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 423 3.3. The ATP synthase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 425 3.4. Possible models of alkaliphile oxidative phosphorylation . . . . . . . . . . . . . 427 Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 432 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 432

1. INTRODUCTION

Investigations of microbial diversity are motivated by both the lessons and the applications that may be anticipated to emerge from comparative studies. These expectations are enhanced with organisms that are now generally referred to as ‘extremophiles’, i.e. those microorganisms that thrive under conditions at which the vast majority of organisms would fail to grow or would even perish. Bacterial extremophiles have become the focus of greatly enhanced attention during the past few years. With the launching of projects that probe environments that were heretofore inaccessible, there has been an expansion of the kinds of extreme environments from which bacteria are isolated. With better tools for identifying organisms that cannot yet be cultured as well as those that can, there has also been a major expansion in the number of different organisms that have been catalogued in given types of extreme environments. Studies of individual genes and their protein products have begun, in some cases, to yield generalizations about adaptations that serve particular extremophiles. Furthermore, during this same period, direct comparisons of the genomes of extremophiles and non-extremophiles, including at least some among members of the same genus, have been initiated. The data from these comparisons should provide important

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new information about aspects of gene organization that promote adaptability to extreme environmental challenges. The genomics of extremophiles will also lead to new genes that play roles in extreme adaptability in general and in connection to specific kinds of extreme environments. Extremely alkaliphilic bacteria, like other extremophiles, have attracted attention because of the utility of their extracellular enzymes in a variety of industrial settings (Horikoshi, 1996). There is, moreover, the expectation that an understanding of the basis for the alkali-stability of those alkaliphile proteins and of the alkaliphile surface components that are exposed to the external pH will lead to cell and protein engineering possibilities of applied interest. One generalization about the alkaliphile proteins that are exposed to the external milieu is that they have a pronounced replacement of basic amino acids with neutral or acidic amino acids; this is particularly evident in polytopic membrane proteins whose extracellular hydrophilic segments exhibit a pronounced deviation from homologous segments of neutralophilic bacterial proteins (Van der Laan et al., 1991; Kang et al., 1992; Quirk et al., 1993; Ito et al., 1997b). Interestingly, it has been noted that over 70% of the proteins predicted from the genome sequence of Helicobacter pylori, a gastric pathogen that grows in a highly acidic environment, have a calculated isoelectric point above 7.0, with a much higher lysine and arginine content than homologues from neutralophilic bacteria (Tomb et al., 1997). Thus, it appears that at both extremes of pH, adapted extremophiles minimize the use in their proteins, and for alkaliphiles especially those exposed to the external milieu, of amino acids whose free acid or base groups might be subjected to pH changes in a range near their pK values. It is likely that there are general properties of the alkaliphile coupling membrane as well, e.g. the presence of neutral isoprenoid lipids and of a high content of branched-chain fatty acids in the membrane phospholipids (Clejan et al., 1986). However, the current state of comparative work on alkaliphile and neutralophile membranes is not yet sufficiently broad or sufficiently bolstered by genetic analyses to support specific hypotheses about alkaliphile membrane lipids. On the other hand, some but not all groups of alkaliphiles do appear to require negatively charged cell wallassociated polymers for growth at the high end of their pH range (Aono and Ohtani, 1990; Aono et ul., 1993, 1995), as will be discussed further because of a possible role of such polymers in cellular energetics. Alkaliphiles have also attracted attention because of general interest in the central physiological processes that are particularly challenged at highly alkaline pH, i.e. cytoplasmic pH regulation and bioenergetic work that depends upon an electrochemical proton gradient (Ap) whose energetically productive orientation is acid and positive out (Ivey et al., 1998). The specially adapted mechanisms that are used to meet central physiological challenges by extremophiles often illuminate the mechanisms used in the

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normal, less extreme setting. Thus, the intrinsically intriguing bioenergetic problems of life at high pH are of further interest because the alkaliphiles' solutions may affect more general paradigms about pH homeostasis and energy coupling phenomena. Alkaliphilic bacteria are diverse and are often adapted to multiple environmental challenges, e.g. either to high temperature or high Na' concentrations in addition to high pH (Grant et al. 1990). In order to isolate the response to high pH, it is important to work with alkaliphiles that are not specially adapted to optimal growth at some other extreme condition, i.e. do not require an extreme of temperature or salt concentration or light. In order to take optimal advantage of mutations to non-alkaliphily and slowly emerging genetic systems in alkaliphilic bacteria, it is also best to employ a strain that is capable of growth at near-neutral as well as extremely alkaline pH. Several alkaliphiles that meet both of these criteria, i.e. are not obligately extremophile in either their alkaliphily or with respect to some other environmental factor, have been studied extensively. They are all Bacillus species, including Bacillus ,firmus OF4 (Guffanti et al., 1986; Krulwich, 1995), Bacillus Ientus C-125 (Aono, 1995) and Bacillus YN-2000 (Yumoto et al., 1991; Koyama and Nosoh, 1995). This discussion will focus on the work conducted with such organisms. Such an emphasis will permit a detailed development of some of the major issues of alkaliphile energetics for this one type of alkaliphile, for which the most information is available. It is important to keep in mind, however, that future studies of other distinct genera of alkaliphiles, particularly those with strong adaptive pressures in addition to high pH, or of alkaliphiles among the archaebacteria, that may have significantly different membrane properties, could reveal very different solutions than those employed by the alkaliphilic Bacillus species.

2. ENERGETICS OF pH HOMEOSTASIS

2.1. The Magnitude of the Challenge

Two different questions are usually asked in order to assess the capacity of an alkaliphilic bacterium for pH homeostasis. First, what is the relationship between the cytoplasmic pH of energized cells and the external pH, i.e. how big is the pH gradient that is generated and sustained? Using cells growing logarithmically on malate in continuous culture at various, carefully controlled pH values, Sturr et al. (1994) showed that the cytoplasmic pH of B. firmus OF4 remained below pH 8.0 in cells growing at pH values of 9.5 and below, and remained below pH 8.5 at values of the growth pH up to about 10.6 (Fig. 1). That is, this alkaliphile exhibited values of the pH gradient

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PHin 200

700 10.0

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525

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125 8.0 100

350

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8.0 25

0

0 7.5

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PHout

Figure 1 Generation times and bioenergetic parameters of Bacillus.firmus OF4 growing in continuous cultures at different, controlled values of external pH. [Reproduced, with permission, from Sturr el al. (1994), which provides details of measurements made.]

(ApH) of 2-2.3 units, acid in, under optimal highly alkaline conditions on a non-fermentative carbon source. Similar values for the magnitude of the ApH were found in batch cultures of B. firmus OF4 growing on malate (Guffanti and Hicks, 1991). Glucose-grown cells of the same organism have recently been shown to exhibit ApH values of comparable magnitude (Gilmour and Krulwich, 1997). It is important to use energized cells rather than cells suspended in buffer. Washed cells in buffer exhibit a lower capacity for acidifying their cytoplasm relative to an alkaline external pH than comparable energized cells. This is why some of the earliest experimental determinations of cytoplasmic pH on alkaliphilic Bacillus species yielded values that were above 8.5 at an external pH of 10.5 (Guffanti et al., 1978). The assays used for most measurements of cytoplasmic pH conducted on alkaliphiles thus far involved calculation of the cytoplasmic pH from the directly measured external pH and the transmembrane ApH calculated from the distribution of a weak base and weak acid. Using an assay that takes

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advantage of a pH-sensitive fluorescent probe that can be loaded into live bacterial cells, 2’,7’-bis-(carboxymethyl)-5carboxyl-fluorescein (BCECF), Aono et al. (1997) have studied the cytoplasmic pH of B. lentus C-125 as a function of buffer pH in glucose-energized cells. In these studies the cytoplasmic pH remained below 8.0 up to an external pH of about 10.7 as long as the test cells had been grown at pH 10. The cytoplasmic pH values of extremely alkaliphilic Bacillus species found in the above studies at the alkaliphiles’ optimal external pH values for growth are not very different from values of cytoplasmic pH for optimally growing non-alkaliphiles (Guffanti and Hicks, 1991; Sturr et a]., 1994; Padan and Schuldiner, 1996; Aono et al., 1997). A rapid fall-off in the growth rate of alkaliphilic Bucillus species is observed when the cytoplasmic pH starts to rise above 8.5; at values of cytoplasmic pH nearing 9.5, growth of B..firmusOF4 is 20-fold slower than at optimal values of cytoplasmic pH, i.e. below pH 8.5 (Sturr er al., 1994). Quite likely, a cytoplasmic pH of about 9.5 is the upper edge that is compatible with growth for most, if not all, bacteria, including alkaliphiles. A bacterium that is apparently devoid of active mechanisms for pH homeostasis, e.g. Clostridium fervidus, has a narrow range of pH for growth that ends below pH 9.5 (Speelmans et al., 1993). There is no evidence thus Par for an alkaliphile that is enhanced in its capacity to survive at unusually high values of the cytoplasmic pH. Nor are the most sensitive cytoplasmic processes or molecules yet known, i.e. molecules that might especially limit growth at the upper edge of cytoplasmic pH; quite possibly, too many processes become limiting at pH 9.5 for adaptations to higher cytoplasmic pH to be a feasible route to alkaliphily. Thus, for known alkaliphiles, a major feature in meeting the challenge of life at high pH is in the maintenance of a cytoplasmic pH that is little different from that of neutralophiles, while the external pH that is optimal for growth is in the range of pH I(b-11. Alkaliphiles may live in niches that are subject to large, rapid pH changes, requiring them to have protective adaptations for such transient events. The second question generally posed in assessment of the pH homeostasis capacity of an alkaliphile is whether it can retain a relatively constant cytoplasmic pH upon a sudden, large alkaline shift in pH (Kroll, 1990). Booth (1985) has suggested that a capacity to withstand a sudden change is a more true measure of homeostatic capacity, and work on Bacillus suhtilis has indicated that it is a more rigorous challenge than is the capacity to exhibit a particular cytoplasmic pH during logarithmic growth (Cheng et al.. 1996a). Such pH shift experiments have been conducted on B..firmus OF4 cells that were first equilibrated in buffer to which an energy source was added at pH 8.5. The pH of the suspension was then rapidly raised to 10.5 and the cytoplasmic pH was assayed 10 min after the shift. In these experiments, the alkaliphilic Bacillus was found capable of resisting any significant

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change in cytoplasmic pH in buffers containing sufficient Na+ and either malate or glucose to energize (Krulwich e f al., 1985b, 1994; Gilmour and Krulwich, 1997).

2.2. Evidence that this Process is Directly Related to the Upper pH Limit for Growth

A major category of mutants to non-alkaliphily are mutants that have lost the capacity to acidify their cytoplasm relative to the external medium during a pH shift experiment (Koyama et al., 1986; Kitada ef al., 1989; Krulwich ct al., 1996). Additional mutations to non-alkaliphily involve the primary energy production mechanisms of the respiratory chain that are coupled to the active mechanisms for pH homeostasis discussed below (Aono et a/., 1996; Krulwich ef al., 1996). This is consistent with the expectation that pH homeostasis is a critical feature of life at extremely alkaline pH. That this capacity is limiting with respect to the upper pH limit for growth is indicated by the continuous culture experiments on B.firmus OF4 in which the fall-off in growth rate paralleled the rise in cytoplasmic pH as the external pH increased above about pH 10.7. That is, the diminishing growth rate directly reflected the failure of the pH homeostatic mechanism to achieve a ApH above about 2.3 (Sturr et al., 1994; Krulwich, 1995). Other bioenergetic parameters correlated much less well with the diminishing growth rate that was observed above an external pH of 10.7 (Kruiwich et al., 1997).

2.3. Overview of the Active Cycle necessary for pH Homeostasis and other features that may support this function

The cartoon in Fig. 2 illustrates the components that are currently considered definite or likely parts of interlocking Na+ and H+ cycles that allow respiring alkaliphiles that extrude H+ during respiration to accumulate H+ and maintain a cytoplasmic pH that is more acidic than the external pH. The H+ pathway involves primary extrusion by the respiratory chain. Respiration establishes an initial, conventionally oriented pH gradient (ApH, acid out), and transmembrane electrical potential (A*, positive out) (Mitchell, 1961). Net proton accumulation is then achieved by rapid, secondary, electrogenic exchange of protons from the external medium for cytoplasmic Na'. A group of highly active Na+/H+ antiporters is used for this exchange. The exchange catalyzed by the antiporters that produce net proton accumulation must be energized by the A W and hence the exchange must occur with a H+/

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T.A. KRULWICH, M. ITO, R. GILMOUR, D.B. HICKS AND A.A. GUFFANTI

-0ocooc-

Acidic well layers needed for glucose-

-qloplasrnic buffer 3 - metabdc ecids?

omwn cells all-alkallphiles? maIal6-gmw?

the,

mP

ha

Figure 2 A representation of the active H+ and Na' cycles that together may constitute the active mechanism for net acidification of the cytoplasm in respiring alkaliphilic Bocillus species. The active pH homeostatic mechanism is shown as including: respiration. a group of three or more Na+/H+ antiporters (Nha), and Na+ re-entry pathways. R =the H'-extruding respiratory chain (electron donors/acceptor not shown). The Na+/H+ antiporters (Nha) include NhaC, an unknown designated as NhaN that could represent the Mrp-antiporter, and additional unknown antiporters. The antiporters are probably more Na+-specific than those involved in p H homeostasis in neutralophilic bacteria and they may have other special features as indicated by @ interacting with NhaC. The Na' reentry pathways are shown as a channel associated with the flagellum and two representative Na+/solute symporters. Other participants that have been suggested to function in the establishment and/or maintenance of an acidified cytoplasm relative to the medium are indicated and are discussed in the text.

Na' ratio > 1, i.e. is an electrogenic exchange that catalyzes inward movement of positive charge as protons are accumulated (McNab and Castle, 1987). The Na' cycle that is part of the active homeostatic mechanism includes the outward movement of Na' through the antiporters and mechanisms that complete the cycle by catalyzing re-entry of Na'. In Fig. 2, the reentry mechanisms are shown as a variety of Na'lsolute symporters and Na' channel(s) such as the one associated with the Na' flagellar rotation that is exhibited by alkaliphilic bacilli (Hirota and Imae, 1983; Atsumi et al., 1990). Each of the parts of this active mechanism for pH homeostasis will be discussed in greater detail below. Also considered will be additional features of alkaliphiles that could play important roles in pH homeostasis at least under some growth conditions or in instances of sudden upward pH changes, i.e. before induction of the full antiporter complement of high pH-grown cells.

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Alkaliphilic bacilli can extrude H+ via the proton extrusion-coupled hydrolysis of ATP by the F, Fo-ATPase under more fermentative conditions at which respiration and its associated H+ extrusion might be insufficient to support requisite antiporter activity. However, respiration may be considered as an essential part of the interlocking Na' and H' cycles for aerobically grown alkaliphilic Bacillus strains growing on non-fermentable carbon sources. Respiration is indicated as an aggregate that results in H+ extrusion in Fig. 2. For extremely alkaliphilic, non-marine, Bacillus species, there is substantial and compelling evidence for respirationcoupled H' extrusion and no evidence to date for primary, respirationcoupled Na+ extrusion systems. Such primary Na+ extrusion systems have been found or proposed for several marine bacteria (Tokuda and Unemoto, 1984; Unemoto and Hayashi, 1993; Beattie et al., 1994; Skulachev, 1994; Pfenninger-Li et al., 1996) as well as for Vitreoscilla (Park et al., 1996) and are often first detected as Na' extrusion activities that are stimulated by reagents that abolish or reduce the AW. The stimulatory effect is understood to release the rate of ion extrusion from the constraint of the back-pressure constituted by the transmembrane potential that develops. By contrast, under normal cultivation conditions for B. firmus RAB (an obligately alkaliphilic strain related to strain OF4) (Guffanti et al., 1986), all Na' extrusion was found to be inhibited by treatment of the respiring cells with agents that collapse the AW. Such inhibition would be anticipated for secondary, electrogenic antiporters whose energization depends upon the A W . Thus, cells grown at pH 10.5, energized with malate under highly aerobic conditions with moderate concentrations of Na' in the medium (e.g. 25-200 mM) exhibit no Na' extrusion in the presence of valinomycin and sufficient K+ to abolish any transmembrane potential that is developed during respiration (Krulwich and Guffanti, 1989). Quite possibly, primary Na' pumping would be inhibitory to extremely alkaliphilic bacteria that grow under conditions in which the Na' concentration is not especially high, e.g. in some conventional soils (Guffanti et al., 1986). While potentially contributing to the capacity to develop a productive AW, such Na' extrusion without concomitant H+ uptake would reduce the cytoplasmic Na' concentration that must be available for the sustained, secondary antiporter activity that catalyzes net proton accumulation during respiration and sets the upper limit of pH for growth. In Fig. 2, the possibility is indicated that alkaliphiles may yet be shown to possess ABC-type primary transport systems for Na+ similar to the NatAB system of Bacillus subtilis (Cheng et al., 1997). If so, it would presumably be induced only under conditions of excess and even inhibitory Na' concentrations.

41 0

T.A. KRULWICH, M. ITO, R. GILMOUR, D.B. HICKS AND A.A. GUFFANTI

2.4. The Antiporters

Na' is required for generation and maintenance of a cytoplasmic pH that is lower than the external pH during alkaliphile growth on non-fermentative carbon sources at highly alkaline pH or upon an alkaline pH shift. K+ ions are unable to substitute for this function (Krulwich et ul., 1985b, 1994). This specific importance of Na' for pH homeostasis, in bacteria that do not generally show a requirement for high concentrations of N a f , is thus far a general feature of extreme alkaliphily even outside of the genus Bucillus (e.g. in E.uiguuhacterium uurunticum; McLaggan et ul., 1984). At least several alkaliphilic Bucillus species possess K f / H f antiporters whose activity can be shown in membrane vesicles (Mandel et ul., 1980; Kitada et ul., 1997). However, in the extreme alkaliphiles, there is no evidence that K f / H+ antiporters contribute to pH homeostasis. It is not yet known whether the K+/H+ antiport activity is catalyzed in alkaliphiles by transporters that are distinct molecular species from the Na+/H+ antiporters and that play a minor role or some different role. Na+/H+ antiporters might also have a low capacity for K+/Hf antiport activity that would be insufficient in itself to support pH homeostasis and would not occur in the presence of the requisite Na+ concentration. Resolution of this issue will await the clarification of the major antiporter proteins and their study in purified, reconstituted form. This has been done for two of the Na+/H+ antiporters from E. coli (Padan and Schuldiner, 1996) and one of the antiporters involved in pH honieostasis in B. suhtilis (Cheng et nl., 1996b) but not yet for an alkaliphile antiporter. Interestingly, the multifunctional TetA(L) antiporter of B. suhtilis catalyzes, in addition to tetracycline-divalent metal ion/H+ antiport, both Na+/H+ and Kf/H' antiport. The monovalent cation/H+ antiport supports cytoplasmic pH acidification and growth at pH 8.3 in the presence of 50-100 mM concentrations of either Na+ or Kf (Cheng et a/., 1994, 1996a). Consistently, either Na' or K+ can support maintenance of a cytoplasmic pH in B. suhtilis close to 7.5 upon an alkaline shift in the medium pH from 7.5 to 8.5 (Cheng et ul., 1994, 1996a). Thus, for the neutralophilic B. suhlilis and quite likely for other neutralophilic bacteria as well, pH homeostasis can be supported either by interlocking H+ and Naf cycles as described above for alkaliphiles or by antiport for H + using the abundant cytoplasmic K+, as long as there is an adequate Kf concentration in the medium for replenishment. Interestingly, alkaliphilic B. firrnus OF4 is Na+-specific for this capacity even when challenged in the same range of pH as used for the B. subrilis challenge. That is, upon a shift from pH 7.5 to 8.5 as well as in a shift from pH 8.5 to 10.5, alkaliphile cells exhibited cytoplasmic pH homeostasis in the presence of Na+ but not in the presence of Kf alone (Krulwich et ul., 1994).

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41 1

It is notable that under conditions that were at least reasonably comparable with respect to the available energetic driving force, the aggregate Na+/ H+ antiporter activity of B.firrnus OF4 was about 10-fold higher than that of B. suhrilis (Krulwich et a/., 1994). This led to the hypothesis that the specific requirement by alkaliphiles for Na+ to achieve pH homeostasis relates to the avoidance of the risk that too much K+ would be lost from the cytoplasm. Loss of K+ could create a suboptimal cytoplasmic milieu for enzyme activity and protein stabilization, upon use of K+ for the large amount of ongoing antiport required for pH homeostasis in respiring alkaliphile cells (Ito et al., 1997b). In any event, the Na+-specificity of the pH homeostasis mechanisms of alkaliphiles makes it likely that the cation specificity of the antiporters involved will be different from those of neutralophiles whose pH homeostasis mechanisms are less cation-specific. In instances, as will be described below, where there are apparent homologues involved in alkaliphiles and non-alkaliphiles, the basis for the cation preference will be of interest. As already noted, monovalent cation/H+ antiporters that catalyze net accumulation of protons during respiration are expected to be electrogenic, and evidence for such electrogenicity has been presented in various bacteria, including alkaliphiles (Garcia et ul., 1983; Bassilana et al., 1984; Cheng et al., 1996b). The stoichiometry of the antiport has been determined for two E. coli Na+/H+ antiporters thus far, where H+/Na+ stoichiometries of 2/1 and l.S/l were found for NhaA and NhaB, respectively (Padan and Schuldiner, 1996). Mechanistic stoichiometries have not yet been determined for any physiologically important alkaliphile antiporter. This will be of importance. The most detailed and controlled studies of the ApH suggested that a gradient of up to 2.3 units was generated, but no larger. The mechanistic stoichiometry is one of the parameters that will be needed to interpret this limitation. In extreme alkaliphiles, the available driving force, the total electrochemical proton gradient (Ap), for electrogenic antiporters that catalyze inward H+ movement may limit the net proton accumulation and could necessitate a high coupling ratio of Hf/Na+ for the antiporters used. Alternatively, or in addition, there may be special mechanisms in the alkaliphile to maximize the antiport that can occur. Finally, with respect to general considerations about the antiporters involved in pH homeostasis, it would be critical for alkaliphiles, as has been indicated for bacteria in general (Booth, 1985), to possess a significant constitutive complement of antiporters in order to adjust to sudden changes of pH. In addition, there may be changes in the Na' concentration available. For the Na+-specific homeostasis in alkaliphiles, these bacteria would need a number of antiporters that can function over different ranges of Na+ concentration. Both of these conditions appear to have been met although detailed information about the gene products involved is still incomplete.

41 2

T.A. KRULWICH, M. ITO, R. GILMOUR, D.B. HICKS AND A.A. GUFFANTI

2.4.1. nhaC The first gene encoding an alkaliphile Na+/H+ antiporter, the nhaC gene from E. jirmus OF4, was cloned and sequenced upon partial functional complementation of a Na+-sensitive mutant strain of E. cofi that was deleted in its major antiporter-encoding gene nhaA (Ivey et al., 1991). That the nhaC gene product catalyzes Na+/H+ antiport activity was demonstrated initially in membrane vesicle preparations from the complemented E. cofi strain. Interestingly, the original complementing fragment of DNA from the alkaliphile is a highly truncated form of the full-length gene and even the form first studied in the vesicle assays, and thought to be complete, is truncated significantly at the N-terminal end of the gene product. This leads to the likelihood that an appreciable part of the N-terminal part of NhaC is not required for at least minimal catalytic activity (Ito ef a f . , 1997a). Using approaches developed for other Gram-positive bacteria and other alkaliphilic Bacillus species (Biswas et af., 1993; Ito et af., 1994) single cross-over and double cross-over insertions into the nhaC locus of B. ,firmus OF4 were achieved, respectively facilitating an extension of the sequence analysis of the region and the construction of an alkaliphile strain in which the locus contained a deletion in the nhaC coding region. A diagram of the nhaC locus is shown in Fig. 3. Both predictions from the sequence and initial Northern analyses suggest that nhaC is part of a two-gene operon that also contains the small downstream gene, nhaS; the putative promoter region for the operon contains two direct repeat sequences (Ito et a f . , 1997a). These sequences also appear in the chromosome at several other places and could represent a mechanism for gene rearrangements involving the nhaC locus. It is also possible that the direct repeat sequences play a role in up-regulation of gene expression since clones containing both the repeat sequences and a significant part of the nhuC gene (e.g. pGCUP1 in Fig. 3) are toxic to E. cafi whereas clones containing either the repeat sequences alone (e.g. pDRS1) or a significant part of the nhaC gene without the repeat sequences (e.g. pJX5) are not (M. Ito, unpublished data). The nhaS gene product is predicted to be a small membrane-associated protein with sequence similarity to a region of the eukaryotic Na', K+-ATPase, as well as a small, highly basic region that could be a DNA- or RNA-binding domain. Thus, NhaS is a reasonable candidate for a cation or pH sensor that might also have direct gene regulatory functions (Ito et al., 1997a). Some sort of sensing device for low Ap and for high internal or external pH is anticipated; sensing devices, e.g. 'protometers' that sense Ap, have long been proposed (Glagolev, 1980). Another hypothetical possibility for a function of a small Na+-binding protein that is membrane-associated is that it concentrates Na+ in the binding site region of NhaC in such a way as artificially to increase the concentration of that substrate near the inner surface of that antiporter.

413

ENERGETICS OF ALKALlPHlLlC BAClLLUS SPECIES 0

or/A

1

2

3

4

5

6 kb

I

I

I

I

I

1

DRSl DRS2

nhaC

ScaI

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orp

or/c

orp

ScaI II

Figure 3 The nhaC region of the B.firrnus OF4 chromosome. Direct repeat sequences (DRSI and 2) that are 95% identical are shown upstream of an operon containing the antiporter-encoding nhaC and a small downstream gene, nhaS, that encodes a putative sensor/gene regulator. Several downstream genes have been sequenced but are not thought to be expressed coordinately with nhaCnhaS (Ito et a/., 1997a). The toxicity of the cloned regions indicated at the bottom of the figure is discussed in the text.

Conceivably, such a function might serve to maximize antiport as the protonic driving force dropped (see the speculation depicted in Fig. 2). Several genes from other bacteria are predicted to encode products with strong sequence similarity to that of NhaC but no function has been established for any of these genes. The functions of NhaC in B. firmus OF4 have been clarified in studies of the nhaC deletion strain N13 (It0 e f al., 1997a). Since N13 is able to grow well on both fermentable and non-fermentable carbon sources at pH 10.5, NhaC is clearly not the major Na+/H+ antiporter, nor even a necessary one under most culture conditions. However, studies of growth, response to a pH shift, and direct studies of Na' efflux support the following conclusions. NhaC is virtually the only Na+/H' antiporter in pH 7.5-grown cells that can carry out antiport at low concentrations of Na', i.e. 25 mM at pH 7.5. In pH 10.5-grown cells, NhaC is an important part of the complement of antiporters that functions at Na' concentrations of 10 mM and below, but additional high-affinity antiporters are present in the high pH-grown cells. Consistently, NhaC plays an important role when cells are shifted from pH 8.5 to pH 10.5, especially if the cells had been growing at pH 7.5 before the experiment. 2.4.2. mrp/pha Type Studies by Horikoshi and colleagues (Hamamoto et al., 1994; Hashimoto et al., 1994) in B. lentus C-125 were the first to use a non-alkaliphilic mutant

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T.A. KRULWICH, M . ITO, R. GILMOUR, D.B. HICKS AND A.A. GUFFANTI

strain of an alkaliphilic species as the host for cloning genes related to alkaliphily, presumably including antiporter-encoding genes. A pair of overlapping cloned fragments of B. lentus C-125 DNA were found to restore alkaliphily and Na+/H+ antiport activity to mutant strains of the alkaliphile that were non-alkaliphilic in growth phenotype and deficient in antiporter activity (Kudo et al., 1990; Hamamoto et al., 1994). The gene locus, not named by the investigators, was apparently an incomplete operon that contained at least three to four genes encoding hydrophobic products. The putative ORFl of the operon was the gene which complemented the nonalkaliphilic mutant by correcting a point mutation in the middle of the coding sequence. The investigators noted that the predicted gene product of ORFl was a large polytopic membrane protein with sequence similarity to membrane-embedded regions of NADH dehydrogenases and more modest sequence similarity to NhaC. They proposed that ORFl encoded an electrogenic Na+/H+ antiporter (Hamamoto et al., 1994) and that at least one of the downstream ORFs in the incomplete operon encompassed in their clones had a role in alkaliphily, albeit not directly in antiporter activity (Hashimoto et al., 1994). The ORFs found in the B. lentus C-125 region that was thus implicated in a major Na+/H+ antiporter activity, required for alkaliphily, are strongly homologous to ORFs found in a larger operon in Rhizohium meliloti (Putnoky el af., 1996) and in B. suhtilis (Ito et a)., 1998). These operons contain additional downstream ORFs such that the entire putative operon is apparently composed of seven genes, each of which is predicted to encode a membrane-associated product. The operon in R. meliloti was identified in nitrogen fixation-defective 'Fix-minus' mutants that were also K+-sensitive; it was proposed that the operon encoded a K+/H+ antiporter activity. Sequence similarity both to NADH dehydrogenases and to Na+/H+ antiporters was noted among the predicted products of the pha genes, a name given because of a suggested role in pH adaptation (Putnoky et al., 1996). The operon from B. suhtilis is strikingly similar to the pha operon, especially at the level of hydropathy predictions for the gene products (Fig. 4). This similarity extends to the presumably partial data currently available for two alkaliphiles (Fig. 4); the sequences of these alkaliphile operons are being completed currently. Studies on the B. subtilis operon utilized targeted disruptions of the first and sixth genes. Studies of growth at different pH values, cation sensitivities, pH homeostasis and bile salt resistance led to the suggestion that this operon might be even more multifunctional than thus far appreciated. Proposed functions included: Na+- as well as K+dependent pH homeostasis in a moderate range of cation concentrations; Na+-resistance over a broad range of pH values; and bile salt resistance. The likelihood of additional roles, yet to be clarified, was also noted in connection with poor growth of mutants in the locus on defined media. Because of

415

ENERGETICS OF ALKALlPHlLlC BAClLLUS SPECIES Bacillus subtilis mrp o p m n

Rhizobium melilotipha operon

I,

4 PhaA

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=1'

Bacillus sp.C-125 mrp operon

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MrpA

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Boci/llrrfnnvr OF4 mrp operon II

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I: Figure 4 The hydropathy plots of predicted rnrp or pha genes. Analyses of fragmentary data available for two alkaliphile mrp operons and complete sequence data for the pha operon of R . meliloti (GenBank accession no. X93358) and the rnrp operon of B. subtilis (GenBank accession no. 293937 and 293932). The data are presented as hydropathy plots according to the Kyte and Doolittle (1982) algorithm using a window of 19 residues.

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T.A. KRULWICH, M. ITO, R. GILMOUR, D.B. HICKS AND A.A. GUFFANTI

these findings, the operon from B. subtilis was called mrp, for the multiple resistance-pH involved operon (It0 et al., 1998). It is interesting to note that in R. meliloti, the pha operon may catalyze its antiport activity with a strong K+ preference whereas the B. suhtifis mrp-associated antiport activity is implicated in both Na' and Kf fluxes but with a preference for the former; the Na+-sensitivity of a B. subtilis strain with a disrupted mrpA is particularly striking (Ito et al., 1998). If the alkaliphile mrp operons are a major antiporter-encoding locus for pH homeostasis, it is anticipated that their Naf-specificity may be even greater (Krulwich et al., 1997). Moreover, it will be of great interest to elucidate the concentration range for Na' at which the mrp genes of the alkaliphile function. In B. subtilis, mrp deletion strains exhibited normal pH homeostasis in the high range of Na' concentrations at which TetA(L) is optimally functional as a monovalent cation/H+ antiporter, e.g. around 100 mM, whereas the mrp genes were important in a somewhat lower range. It is possible that the B. subtilis nhaC gene will have a role at even lower concentrations of Na', analogous to its role in the alkaliphile (It0 et al., 1997a). Perhaps the Mrp antiporter of alkaliphiles will have a somewhat higher range of Na' concentrations for its optimal activity, accounting for the observations in B. fentus C-125 that are suggestive of a necessary if not sufficient role in alkaliphily and pH homeostasis (Hamamoto el af., 1994). Another interesting and incompletely understood feature of the mrplphalike operons is the apparent need for the intact operon or almost all thereof for any and all of the activities thus far attributed to the operon (Ito el al., 1998). It will be of great interest to determine the basis for this interdependency of the gene products, be it a common requirement for one of the downstream genes for sensing, expression, or some cooperative phenomenon at the level of protein stabilization, insertion or catalysis. The interdependency might have the effect of coordinating a response to multiple stresses that often are experienced together, and the relevance of that consideration to alkaliphiles will have to be examined specifically as more data emerge. 2.4.3. Additional Antiporters are Anticipated The studies of the nhaC deletion strain of B.$rmus OF4, N13, indicated that there are at least three physiologically important Na+/H+ antiporters, and probably more. There is a constitutive dominant, low-affinity (high flux) antiporter that is active at both near-neutral and alkaline pH, or two antiporters that function optimally in one of the two pH ranges. The Mrp antiporter could be part of this constellation, e.g. could be the 'NhaN' in

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Fig. 2. There is another component of the low affinity antiporter complement that is inducible by, and functional at, high pH. Then, in addition to NhaC, there is a high pH-inducible, high-affinity antiporter. Similarly, studies with other alkaliphilic Bacillus species indicate the presence of multiple Na'/H+ antiporter species (Kitada and Horikoshi, 1992; Kitada et al., 1994). It is likely that a set of antiporters that are homologous to the napA-encoded antiporter of Enterococcus hirae (Waser et al., 1992) and Bacillus megaterium (Tani et al., 1996) will be part of this complement inasmuch as homologues of as yet undetermined function have been found in several bacteria, including B. subtilis (GenBank Accession no. Y14080), H. pylori (Tomb et al., 1997), and recently, in B. firmus OF4 (M. Ito, unpublished results).

2.5. Considerations and Hypotheses with Respect to Na+ Reentry

At moderate to high concentrations (e.g. > 25 mM) of Na', the re-entry of Na' may not be a major problem, although it could still be rate-limiting for the Na' cycling that is part of the alkaliphile pH homeostatic mechanism. Clearly, at low concentrations of Na', the re-entry of the cation is indeed rate-limiting as was shown over a decade ago. In those experiments (Krulwich et al., 1985b), the presence of a non-metabolizable amino acid analogue whose uptake was coupled to Na' uptake markedly improved pH homeostasis during an alkaline shift in the presence of sub-optimal concentrations of Na'. Since the uptake itself would be AWconsuming, and in that respect potentially competitive with antiport, the positive overall effect on pH homeostasis strongly indicated that, at low Na', the assistance given by the symport to cation re-entry was the dominant effect. It has been observed, however, that appreciable pH homeostasis during an alkaline shift is shown by alkaliphiles suspended in buffer without added solutes that enter with Na' (McLaggan et al., 1984; Krulwich et al., 1985b). This led to the suggestion that as the cytoplasmic pH rises, alkaliphiles might have mechanisms triggered to greatly increase Na' entry to support the antiport that is essential for pH homeostasis. As indicated in Fig. 2, one possibility is that the Na'lsolute symporters themselves might have a mode whereby they allow Na' influx in the absence of solute above some threshold cytoplasmic pH (Krulwich et al., 1997; Ivey et al., 1998). Another suggestion (McLaggan et al., 1984) was that there might be specific pHregulated Na' channels. A candidate for such a channel, as shown in Fig. 2, would be the channel associated with flagellar rotation. Sugiyama (1995) has calculated that Na' flux through that channel would indeed be competent to support antiport activity at high pH. It is further notable that

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motility, which is Na+-coupled in alkaliphilic Bacillus species (Hirota and Imae, 1983), is appreciable only in the most alkaline range of pH for alkaliphile growth (Aono ef al., 1992; Sturr et al., 1994).

2.6. Glucose- and Malate-grown Cells have Different Extents of Dependence upon a Na+-dependentActive Cycle

Upon a sudden shift in the external pH, to 10.5, of E . firmus OF4 cells grown on malate at pH 10.5 and then equilibrated at pH 8.5, there is excellent pH homeostasis as long as an energy source and Na' are both present. This has already been extensively discussed. However, if the same experiment is conducted with glucose-grown cells, using glucose as the energy source during and after the shift, then there is a significant component of the pH gradient, about one pH unit, that is found even in the absence of Na' (Gilmour and Krulwich, 1997). For technical reasons concerning non-inhibitory but effective buffers in the high pH range, it is difficult to eliminate K+ from the experiment at the same time that Na' is omitted. Thus, it is possible that K+/H' antiport takes over partially in glucose-grown cells. This seems less likely to us, however, than the possibility that other types of mechanisms play some role in glucose-grown cells but not in cells growing on non-fermentable carbon sources. This hypothesis will be testable as the full panoply of defences against sudden shifts in external pH are identified. Some of the possible defences and the data supporting them follow.

2.7. Outer Cell Wall Layer Involvement in Glucose-grown Cells Work by Aono and colleagues (Aono and Ohtani, 1990; Aono et al., 1993, 1995) on a variety of alkaliphilic Bacillus species, grown on glucose, has shown that some groups of alkaliphiles produce two acidic cell wall-associated polymers in high quantity, a teichuronic acid and a glutamate-rich teichuronopeptide. The presence of these polymers is indicated in the cartoon in Fig. 2. Mutational loss of these polymers leads to poor growth on glucose at highly alkaline pH (Aono and Ohtani, 1990; Aono et al., 1995). The investigators have suggested that alkaliphily in the parent strains of these mutants may depend upon a highly negatively charged cell surface to restrict entry of hydroxide ion. I t has not yet been shown whether this sort of passive barrier is also important under other growth conditions, e.g. during growth on non-fermentable carbon sources. Possibly, an initial barrier of this type is the basis for the difference between the extent of Na+-

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dependence of a pH homeostasis in glucose- versus malate-grown B. firmus OF4; in that context it should be noted that this alkaliphile does not possess significant or high pH-induced quantities of these polymers during growth on malate (Guffanti and Krulwich, 1994), but glucose-grown cells have never been examined. In any event, it is clear that the acidic polymers in those alkaliphiles where an important role has been found d o not replace the need for an active mechanism of H+ accumulation since single mutations that reduce energy-dependent antiporter activity produce a non-alkaliphilic phenotype (Hamamoto ef a/., 1994).

2.8. Some other Possible Mechanisms of Defence against Profound pH Change

As indicated in Fig. 2, there are other features of alkaliphile cells that may contribute, at least transiently, to pH homeostasis upon upward changes in pH. These include, for cells growing on fermentable carbon sources, the production of metabolic acids. In further connection with glucose growth, it is of interest that alkaliphilic B. firmus OF4 has been found to have homologues (GenBank accession no. U91841) of the motAB-like genes that are found in a catabolite repression locus of the B. suhtilis chromosome (Grundy et al., 1993); these genes are expected to encode a H+translocating channel structure. The alkaliphile region was cloned as a DNA fragment that weakly improved the growth of a non-alkaliphilic mutant of B. firmus OF4 at pH 10.5 (M. Ito, unpublished data). This finding raises the possibility that proton flux through a proton-translocating element could provide some useful, inward proton flux in fermenting cells that are generating a high transmembrane potential at moderately alkaline external pH. Additional features of alkaliphile cells that might have some role in pH homeostasis include the possibility that observed, high buffering capacity of the alkaliphile cytoplasm (Krulwich et al., 1985a) might offer some transient protection against a change in pH. Alternatively, the overall buffering capacity might reflect a high concentration of buffering protectants that surround particularly sensitive or important structures or molecules. Further, there may be specific membrane lipids required to produce a sufficiently ionimpermeable coupling membrane at high pH while also preserving optimal fluidity. Some polar isoprenoid lipids have been found in alkaliphilic Bacillus species and may have a modulatory role in ion permeability (Clejan et al., 1986; Clejan and Krulwich, 1988). Finally, the report of generally higher PI values for a large percentage of the proteins in H . pylori than in neutralophile homologues (Tomb et al., 1997) raises the question of whether alkaliphile cytoplasmic proteins might

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T.A. KRULWICH,

M. ITO, R. GILMOUR, D.B. HICKS AND A.A. GUFFANTI

have some modest, overall tendency to have lower PI values than comparable proteins from neutralophiles, so that they are less vulnerable to changes in charge upon transient upward changes in pH. As the sequences of more cytoplasmic enzymes from alkaliphiles are reported, such an analysis might be of interest.

3. ENERGETICS OF OXIDATIVE PHOSPHORYLATION

The chemiosmotic model for the energization of oxidative phosphorylation posits that the bulk Ap across the coupling membrane of mitochondria or respiring bacteria is the direct and complete source of energy for a variety of bioenergetic work functions (Mitchell, 1961). Figure 5 depicts several examples of bioenergetic work that can be energized by the Ap that is established by primary H+ extrusion from a bacterial cell during respiration. The work processes shown in the figure include antiporters and

out

Resplration generates a ApH, acid out, and AY, positive out, via primary proton translocation that accompanier electron transport ApH and AY comprise the Ap, the electrochemlcal proton gradient

0 Bioenergetic work can be energized

by ApH andlor AY 0 Predict: Work functlonr vary directly with magnitude of Ap

Artiflciaily imposed gradienta should substitute for respiration-generatedones

Figure 5 Chemiosmotic energization of bioenergetic work: ion-coupled symport, antiport and FIFOATPase-mediated ATP synthesis.

ENERGETICS OF ALKALlPHlLlC BAClLLUS SPECIES

42 1

cation/solute symporters as well as the proton-translocating F, Fo-ATPase (synthase) that catalyzes ATP synthesis. As indicated, the bioenergetic work processes that are coupled to one or both components of respiration-generated Ap should show dependence on the magnitude of the Ap, in some instances above a particular threshold value. In addition, artificially imposed gradients should be as effective as respiration-generated ones of the same orientation and magnitude if they are of sufficient duration.

3.1. Overview of the Problem

The data in Fig. 1 illustrate a consistently observed divergence of the rate and extent of ATP synthesis via alkaliphile oxidative phosphorylation from the anticipated, direct quantitative relationship to the magnitude of the putative driving force, the bulk Ap. As shown in the data taken from carefully pH-controlled continuous cultures of B. finnus OF4, the Ap drops substantially between pH 7.5 and 9.5 as a large ApH is generated to keep the cytoplasmic pH in the optimal range; there is a partially compensatory increase in the AW over this pH range but the fall in the Ap is nonetheless appreciable and continues as the growth pH is raised to pH 10.7 (Sturr et af., 1994). The phosphorylation potential, AGp, which reflects the [ATPI/ [ADP][Pi] ratio, would be expected to show the same general pattern as the Ap, at least qualitatively if not perfectly quantitatively. By contrast, over the same range of increasing pH for growth in which the Ap falls, the AGp rises (Fig. I). In highly buffered batch cultures of B.$rmus OF4 grown at pH 7.5 versus 10.5, ATP synthesis was also at least as robust at the higher pH, as reflected by phosphorylation potentials, although the Ap was about 3-fold higher during growth at the lower pH (Guffanti and Hicks, 1991). Moreover, in both the batch cultures and continuous cultures, the molar growth yields calculated in the malate-containing medium were as high or higher at pH 10.5 than at pH 7.5, indicating that the ATP synthesis was not simply supported by a much higher rate of growth substrate consumption (Guffanti and Hicks, 1991; Sturr et af., 1994). Extensive work has been done to validate the methods used for determining the Ap and AGp values, as has been discussed in detail elsewhere (Guffanti and Krulwich, 1994; Ivey et af., 1998). So far, methods in which direct probes have replaced more indirect probing methods, have supported the conclusion that the total Ap at pH values above 10 is significantly lower, probably at least 3-fold, than at pH 7.5 (e.g. Aono et a f . , 1997). One divergent study was conducted on B. alcafophifus (Hoffmann and Dimroth, 1991b). in which the Ap parameters were monitored in batch cultures whose pH was allowed to decline over time from an initial value above 10 to near neutral. The values of the reversed ApH, measured on separate

422

T.A. KRULWICH, M. ITO, R. GILMOUR, D.B. HICKS AND A.A. GUFFANTI

suspensions prepared at various intervals, were slightly lower than generally found. Using unconventionally low probe concentrations (that may have failed to saturate the binding component), the investigators also reported somewhat higher values for the AW than have been reported by several groups working with logarithmically growing cells and more conventional measurements. Notwithstanding those differences, ATP synthesis was highest at the initial high pH at which the Ap was lowest (Hoffmann and Dimroth, 1991b). The inverse correlation between optimal ATP synthesis and the magnitude of the Ap was thus preserved and requires some accommodation with respect to proposed energetics. One possible accommodation would be the use of a different mechanistic coupling of protons to ATP synthesis, as was suggested by Hoffmann and Dimroth (1991b). Using values of the Ap that have most often been reported, in experiments with cells that are in fast, logarithmic growth at either near-neutral or at the extreme alkaline end of their range, the H+/ ATP ratio would have to vary from between 3 and 4 up to about 13 (Krulwich, 1995) in order to resolve the energy-coupling issue in this manner. That is. a much larger requirement for inward translocation of protons at the external surface would obtain at pH 10.5 than at pH 7.5. An independent line of experiments suggests that this sort of flexible stoichiometry capacity is not the one that alkaliphiles adopt. Were a variable coupling stoichiometry to account for the increasing efficacy of a decreasing Ap in energizing ATP synthesis, an imposed transmembrane electrical potential of about the same magnitude as that generated by respiration should support synthesis at the same coupling stoichiometry that occurs at any given pH during respiration. On the contrary, experiments indicate that artificially imposed, valinomycin-mediated K+ diffusion potentials energize both ioncoupled transport and ATP synthesis at external pH values below about 9.2 but do not energize ATP synthesis above this pH, although the imposed gradients are still competent in energizing transport (Guffanti et al., 1984; Guffanti and Krulwich, 1992). Control experiments showed the following. Electron donors energize the same cells that failed to respond to the diffusion potential, so that the conditions of the experiment had not inhibited ATP synthetic capacity in some global way. The ATP synthesized once electron donors are added, and the diffusion potentials that can be generated at high pH, decayed no faster than at low pH. The inefficacy of the diffusion potential at high pH thus could not be explained by fast decay of the potential or fast decay of transiently made ATP (Guffanti and Krulwich, 1994).These experiments suggest that the path of protons that are generated during respiration, or some other aspect of this energization, is not fully equivalent to the response to an imposed potential of the same measured magnitude but generated as a K+ diffusion potential.

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3.2. The Respiratory Chain

Evidence that respiration-dependent Na+-extrusion is entirely sensitive to abolition of the A Q under routine growth conditions at very alkaline pH, has already been cited (Krulwich and Guffanti, 1989). Those observations support the conclusion that primary Na+-extrusion is not a normal function of the respiratory chain of extreme, non-marine alkaliphiles. Rather, direct evidence from several lines of experiments that have recently been reviewed (Hicks and Krulwich, 1995) show that there is primary H+-extrusion by the alkaliphile respiratory chain. It might be expected that the energetic costs of growth of extremely alkaliphilic Bacillus species (e.g. pH homeostasis) would force cells growing on non-fermentative carbon sources to maximize the energy conservation that can be achieved by H+ during respiration. It is not yet clear whether this is the case, inasmuch as the evidence for a protontranslocating complex I is equivocal albeit suggestive that B. firmus OF4 has both an energy-conserving ndh-I and non-energy-coupled (ndh-11) type of NADH dehydrogenase (Hicks and Krulwich, 1995). Evidence for a Complex I11 is compelling (Lewis et al., 1981; Riedel et al., 1993), but a complete characterization in purified form has yet to be achieved for an alkaliphile complex and the genes are also yet to be characterized. Other respiratory chain complexes that have been more completely characterized tend to be present in the membranes in high concentration, sometimes in a high pH-induced manner (Lewis et al., 1981; Qureshi et a/., 1990, 1996; Yumoto et al., 1991,1993; Quirk et al., 1993; Hicks and Krulwich, 1995; Gilmour and Krulwich, 1996). There is also an increase in the catalase content of high pH-grown alkaliphile cells (Yumoto et al., 1990; Hicks, 1995). As shown in Fig. 6, however, current evidence indicates no special overall complexity or elaborate terminal branching of the respiratory chain of a typical alkaliphilic Bacillus. Both of the alkaliphilic Bacillus species whose respiratory chains have been studied extensively have been found to have a single major, high pHinduced terminal oxidase. In B. firmus OF4 that oxidase is a caa3-type oxidase (Quirk et al., 1993); disruption of the cta operon encoding this complex in B. ,firmus OF4 shows that, unlike B. subtilis (Lauraeus et al., 1991), the alkaliphile does not possess a second heme A-containing oxidase (Gilmour and Krulwich, 1997). On the other hand, the alkaliphile does possess a cytochrome bd-type terminal oxidase that is generally expressed only during the stationary phase in high pH-grown cells (Hicks et al., 1991). The bd-type cytochrome oxidase has been purified from the cta-disrupted strain of B. firmus OF4 (Gilmour and Krulwich, 1997). It is catalytically active as a quinol oxidase and has subunit molecular weights similar to those found for the enzyme from B. stearothermophilus (Sakamoto et al., 1996). The bd-type oxidase of B.firmus OF4, while elevated upon disruption of the

424

T.A. KRULWICH, M. ITO, R. GILMOUR, D.B. HICKS AND A.A. GUFFANTI

h c t C d d klpintol'y ChdM

Schematic illustration of major complexes of the respiratory chains of E. (Garcia-Horsman c't a/., 1994). Purucoccus denitr~ficans(Richter et a / . , 1994; DeGier et al., 1996) and B..firrnu.s OF4 (Hicks and Krulwich, 1995). The quinone of the alkaliphile is menaquinone (Hicks and Krulwich, 1995). Figure 6

co/i

operon, did not support growth of the mutant strain on non-fermentative carbon sources even at near-neutral pH. The basis for this observation is not yet clear but may relate to an overall decrease in the levels of cytochromes of various types (Gilmour and Krulwich, 1997). It is notable, though, that even partial mutational loss of the cau3-type oxidase, unaccompanied by significant decreases in other cytochromes, leads to a nonalkaliphilic phenotype (Krulwich ef a f . , 1996). The single major terminal oxidase of alkaliphilic Bacilfus YN-2000 is a cytochrome aco-type oxidase which has been purified and shown to be an oxygen- and CO-binding oxidase that contains two copper atoms (Qureshi et al., 1990). cta

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3.3. The ATP Synthase It was anticipated when studies of extremely alkaliphilic Bacillus species were undertaken in our laboratory, that these organisms would be found to have resolved the problem of ATP synthesis at low Ap by utilization of a Na+-translocating ATPase for synthesis. These bacteria use Na'lsolute symport rather than H+/solute symport as their ion-coupling mode of transport precisely to by-pass the same problem, i.e. that the reversed ApH component of the A p at the optimal, highly alkaline pH values for growth lowers the available total driving force. In spite of the fact that many of the strains grow on low or moderate Na+ concentrations, the total electrochemical gradient of Na' is much higher than the Ap when the bacteria are growing around pH 10.5, so use of Na+ instead of H+ would seem a logical way to circumvent the low Ap both for solute transport and ATP synthesis (Guffanti et al., 1978; Ivey et al., 1998). However, whereas this solution has been adopted for ion-coupled solute transport, it is not the solution used for ATP synthesis. In both B.firmus OF4 (Hicks and Krulwich, 1990) and B. alcalophilus (Hoffmann and Dimroth, 1991a), the FIFO-ATPasehas been purified, functionally reconstituted in proteoliposomes, and shown to be a H+-translocating enzyme that does not also translocate Na'. The alkaliphile enzyme is distinct from bona fide Naf -translocating FIFo-ATPases that have been documented in a few non-alkaliphilic bacteria, especially anaerobic ones (Kluge et al., 1992; Forster et al., 1995). Subsequent studies, after the genes encoding the FIF,-,-ATPase from B. firmus OF4 had been identified (Ivey and Krulwich, 1991), demonstrated that there was a single species of the enzyme expressed in both pH 7.5- and pH 10.5-grown cells and with no apparent change in the c-subunit/holoenzyme stoichiometry (hey et al., 1994). The deduced amino acid sequence of the a and c subunits of the alkaliphile Fo possess intriguing variations in regions that are thought to be of importance with respect to the path of protons through this part of the synthase assembly. These 'alkaliphile-specific motifs' have been found in B. firmus OF4, B. alcalophilus (Ivey and Krulwich, 1992), and a newly isolated alkaliphilic Bacillus species (D.M. hey, personal communication). These differences are shown diagrammatically in Fig. 7. They include an unusual lysine residue in a putative transmembrane region of the a subunit and a second proline in the region of an important carboxylate-bearing residue of the c subunit. The importance, if any, of these deviations in sequence is yet to be determined, but at least some of them might be expected to confer different properties upon these important functional regions of the Fo as compared with the neutralophilic enzyme.

426

T.A. KRULWICH, M . ITO, R. GILMOUR, D.B. HICKS AND A.A. GUFFANTI

IN

E Q

OUT

a4

a5

c2

cl

Figure 7 Alkaliphile ‘Fo motifs’ in the intramembrane helices of the a and c subunits. The data for the BadlusJirtnus OF4 sequence are shown in a model of the corresponding E. coli subunits that is based on the presentation of Hatch ei a/. (1995). Only the two helices, a4 and 0 5 , of the putative 5-helix a subunit are shown. The shift in the a-helix at the proline residues 58 and 64 is shown to indicate a possible change in the direction of the a-helix as has been determined for proline-64 of the E. mli c-subunit (Girvin and Fillingame, 1995). Residues in bold are those relatively conserved in Fo subunits that are different in the alkaliphile Fo subunits, the first letter indicating the residue found in the E. coli subunit and the second letter showing the alkaliphile residue. The numbering is taken from the E. coli subunits.

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3.4. Possible Models of Alkaliphile Oxidative Phosphorylation

The finding that the ion specificity of oxidative phosphorylation in E. coli can be converted to Na' by substitution of the Fo and 6 genes of the arp operon from the anaerobe Propionigenium modestum demonstrates that Na+-coupling is capable of supporting aerobic growth of eubacteria (Kaim and Dimroth, 1995). It is surprising, then, that alkaliphiles such as B.jirmus OF4 do not utilize Na+-coupling at the very alkaline growth pH at which the Ap is just a fraction of that observed at near-neutral growth pH whereas the electrochemical Na' gradient is high. Also important in evaluating any model for how oxidative phosphorylation is energized in the upper end of the pH range for growth is the finding that the molar growth yields are not compromised compared with growth at lower pH values. This suggests that the resolution to the energization problem may represent an energy-sparing solution since there is an energetic cost of pH homeostasis at high pH and the intrinsic energy cost of producing ATP from ADP and Pi is a pH-dependent function that is elevated as the pH rises. There is support for a particular energy-sparing aspect to oxidative phosphorylation in the more alkaline part of the alkaliphiles' pH range for growth. The experiments were conducted on (ADP + Pi)-loaded, right-side-out membrane vesicles prepared from pH 10.5-grown cells. The vesicles were energized by adding ascorbate in the presence of phenazine methosulfate as electron donor, and nigericin was included in order to assure that the Ap generated was all in the form of a AW, positive out. Energization was carried out at either pH 7.8 or 9.5. These pH values were chosen because they are, respectively, below and above the pH at which an artificially imposed diffusion potential ceases to be effective in energizing ATP synthesis by the same preparations (Guffanti and Krulwich, 1994). Two different protocols were used to titrate the AW downward, as described in the legend to Fig. 8, where the resulting ATP synthesis is shown. Downward titration of the transmembrane potential diminished the ATP synthesized at pH 7.8 even at levels of the AW that were still well above -100 mV. At pH 9.5, by contrast, reduction of the AW was comparatively without effect until it had been lowered beyond -100 mV. These observations supported the possibility that at the higher pH values at which growth is as good or better than at near-neutral pH, there is some sparing component that is not reflected in the measured steady-state AW.

Perhaps there is some manner in which this system is not functioning in full equilibrium. No explicit models have been presented that suggest a mechanism whereby the synthase itself might be functioning in a way in which displacement from equilibrium result in the observed deviations from a direct relationship between the AGp or rate of ATP synthesis and the magnitude of the Ap. However, several different models have been proposed

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T.A. KRULWICH, M. ITO, R. GILMOUR, D.B. HICKS AND A.A. GUFFANTI

A

-150

-100

-50

0

AY (mV) Figure 8 The effect on ATP synthesis of downward titration of the A@ across respiring, ADP t P,-loaded right-side-out membrane vesicles from B. firmus OF4. Membrane vesicles prepared from pH 10.5-grown cells were loaded with ADP + Pi in the presence of K', both inside and outside the vesicles, and nigericin (to prevent development of a ApH). Vesicles were energized at either pH 7.8 ( 0 )or 9.5 (0). In (A), energization was achieved by addition of various concentrations of ascorbate indicated in the paren-

whereby partial sequestration of the protons that are translocated outward by respiration may allow that proton component to reach the ATP synthase before it can fully equilibrate with the bulk. Five such models are illustrated in Fig. 9. In Fig. 9(A), the possibility is raised, as suggested by Skulachev (1991), that the low bulk Ap might not matter because the oxidative phosphorylation activity of the cell is sequestered in membrane-associated or some other kind of organelles. This suggestion has not been supported in a variety of structural examinations of cells and cell surfaces (Rhode et a/.,

ENERGETICS OF ALKALlPHlLlC BACLLUS SPECIES 2.5

429

B

2.0

x

:

E 1.5

il a -E

B .-

$z 1.0 v)

a l-

a

0.5

0

theses, together with 0.1 mM phenazine methosulfate. In (B), energization was achieved with 10 mM K-ascorbate + 0.1 mM phenazine methosulfate but the vesicles were preincubated for 5 min with the concentrations of valinomycin indicated in parentheses. Samples were taken for measurements of the A* via the distribution of tetraphenylphosphonium and for measurements of ATP. [Details of the methods are provided in (Guffanti and Krulwich (1994) from which this figure is reproduced with permission.]

1989; Sturr el al., 1994; Ivey et al., 1998). Figure 9(B) depicts the possible involvement of the acidic cell wall polymers that have been described by Aono and colleagues (Aono and Ohtani, 1990; Ito r t al., 1994; Aono et al., 1995) in binding divalent cations such that protons become trapped within the space between the coupling membrane and the polymers. Some variation on this theme has been proposed in the Aono group’s work as well as by Koch and colleagues (Koch, 1986; Kemper et al., 1993). However, as shown

A

C H+ 4

Figure 9 Models of various modes of sequestration of some of the protons translocated outward by the respiratory chain during oxidative phosphorylation so that they are utilized for energization of ATP synthesis without full equilibration with the outside bulk phase. These speculative models are described in the text.

ENERGETICS OF ALKALlPHlLlC BACILLUS SPECIES

43 1

in Fig. 8, the energetic dilemma with respect to oxidative phosphorylation is demonstrable in vesicle preparations that do not have significant content of cell wall material (Guffanti and Krulwich, 1994). In addition, as noted elsewhere (Krulwich, 1995), the pH near the external surface may be inferred to be high, even though it has not yet been measured directly. The pattern of sequence diversion (i.e. the low basic amino acid composition) of short external loops of alkaliphile polytopic membrane proteins indicates that the surface pH must be strikingly different from that encountered, for example, by B. suhtilis or B. siearothermophilus. Figure 9(C) depicts a sequestration in the form of sufficiently fast movement between a respiratory chain complex and the ATP synthase on the surface of the membrane, that a part of the driving force for synthesis is in the form of protons that have never equilibrated with the bulk. Several groups have reported faster movements along the surface of membrane phospholipids and/or membrane protein surfaces than in the extrusion of protons out into the bulk phase (Heberle et al., 1994; Scherrer et ul., 1994; Alexiev et al., 1995; Gutman and Nachliel, 1995; Gabriel and Teissie, 1996). That such movements are of bioenergetic significance has yet to be shown. It is notable that ATP synthesis by right-side-out vesicles from the alkaliphile at high pH is not adversely affected by high bulk buffering capacity or ionic strength (Guffanti and Krulwich, 1994). Figures 9(D) and 9(E) depict a model in which the proton pathway into the synthase above about pH 9.2 does not start at the external bulk phase or surface; in fact these models explain the inefficacy of an imposed diffusion potential above such pH values by positing a pH-regulated closing of the outer entry point for protons. Above that gating pH, protons are suggested to move into the Fo either by a direct proton hand-off from a respiratory chain complex such as the cytochrome cuu3-type oxidase or via the intermediary involvement of a coupling factor. In such a model, the ‘alkaliphilespecific’ motifs of the Fo might be involved in modulating an important pK in connection with the gating and/or proton pathway and in making the conformation of the crucial c subunit capable of accepting protons from the ‘alternate’ intra- or perimembrane route from the interacting proton partner. Features of these types of models are currently being tested directly. It is further possible that different kinds of alkaliphiles will utilize strategies to support oxidative phosphorylation that will not depend upon any sequestration. For example, there may emerge organisms that generate a much higher steady-state A W either because of their coupling membrane properties or their proton extrusion properties. Perhaps, as alluded to earlier, there will turn out to be bacteria that have global cytoplasmic adaptations that enable them to grow well with cytoplasmic pH values greater than 9.5; in that case, a higher bulk Ap might be generated because the magni-

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tude of the adverse pH gradient need not be as great as in the alkaliphilic Bacillus strains discussed here.

ACKNOWLEDGEMENTS Work from the authors' laboratory was supported in part by research grants GM28454 from the National Institutes of Health and DE-FG0286ER13559 from the Department of Energy.

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Cheng, J., Guffanti, A.A. and Krulwich, T.A. (1994) The chromosomal tetracyclineresistance locus of Bacillus subrilis encodes a Na+/H+ antiporter that is physiologically important at elevated growth pH. J . Biol. Chem. 269, 27 365-27 371. Cheng, J., Guffanti, A.A., Wang, W., Krulwich, T.A. and Bechhofer, D.B. (1996a) Chromosomal terA(L) gene of Bacillus subtilis: regulation of expression and physiology of a tetA(L) deletion strain. J. Bacteriol. 178, 2853-2860. Cheng. J., Hicks, D.B. and Krulwich, T.A. (1996b) The purified Bacillus subtilis tetracycline eftlux protein TetA(L) reconstitutes both tetracycline-cobalt/H+ and Na+(KC)/ H + exchange. Proc. Nail. Acad. Sci. USA 93, 14446-14451. Cheng, J., Guffanti, A.A. and Krulwich, T.A. (1997) A two gene ABC-type transport system involved in Na' extrusion by Bacillus subtilis is induced by ethanol and protonophore. Mol. Microbiol. 23, 1107-1 120. Clejan. S. and Krulwich, T.A. (1988) Permeability studies of lipid vesicles from alkalophilic Bacillus,firmus showing opposing effects of membrane isoprenoid and diacylglycerol fractions and suggesting a possible basis for obligate alkalophily. Biochim. Biophys. Acta 946, 4 M 8 . Clejan, S., Krulwich, T.A., Mondrus, K.R. and Seto-Young, D. (1986) Membrane lipid composition of obligately and facultatively alkalophilic strains of Bacillus. J . Bacteriol. 169, 44694478. DeGier, J.W., Schepper, M., Reijnders, W.N., van Dyck, S.J., Slotboom, D.H., Warne, A., Saraste, M., Krab, K., Finel, M., Stouthamer, A.H., van Spanning, R.J. and van der Oost, J. (1996) Structural and functional analysis of aa3-type and cbb3-type cytochrome c oxidases of Paracoccus denitrificans reveals significant differences in protonpump design. Mal. Microbial. 20, 1247-1260. Forster, A., Daniel, R. and Muller, V. (1995) The Na(+)-translocating ATPase of Acetobacterium woodii is a F IFo-type enzyme as deduced from the primary structure of its beta, gamma and epsilon subunits. Biochim. Biophys. Acta 1229, 393-397. Gabriel, B. and Teissie, J. (1996) Proton long-range migration along protein monolayers and its consequences on membrane coupling. Proc. Natl. Acad. Sci. USA 93, 14521I4 526. Garcia, M., Guffanti, A.A. and Krulwich, T.A. (1983) Characterization of the Na+/H+ antiporter of alkalophilic bacilli in vivo: a W-dependent "Na+ efflux from whole cells J. Bacteriol. 156. 1 151-1 157. Garcia-Horsman, J.A., Barquiera, B., Rumbley, J., Ma, J. and Gennis, R.B. (1994) The superfamily of heme-copper respiratory oxidases. J. Bacteriol. 176, 5587-5600. Gilmour, R. and Krulwich, T.A. (1996) Purification and characterization of the succinate dehydrogenase complex and CO-reactive b-type cytochromes from the facultative alkaliphile Bacillus firmus OF4. Biochim. Biophys. Acta 1276, 5743. Gilmour, R. and Krulwich, T.A. (1997) Construction and characterization of a mutant of alkaliphilic BacillusJirmus OF4 with a disrupted cta operon and purification of a novel cytochrome bd. J. Bacteriol. 179, 863-870. Girvin, M.E. and Fillingame, R.H. (1995) Determination of local protein structure by spin label difference 2D NMR: the region neighboring Asp61 of the subunit c of the F I F OATP synthase. Biochemistry 34, 1635-1645. Glagolev, A.N. (1980) Reception of the energy level in bacterial taxis. J. Theor. Biol. 82, 171.~185. Grant, W.D., Mwatha, W.E. and Jones, B.E. (1990) Alkaliphiles: ecology, diversity, and applications. FEMS Microbiol. Lett. 75, 255-270. Grundy, F.J., Waters, D.A., Takova, T.Y. and Henkin, T.M. (1993) Identification of genes involved in utilization of acetate and acetoin in Bacillus subtilis. Mol. Microbiol. 10. 259-271.

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T.A. KRULWICH, M. ITO, R. GILMOUR,

D.B.HICKS AND A.A. GUFFANTI

Guffanti, A.A. and Hicks, D.B. (1991) Molar growth yields and bioenergetic parameters of extremely alkalophilic Bacillus species in batch cultures and growth in a chemostat at pH 10.5. J . Gem Microhiol. 137, 2375--2379. Guffanti, A.A. and Krulwich, T.A. (1992) Features of apparent non-chemiosmotic energization of oxidative phosphorylation by alkaliphilic Bacillus .firniu.s OF4. J . Biol. Chem. 267, 9580 9588. Guffanti, A.A. and Krulwich, T.A. (1994) Oxidative phosphorylation by ADP -t Piloaded membrane vesicles from alkaliphilic Bacillus .firmus OF4. J . B i d . Clicpm. 269. 21 576 21 582. Guffanti. A.A., Susman, P., Blanco, R. and Krulwich, T.A. (1978) The protonmotive force and a-aminoisobutyric acid transport in an obligately alkalophilic bacterium. J . B i d . Cliein. 253, 708-715. Guffanti. A.A., Fuchs, R.T., Schneier, M., Chiu, E. and Krulwich, T.A. (1984) A A 9 generated by respiration is not equivalent to a diffusion potential of the same magnitude for ATP synthesis by BacilIu.s,/irtnus RAB. J . Biol. C h m . 259. 2971-2975. Guffanti, A.A., Finkelthal, O., Hicks, D.B., Falk, L., Sidhu, A,, Garro, A. and Krulwich. T.A. (1986) Isolation and characterization of new facultatively alkalophilic strains of Bacillu.~.J . B~crrriol.167. 76&773. Gutman, M. and Nachliel, E. (1995) The dynamics of proton exchange between bulk and surface groups. Biochim. Biophys. Acru 1231, 123-1 38. Hamarnoto, T., Hashimoto, M.. Hino. M.. Kitada. M., Seto, Y., Kudo, T. and Horikoshi, K . (1994) Characterization of a gene responsible for the N a t / H f antiporter system of alkalophilic Bacillus species strain C-125. Mol. Microhiol. 14. 939 -946. Hashimoto, M., Haniamoto, T., Kitada, M . , Hino, M., Kudo, T. and Horikoshi, K. (1994) Characteristics of alkali-sensitive mutants of alkaliphilic Bacillus sp. strain C125 that show cellular morphological abnormalities. Biosci. Biorech. Biocheni. 58. 2090-2092. Hatch, L.P., Cox, G.B. and Howitt, S.M. (1995) The essential arginine residue at position 210 in the a subunit of the Escherichia c d i ATP synthase can be transferred to position 252 with partial retention of activity. J . Biol. Chivn. 270, 29407-29412. Heberle, J., Riesle, J., Thiedemann, G., Oesterhelt, D. and Dencher, N.A. (1994) Proton migration along the membrane surface and retarded surface to bulk transfer. Nufurr. 370, 379-382. Hicks, D.B. (1995) Purification of three catalase isozymes from facultatively alkaliphilic Bacil1usfirmu.r OF4. Biochim. Biciphys. Acta 1229, 347-355. Hicks, D.B. and Krulwich. T.A. (1990) Purification and reconstitution of the F,F,-ATP synthase from alkaliphilic BacillusJirmus OF4: evidence that the enzyme translocates H t but not Na'. J . B i d . Ckr~ni.265, 20547-20554. Hicks, D.B. and Krulwich. T.A. (199.5) The respiratory chain of alkaliphilic bacilli. Biochim. Biophys. Acta 1229, 303--314. Hicks, D.B., Plass, R.J. and Quirk, P.G. (1991) Evidence for multiple terminal oxidases, including cytochrome d, in facultatively alkaliphilic Bacil1usJirniu.s OF4. J . Bacreriol. 173, m o - - ~ n 1 6 .

Hirota, N. and lmae, Y. (1983) Na'-driven flagellar motor of an alkalophilic Btrcillus strain YN-1. J . Biol. Chem. 258, 10577-10581. HolTmann, A. and Dimroth. P. (1991a) The ATPase of Bucillus a1calophilu.s. Reconstitution of energy-trdnsducing functions. Eur. J . Biochetn. 196, 493497. Hoffmann, A. and Dimroth, P. (1991 b) The electrochemical proton potential of Bmdlus alcalophilus. Eur. .I. Biocheni. 201, 4 6 7 4 7 3 . Horikoshi, K. (1996) Alkaliphiles from an industrial point of view. FEMS Microhiol. Lert. 18, 259- 270. ~

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Author Index Note

~-Page

numbers in ifalics indicate where a reference is given in full.

Aasd, R. 198, 226, 230 Abate, L. 145, 187 Aberg, A. 321, 341 Abshire, K.Z. 271, 273 Acuna, G. 196, 206, 210, 218, 224, 228 Adachi, 0. 6, 7, 8, 9, 10, I I , 13, 14, 16, 17, 18, 19,20,21, 22,24,25, 39,40, 41,42,43, 44,47, 50, 58,60,67, 71, 73, 74, 75, 77, 78, 80 Adam, E. 141, 183 Adamowicz, M. 42, 49, 67 Adarns, M.D. 140, 145, 146, 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, I 74, 175, 176, 188, 304, 343, 403, 41 7, 419, 438 Adams, M.W.W. 161, 180 Adler, M. 17, 72 Aguilar. G.R. 219, 220, 221, 222, 229 Aguilar, J . 244, 277 Agus. R. 419, 436 Ahringer, J. 94, 132 Al-Basseet, J. 298, 299, 301, 317, 324, 343 Alben. J.O. 197, 225 Albertini, A. 305, 346 Albracht, S.P.J. 164. 185, 409, 437 Alderson, J. 174, 180 Alexiev, U. 431, 432, 437 Aliabadi, Z. 237, 238, 239, 278 Allewell. N.M. 288. 350 Al-Massad, F. 292, 298, 299, 301, 302, 311, 312, 317, 324, 343, 347 Alrniron, M. 238, 257, 259, 273 Altschul, S.F. 99, 132 Altuvia, S. 257, 259, 265, 268, 273, 278 Amako, K . 304, 333. 350 Amaratunga, K. 10, 64, 67 Ames, B.N. 250, 276

Ameyama, M. 6, 7, 8, 9, 11, 13, 14, 16, 17, 18. 19,20,21,22,24,25, 39,40,41,42,43, 44,47, 67, 73, 74, 75, 77, 78 Anazawa, H. 64, 66, 72, 79 Andersen, L.P. 153, 173, 180, 186, 187 Anderson, D. 10, 64, 67 Anderson, R.F. 207, 227 Andersson, L. 337, 340 Andrews, S.C. 151, 189, 285, 291, 292, 295, 297, 298, 299, 300, 301, 302, 303, 304, 305, 310, 317, 319, 320, 323, 324, 325, 326, 328, 329, 332, 333, 334, 335, 337, 340, 341, 342, 343, 345, 346, 348, 349 Annear, D.I. 139, 185 Anraku, Y. 193, 222 Anthamatten, D . 174, 187, 195,210, 21 I , 212, 213, 214, 215, 216, 217, 218, 222, 224, 225, 227 Anthony, C. 3, 6, 8, 9, 10, 11, 16, 17, 18, 19, 20,21,24,25, 26,27, 28, 29, 30, 31, 32, 33, 35, 36, 37, 38,41,42,43,48,60,64, 67, 68, 69, 70, 71, 72, 73, 74, 76, 77 Aono, R. 403,404,406,407,412,418,421, 428, 432, 435 Aoyarna, H. 214, 230 Appleby, C.A. 194 198, 206, 207, 209, 210, 213, 222 Appleby, C.A. 196, 198, 206, 209. 210, 213, 222, 223, 224, 228, 230 Applernan, M.D.143, 182 Arbinger, B. 96, 133 Archibald, A.R. 387, 398 Archibald, F. 283, 341 Arents, J.C. 58, 75 Arigoni, F. 217, 218, 224, 225 Armellini, D. 145, 187 Armstrong, J.A. 139, 140, 141. 142, 144, 177. 183, 185

440 Arosio, P. 285, 288, 290, 305, 320, 321, 323, 325, 326, 341, 342, 345,346 Artymiuk, P. 319, 341 Artymiuk, P.J. 288, 291, 305, 320, 323, 326. 345, 346 Asai, T. 50, 68 Asaoka, S. 17, 18, 48, 49, 62, 7Y Aspedon, A. 49, 76 Atsumi, T. 408, 432 Atta, M. 321, 341 Attwood, M.M. 62, 70 Au, D.C.T. 198, 208, 222, 223 Auger, E.A. 237, 238, 239, 252. 263, 276 Austin, J.W. 147. 155, 182, 184, 303, 343 Auton, K.A. 37, 68 Avezoux, A. 21, 24, 25, 29, 30, 68. 69, 71 Axon, A.T.R. 143, 153. 188 Aziz, J. 145, 187 Azzi, A. 202, 223 Babcock, G.T. 197, 222, 225 Babel, W. 42, 47. 76 Babst, M. 210, 218, 224, 225 Babu-khan, S. 58, 68 Bickstrorn, D. 337, 34Y Bader, R. 58, 75 Badia, J. 244, 277 Baer, W. 170, 171, 180 Bagg, A. 337, 341 Baker, M.E. 100, 133 Bakkeren, D.L. 337, 341 Baldoma, L. 244, 277 Baldwin. S.A. 100, 108, 132, I33 Balny. C. 9, 10, 71 Bamforth, C.W. 8. 68 Banfalvi, Z. 219, 220, 226 Banfieid, M.J. 320, 323. 345 Bannister, J.V. 292, 298, 300, 326, 329, 347 Banyard, S.H. 288, 341 Baoguang, Z. 294. 341, 345 Barassi, C. 198, 227 Barclay. R. 150. I84 Bare, R.E.298, 317, 344 Barer, M.R. 146, I83 Barker, J . 236, 273 Barker, P.D. 298, 341 Barnickel, G . 390, 3Y8 Barquera, B. 174,183, 195, 197.207,208,212. 223, 225 Barquiera, B. 424, 433 Barra, D. 316, 342

AUTHOR INDEX

Barrett, E.L. 247, 273 Barrett, L.J. 177, 185 Barta, T.M. 60, 68 Barth, M . 258, 273 Bartlett, J.A. 236, 274 Bartsch, R.G. 292, 298, 341 Barynin, V. 297, 301, 302, 317, 319, 324, 341 Basagni, C . 145, 187 Bassilana, M. 41 1, 432 Batut, J. 195, 210, 211, 212, 216, 217, 218, 223. 225, 228 Bauerfiend, P. 178, 180, I86 Baugh, C.L. 168, 181 Baum, H. 283, 350 Bauminger, E.R. 292, 294, 295, 296. 320, 325, 326, 327, 328, 337, 338, 342, 350 Baumler, A.J. 334, 335, 349 Bayeli. P.F. 145, 187 Beardmore-Gray, M. 17, 18, 42, 68 Bearson, S. 245 Bearson, S.M.D. 260, 269, 270, 273 Beattie, P. 409, 432 Bechhofer, D.B. 406, 410, 433 Becker, S.A.W.E. 298, 342 Behlau, 1. 250, 273 Behrendt, M.C. 149, 182 Beier, D. 148, 180 Bell, S.H. 326, 328, 342, 349 Beness, A.M. 135 Benjamin, W.H. Jr. 260, 269, 270, 273 Bennett, M.J. 130, 132 Beppu, S. 9, 13, 14, 15, 78 Beppu,T. 9.13, 14, 15, 19.68, 74. 78, 198.224 Berg, D.E. 140, 145, 146, 148, 149. 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176, 188, 403, 417, 419, 438 Bergensen, F.J. 198, 213, 222 Berger, E.M.284. 348 Bergersen, F.G. 209, 223 Berka, T. 44, 74 Berlyn, M.B. 295, 342 Berlyn, M.K.B. 243, 274 Berry, E. 197, 225 Best, L.M. 141, l8Y Betts, J.D. 147, I84 Beveridge, T.J. 429, 435 Bezanson. G.S. 141, 189 Bianciardi, S. 145, 187 Biedermann, G . 284, 342 Bill, E. 337, 338, 347 Birkholz, S. 164, 165, 180

AUTHOR INDEX

Bimstiel, M.L. 293, 345 Bishop, R. 333, 334. 335, 341 Bisseling, T . 219, 228 Biswas, I. 412, 432 Bittinger, M . A . 194, 204, 205, 210, 224 Bittner. R. 238, 257, 275 Biville, F. 52, 53, 54, 57, 58.64, 68, 69, 71, 74, 75, 76, 78

Blair, A. 126, 132 Blake, C.C.F. 24, 26. 27. 28, 32, 68, 69, 71 Blarney, J.M. 161, I8U Blanco, R. 405, 425, 434 Blaser, M.J. 142, 143, 181, 188, 235, 274 Blasi, U. 129, 136 Blattner, F.R. 129, 136 Blaut, M. 366, 395 Bloch, P.L. 295, 347 Blom, J. 153, 180 Blount, P. 129, 136 Blows, W.D. 158, 159, 160, 170, 181 Blum, P.H. 237, 238, 239, 252, 263, 276 Bockman, A.T. 238, 277 Bode, G. 146, 180 Bogusz, D. 209, 223 Bohnke, R. 292, 298, 300, 311, 312, 337, 338, 342

Boiardi, J.L. 7, 18, 22, 50, 51, 69, 71 Boistard, P. 212, 218, 223, 228 Bolliger, M . 194, 196, 202, 203, 206, 210, 223 Bonina, L. 235, 274 Bonnet, F. 155, 156, 170, 185 Bonomi, F. 291, 342 Boos, W. 241, 274 Booth, I.R. 406, 41 I , 432 Bork, P. 329, 331. 348 Borodovsky, M. 140, 142, 145, 146, 148, 149, 150, 152, 154, 155. 162, 166, 167, 168, 169, 174, 175, 176, 181, 188, 403, 417, 419, 438 Borrias, M. 285, 342 Borrielo, S.P. 148, 189 Borys, A . 246, 277 Bossewitch, J.S. 412, 435 Bossie, S. 250, 274 Botsford, J.L. 238, 252, 274 Botstein, D . 245, 278 Bott, M. 194, 196, 202, 203, 204, 205, 206, 210, 218, 219, 220, 223, 225, 226, 228 Bottke, W. 285, 341 Bouchard, L. 334, 349 Bourne, R.M. 409, 432

44 1 Bovy, A . 285, 342 Bowler, L . D . 398 Bowman,C. 140, 145, 146, 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176, 188,403, 417, 419, 438 Boyd, G. 24, 79 Boyd, G . D . 24, 26, 79 Bozzi, M. 316, 342 Bradaczek, H. 390, 398 Bradley, K.L. 305, 347 Bradley, W.F. 291, 326, 349 Brady, L. 147, 183 Brdkenhoff, G.J. 387, 399 Brandsch, R. 55, 75 Braun, V. 294, 345 Bremer, E. 258, 274 Brennan, P.J. 329, 348 Briat, J.-F. 285, 292, 298, 300, 304, 312, 335, 341, 342, 346

Briffeuil, P. 298, 343 Britt, B.M.51, 78 Brock, J.H. 302, 303, 348 Broer, A . 92, 132 Broer, S. 92, 93, 129, 132, 135 Broger, C. 202, 223 Brooker, R.J. 107, 133 Brookes, C.L. 177, 182 Brooks, B.W. 298, 342 Brooks Low, K. 295, 342 Broome-Smith, J.K. 398 Brouwer, P. 218, 224 Brown, M.H. 100, 104, 107, 129, 130, 134 Brown, M.R.W. 236,273 Brown, S. 197, 223 Brubaker, R.R. 336, 348 Bruhn, D . F . 93, 132 Brutovetsky, N. 93, 132 Bryant, M.P. 366, 395 Buchmeier, N. 250, 274 Buchmeier, N . A . 239, 248, 249, 270, 275 Buck, G.E. 145, 180 Bugnoli, M . 145, 187 Buhman, A . 152, 178, 185 Bulder, I. 56, 57, 59, 79 Bulen, W.A. 293, 342 Bullerjahn, G.S. 315, 348 Bult, C.J. 304, 343 Bunn, C.R. 193, 224 Burdett, I.D.J. 384, 390, 397, 398 Burdett, I.D.K. 387, 395 Burnens, A.P. 144. 188

442 Burns, B.P. 157, 158, 159, 164, 168, 180, I86 Burr, D.H. 147, I89 Burton, S.M. 62, 73 Butt, J.N. 332, 333, 348, 349 Buurman, E.T. 18, 22, 47, 69 Byler, R.M. 284, 342 Cabello, F. 235, 274 Calam, J. 140, 143. 181, 186 Caldeira, J. 9, 12, 21, 70 Calderwood, S.B. 285, 347 Calhoun, M.W. 197, 225 Capaldi, R.A. 198, 223 Carrano, C.J. 292, 298, 300, 311, 312, 342 Cashel, M. 238, 257. 261. 274, 275 Castle, A.M. 408, 436 Castresana, J. 212, 223 Catrenich, C.E. 145, 181 Censini, S. 142, 181 Cesareni, G . 288, 305, 320, 323, 326, 346 Cha, J. 17, 69 Chakrabarti, S. 194, 199, 223 Chakrabartty, P.K. 194, 199, 223 Chalk, P.A. 156, 158, 159, 160, 161, 162, 164, 165, 166, 167, 168, 169, 170, 171, 172, 173, 174, 175, 181, 184, 185, I88 Chalker, R.B. 235, 274 Chan, H.T.C. I I , 16, 20, 21, 41, 69 Chan, S.I. 198, 227 Chance, B. 198, 227 Chang, H. 162. 170, 171, I81 Chang, H.-T. 173, 174, 175, 185 Chang, S. 356, 395 Change, Y.S. 53, 58, 74 Chapman, T. 173, 174, 175, 185 Charloteaux-Wauters, M. 337, 343 Chasteen, N.D. 326, 337, 347 Chater, K.F. 100, 133 Chaudhry, G.R. 297, 342 Chauhan, S. 195, 223 Chaut, J.-C. 128, 133 Chavas-Alba, 0. 288, 344 Chawla, A. 227 Cheesman, M.R. 298, 299, 300, 30 I, 3 17, 3 19, 324, 343, 346, 347 Chelm, B.K. 207, 225 Chen, C.-Y. 250, 274 Chen, C.Y. 239. 243. 245, 246, 248, 251, 253, 256, 261, 264. 265, 269, 275 Chen, J.W. 221, 227 Chen, M. 292, 297, 343

AUTHOR INDEX

Cheng, A.F.B. 140, 184, 188 Cheng, J. 406, 407, 409, 410, 411,433, 436 Chepuri, V . 198, 223 Chiancone, E. 316, 342 Chiao, E. 250, 270, 275 Chilvers, T. 140, 144, 183 Chinnery, R. 148, 189 Chippendale, G.R. 177, 187 Chistoserdova, L. 60, 69 Chiu, E. 422, 434 Cho, G. 105, 133 Chou, H.-H. 64. 77 Christoserdova, L. 53, 54, 55, 56, 78 Chung, S.C.S. 140, 184, 188 Clark, D.P. 244, 263. 274 Clayton, C.L. 148, 151, 161, 162, 164, 165, 167, 168. 171, 172, 174, 177, 180. 181. 184, 187, 189 Clayton, R.A. 140, 145, 146, 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176, 188, 304. 343, 403, 417, 419. 438 Clegg, G.A. 289, 345 Clejan, S. 403, 419, 433 Cleton, M.I. 327, 349 Cleton-Jansen, A. 17, 18, 19, 61, 69 Cockdyne, A. 148, I89 Cohen, H. 143, 182 Cohen, J.I. 236, 274 Cohen, S.G. 292,294,295, 296, 326, 327, 328, 337, 338, 342, 350 Cokie, P. 409, 436 Cole, R.M. 387, 395 Cole, S.T. 218, 224, 329, 348 Collier, J.R. 262, 270, 276 Collins, M.D. 140, 144, 171, 173, 181, I83 Colwell, R.R. 146, 187, 235, 278 Conway, T. 42, 49, 67, 71, 130, 135 Cooper, S. 390, 395 Cooperberg, B. 403, 41 I , 435 Corey, G.R. 236, 274 Correa, P. 143, 144, 182 Costas, M. 144, 171, 173, 181, 188 Cotton, M.D. 140, 145, 146, 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176. 188, 403, 417, 419, 438 Coucoux, P. 177, I84 Covacci, A. 142. 181 Cover, T.L. 142, 143, 181, 188 Coves, J. 337. 339, 344 Cox, D.L. 168, 181

AUTHOR INDEX Cox, G.B. 426,434 Cox, J.M. 10, 69 Coynault, C. 256, 271, 276 Cozier, G.E. 17, 24, 26, 31, 32, 33, 69, 70 Cozzi, A. 305,346 Crabtree, J. 153, I88 Crabtree, J.E. 142, 181 Cragoe, E.F., Jnr. 408, 432 Crichton, R.R. 288, 292, 297, 314,337.343, 344, 348 Crick, F.H. 393, 399 Croes, L.M. 63, 70 Cronan. J.E. Jr. 244, 263, 274 Cross, A.R. 36, 70 Cross, R. 149, 182 Cruse, W.B.T. 3, 77 Cubitt, C.L. 238, 239, 240, 245, 246, 247, 248, 253, 257, 261, 264. 278 Cui, X. 291, 342 Cui. X.Y. 329, 332, 344 Curry, A. 144, 184 Curtiss, R. 236, 275 Cusdnovich, M.A. 292, 298,347 Cussac, V. 177, 178, 181, 184 Dai, W. 24, 26. 79 Dai, W.W. 24, 79 Daldal, F. 197, 225 Dales, S.L. 11, 21, 24, 25, 70, 71 Damiano, E. 41 I , 432 Danbara, H. 249,253,257,259,261, 270,271, 275 Danchin, A. 198, 228 Daniel, R. 425, 433 Daniel, R.M. 206, 224 DAri, R. 259, 277 Dassa, E. 112, 119, 121, 134. 135 Dautant, A. 288, 319, 341, 344 Daveran, M.L. 216, 218, 225 Daveran-Mingot, M.-L. 195, 210, 21 I , 217, 218.223 David, C.N. 293, 343 David, M. 195, 210, 211, 212, 216, 217, 218, 223. 224, 225, 228 David, S. 56, 57, 59, 79 Davidson, V.L. 6, 1 I , 21, 24, 70, 72, 79 Davies, A.E. 194, 199,20I , 202,203. 208.2 19, 220, 224, 230 Davies, K. 417. 438. Davis0n.A.A. 156, 162, 164, 165, 166, 167, 170, 171, 172, 181

443 Davy, S.L. 292, 298, 316, 336, 348 Dawes, E.A. 46, 60, 61, 75, 76, 79 Dawson, A.P. 410, 417, 436 Day, D.J. 10, 64, 69, 76 Deamer, D.W. 356, 359, 370, 396, 398 Dean, D.R. 329, 344 de Boer, A.P.N. 197, 212, 230 de Bont, J.A.M. 79 DeBoer, T. 64,79 De Bruijn, W.C. 327, 349 De Cherchi. M.I. 337, 344 Deeb, S.S. 292, 295, 343, 344 Defago, G. 52, 53, 54, 77 DeGier, J.W. 424, 433 Degraaff, J. 150, 189 Degregorio, L. 145, 187 de Grier, J.W. 197, 212, 230 De Hollander, J.A. 194, 195, 199, 208, 224 Deiana, G. 337, 344 De Jong, G.A.H. 9, 12, 21, 70 De Jong, R. 7, 49, 70 Dekker, R.H. 6, 70 Dekker, S. 17, 18, 69 Delgado, M.J. 194, 199, 201, 202, 203, 208, 209, 215, 216, 219, 220, 221, 224, 230 deMare, F. 291, 323, 326, 343 Demattos, M.J.T. 18, 42, 46, 47, 69, 72, 76 Demchich, P. 380, 381, 396 de Mooy, O.H. 47, 79 Dencher, N.A. 431, 434 Denoel, P.A. 298, 343 Dent, J.C. 155, 156, 170, 185 De Pedro, M.A. 387, 396 DeReuse, H. 178, 182 Dersch, P. 258, 274 Desnoues, N. 214, 217, 218, 225, 226 Despied, S. 334, 349 Dessi, A. 337, 344 De Vries, G. 64. 72 De Vries, G.E. 60, 62, 70 De Vries, S. 9, 12, 21, 24, 31, 70, 78, 198, 230 de Vries, W. 194, 195, 199, 208, 229 de Vrieze, G . 285, 342 Devereux, J. 309, 343 Dewanti, A. 18, 22, 25, 47, 75 Dewhirst, F.E. 141, 144, 183, 188 Dey, S. 92, 93, 129, 135 Dhaenens, L. 150, 182 Diavolitsis, S. 143, 182 Diaz del Castillo, L. 193, 226 Dick, J.D. 156, 182

444 Dickson, D.P. 329, 345 Dickson, D.P.E. 292, 294, 298, 326, 327, 328, 329, 336, 337, 342, 348, 349 Dierks, T. 93, 132 Diez, A.A. 246, 259, 275 Diffin, F.M. 295, 349 Dijkhuizen, L. 63, 70, 235, 275 Dijkstra, M. 9, 10, 71 Dimroth, P. 409,421,422,425,427,434,435, 437 Dinh,, T. 100. 133 Dinh, D. 104, 133 DiRusso, C.C. 244, 246, 259, 275 Dittmer, D.S. 168, I82 Dixon, M. 84, 133 Dixon, M.F. 142, 143, 153, 182, 188, 189 D’Mello, R. 213, 224 Dobereiner, 3. 50, 51, 78 Dodson, R. 140, 145, 146, 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176, 188, 403, 417, 419, 438 Doig, P. 155, 182, 303, 343 Dokter, P. 17, 18, 19, 70, 72 Dombroski, D.M. 236, 274 Domergue, 0. 212, 216. 218, 224, 225 Donachie, W.D. 390. 396 Dooley, C.P. 143, 181, 182 Doolittle, R.F. 96, 105, 133, 415, 436 Dorman, C.J. 257, 258, 271, 277 Dou, D. 92, 93, 129, 135 Dougan, G. 241, 244, 245 Dougherty, B. 140. 145, 146. 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176, 188 Dougherty, B.A. 304, 343,403, 417, 419, 438 Dougherty, T.J. 238, 274 Douglas, E.T. 195, 223 Dover, S. 244, 274 Downie, J.A. 194, 199, 201, 202, 203, 208, 209, 215, 216, 219. 220, 221, 224, 230 Doyle, R.J. 372, 374, 383, 384, 387, 388, 390, 396, 397, 398, 429, 435 Doyle, T.J. 259, 262. 271. 272, 279 Drabble, W.T. 43, 60, 71 Drexler, M. 241, 274 Dreyfus, B.L. 214, 218, 226 Druilhet. R.E. 238, 274 Duchars, M.G. 62, 70 Duine, J.A. 3, 6, 7, 8, 9, 10, 11, 12, 16, 17, 18, 19, 20, 21,22,24,25. 31, 31,43,44,47,

AUTHOR INDEX 51, 59, 60, 70, 71. 72, 73, 75, 76. 77, 78,

79 Dunlap, J.C. 94, 134 Dunstan, P.M. 43, 60, 71 Duron, M.R. 58, 68 Dutton, P.L. 423, 436 Earhart, C.F. 285, 339, 343 Easterbrook, K. 293, 343 Eaton, K.A. 147, 177, 182 Ebisuya, H. 198, 224 Edelman, A. 398 Ehrenberg, A. 337, 349 Ehrlich, S.D. 412, 432 Eiglmeier, K. 218, 224 Eklund, H. 321, 323, 341, 347, 348 Elferink, M.G.L. 42, 46, 49, 79 Elkan, G.H. 193,224 El-Khani, M.A. 238, 274 EIKurdi, A.B. 175, 183 Ellermann, K. 52, 53, 54, 76 Elliott, E.J. 16, 41, 71 Elmerich, C. 194, 195, 209, 211, 214,215, 216, 217, 218, 225, 226, 227 El Mokadem, M.T. 203, 224 Endre, G. 414,437 Engstrand, L. 178, 182 Ernst, R.K. 236, 274 Escamilla, E. 194, 199, 221, 229 Escamilla, J.E. 195, 207, 208, 223 Esposito, E. 145, 187 Evans,D.G. 141, 145, 163, 178,182,183,315. 3 16,343 Evans, D.J. 141, 145, 154, 178, 182, 183 Evans, D.J. Jr. 163, 183, 315, 316, 343 Fabianek, R.A. 219, 220, 224 Falk, L. 404, 409, 434 Falk, P. 155, 182, 303, 326, 344 Falkow, S. 58, 78. 236, 275, 276 Fan, W.-H. 52, 53, 54, 76 Fang, F. 246 Fang, F.C. 236, 238, 239, 243, 245, 246, 248, 249, 250, 251, 253, 256, 257, 261, 262. 264, 265, 269, 270. 274, 275, 277 Farewell, A. 246, 259, 275 Farmery, S. 141, 184 Farrar, J. 298. 299, 301, 317. 324, 343 Fath, M.J. 112, 133 Fayet, 0. 17, 18, 61, 69 Federbush, J. 407, 410, 417, 436

AUTHOR INDEX Fee, J.A. 153, 182, 197, 230, 305, 347 Feldmann, K.A. 130, 132 Felsenstein, J . 310, 343 Fen, W.-H. 64, 77 Feng, D.F. 105, 133 Ferguson-Miller, S. 197, 198, 225, 229 Fernandez, E. 94, 133 Fernandez, M. 94, 133 Fernie, A.R. 288, 345 Ferrero, R.L. 159. 177, 178, 181, 182 Fetter, J . 197, 225 Fiedorek, S.C. 141, 183 Fields, C . 304, 343 Fierer, J. 236, 239, 275 Fiermonte, G . 94, I33 Figura, N. 145, 187 Fikumori, Y . 423, 438 Fillingame, R.H. 426, 433 Filser, M. 199, 228 Findlay, J.B.C. 292, 297, 301. 311, 312, 341, 34 7 Finel, M. 424, 433 Finkelthal, 0. 404, 409. 434 Finlay. B.B. 236, 275 Fischbach, F.A. 291. 326, 343 Fischer, D. 258, 273 Fischer, K. 95, 96, 133, 134, 135 Fisher, H.M. 199,210,217,218,224,225,228 Fitzgerald, L.M. 140, 145, 146, 148. 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176, 188, 403,417, 419,438 Fitzgibbons, P.L.143, 182 FitzHugh, W. 304, 343 Flatmark, T. 337, 349 Fleischaker, G.R. 356, 3Y6 Fleischmann, R . D . 304. 343, 403, 417, 419, 438 Fleishmann, R.D. 140. 145, 146, 148. 149, 150, 152, 154, 155, 162, 166. 167, 168, 169, 174, 175, 176, 188 Fliege, R. 49, 71 Flitter, W. 284, 344 Floyd, R.A. 284, 344 Fluckiger, R. 6, 71 Flugge. U.-I. 95, 96, 133. 134, 135, 136 Fomchenkov, V.M. 238, 276 Fontecave, M . 321, 337, 339, 341, 344 Fontham, E. 143, 144, 182 Ford, G.C. 288, 295. 317, 326, 344, 345, 348, 349 Forman. D. 143, 182

445 Forrest, H.S. 3, 77 Forster, A . 425, 433 Forward, J.A. 149, 182 Foster, J. 245 Foster, J.W. 235, 236,237, 238, 239, 240, 245, 246, 247, 251, 257, 260, 261, 263, 265, 269, 270, 273, 275, 278 Fox, A.J. 144, 184 Fox, J.G. 141, 143, 144, 182, 183 Fox, R.B. 284, 348 Franceschinelli, F. 305, 346 Frank, J . 3, 6, 8, 9, 10, 11, 16, 17, 18, 20, 21, 22, 37. 43, 70, 71, 72, 76, 77, 78 Frankel. R.B. 292, 294, 301, 302, 324, 325, 326, 327, 329, 350 Fraser,C.M. 140, 145, 146, 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176, 188,403,417,419, 438 Fraser, G.J. 141, 183 Frazier, B.A. 155, 182, 303, 326, 344 Frebortova, J . 60, 71 Frederick, C . A . 321, 323, 348 Frederick, K.L. 149, 183 Frehel, C. 238, 276 Freund, S.M.V. 298, 341 Frohlich, 0. 95, 133 Frommer, W.B. 95. I35 Frowlow, F. 299, 301, 317, 318, 323, 324, 344 Frustaci, J.M. 195, 207, 224 Fu, C. 172, 173, 175, 185 Fu, W. 329, 344 Fuchs, R.T. 422, 434 Fujii, A. 9, 11, 13, 39. 43, 44, 78 Fujii, C. 140, 145, 146, 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176. 188,403, 417, 419, 438 Fujita, T. 292, 295, 344 Fujiwara, T. 423, 424, 437 Fukaya, M . 9, 13, 14, 15, 19, 78, 198, 224 Fukuda. M. 9, 14, 73 Fukumori, Y. 292, 297, 300, 346, 423, 424, 437 Fukumoto, Y. 404, 423, 438 Fukuzumi, A. 4. 73 Fullerton, F. 143, 182 Furlong, D. 238, 273 Fusamaoto, H. 144, 188 Gabazza, E.C. 150, 186 Gabel, C. 194, 196, 203, 204. 205, 210, 224, 226

446 Gabriel, B. 431, 433 Gabriel, N.E. 198, 227 Gabriel, W.M. 238, 246, 248, 253, 256, 257, 261, 262, 264, 277 Gagnon, J. 292, 298, 300, 312, 335, 346 Gaillour, A. 141, 184 Galan, J.E. 236, 275 Galar, M.L. 7, 50, 51, 69, 71 Galibert, F. 128, 133 Gallant, J. 238, 257, 275 Gallois, B. 288, 344 Gallop, P.M. 6, 71 Gdmcsik, M. 156, 182 Garcia, M. 41 I , 433 Garcia-Horsmdn, A. 195, 207, 208, 223 Garcia-Horsman, J.A. 174, 183, 197,212,225, 424, 433 Garg, J. 129, 135 Garg, R.P. 329, 332, 344 Garland, D. 284, 350 Garlid, K.D. 93, 134 Garner, R.M. 178, 180, 186 Garnerone, A . M . 195.210,211,212,217,218, 223, 228 Gamier. M. 155, 156, 170, 185 Garro, A. 404,409, 434 Gasser, F. 52. 53. 54, 57, 58, 64. 68,69,71, 74, 75, 78 Gassmann, W. 130, 133 Cause, G . F . 356, 396 Gea, Y.298, 317, 344 Geerlof, A. 9, 12. 21, 70. 71 Geiger, 0. 6, 17, 18, 19, 22, 24, 25, 71, 72 Geis, G. 147, 185 Geisler. V. 164, I84 Gennis. R.B. 174, 183, 197, 198,208,212,222, 223, 225. 227, 230,247,275, 424, 433 Gentry, D.R. 238, 257, 261, 274, 275 George, G.N. 298, 317, 344 Georgiou, C. 197, 225 Geren, L. 202, 229 Gergious, C.D. 198, 227 Gero. A . M . 169, I136 Gerrits, M.M. 146, 184 Gerritse, G. 403, 4313 Gerstenecker, B. 153, 154, I88 Gessa, C. 337, 344 Ghai, J. 212, 216, 218. 225, 228 Gherdrdi, M. 212, 223. 228 Ghirra, P. 142, 1x1 Ghosh, M . 24. 26, 27, 28, 32, 68, 69. 71

AUTHOR INDEX Giangregorio, N. 94, 134 Giberman. E. 294, 342 Gibson, G. 141, 184 Gibson, J.F. 327, 328, 342, 350 Giesbrecht, P. 390, 398 Gilbert, J. 172, 173, 175, 185 Gilbert, J.V. 172, 183 Gilden, N. 238, 278 Giles, I.G. 31, 32, 70 Gill, S. 140, 145, 146, 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176, 188, 403, 417, 419, 438 Gilmour, R. 403,405,407,416,417,418,421, 423, 424, 425, 428, 433, 435, 436 Girvin, M.E. 426, 433 Gish, W. 99, 132 Glagolev, A.N. 412, 433 Glancy, R.J. 141, 142, 185 Glaser, P. 198, 228 Glauner, B. 380, 385, 396 Glerum, D.M. 94, 136 Glockshuber, R. 219, 220, 224 Glodek, A. 140, 145, 146, 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176, 188, 304, 343, 403,417,419, 438 Gloudemans, T. 219, 228 Gocayne, J.D. 140, 145, 146, 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176, 188, 304, 343, 403, 417, 419, 438 Goffeau, A. 100, 133 Goggin, P.M. 144, 183 Goldberg, E.B. 130, 135 Goldstein, A.H. 46, 53, 58, 68, 71, 74 Gomelsky, M. 52, 53, 54, 57, 71 Gonda, M. 197. 225 Gonzalez, T. 237, 238. 239, 245, 247, 251,278 Goodell, A. 384, 398 Goodell, E.W. 384, 392, 396, 396 Goodhew, C.F. 175, I83 Goodwin, A. 165, 167, 168, 184 Goodwin, C.S. 139, 140, 144, 177, I83, I135 Goodwin, M.G. 21, 24, 25, 26, 28, 29, 30, 32, 68. 71 Goodwin, P. 64, 74 Goodwin. P.M. 10, 62, 64, 67, 77 Goosen, N. 17, 18, 19. 52, 53, 54. 55, 57, 61. 69. 72 Gorisch, H. 4, 6, 9, 1 I , 17, 18, 19, 21, 22, 24, 25, 43, 71, 72, 76

AUTHOR INDEX

447

Goswitz, V.C. 107, 133, 198, 227 Gulig, P.A. 249, 253, 257, 259, 261, 262, 270, 271, 272, 275, 279 Gottschalk, G. 366, 395 Gustafsson. L. 246, 248, 277 Gouaux, E. 129, 133 Gutensohn, M. 96, 133 Gould, S.J. 355, 396 Gutman, M. 431, 434 Graham, D.Y. 141, 145, 154, 163, 178, 182, Gutteridge, J.M.C. 152, 183, 284, 344 183, 184, 185 Guy, H.R. 129, 136 Granier, T. 288, 344 Grant, W.D. 404, 433 Ha, Y. 288, 350 Grass, A. 412, 432 Haas, D. 52, 53, 54, 77 Gray. H.B. 198, 228 Haas, R. 147, 154, 185. 187 Gray, K.A. 197, 225 Hadzija, 0. 337, 349 Green, H.G. 130, 132 Hager, L.P. 292. 295, 343, 344 Greenman, J . 145, 176, 188 Haggerty, D.A. 129, 135 Greenwood, C. 292, 298, 299, 300, 301, 311, Hagihara, B. 295, 344 312, 317, 324, 329, 343, 345, 347, 348 Hahn, H. 172, 189 Greenwood, J.A. 62, 72, 73 Hahn, J.J. 387, 395 Gregory, D.W. 291, 326, 343 Hahn, M. 199, 228 Gribbon, L. 146, 183 Haines, T.H. 370, 398 Griffith. J.K. 100. 133 Haldane, J.B.S. 356, 396 Griffith, P.L. 150, 184 Hall, H.K. 236, 239, 265, 269, 275 Grinius, L.L. 130, 135 Halliwell, B. 152, 183, 284, 344 Grodsitski, D. 423, 437 Halpern, Y.S. 244, 274 Groen, B.W. 6, 7,9, 11, 12, 21, 24,44, 72, 78 Haltia, T. 198, 205, 225, 226. 423, 436 Grogan. J.M. 332,333,334,335,341,348,349 Hamamoto, T. 413, 414, 416, 419, 434 Groisman, E.A. 250, 270, 275 Hamasaki, N. 304, 350 Grooms, M. 197, 225 Hammar, M. 303, 326, 344 Gross, A. 95, 135, 136 Hammer, M. 155, 182 Gross, C.A. 265, 268, 275 Han, H.M. 154, 185 Grosskopf, E. 414, 437 Hancock, I.C. 387, 398 Grossman, M.J. 299, 305, 310, 319, 344 Handt, L.K. 141, 183 Grundy, F.J. 419, 433 Hanna, P.C. 262, 270, 276 Grutter, M.G. 18, 19, 77 Hanners, J.L. 51, 73, 78 Guerinot, M.L. 207, 225, 285, 344 Hanson, R. 64,74 Guerrant, G . 173, 186 Hanson, R.S. 60, 63, 64,68, 74, 76, 79 Guerrant, R.L. 177, 185 Hantke, K. 334, 335,349 Guerry, P. 147, 189 Harayama, S. 8. 77 Guest, J.R.217, 229, 292, 295, 291, 298, 299, Harder, W. 235. 275 300, 301, 302, 303, 304, 305, 310, 317, Hardy, G.P.M.A. 46, 72 319, 320, 323, 324, 325, 326, 328, 329, Harker, A.R. 293, 344 332, 333, 334, 335, 337, 340, 341, 342, Harlos, K. 24, 26, 28, 32, 69, 71 343, 345, 346, 348, 349 Harms, N. 60, 62, 64,66, 70, 72, 74, 79, 80 Guffanti, A.A. 403, 404, 405, 406, 407. 409, Harper, W.E.S. 140, 144, 183 410, 411, 412, 413, 414, 416, 417, 418, Harris, T.K. 11, 21, 72 419, 421, 422, 423, 424, 425, 427, 428, Harrison, P.M. 285, 288, 289, 290, 291, 292, 429, 431, 433, 434, 435, 436, 437 293, 295, 297, 298, 299, 300, 301, 302, Guillain, F. 179, I84 303, 304, 305, 310, 315, 317, 319, 320, Guiney, D.G. 236, 239, 243, 245, 246, 248, 321, 323. 324, 325, 326, 327, 328, 329, 249, 250, 251, 253, 256, 257, 259, 261, 333, 334, 335, 337, 340, 341, 342, 343. 264, 265, 269, 270, 211, 274, 275 344, 345, 346, 347, 348, 349 Gulig, P. 236 Hartman, H. 283, 345

448 Hartmann, A. 294,345 Hanvood, C.R. 387,398 Hanvood, J. 239, 248, 249, 270, 275 Hashimoto, M. 413. 414, 416, 417, 419. 434, 435 Hassan, 1.J. 145, 176, I88 Hatch, L.P. 426, 434 Hauge, J.G. 16, 72 Hausinger, R.P. 178, I86 Hauska, G. 423,437 Hausler, R.E. 96, 133 Hawkey, C.J. 148, 189 Hawkins, C. 297, 299, 300, 301, 303, 304, 325, 328, 334, 337, 341,345 Hawkins, J.M. 333 Hayashi, M. 7. 20, 22, 67, 409, 438 Hayashi, N. 144, 188 Hayes, J.M. 364. 396 Hayes, W.S. 140. 145, 146, 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176, 188,403,417,419, 438 Hazelaar, M.J. 64,79 Hazell, S.L. 145, 147, 154, 156, 157, 158, 159, 160, 164, 166, 168, 169, 170, 176, 177, 180, 183, 186 Heatley, R.V. 143, 153, 188. 189 Heberger, C. 93, 132 Heberle, J . 431, 434 Hechel, D. 320, 325, 342 Hedblom, E. 304, 343 Heffron, F. 250, 270, 275 Hefta, S.A. 329, 348 Heijenoort, J.V. 368, 370, 372, 396 Heldt, H.W. 96, 133 Hellingwerf, K.J. 42, 46, 49, 79 Helman, J.D. 149, 183 Hempstead, P.D. 288,291,305, 320, 323.326, 345 Henderson, P.J.F. 100, 133 Hendricks, J.K. 178, 183 Hengge-Aronis, R. 238, 240, 246, 248, 257, 258, 259, 261, 273, 275, 276 Henkin, T.M. 419, 433 Hennecke, H. 174, 187, 194, 195, 196, 199, 200, 201, 202, 203, 204, 205, 206, 210, 211, 212, 213, 214, 215, 216, 217, 218, 219, 220, 222, 223, 224, 225, 226, 227, 228, 229. 231 Hennessy, W. 147, I83 Henninger, H. 55, 75 Herbas, A. 288, 344

AUTHOR INDEX

Herman, P.T.D. 59, 73 Hernandez, V.J. 238, 257, 261, 274, 275 Herrmann, R. 345 Hess-Bienz, D. 417, 438 Hessel, A. 243, 278 Heul, C. 337, 341 Heyn, M.P. 431, 432, 437 Hickey, E. 140, 145, 146, 148, 149. 150, 152. 154, 155, 162. 166, 167, 168, 169. 174, 175, 176, 188 Hickey, E.K. 403, 417, 419, 438 Hicks, D.B. 403,404,405, 406,407, 409, 410, 41 I , 417, 421, 423. 424, 425, 428, 433, 434, 435, 436, 437 Higgins, D.G. 212, 223 Higgins, M.L. 383, 387, 395, 396, 398 Higins, M.L. 374, 397 Hilbert, H. 345 Hildebrand, M. 130. 133 Hill, B.C. 198, 225 Hill, J. 197, 225 Hill, J.J. 198, 227 Hill, S. 42,48, 72, 73, 77, 213, 224 Hille, B. 133 Hillebrand, H. 214, 218, 226 Himmelreich, R. 345 Hino, M. 413, 414, 416, 419,434, 436 Hinton, S.M. 299, 305, 310, 319, 344 Hirota, N . 408, 418, 434 Hirsch, P.R. 216, 218, 225 Hoare, R.J. 293, 345 Hobot, J.A. 387, 398 Hochberg, M.L. 384, 398 Hodge, A.J. 293, 345 Hodson, N . W . 320, 325,342 Hoffman, M.E. 284,347 Hoffman, P.S. 155, 165, 167, 168, 184 Hoffmann, A. 42 I , 422,425,434 Hofnung, M. 112, 119, 121, 134 Hoj, P.B. 176, 188 Hollanders, D. 139, 185 Holm, L. 198, 228 Holmes, M.A. 321, 323, 345 Holtje, J.-V. 372, 380, 384, 385, 386, 387, 396, 397,398 Holtje, J.V. 387, 396 Hommes, R.W.J. 18.42.47.49, 59.61, 72, 73, 76 Honor&,N. 218, 224 Hoogenkijk, J. 64,72 Hook-Nikanne, J. 162, 163, 187

AUTHOR INDEX

Hopper, D.J. 10, 73 Horikoshi, K. 403,406, 407, 410, 413, 414, 416, 417, 418, 419, 421, 428, 432, 434, 435, 436 Horinouchi, S. 9, 13, 14, 15, 19, 73, 74, 78, 198, 224 Horio, T. 292, 298, 341 Hormaeche, C. 235, 274 Horne, R. 176, 188 Horowitz, N.H. 368, 396 Horsman, H.P.A. 52, 55, 57, 72 Horwitz, L.D. 284, 342 Horwitz, M.A. 336,347 Hosking, S.W. 140, 184 Hosler, J.P. 197, 225 Houck, D.R. 51, 73, 78 Howitt, S.M. 426, 434 Hoy, T.G. 291, 293, 326, 343, 345 Huang, H. 324, 350 Huang, M.-E. 128, 133 Huber-Wunderlich, M. 219, 220, 224 Hudson, A.J. 303, 304, 320, 323, 325, 333, 337, 342, 345 Huesca, M. 154, 185 Hughes, N.J. 151, 161, 162, 164, 167, 168, 171, 172, 181, 184, 189 Huguet, M. 212, 223 Huguet, T. 212, 223, 228 Huinen, R.G.M. 52, 55, 57, 72 Huisman. G. 257, 259, 273 Huisman, G.W. 237, 238, 240, 241, 245, 276 Hullo, M.F. 198, 228 Huls, P. 387, 399 Hummel, S. 95, 135 Hung, C.H. 53, 58, 74 Hunt, C. 292, 298, 336, 348 Hunt, J.C. 46, 73 Hunt, S. 193, 225, 226 Hunter, F. 143, 144, 182 Husson, M.O. 150, 182, 184 Hutchinson, W.J. 129, 135 Hyde, B.B. 293, 345 Hynes, M. 195, 21 I , 216, 218, 229 Hynes, M.F. 216, 228 Iacobazzi, V. 94, 134 Igloi, G. 55, 75 Ikernura, N. 150, 186 Illingworth, C.A. 94, 136 Illingworth, D.S. 150, 184 Irnae, Y. 408,418, 432, 434

449 Imai, C. 9, 14, 73 Imanaga, Y. 18, 22, 33, 73 Imlay, J.A. 334, 337, 346 Irnoto, I. 150, 186 Indiveri, C. 94, 134 Inoue, T. 9, 14, 18, 73, 74 Ishikawa, Y. 407, 436 Ishizuka, M. 197, 225 Islam, Q.T. 326, 348 Island, M.D. 178, 186 Itagaki, E. 292, 295, 344 Ito, M. 403, 406, 407,411,412,413,414,416, 417, 418, 421, 424, 425, 428, 432, 435, 436 Ito, S. 17, 77 Ito, T. 144, 188 Itoh, S. 4, 18, 22, 73, 77 Iuchi, S. 218, 224 Ivanov, A.I. 238, 276 Ivey, D.M. 403,412,413,416, 417, 421, 425, 428,435 Iwata, S. 214, 225 Izuhara, M. 302, 303, 304, 325, 335, 337, 345 Jack, D.L. 129, 135 Jack, R.F. 329, 344 Jackson, C.J. 148, 184, 187 Jacobs, D. 302, 324, 325,350 Jacobs, J. 195, 210,211, 217, 218, 223 Jacobson, G.R. 241,277 Jacques, M. 334, 349 James, P. 200, 206, 212, 222, 229 Jang, J. 94, 136 Janka, J.J. 79 Janney, F. 143, 144, 182 Jensen, D.B. 257, 275 Jernigan, H.M. 284, 350 Jeu-Jaspars, C.M.H. 337,341 Jezek, P. 93, 134 Ji, G. 92, 93, 129, 132, 135 Jimenez, B.M. 169, 186 Jiudi, L. 294, 341, 345 Jiwen, W. 294, 341, 345 Joblin, K.N. 403, 418, 428, 432 John, T.R. 221, 227 Johnsen, J. 165, 167, 168, 184 Johnson, M.K. 329,344 Johnson, R.C. 244, 257,258,279 Johnston, A.W.B. 194, 199, 201, 219, 220, 22 1, 224, 227, 230 Jones, B.E. 404,433

AUTHOR INDEX

Jones, C.W. 62, 72, 73 Jones, D.M. 144, 184 Jones, R.M. 219, 228 Jongejan, J.A. 3, 6, 8.9, 10, 12, 17, 20, 21.22, 24, 31, 70, 71, 78 Jonniaux, J.-L. 100, 133 Jordan, M.P. 219, 228 Jordan, P. 332, 333, 348, 349 Josenhans, C . 147, 182. 188 Juty, N.S. 42, 48, 73 Kaback, H.R. 108, 134 Kadir. F. 298, 300, 317, 343 Kadir, F.H. 329, 345 Kadir, F.H. A. 292, 298, 299, 30 I , 302, 3 1 I , 3 12, 3 17, 3 19, 324, 329, 343, 345, 347, 348 Kado, C.I. 17. 69 Kahn, A. 293, 345 Kahn, D. 195, 210. 211. 212, 216, 217, 218, 223, 224, 225 Kaim, G . 427,435 Kakuno, T. 292, 298,341 Kalb, A.J. 292, 295, 296, 299, 301, 317, 318, 323. 324, 338, 344, 349, 350 Kaluza, B. 219, 228 Kamada, T. 144, I88 Kamen, M.D. 292, 298, 341 Kaminski, P.A. 194, 195, 208, 209, 211, 214, 215, 216, 217, 218,225, 226, 227 Kammerer, B. 96, 133, 134 Kaneko, H. 407, 432 Kang, S.-K. 403, 435 Kansau, I. 179. 184 Karp, P.D. 140, 145, 146, 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176. 188, 403, 417, 419, 438 Kaspar, T. 210, 218, 224, 225 Kato, N . 8, 77 Kawai, F. 8, 73 Kawakami, H. 4, 73 Kawamura, Y. 9, 13, 14, 15, 19, 78 Kawano, S . 144, I88 Keech, A. 317, 324, 349 Keech, A.M. 324, 346 Keefe, R.G. 203,210,212, 213, 226 Keel, C. 52, 53, 54, 77 Keen, J.N. 292, 297, 301, 302, 303, 311, 312, 341, 347, 348 Keilin, D. 295, 346 Keister, D. 196, 206, 210. 223

Keister, D.L. 203,206, 210,224,226,227,230 Kelley, J.M. 140, 145, 146, 148, 149, 150, 152, 154. 155, 162, 166, 167, 168, 169, 174, 175, 176, 188, 304, 343, 403. 41 7, 4 19, 438 Kellner, E. 423, 437 Kelly, D.J. 148, 149, 151, 156, 161, 162, 164, 165, 166, 167. 168, 171, 172. 174, 180, 181, 182. 184, 187, 189 Kelly, M. 198, 226 Kelly, S. 141, 184 Kernper, M.A. 429, 435 Keneszt, A. 414. 437 Kennard, 0. 3, 77 Kereszt, A. 219. 220, 226 Kerlavdge, A.R. 140, 145, 146, 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176, 188, 304, 343, 403, 417, 419, 438 Kersten, A. 153, 154, 188 Kessel, M. 146, 187 Ketchum, K.A. 140, 145, 146, 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176, 188, 403, 417, 419, 438 Keyer, K. 334, 337, 346 Keyhan, M. 151, 152, 185 Khalak, H.G. 140, 145, 146, 148, 149, 150, 152, 154, 155, 162. 166, 167, 168, 169, 174, 175, 176, 188, 403, 417, 419, 438 Khan, M.A. 238,245,247,248,257.259, 260, 261, 266, 268, 271, 278 Khorana, H.G. 431,432, 437 Kier, L.D. 250, 276 Kikuchi, T. 17, 77 Kim, J.G. 141, 185 Kim, S.D. 141, 185 Kim, Y.M. 23, 64, 76 King, N.D. 195, 226 King, R.F.G. 143, 188 Kinosito, K. Jr. 126, 134 Kirchner, G. 390, 398 Kirkness, E.F. 140, 145, 146, 148, 149, 150, 152, 154, 155, 162, 166, 167, 168. 169, 174, 175, 176, 188, 304, 343, 403, 417, 419, 438 Kirshbom, P.M. 203,227 Kiss, P. 414, 437 Kist. M. 153, 154, I88 Kitada, M .407, 410, 413, 414, 416, 417, 419, 434, 435, 436 Kitagawa, T. 423, 438

AUTHOR INDEX Kitamura, Y. 18, 22, 77 Kitts, C.L. 195, 196, 208. 209, 215, 226 Kleanthous, H. 177, 181 Klein, P.D. 141, 183, I84 Klenk, H.P. 140, 145, 146, 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176, 188, 403, 417, 419, 438 Klingenberg, M . 93, 132 Klinman, J.P. 6, 73 Kluge, C. 425, 435 Knaff, D. 423, 436 Knipp, U. 164, 165, 180 Koch, A . 429, 435 Koch, A.L. 235, 276, 356, 361, 364, 368, 371, 372, 373, 374, 375, 379, 380, 381, 382, 383, 384, 385, 386, 387, 388, 389, 390, 392, 393, 396, 397, 398, 429, 435 Koivisto, T. 162, 163, 187 Kolmerer, B. 197, 226 Kolter, R. 112, 133, 237, 238, 240, 241, 245, 257, 259, 273, 276 Kondo, K. 9, 13, 14, 15, 73, 74, 78 Kondo, M. 417,437 Kondorosi, A . 219, 220, 226, 414, 437 Kongings, W.L. 42, 46, 49, 79 Konings. W.N. 406, 437 Koopman, H. 170, 171, 180 Korolyov, A.V. 194, 195, 199, 208, 226 Kortner, C. 164, 165, 184, 185 Kostrzynska, M. 147, 184 Kosunen, T.U. 162, 163, 187 Kowarz, L. 256, 271, 276 Koyama, N . 404,407,436 Kraayveld, D.E. 9, 12, 24, 31, 78 Krab, K. 424, 433 Kraft, A. 380, 398 Krakowka, S . 147, 177, 182 Kramer 195, 211, 216, 218, 229 Kramer, M. 195, 215, 218, 228 Kramer, R. 93, 132 Krause, M. 236, 239, 275 Kreig, N.R. 155, 184 Kretovich, W.L. 194, 195, 199, 208, 226 Kroger, A . 164, 165, 180, 184, 185 Kroll, R.G. 406, 436 Krulwich, T.A. 403, 404,405, 406, 407, 409, 410, 411, 412, 413,414, 416, 417, 418, 419, 421, 422, 423, 424, 425, 427, 428, 429, 43 I , 433, 434, 435, 436, 437 Kuan, G . 112, 119, 121, 134

45 1 Kudo, T. 403,410,413,414, 416, 417, 419, 434, 435, 436 Kuenen, J.G. 18, 22,42, 46,47, 49, 59, 61, 79 Kuh, K. 60, 69 Kuipers, E.J. 150, 189 Kukral, A.M. 236, 250, 270, 277 Kullik, I. 210, 218, 224, 225 Kung, C. 129, 136 Kunow, J. 161, 184 Kunst, F. 198, 228 Kiinzler, P. 219, 220, 224 Kurokawa, T. 292, 297, 300, 346 Kurtz, D . M . 313, 346 Kurtz, D . M . Jr. 329, 332, 344 Kurtz, M. 291, 342 Kusters, J.G. 146, 150, 184, 189 Kutz, D.M. 291, 323, 326, 343 Kuwamura, Y. 198, 224 Kuzma, M. 193, 226 Kyte, J. 415, 436 Labigne, A . 147, 159, 177, 178, 179, 181, 182, 184, 188 Labischinski, H. 368, 384, 390, 398 Laboure, A.-M. 292, 298, 300, 304, 312, 335, 346 Lai, E.C. 129, 135 Lamb, J.W. 199, 228 Lambert-Fair, M.A. 173, 186 Lamouliatte, H. 155, 156, 170, 185 Lampert, H.C. 315, 316, 343 Lange, C. 142, 181 Lange, R. 246, 276 Langlois d’Estaintot, B. 288, 344 LaPlace, P.S. 383, 398 Lappalainem, P. 198, 230 Lappalainen, P. 198, 226 Larsson, C. 246, 248, 277 Laubinger, W. 425, 435 Laufberger, V. 293, 346 Laulhtre, J.-P. 285, 292, 298, 300, 304, 312, 335, 341, 346 Lauraeus. M . 198, 226, 423, 436 Lauterbach, F. 164. 165, 184, 185 Lawson, D . M . 288, 320, 323, 326, 345, 346 Lax, A.J. 249, 253, 257, 259, 261. 270, 271, 275 Layzell, D.B. 193, 225, 226 Le Brun, N.E. 292, 297, 298, 299, 301, 31 I, 312, 317, 319, 324, 326, 328, 343, 346. 347, 349

452 Le, T. 129, 134 Leach, D. 409, 432 Leblanc, G. 41 I , 432 LeBrun, N . E . 297, 301, 302, 317, 324, 341 Leclerc, H. 150, 184 LeComte, J.R. 293, 342 Leduc, M.238, 276 Ledvina, P.S. 1 I I , 135 Lee, A. 147, 159, 177, 182, 183 Lee, A.G. 134 Lee, C.A. 236, 276 Lee, E. 52, 53, 54, 64, 76, 77 Lee, F.D. 143, 188 Lee, L.Y. 53, 58, 74 k , N .140, 145, 146, 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176, 188,403,411, 419, 438 Leenen, E.J.T.M. 9, 12, 71 Lees, H. 297, 342 Legrand, D. 150, 184 Lehmann, Y. 291, 346 Leitch, F.A. 301, 346 Lemieux, L. 197, 198, 223, 225, 228, 230 Lemma, E. 164, 165, 180, 184 Lengeler, J.W. 241, 277 Lenger, R. 164, 184 Lessie, T.G. 44,74 Lesuisse, E. 314, 348 Letesson, J.-J. 298, 343 Leung, V.K.S. 140, 188 Levi, S. 285,288, 305, 320, 323, 325, 326,341, 342, 346 Levy, A. 294, 326, 327, 328, 337, 342 Lewandowski, F.A. 288,350 Lewis, R.J. 423, 436 Lewison, J.F. 177, I87 Lewontin, R.C. 355, 396 Leying, H. 147, 185 Li, A.K.C. 140, 184, 188 Li, B.C. 345 Li, J. 93, 132 Li, M.K.K. 140, 188 Libby, S.J. 238, 239, 246, 248, 249, 250, 253, 256, 257, 261, 262, 264, 210, 274, 275, 277 Lichtfouse, B. 298, 343 Lidstrom, M. 10, 64, 67 Lidstrom, M.E. 10, 23, 52, 53, 54, 55, 56, 57, 59, 60,63, 64, 69, 74, 7s. 76, 77, 78 Liebl, Y. 423, 437 Limet, J.N. 298, 343

AUTHOR INDEX

Lin, E.C.C. 218, 224 Lin, R.T. 259, 277 Lin, T.K.W. 140, 184, 188 Linder, D. 161, 184 Lingwood, C.A. 154, 185 Linial, M. 100, 135 Link, A. 238, 273 Linton, D. 144, 188 Linton, J.D. 47, 79 Lipman, D.J. 99, 132 Lippard, S.J. 321, 323, 348 Lipps, C.J. 250, 270, 275 Little, E. 105, 133 Littlejohn, T.G. 100, 107, I34 Litwin, C.M. 285, 347 Liu, H.J. 129, 135 Liu, L.4. 304, 343 Liu, Q. 94, 134 Liu, S.-L. 243, 278 Liu, S.T. 53, 58, 74 Livingstone, J.C. 288, 320, 323, 326, 346 Lobreaux, S. 285, 341 Loddenkotter, B. 96, 133, 134 Loehr, T.M. 9, 12, 21, 70 Loenen, W.A.M. 53, 54, 55, 57, 75 Loewen, P.C. 239, 246, 248, 249, 253, 270, 275, 276, 277 Loferer, H. 196, 203, 204, 206, 210, 218, 223, 225,226, 228 Loftus, B. 140, 145, 146, 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176, 188, 403, 417, 419, 438 Logan, R.P.H. 148, 189 Long, A.R. 43, 74 Longhi, C. 316, 342 Lorence, R.M. 208, 222 Lough, S. 293, 342 Low, K.B. 243,274 Liibben, M.197, 198, 226, 228 Lubben, M. 194, 196, 198, 199, 206, 207, 229 Liibenn, M. 212, 223 Lucht, J.M. 241, 274 Lucier, T.S. 336, 348 Ludwig, B. 214, 225, 424, 437 Ludwig, R.A. 195, 196, 208, 209, 215, 226 Lugones, L. 285, 342 Lutz, M. 332, 333, 348. 349 Luzzago, A. 288, 305, 320, 323, 326, 346

Ma, J. 197, 212, 225, 424, 433 Macara, I.G. 293, 345

AUTHOR INDEX McCallum, R.W. 177, 185 McConnell, W. 140, 144, 183 McCormick, J. 329, 348 McEwan, A.G. 292,298, 316, 336,348 McFall. E. 244. 245, 276 McGechie, D.B. 141, 142, 185 Machida, K. 197, 225 Machlin, S. 64,74 Mack, R. 356, 395 McKenney, K. 140, 145, 146, 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176, 188, 304, 343, 403, 417, 419, 438 Mackey, J.B. 325, 345 McKnight, J. 298, 317, 347 McLaggan, D. 410,417,436 McLeod, G.I. 246,270,277 McNab, R.M. 408, 436 McNulty, C.A.M. 155, 156, 170, I85 Magasanik, B. 237, 276 Magee, K. 165, 167, 168, 184 Maguin, M. 412, 432 Mahan, M.J. 249, 253, 261, 262, 270, 276 Mai, U . 146, 187 Maidhof, H. 368, 398 Maier, J. 194, 196, 198, 199, 206, 207, 229 Maier, R.J. 172, 173, 175, 185, 194, 196, 199, 203, 204, 205, 206, 207, 208, 210, 212, 213, 224, 226, 227, 229 Makin, K.M. 145, 181 Malaty, H.A. 141, 183 Malaty. H.M. 141, 185 Malfertheiner, P. 140, 146, 180, 185 Malmstrom, B.G. 198, 207, 226, 228. 230 Mandel, K.G. 410,436 Mandon, K. 194, 195,209,21I , 214,215,216, 217, 218, 225, 226, 227 Mann, S. 292, 298, 300, 303, 304, 323, 325, 326, 327, 328, 329, 337, 345, 347, 349 Mansour, A.N. 326, 337, 347 Marcelli, S.W. 162, 170, 171, 173, 174, 175, 181, 185 Marchant, A. 130, 132 Marger, M.D. 100, 134 Markesich, D.C. 145, 183 Marrie, T. 141, 189 Marschall, C. 258, 273 Marsh, S.S. 210, 226 Marshall, B.J. 139, 141, 142, 144, 177, 183, 185 Marti, T. 431, 437

453 Martin, D. 139, 18.5 Martin, L.M. 58, 68 Martinac, B. 129, 136 Martinez-Drets, I . 50, 51, 78 Massey, V. 207, 227 Mastroeni, P. 235, 274 Mathews, F.S. 24, 26, 79 Matin, A. 151, 152, 185, 237, 238, 239, 246, 252, 263, 265, 268, 276, 277 Matsuda, H. 417, 437 Matsuhashi, M. 370, 372, 377, 398 Matsumoto, A. 423, 437 Matsumura, K. 17, 77 Matsushita, K. 6, 7, 8,9, 10, 11, 13, 14, 16, 17, 18, 19, 20, 21, 22, 24, 25, 39,40,41,42, 43,44,47, 50, 58, 60,67, 71, 74, 75, 77, 78,80 Matzanke, B.F. 292, 298, 300, 311, 312, 337, 338,342,347 Mauch, F. 146, 180 Mauk, A.G. 324, 346 Mauk, M.R. 324, 346 Maurer, K. 60, 62, 64, 70, 72 May, K. 289,345 May, S. 139, 185 May, S.T. 130, I32 Mayer, F. 428, 437 Mazodier, P. 53, 54, 75 Mee, B.J. 176, 188 Megraud, F. 140, 155, 156, 170, 185 Meijer, W.G. 63, 70 Meile, L. 291, 346 Mekalanos, J.J. 236, 249, 250, 253, 261, 262, 270, 276, 277 Melchers, K. 151, 152, 178, 185, 186 Meldrum, F.C. 303, 304, 325, 337, 345 Mello Fihlo, A.C. 284, 347 Membrillo-Hernandez, J. 221, 227, 229 Mendelson, N.H. 383, 399 Mendz,G.L. 145, 156, 157, 158, 159, 160, 164, 166, 168, 169, 170, 176, 180, 186 Meneghini, R. 284, 347 Menendez, C. 55, 75 Mengaud, J.M. 336.347 MenginLecreulx, D. 178, 182 Merad, T. 387, 398 Merrick, J.M. 236, 274, 304, 343 Merrick, M. 42, 48, 73 Mervarech, M. 294, 342 Meulenberg, J.J.M. 52, 53, 54, 55, 56, 57, 59, 75, 79

454

AUTHOR INDEX

Meyer, J.-B. 319, 341 Meyer, L. 199, 228 Meyer, T.E. 292, 298, 347 Meyer-Rosberg, K . 151, 152, 186 Micera, G . 337, 344 Michaux, M.-A. 288, 344 Michel, H. 214, 225 Michetti, P. 140, 185 Midgley, M. 46, 61, 75, 79 Mignogna, G. 316, 342 Miles, R.J. 162, 170, 171, 173, 174, 175, 181, 185

Millar, M.R. 141, 145, 176, 188, 189 Miller, C.G. 238. 277 Miller, J. 139, 185 Miller, S.I. 236, 250, 270. 273. 277 Miller, S.L. 356, 395, 398 Miller, W. 99, 132 Millet, F. 202, 22Y Millner, P.A. 130, I32 Minak-Bernero, V. 299, 305, 310, 319, 344 Minchin, F.R. 193. 230 Minghetti, K.C. 198, 227 Minnaert, K. 195, 229 Miranda, J . 221. 227 Misaki, M . 150, 186 Mishra. A.K. 194, 199, 223 Mishra, P.V. 238,245,247,248,257,259,260, 26 I , 266, 268, 27 I , 278 Mitchell, B.A. 100, 107, 134 Mitchell, P. 85, 134, 407, 420, 436 Mitchell, R. 197, 223 Mizote, T. 148, 186 Mobley, H.L.T. 177, 178, 180, 183, 186, 187, 387, 388, 390, 398 Modriansky, M. 93, 134 Mogi, A . 197, 225 Mollaaghababa, R. 431, 432 Mondrus, K.R. 403, 419,433 Monkara, F.A. 292, 298, 336, 348 Moody, J . 197, 223 Moomaw, C. 197, 225 Moon, J.Y. 409,436 Moore, G.R. 292, 297, 298, 299, 300, 301, 302, 311, 312. 316, 317, 319, 324, 326, 328, 329, 336, 341, 343, 345, 346, 347, 348, 349 Mordant, P. 100, 133 Morgan, D.R. 147, 177, I82 Morgan, T.V. 329, 344 Mori, A . 9, 14. 73

Moriya, T. 304, 333, 350 Morotomi. S.410, 435 Morowitz, H. 356, 398 Morris, C. 10, 64, 67 Morris, C.J. 23, 52, 53, 54, 63, 64,75, 76, 77 Moshiri, F. 42.48, 73, 172, 173, 175, 185. 194, 196, 203, 206, 210, 226, 227, 229 Moss, C.W. 173, 186 Moss, S. 143, 186 Mougel, C. 214, 218, 226 Moura, 1. 9. 12, 21, 70 Moura, J.J.G. 9, 12, 21. 70 Moyle, J. 85, I34 Mueller, R.H. 42, 47, 76 MufRer, A. 258, 273 Mullany, P. 177, 181 Mulleners, L.J.S.M. 403, 438 Muller, A. 423, 437 Miiller, G. 337, 338, 347 Miiller, V. 366, 395 Muller, V. 425, 433 Mulvey, M.R. 246, 248, 253, 277 Miinck, E. 197, 230 Murphy, A.M. 194, 196, 206, 229 Murray, E. 130, 135 Musacchio, A. 198, 230 Mutzel, A . 4. 9, 11, 21, 24, 76 Mwatha, W.E. 404, 433 Myers, E.W. 99, 132 Myllicallio, H. 197, 225 Nachliel, E. 431, 434 Nadler, K . D . 221. 227 Nagano, K. 144, 188 Nagata, K . 173, 174, 186 Nagatani, Y. 17, 74 Nagy-Czako, I. 337. 349 Nakano, H. 315, 316,343 Nakao, K . 150, 186 Nakashima, R. 214, 230 Nakayama, K . 304, 333,350 Nakazawa, T. 148, I86 Nakshabendi, 1. 143, 188 Namba, T. 364, 399 Nanninga, N. 3x6, 387, 398. 399 Naranjo, C.M. 305, 347 Nasu, M. 417, 437 Nautiyal, C.S. 203, 227 Nedenskov, P. 145, 169, 186 Neidhardt, F.C. 265, 268, 271, 273, 277. 295, 347

AUTHOR INDEX Neijssel. O.M. 7, 18, 22,42,46, 47.49, 59, 61, 69, 70, 72, 73, 76, 235, 278 Neilands. J.B. 285, 337. 341, 347 Nelson, K. 140, 145, 146. 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176. 188, 403, 417, 419, 438 Nelson, O.E. Jr. 94, 136 Newell. D.G. 143, I82 Newman. E.B. 244, 245, 259, 276, 277 Ng. F.M.W. 46, 60, 61, 76 Ngo, L. 126, 132 Nguyen. D.E. 257, 275 Nicholson, M . A . 173, 186 Nickerson, K.W. 42, 49, 67, 71, 76 Niederhoffer, E.C. 305, 347 Nielsen, H. 153, 180, I86 Nielsen, K . H . 298, 342 Nilsen, T. 337, 347 Nishi, K . 8, 77 Nishiyama, M. 9, 13, 14, 15, 78 Nitsche, C.I. 197, 230 Nitschke, W. 423,437 No, E.K.W. 140, 188 Noji, H. 126, 134 Nonobe, M. 17, 18, 20. 22, 42, 67, 74, 77 Norel, F. 249, 253, 256, 257, 259, 261, 270, 271, 275, 276 Norlund, P. 291, 321. 323, 326,341, 343,347, 348 Normark. S.J. 155, 182, 303, 326, 344 North, R.A. 134 Northfield, T.C. 144, 183 Nosoh. Y. 404, 407. 436 Nowik, I. 294, 320, 325, 342 Nunn, D. 10, 64,67 Nunn, D.N. 10, 23, 64,76 Nystrom, T. 246, 248, 259, 275, 277 OBrian, M.R. 195, 199, 203, 207, 208, 210, 223, 224, 226, 227 OBrien. P. 202, 229 OByrne, C.P. 257, 258, 271, 277 O’Connor. D.C. 10, 64,67 OConnor, H.J.144 143, 187 O’Connor, M.L. 60, 63, 76 Odenbreit, S. 154, 187 Odle, G. 69 O’Donnell, C. 174, 181 Odum, L. 173, 187 Oesterhelt, D. 431, 434

455 Ofer, S. 292,294,295, 296. 326, 327, 328, 337, 338, 342, 350 Ogawa, W. 128, 135 Ogden. G.B. 388, 398 Ogino, H. 418, 432 Ohmori, T. 197, 225 Ohno. Y. 17, 18. 74 Ohshima, T. 167, 187 Ohshiro, Y. 18, 22, 77 Ohtani, M. 403, 418, 428, 432 Okada, Y. 298, 348 Okamoto, K. 6, 67 Okumura, H. 9, 13, 14, 15, 19, 78, 198, 224 Okunuki, K. 298, 348 Oliveira, M. 50, 5 I , 78 Oliver, J.D. 146, 187 Olivieri, R. 145, I87 Olsen, A.N. 155. 182 Olsen, A.N. 303, 326, 344 Olson, J . 172, 173. 175, 185 Olsthoorn, A.J.J. 18, 19, 22, 25, 76, 77 Oltmann, L.F. 64, 66, 72, 79 Ornar, A. 129, 135 On, S.L.W. 144, I88 Onda, K. 407,435 ONeal, C.R. 238, 246, 248, 253, 256, 257, 261, 262. 264, 277 Onishi, S. 8, 77 Oobuchi, K. 423, 437 Oparin, A.I. 356, 398 Opekun, A.R. 141, 184 Opferkuch, W. 164, 165. 180 Oresnik, 1. 193, 226 Orgel, L.E. 356, 398 Orosz, D.E. 93, 134 Orosz, R.D. 298, 317,347 Ostermeier, C. 214, 225 O’Sullivan, W.J. 169, 186 Otto, B.R. 150, 189 Ouzounis, C. 329, 331, 348 Owen, R.J. 144, 188 Owens, R.J. 171, 173, 181 Padan, E. 406, 410. 411, 412, 435, 436 Page, M.G.P. 100, 133 Pallen, J.M.177, 181 Pallier, M. 241, 244, 245 Palmieri, F. 94, 133, 134 Pao, S.S. 84, 100, 109, 110, 129, 134, 135 Papaefthymiou, G.C. 292, 294. 301, 324, 326, 327. 329. 350

456

AUTHOR INDEX

Papendrecht, A. 60, 62, 70 Park, C. 409, 436 Park, J. 100, 126, 132, 133 Park, J.H. 98, 127, 134 Park, Y.K. 237, 239, 245, 247, 251, 278 Pas, E. 387,399 Pascher, T. 198, 230 Paster, B.J. 141, 183 Paszko-Kolva, H. 146, 187 Patschkowski, R. 195, 211, 216, 218, 229 Patschkowski, T. 216. 217, 218, 227, 228 Paula, S. 370, 398 Paulsen. I.T. 84, 100, 104, 107, 109, 110, 112, 121, 122, 126, 129, 130, 132. 133, 134, I35

Pavia, A.T. 236, 277 Paz, M.A. 6, 71 Pearman, J.W. 139, I85 Peekhaus, N. 130, 135 Pelzer, H. 374, 399 Pena, M.M.O. 315, 348 Penfold, C.N. 292, 316, 348 Penn, C.W. 145, 146, 169, 187 Perkins, K.E. 23, 64,76 Perry, R.D. 336, 348 Pesci, E.C. 153, 187 Pessolani, M.C.V. 329, 348 Peters, M. 140, 144, 183 Peters, T.J. 327, 328, 342, 350 Peterson, J.D. 140, 145, 146, 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176, 188. 403, 417, 419, 438 Peterson, S. 140, 145, 146, 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176, 188, 403, 417, 419, 438 Pettigrew, G . 175, 183 Pettigrew, G.W. 301, 346 Pfeifer, J.D. 155, 182, 303, 326, 344 Pfenninger-Li, X.D. 409, 437 Phibbs, P.V. 46, 73 Phillips, C.A. 304, 343 Phillips, R.L. 94, 136 Phillips, T.A. 295, 347 Pickett, C.L. 153, 187 Pierre, J.-L. 337, 339, 344 Pirkl, E. 345 Pitcher, M. 141, 184 Pittman, M.S. 148. 187 Pitts, K.R. 302, 348 Plagens, H. 345 Plank, R.W.H. 295, 349

Plass, R.J. 423, 434 Plaut, A.G. 172, 173, 175, 183, 185 Pognonec, P. 212, 224 Pollack, S. 337, 350 Pollak, P.T. 141, 189 Poole, R.K. 36, 76, 162, 170, 171, 173, 174, 175, 181, 185, 194. 195, 198, 199, 201, 202, 203, 208, 209, 210, 213, 215, 216, 219, 220, 221, 222, 224, 227, 229, 230 Poolman, B. 406, 437 Porath, J. 337, 340 Porter, A. 144, 188 Postma, 18, 42, 76 Postma, P.W. 18,22,47,49, 52, 53, 54, 55, 56, 57, 58, 59, 61, 72. 73, 75, 76, 79, 241, 277

Prakash, C. 177, 185 Pratt, M.J. 388, 398 Precigoux, G. 288, 319, 341, 344 Preisig, 0. 174, 187, 194, 195, 196, 205, 206, 210, 211, 212, 213, 214, 215, 216, 217, 218, 223, 224, 225, 227, 228. 229, 231 Price, A. 140, 185 Prickril, B.C. 313. 346 Priefer, U. 195, 215, 216, 218, 228 Priefer, U.B. 195, 21 I , 216, 217, 218, 227, 229 Prince, R. 423, 436 Prince, R.C. 298, 317, 344 Pronk, J.T. 70, 79 Pucci, M.J. 238, 274 Pujol, C. 17, 69 Putnoky, P. 219, 220, 226, 414, 437 Quackenbush, J. 140, 145, 146, 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176, 188, 403, 417, 419, 438 Quail, M.A. 332, 333, 334, 335, 341,348,349 Quan, J.A. 129, 135 Quandt, J. 195. 21 I , 216, 218, 229 Quax. W.J. 403, 438 Quayle, J.R. 8, 68 Queller, D.C. 355. 398 Quintella, J.C. 387, 3Y6 Quiocho, F.A. I 1 I . 135 Quirk, A.V. 295, 349 Quirk, P.G. 403, 423, 434, 437 Quitilla, F.X. 244, 277 Qureshi, M.H. 423, 424. 437 Rabinowitz, J.C. 161, 187 Raeburn, S. 161, 187

AUTHOR INDEX

Raguzzi, F. 314. 348 Ramakrishna, J. 172, 183 Ramamoorthi, R. 52, 53, 54, 55, 57, 59,60, 76, 77 Ramirez, B.E. 198, 228 Ramseier, T. 218, 225 Ramseier, T.M. 219, 220, 228, 238, 248, 249, 252, 278 Rapoport, G . 198, 228 Rappuoli, R. 142, 145, 148, 180, 181, 187 Rathbone, B.J. 143, 153, 188, 189 Ravenscroft, M. 139, 185 Read, N.M. 329,345 Reddy, V.N. 284,350 Reed, C.A. 298, 317, 347 Reeve, C.A. 238, 277 Regensburger, B. 199, 228 Reid, N.M.K. 329, 348 Reijnders, W.N. 424, 433 Reijnders, W.N.M. 64, 66, 72, 80 Reitzer, L.J. 244, 278 Reizer, A. 98, 128, 129, 135 Reizer, J. 98, 128, 129, 130, 135,238,248,249, 252, 278 Renalier, M.H. 212, 228 Repine, J.E. 284, 348 Resau, J.H. 177, 187 Rex, D. 151, 152, 186 Reynolds, D.J. 145, 146, 169, I87 Rhen, M. 249, 253, 257, 259, 261, 270, 271, 2 75 Rhode, M. 428,437 Rice, D.W. 288, 326,344,345 Rich, P.R. 197, 223, 409, 432 Richards, T. 298, 317, 344 Richards, T.D. 302, 348 Richardson, D. 140, 145, 146, 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176, 188, 403, 417, 419, 438 Richardson, I.W. 20, 21, 24, 76 Richter, O.M. 424, 437 Riedel, A. 423, 437 Riegman, N.H. 52, 53, 54, 55, 57, 75 Riesle, J. 431, 434 Riggs, D.L. 247, 273 Ringeling, P.L. 292, 298, 316, 336, 348 Ritz, D. 196,202,210,218,219,220,223,225, 228. 229 Rivoire, B. 329, 348 Robbe-Saule, V. 256, 271, 276 Roberto, F. 93, 132

Roberts, A.D. 156, 158, 159, 160, 164, 165, 170, 171, 181 Roberts, C. 236,278 Robertson, R.H. 298,342 Rocha, E.R. 302, 303, 348 Rodicio, R. 94, 133 Rodriguez, E. 143, 182 Rogers, R. 53, 58, 74 Rogers, R.D. 58, 68 Rogozinski, J. 10, 73 Rohrer, J.S. 326, 348 Roine, R.P. 162, 163, 187 Roitsch, T. 60,63, 77 Roland, F. 288, 344 Romanov, V.L. 194, 195, 199, 208, 226 Romslo, A. 337, 347 Rosen, B.P. 92, 93, 129, 132, I35 Rosenzweig, A.C. 321, 323, 348 Rossbach, S. 196, 206, 210, 223, 228 Roszak, D.B. 235, 278 Rouch, D.A. 100, 133 Roudier, C. 236, 239, 275 Rouf, M.A. 283, 348 Rouwenhorst, R.J. 79 Rowley, D.A. 284, 344 Royce, H. 139, 185 Rozmiarek, H. 141, I83 Riiberg, S. 195, 215, 218, 228 Rudd, K.E. 243, 274, 278, 295, 342 Rufo, R. 141, 183 Ruiz, B. 144, 182 Rumbley, J. 197, 198, 212, 225, 230, 424, 433 Rumley, M.J. 174, 183 Runswick, M.J. 94, I33 Rupp, M. 9, 11, 43, 72 Rusnak, F. 197, 230 Russell, D.G. 155, 182, 303, 326, 344 Russell, R.I. 143, 188 Rutherford, A.W. 423, 437 Sachs, G. 151, 152, 178, 185, 186 Sadowsky, M.J. 203, 227 Sadowsky, M.J. 206, 230 Saier, M.H. 17, 18, 48, 49, 62, 79 Saier, M.H. Jr. 84, 95, 96, 98, 100, 104, 107, 109, 110, 112, 119, 120, 121, 122, 126, 127, 128, 129, 130, 131, 132, 133, 134, 135, 136, 238, 248, 249, 252, 278 St Pierre, T.G. 326, 328, 349 Saitoh, M. 364, 399 Sakamoto, J. 423, 437

AUTHOR INDEX Sakazawa, C. 8, 77 Salardi, S . 202, 223 Salaspuro. M. 162, 163, 187 Salentin, A. 93, 132 Salisbury, S.A. 3, 77 Salmela, K.S. 162, 163, 187 Sanchez, F. 219, 221. 229 Sander, C. 329, 33 I , 348 Sanderson, K.E. 243, 278 Sangwan, 1. 195, 207, 224 Sano, H. 17, 77 Santana, M. 198, 228 Santos, H. 62, 73 Saraste, M. 197, 198, 205, 212, 223, 225, 226, 228, 230, 423, 424, 433, 436 Sarker, R.I. 128, 135 Sato, N. 295, 344 Sato, R. 292, 295, 344 Saurin, W. 112, 119, 121, 134, 135 Sayers, D.E. 326, 337, 347, 348 Scarlato, V. 148, 180 Schaechter, M. 388, 398 Schafer, K.P. 152. 178, 185 Schepper, M. 424, 433 Scherb, B. 217, 218, 222. 224 Schemer, P. 431, 432. 437 Schiltz, E. 153, 154, 188 Schindler, P. 284, 342 Schlunegger, M.P. 18, 19, 77 Schliiter, A. 195. 21 I , 215, 216, 217, 218, 227. 228, 229 Schmidt, K. 258, 274 Schmidt, T. 361. 397 Schneier, M. 419, 422, 434, 436 Schnider, U. 52, 53, 54, 77 Schopfer, L.M. 207, 227 Schorah, C.J. 153, 188 Schroeder, J.1. 130, 133 Schrover, J.M.J. 9, I I , 21, 37. 43, 77 Schuldiner, S. 100, 135, 406, 410, 411, 436 Schultz, J.E. 237, 238, 239, 252, 263, 276 Schulz, B. 95. 130, 132, 135 Schwarz, H. 387, 396 Schwdrz, M. 95, 135 Schwarz, U . 380, 385, 392, 396 Scott, D.R. 151, 152, 186 Scott, J . 304, 343 Segal, E.D. 177, 187 Seigele, D.A. 237, 238, 240, 241, 245, 276 Selinger, B. 195, 21 I , 216, 218. 229 Sellink, E. 52, 53, 54, 55, 56, 57. 59, 75, 79

Selwyn. M.H. 410, 417, 436 Seto, Y. 413, 414, 416, 419, 434 Seto-Young, D. 403, 419, 433 Seymour, R.L. 238, 245, 247, 248, 257, 259, 260, 261, 266, 268. 271, 278 Shahamat, M. 146, 187 Shapleigh, J.P. 197, 225 Shapleigh, M. 197, 225 Sharma, S.A. 142, 188 Sharp, G.A. 328, 342 Shaw, W.V. 288, 320, 323, 326, 346 Shell, L. 238, 257, 275 Sherman, N.A. 284, 342 Shibata, A. 49, 71 Shibata, T. 150. 186 Shimada, S. 197, 225 Shimamoto, T. 128, 135 Shimao, M. 8. 77 Shinagawa, E. 6, 7. 9, I I , 13, 14, 16, 17. 18, 19, 20, 21, 22, 24, 39, 40,42,43, 44.67, 74, 75, 77, 78 Shinagawa, E.A. 8 , 73 Shinzawa-ltoh, K. 214, 230 Shirley, R. 304, 343 Shirvan, A. 100, 135 Shockman, G.D. 372, 387, 396, 398 Shon, J. 177. 187 Sidhu, A. 4 0 4 , 4 0 9 , 4 3 4 Siegel, E. 238, 276 Sikkema, D.J. 336. 348 Silver. S. 92, 93, 129, 132, 135 Simons, J.A. 49, 61, 73 Singer, M. 243. 274 Sipponen, P. 144, 187 Sitas, F. 143, 182 Sjoberg, B.M. 321, 323, 348 Skulachev, V.P. 409. 428, 437 Skurray, R.A. 100, 104, 107. 129, 130, 133. 134, 135

Slaska-Kiss, K. 219, 220. 226 Slater, E.C. 195, 229 Slauch, J.M. 249, 253, 261, 262, 270, 276 Slaughter, C. 197, 225,299, 305, 310, 319.344 Sliwinski, M. 129, 135 Sliwinski, M.K. 84, 107, 112, 121, 122, 135 Slotboom, D.H. 424. 433 Sly, L. 140, 144, 183 Smith, A. 42, 48, 77 Smith, A.T. 48, 72 Smith, D.R. 329, 348 Smith, E.O. 141, 184

AUTHOR INDEX Smith.H.0. 140, 145, 146, 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176, 188, 403, 417,419, 438 Smith. J.M.A. 288, 292, 295, 297, 298, 299, 300, 301, 302, 305, 310, 317, 319, 320, 323, 326, 328, 334, 341, 343, 344, 346, 348. 349 Smith, J.S. 145, 180 Smoot, D.T. 177, 187 Snoep, J.L. 49, 61, 73 Sobala, G . M . 143, 153, 188 Sobek, J.M. 238, 274 Soberon, M. 194, 199,219,220,221,222,227, 229 Sode, K. 17, 77 Solioz, M. 417, 438 Sone, N. 173, 174, 186, 197, 225,423,437 Soruco, J . 288, 344 Sosnick, T.R. 51, 78 Souma, Y. 197, 225 Southgate, G . 62, 77 Sowerby, J.M. 325, 349 Sparrius, M. 150, 189 Spartalian, K. 292, 294, 301, 326, 327, 350 Spector. M. 236, 241, 244, 245, 246 Spector, M.P. 235, 237, 238, 239, 240, 245, 246, 247, 248, 251, 253, 256, 257, 259, 260, 261, 262, 263, 264, 265, 266, 268, 269, 270, 271, 275, 277, 278 Speelmans, G. 406,437 Sperling, R. 292, 295, 296, 338, 350 Spiegelhalder, C. 153. 154, 188 Spik, G . 150, 184 Spiro, S. 217, 229, 292, 316, 348 Spohn, G. 148, 180 Spratt, B.G. 398 Spriggs, T . 304, 343 Springer, A.L. 52, 53, 54, 55, 57,60,64, 76, 77 Spychal, R.T. 144, 183 Srinivasan, S. 169, 186 Stacey, A.R. 143. I82 Stalls, K. 141. 183 Stam, H. 194, 195, 199, 208, 229 Stammers, D . K . 288, 341 Stanley, J. 144, 188 Stark, R.M. 145, 176, 188 Stax, D. 196, 200, 201, 202, 229 Stefanini, S. 3 16. 342 Steidl, J . 202, 229 Steinhilber, W. 152, 178, 185 Steinkamp, T. 95, 135

459 Stenkamp, R.E. 321, 323, 345 Stephan, M.P. 50, 51, 78 Stewart, V. 247, 275 Stiefel, E. 292, 293, 294, 300, 326, 339, 349 Stiefel, E.I. 292, 294, 298, 299, 301, 305, 310, 317, 319, 326, 327, 344, 350 Stock, J.B. 7, 49, 70 Stojiljkovic, I. 334, 335, 349 Stolp, H.60, 63, 77 Stonehuerner, J. 202, 229 Stoorvogel, J. 9, 12, 21, 24, 31, 70, 71, 78 Storz, G. 257,259, 265, 268, 273, 278 Stouthamer, A . 64, 72 Stouthamer, A . H . 60, 62, 64, 66, 70, 72, 79, 80, 194, 195, 197, 199, 208, 212, 224, 229, 230,424, 433 Stratheran, S.L. 356, 395 Strauss, E.J. 58, 78 Streiff, M.B. 18, 19, 77 Streips, U.N. 387, 388, 390, 398 Strelow, L.I. 94, 136 Stretton, R.J. 238, 274 Studer, D. 199,219, 228 Sturr, M.G. 403, 404, 405, 406, 407, 417, 418, 421,424,425,428,435, 436,437 Sturrock, R.D. 143, 188 Subhan, H. 144, 182 Suerbaum, S. 147, 182, 185, 188 Sugden, E.A. 298, 342 Sugiyama, S. 408, 417, 432, 437 Sukharev, S.I. 129. 136 Suleiman, M.S. 145, 176, 188 Sulliven, T . D . 94, 136 Sumi, K . 80 Sun, J . 9, 12, 21, 70 Sunagawa, M . 9, 14, 73 Sunderman, F.W. Jr. 172, 183 Sung, J.J.Y. 140, 184 Sung, J.Y. 140, 188 Surpin, M.A. 194, 196, 198, 199,206,207,229 Susman, P. 405, 425, 434 Sutton, G. 304, 343 Sutton, G.G. 140, 145, 146, 148, 149, 150. 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176, 188, 403, 417, 419, 438 Sutton, H.C. 284, 349 Suzuki, I. 297, 342 Swings, J . 50, 78 Switala, J. 239, 246, 248, 249, 270, 275, 276, 277 Swords, W.E. 260, 269, 270, 273

460 Szczebara, L. 150, 182 Tabata, K. 412, 428, 435 Tabche, M.L. 221, 227 Taha, A S . 143, 188 Taha, S.M.198, 229 Tai, C.Y. 53, 58, 74 Tajuchi, Y. 150, 186 Takade, A. 304,350 Takagi, M. 9, 14, 73 Takahashi, K. 10, 16.41, 75 Takahashi, M. 16, 41, 75 Takahashi, S. 423, 438 Takaji, S. 150, 186 Takaki, Y. 14, 16, 40, 75 Takamune, K. 302, 335, 345 Takata, R. 302, 335,345 Takata, T. 304,350 Takemura, H. 9, 13, 14, 15, 19, 78 Takimoto, K. 18, 22, 67 Takova, T.Y. 419, 433 Talbo, G. 198, 226 Tam, R. 112, 119, 120, 129, 130, 135, 136 Tamaki, T. 9, 13, 14, 15, 19, 78, 198, 224 Tamogami, T. 8, 77 Tamura, K. 144, 188 Tamura, T. 173, 174, 186 Tan, K. 409,432 Tanaka, S. 167, 187 Tangeras, A. 337, 349 Tani, K. 417,437 Tao, J.S. 424, 437 Tardat, B. 334, 349 Tauxe, R.V. 236, 277 Tay, A. 174, 181 Tayama, K. 9, 13, 14, 15. 78, 198, 224 Taylor, J.D. 153, 188 Taylor, N.S. 143, 144, 182 Tecklenburg, M.J.197, 225 Tehrani, H.A. 333, 349 Teissie, J. 431, 433 Teixeira, K.R.S. 50, 51, 78 Teixeira-de-Mattos, M.J. 7, 49, 70 Tempest, D.W. 18, 22. 42, 47, 49, 59, 61, 72, 73, 76, 235, 278 Tenvoorde, G.J. 18, 47, 69 Terzaghi, B. 212. 223, 228 Terzaghi, E. 212, 223 Teuber, M. 291. 346 Texeira de Mattos, M.J. 18, 22, 69 Thauer, R.K. 161, I84

AUTHOR INDEX Theil, E.C. 285, 288, 326, 337, 347, 348, 349, 350 Theis, F. 164, 184 Thiberge, J.M.179, 184 Thiedemann, G. 431,434 Thomas, C.D. 288, 320, 323, 326, 346 Thomas, J.W. 197,225 Thompson, A. 329, 345, 348 Thompson, C. 326, 337,347 Thompson, N. 144, 182 Thomson, A.J. 292, 297,298, 299, 300, 301, 302, 311, 312, 317, 319, 324, 326, 332, 333, 341, 343,346, 347, 348, 349 Thony, B. 218, 225 Thony-Meyer, L. 195, 196,200,201,202,206, 210, 212, 213, 214, 216, 217, 218, 219, 220,223, 224, 225,228, 229, 231 Thwaites, J.J. 383, 399 Tibor, A. 298, 343 Timms, A.R. 333 Tipper, D.J.383, 399 Tirgari, S. 231, 239, 245, 247, 251, 278 Tobaqchali, S. 177, 181 Tobias, J.W. 262, 270, 276 Toczko, M. 10, 73 Tokuda, H. 409,438 Tomb, J.-F. 140, 145, 146, 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176, 188, 304,343, 403, 417, 419, 438 Tomizaki, T. 214, 230 Tompkins, D.S.141, 189 Tompkins, L.S. 177, 187 Tong, S. 130, I35 Tong, S.X.49, 71 Tonkovic, M. 337,349 Topping, A. 177, 181 Touati, D. 334, 349 Towe, K.M. 291, 326,349 Toyama, H. 8,9, 11, 13, 14, 16, 17, 18,22. 25, 39,40,43,44,47, 50, 53, 54, 55, 56, 75, 78 Tragesser, M. 21 7, 230 Trautwein, A.X. 337, 338, 347 Treffry, A. 288, 292, 320, 323. 325, 326, 327, 328, 342,344,345, 346, 347, 349 Triggs-Raine, B.L. 246, 276 Trikha, J. 288, 350 Tripier, D. 165, 184 Truchet, G. 212. 228 Trueba, F.J. 387, 399

46 1

AUTHOR INDEX Trumpower, B.L. 195, 229 Trust, T.J. 147, 155, 182, 184, 303, 343 Tsang, S. 105, 133 Tsieng, T.-T. 129, 135 Tsuchida, T. 19, 78 Tsuchiya, T. 128, 135, 197, 225 Tsugita, A. 295, 350 Tsujii, M. 144, 188 Tsujii, T. 144, 188 Tsukihara, T. 214, 230 Tsukita, S. 173, 174, 186 Tsygankov, Y.D. 52, 53, 54, 57, 71 Tuckwell, A. 169, 188 Tully, R.E. 196, 206, 210, 223, 230 Tummuru, M.K.R. 142, 188 Turbd, A. 424, 437 Turbett, G.R. 176, I88 Turk, A. 236 Turk, A.M. 238, 246.248, 253, 256,257, 261, 262,264, 277 Turlin. E. 52, 53, 54, 57, 58, 68, 69, 75. 76, 78 Turner, D.L. 62, 73 Turner, G.A. 148, 189 Turner, G.L. 209, 223 Turner, R.J. 130, 135 Tweedle, D. 139, 185 Tzagoloff, A. 94, 136 Umene, K. 304, 333, 350 Unden, G . 164, 165. 184, 185, 216, 217, 228, 230 Unemoto, T. 409, 438 Unkefer, C.J. 51, 73, 78 Urrutia, M.M. 429, 435 Utterback, T.R. 140, 145, 146, 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176, 188, 403, 417. 419, 438

van der Oost, J. 197, 198, 212, 226, 228, 230, 424, 433 van der Palen, C.J.N.M. 64, 66, 72 van Dijken, J.P. 18, 22, 42, 46, 47, 49, 59, 61, 70, 79 van Dijl, J.M. 42, 46, 49, 79 Vandooren, T.J.G.M. 9, 12, 71 van Dyck, S.J. 424, 433 Van Eijk, H.G. 337, 341 van Gelder, B.F. 195, 229 Van Gorkom, L. 160, 186 Van Heijenoort, J . 238, 276 van Hell, B. 47, 61, 72 Van Hock, A.N. 370, 398 van Horssen, P. 285, 342 van Kleef, M.A.G. 6, 7, 9, 12, 19, 21, 24. 44, 51, 53, 54, 55, 57, 59, 60, 70, 72, 73, 75, 78, 79 van Schie, B.J. 18, 22, 42, 46, 47, 49, 59, 61, 70, 79 Vansluis, C.A. 9, 12. 24, 31, 78 van Spanning, R.J.M. 64, 66, 72, 79, 80, 197, 212, 230, 424, 433 Van Strijp, J.A.G. 146, 184 Van Verseveld, W.H. 194, 195, 199, 208, 229 van Wielink, J.E. 9, 11, 19, 21, 37, 43, 70, 77 Van Wuytswinkel, 0.292, 298, 300, 312, 335, 346 van Zeeland, J.K. 16, 18, 70 Varela, M.F. 108, 136 Vargas, C. 194, 199, 201, 202, 203, 208, 209, 215, 216, 219, 220, 224, 230 Vargo, C.J. 329, 332, 344 Vasse, J. 212, 228 Vaughn, V. 295,347 Velayudhan, J. 151, 189 Veldhuyzen van Zanten, S.J.O. 165, 167, 168,

Valenti, P. 316, 342 van Belzen, R. 409, 437 van Berkum, P. 203, 227 Van Bogelen, R.A. 265. 268, 277 van Cauwenberhe, 0 . 193, 226 van den Berg, C. 285, 342 Vandenbroucke-Grauls, C.M.J.E. 146, 184 Vandentweel, W.J.J. 9, 12, 71 van de Putte, P. 17, 18. 19, 52, 53, 54, 55, 57. 61, 69, 72 Van der Hock, R.A.C. 403, 438 Van der Laan, J.C. 403, 438 van der Meer, R.A. 6, 78

Veldhuyzen van Zanten, S.Q.J. 141, 189 Velterop, J.S. 56, 57, 59, 79 Venter, C. 403, 417, 419, 438 Venter, J.C. 140, 145, 146, 148. 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176, 188 Vermaas, D.A.M. 53, 54, 72 Venver, R.W.H. 386, 398 Venviel, P. 6, 70 Venviel. P.E.J. 9, 10, 71 Viebahn, M. 60.63, 64. 79 Vilkov, A.G. 370, 398 Vinella, D. 238, 257, 261. 274

184

462 Vink, K. 17, 18, 19. 69 Viollet, S. 48, 72 Vogt, K. 172, I89 Voisard, C. 52, 53, 54, 77 Volcani, B.E. 130. 133 Von Dad, M. 285, 341 von Wachenfeldt. C . 198, 230 Wachi, S. 129, 135 Wagner, R. 95, 135 Wagner, S. 170. 171. 180 Wai, S.N. 304, 333,350 Wald, N. 143, 182 Waldo, G.S. 288, 350 Walker, A.R. 130, I32 Walker, J.E. 94, 133 Wallin, E. 140, 145. 146, 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176. 188, 403. 417. 419, 438 Wallmeier, H. 95, 96. 133, 136 Wallner, J.S.284, 342 Walsh, C.T. 245, 278 Walter. K.S. 150, I84 Wang, W. 406, 410, 433 Wanner, B.L. 237, 278 Wansell. C.W. 64, 79 Ward. F.B. 409, 432 Ward. S.P. 130, 132 Warne, A. 198, 230, 424, 433 Warrelmann, M. 172, 189 Warren, J.R. 139, 144, 185, 189 Waser, M. 417, 438 Wasfy, G . 154, 185 Wasserman, S.A. 245, 278 Watanabe, H. 364, 399 Watanabe, K. 17, 77 Watanabe, T. 417, 437 Waters, D . A . 419, 433 Watson, D.C. 298. 342 Watson, J.D. 393, 399 Watt, G.D. 292, 293, 294, 298, 300, 301, 302, 317, 324, 325, 326. 327, 329, 339, 344, 348, 349, 350 Watthey, L. 140, 145, 146, 148, 149, 150, 152. 154, 155, 162, 166, 167, 168, 169, 174, 175, 176. 188, 403. 417, 419, 438 Weaver, J . 337, 350 Webb, E.C. 84, 133 Webb, J. 326, 328, 349 Weber, A. 95. 96, 133, 136 Weber, J. 199, 228

AUTHOR INDEX Weber, P.C. 288, 350 Webster, D . A . 409, 436 Weidel, W. 374, 399 Weidenhaupt, M. 210, 218, 224, 225 Weidman, J.F. 304, 343 Weidman, J.M. 140, 145, 146, 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176, 188, 403,417, 419, 438 Weidner, S. 195, 211, 215, 216, 218, 228, 229 Weiner, J.H. 130, 135 Weir, M.P. 327, 328, 342, 350 Weisbeek, P. 285, 342 Weitzenegger, T. 152, 178, 185 Wensink, J. 238, 278 Wenzel, T.J. 17. 18, 69 Weppelman, R. 250, 276 Werber, M.M. 294. 342 West, A.P. 141, I89 Westblom, T.U. 155, 182, 303, 326, 344 Westerling, J . 6. 70 Wexler, M. 220, 221. 230 Weynants, V. 298, 343 White, J.L. 288, 326, 344 White, 0. 140, 145, 146, 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169. 174, 175, 176, 188, 304, 343, 403. 417, 419, 438 White. P.J. 156, 164. 165, 166, 167, 171, 172, 181

White, S. 24, 79 White, S.A. 24, 26, 79 Whiting, P.H.46, 79 Wieland, B. 154, 187 Wikstrom, M. 197, 198, 205, 222. 225, 226, 423, 436 Wilkens, E.G.L. 236, 278 Williams, H.D. 194, 199, 210, 221, 229, 230 Williams, J.M. 291, 292, 297, 299, 300. 301, 303, 304, 323, 325, 326, 328, 333, 334, 337, 341, 343, 345, 347 Williams, R.J.P.283, 326, 328. 337, 347, 349. 350 Willmitzer, L. 95, 135 Wilson, J.A. 259, 262, 271, 272, 279 Wilson, M.T. 319, 324, 346 Wilson, T.H. 108, 136 Winkler, J.R. 198. 228 Winter, H.W. 219, 220, 228 Winterbourn, C.C. 284, 349 Witholt, B. 238, 278 Witty, J.F. 193, 230

AUTHOR INDEX Woese, C. 364, 399 Woeste, S.W. 382, 383, 397 Woldringh, C.L. 387, 397, 399 Wolfe, R.S.366, 395 Wolfram, J.H. 53, 58, 74 Wolin, E.A. 366, 395 Wolin, M.J. 366, 395 Worst, D.J. 150, 189 Wren, B.W. 177, 181 Wrigglesworth, J.M. 283, 350 Wright, A. 172. 183, 383, 399 WU, G . I 94, I 99,20 I , 202.203,20n,209,2 IS, 2 16, 2 19, 220, 224, 230 Wu, M. 94, 136 WU, Y.-Q. 209, 223 Wullstein. L.H. 293, 344 Wyatt, J.I. 142, 143, 189 Wyborn, N.R. 149, 182 Xia, 2. 24, 26, 79 Xia, Z.X. 24, 79 Xiang, Z.Y. 142, 181 Xu, H.H. 60, 63, 64, 7Y XU, J . 244. 257. 258.279 Xu, Y. 239, 243, 245. 246, 248, 251, 253, 256. 261, 264, 265, 269, 275 Ya, T. 294. 341. 345 Yakushi, T. 14, 16, 75 Yarn, L.L. 141, 183 Yamada, M . 17, 18. 48, 49, 58, 62, 75. 79, 80 Yamada, Y. 8, 17, 18, 48, 49, 62, 73, 79, 80 Yamagata, Y. 364, 399 Yamaguchi, H . 214, 230 Yamanaka, K. 8, 80 Yamanaka, T. 292, 295, 297, 300, 344, 346, 404. 423, 424, 437, 438 Yarnashita, E. 214, 230 Yarnauchi, K. 150, 186 Yamazuki, N. 150, 186 Yang, H . 66, 80 Yano, K. 9, 14, 73 Yao, R. 147, 189 Yaono, R. 214,230 Yariv, J . 288, 292, 294, 295, 296, 299, 301, 317, 318, 323, 324, 326, 327, 328, 334. 337, 338, 342, 344, 349, 350 Yarnell, J.W.G. 143, 182 Yasuda, R. 126, 134

463 Yasutake, N. 17, 77 Yeo, T.C. 58, 68 Yeoman, K.H. 194, 199, 201, 219. 220,221, 224, 230 Yewdall, S.J. 285, 288, 298, 299, 302, 305, 310, 317, 319, 320, 323, 326, 341, 343, 345, 346 Yoshida, H . 17, 77 Yoshida, M . 126. 134 Yoshikawa, S. 214, 230 Yoshinga, F. 19, 78 Yoshiyama, H . 148, 186 Young, G.B. 129. 135 Young, J.P.W. 193, 230 Young, N.M. 298,342 Young, R. 129, 136 Youqi, T. 294, 341 Yu, C. 202, 229 Yu. L. 202, 229 Yumoto, I. 404, 423, 424, 437, 438 Yung, M.Y. 140. 184, I88 Zajic, J.E. 283, 350 Zamanigian, S. 44, 74 Zarnbrano, M.M. 237,238,240,241,245,276 Zatman, L.J. 3, 6, 9, 68 Zavala, D. 143, 182 Zemsky, J . 403, 412, 413, 416, 417, 421, 425, 428, 435 Zepu, Z. 294, 345 Zevenboom, W. 235.278 Zhang, Y. 24, 26, 79 Zhang, Y.F. 24, 79 Zhao, Z. 349 Zhengjiong, L. 294, 341 Zhou, L. 140, 145, 146, 148, 149, 150, 152, 154, 155, 162, 166, 167, 168, 169, 174, 175, 176, 188, 195, 211, 216, 218, 229, 403, 417, 419, 438 Zigler, J.S. 284, 350 Zimmerman, 2. 195, 208, 209, 215, 226 Zimmermann, B.H. 197, 230 Zuber, M. 219, 228 Zufferey, R. 213, 214, 216, 217, 218, 228, 229, 23I Zumft, W.G. 197, 212, 230 Zychlinsky, E. I5 I , 152, 185 Zygmunt, M.S. 298, 343

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Subject Index Figures and table references are shown in italic ABC see ATP-binding cassette (ABC) superfamily Absidia spinosa 286. 292, 300, 309, 31 1-13 acetic acid bacteria 39 roles of alcohol and glucose dehydrogenases 5&1 Acetobacter 9, 13, 19, 39, 50 Acetobacter diazotrophicus 50 Acetobacter methanolicus 10, 16, 38 Acetobacter pasteurianus I5 Acetobacter polyoxogenes 14, 15 N-acetylglucosamine 368, 375, 377, 379 N-acetylmuramic acid 368. 375, 376, 377, 379 Acinetobacter 17, 42 Acinetobacter calcoaceticus 1619, 18, 22, 25, 41, 52, 53, 54, 59, 61 glucose dehydrogenase in 47 adaptive acid tolerance response (ATR) 269 agrobacterium 215 Alcaligenes eutrophus 104, 309 alcohol dehydrogenase (ADH) 39, 162-3 calcium in 2 0 4 in acetic acid bacteria 5&1 secondary structure 32 structure and mechanism 3&5 type 1 1 I , 37 type I1 12-13, 24, 39 type 111 13-16, 15, 23, 24, 30-5, 39 alcohols electron transport chains in oxidation of 39 periplasmic quinoproteins that oxidize 4 3 4 aldehyde dehydrogenases 19-20 alkaliphile oxidative phosphorylation 427-32 alkaliphilic Bacillus species energetics 401-38 pH homeostasis 404-20, 405, 408 amine dehydrogenases 4

amino acid-auxin permeases (AAAP) 130 amino acid sequences of proposed polypeptide precursor of PQQ 52 amino acids 145, 369 amino acyl residues 99 8-aminolevulinic (ALA) synthase 207 ammonia assimilation in Helicobacter pylori 176 Anabaena 331 Anabaena variabilis 3 16, 331 anabolic pathways in Helicobacter pylori 169 animal ferritins iron core 3 2 6 9 sequence alignment 309 antimycin A 172 antiporters 41&17 Archaea 124, 353, 361-3, 366 classification of transport proteins 8 1-1 36 origin 3 6 4 5 archaebacteria 285 Archaeoglobus fulgidus 161, 286, 304 arsenical (Ars) efflux family 92, 93 arsenical (Ars) family transporters 129 ATP-binding cassette (ABC) superfamily 107, 109-21, 111-19, 123, 129 ATP-driven active transporters 87, 91. 131 ATP-driven permeases 124 ATP-gated cation channels (ACC) 129 autolysis 373 auxiliary transport proteins 87-8, 92, 13I Azorhizobium 193 Azorhizobium caulinodans 191, 194, 196. 199 Azorhizobium caulinodansJixGHIS operon 216-17 Azorhizobium caulinodansJixNOQP genes 21415 Azorhizobiuni tumefaciens 195 Azorhizobium tumefaciensJixNOQP genes 2 15

466

SUBJECT iNDEX

Azorhizobium caulinodans multiple oxidases

208-9 Azospirillum brmilense 331 Azorobacter chroococcirm 286. 292, 331 Azotobacter vinelandii 286, 292, 2934,300, 309, 315, 323,327, 328,329,331. 332 azurin oxidase 37 Bacillus, see also alkaliphilic Bacillus species Bacillus alcalophilus 42I , 425 Bacillusfirmus 404,405,405, 407,410-13, 413, 416,419,421,423,424, 425,426,

429 Bacillus lenrus 404,406,413,414 Bacillus megaterium 417 Bacillus stearorhermophilus 423,431 Bacillus subtilis 98, 148,286, 316,331. 369, 375,375, 380,381. 384.387,388, 409-1I , 414,415.416, 417,419,423,

43I Bocillus YN-2000404,424 Bacteria 124,129,353-99 classification of transport proteins 8 1-1 36 iron storage in see iron storage in Bacteria bacterial ferritins 302-5 iron core 326-9 sequence alignment 309 structure 320 bacterial wall elasticity 382 first bacterium 388-92 formation 367-74,369 growth aspects 382-8 non-growth aspects 374-82 porosity 380-2 bacterial wall fabric 373-9,376--9 in plane of stress 380 bactenoferritin-associated ferredoxin (Bfd) 281-2,286-7. 329-33 bacterioferritin heteropolymers 311-1 3 bacterioferritins 28I , 286-7. 293-302,296. 29Y. 340 haem group of 298-302 iron core 32&9 iron uptake in 323-6 native core properties 327 sequence alignment 309 properties of 292 structure 316-20,318 ubiquity 314

see also ferritin-bacterioferritinrubrerythrin (F-BR) superfamily Bocreroides fragilis 286, 302,303,309 bactoprenol 37&2, 371 bactoprenyl 373 Bfd-NifU-nitrite reductase Family 332 bioenergetic work 420,420 2',7'-bis-(carboxymethyl)-S carboxylfluorescein (BCECF) 406 BLAST program 99 Borderella pertussis 154 Bradyrhizobium 193 Bradyrhizobium japonicum 7,172,173,191-2, 194, 196, 198,200, 331 Bradyrhizobium japonicum aa,-type cytochrome c oxidase 2034 Bradyrhizobium japonicum coxA expression

2045 Bradyrhizobium japonicum CoxMNOP oxidase 205-6 Bradyrhizobium japonicum CoxWXYZ oxidase 206-7 Bradyrhizobium japonicum cytochrome bc, complex 199-201 Bradyrhizobium japonicum cytochrome ebb,type oxidase 212-13 Bradyrhizobium japonicum cytochrorne CycM

201-2 Bradyrhizobium japonicum fixNOQP genes

210-12 Bradyrhizobiurn japonicum JixNOQP-related JixGHlS operon 216-17 Brucella melitensis 286, 309 butanol 44

Ca2+ : cation antiporter (CaCA) family 129 calcium in alcohol dehydrogenase 2 M in methanol dehydrogenase 2 M CAMP : CRP complex 252-3 Campylobacter jejuni 139,282,286, 304,305. 30Y, 333,339 Campylobacter mucosalis 175 Campylobacter pyloridis 139 cancer promoters 144 carbon (C)-compound catabolic enzymes

241-5 carboxylic acid-dependent decarboxylationdriven active transporters 87 carcinogenesis 144 carrier-type facilitators 87.89-90

SUBJECT INDEX

467

cell division 387-92 channel-forming colicins 129 channel protein families 129 channel-type Facilitators 86-8. 89 chemiosmotic energization of bioenergetic work 420 Chlorella kessleri 100 chloride channel (CIC) family 128 citric acid cycle 160 in Helicohacrer pylori 1 6 6 8 Clostridium acetohutylicwn 286, 304 Clostridium fervidus 406 Clostridium perfringens 286, 309 COz requirement of Helicobacter pylori 168-9 Comamonas testosteroni 7, 9. 12, 21, 3 I , 39. 44

Competitive Exclusion Principle 356 copper-containing amine oxidases 4 coxA 2 0 3 4 Cu, redox centre 198 Cyanophora paradoxa 286. 309. 3 13 cyclic AMP (CAMP)238 cyclic AMP (CAMP)receptor protein (CRP) 248 cystic fibrosis transmembrane conductance regulator (CFTR) I I 1 cytochrome be, - aa, respiratory chain 199 cytochrome c biogenesis 218-21 cytochrome c oxidase 197 cytochrome chhl-type oxidase 21&14 cytoplasmic energy coupling, phylogenetic tree for 121 cytoplasmic membrane 370, 371, 372. 373 Darwinian evolution 356 decarboxylation-driven active transporters 9 1 Deinococcus radiodurans 286 Desulphovihrio vulgaris 285, 309 diaminopimelic acid 369 2,6-dichlorophenol-indophenol(DCPIP) 207 dimethyl sulphoxide (DMSO) 175 disaccharide pentapeptides 368-7 1 371, 376 divalent anion : sodium symporter (IIASS) family 129 divalent metal ions in glucose dehydrogenase 25 in PPQ-containing quinoproteins that oxidize alcohols 21 in PQQ-containing quinoproteins 20-6 in relation to glucose dehydrogenase 22 diversity. development 359-60 I

DNA replication 391, 391 Domains 353, 357, 358, 359 evolution 356-67 formation 3 9 4 5 splits 361-3, 365-6, 374 Dps families 3 1S 16 duodenal ulceration 143 electron flow-driven active transporters 87 electron transport chains in glucose dehydrogenase 41-2 in oxidation of alcohols 39 involved in oxidation of glucose 41 involving soluble alcohol dehydrogenases 37 Emericella nidulans 331 endoskeleton 362, 363 energy coupling mechanisms 1246, 126 energy transduction, advances 36&l Enrerococcus hirae 387, 388. 417 Entner-Doudoroff enzymes I59 Entner-Doudoroff pathway 44,48, 49 enzyme classification 84, 85 Enzyme Commission (EC) 8 I , 84, 86 classification system 85 epithelial Na' channels (ENaC) 129 Escherichia 42 Escherichiacoli7. 17, 18.22.25,31,41,55,61, 88, 98, 100. 104, 111, 112, 122, 123, 154, 155, 236, 282, 285, 286, 292. 2 9 4 7,296,299,300,303, 304,309, 315-20, 318. 323, 325-6. 327, 331, 332-5, 33740, 369, 375, 375, 380. 381, 382, 384, 387, 41 I , 424, 426 glucose dehydrogenase in 48-8 pqq genes 57-9 ethanol 44 ethanol dehydrogenases see alcohol dehydrogenases, type I eubacteria 285 Eukarya 353, 361-3, 365, 367 classification of transport proteins 8 1-1 36 Eukaryotes 124, 129, 285 eukaryotic emux systems 109 eukaryotic ferritins 288-91 eukaryotic-specific carrier families I30 evolution of Domains 35&67 evolutionary creative breakthroughs 393-4 evolutionary tree 357, 358 Exiguabacterium auranricum 4 10 exoskeleton 362, 363, 365, 368. 373

SUBJECT INDEX extracytoplasmic receptors, phylogenetic families of 120 extrusion, biophysics 370-2 fbcFH genes 199-201 ferritin-bacterioferritin-rubrerythrin (F-B-R) superfamily 305-1 I , 310 and Dps family 315-16 dinuclear iron sites 321 evolution from two-helix protein 313 ferritins 155, 285, 286-7 bacterial see bacterial ferritins eukaryotic 288-91 H-subunits 290 iron core formation 289, 3 2 5 4 prokaryotic 302 subunits 291 ubiquity 314 see also ferritin-bacterioferritinrubrerythrin (F-B-R) superfamily ferroxidase centres 321-3 &CHIS operon 21618 fixNOQP operon 217-18 flavin adenine dinucleotide (FAD) 99 flavoprotein dehydrogenases 4 fluorescein-labeled dextran 381 formate-nitrite porter (FNP) family 129 fumarate metabolism in Helicobacier pylori 163-6

GABA 244 gastric cancer 1 4 3 4 gastric metaplasia 143 gastric ulceration 143 gastritis 140, 142, 147 gastrointestinal ulceration 139, 140 Cause principle 361-3 genome analyses 1 3 1 genome sequence of Helicobacier pylori strain 26695 175 Gluconobacier 9. 13, 17, 39, 41. 42, 50 Gluconobacter suboxydans 9, 14, 15, 17, 21 glucose, electron transport chains involved in oxidation 41 glucose dehydrogenase (GDH) 15, 16-19. 23 divalent metal ions in 22, 25 electron transport chains in 41-2 in acetic acid bacteria 50-1 in Acineiobacter calcoaceiicus 47 in Escherirhia cofi 48-8 in Klebsiella pneumoniae 47-8

in pseudomonads 4 5 4 secondary structure 32 stacking interactions 33 structure and mechanism 3&5 synthesis 60-2 glucose metabolism 45 Helicobacier pylori 1 5 6 9 glucose 6-phosphate 156 glucose 6-phosphate dehydrogenase 156 glycolysis 357 Gram-negative bacteria 3, 35, 121, 123, 137, 175, 193, 388, 392-3 Gram-negative organisms, insertion into stress-bearing wall 3 8 4 6 Gram-negative sacculus 383 Gram-negative wall 355 Gram-positive bacteria 105, 121, 123, 304, 388 Gram-positive organisms 379 inside-to-outside growth 384 Gram-positive wall 355 GTP 337 guanosine 3’,5’-bis(diphosphate) (ppGpp) 238 haem-copper respiratory oxidases 195-8 haem-CuB bimetallic centre 197-8 haem group of bacterioferritins 298-302, 300 Haemophilus injluenzae 286, 304, 309 haemoquinoprotein lupanine hydroxylase 10 Hansenula polymorpha 331 heat shock protein (HSP) 178 Helicobacter pylori 137-89,172,173,286,303, 304, 309. 316, 331, 403. 417, 419 amino acid requirements 145 anabolic pathways in 169 as gastric pathogen 1 4 W associated disease 1 4 2 4 biology of 140 cagA gene 142 cellular features 144-5 characteristics 144-9 chemotaxis 147-9 citric acid cycle in 166-8 CO2 requirement of 168-9 composition of respiratory chain 1 7 1 4 early studies of metabolism 155-6 epidemiology 140-2 evolutionary 144 PaAB genes 147 fumarate metabolism in 163-6

SUBJECT INDEX glucose metabolism 156-9 growth requirements 144-5 ion homeostasis and its relationship to' acid tolerance 151-2 iron acquisition mechanisms 149-51 katA gene 154 microaerophilic nature 152-5 motility 147-9 nitrogen assimilation in 176 nitrogen metabolism in 176-9 . oxidative stress 153-5 pathogenicity 142 pH homeostasis in 152 POR and OOR 161-2 pyruvate metabolism 15p-61,163 respiratory chain 169-75. 173 seroprevalance 141 serum antibody tests 141 spiral to coccoid cell transition 145-7 substrate oxidation 16%71 succinate respiration in 171,172 taxonomy 144 terminal oxidase(s) in 1745 transmission 14&2 transport systems 149 two-component families 149 urease of 17&9 virulence factors 142 holobacterioferritin 338 Homo sapiens 100, 309 horse-spleen apoferritin 288 horse-spleen ferritin 294,328 Hyphomicrohium 21, 37,5 I , 62 integral membrane transport protein families

95-107 interleukin-8 (IL-8) 142 intracellular ferric reductases 339 intracellular iron metabolism 333-9 intracellular salts medium (ISM) 234 intramembrane helices 426 in vieo expression technology (IVET) 262 iron beneficial properties 283 biologically relevant features 283-5 countering problems of dependence 284-5 detrimental properties 2834 storage in bacteria 281-350 uptake in bacterioferritins 3234 iron core animal ferritins 326-9

469 bacterial femtin 3269 bacterioferritin 326-9 formation in ferritin 325-6 iron-storage proteins 305-16,333 within bacteria 314-15

2-keto-3-deoxy-6-phosphogluconate aldolase 159 2-ketogluconic acid 50 Klebsiefh 42 Klebsiella aerogenes 245 Klebsiella pneumoniae 18, 22. 51,52,53, 54, 54. 55,57. 5941,154,329,331 glucose dehydrogenase in 47-8 Kluyveromyces lactis 100 D-laCtiC acid 369 Last Universal Ancestor (LUA) 353,356,357, 358, 359,361, 363 Legionella pneumophifu 154,336 light-driven active transporters 87,91 lipid membrane 37&2 Listeria innocua 316,338 Listeria ivanovii 153 Listeria monocytogenes 58,1534 low-molecular mass molecules (LMMM) 335 low-molecular weight (LMW) iron pool 337-9 Magnetospirillum magnetotacticum 286, 309,

311-13 major facilitator superfamily (MFS) 97, J O S 5 , 107-9, 107, 108, 110. 123,128 major intrinsic protein (MIP) family 97, 98-9. 99,10.5. 127 malate-grown cells 418 mechanically driven active transporters 87,91 mechanosensitive channels with large conductance (MscL) I29 mesophilia 364 metabolic energy, early sources 359 Methanobacillus omelianski 366 Methanobacterium bryantii 366 Methanobacterium thermoautotrophicum 286, 304,309 Methanococcus jannaschii 122,123.286, 309,

314 methanogenesis 358,359,364,366,366 methanol dehydrogenase 1G11, 23,43 a2 dimer 34 azbz tetrameric structure 30 cub unit 28

47 0 methanol dehydrogenase (continued) calcium in 20-4 disulphide ring 31 electron transport systems 37, 38 model for expression 65 molecular mechanism of synthesis regulation 63-6 stacking interactions 33 structure and mechanism 26-30 synthesis 62-3 methyl-accepting chemotaxis proteins (MCPs) 147. 148 methyl transfer-driven transporters 13 1 Mefhylohacil1u.r 2 I Methylohacillus .f~agellatum53, 54. 57 Methylohacillus gl.vcogenes 20 Methylobacterium 21. 62. 63, 64. 66 Methylobacterium estorquens 10, 20, 26, 38, 51. 52. 54. 5 4 , 55, 56. 57, 59, 6 0 , 6 4 , 6 5 Methyloharterium orgunophilum 53, 54, 54, 57, 60. 6 4 , 66 Mefh.vlophaga murina 20, 21 Methylophilus 21 Me fhylophilus methylotrophus 38, 62 methylotrophs 9 electron transport chains 38 methyltransferase-driven active transporters 88, 92 middle finding strategies 387-92 mitochondria1 carrier family (MCF) 93, 94-5, 97, 105, 130 molecular phylogeny 81-136 monophyletic period 356-8, 357, 360 nirp gene 4 13- 16, d l 5 MxaA 2 3 4 MxaD 23 Mycohacrerium avium 286. 309 Mvcohacterium lrprae 286. 309. 329, 331 Mycohacterium paratuherculosis 286, 309 Mycohacrerium tuberculosis 286. 304 Mycoplasma 3 I4 Mycoplamia genifaliuni 122, 123, 286 Mycoplasma pneumoniae 287 myxothiazol 172 Na+-dependent active cycle 418 Na+-extrusion, respiration-dependent 423 Na' re-entry 417-18 N A D 162, 167, 175 NADH 43, 56, 126, 167, 171, 414, 423 NADP 162

SUBJECT INDEX NADP' 156 NADPH 44, 56, 156, 162, 167, 171 Neisseria gonorrhoear 287, 309, 31 1-13, 323, 323 Neisseria meningitidis 287 Neurosporu crassa 100, 331 nhaC gene 412-13. 413 NifU proteins 329-33, 331 Nitrohacrer winograd.vkyi 292. 300, 309 nitrogen fixation 210-12 nitrogen metabolism in Helicohacter pylori I7&9 non-steroidal anti-inflammatory drugs (NSAIDs) 143 Nostoc commune 331 oligopeptidoglycans 375, 386 ORFs 414 osmotic challenges 363 osmotic forces 363 osmotic stress 362 outer membrane channel-type facilitators (porins) 87 outer membrane porins (B-type) 91-2 oxidative phosphorylation 430 energetics 42&32 2-oxoglutarate : acceptor oxidoreductase (OOR) 161-2 Paracoccus 2 I Paracoccus denitrificans 31, 38, 62, 6 4 , 6 5 . 66, 206, 424 PBPs (penicillin binding proteins) 372, 377, 389 pentapeptide 385 pentapeptide disaccharides 375 PEP-dependent phosphoryl transfer-driven group translocators 87 peptidoglycan 367 peptidoglycan disaccharide 373 periplasmic alcohol dehydrogenases, type I and type I1 4 3 4 periplasmic quinoproteins that oxidize alcohols 4 3 4 permeases classification 8 4 6 primary categories 86 pH homeostasis 404-20, 405. 408. 427 in Helicobacter pylori I52 pha gene 4 13- 16, 415 phosphate-buffered saline (PBS) 159-60

SUBJECT INDEX phosphenolpyruvate (PEP) carboxykinase I68 phosphenolpyruvate (PEP) carboxylase 168 6-phosphogluconate 156 6-phosphogluconate dehydratase I59 6-phosphogluconate dehydrogenase I 56 phosphogluconolactonase I56 phosphotransferase systems 91 photosynthesis 359 phylogenetic families of extracytoplasmic receptors 120 phylogenetic trees I I I , I 19 for cytoplasmic energy coupling 121 Plectoneme boryanum 331 polymorphonuclear granulocytes (PMNs) I53 polyvinyl alcohol dehydrogenase 8 prokaryotic efRux systems 109 prokaryotic ferritins 302 prokaryotic genome sequence analyses 1 2 1 4 prokaryotic uptake systems 109 proteins, extrusion 372 protoporphyrin IX 293, 295, 298 pseudomonads, glucose dehydrogenase in 4 M Pseudomonas 7, 8, 10, 17, 18, 39. 41 Pseudomonas aeruginosa 9, I I , 21. 24, 37, 42, 43, 46, 61, 287, 292. 300. 309. 31 1-13, 323, 327, 328, 329 Pseudomonas cepacia 44, 46 Pseudomonaspuorescens 22, 53, 54, 331 Pseudomonas putida 9 , I I , 13, 39.43, 44, 287, 309 Pyrococrus furiosus I6 I pyrophosphate bond hydrolysis-driven transporters 13I pyrrolo-2 carboxylic acid 5 I pyrrolo-quinoline quinone (PQQ) 1-80 adducts 5 amino acid sequences of proposed polypeptide precursor of 5 2 biosynthesis genetics 52-7 exogenous, effect on bacterial growth 7 genes required for synthesis 53 identification 6-7 in bacteria 6 8 isolation 3 organization of genes in bacteria 54 origin of backbone 51-2 origin of carbon atoms 51 structure 4, 4 synthesis 51-9 model 55, 56

47 1 regulation 5 9 4 6 pyrrolo-quinoline quinone (PQQ)-containing quinoproteins importance of divalent metal ions in structure and function 2 M structure and mechanism 2 6 3 5 that oxidize alcohols 9 that oxidize glucose 18 pyruvate : acceptor oxidoreductase (POR) 161-2 pyruvate metabolism in Helicobacier pylori 159-61, 163 quinohaemoprotein 8, 24 quinohaemoprotein alcohol dehydrogenases (type I1 alcohol dehydrogenases) 12-13, 39 quinohaemoprotein alcohol dehydrogenases (type Ill alcohol dehydrogenases) 13-16, 39 quinol oxidase 197 quinoprotein alcohol dehydrogenases 10-1 3 type I 39 quinoprotein dehydrogenases 3 amino acid sequence alignment 27 factors affecting synthesis 60-6 physiological functions 42-5 1 PQQ-containing 7-20 quinoproteins in energy transduction 35-42 prosthetic groups 4, 6-7 wrongly identified as PQQ-binding domain 35 Rattus norvegicus 100 resistance-nodulation-cell division (RND) family 97, 104, 105, 129 respiration-dependent Na+-extrusion 423 respiratory chain 4 2 3 4 , 424, 430 rhizobia 191-231 non-cytochrome-containing branch of respiratory chain 207 respiratory chains 198-205 respiratory pathways 195 symbiosis-specificcytochromes 209-14 terminal oxidases 194 rhizobiaceae 193 rhizobial genes in cytochrome c assembly 220 in free-living respiration 196 in symbiosis-specificoxygen respiration 211

47 2 rhizobial mutants with altered oxidase activity and improved symbiotic nitrogen fixation 221 Rhizobium 193 Rhizobium etli 191, 199, 208 Rhizobium-legume symbiosis 195 Rhizobium leguminosarum 191 Rhizobium leguminosarum biovar viciae 194, 196, 220 Rhizobium leguminosarum biovar viciae and trifolii 199 Rhizohium leguminosarum biovar viciae and frifolii cytochrome d 208 Rhizobium leguminosarum biovar viciae cytochrome bc, complex 201 Rhizobium leguminosarum biovar viciae cytochrome CycM 202 Rhizobium leguminosarum biovar viciae symbiosis-specific oxidase 21 5-16 Rhizohium meliloti 7, 104, 195, 210, 212, 220, 414, 415, 416 Rhizobium trifolii 194 Rhizobium tropici 194. 196 Rhizobium tropici coxA 204 Rhodobacter capsulatus 287, 292. 309, 3 3 5 4 Rhodobacter etli 220 Rhodopseudomonas acidophila 8 , 43 Rhodopseudomonas sphaeroides 201, 287. 292, 300 Rhodospirilluni rubrum 287, 292, 361 ribulose 5-phosphate 156 rod elongation 3 8 2 4 rod-shaped bacteria 386-7 rpoS-regulon 2 5 3 4 rubrerythrins 286-7. 291-3 sequence alignment 309 see also ferritin-bacterioferritinrubrerythrin (F-B-R) superfamily ryanodine-inositol 1,4,5-triphosphate receptor Ca2+ channels (RIR-CaC) 129 Saccharomyces cerevisiae 98. 100, 104. 122, 314 Salmonella 267 Salmonella enterica 235 Salmonella typhimurium 2 3 5 4 see also starvation-stress response (SSR) Schistosoma mansoni 309 Shigella Jexneri 58 small multidrug resistance (SMR) family 130

SUBJECT INDEX

socioeconomic conditions 141 solute : sodium symporter (SSS) family 128, I28 Sphaerotilus natans 283 Staphylococcus aureus 92, 108 starvation-stress response (SSR) 233-79 acid tolerance 269-70 and long-term starvation survival 263-5 and resistance to other environmental stresses 265-70, 267 and Salmonella virulence 27&2 carbon-starvation-inducible crossresistance 266 H202 resistance 266-8 osmotolerance 269 physiologic changes during 237-8 polymyxin resistance 270 thermotolerance 268-9 starvation-stress response (SSR) loci C-starvation-inducible regulation 254-5 carbon-starvation-inducible loci 242-3 core 264-5 defined stresses/conditions 261 environmental and physiologic regulation 26&3 genetic regulation 252-60 intracellular environments 261-3 starvation-stress response (SSR) stimulon 239-51 carbon (C)-compound catabolic enzymes 241-5 known protective enzymes 246 regulatory proteins 248-9 respiratory enzyme systems 2 4 6 8 transport systems 240-1 unclassified 251 virulence functions 249-50 Streptococcus pyogenes 287, 3 14 succinate respiration in Helicobacter pylori 171, 172 Sulfolobus acidocaldarius 197 superoxoide dismutases (SODS) 153-4 symbiotic nitrogen fixation 221 Synechococcus 98. 3 15 Synechocystis 292, 300. 31 1-13, 329 Synechocystis PCC 6803 122, 124, 287, 30Y. 331, 335 synthesis 373 TCA cycle 47, 48 TCA cycle enzymes 246

SUBJECT INDEX

terminal oxidases 205-9 in Helicobacter pylori 174-5 thermophilia 364 Thermotoga maritima 161, 287, 304 Thermus thermophilus 197 TMPD 200, 213 TPQ (6-hydroxyphenylalanine or topa quinone) 4 transmembrane a-helical spanners (TMSs) 99, 105, 106, 106, 110, 123 Transport Commission (TC) 81, 86 transport modes 124-6 transport protein classification system 8 6 9 5 expanded and updated 131 transport protein families 88, 89-92 reconstructed histories 105 transport proteins, classification 81-136 transport systems, starvation-stress response (SSR) stimulon 24&1 transporter families substrate ranges 127, 127 topological features 97, 97 transporters of unknown classification92, 131 TRAP transporters 149

473 Treponema pallidum 287, 314, 316 trimer formation 385 triose phosphate translocator (TFT) family 93,96 tryptophan residues 29 TTQ (tryptophan tryptophylquinone) 4 turgor pressure 358

ubiquinol-cytochromec oxidoreductase I95 urease of Helicobacter pylori 1 7 6 9 vacuolating cytotoxin VacA 144 viable but non-culturable (VBNC) cells 146 Vibrio cholerae 287, 304 Vitreoscilla 409 voltage-sensitiveion channel (VIC) family 127 Wolinella succinogenes 164-5, 172 Xanthobacter 62 Yersinia pestis 336 Zymomonas mobihs 100

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