Handbook of Corynebacterium glutamicum

HANDBOOK OF

Corynebacterium glutamicum

1821_C000.fm Page ii Friday, February 25, 2005 11:17 AM

HANDBOOK OF

Corynebacterium glutamicum Edited by

Lothar Eggeling Michael bott

Boca Raton London New York Singapore

A CRC title, part of the Taylor & Francis imprint, a member of the Taylor & Francis Group, the academic division of T&F Informa plc.

Published in 2005 by CRC Press Taylor & Francis Group 6000 Broken Sound Parkway NW, Suite 300 Boca Raton, FL 33487-2742 © 2005 by Taylor & Francis Group, LLC CRC Press is an imprint of Taylor & Francis Group No claim to original U.S. Government works Printed in the United States of America on acid-free paper 10 9 8 7 6 5 4 3 2 1 International Standard Book Number-10: 0-8493-1821-1 (Hardcover) International Standard Book Number-13: 978-0-8493-1821-4 (Hardcover) Library of Congress Card Number 2004057912 This book contains information obtained from authentic and highly regarded sources. Reprinted material is quoted with permission, and sources are indicated. A wide variety of references are listed. Reasonable efforts have been made to publish reliable data and information, but the author and the publisher cannot assume responsibility for the validity of all materials or for the consequences of their use. No part of this book may be reprinted, reproduced, transmitted, or utilized in any form by any electronic, mechanical, or other means, now known or hereafter invented, including photocopying, microfilming, and recording, or in any information storage or retrieval system, without written permission from the publishers. For permission to photocopy or use material electronically from this work, please access www.copyright.com (http://www.copyright.com/) or contact the Copyright Clearance Center, Inc. (CCC) 222 Rosewood Drive, Danvers, MA 01923, 978-750-8400. CCC is a not-for-profit organization that provides licenses and registration for a variety of users. For organizations that have been granted a photocopy license by the CCC, a separate system of payment has been arranged. Trademark Notice: Product or corporate names may be trademarks or registered trademarks, and are used only for identification and explanation without intent to infringe.

Library of Congress Cataloging-in-Publication Data Handbook of corynebacterium glutamicum / edited by Lothar Eggeling and Michael Bott. p. cm. ISBN 0-8493-1821-1 (alk. paper) 1. Corynebacterium glutamicum--Handbooks, manuals, etc. 2. Glutamic acid--Biotechnology--Handbooks, manuals, etc. 3. Amino acids--Biotechnology--Handbooks, manuals, etc. I. Eggeling, L. II. Bott, Michael. QR82.C6H26 2004 579.3′73--dc22

2004057912

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and the CRC Press Web site at http://www.crcpress.com

Foreword The era of microbial amino acid production with Corynebacterium glutamicum and the scientific study of this organism began almost 50 years ago with its discovery as a glutamate-secreting bacterium. This microorganism is today one of the most important organisms in biotechnology and is used to produce about 2 million tons of amino acids per year, of which more than 1 million tons are accounted for by sodium glutamate, used as a flavor enhancer in the food industry, and more than 0.6 million tons by L-lysine, employed as a feed additive. This market volume is constantly expanding. In the case of L-lysine, the increase currently amounts to as much as 10% each year. Whereas until the early nineties, the producer strains for the various amino acids were largely obtained and improved empirically by random mutagenesis and selection, today detailed knowledge of the metabolic pathways and their regulation permits selective improvement of strains by metabolic engineering making use of genetic engineering tools. These studies yielded in part surprising and novel information for science and applications. Examples are the existence of specific amino acid export carriers as well as of cyclic fluxes within the anaplerotic reactions — findings that go far beyond C. glutamicum and amino acid production. One milestone was the genome sequencing of C. glutamicum and closely related bacteria. This means that genome-wide transcription and proteome studies can now be performed in order to increasingly elucidate global regulatory mechanisms and interactions within the cell. It is to be expected that findings from such studies will considerably accelerate the development of even more efficient producer strains. In view of the profound data basis, the extraordinary industrial significance, and the existing global analyses of C. glutamicum, this organism is ideal for further analysis and modeling in order to ultimately understand and further exploit its entire metabolic and regulatory potential. Those wishing to inform themselves about C. glutamicum have until now had to take recourse to individual papers, some of which are difficult to locate. It is therefore especially gratifying that all major findings on C. glutamicum are now available in a monograph for the first time. Since the past has demonstrated that research on C. glutamicum is profitable for both industry and science, and also in terms of "systems biology,” its continuation will certainly promote this success story, to which this book will undoubtedly make a major contribution. Hermann Sahm Institute of Biotechnology Forschungszentrum Jülich GmbH Jülich, Germany

Editors Lothar Eggeling is a member of the Institute of Biotechnology, Forschungszentrum Jülich, Germany. He directs the microbial amino acid production activities at the institute. Dr. Eggeling studied microbiology at the German Research Center for Biotechnology in Braunschweig, Germany, and obtained a doctorate in biotechnology from the same institution in 1977. After a postdoctoral fellowship at the University of Sheffield, U. K., he joined the Institute of Biotechnology, Forschungszentrum Jülich, in 1978. Dr. Eggeling is a member of the American Society for Microbiology, the German Society for Biochemistry and Molecular Biology, the Society for General Microbiology, the German Association for General and Applied Microbiology, and the Society for Bioscience and Bioengineering. He has acted as an international editor for the Japanese Journal of Bioscience and Bioengineering. Dr. Eggeling has received research grants from the European Union and the German Federal Ministry of Education and Research, as well as from private industry. He is the author or coauthor of more than 100 publications and book chapters and holds several patents. His current interests include molecular physiology of Corynebacterium and Mycobacterium, amino acid and vitamin production, metabolic engineering, export of metabolites, and microbial cell wall synthesis. Michael Bott is professor of biochemistry at the University of Düsseldorf and head of the biochemistry group at the Institute of Biotechnology, Forschungszentrum Jülich (Germany). Dr. Bott studied biology at the University of Marburg (Germany), where he graduated in 1985 with microbiology as his major subject. In 1987 he received his doctorate from the University of Marburg with a thesis on carbon monoxide metabolism in methanogenic bacteria. In 1989, with a fellowship from the Deutsche Forschungsgemeinschaft DFG (German Research Association), Dr. Bott joined the Institute for Microbiology at the Swiss Federal Institute of Technology (ETH) in Zürich. After genetic studies on the respiratory chain of rhizobia, he focused on the regulation of anaerobic citrate metabolism in enterobacteria. In 1998, he finished his Habilitation in microbiology at the ETH Zürich. In the same year, he was appointed Professor of Biochemistry at the University of Düsseldorf and head of the biochemistry group at the Institute of Biotechnology of Forschungszentrum Jülich. Dr. Bott is a member of the Vereinigung für allgemeine und angewandte Mikrobiologie (VAAM) and of the Zukunftsforum Biotechnologie der DECHEMA e.V. He has received research grants from the Bundesministerium für Bildung und Forschung BMBF (German Federal Ministry of Education and Research), the Deutsche

Bundessstiftung Umwelt (DBU — German Environmental Foundation), the DFG, and from private industry. He is the author or coauthor of more than 40 research papers and book chapters. His current research interests focus on the elucidation of central regulatory pathways in biotechnologically relevant bacteria and in the development of sustainable microbial production processes.

Contributors Dr. Brigitte Bathe Degussa AG, FA-FE-B Halle, Germany

Dr. Thomas Hermann Degussa AG, FA-FE-B Halle, Germany

Prof. Dr. Michael Bott Institute of Biotechnology Forschungszentrum Jülich GmbH Jülich, Germany

Dr. Masato Ikeda Department of Bioscience and Biotechnology Faculty of Agriculture Shinshu University Nagano, Japan

Dr. Andreas Burkovski Institute of Biochemistry University of Cologne Cologne, Germany Dr. Mamadou Daffé Molecular Mechanisms of Mycobacterial Infections UMR CNRS — Université Paul Sabatier Toulouse, France

Dr. J. Kalinowski Zentrum für Genomforschung Universität Bielefeld Bielefeld, Germany Dr. Ralf Kelle Degussa AG, FA-FE-B Halle, Germany

Dr. L. Eggeling Institute of Biotechnology Forschungszentrum Jülich GmbH Jülich, Germany

Dr. Eiichiro Kimura Ajinomoto Co., Inc. Institute of Life Science Kawasaki-shi, Japan

Prof. Dr. Bernd Eikmanns Angewandte Mikrobiologie Universität Ulm Ulm, Germany

Dr. Shukuo Kinoshita Tokyo Research Labs Kyowa Hakko Kogyo Co. Ltd. Tokyo, Japan

Dr. Jeremy Felce Division of Biological Sciences University of California at San Diego La Jolla, California, USA

Prof. Dr. R. Krämer Institute of Biochemistry University of Cologne Cologne, Germany

Dr. Albert A. De Graaf Department of Surgery Maastricht University Maastricht, The Netherlands

Prof. Dr. Heung-Shick Lee Department of Biotechnology Korea University, Jochiwon Choongnam, Korea

Philip A. Lessard Department of Biology Massachusetts Institute of Technology Cambridge, Massachusetts, USA

Dr. Steffen Schaffer Institute of Biotechnology Forschungszentrum Jülich GmbH Jülich, Germany

Prof. Dr. W. Liebl Microbiology and Genetics Georg-August-Universität Göttingen, Germany

Anthony J. Sinskey Department of Biology Massachusetts Institute of Technology Cambridge, Massachusetts, USA

Prof. Dr. Nic D. Lindley Centre de Bioingenierie Gilbert Durand Institut National des Sciences Appliquees Toulouse, France

Dr. A. Tauch Zentrum für Genomforschung Universität Bielefeld Bielefeld, Germany

Dr. Axel Niebisch Institute of Biotechnology Forschungszentrum Jülich GmbH Jülich, Germany Dr. Susanne Morbach Institute for Biochemistry University of Cologne Cologne, Germany Dr. Miroslav Pátek Institute of Microbiology Academy of Sciences of the Czech Republic Praha, Czech Republic Dr. Oscar Reyes Institute of Genetics and Microbiology University of Paris XI Orsay, France

Dr. Volker F. Wendisch Institute of Biotechnology Forschungszentrum Jülich GmbH Jülich, Germany Dr. Laura B. Willis Department of Biology Massachusetts Institute of Technology Cambridge, Massachusetts, USA Dr. Brit Winnen Institute for Microbiology ETH Zentrum Zürich, Switzerland Dr. Christoph Wittmann Biochemical Engineering Saarland University Saarbruecken, Germany

Prof. Dr. Milton H. Saier Jr. Division of Biological Sciences University of California at San Diego La Jolla, California, USA

Dr. Atsushi Yokota Lab of Microbial Resources and Ecology Graduate School of Agriculture Hokkaido University Sapporo, Japan

Prof. Dr. G. Sandmann Botany Institute Goethe Universität Frankfurt, Germany

Dr. Hideaki Yukawa Research Institute of Innovative Technology for the Earth Kyoto, Japan

Introduction This book is concerned with Corynebacterium glutamicum, a bacterium that was discovered because it has the pleasant characteristic of excreting a substance that enhances the flavor of many foodstuffs, namely the amino acid L-glutamate. In view of the almost 1,000 scientific studies that have been published since the first description of C. glutamicum and the major significance of this bacterium in industrial amino acid production, we are convinced that it deserves a whole monograph of its own. We have been fortunate enough to persuade scientists from both industry and research, all of them acknowledged experts in their fields, to contribute to this book and we would like to take this opportunity to thank them for their willingness to become involved. The aim of this book is to provide, for the first time, a comprehensive representation of C. glutamicum and its special properties comprising genetics, biochemistry, physiology, and applications. Two concerns were of particular importance to us, namely the most exhaustive possible inclusion of the literature on the subjects presented and also intensive utilization of the genome sequence, especially for aspects that have not yet been analyzed experimentally. One example of this is the bioinformatic analysis and classification of all the transport proteins of C. glutamicum. The book is aimed at all those who wish to become acquainted with C. glutamicum and also at those readers whose purpose is to obtain an overview of a specific area. Due to the special significance of C. glutamicum for biotechnology, one of our other concerns was that the book should include an experimental section. By making use of the instructions it is not only possible to present C. glutamicum theoretically but also to employ it for practical teaching purposes. The experiment on glutamate secretion is undoubtedly a classic for laboratory courses in biotechnology. Other experiments describe the practical refinements in handling C. glutamicum, for instance in order to generate directed mutants. The gratifying "user friendliness" of C. glutamicum is immediately apparent. C. glutamicum is fast-growing and achieves incredibly high cell densities, which are initially rather astonishing for those of us used to working with E. coli. Even more important, however, is the wide range of reliable techniques that are available for genetic modifications with C. glutamicum. Furthermore, the organism is apathogenic and is classified as GRAS (generally regarded as safe). Apart from the biotechnological aspect, research with C. glutamicum is of significance for another reason. This species belongs to the suborder Corynebacterianeae, which includes such important and difficult-to-handle bacteria as Mycobacterium tuberculosis, as well as the comparatively little-studied Rhodococcus genus. The relatively small genome (3,000 kb) combined with the genome sequences of the related Corynebacterium and Mycobacterium species that are now available, as well as the above-mentioned advantages for experimental work, make C. glutamicum an

ideal model organism for investigating fundamental properties of the Corynebacterianeae, such as the synthesis and function of the outer membrane, which is otherwise found only with Gram-negative bacteria. We hope that we have attracted your attention to both the book and C. glutamicum itself, and that you will enjoy reading all about this fascinating organism.

Table of Contents PART I History Chapter 1

A Short History of the Birth of the Amino Acid Industry in Japan ..................................................................................................3

S. Kinoshita

PART II Taxonomy Chapter 2

Corynebacterium Taxonomy .............................................................9

W. Liebl

PART III Chapter 3

Genome, Plasmids, and Gene Expression The Genomes of Amino Acid–Producing Corynebacteria .............37

J. Kalinowski Chapter 4

Native Plasmids of Amino Acid–Producing Corynebacteria..........57

A. Tauch Chapter 5

Regulation of Gene Expression.......................................................81

M. Pátek Chapter 6

Proteomics........................................................................................99

S. Schaffer and A. Burkovski

PART IV Transport Chapter 7 M. Daffé

The Cell Envelope of Corynebacteria ...........................................121

Chapter 8

Genomic Analyses of Transporter Proteins in Corynebacterium glutamicum and Corynebacterium efficiens ..................................149

B. Winnen, J. Felce, and M.H. Saier Jr. Chapter 9

Export of Amino Acids and Other Solutes ...................................187

L. Eggeling

PART V

Physiology and Regulation

Chapter 10

Central Metabolism: Sugar Uptake and Conversion ....................215

A. Yokota and N.D. Lindley Chapter 11

Central Metabolism: Tricarboxylic Acid Cycle and Anaplerotic Reactions........................................................................................241

B. Eikmanns Chapter 12

Metabolic Flux Analysis in Corynebacterium glutamicum ..........277

C. Wittmann and A.A. De Graaf Chapter 13

Respiratory Energy Metabolism....................................................305

M. Bott and A. Niebisch Chapter 14

Nitrogen Metabolism and Its Regulation ......................................333

A. Burkovski Chapter 15

Sulfur Metabolism and Its Regulation ..........................................351

H.-S. Lee Chapter 16

Phosphorus Metabolism ................................................................377

V.F. Wendisch and M. Bott Chapter 17

Vitamin Synthesis: Carotenoids, Biotin, and Pantothenate ..........397

G. Sandmann and H. Yukawa Chapter 18

Osmoregulation..............................................................................417

S. Morbach and R. Krämer

PART VI Synthesis and Production Chapter 19

L-Glutamate

Production .................................................................439

E. Kimura Chapter 20

L-Lysine

Production .......................................................................465

R. Kelle, T. Hermann, and B. Bathe Chapter 21

L-Tryptophan

Production ...............................................................489

M. Ikeda Chapter 22

Synthesis of L-Threonine and Branched-Chain Amino Acids......511

L.B. Willis, P.A. Lessard, and A.J. Sinskey

PART VII Experiments Chapter 23

Experiments ...................................................................................535

L. Eggeling and O. Reyes Index......................................................................................................................567

Part I History

1

A Short History of the Birth of the Amino Acid Industry in Japan S. Kinoshita

CONTENTS Introduction................................................................................................................3 Role of Monosodium Glutamate...............................................................................4 References..................................................................................................................5

INTRODUCTION In 1956, we started a research program at Kyowa Hakko Kogyo Co., Ltd., Tokyo, that was aimed at obtaining a microorganism that could accumulate glutamic acid extracellularly. Among many isolates we found a colony that might be fit for the purpose. We named this isolate Micrococcus glutamicus No. 534. Further study revealed that this microorganism could accumulate glutamic acid at a limiting concentration of biotin present in the medium. This suggested that biotin must play a key role in the physiology of the cells and their glutamate-forming capability. By microscopic observation of cultures at various stages, we found that the cell form can change considerably. For this reason, and due to further taxonomical studies, we renamed the bacterium Corynebacterium glutamicum. From mutational work on this organism, together with discoveries regarding key regulatory features, it was found that many amino acids, such as lysine, arginine, ornithine, threonine, etc., could be accumulated. Most of these amino acids are now produced commercially. Amino acids produced by such a process are all in their natural (L) form, and this gives microbial production a big advantage over chemical synthesis. Thus, a new industry called amino acid fermentation was born. The commercial production of amino acids up to the discovery of C. glutamicum had relied on the decomposition of natural protein and the isolation of its constituent amino acids. Our new process, on the contrary, was a biosynthetic process using carbohydrate and ammonium ions. Therefore our process can contribute to the amino acid supply, and also helps to increase the absolute amount of protein in the world. Since the world population continues to increase year by year, so will the demand for amino acids and protein.

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Handbook of Corynebacterium glutamicum

After World War II, two new fermentation industries were born in Japan. These are the amino acid and nucleotide fermentation industries.

ROLE OF MONOSODIUM GLUTAMATE I would like to explain briefly why amino acid production was born in Japan, and to do so we have to go back to the year 1908. At that time Prof. Kikunae Ikeda at the University of Tokyo found that monosodium glutamate (MSG) had a potent taste-enhancing power [1]. He found this phenomenon through a careful examination of the decomposition products of konbu, a type of seaweed. During these studies he found a small crystal. This was glutamic acid, which he discovered had a sour taste. Then he added NaOH to a glutamic acid solution and tasted again. Surprisingly, it had changed into a beautiful taste. That was the aim of his studies, since he was searching for the potent essence of a flavor or taste enhancer. By the addition of only a few milligrams of MSG to various foods, their taste was noticeably improved. What a splendid achievement this was! Here, we have to consider the original ideas that led him to conduct such research. His real intention was to improve nutrition and increase the short life expectancy of the Japanese at that time. However, to provide large amounts of microbial proteins competitive with natural protein sources like soy or wheat protein was economically impossible. He thought it over, searching for a good idea to relieve malnutrition in Japan. He finally got the idea that even if the same food was eaten, its value might be increased if the taste is enhanced. In this sense, an improvement in taste might contribute to relieving malnutrition. Therefore, he began to search for the essence of good taste. Konbu had been traditionally used in Japanese food as a taste enhancer, so he believed it should contain the essence of flavor. This led to the discovery of MSG, whose commercial production was essential to make use of its taste-enhancing properties for the daily food of the Japanese. Mr. Saburosuke Suzuki was the man who supported Prof. Ikeda’s desire. Wheat gluten was chosen as the raw material to obtain MSG. But this task was very difficult. Concentrated HCl must be used for decomposition of gluten, but no anticorrosive vessels were available in those days. So clay pots were used, but they were fragile and their use was very dangerous. Moreover, the gas from HCl caused serious damage to the health of the residents living near the factory. He had to face an onslaught of accusations and complaints. Consequently, he had to move his factory to a remote location. His struggle to produce MSG continued for ten years, before he finally became confident of commercial success. Once MSG appeared in the market, its miracle power overwhelmed the food market and it became an essential food additive. Mr. Suzuki’s company is now known as Ajinomoto Co., Inc. After World War II, Dr. Benzaburo Kato set up Kyowa Hakko Co., Inc. in 1945. Because of the shortage of food, the Japanese suffered great hunger. Everywhere malnourished patients were seen. Dr. Kato was deeply worried by this situation and thought of an idea for relieving the miserable situation by supplying plenty of protein as food. To implement his idea, he asked me to establish a commercial process that could supply food protein by a fermentation process. “To produce food protein by a fermentative process?” I couldn’t believe my ears. I was deeply impressed by his

A Short History of the Birth of the Amino Acid Industry in Japan

5

sincere desire to relieve malnutrition in Japan, but it was impossible to produce protein in a price range that was competitive with natural proteins. If protein production was not feasible, then how about amino acids? Their nutritive value is very similar, so my judgment was that an attempt to produce amino acids may not be the wrong choice. Thus, our challenging program started and was finally successful as described above. A review of the amino acid and nucleotide fermentation industry in Japan is available in the literature [2]. This is the background of the birth of the amino acid industry in Japan. As was shown, Dr. Ikeda’s and Dr. Kato’s original motivation was the same: relieving the malnutrition of the Japanese. It is interesting to note that the answer to this problem comes out as two entirely opposite processes, one decomposition and the other biosynthesis.

REFERENCES 1. Ikeda, K. New seasonings [translation]. Chem. Senses 27:847–849 (2002). 2. Kinoshita, S. Thom Award Address. Amino acid and nucleotide fermentations: From their genesis to the current state. Developments in Industrial Microbiology 28:1–12 (1987).

Part II Taxonomy

2

Corynebacterium Taxonomy W. Liebl

CONTENTS 2.1

Position of Corynebacterium and Closely Related Genera within the Class Actinobacteria ........................................................................................9 2.2 Taxonomy and Characteristics of the Genus Corynebacterium ...................13 2.3 Methods for the Isolation, Identification, and Differentiation of Corynebacteria ...............................................................................................16 2.3.1 Morphology, Microscopic Appearance, and Staining Properties......20 2.3.2 Cell Wall.............................................................................................20 2.3.3 Mycolic Acids and Other Lipids .......................................................20 2.3.4 DNA and 16S rRNA Gene Analysis .................................................21 2.3.5 Physiological Properties.....................................................................21 2.4 Isolation, Classification, and Taxonomy of Industrially Relevant Corynebacteria ...............................................................................................21 2.4.1 Corynebacterium glutamicumAL ........................................................22 2.4.2 Corynebacterium callunaeAL..............................................................24 2.4.3 Corynebacterium efficiensVP ..............................................................24 2.4.4 ‘Corynebacterium thermoaminogenes’..............................................25 2.4.5 Corynebacterium ammoniagenesVP....................................................25 2.5 Conclusion......................................................................................................26 Acknowledgments....................................................................................................26 References................................................................................................................27

2.1 POSITION OF CORYNEBACTERIUM AND CLOSELY RELATED GENERA WITHIN THE CLASS ACTINOBACTERIA The large group of Gram-positive bacteria that have DNA with a high G+C content (above 50 mol %, with a few exceptions) are subsumed under the actinomycetes subphylum of Gram-positive eubacteria [111]. This group, which represents one of the main lines of descent within the domain Bacteria, was originally described on the basis of 16S rRNA cataloging [112] and represents an evolutionary line phylogenetically distinct from the low G+C Gram-positive bacteria. 9

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Handbook of Corynebacterium glutamicum

Early chemotaxonomic studies based on cell wall composition (cell wall chemotype IV sensu Lechevalier and Lechevalier 1970 [67]: peptidoglycan structure with meso-diaminopimelic acid (meso-A2pm); a polysaccharide fraction of the wall rich in arabinose and galactose), the occurrence of mycolic acids (2-alkyl-3-hydroxy acids), and lipid profile analysis suggested that the genera Corynebacterium, Mycobacterium, Nocardia, and Rhodococcus are closely related, which led to the proposal to combine the four in the so-called “CMN group” [2]. Different classification concepts were used later to accommodate these taxa. It was suggested to join these genera and Caseobacter (later transferred to Corynebacterium [26]) in the family Mycobacteriaceae [3,51]. In a different classification scheme, the mycolate-containing, cell wall chemotype IV actinomycetes were combined in the family Nocardiaceae while the genera Corynebacterium and Mycobacterium were treated separately (see [44]). Despite some variation in the classification models used over the decades, in principle, the chemotaxonomy-based classification using the markers mentioned previously (type IV cell wall chemistry, mycolic acids) proved to be correct. It is in agreement with phylogenetic analysis using methods of modern molecular systematics, i.e., 16S rDNA/rRNA sequence comparison, and now it is clear that the CMN group, which from today’s standpoint encompasses the genera Corynebacterium, Dietzia, Gordonia, Mycobacterium, Nocardia, Rhodococcus, Skermania, Tsukamurella, Williamsia, and the mycolate-less Turicella forms a robust monophyletic taxon [10,52,97,102]. One of the major problems that repeatedly led to different groupings of actinomycetes at the level of families and higher taxa was that various phenotypic characteristics such as morphological, physiological, and chemotaxonomic properties were used in various combinations as the basis for the proposal of the higher taxa. However, although the use of the mentioned properties is very useful and in general in accord with molecular phylogenetic clustering at the genus level, the diversity of morphology, physiology, and chemical composition can be very high between different genera or higher taxa, even between phylogenetically closely related genera. As a consequence, the description of higher taxa on this basis is problematic because (i) an exceedingly broad description of a family or higher taxon can become meaningless for the description of the enclosed taxa [111], and (ii) this sometimes can result in a relatively high degree of ambiguity in the grouping of genera into families or of families into higher taxa. Today’s picture of the position of the corynebacteria in the classification of bacteria is based on a new hierarchal classification structure for the taxa of the actinomycete line of descent that was brought forward a few years ago. Stackebrandt et al. [111] proposed that the delineation of actinomycete taxa should be based solely on 16S rRNA/rDNA sequence–based phylogenetic clustering and the presence of taxon-specific 16S rDNA/rRNA signature motifs. In this translucent and phylogenetically meaningful classification concept, the new class Actinobacteria, whose members share >80% 16S rDNA/rRNA sequence identity, and a hierarchically branched system of lower taxa was proposed, including subclasses, orders, suborders, and families containing one or more genera. The hierarchy leading to the genus Corynebacterium is (Figure 2.1) class Actinobacteria — subclass

Corynebacterium Taxonomy

Class Actinobacteria

Subclass Acidimicrobidae Rubrobacteridae Coriobacteridae Sphaerobacteridae Actinobacteridae

11

Order

Suborder

Family

Acidimicrobiales Rubrobacterales Coriobacteriales Sphaerobacterales Bifidobacteriales Actinomycetales Actinomycineae Micrococcineae

Acidimicrobiaceae Rubrobacteriaceae Coriobacteriaceae Sphaerobacteriaceae Bifidobacteriaceae Actinomycetaceae Micrococcaceae Brevibacteriaceae Cellulomonadaceae Dermabacteriaceae Dermatophilaceae Intrasporangiaceae Jonesiaceae Microbacteriaceae Promicromonosporaceae Corynebacterineae Corynebacteriaceae Dietziaceae Gordoniaceae Mycobacteriaceae Nocardiaceae Tsukamurellaceae Micromonosporineae Micromonosporaceae Propionibacterineae Propionibacteriaceae Nocardioidaceae Pseudonocardineae Pseudonocardiaceae Streptomycineae Streptomycetaceae Streptosporangineae Streptosporangiaceae Nocardiopsaceae Thermomonosporaceae Frankiaceae Frankineae Acidothermaceae Geodermatophilaceae Microsphaeraceae Sporichthyaceae Glycomycineae Glycomycetaceae

FIGURE 2.1 Hierarchic classification system of the class Actinobacteria according to Stackebrandt et al. [111].

Actinobacteridae — order Actinomycetales — suborder Corynebacterineae — family Corynebacteriaceae. The cell wall chemotype IV, mycolic acid–containing actinomycetes genera fall into the suborder Corynebacterineae which consists of the families Corynebacteriaceae (consisting of the genera Corynebacterium and Turicella), Dietziaceae (consisting of the genus Dietzia), Gordoniaceae (consisting of the genus Gordonia), Mycobacteriaceae (consisting of the genus Mycobacterium), Nocardiaceae (consisting of the genera Nocardia and Rhodococcus), Tsukamurellaceae (consisting of the genus Tsukamurella), and the genera Williamsia and Skermania [10,52,111]. Patterns of 16S rDNA signatures characteristic for the families just mentioned were described by Stackebrandt et al. [111]. The following pattern of 16S rDNA/rRNA signature nucleotides and nucleotide pairs defines the family Corynebacteriaceae: 293-304 (G-U), 307 (A), 316-337 (U-G), 468 (U), 508 (U), 586-755 (U-G), 631 (G), 661-744 (G-C), 662-743 (U-G), 771-808 (A-U), 824-876 (C-G), 825-875 (G-C), 837-849 (G-U), 843 (C), and 1059-1198 (U-A). The important role that the cell wall chemistry and lipid composition (in particular mycolic acid prevalence) played in the development of meaningful classification concepts for corynebacteria and related genera, long before molecular systematic

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Handbook of Corynebacterium glutamicum

methods were routinely available, is striking and deserves further attention. The conservation of cell walls of chemotype IV in concert with the occurrence of unique cell wall lipids — the mycolic acid esters — throughout a whole phylogenetically defined taxon at the subclass level is intriguing because there may be a connection between this phylogenetically conserved cell wall chemistry and an important physiological function. In this context, recent findings about the cell wall ultrastructure of corynebacteria and mycobacteria, which led to the idea that the members of Corynebacterium and related genera have an unusual cell envelope structure when compared to other Gram-positive bacteria, are noteworthy. This is discussed in detail in Chapter 7. The cell wall of corynebacteria contains an arabinogalactan polysaccharide, which is partially esterified by mycolic acids, and is covalently linked to the A1γ-type [103], directly cross-linked peptidoglycan. The Corynebacterium arabinogalactan may contain significant amounts of mannose and glucose. Additionally, high- and low-molecular mass glucan, arabinomannan, lipoglycans, and a protein surface layer are present in the cell walls of corynebacteria [41,98,100,108,113]. Even more important, the cell walls of Corynebacterium and related Grampositive genera contain a hydrophobic layer [85,91] that has been shown to play an important role in drug and substrate permeability [50,99]. According to recent data, this outer lipid layer apparently forms a structure reminiscent in architecture and function of the characteristic outer membrane of Gram-negative bacteria, although the molecular details are strikingly different. Whereas the outer membrane in the envelope of Gram-negative bacteria is composed of phospholipids and lipopolysaccharides, the predominant constituents of the hydrophobic diffusion barrier in the corynebacterial cell envelope are covalently linked mycolic acids, together with free corynomycolic acid esters (trehalose dicorynomycolate, trehalose monocorynomycolate) and phospholipids [99]. Mutants defective in protein components involved in extracytoplasmic lipid metabolism (mycoloyl transferases) display a decreased mycolate content and an altered cell wall permeability; this points to an important role of the mycolic acids for the outer membrane-like barrier [49,99]. Freeze-etch electron microscopy techniques indicate that this hydrophobic barrier represents a true lipid bilayer [100]. In accordance with an outer membrane bilayer, ion-permeable channels and pore-forming proteins, albeit different in monomer size and multimeric association from the trimeric porins of Gram-negative bacteria, have been found in corynebacteria and related bacteria [28,55,70,71,83,90,101]. The mycolate-less cells of C. amycolatum strains, on the other hand, apparently lack an outer membrane-like lipid bilayer [100]. The important role of the mycolic acids in the outer membrane-like structure of the cell envelopes of most species of Corynebacterium and related genera underscores the relevance of these lipids as chemotaxonomic markers for classification purposes. Extrapolating from the aforementioned results concerning molecular biological, biochemical, and ultrastructural studies on the cell walls of selected Corynebacterium and Mycobacterium strains, it seems clear that the presence of an outer membrane-like cell wall structure containing mycolic acid esters is an evolutionary conserved feature of the vast majority of genera, species, and strains grouped in the actinomycetes suborder Corynebacterineae.

Corynebacterium Taxonomy

13

2.2 TAXONOMY AND CHARACTERISTICS OF THE GENUS CORYNEBACTERIUM The genus Corynebacterium was originally defined in 1896 by Lehmann and Neumann [69] to accommodate nonmotile species that were pathogenic or at least parasitic to animals, in particular the diphtheroid bacilli. Thereafter, a number of morphologically similar plant pathogenic and soil-borne species were included into the genus (see [123]). For several decades after creation of Corynebacterium, the genus comprised an extremely diverse collection of microorganisms, accommodated together in one group mainly on the basis of their cell morphology, staining properties, and respiratory metabolism (see [72]). Mainly during the 1970s and 1980s, the use of chemotaxonomic markers (mainly the cell wall chemistry and lipid composition, as mentioned previously, and DNA base composition) helped to clarify the taxonomy and enabled researchers to redraw the borderline of the genus Corynebacterium. As a consequence, various Corynebacterium species were transferred to other genera, and other species previously placed elsewhere were included in the genus (e.g., [11–14,25], see [15,73]). During the last ten years, a large number of new species of Corynebacterium were isolated and classified. Now, phylogenetic approaches (mainly 16S rDNA sequence analysis) are used on a routine basis for classification, in addition to classical chemotaxonomic markers. In the course of this process of redefining the genus Corynebacterium, it turned out that the “plant pathogenic coryneform bacteria” that had been treated as members of the genus Corynebacterium for many years (e.g., see [29]), had to be removed from the genus. These bacteria are clearly not true Corynebacterium species sensu stricto (see [13]) and therefore were reclassified in other genera, mainly in Curtobacterium and Clavibacter [19,20,22,23,31,42,43]. In the new phylogenetic classification system of the class Actinobacteria [111], these genera are accommodated in a different suborder (Micrococcineae) of the order Actinomycetales than Corynebacterium (suborder Corynebacterineae). Although, as mentioned before, comparative 16S rDNA sequence analysis of the Actinomycetales taxa with cell wall chemotype IV and mycolic acids revealed that the species of the genus Corynebacterium form a monophyletic group [97,102], it is noteworthy that the genus exhibits considerable phylogenetic depth. This is also reflected by the observation that the genus Corynebacterium displays significant chemical heterogeneity in terms of lipid composition: in contrast to the other genera of the CMN group, Corynebacterium is not characterized by the presence of a unique major menaquinone and fatty acid type [15]. Interestingly, in phylogenetic analyses, the mycolic acid-less Corynebacterium species C. amycolatum [14], C. kroppenstedtii [16], and C. atypicum [47], and the single species of Turicella, T. otitidis [37], which also lacks mycolic acids and whose separate genus status may need to be reevaluated [102], apparently form relatively deeply branched distinct sublines in the phylogenetic tree of the genus Corynebacterium [16,47,102], but the deep branching position of T. otitidis has been questioned by Pascual et al. [97]. A maximum parsimony tree based on 16S rDNA sequence data demonstrating the radiation of the species of the genus Corynebacterium is shown in Figure 2.2.

14

Handbook of Corynebacterium glutamicum

Corynebacterium

Dietzia

Nocardia–Rhodococcus

Tsukamurella

Williamsia

Mycobacterium Gordonia 10%

(a)

FIGURE 2.2 (a) Maximum parsimony tree based on 16S rDNA sequence data showing the positions and phylogenetic depths of mycolic acid-containing Actinomycetales genera. The scale bar indicates 10% sequence divergence. (b) Maximum parsimony tree showing the positions of the industrially relevant species C. glutamicum, C. callunae, C. efficiens, and C. ammoniagenes within the radiation of species of the genus Corynebacterium. The scale bar indicates 5% sequence divergence. Both trees were kindly supplied by W. Ludwig (Freising, Germany).

As of July 2003, there were 67 validly published species of Corynebacterium, making this genus one of the top ten prokaryotic (Bacteria and Archaea) genera with regard to the number of known species. This observation could indicate that for unknown reasons this genus is evolutionarily differentiated into more species than many other genera, but alternatively the large number of known species may simply reflect an increased medical and industrial interest in corynebacteria. Many new species of Corynebacterium were described relatively recently, with an annual average of about four new species descriptions over the past eight years. A large number of corynebacterial species were described on the basis of strains isolated

Corynebacterium Taxonomy

15

C. cystitidis

C.

C. pilosum C. ammoniagenes C. amycolatum

es flc

ce

ns

um ur .d C le is na tid re asti C. . m C

C. xerosis C. glutamicum C. callunae C. efficiens C. auriscanis C. urealyticum C. jeikeium C. falsenii C. bovis C. variabillis C. terpenotabidum C. pseudotuberculosis C. diphtheriae C. vitaeruminis C. kutscheri

C. imitans ecis s e pend C. ap mycetoid m C. flavu m o il h op ucu C. lip C. gla C. thomssenii se C. sundsvallen ae C. coyle C. mucifaciens C. pseudodiphtheriticum C. confus C. propinquum um C. fastidiosum C. striatum C. simulans C. minutissimum C. nigricans 5%

C. glucuronolyticum (b)

FIGURE 2.2 (continued).

from human clinical samples or from animals, while others were isolated from various samples such as soil, feces, cheese smear, dairy products, vegetables, fruits, animal fodder, and other sources (see [73]). The following observations make it seem likely that the number of corynebacterial species will continue to expand rapidly. In an rRNA-based molecular phylogenetic approach aimed at the identification of bacteria from specimens from prostatitis patients, Tanner et al. [117] found a wide diversity of 16S rRNA sequences resembling Corynebacterium, a subset representing sequences from undescribed species on the basis of their positions in phylogenetic trees. Also, the increasing number of novel Corynebacterium species isolated from various animals (e.g., [18,35,45,96]) indicates that much new corynebacterial species diversity remains to be discovered not only from human, but also from animal sources. There also appears to be considerable unexplored Corynebacterium diversity in other fields than human- or animal-associated origin. It was reported that some coryneform bacteria isolated from the rind of different cheese varieties clustered close to Brevibacterium ammoniagenes (C. ammoniagenes) [105]. Recent studies on the microflora of smear-ripened cheeses showed that while in some cases the organisms isolated were found to be members of the already known species C. ammoniagenes and C. variabile [34,119], in other cases some new Corynebacterium species could be described [8]. Finally, the observation that significant numbers of bacteria with properties characteristic of the genus Corynebacterium have been isolated from marine samples [6] is noteworthy. However, these isolates were neither differentiated to the species level nor were they investigated with molecular systematics methods. In conclusion, even though only few data are available concerning the numbers of corynebacteria present in different habitats, it seems clear that nonmedical corynebacteria as well as medically relevant representatives of this

16

Handbook of Corynebacterium glutamicum

important group of bacteria are widely disseminated in nature, and that much species diversity of the genus Corynebacterium remains to be defined. The characteristic features of the genus Corynebacterium as described by Collins and Cummins [15] are the following: Gram-positive (sometimes unevenly stained); nonsporing; nonmotile; not acid-fast; straight or slightly curved rods, ovals, or clubs, often with metachromic granules; often exhibit typical V-shaped arrangement of cells (see Figure 2.3); facultatively anaerobic to aerobic; catalase-positive; chemoorganotrophic; peptidoglycan directly cross-linked of the type A1γ (cross-linkage of adjacent peptide chains via positions 3 and 4, peptide bridge absent, meso-A2pm at position 3 of tetrapeptide subunits [103]); predominant cell wall sugars are arabinose and galactose; mycolic acids (corynomycolic acids = short-chain α-substituted-βhydroxy acids with 22 to 36 carbon atoms) are present with rare exceptions (C. amycolatum, C. kroppenstedtii, and C. atypicum); straight-chain saturated or monounsaturated fatty acids are present; 10-methyl branched-chain acids may be present; and eight- and/or nine-isoprene-unit dihydrogenated menaquinones (MK8, MK9) are present. The DNA base composition of the genus Corynebacterium covers the wide range of approximately 46 to 71 mol % G+C, but most species have between 51 and 68 mol % G+C.

2.3 METHODS FOR THE ISOLATION, IDENTIFICATION, AND DIFFERENTIATION OF CORYNEBACTERIA A large number of different Corynebacterium species have been isolated from various human clinical and veterinary sources. In addition, nonmedical corynebacteria are found in a broad variety of different habitats such as soil, plants, animal fodder, and dairy products (see [73]). However, no selective media or enrichment procedures are known that are specifically suited for this group of organisms. During growth, bacteria obviously must be supplied with an energy source, a carbon source, a nitrogen source, and all other macronutrients and trace-elements (all in formulations suited for assimilation by the organisms) to meet the requirements defined by the cellular chemical composition (the elemental composition of C. glutamicum cells is shown in Table 2.1). In addition, variably, other organic growth supplements like amino acids, nucleotide bases, or vitamins are needed for growth. Most strains of nonmedical Corynebacterium species grow well at 30˚C in standard peptone-yeast extract media like the Corynebacterium medium (DSMZ, Deutsche Sammlung von Mikroorganismen und Zellkulturen GmbH, containing: 1% casein peptone, 0.5% yeast extract, 0.5% glucose, and 0.5% NaCl; pH 7.2–7.4), although growth on very rich media such as Brain Heart Infusion (Difco) is often faster and more abundant. For some industrially relevant Corynebacterium species like C. ammoniagenes and C. glutamicum, chemically defined media were described [75,85]. The addition of certain substances such as 0.1% citrate or low concentrations (10–5 M) of certain dihydroxyphenolic compounds (catechol, protocatechuate) greatly stimulates the growth of C. glutamicum in synthetic broth, presumably by assisting in the assimilation of iron by this organism [75,120]. In accordance, a cyclic catecholate named corynebactin was reported from C. glutamicum strain DSM

Corynebacterium Taxonomy

17

(a)

200 μm

(b)

FIGURE 2.3 (a) Phase-contrast micrograph of C. glutamicum cells grown on complex medium. Note frequent V-type arrangement of cell pairs, due to “snapping division.” (b) Same cells placed on a nucleopore membrane and viewed by scanning electron microscopy.

20411. Corynebactin is structurally similar to the enterobacterial siderophore enterobactin but displays opposite chirality [5,9]. Importantly, strains of glutamic acid producing corynebacteria (C. glutamicum and similar bacteria) are dependent upon the presence of biotin in the growth medium; some additionally require thiamine or p-amino benzoic acid [1]. A recipe for C. glutamicum synthetic broth BMCG is shown in Table 2.2. An alternative synthetic medium in use is CGXII as described

18

Handbook of Corynebacterium glutamicum

TABLE 2.1 Elemental Composition of C. glutamicum ATCC 13032 Cellsa Concentration (mg/g) Cells Grown in Element

BMCG

CGXII

CGIIIb

C O H N P S K Mg Ca Al Mn Fe Co Ni Cu Zn Mo

400 384 59.4 78.7 21.3 2.60 32.2 2.55 0.101 0.017 0.108 0.310 <0.00002 0.00046c 0.056 0.064 0.0009

413 373 61.4 84.5 14.5 4.00 23.6 1.59 0.035 0.007 0.201 0.510 <0.00002 0.00043 0.012 0.046 0.0010

411 368 60.9 97.4 17.3 3.10 28.2 2.26 0.028 <0.002 0.0039 0.040 0.00008 0.00009 0.003 0.075 0.0011

a

C. glutamicum strain ATCC 13032 was grown in BMCG synthetic broth or in CGIII complex medium containing 4% glucose. The cells were washed twice with 0.9% NaCl and dried before analysis with the following methods: IR spectroscopy (C, H, S after combustion with oxygen; O after combustion to CO2), heat conductance of N2 (N), inductively coupled argon plasma mass spectroscopy (ICP-MS) (other elements and trace elements). Standard errors of multiple measurements were generally less than ±15% of the values shown. b Interestingly, the cells grown in CGIII complex medium showed a significantly reduced content of some trace elements. c It is noteworthy that BMCG-grown and CGXII-grown C. glutamicum cells had a fourfold higher Ni content than E. coli cells grown in minimal medium (data not shown).

by Keilhauer et al. [58] but with 30 mg/l protocatechuic acid. For more details on growth conditions for C. glutamicum, also see the experimental section of this book. Several nongenotypic rapid identification and differentiation methods for corynebacteria are available, for example, some commercially available identification systems based on the analysis of physiological traits: the API CORYNE system (bioMerieux), the BIOLOG identification system (Biolog), and the RapID CB Plus system (Remel). A comparison of the performance of these systems was reported by Oberreuter et al.

Corynebacterium Taxonomy

19

TABLE 2.2 BMCG Synthetic Broth for C. glutamicum (NH4)2SO4 Distilled water 10× M9 solution Autoclave 20 min at 121˚C Then add aseptically: 200× salt solution Trace element solution 1 M CaCl2 Vitamin stock solution 20% glucose 10 mM catechol

7g 850 ml 100 ml

5 ml 2 ml 0.05 ml 1 ml 50 ml 1 ml

10× M9 stock = 60 g Na2HPO4, 30 g KH2PO4, 5 g NaCl, 10 g NH4Cl, 1000 ml H2O, pH 7.3; 200× salt solution = 80 g MgSO4•7H2O, 4 g FeSO4•7H2O, 0.4 g MnSO4•H2O, 5 g NaCl, and 1000 ml H2O; trace element solution = 88 mg Na2B4O7•10H2O, 40 mg (NH4)6Mo7O24•4H2O, 10 mg ZnSO4•7H2O, 270 mg CuSO4•5H2O, 7.2 mg MnCl2•4H2O, 870 mg FeCl3•6H2O, and 1000 ml H2O; vitamin stock solution = 1 mg biotin, 10 mg thiamine HCl per 1 ml H2O; 200× salt solution, trace element solution, and 1 M CaCl2 are autoclaved separately; vitamin and glucose solutions are filter sterilized; the catechol stock is adjusted to a neutral pH, sterilized by filtration, stored in aliquots at –20˚C and added aseptically to the medium just before inoculation.

[94]. However, it should be noted that some of the commercial systems are focused on the identification of clinical isolates and therefore do not cover the whole species diversity of the genus Corynebacterium. In addition to the commercial identification kits, Fourier-transform infrared (FT-IR) spectroscopy has recently been described as a useful method for the identification of actinomycete and corynebacterial species and strains [46,93,94]. Interestingly, Oberreuter et al. [93] have analyzed a number of amino acid-producing corynebacteria with FT-IR spectroscopy. In this study it was found that a number of suspected C. glutamicum strains with nonvalidated ‘Brevibacterium’ and ‘Micrococcus’ species names that had previously been included in ‘Corynebacterium’ or ‘Brevibacterium/Corynebacterium’ clusters in numerical studies by others [53,54] all revealed a high degree of spectral similarity. Comparative 16S rDNA sequence analysis confirmed the allocation of these strains to C. glutamicum [93]. For the in-depth characterization of new isolates as members of Corynebacterium and their description as species of this genus, normally a combination of physiological analysis (either classical methods or miniaturized kits for biochemical characterization), chemotaxonomic methods, and 16S rDNA sequence analysis is used. In

20

Handbook of Corynebacterium glutamicum

the following paragraphs, the most commonly used methods for the classification and differentiation of corynebacterial strains are listed.

2.3.1 MORPHOLOGY, MICROSCOPIC APPEARANCE, AND STAINING PROPERTIES For classification of bacteria by morphology, the cell morphology should be inspected carefully and repeatedly after obtaining a pure culture, as usual in good microbiological practice. Under the light microscope, at about 400- to 1,000-fold magnification, bacteria from species of the genus Corynebacterium appear as rodshaped cells with a typical, somewhat irregular (“coryneform”) morphology. Cells are often arranged in V-formations, due to their “snapping” mode of cell division (Figure 2.3). Sometimes, packages of several cells in parallel arrangement (as “palisades”) are also found. Although some observed morphological differences can depend on the media used and the culture age (see [1,59,72,122]; author’s unpublished observations), the typical morphological features of Corynebacterium are normally recognized without difficulty when C. glutamicum is grown in various media or when inspected after different incubation periods (see [72]). Therefore, the use of special growth media for this purpose, such as EYGA medium as recommended by Cure and Keddie [30], is normally not necessary. Corynebacterium cells are not motile and do not sporulate. The cells are Gram-positive and non-acid-fast.

2.3.2 CELL WALL One important facet of the classification process is the analysis of bacterial cell walls and in particular the determination of the cross-linkage type [103]. After mechanical disruption of the bacterial cells and isolation of murein [103], whole-cell sugars can be determined by the method of Lechevalier [66]. The isomeric form of diaminopimelic acid can be identified by paper chromatography of cell wall hydrolysates [4,103].

2.3.3 MYCOLIC ACIDS

AND

OTHER LIPIDS

Due to the fact that mycolic acids are unique lipophilic components of the cell envelopes of the genera of the suborder Corynebacterineae (see earlier discussion), they represent diagnostic chemotaxonomic markers for Corynebacterium and related taxa [15,17,23,24,56,80]. The most convenient method for the determination of mycolic acids (see [57] for an evaluation of methods) is the acid methanolysis of the cells and subsequent thin layer chromatographic analysis of the resulting mycolic acid methyl esters [79–81]. An alkali procedure for the preparation and two-dimensional TLC separation of mycolic acid methyl esters was described by Minnikin [78]. The presence of mycolic acids can also be investigated by GLC analysis of trimethylsilylated mycolic acid methyl ester derivatives [64]. Nonhydroxylated fatty acids are usually identified as the corresponding methyl esters using gas chromatographic analysis [53]. Methods for the purification and analysis of menaquinones were described by Kroppenstedt [65] and Collins et al. [24].

Corynebacterium Taxonomy

2.3.4 DNA

AND

21

16S RRNA GENE ANALYSIS

The DNA base composition of total DNA can be determined by thermal denaturation [40,48] or via HPLC analysis [77,115]. DNA-DNA hybridization [32,48] is an extremely important method for the delineation of species [109] and to study intraand inter-species relationships. For 16S rDNA sequence analysis and phylogenetic studies, a large fragment of the 16S rRNA gene of the bacterium under investigation is sequenced after amplification via PCR, using universal primers that bind near the 5 and 3 ends of the rRNA gene. Alignment of the resulting sequence with known 16S rDNA sequences, and phylogenetic analyses are conveniently carried out using the ARB software package [76].

2.3.5 PHYSIOLOGICAL PROPERTIES Corynebacteria are catalase-positive, aerobic, or facultatively anaerobic bacteria. A number of biochemical characteristics that may be useful for differentiating some saprophytic Corynebacterium species were compiled by Liebl [72,73]. For various tests such as acid formation from carbohydrates and certain enzyme tests, commercially available, miniaturized systems may be very useful, e.g., API CORYNE (bioMerieux), BIOLOG identification system (Biolog), RapID CB Plus (Remel), MINITEK (Becton Dickinson), API ZYM (bioMerieux), etc. The optimum growth temperature of the amino-acid-producing corynebacteria, the main focus of this monograph, is between 25 and 37˚C [1]. It was recently reported that representative wild-type strains of C. glutamicum also grew rapidly at 40˚C, but apparently this trait of thermotolerance is easily lost during the breeding of lysine production strains, presumably due to secondary mutations during repeated rounds of random mutagenesis and selection [95].

2.4 ISOLATION, CLASSIFICATION, AND TAXONOMY OF INDUSTRIALLY RELEVANT CORYNEBACTERIA Certain corynebacteria have a multidecade tradition as industrial microorganisms in biotechnological production processes, in particular for the production of the amino acids L-glutamic acid and L-lysine, which are mainly used as a flavor enhancer and a feed additive, respectively. Saprophytic corynebacteria are also used in the fermentative production of nucleotides, which are of interest primarily as flavor-enhancing additives in foods. The following industrially relevant Corynebacterium species will be discussed from a taxonomic point of view: C. glutamicum, C. callunae, C. efficiens, ‘C. thermoaminogenes,’ and C. ammoniagenes. Abbreviations in superscript are used to indicate the source of the recognition of the name: AL indicates that the name is included in the “Approved Lists of Bacterial Names” [107], and VP indicates that the name has been validly published in the International Journal of Systematic Bacteriology (now the International Journal of Systematic and Evolutionary Microbiology) or published elsewhere and quoted in the validation lists of that journal. All genera and species not validly published are written in single quotation marks.

22

2.4.1 CORYNEBACTERIUM

Handbook of Corynebacterium glutamicum GLUTAMICUMAL

First reported by Kinoshita et al. in 1958 [61], C. glutamicum is included in Abe et al. [1] and in the “Approved Lists of Bacterial Names” [107]. Two of the most important amino acids in terms of quantity, i.e., l-glutamic acid and l-lysine, are produced fermentatively on a large scale by strains of C. glutamicum. In addition, mutant strains of C. glutamicum have been selected which excrete the purine ribonucleoside 5-monophosphates 5-inosinic acid (IMP), 5-xanthylic acid (XMP), and 5-guanylic acid (GMP) (see [101]). The history of the discovery of C. glutamicum (synonym Micrococcus glutamicus) as a potent amino acid producer and the beginning of industrial amino acid production by microbial fermentation can be redrawn as follows (based on the following sources: a recapitulation of events given in a lecture by S. Kinoshita at the Congress of the International Union of Microbiological Societies at Osaka, Japan, September 16, 1990 [60,118]). Near the beginning of the twentieth century, monosodium glutamate (MSG) was discovered in Japan as a new taste (“umami”) and used as a flavoring compound for food. In search of a microorganism that could produce and excrete amino acids, a group at the Japanese company Kyowa Hakko Kogyo in Tokyo initiated a screening program headed by S. Kinoshita and S. Udaka in 1955. The screening method [118], which in essence represented a bioassay for glutamic acid, was carried out by replica plating bacterial isolates of various origins onto a series of plates containing various defined test media. After growth, the cells on the test plates were inactivated by a strong dose of UV radiation, and the plates were overlayed with basal agar containing a glutamate-auxotrophic bacterium, Leuconostoc mesenteroides strain P-60. After incubation at 37˚C, the test plates were scored for the appearance of a halo of growth of the assay organism around some of the UV-killed colonies. The glutamic acid-producers were then recovered from a non–UV-irradiated nutrient medium master plate. Among about 500 isolates tested on a simple synthetic medium containing relatively high concentrations of sugar and nitrogen source (50 g glucose, 0.5 g K2PO4, 0.1 g MgSO4·7H2O, 8 g urea, and 40 mg FeCl3·6H2O, per liter H2O, pH 7.2), researchers at Kyowa Hakko found that several percent of the tested strains gave rise to halo formation in the plate bioassay. Most of them produced between 0.1 and 0.5 mg/ml glutamic acid in the culture filtrates of aerobically grown liquid cultures, but one strain (isolate No. 534) accumulated about 10 mg/ml after two days of incubation. This strain, which was found in early 1956, and further strains isolated thereafter, were named ‘Micrococcus glutamicus’ by Kinoshita et al. [61–63]. The strains were found to be biotin auxotrophs and only produced glutamic acid within a relatively narrow concentration range of biotin in the medium. Subsequently, a large number of glutamic acid-producers were isolated and classified in different genera (mainly Brevibacterium, Corynebacterium, Microbacterium, Micrococcus, and Arthrobacter), but most of them carried invalid names. One of the largest numerical taxonomic studies on glutamic acid–producing bacteria was performed by Abe et al. [1], who investigated a total of 208 glutamic acidproducing strains designated as ‘Micrococcus glutamicus’ (= Corynebacterium

Corynebacterium Taxonomy

23

glutamicum), ‘Brevibacterium aminogenes’, ‘Brevibacterium divaricatum,’ Brevibacterium ammoniagenes, ‘Brevibacterium flavum,’ ‘Brevibacterium lactofermentum,’ ‘Brevibacterium roseum,’ ‘Brevibacterium saccharolyticum,’ ‘Brevibacterium immariophilum,’ ‘Corynebacterium acetoacidophilum,’ Corynebacterium lilium, Corynebacterium callunae, and ‘Corynebacterium herculis.’ All were Gram-positive, nonsporulating, nonmotile, ellipsoidal spheres to short rods that displayed biotin requirement and were able to produce >30 g/l of l-glutamic acid from carbohydrates and ammonia in their culture broth under aerobic conditions. Numerous data exist ([1,80,114]; Liebl et al. unpublished data) indicating their close relatedness or identity with C. glutamicum. Numerous other strains originally assorted to the genera Brevibacterium, Corynebacterium, Microbacterium, Micrococcus, or Arthrobacter, e.g., ‘Brevibacterium chang-fua’, Brevibacterium divaricatum, ‘Brevibacterium flavum’, ‘Brevibacterium glutamigenes’, ‘Brevibacterium lactofermentum’, ‘Brevibacterium roseum‘, ‘Brevibacterium seonmiso, Brevibacterium sp., ‘Brevibacterium taipei, ‘Brevibacterium thiogenitalis’, Corynebacterium lilium, ‘Corynebacterium herculis’, ‘Microbacterium ammoniaphilum’, Microbacterium sp., ‘Micrococcus maripuniceus’, Arthrobacter sp. which have been included in numerical taxonomic studies were found to group into C. glutamicum clusters [53,54,114], indicating that they were misnamed. Also, some glutamic acid-producing strains designated as Brevibacterium ammoniagenes were shown to actually belong to C. glutamicum [1,114]. Importantly, none of the amino acid-producing, nomenclatural Brevibacterium species turned out to represent a true member of the genus Brevibacterium. Consequently, numerous strains have been reclassified as C. glutamicum, e.g., strains misclassified as Brevibacterium species (e.g., ‘B. chang-fua’ ATCC 14017, B. divaricatum DSM 20297T, ‘B. flavum’ DSM 40411, ‘B. glutamigenes’ ATCC 13747, ‘B. lactofermentum’ DSM 20412 and DSM 1412, ‘B. roseum’, ‘B. saccharolyticum’ ATCC 14066, ‘B. seonmiso’ ATCC 14915, Brevibacterium sp. ATCC 19165, ‘B. taipei’ ATCC 13744, ‘B. thiogenitalis’ ATCC 19240), Corynebacterium species (e.g., C. lilium DSM 20137T), Microbacterium species (e.g., Microbacterium sp. ATCC 15283), Micrococcus species (e.g., ‘Mc. maripuniceus’ ATCC 14399), Arthrobacter species (e.g., Arthobacter sp. NCIB 9666) [74,93]. Among the strains reclassified as C. glutamicum are also the type strains of two species, i.e., of Brevibacterium divaricatum and Corynebacterium lilium [74]. Values reported in the literature for the genomic G+C content of C. glutamicum strains are in the range of about 53 to 58 mol %. The precise G+C content of the genome sequence available from Kyowa Hakko for C. glutamicum ATCC 13032 is 53.8 mol %. Recently, the intraspecific diversity of C. glutamicum was studied with 16S rDNA sequence analysis and Fourier-transform infrared (FT-IR) spectroscopy [93]. The values for pairwise 16S rDNA sequence identity between the 27 strains of C. glutamicum investigated ranged from 95.7 to 100%. No significant correlation between the FT-IR spectral similarity and the 16S rDNA sequence similarity between the strains of this species could be detected, which indicates that FT-IR spectral comparison, although useful for strain identification at the species and genus level, is not suited to assess evolutionary relationships [93].

24

Handbook of Corynebacterium glutamicum

The natural habitats of C. glutamicum strains reported so far are soil, soils contaminated with bird feces, sewage and manure, vegetables, and fruits [72,73,93,106,121]. One strain appears to be from a marine source (ATCC 14399, ‘Micrococcus maripuniceus’; see [93]). The type strain (DSM 20300T = ATCC 13032T) was originally isolated from sewage. Most strains of C. glutamicum form pale yellow or yellow colonies, some are cream-white, but spontaneous color variants are easily found [1].

2.4.2 CORYNEBACTERIUM

CALLUNAEAL

C. callunae was described as an amino acid-producing bacterium in a U.S. patent by Lee and Good [68] and included in Yamada and Komagata [122]. In the numerical study of Abe et al. [1] which included the type strain of C. callunae (NRRL B-224T = DSM 20147T = ATCC 15991T), this organism was grouped separately from most typical glutamate-producing corynebacteria (Micrococcus glutamicus = C. glutamicum and other misnamed strains) because it was urease-negative (however, this trait was not consistently reported to be negative in the literature; i.e., Yamada and Komagata [122] reported C. callunae to be ureasepositive) and did not produce nitrite from nitrate. Nonetheless it was suggested that the organism could be named C. glutamicum [1]. However, more recent studies do not question the separate species status of C. callunae. DNA-DNA hybridization studies revealed a genomic DNA relatedness of about 37% between C. callunae and C. glutamicum [74]. As to be expected from this relatively high DNA-DNA similarity value, phylogenetic studies based on 16S rDNA sequence comparison including a large number of Corynebacterium species (e.g., [8,97,102]) confirmed that the closest relative of C. callunae is C. glutamicum, revealing a 97 to 98% 16S rDNA sequence identity between these two species [97,102]. Phenotypically, C. callunae can be distinguished from C. glutamicum by its inability to produce nitrite from nitrate, and its ability to produce acid from salicin [36], although also contradictory data are reported for acid production from salicin by these two species [36,51,122]. Also, the G+C content of C. callunae (about 51 mol %) is lower than reported for C. glutamicum strains (about 53 to 58 mol %) (see [73]). The habitat for the only strain of C. callunae investigated so far is heather [51].

2.4.3 CORYNEBACTERIUM

EFFICIENSVP

Fudou et al. reported this species in the International Journal of Systematic and Evolutionary Microbiology in 2002 [36]. Strains of this species were first described and tentatively named ‘Corynebacterium thermoaminogenes’ in the 1980s in a Japanese patent application by Yamada and Seto [124] after having been isolated from onion bulbs and soil samples collected at Kanagawa, Japan [36]. The strains were isolated on complex media incubated aerobically at 45˚C in search for bacteria which are more heat-tolerant than C. glutamicum and could be used in amino acidproduction at elevated temperature, with the potential advantage of reducing the costs needed for cooling during the fermentation process. The strains fulfilled all the chemotaxonomic prerequisites to be assigned to the genus Corynebacterium

Corynebacterium Taxonomy

25

sensu stricto as defined by Collins and Cummins [15]. Also, the three isolates studied and proposed as members of the novel species Corynebacterium efficiens by Fudou et al. [36] revealed >80% DNA similarity among each other by DNA-DNA hybridization, whereas the DNA-DNA relatedness with the other amino acid-producing Corynebacterium species, C. glutamicum and C. callunae, was less than 20%, thus clearly indicating separate species status. Comparative 16S rDNA sequence analysis revealed 95.3% identity of C. efficiens with its closest relative, C. glutamicum, which is lower than the 97% value proposed by Stackebrandt and Goebel [110] to define a species [36]. The genome sequences of C. glutamicum and C. efficiens confirm their close relatedness. Physiologically, the new species C. efficiens differs from the other amino acid-producing species, C. glutamicum and C. callunae, by its ability to produce acid from dextrin, its inability to assimilate D-lactate and succinate, its inability to grow at pH 6 or with 30% glucose, and its ability to grow at 45˚C. Finally, the DNA G+C content of C. efficiens as determined by whole-genome analysis [92] is 63.8 mol % and thus is considerably higher than the G+C contents of C. glutamicum (53.8 mol %) and C. callunae (about 51 mol %) [36]. The natural habitats of C. efficiens strains reported so far are soil and vegetables (onion bulbs; see [36]).

2.4.4 ‘CORYNEBACTERIUM

THERMOAMINOGENES’

Strains with this nonvalidated species designation have been reported in patent applications dealing with amino acid production employing thermophilic corynebacterial isolates, by Yamada and Seto [124] and Murakami, Miwa, and Nakamori [84]. Three strains that had tentatively been named ‘C. thermoaminogenes’ were recently classified as the new species C. efficiens [36].

2.4.5 CORYNEBACTERIUM

AMMONIAGENESVP

First reported by Cooke and Keith in 1927 [27], C. ammoniagenes was transferred from its basonym ‘Brevibacterium ammoniagenes’ to the genus Corynebacterium in 1987 [12]. Strains of this species were first isolated, described, and designated as ‘Bacterium ammoniagenes’ in the 1920s. This organism was found as a ureasplitting bacterium during the investigation of ammonia dermatitis of the gluteal region of infants and was isolated from the stools of infants and children [27]. The species was assigned to the genus Brevibacterium by Breed [7]. However, after redefinition of the genus Brevibacterium by Collins et al. [21], B. ammoniagenes was transferred to the genus Corynebacterium as C. ammoniagenes [12]. The main features which led to the reclassification were the presence of arabinogalactan in the cell wall, the presence of mycolic acids (chain length 32 to 36 carbon atoms), the polar lipid composition, and the G+C content of about 54 to 56 mol %, all features incompatible with the classification concept of true brevibacteria. In the 1960s it was found that adenine-requiring mutants of B. ammoniagenes (C. ammoniagenes) accumulated the purine ribonucleoside 5-monophosphate 5-inosinic acid (IMP) under certain fermentation conditions [86]. Nucleotide production took place at certain concentrations of manganese ions (Mn2+) [39,88] or,

26

Handbook of Corynebacterium glutamicum

under Mn2+-excess conditions, upon addition of certain antibiotics or surfactants or with the use of mutants in which manganese did not inhibit production [38,89]. Remarkably, intact wild-type cells of C. ammoniagenes are able to use “salvage synthesis” reactions to convert the bases hypoxanthine, guanine, and adenine to the respective nucleoside monophosphate IMP (for hypoxanthine) or the mono-, di-, and triphosphates GMP, GDP, and GTP (for guanine) and AMP, ADP, and ATP (for adenine), respectively [86,87,116]. These observations stood at the beginning of the development of nucleotide fermentation processes based on C. ammoniagenes. Various alternative processes for the production of IMP have been described. In addition, C. ammoniagenes has been used for the production of inosine, sugar nucleotides, GMP, ATP, NAD+, and, recently, riboflavin (vitamin B2) (see [73]). Strains classified as ‘Brevibacterium ammoniagenes’ (C. ammoniagenes) and C. glutamicum with numerical taxonomy methods were isolated from piggery wastes by Seiler and Hennlich [106]. In a numerical taxonomic study of cheese-smear coryneform bacteria isolated from the rind of different cheese varieties, Seiler [105] found that about 20% of the white and yellow colored coryneform isolates clustered in a group closely resembling the “Brevibacterium ammoniagenes group” of Seiler [104]. A number of cheese-smear isolates were classified as C. ammoniagenes and C. variabile by Eliskases-Lechner and Ginzinger [34] and Valdes-Stauber et al. [119]. The natural habitats of C. ammoniagenes strains reported so far are feces from humans and animals, and smear-ripened cheeses [12,34,105,106,119].

2.5 CONCLUSION Modern biochemical and molecular methods of strain characterization and the meaningful taxonomic concepts available now aid in the rapid and reliable identification and differentiation of corynebacteria and related organisms. It is of great value that the classification concept of the genus Corynebacterium and related actinomycete bacteria has been significantly improved over the past decades. Especially the now widely recognized picture of the phylogenetic relationship between Corynebacterium and related genera is of importance for future studies, because close phylogenetic relatedness also often means close genetic, biochemical, and physiological similarity, and results obtained from studies in one organism can often be transferred to another related organism. Therefore, it is likely that our knowledge about the biology of medical and nonmedical corynebacteria will rapidly expand in the near future. For this, the nonpathogenic corynebacteria may play a particularly important role due to their ease of handling and due to the availability of sophisticated tools for their genetic manipulation. As is evident from other chapters of this book, the characterization of corynebacterial species to the molecular level is already rapidly advancing.

ACKNOWLEDGMENTS The author gratefully acknowledges the calculation of phylogenetic trees by W. Ludwig (Freising, Germany) and thanks M. Hoppert (Göttingen, Germany) for his aid in electron microscopy.

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86. Nara T, Misawa M, and Kinoshita S. (1967) Production of 5-inosinic acid by an adenine auxotroph of Brevibacterium ammoniagenes. Agr. Biol. Chem. 31:1351–1356. 87. Nara T, Misawa M, and Kinoshita S. (1968) Fermentative production of 5-purine ribonucleotides by Brevibacterium ammoniagenes. Agr. Biol. Chem. 32:561–567. 88. Nara T, Misawa M, and Kinoshita S. (1968) Pantothenate, thiamine and manganese in 5-purine ribonucleotide production by Brevibacterium ammoniagenes. Agr. Biol. Chem. 32:1153–1161. 89. Nara T, Misawa M, Komuro T, and Kinoshita S. (1969) Effect of antibiotics and surface-active agents on 5-purinenucleotide production by Brevibacterium ammoniagenes. Agr. Biol. Chem. 33:1198–1204. 90. Niederweis M, Maier E, Lichtinger T, Benz R, and Krämer R. (1995) Identification of channel-forming activity in the cell wall of Corynebacterium glutamicum. J. Bacteriol. 177:5716–5718. 91. Nikaido H, Kim SH, and Rosenberg EY. (1993) Physical organization of lipids in the cell wall of Mycobacterium chelonae. Mol. Microbiol. 8:1025–1030. 92. Nishio Y, Nakamura Y, Kawabarayasi Y, Usuda Y, Kimura E, Sugimoto S, Matsui K, Yamagishi A, Kikuchi H, Ikeo K, and Gojobori T. (2003) Comparative complete genome sequence analysis of the amino acid replacements responsible for the thermostability of Corynebacterium efficiens. Genome Res. 13:1572–1579. 93. Oberreuter H, Charzinski J, and Scherer S. (2002) Intraspecific diversity of Brevibacterium linens, Corynebacterium glutamicum and Rhodococcus erythropolis based on partial 16S rDNA sequence analysis and Fourier-transform infrared (FT-IR) spectroscopy. Microbiology 148:1523–1532. 94. Oberreuter H, Seiler H, and Scherer S. (2002) Identification of coryneform bacteria and related taxa by Fourier-transform infrared (FT-IR) spectroscopy. Int. J. Syst. Evol. Microbiol. 52:91–100. 95. Ohnishi J, Hayashi M, Mitsuhashi S, and Ikeda M. (2003) Efficient 40˚C fermentation of L-lysine by a new Corynebacterium glutamicum mutant developed by genome breeding. Appl. Microbiol. Biotechnol. 62:69–75. 96. Pascual C, Foster G, Alvarez N, and Collins MD. (1998) Corynebacterium phocae sp. nov., isolated from the common seal (Phoca vitulina). Int. J. Syst. Bacteriol. 48:601–604. 97. Pascual C, Lawson PA, Farrow JAE, Gimenez MN, and Collins MD. (1995) Phylogenetic analysis of the genus Corynebacterium based on 16S rRNA gene sequences. Int. J. Syst. Bacteriol. 45:724–728. 98. Peyret J-L, Bayan N, Joliff G, Gulik-Krzywicki T, Mathieu L, Schechter E, and Leblon G. (1993) Characterization of the cspB gene encoding PS2, an ordered surfacelayer protein in Corynebacterium glutamicum. Mol. Microbiol. 9:97–109. 99. Puech V, Bayan N, Salim K, Leblon G, and Daffe M. (2000) Characterization of the in vivo acceptors of the mycoloyl residues transferred by the corynebacterial PS1 and the related mycobacterial antigens 85. Mol. Microbiol. 35:1026–1041. 100. Puech V, Chami M, Lemassu A, Laneelle M-A, Schiffler B, Gounon P, Bayan N, Benz R, and Daffe M. (2001) Structure of the cell envelope of corynebacteria: importance of the non-covalently bound lipids in the formation of the cell wall permeability barrier and fracture plane. Microbiology 147:1365–1382. 101. Riess FG, Lichtinger T, Cseh R, Yassin AF, Schaal KP, and Benz R. (1998) The cell wall porin of Nocardia farcinica: biochemical identification of the channel-forming protein and biophysical characterization of the channel properties. Mol. Microbiol. 29:139–150.

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102. Ruimy R, Riegel P, Boiron P, Boivin V, Monteil H, and Christen R. (1995) A phylogeny of the genus Corynebacterium deduced from analysis of small-subunit ribosomal DNA sequences. Int. J. Syst. Bacteriol. 45:740–746. 103. Schleifer KH and Kandler O. (1972) Peptidoglycan types of bacterial cell walls and their taxonomic implications. Bacteriol. Rev. 35:407–477. 104. Seiler H. (1983) Identification key for coryneform bacteria derived by numerical taxonomic studies. J. Gen. Microbiol. 129:1433–1471. 105. Seiler H. (1986) Identification of cheese-smear coryneform bacteria. J. Dairy Res. 53:439–449. 106. Seiler H and Hennlich W. (1983) Characterization of coryneform bacteria in piggery wastes. System. Appl. Microbiol. 4:132–140. 107. Skerman VBD, McGowan V, and Sneath PHA. (1980) Approved lists of bacterial names. Int. J. Syst. Bacteriol. 30:225–420. 108. Soual-Hoebeke E, de Sousa-D’Auria C, Chami M, Baucher M-F, Guyonvarch A, Bayan N, Salim K, and Leblon G. (1999) S-layer protein production by Corynebacterium strains is dependent on the carbon source. Microbiology 145:3399–3408. 109. Stackebrandt E, Frederiksen W, Garrity GM, Grimont PAD, Kämpfer P, Maiden MCJ, Nesme X, Rosselo-Mora R, Swings J, Trüper HG, Vauterin L, Ward AC, and Whitman WB. (2002) Report of the ad hoc committee for the re-evaluation of the species definition in bacteriology. Int. J. Syst. Evol. Microbiol. 52:1043–1047. 110. Stackebrandt E and Goebel BM. (1994) Taxonomic note: a place for DNA-DNA reassociation and 16S rRNA sequence analysis in the present species definition in bacteriology. Int. J. Syst. Bacteriol. 44:846–849. 111. Stackebrandt E, Rainey FA, and Ward-Rainey NL. (1997) Proposal for a new hierarchal classification system, Actinobacteria classis nov. Int. J. Syst. Bacteriol. 47:479–491. 112. Stackebrandt E and Woese CR. (1981) The evolution of prokaryotes. In Carlile MJ, Collins JF, and Moseley BEB (Eds.), Molecular and Cellular Aspects of Microbial Evolution. Cambridge University Press, Cambridge, pp. 1–31. 113. Sutcliffe IC. (1995) Identification of a lipoarabinomannan-like lipoglycan in Corynebacterium matruchotii. Arch. Oral Biol. 40:1119–1124. 114. Suzuki K, Kaneko T, and Komagata K. (1981) Deoxyribonucleic acid homologies among coryneform bacteria. Int. J. Syst. Bacteriol. 31:131–138. 115. Tamaoka J. (1994) Determination of DNA base composition. In Goodfellow M and O’Donnell AG (Eds.), Chemical Methods on Prokaryotic Systematics. Wiley, Chichester, pp. 463–470. 116. Tanaka H, Sato Z, Nakayama K, and Kinoshita S. (1968) Formation of ATP, GTP, and their related substances by Brevibacterium ammoniagenes. Agr. Biol. Chem. 32:721–726. 117. Tanner MA, Shoskes D, Shahed A, and Pace NR. (1999) Prevalence of Corynebacterial 16S rRNA Sequences in Patients with Bacterial and “Nonbacterial” Prostatitis. J. Clin. Microbiol. 37:1863–1870. 118. Udaka S. (1960) Screening method for microorganisms accumulating metabolites and its use in the isolation of Micrococcus glutamicus. J. Bacteriol. 79:754–755. 119. Valdes-Stauber N, Scherer S, and Seiler H. (1997) Identification of yeast and coryneform bacteria from the surface microflora of brick cheese. Int. J. Food. Microbiol. 34:115–129. 120. Von der Osten CH, Gioannetti C, and Sinskey AJ. (1989) Design of a defined medium for growth of Corynebacterium glutamicum in which citrate facilitates iron uptake. Biotechnol. Lett. 11:11–16.

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121. Woodruff HB. (1981) A soil microbiologist’s odyssey. Annu. Rev. Microbiol. 35:1–28. 122. Yamada K and Komagata K. (1972) Taxonomic studies on coryneform bacteria. IV. Morphological, cultural, biochemical, and physiological characteristics. J. Gen. Appl. Microbiol. 18:399–416. 123. Yamada K and Komagata K (1972) Taxonomic studies on coryneform bacteria. V. Classification of coryneform bacteria. J. Gen. Appl. Microbiol. 18:417–431. 124. Yamada and Seto, Japanese Patent JP 63-240779, October 6, 1987.

Part III Genome, Plasmids, and Gene Expression

3

The Genomes of Amino Acid–Producing Corynebacteria J. Kalinowski

CONTENTS 3.1 Introduction ....................................................................................................37 3.2 Mapping and Sequencing of the C. glutamicum Genome ............................38 3.3 Sequencing Other Corynebacterial Genomes................................................39 3.4 Annotation of the C. glutamicum Genome ...................................................39 3.5 The Overall Structure of the C. glutamicum Genome ..................................41 3.6 Prophages in the C. glutamicum Genome.....................................................44 3.7 The Gene Inventory of C. glutamicum..........................................................46 3.8 Comparative Corynebacterium Genome Analysis ........................................47 3.9 Conclusions ....................................................................................................51 Acknowledgments....................................................................................................52 References................................................................................................................53

3.1 INTRODUCTION In the mid-1950s, Kinoshita and co-workers in Japan isolated a bacterium that was shown to excrete large quantities of L-glutamic acid into the culture medium [23]. This bacterium, now named Corynebacterium glutamicum, was described as a short, aerobic, Gram-positive rod capable of growing on a variety of sugars or organic acids. Following the first description as ‘Micrococcus glutamicus,’ a number of different names were assigned to various isolates of C. glutamicum, e.g., ‘Brevibacterium lactofermentum,’ ’Brevibacterium flavum,’ ’Brevibacterium divaricatum,’ and ’Corynebacterium lilium.’ This naming confusion was first clarified by the publication of Liebl et al. [26], who showed by modern taxonomic classification methods that the above-mentioned species designations were invalid and that all strains belong to the species Corynebacterium glutamicum. The type strain of this species is C. glutamicum ATCC 13032 (alternative designations: DSM 20300, IMET 10482, and NCIB 10025). DNA sequencing of individual C. glutamicum genes started in the mid-1980s, when several genes from amino acid–biosynthetic pathways in C. glutamicum were 37

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cloned and analyzed. These genes were mainly identified by heterologous complementation of Escherichia coli mutants and, occasionally, in the homologous system, e.g., by isolating genes of producer strains conferring resistance of a C. glutamicum strain against an amino acid analog [44] or by complementation of an export deficiency [46]. These studies already led to a general understanding of metabolic pathways, but a complete picture of the complex interactions could not be achieved owing to the lack of comprehensive genetic information. The sequencing of the complete corynebacterial genome turned out to represent the ideal method to obtain the missing genetic information and thus the basis for an efficient rational development of industrial producer strains. This chapter is devoted to the description of the current knowledge on corynebacterial genomes. The genome sequences presented have been determined recently, and first analyses using bioinformatics have already provided interesting and valuable information for basic science as well as for industrial application. Because new genome information as well as novel bioinformatics tools are generated at high speed, a large number of fascinating discoveries on corynebacteria can be expected in the near future.

3.2 MAPPING AND SEQUENCING OF THE C. GLUTAMICUM GENOME The first steps toward a complete genome sequence of C. glutamicum were made by determining the genome size and establishing a linked physical and genetic map [3]. The genome size was determined by pulsed-field gel electrophoresis of large DNA fragments generated by digestion with rare-cutting restriction enzymes. By this, the genome size was estimated to be 3.1 megabase pairs (Mbp). The large DNA fragments were arranged by Southern hybridizations using an established cosmid library of C. glutamicum. These studies revealed that the C. glutamicum ATCC 13032 genome consists of one circular chromosome. Later, the cosmid library was used to generate an ordered minimal set of clones for the sequencing of the C. glutamicum genome by the same research team. In the study, 95 cosmid clones were arranged in a minimal set, but unfortunately the genome sequence coverage by cosmids was only 87% [42]. Therefore, an additional library in bacterial artificial chromosomes (BACs) was generated, and 21 of these BACs helped to cover the whole chromosome. With the higher resolution obtained in this mapping project, the estimated genome size was corrected to 3.28 Mbp. In a collaborative research project led by the Degussa Company (Düsseldorf, Germany) and the Department of Genetics of Bielefeld University (Bielefeld, Germany), the 116 overlapping BAC and cosmid clones were sequenced individually by the shotgun method up to coverages of five- to ninefold and assembled by bioinformatics software. After finishing the cosmid and BAC sequences by PCR and primer-walking sequencing methods, contiguous high-quality sequences were available early in the project. The nucleotide sequences of the individual cosmid and BAC clones were finally assembled to a whole-genome sequence of 3,282,708 base pairs, harboring 3,002 potential genes [21].

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39

The complete genome sequence of C. glutamicum ATCC13032 was determined in independent approaches by at least two other research teams. A Japanese team consisting of a collaboration between the Kyowa Hakko Company (Machida, Japan) and Kitasato University (Sagamihara, Japan) applied the whole-genome shotgun method based on plasmid and cosmid libraries [20]. These two libraries were sequenced to a five- to sixfold redundancy, and the sequences were assembled by bioinformatics software. The cosmid library was later used to close the sequence gaps resulting in a contiguous sequence of 3,309,401 base pairs and the identification of 3,099 genes. This genome sequence was the first that appeared in public databases. A third project was carried out by the BASF company (Ludwigshafen, Germany) together with Integrated Genomics Inc. (Chicago, IL, U.S.A.). Information on how this project was carried out is scarce, but 2,900 genes were reported, of which 1,400 found entry into several patent applications. Because at the time of this writing only the first two complete genome sequences have found entry into public databases, the BASF/Integrated Genomics Project will not be discussed in this text.

3.3 SEQUENCING OTHER CORYNEBACTERIAL GENOMES In parallel with the tight race between the different consortia that were sequencing C. glutamicum ATCC 13032 at almost the same time, the Ajinomoto Co. (Kawasaki, Japan) isolated a closely related species that was also sequenced. A special feature of this strain is its adaptation to higher temperatures around 40°C, a fact that led to its initial naming as ‘Corynebacterium thermoaminogenes.’ A deeper taxonomic analysis of this soil-inhabiting and glutamate-producing Corynebacterium strain led to the proposal of a new species clearly distinct from C. glutamicum, named C. efficiens [13] (see Chapter 2). The complete genome sequence of C. efficiens [33] was determined by the whole-genome shotgun method using two plasmid clone libraries with different insert size classes (0.8 to 1.2 kbp, 2.0 to 2.5 kbp). Sequences were assembled with standard bioinformatics software. The sequence was deposited in public databases and displays 3,147,090 bp for the main chromosome. In addition to the chromosome, the sequenced strain inherits two plasmids (Table 3.1). The GC content of the C. efficiens chromosome is unexpectedly high (63.4%), and 2,950 genes were predicted [33]. The other corynebacterial genome sequenced because of its great medical interest was that of Corynebacterium diphtheriae. This species is the causative agent of diphtheria, and the NCTC 13129 strain is a recent U.K. clinical isolate representative of an epidemic clone currently circulating in Eastern Europe. The genome is 2,488,635 bp long, with a GC content of 53.5% (Table 3.1), and was assembled from 66,099 sequencing reads [8].

3.4 ANNOTATION OF THE C. GLUTAMICUM GENOME The initial step in genome annotation is gene finding, which can be carried out using a variety of bioinformatics tools that work reasonably well but at a certain level fail

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TABLE 3.1 Features of Corynebacterium Genome Sequences Strain Accession number Data produced by Size (bp)

Average G+C content (%)

Number of ORFs

Number of rRNA operons Number of tRNAs Coding regions (%) Mean ORF length (bp) Start codon usage (%) AUG GUG UUG Other (pseudogenes)

C. glutamicum ATCC13032a BA000036/NC003450b

C. glutamicum ATCC13032a BX927147

C. efficiens YS-314 BA000035

C. diphtheriae NCTC13129 BX248353

Kyowa Hakko and Kitasato University 3,309,401a

Degussa AG and Bielefeld University 3,282,708a

Ajinomoto

Sanger Institute 2,488,635

53.8

53.8

3,147,090 23,743 (pCE1) 48,672 (pCE2) 63.4

53.5

3,002

54.4 (pCE1) 56.4 (pCE2) 2,950

2,320

6

6

15 (pCE1) 41 (pCE2 5

5

60

60

56

54

86.8

88.3

90.4

89.6

933

916

979

964

62.5 24.3 13.2 —

66.5 23.1 10.3 0.1

53.8 32.5 13.7 —

68.0 23.7 7.5 0.8

3,099/2,993b

a

The C. glutamicum strains are variants differing with respect to point mutations, insertion elements and a putative prophage (see text). b The submitted genome sequence (GenBank Acc. No. BA000036) was reannotated by the National Center for Biotechnology Information and deposited under Acc. No. NC003450.

to predict some genes or falsely predict nonexistent genes. The numbers of falsepositives and false-negatives depend on the software tool used, its parameter settings, or training sets as well as on the composition of the genomic DNA to be analyzed. Therefore, it is preferable to use different tools for gene prediction in conjunction with visual inspection of each anticipated gene. This holds true also for the predicted gene starts, which in some instances have to be modified.

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41

The C. glutamicum genome sequence established by the Degussa–Bielefeld University consortium was consistently annotated in the software GenDB [27]. Gene finding was performed by combining two bioinformatics tools: CRITICA [2] was used to define a gene set, which was subsequently used by GLIMMER [11] to construct a training model and to perform the final gene finding. This combination makes effective use of the selectivity of CRITICA (very few false-positives) and the sensitivity of GLIMMER (very few true-negatives). Genes were validated and coding sequence starts were checked by visual inspection after TBLASTN comparisons of the protein sequences deduced from all C. glutamicum ORFs against four other genome sequences from the Actinomycetales phylogenetic lineage, comprising C. diphtheriae, C. efficiens, Mycobacterium tuberculosis, and Streptomyces coelicolor. The C. glutamicum genome sequence established by the Kyowa Hakko–Kitasato University consortium was first analyzed by automatic gene prediction using GLIMMER, and the results were intensively controlled and modified by manual intervention [20]. These researchers identified 3,099 putative protein-coding genes. The National Center for Biotechnology Information has independently searched the deposited sequence with another gene-finding tool, GeneMarkS [4], and has predicted 2,993 genes. It must be kept in mind that automatic prediction is by no means perfect and gene numbers in C. glutamicum may vary in the future between the reported extreme values of 2,900 and 3,100. Variations are also expected for other values deduced from the first annotation of a genome sequence, e.g., the genomic coverage by open reading frames, the mean ORF length, as well as the distribution of start codons depend heavily on the gene-finding strategies employed (Table 3.1). For annotation, additional databases were used in each of the genome projects but only reported in detail for the Degussa–Bielefeld University project [21]. These authors used additional databases for gene function analysis, the nonredundant protein sequence database (nr), SWISSPROT, and INTERPRO, including several protein pattern databases. Additionally, SignalP [32] and TMHMM [24] were used to identify proteins that are potentially secreted or located in the cytoplasmic membrane, respectively.

3.5 THE OVERALL STRUCTURE OF THE C. GLUTAMICUM GENOME As mentioned, two complete genomic sequences for the strain C. glutamicum ATCC 13032 are available to date. However, these genomes are not identical, a fact that is first reflected by the differing genome sizes, 3,282,708 bp and 3,309,401 bp. The difference of roughly 27 kbp in size is mainly due to additional copies of insertion elements and an additional putative prophage inserted in the larger genome (or deleted from the smaller one). It must be concluded that these mobile elements are capable of changing the C. glutamicum genome sequence by insertion or recombination in relatively short time periods, leading to two different genome sequences of the same ATCC 13032 strain. The general features of the C. glutamicum genome sequences are shown in Table 3.1 and Figure 3.1. The C. glutamicum genome is represented by a circular

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Handbook of Corynebacterium glutamicum

0 kb

3200 kb

200 kb

3000 kb 400 kb

HGC1 2800 kb LG C1

600 kb

2600 kb Corynebacterium glutamicum ATCC 13032 3.3 Mb

800 kb

2400 kb 1000 kb

2200 kb

CG

P4

P1

CGP

CG

3

1200 kb

CGP2 2000 kb 1400 kb 1800 kb

1600 kb

FIGURE 3.1 (Color insert follows page 208.) Circular representation of the C. glutamicum ATCC 13032 genome (GenBank Acc. No. NC003450). The plot was generated by GenDB version 2.0 [27]. Circles denote (outward to inward) the following: coding regions transcribed in clockwise and counterclockwise direction, respectively; GC content and GC skew. Bars pointing outward indicate values positively deviating from the median, and bars pointing inward indicate values negatively deviating from the median. The locations of low-GC and high-GC genomic regions as well as the prophages described in the text are represented by red, black, and green bars, respectively.

chromosome of 3.3 Mbp in size, which is a little larger than that of its close relative, C. efficiens (3.1 Mbp), and significantly larger than that of C. diphtheriae (2.5 Mbp). The G+C content of the genome is 53.8%, which is close to that of E. coli and at the lower boundary for the taxonomic class of the Actinobacteria, referred to as high-G+C Gram-positive bacteria. However, C. diphtheriae also has a similar G+C content of 53.5%, whereas the genome of C. efficiens has one of 63.4%. In contrast to the other corynebacterial genomes, C. efficiens inherits two plasmids, pCE1 and pCE2, which display G+C contents of 54.4 and 56.4%, respectively. These numbers are clearly different from that of the chromosome and might indicate a recent acquisition of these plasmids by C. efficiens. The GC skew analysis [15], which is generally applicable to identify the leading and the lagging strand in DNA replication, indicated a bidirectional replication that

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TABLE 3.2 Genomic Regions Differing in G+C Content and Prophages in C. glutamicum Name

Locationa

Size (kbp)

Genesb

Description

NCgl0335-0357 Cgl0342-0368

Low-G+C region containing genes involved in cell wall formation and lipopoly– saccharide synthesis Putative incomplete prophage inserted at a tRNA-Leu carrying an integrase gene Putative prophage remnant carrying integrase and lysin genes

LGC1

NB 360.111-390.733

26.9

CGP1

CG 363.825-390.734 NB1.400.182-1.413.641

13.5

cg0414-0443 NCgl1281-1298 Cgl1336-1352

CGP2

CG 1.401.510–1.415.069 NB 1.637.081-1.641.004

3.9

cg1507-1524 NCgl1490-1492 Cgl1548-1553

CGP3

CG 1.638.548–1.642.471 NB 1.776.613-1.995.294c

(219.7)c

cg1746-1752 NCgl1611-1816 Cgl1675-1891

CG 1.778.085–1.965.342

187.3

cg1890-2071

CGP4c

NB 1.963.136-1.986.590

23.5

NCgl1773-1806 Cgl1847-1881

HGC1

NB 3.156.304-3.176.905

20.6

NCgl2851-2871 Cgl2952-2974

CG 3.129.610-3.150.211

cg3267-3295

Putative prophage inserted at a tRNA-Val gene carrying integrase, primase, restriction/modification, and lysin genes Putative prophage inserted into CGP3 and carrying integrase, nuclease, singlestrand–binding protein, and lysin genes. Terminal duplication of ca. 4.5 kb High-G+C region with a 7-kbp region highly conserved in C. diphtheriae encoding a putative copper transport and chaperone system

a

NB refers to GenBank Acc. Nos. NC003450 and BA000036, CG refers to BX927147 NCgl refers to NC003450, Cgl refers to BA000036, and cg refers to BX927147. c The CGP4 prophage is inserted into CGP3 and is only present in the NC003450/BA000036 genome sequence. b

starts at the proposed oriC sequence near the dnaA gene and ends near the calculated replication terminus at around 1.6 Mbp (Figure 3.1). Note that several regions of the C. glutamicum genome deviate significantly in G+C content from the median (Figure 3.1, Table 3.2). There is one region of 20 kbp in size located at around 3,150 kbp, which deviates significantly to a high-G+C content and was named HGC1. The genes of this region have G+C contents of up to 66% and are flanked by defective insertion sequences. The leftward 7 kbp of this region are more than 95% identical at the nucleotide level

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to a segment from the C. diphtheriae genome and contain a putative copper transport system and a two-component sensor–regulator system. The extraordinarily high level of sequence conservation points to a recent horizontal gene transfer. This interpretation is supported by the fact that C. efficiens, which has a similar high mean G+C content, contains orthologs to the genes of this region that are conserved only on the protein sequence level. The puzzling question concerns the source organism in which the genes of the high-G+C region have evolved. Although C. diphtheriae contains a nearly identical gene region, the high-G+C content of this region is also exceptional for this organism. Therefore, parallel gene transfer events into both organisms, or two events with one of both being the source for the second gene transfer, must have occurred. This horizontal gene transfer might have been initiated by flanking insertion sequences and a possibly involved plasmid. Plasmids replicating in different Corynebacterium species have been reported [43] (see Chapter 4). In contrast to the HGC1 region’s being exceptional in having a higher G+C content, a number of genomic regions were identified as exceptionally deficient in GC (Figure 3.1, Table 3.2). One of these regions is located at around 380 kbp, has a size of approximately 27 kbp, and covers around 20 coding regions with G+C contents of 41 to 49% (LGC1). The genes that are located in this region are involved in some aspects of murein formation, e.g., murA and murB, and lipopolysaccharide synthesis. It is interesting to note that two genes encoding the enzymes for the initial steps in murein formation are duplicated in C. glutamicum. Whereas murA and murB located in this region have a low-G+C content of 44%, the second copy of these genes (murA2, murB2) exhibit a G+C content typical for C. glutamicum ORFs. Also in other organisms, exceptionally low-G+C regions carrying genes involved in cell wall and cellular surface formation have been found (e.g., Bacillus subtilis), indicating a preferred horizontal gene transfer or special selective benefit from receiving such functions by horizontal gene transfer. This is also the case in the so-called pathogenicity islands, which often confer functions involved in bacterial surface modifications [25]. Horizontal gene transfer is generally mediated by transposable elements, plasmids, and bacteriophages. In fact, at least 24 insertion elements are present in the C. glutamicum genome [21]. These elements are frequently found at the borders of regions of unusual G+C content, e.g., the HGC1 region (Table 3.2).

3.6 PROPHAGES IN THE C. GLUTAMICUM GENOME Ubiquitous to bacterial genomes are bacteriophages, and their genomically integrated forms are referred to as prophages [6]. Integration of prophages into a bacterial genome is generally recognizable by a discontinuity in the DNA composition (mean G+C, GC skew). Prophages are diverse in size and are found in bacterial genomes in various stages of degeneration. In the search for prophages in the C. glutamicum genome, the criteria for their identification were the above-mentioned irregularities in G+C content or GC skew, the presence of genes with bacteriophage homologs of known function (e.g., integrases), or the presence of gene regions lacking homologs, especially in the closely related strains (C. efficiens, C. diphtheriae).

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Integrase genes (int) encoding the vital enzyme responsible for integration of the bacteriophage genome into the genome of its host are particularly easy to detect by similarity searches. Altogether, five hypothetical phage integrase genes (′int, int1, int2′, ′int2, int3) were found in the Degussa–Bielefeld University sequence, with ‘int being partially deleted and int2 appearing to be disrupted by a frame-shift mutation. A closer inspection revealed a large low-G+C region, including the int2 gene fragments as the putative C. glutamicum prophage CGP3, whereas the int1 gene is part of the CGP1 putative prophage and the ‘int gene might be part of another defective prophage named CGP2 (Table 3.2). The CGP3 element present in the larger genome sequence is extended by approximately 20 kbp and displays a 5-kbp duplicated gene region containing a second ‘int2 gene at its end. The putative integrase gene int3 is not flanked by genes fulfilling the above-mentioned criteria for a prophage origin. The largest prophage region CGP3 spans more than 180 kbp. It covers approximately 200 coding regions, most of which lack any significant similarities to known bacterial genes. However, there are a few exceptions, including the three genes of the already known restriction-modification system (cglIM, cglIR, cglIIR) [39], genes encoding transposases, putative recombination enzymes, and a number of homologs to known bacteriophage proteins, especially a phage primase and the phage-type integrase ‘int2. It is interesting to note that the left border of this region is formed by a cluster of tRNA genes, whereas the phage-type integrase gene is located near the right border of the insertion. These observations might be explained by the integration of one or more prophage-like elements at a specific tRNA locus, a mechanism that is common for phages and integrative plasmids [47]. The putative prophage region CGP3 represents the major difference in the two genome sequences. In the left flank of this region, the Kyowa Hakko–Kitasato University sequence has a large insertion of another prophage (CGP4) and a duplicated gene region containing the committed phage-type integrase gene. The assumption that this C. glutamicum ATCC 13032 genome inherits an additional prophage is supported by the fact that the integrated region carries another serine protease as a putative cellulytic enzyme. An alternative explanation is that the unique fragment of the original prophage has been deleted in the Degussa–Bielefeld University sequence. The potential prophages found in C. glutamicum are diverse in size. Whereas the CGP3 element is larger than most known prophages, the smaller putative prophages CGP1 and CGP2 are presumably highly degenerated remnants of former functional bacteriophages. In the case of CGP1 and CGP3, the place of insertion is a tRNA gene, and the insertion site sequence is detectable as a direct repeat flanking the element. A flanking tRNA gene and direct repeats are missing in case of the apparently highly degenerated CGP2 element. The origin of these prophages is not known. Isolations of bacteriophages infecting different C. glutamicum strains have been described several times [29,35,40,45], but no report on the successful induction of a prophage from strain ATCC 13032 is available. Also, nucleotide sequences of genes from these corynephages were obtained only in a few cases, thus leaving the relationship between the presumed prophages and known infectious corynebacterial phages unclear.

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3.7 THE GENE INVENTORY OF C. GLUTAMICUM From a soil bacterium, it can be expected that its genome has to encode all necessary functions for primary metabolism, for catabolism of a wide variety of nutrients, and for optimal adaptation to changes in the environment. Corynebacterium glutamicum was the first completely sequenced Gram-positive soil bacterium from the Corynebacterianeae. The other members of this group whose genomes are known are C. efficiens, C. diphtheriae, Mycobacterium tuberculosis, M. leprae, M. bovis, and M. marinum — most of them important pathogens. Since nonpathogenic model systems are necessary, C. glutamicum may serve as an ideal system for studying the cell wall structure and synthesis and, especially, mycolic acid synthesis. Corynebacterium glutamicum is capable of growing in a simple mineral salts medium; i.e., it is able to synthesize from simple precursors all cell constituents, including metabolites, cofactors, and vitamins, except for D-biotin. This defect is most probably due to the fact that the gene bioF, encoding the biotin biosynthetic enzyme 7-keto-8-aminopelargonic acid synthetase, is missing in C. glutamicum [17,18]. All of the genes already described for various C. glutamicum strains and represented as nucleotide sequences in public databases were found also in the genome sequences, with one important exception. The gene for the para-crystalline surfacelayer protein cspB [36] from C. glutamicum ATCC 17965, which is synthesized in extremely large amounts and has a possible function in protecting the bacterium in soil against rough conditions, is missing in both C. glutamicum ATCC 13032 sequences. It is not clear why this gene is absent, but it can be speculated that bacterial strains in laboratories adapt to the specific growth conditions by losing functions that provide a heavy metabolic load and do not confer any advantage under optimal growth conditions [14]. Because also the immediate cspB flanking regions are not represented in the C. glutamicum ATCC 13032 sequence, it is impossible to determine its original place in the genome. The conjunction of automated and manual annotation of coding regions by similarities to known genes in public databases helped to annotate 82% of the C. glutamicum ORFs. However, annotation sometimes consisted only of a global functional characterization based on minor similarities with other ill-characterized genes or proteins. By accepting the huge bandwidth in functional assignments of C. glutamicum genes, only a small fraction of genes remained hypothetical (9%), meaning not similar to a database entry, or conserved hypothetical (9%), meaning similar to a hypothetical protein from another organism. In the case of a conserved hypothetical protein, the assumption is that it represents a real gene, whereas the hypothetical genes must be confirmed by further studies, such as by proteome analysis. Annotated genes can be assigned to functional classes with the widely used COG (cluster of orthologous groups) system [41]. Figure 3.2 shows that about 500 genes of C. glutamicum fall into the COG categories S (function unknown) and R (general function prediction only). About 900 genes cannot be classified by the COG system. However, the numbers of the members within the other classes are more or

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FIGURE 3.2 Functional classification of proteins from C. glutamicum, C. efficiens, and C. diphtheriae. The assignments have been made by using the COG functional classification scheme. The bars represent the numbers of genes of a functional class in a genome.

less the same in all three corynebacterial genomes, differing only in the three categories mentioned. The ill-characterized genes, which cannot be classified, might be interpreted as additional ones that carry probably nonessential genetic information whereas a common core of around 1,600 genes is found in all three genomes. Expert-manual annotation already provided a deeper understanding of gene function and helped to reconstruct most parts of the central metabolism, starting from sugar consumption and ending with produced amino acids [21]. Other functional complexes have also been analyzed and reconstructed using the information provided by the genome projects, e.g., in the excellent review on the respiratory chain by Bott and Niebisch [5]. Further reconstructions of other parts of metabolism can be expected in the near future.

3.8 COMPARATIVE CORYNEBACTERIUM GENOME ANALYSIS Up to now, four genome sequences from corynebacteria are available. These sequences represent the three species C. glutamicum, C. efficiens, and C. diphtheriae, of which C. glutamicum and C. efficiens represent both natural producers of

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L-glutamic acid. Since C. diphtheriae is a human pathogen and more distantly related to the biotechnologically relevant corynebacteria, it is especially interesting to compare the C. glutamicum and C. efficiens genomes. This comparison is of even higher relevance because both species differ by approximately 10% in their overall G+C content. Irrespective of this, the amino acid sequences of the proteins are fairly well conserved, facilitating the discrimination between protein-coding and noncoding DNA. Nishio and co-workers [33] have studied the connections among overall G+C content, codon preference, and phenotype in C. efficiens in comparison to C. glutamicum. These authors discovered that the differences in G+C content between both genomes were accompanied by a significant bias in amino acid substitutions. Three major substitutions were identified in C. efficiens, from lysine to arginine, serine to alanine, and serine to threonine. These substitutions are suggested to be the cause of the higher thermostability of C. efficiens proteins and the organism’s attribute for growing at temperatures above 40°C. Although some of the numbers deduced from the genome sequences are dependent on the gene-finding strategies, a rough comparison can be made. Corynebacterium glutamicum and C. efficiens have comparable genome sizes and gene numbers (Table 3.1). The numbers of six (C. glutamicum) and five rRNA operons (C. efficiens) of the order 16S-23S-5S and of 60 and 56 tRNA genes, respectively, are typical for fast-growing environmental bacteria. However, C. diptheriae as a human pathogen displays similar figures. Other genome features, such as the fractional genomic coverage by ORFs and the mean ORF length, are also rather similar and close to the values determined for many other bacterial genomes. In contrast to this, the usage of start codons is affected by the G+C content. Therefore, C. efficiens has a slightly higher fraction of ORFs starting with GUG in comparison to the two other corynebacterial species. An evaluation of gene-order conservation revealed an astonishing degree of synteny between all three Corynebacterium species (Figure 3.3). A possible reason for this unusual genome stability is given by Nakamura and co-workers [30]. These researchers found out that corynebacteria did not contain recBCD genes, encoding the recombinational repair system. It was suggested that the absence of this system prevented gene shuffling and retained an ancestral gene order in corynebacteria. The gene order analysis performed on the three genomes also clearly showed that the putative bacteriophage insertion regions are of alien origin. Additionally, there are several smaller regions carrying genes with no counterpart in one of the other species. A closer inspection of these regions might reveal genes that are also either horizontally transferred or are necessary only in a certain ecological niche. An example for this is the LGC1 region, which carries genes possibly involved in lipopolysaccharide synthesis, having their closest homologs in Gram-negative pathogens such as Neisseria and Campylobacter. In addition to putative prophages and the large number of different insertion elements that are present in both genomes, further differences in gene content between the two soil-inhabiting corynebacteria were detected. Only slight differences are detectable between the C. glutamicum and C. efficiens genomes if COG classes are compared (Figure 3.3). The differences are found mainly in the unclassified

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ORF # C. diphtheriae

2000 1500 1500 1000 1000 500

ORF # C. efficiens

2500

2000

500

0

0 0

500

1000

1500

2000

2500

3000

ORF # C. glutamicum

FIGURE 3.3 (Color insert follows page 208.) Gene order comparison for three corynebacterial genomes. The dots give the results of a bidirectional best hit generated by comparing the amino acid sequences with BLASTP [1]. The gene numbering refers to the BX927147 sequence.

genes, confirming the notion that they might carry nonessential information. However, it must be noted that C. glutamicum contains a larger number of transcriptional regulator genes (class K), whereas the C. efficiens genome is richer in transposases (class L). A more detailed analysis revealed a number of genes from C. glutamicum ATCC 13032 where the functions are known to be due to detailed characterization of mutant strains and functional analyses of the encoded proteins and that are absent from the C. efficiens YS-314 genome (Table 3.3). Significant is the missing of the brnEF

TABLE 3.3 C. glutamicum Genes Encoding Known Functions without Homologues in C. efficiens Gene

Function of Gene Product

Reference

brnE brnF lrp cma prpD2 prpB2 prpC2 porA

Component of a branched-chain amino acid exporter Component of a branched-chain amino acid exporter transcriptional Regulator, Lrp family Cyclopropane-fatty-acyl-phospholipid synthase Component of the methylcitrate cycle for propionate degradation Component of the methylcitrate cycle for propionate degradation Component of the methylcitrate cycle for propionate degradation Porin

[22]

[31] [7]

[9]

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Handbook of Corynebacterium glutamicum

genes encoding the components of a branched-chain amino acid exporter [22]. This transporter was shown to export the amino acids L-isoleucine, L-leucine, and L-valine. In addition, the adjacent lrp gene encoding a transcriptional regulator, is missing also in C. efficiens. Whether this means that C. efficiens is not capable of excreting branched-chain amino acids should be clarified by experimental studies. A second gene cluster missing in C. efficiens is the prpDBC2 cluster, whereas the prpDBC1 cluster is present. The function of the latter cluster is unknown, whereas the prpDBC2 gene cluster has been shown to encode the enzymes of the methylcitrate cycle essential for propionate degradation in C. glutamicum [7]. In agreement with the absence of prpDBC2 in C. efficiens is the characterization of this species by Fudou and co-workers [13] showing that C. efficiens is unable to degrade propionate as carbon source. The cma gene possibly encoding a mycolic acid cyclopropane synthase [31] is also missing in C. efficiens. Cyclopropanation of mycolic acids is a general feature of mycobacteria. At present, it is not known whether C. glutamicum possesses cyclopropanated mycolic acids, but the difference with respect to the cma gene might result in a difference in mycolic acid structure between the two species. Another structural difference with respect to the outer cell layers is also implicated by the missing of a clear porA gene homologue in C. efficiens. The porA gene encodes the major porin of C. glutamicum [9], which forms a channel necessary to allow the transport of hydrophilic substrates through the highly hydrophobic outer layer of C. glutamicum. Because it can be expected that the C. efficiens surface is hydrophobic enough to require the presence of a porin, an unrelated porin gene might be present in C. efficiens. Such a gene is probably not detected by automated gene-finding methods owing to the small size of the protein (PorA: 45 amino acids). In comparison to C. glutamicum, much less is known on the biology and the biochemistry of C. efficiens. Therefore, the C. efficiens genome is annotated in a rather conservative way. However, the comparison of both genomes clearly identified a number of C. efficiens genes without homologs in C. glutamicum ATCC 13032. Among these are the examples displayed in Table 3.4. It is striking that C. efficiens carries gene equipment for the degradation of a number of aromatic compounds from biological sources. One example is the gene cluster for the CoA-ligase and the monooxygenase enzymes degrading phenylacetic acid (CE0663-CE0672). In addition, the enzymes for the release of inorganic sulfate from aromatic sulfate esters (arylsulfatase, CE2198) as well as for the degradation of phenols (tyrosinase, CE1756) and amides (formamidase, CE2198) seem to be present. However, tests for tyrosinase enzyme activity proved negative for C. efficiens [13], indicating that all hypotheses deduced via bioinformatics have to be verified by additional experimental approaches. However, disagreements between predicted and observed phenotypes might result from incorrect annotations based on sequence similarity but also from possible difficulties in finding the right parameters for the expression of a regulated gene. Beside degradative functions, interesting biosynthetic functions are predicted to exist in C. efficiens. By judging from sequence similarities, the genes CE1202 and CE1203 might encode the subunits of a cellulose synthase. Up to now, such enzymes

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TABLE 3.4 Examples of Annotated C. efficiens Genes without Homologs in C. glutamicum Gene Number

Annotated Function

Comment

CE0663-CE0672

Phenylacetic acid degradation gene cluster

CE1202, CE1203 CE1568

Subunits of cellulose synthase Arylsulfatase

CE1756 CE2198 CE2362, CE2363

Tyrosinase Formamidase Ribonucleotide reductase type III

CE2454-CE2458 CE2737-CE2741

Fimbrial proteins and export system

Usage of phenylacetate as carbon source Cellulose synthesis Usage of aromatic sulfate esters as sulfur source Degradation of monophenols Usage of amides as nitrogen source Enzyme only active under anaerobic conditions Important for attachment to eukaryotic cells and biofilms

have been found only in some Gram-negative bacteria [37], and it will be fascinating to find out whether C. efficiens is able to produce cellulose. Additional genes might give predictions on the habitat and lifestyle of C. efficiens. Examples of such genes are CE2454–CE2458 and CE2737–2741 predicted to encode fimbriae proteins as well as their specific transport mechanisms. Fimbriae are often found in bacteria of medical relevance and are important for attachment to eukaryotic cells. In addition, they might be needed for the buildup of bacterial biofilms that also occur in the environment [10]. Another example is the possession of a type III ribonucleotide reductase encoded by CE2362 and CE2363. Ribonucleotide reductases of type III are oxygen sensitive and found exclusively in anaerobic or facultatively anaerobic microorganisms [12]. Corynebacterium efficiens possesses an additional oxygen-insensitive ribonucleotide reductase of type 1b (encoded by nrdF and nrdE), very similar to that of C. glutamicum. This is in agreement with the strain description for C. efficiens as being capable of life under aerobic and facultatively anaerobic conditions [13]. The genes mentioned in the examples given above as well as other genes encoding interesting functions in C. efficiens now await experimental proof. Therefore, genetic engineering techniques for C. efficiens should be developed or C. glutamicum could be used as a model system in heterologous expression experiments. However, owing to their much higher GC-content and differences in codon usage [33], expression of C. efficiens genes in C. glutamicum is not guaranteed.

3.9 CONCLUSIONS The establishment of complete annotated genome sequences of Corynebacterium strains is a big leap forward in the understanding of these organisms and will boost genetic analyses as well as metabolic engineering to overproduce compounds of commercial relevance.

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The C. glutamicum genome sequence has already helped to directly identify missing genes along close biosynthetic pathways of interest or to provide a limited number of candidate genes for testing. An example of this is the work of Hartmann and co-workers [16], in which the genes dapF and dapC completing the lysine biosynthetic pathway have been successfully located. Another example is provided by Rückert and co-workers [38], who studied the biosynthesis of methionine. In this study, the C. glutamicum genome was first scanned for candidate genes involved in methionine biosynthesis. Then, by generating deletion mutant strains for all candidate genes and auxanography, a complete branched pathway for L-methionine synthesis was reconstructed. Another issue for which a genome sequence is essential is the discovery of novel or unexpected functions. An example is the discovery of two genes encoding carbonic anhydrases in this organism [28] or the discovery of a second glyceraldehyde-3dehydrogenase gene, which is expressed exclusively under gluconeogenetic conditions [19]. As mentioned before, the C. efficiens genome awaits the development of genetic engineering techniques to prove some fascinating hypotheses deduced via bioinformatics. The information generated by the joint application of high-quality bioinformatics and broad biochemical and biotechnological knowledge — as well as genetic engineering will allow the creation of highly efficient production strains for well-established and for novel products. Of particular importance with respect to production strain development is the genome analysis of the highly efficient production strains developed traditionally. Through sequence comparison of alleles relevant for the production process with their wild-type counterparts and subsequent introduction of the identified mutations into a wild-type strain, a minimally mutated production strain with improved properties can be derived. This technique of “genome breeding” was introduced by Ohnishi et al. [34] with the development of an efficient L-lysine– producing strain of C. glutamicum by the introduction of only three different mutations. The engineered strain reached a high production level but showed far better growth than the traditionally developed production strain, which allows fermentation times to be shortened by about 50%. The complete genome sequence also forms the basis for most methods of global expression analyses, e.g., proteome and transcriptome studies. Such analyses will lead to a comprehensive systemic understanding of gene expression and regulatory networks in corynebacteria in the future.

ACKNOWLEDGMENTS The help of Christian Rückert, Alexander Goesmann, Burkhard Linke, Oliver Rupp, Alice McHardy, and Daniela Bartels during various steps of bioinformatics analyses and generation of figures is thankfully acknowledged.

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31. Nampoothiri KM, Hoischen C, Bathe B, Möckel B, Pfefferle W, Krumbach K, Sahm H, and Eggeling L. (2002) Expression of genes of lipid synthesis and altered lipid composition modulates L-glutamate efflux of Corynebacterium glutamicum. Appl. Microbiol. Biotechnol. 58:89–96. 32. Nielsen H, Engelbrecht J, Brunak S, and von Heijne G. (1999) A neural network method for identification of prokaryotic and eukaryotic signal peptides and prediction of their cleavage sites. Int. J. Neural. Sys. 8:581–599. 33. Nishio Y, Nakamura Y, Kawarabayasi Y, Usuda Y, Kimura E, Sugimoto S, Matsui K, Yamagishi A, Kikuchi H, Ikeo K, and Gojobori T. (2003) Comparative complete genome sequence analysis of the amino acid replacements responsible for the thermostability of Corynebacterium efficiens. Genome Res. 13:1572–1579. 34. Ohnishi J, Mitsuhashi S, Hayashi M, Ando S, Yokoi H, Ochiai K, and Ikeda M. (2002) A novel methodology employing Corynebacterium glutamicum genome information to generate a new L-lysine-producing mutant. Appl. Microbiol. Biotechnol. 58:217–223. 35. Patek M, Ludvik J, Benada O, Hochmannova J, Nesvera J, Krumphanzl V, and Bucko M. (1985) New bacteriophage-like particles in Corynebacterium glutamicum. Virology 140:360–363. 36. Peyret JL, Bayan N, Joliff G, Gulik-Krzywicki T, Mathieu L, Shechter E, and Leblon G. (1993). Characterization of the cspB gene encoding PS2, an ordered surface-layer protein in Corynebacterium glutamicum. Mol. Microbiol. 9:97–109. 37. Ross P, Mayer R, and Benziman M. (1991) Cellulose biosynthesis and function in bacteria. Microbiol Rev. 55:35–58. 38. Rückert C, Pühler A, and Kalinowski J. (2003) Genome-wide analysis of the L-methionine biosynthetic pathway in Corynebacterium glutamicum by targeted gene deletion and homologous complementation. J. Biotechnol. 104:213–228. 39. Schäfer A, Tauch A, Droste N, Pühler A, and Kalinowski J. (1997) The Corynebacterium glutamicum cglIM gene encoding a 5-cytosine methyltransferase enzyme confers a specific DNA methylation pattern in an McrBC-deficient Escherichia coli strain. Gene 203:95–101. 40. Sonnen H, Schneider J, and Kutzner HJ. (1990) Corynephage Cog, a virulent bacteriophage of Corynebacterium glutamicum, and its relationship to phi GA1, an inducible phage particle from Brevibacterium flavum. J. Gen. Virol. 71:1629–1633. 41. Tatusov RL, Natale DA, Garkavtsev IV, Tatusova TA, Shankavaram UT, Rao BS, Kiryutin B, Galperin MY, Fedorova ND, and Koonin EV. (2001) The COG database: new developments in phylogenetic classification of proteins from complete genomes. Nucleic Acids Res. 29:22–28. 42. Tauch A, Homann I, Mormann S, Rüberg S, Billault A, Bathe B, Brand S, BrockmannGretza O, Rückert C, Schischka N, Wrenger C, Hoheisel J, Möckel B, Huthmacher K, Pfefferle W, Pühler A, and Kalinowski J. (2002a) Strategy to sequence the genome of Corynebacterium glutamicum ATCC 13032: use of a cosmid and a bacterial artificial chromosome library. J. Biotechnol. 95:25–38. 43. Tauch A, Kirchner O, Löffler B, Götker S, Pühler A, and Kalinowski J. (2002b) Efficient electrotransformation of Corynebacterium diphtheriae with a mini-replicon derived from the Corynebacterium glutamicum plasmid pGA1. Curr. Microbiol. 45:362–367. 44. Thierbach G, Kalinowski J, Bachmann B, and Pühler A. (1990) Cloning of a DNA fragment from Corynebacterium glutamicum conferring aminoethyl cysteine resistance and feedback resistance to aspartokinase. Appl. Microbiol. Biotechnol. 32:443–448.

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Handbook of Corynebacterium glutamicum 45. Trautwetter A, Blanco C, and Sicard AM. (1987) Structural characteristics of the Corynebacterium lilium bacteriophage CL31. J. Virol. 61:1540–1545. 46. Vrljic M, Sahm H, and Eggeling L. (1996) A new type of transporter with a new type of cellular function: L-lysine export from Corynebacterium glutamicum. Mol. Microbiol. 22:815–826. 47. Williams KP. (2002) Integration sites for genetic elements in prokaryotic tRNA and TmRNA genes: sublocation preference of integrase subfamilies. Nucleic Acids Res. 30:866–875.

4

Native Plasmids of Amino Acid–Producing Corynebacteria A. Tauch

CONTENTS 4.1 4.2 4.3

Introduction ....................................................................................................57 Isolation of Plasmids from Amino Acid–Producing Corynebacteria............61 Structural Organization of the pBL1 Family of Corynebacterial Plasmids .62 4.3.1 The Archetype Plasmid pBL1 from B. lactofermentum ATCC 13869 ......................................................................................62 4.3.2 Other Members of the pBL1 Plasmid Family...................................66 4.4 Structural Organization of the pCG1 Family of Corynebacterial Plasmids ..67 4.4.1 The Archetype Plasmid pCG1 from C. glutamicum ATCC 31808....67 4.4.2 The Small Cryptic Plasmid pGA1 from C. glutamicum LP-6 .........68 4.4.3 Large (Antibiotic Resistance) Plasmids of the pCG1 Family ..........69 4.5 Structural Organization of pXZ10142 and pXZ10145 from C. glutamicum 1014.......................................................................................72 4.6 Genetic Organization of the Basic Replicon of pCRY4 from C. glutamicum LP-6.......................................................................................73 4.7 Host Range of Plasmids from Amino Acid–Producing Corynebacteria.......73 4.8 Concluding Remarks and Perspectives..........................................................74 References................................................................................................................75

4.1 INTRODUCTION Plasmids are extrachromosomal DNA elements that replicate in an autonomous, self-controlled way and that exist with characteristic copy numbers within a host organism [7,10]. They are mostly organized as double-stranded circular DNA molecules with closed strands that can be isolated in the form of supercoiled DNA. Concerning their genetic organization, plasmids contain an essential DNA region, the basic replicon, which comprises the genes and loci involved in autonomous replication and replication control. Additionally, plasmids may encode auxiliary inheritance mechanisms ensuring their stable maintenance and faithful segregation

57

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during cell division [18,15]. Plasmids may also carry genes that could be considered dispensable, although they might play an important or beneficial role for the plasmid itself and/or its host organism. Furthermore, plasmids favor the genetic exchange in bacterial populations since they cannot only incorporate, but also deliver additional genetic material by recombination processes or transposition. This so-called gene load may be involved, for instance, in antibiotic or toxic heavy metal resistance or it may confer favorable physiological traits on the host [68]. Since plasmids can be introduced into new host species by a variety of transfer mechanisms, they can be considered to represent an easy, accessible pool of horizontal mobile DNA, which is shared among bacterial populations. Consequently, plasmids play a major role in enhancing the genetic diversity and adaptation of bacteria by transferring their gene load into new host species [31]. Originally, plasmids were characterized by their incompatibility (Inc) group, which is a property by which plasmids control replication initiation and stable inheritance [18]. Plasmids of the same Inc group share these replication and inheritance functions and cannot be stably co-inherited. Nowadays, plasmids can be characterized also according to conserved structural and genetic features deduced from their nucleotide sequences. In particular, the molecular mechanism of plasmid DNA replication and the degree of amino acid sequence similarity between replication initiator proteins are criteria that can be applied for plasmid classification [6,20]. Despite their autonomous replication, plasmids extensively use the replication machinery of the host organism [10]. However, the requirement of a plasmid-encoded initiator in most plasmid families is reflected by the presence of DNA cognate sites in the origin of replication, where specific protein–DNA interactions take place during replication initiation [7,10]. Circular bacterial plasmids use three modes of DNA replication, known as rolling circle (RC) replication, theta replication, and strand displacement replication [7]. Rolling circle replication is most commonly found in plasmids from Gram-positive bacteria. A comprehensive model of plasmid rolling circle replication is illustrated in Figure 4.1. This mechanism of plasmid DNA replication is unidirectional and asymmetric because syntheses of the leading and lagging strands are uncoupled. Theta replication involves the melting of the parental DNA strands and synthesis of a primer RNA (Figure 4.2). Plasmid DNA synthesis is begun subsequently by covalent extension of the RNA primer and is continuous on the leading strand and discontinuous on the lagging strand. Theta replication can be either uni- or bidirectional [7]. Whereas plasmids replicating according to the rolling circle and the theta mechanism are present in amino acid–producing corynebacteria, the third type of replication by strand displacement has not yet been found in corynebacterial species. Based on homology among their basic replicons, plasmids using rolling circle replication have been divided into four groups (Figure 4.3A), which are represented by the archetype plasmids pT181, pC194, pMV158, and pSN2 [7]. In this context, it is noteworthy that a number of plasmids have been isolated from amino acid–producing and pathogenic corynebacteria, whose replication initiator proteins share no significant amino acid sequence similarity with the archetypal plasmid families [38,66,77]. These corynebacterial plasmids were already proposed to represent a fifth basic group of plasmids using the rolling circle mode of DNA replication [38].

Native Plasmids of Amino Acid–Producing Corynebacteria

Rep homodimer

3′-OH

dso

59

ccc form

SSB DNA Pol lll Helicase

DNA gyrase

DNA gyrase

Rep heterodimer

DNA Pol l DNA Pol lll

DNA Pol l

sso

RNA Pol

sso

sso

FIGURE 4.1 Model for rolling circle (RC) replication of plasmid DNA. Rolling circle replication is initiated by the plasmid-encoded replication protein at the double-strand origin dso (upper left). Two loci have been identified within the dso sequence, the bind and the nic region. The former is the binding site for the replication initiator protein, and the latter contains the nick site where the plasmid DNA is cleaved. DNA cleavage results in a site-specific nick in the parental (+)-strand and leaves a 3′-OH end, which is used as primer for leading strand synthesis. Elongation is probably performed by host proteins and is accompanied by displacement of the parental (+)-strand. A double-stranded plasmid molecule is consequently generated, consisting of the parental (−)-strand and the newly synthesized (+)-strand (middle right). The released Rep heterodimer is composed of an intact replication initiator protein and a modified subunit, which is inactivated by a short oligonucleotide derived from a reconstituted nic locus. A further replication product is a single-stranded DNA intermediate, which corresponds to the parental (+)-strand (lower right). Lagging-strand synthesis, initiating at the single-strand origin sso, converts the single-stranded DNA intermediate into a double-stranded plasmid molecule. Finally, the replication products will be supercoiled by the host DNA gyrase. For details, see [7,10].

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Handbook of Corynebacterium glutamicum

ori

Rep protein

pRNA

DNA Pol l

primosome

replication fork movement

DNA Pol lll

FIGURE 4.2 Early intermediates during theta replication of plasmid DNA. Theta replication involves DNA strand opening at an origin sequence, followed by initiation of leading-strand synthesis. This replication mode can start from one origin or from several, and replication can be either uni- or bidirectional [10]. To simplify, only a unidirectional mode of plasmid DNA replication from a single origin (ori) is shown. During replication of plasmids belonging to the ColE2-P9 family, the replication protein Rep binds specifically to the origin, synthesizing a unique primer RNA [58]. The RNA primer (pRNA) is used by DNA polymerase I to initiate leading-strand synthesis (DNA Pol I-dependent replicon). DNA synthesis progresses in the same direction as rep transcription. Primer synthesis for initiation of lagging-strand replication is directed by the primosome complex, followed by switching of replication from DNA polymerase I to DNA polymerase III holoenzyme [10].

The name pNG2 family was suggested for this group of plasmids, according to the representative replicon from C. diphtheriae [66]. Likewise, plasmids using theta replication can be subdivided into various classes based on specific components participating in the replication process [10]. With some exceptions (e.g., ColE1 family plasmids), the theta mechanism of DNA replication requires a plasmidencoded initiator protein. Additionally, some plasmids may require host DNA polymerase I during early stages of leading strand synthesis. Numerous DNA polymerase I-dependent plasmids have been identified and are tentatively classified into five

Native Plasmids of Amino Acid–Producing Corynebacteria

A

61

Rolling circle (RC) replication

pT181

pC194

pMV158

pSN2

pBL1

pNG2

pCG1

Theta-type replication

B

DNA Pol I-independent replicons iteroncontaining replication

pCRY4

DNA Pol I-dependent replicons

ColE1

ColE2-P9

pAMβ1

pCU1

pJDB23

pXZ10142

FIGURE 4.3 Classification of plasmids from amino acid–producing corynebacteria. Plasmids from amino-acid producing corynebacteria are assigned to rolling circle (RC) replicons (A) or theta replicons (B) according to their mechanism of DNA replication. Based on sequence homologies in the replication control regions and in the respective replication initiator proteins, RC replicons are subdivided into five plasmid groups [7,66]. The archetype plasmids representing each basic group are shown in boxes. The assignment of corynebacterial plasmid families (black boxes) to a basic group of RC replicons is indicated by arrows. Theta plasmids are further subdivided according to their replication requirements. DNA polymerase I–dependent replicons are classified into five groups [10], whereas DNA polymerase I–independent replicons are loosely characterized by iteron sequences [7]. The iteron-containing theta plasmid pCRY4 from C. glutamicum is therefore tentatively classified into the DNA polymerase I–independent group of plasmids.

families according to sequence homologies and replication requirements. These families are represented by the archetype plasmids ColE1, ColE2-P9, pAMβ1, pCU1, and pJDB23 [10].

4.2 ISOLATION OF PLASMIDS FROM AMINO ACID–PRODUCING CORYNEBACTERIA Endogenous plasmids are an important prerequisite for the development of recombinant DNA techniques for amino acid–producing corynebacteria. Therefore, the presence of plasmids in corynebacteria was investigated in the early 1980s [33,47,48], and the identified replicons were used subsequently for the construction of cloning

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vectors and the development of methods enabling efficient DNA transfer [25]. A tabular survey of cloning vectors for Corynebacterium glutamicum genetics is provided in Chapter 23 (Section 23.6) of this book. Meanwhile, a large number of corynebacterial isolates was screened for the presence of plasmid replicons [67]. The most salient features of the identified plasmids from C. glutamicum, C. callunae, C. efficiens, and Brevibacterium stationis are listed in Table 4.1. A total of 30 endogenous plasmids have been discovered, ranging in size from 2.4 to 95 kbp. A few strains such as C. glutamicum LP-6 [56,63] and B. lactofermentum ATCC 13869 [48,55] have been reported to carry multiple plasmids, ranging widely in size (Table 4.1). The smaller plasmids are generally cryptic since they do not possess any known phenotypic marker, with the exception of pXZ10145 from C. glutamicum 1014, which carries a chloramphenicol resistance determinant [78]. Antibiotic resistances in amino acid-producing corynebacteria are thus mostly encoded by larger plasmids (Table4.1). Other types of genetic markers were identified on pCGL500 from C. melassecola ATCC 17965 and on pBT40 from B. thiogenitalis ATCC 19240. Plasmid pCGL500 carries the DNA restriction-modification system CmeI, representing an isoschizomer of the well-characterized EcoRI system [4,67], whereas pBT40 confers the ability on its native host strain to utilize fatty acids as sole carbon source [46]. Meanwhile, considerable genetic information is available on the global organization of plasmids from amino acid–producing corynebacteria [67]. Owing to the growing number of completely sequenced plasmids, comparative genetics is a promising approach for identifying conserved nucleotide sequence motifs within noncoding regions of the plasmid genomes and to characterize conserved amino acid sequence motifs in the replication initiator proteins. Conserved structural features of plasmid genomes might be relevant for DNA replication, stable plasmid maintenance, incompatibility, and copy number control in corynebacteria. Very recently, a molecular genetic classification by comparative genomics was performed with sequenced plasmids from C. glutamicum, which were divided into four distinct plasmid families [67]. An assignment of the C. glutamicum plasmid families to the basic groups of plasmid replicons using either rolling circle or theta replication is shown in Figure 4.3. According to conserved structural motifs deduced from nucleotide and protein sequences, plasmids pCE2, pCE3, and pYM2 from C. efficiens [14,40] and pBY503 from B. stationis [49] are similar to plasmids from C. glutamicum and classified into the pCG1 family, whereas pCC1 from C. callunae [45] is a member of the pBL1 family of corynebacterial plasmids.

4.3 STRUCTURAL ORGANIZATION OF THE pBL1 FAMILY OF CORYNEBACTERIAL PLASMIDS 4.3.1 THE ARCHETYPE PLASMID PBL1 ATCC 13869

FROM

B.

LACTOFERMENTUM

The small cryptic plasmid pBL1 was originally isolated from B. lactofermentum ATCC 13869 and its derivatives [47]. Plasmids with very similar or even identical restriction pattern were also described as pAM330 [33], pBB1 [35], pBL25 [22],

pC194

pNG2

ColE2-P9

Unclassified

RC plasmid

Theta plasmid

Theta plasmid

Plasmid Group

RC plasmid

Plasmid Type

pCRY4

pXZ10142

pCG1

pBL1

Corynebacterial Plasmid Family 4,447 bp 4,448 bp 4,457 bp 4,603 bp 4,109 bp 6,758 bp 5,949 bp 3,069 bp 3,054 bp 19,751 bp 15 kbp

23,743 bp 48,672 bp 29,371 bp 4,826 bp 19,218 bp 27,856 bp — 2,444 bp 4,885 bp 48 kbp

pCE2 pCE3 pCG4 pGA1 pGA2 pTET3 pYM2 pXZ10142 pXZ10145 pCRY4

Plasmid Size

pBL1 pAM330 pGX1901 pAG3 pCC1 pCG2 pXZ608 pCG1 pSR1 pAG1 pBY503

Plasmid Name

Cryptic Putative defensin Sm, Spc, Sul Cryptic Cryptic Tc, Sm, Spc, Sul — Cryptic Cm Cryptic

Cryptic Cryptic Cryptic Cryptic Cryptic Cryptic Cryptic Cryptic Cryptic Tc Cryptic

Marker(s)a

TABLE 4.1 Native Plasmids Identified in Amino Acid–Producing Corynebacteria

C. C. C. C. C. C. C. C. C. C.

efficiens YS-314 efficiens YS-314 glutamicum ATCC 31830 glutamicum LP-6 glutamicum LP-6 glutamicum LP-6 efficiens glutamicum 1014 glutamicum 1014 glutamicum LP-6

B. lactofermentum ATCC 13869 B. lactofermentum ATCC 13869 B. lactofermentum ATCC 13869 C. melassecola 22220 C. callunae ATCC 15991 C. glutamicum ATCC 31832 C. glutamicum 227 C. glutamicum ATCC 31808 C. glutamicum ATCC 19223 C. melassecola 22243 B. stationis IFO 12144

Native Corynebacterial Host Strain

AF092037 D00038 X03987 AY172684 AJ308231 AY172685 AF479770 AB027714 Z22927 AF121000 E08498f; D00661f; E08496g; E08497g AP005225 AP005226 AF164956 X90817 AY172687 AJ420072 AB084384 h X72691 U85507 AY172686 h

GenBank Acc. No.

[40] [40] [23,67] [38,56] [56,67] [63] — — [54,78] [63,67]

[47] [33,74] [12,55] [59,67] [45] [42,67] [29] [42] [2,76] [59,62] [26,28,49]

Reference

Native Plasmids of Amino Acid–Producing Corynebacteria 63

h

g

f

e

d

c

b

Unclassified

Corynebacterial Plasmid Family 36 55 48 40 45 55 29

4.1 95 >25 40 24

pBD12b pBL770c pBI68 pBT40 pBY502 pCC2 pCGL500

pCL1d pGX1906 pHM1520e pMA54 pMA90 kbp kbp kbp kbp kbp

kbp kbp kbp kbp kbp kbp kbp

Plasmid Size

Plasmid Name Cryptic Cryptic Cryptic Fatty acid utilization Cryptic Cryptic DNA restrictionmodification system CmeI Cryptic Cryptic Cryptic Cryptic Cryptic

Marker(s)a

C. lilium ATCC 15990 B. lactofermentum ATCC 13869 C. glutamicum ATCC 13058 M. ammoniaphilum ATCC 15354 M. ammoniaphilum ATCC 21490

B. divaricatum ATCC 14020 B. lactofermentum ATCC 13869 B. immariophilum ATCC 14068 B. thiogenitalis ATCC 19240 B. flavum MJ-233 C. callunae ATCC 15991 C. melassecola ATCC 17965

Native Corynebacterial Host Strain

Antibiotic resistance markers; Tc, tetracyline; Sm, streptomycin; Spc, spectinomycin; Sul, sulfamethoxazole; Cm, chloramphenicol. Also identified as pBD14 with a calculated size of 33.6 kb [22]. Also identified as unnamed plasmid with a calculated size of 55.5 kb [21] and with a calculated size of more than 45 kb [75]. The existence of pCL1 in C. lilium ATCC 15990 was not confirmed by other authors [46,63]. The plasmid was originally not named; a plasmid with similar size was detected in C. glutamicum ATCC 19223 [76]. Only the replication region and a partitioning locus were sequenced. Nucleotide sequence of a copy number mutant of the pBY503 replication region [26]. Only the replication region was sequenced.

Unclassified

Unclassified

a

Plasmid Group

Plasmid Type

TABLE 4.1 (continued) Native Plasmids Identified in Amino Acid–Producing Corynebacteria

— — — — —

— — — — — — —

GenBank Acc. No.

[5] [55] [76] [46] [46]

[46] [46] [46] [46] [49] [46] [4,67]

Reference

64 Handbook of Corynebacterium glutamicum

Native Plasmids of Amino Acid–Producing Corynebacteria

65

pBL100 [53], pGX1901 [55], pWS101 [76], and pX18 [75]. Additionally, the complete nucleotide sequences of pBL1 (GenBank Acc. No. AF092037), pAM330 [74], and pGX1901 [12] were determined, revealing only minor differences (Table 4.1). A deduced physical and genetic map of pBL1 is shown in Figure 4.4. Considering the similarity of the restriction maps and of the nucleotide sequence data as well as the source of plasmid isolation, the specifically named plasmids are now uniformly referred to as pBL1. The copy number of recombinant pBL1 derivatives was estimated between 10 and 30 copies per cell [33,47] and alternatively between 8 and 30 copies per C. glutamicum chromosome [43,69]. Analysis of the complete nucleotide sequence of pBL1 revealed the presence of five coding regions, designated ORF1 to ORF5 (Figure 4.4). The minimal plasmid region involved in autonomous replication of pBL1 in C. glutamicum is localized on a 1.8-kbp HindII-SphI DNA fragment [11]. This DNA region contains ORF1 and ORF5 (Figure 4.4), which are essential for pBL1 replication, since deletion of either coding region results in the inability of hybrid plasmids to stably transform C. glutamicum cells. Comparative analysis of the ORF1 nucleotide sequence indicated that it encodes the replication protein of pBL1, which shows homology to the replication initiator proteins of the Streptomyces plasmids pIJ101 and pJV1 [11]. Plasmids pIJ101 and pJV1 represent a distinct subgroup of the pC194 family, which is one of the basic groups of plasmids replicating by the rolling circle mechanism [7]. Furthermore, the ORF1 protein of pBL1 contains conserved amino acid sequences resembling the two His motif and the Yuxk motif of initiator proteins for rolling circle DNA replication [20]. The Yuxk motif includes the DNA-linking tyrosine residue, which forms a covalent link with the nicked plasmid DNA, whereas the conserved histidine residues of the two His motif might be involved in metal ion coordination required for the activity of the replication protein [20]. Further sequence analysis led to the identification of both a homologous stretch of DNA resembling the nick site within the double-strand origin of pIJ101 and a potential secondary structure comprising a putative single-strand origin. The obvious accumulation of single-strand DNA in C. glutamicum finally indicated that pBL1 replicates by a rolling circle mechanism [11]. On the other hand, the predicted amino acid sequence of ORF5 revealed no homology with any known protein sequences in databases and its function remains to be elucidated. However, a promoter for this gene is identified and its transcription-initiation site was determined, suggesting that the ORF5 region is indeed transcribed in C. glutamicum and could be involved in replication or replication control of pBL1 [11]. The ORF3 region is not part of the minimal replicon of pBL1 and is thus dispensable for plasmid replication and stable maintenance of pBL1 in C. glutamicum (Figure 4.4). Interestingly, bifunctional vectors containing pBL1 DNA fragments inhibit the growth of Escherichia coli and cause extensive cell filamentation [16]. Chromosome segregation of the E. coli host strain is also severely affected. Detailed genetic analyses provide evidence that a 1.23-kbp AccI-HindIII DNA fragment comprising ORF3 of pBL1 (Figure 4.4) is ultimately responsible for the observed effects on E. coli cells. The growth inhibition is dependent on the gene dose of ORF3 since the inhibitory effect was negligible in the case of hybrid plasmids with a low copy number [16]. It is noteworthy that the deduced ORF3 protein of

66

Handbook of Corynebacterium glutamicum

Acc I, 4442 Hin dlll, 1 Ssp I, 4311 Hin dll, 60

Bgl Il, 1 Hin dllll, 2888 Hin dll, 464

Sph I, 3898 Sph I, 3591 Sph I, 3582

ORF3

ORF4

sso

Sca I, 828

pBL1 4447 bp ORF2

Mun I, 3236 dso

Hin dlll, 90 Bcl I, 219

ORF5

Bsp HI, 683

pCG1 3069 bp

Acc I, 1226 Hin dll, 1402

Hin dll, 1408

ORF1/repA

repA

Sca I, 1110

per orfA2

Sph I, 2071

Ssp I, 1021 Nco I, 1147

Bcl I, 2496 Acc I, 2487 Nae I, 2339

Hin dll, 1766

Nae I, 1778 Eco RI, 1565

Xba I, 2064

Hin dlll, 4815 Bam Hl, 1

Eco RI, 2442

Xba l, 4354 Xba l, 2068

per Xho l, 3992

dso

Pst l, 727

orfA2 pGA1 4826 bp

repA

aes

Hin dlll, 1102 Pst l, 1293

orfB

orf 1 orf 2

Sac Il, 483 pXZ10142 2444 bp

repB

Hin dlll, 1400

Nae I, 584 ori Nde I, 714

Bam Hi, 1607 Eco Rl, 1788

Xba l, 1979 Hin dlll, 2394 ctRNA l, 2071

repA Not I, 1068

FIGURE 4.4 Genetic organization of small cryptic plasmids from C. glutamicum. Shown are the physical and genetic maps of pBL1 from B. lactofermentum ATCC 13869, pCG1 from C. glutamicum ATCC 31808, pGA1 from C. glutamicum LP-6, and pXZ10142 from C. glutamicum 1014. Predicted coding regions are marked by arrows indicating the direction of transcription. The positions of the double-strand origin (dso) and of the single-strand origin (sso) of pBL1 were deduced from nucleotide sequence annotation [11]. The position of the double-strand origin of pGA1 was localized in the distal part of the repA gene [1]. The countertranscribed RNA (ctRNA) in the upstream region of the repA gene is indicated by an arrow [72]. The position of the origin of replication (ori) of pXZ10142 was deduced from comparative analysis with pAL5000-related plasmids [67].

pBL1 exhibits no homology with any existing protein in databases. Likewise, the molecular functions of ORF2 and ORF4 (Figure 4.4) are currently unknown.

4.3.2 OTHER MEMBERS

OF THE PBL1

PLASMID FAMILY

Very recently, further complete nucleotide sequences of small cryptic plasmids from amino acid–producing corynebacteria were determined (Table 4.1). This set of plasmids includes pAG3 from C. melassecola 22220 [67], pCG2 from C. glutamicum ATCC 31832 [67], pCC1 from C. callunae ATCC 15991 (GenBank Acc. No. AJ308231), and pXZ608 from C. glutamicum 227 [29]. Comparative analyses with the deduced replication initiator proteins revealed significant amino acid sequence similarity to

Native Plasmids of Amino Acid–Producing Corynebacteria

67

the ORF1 protein of pBL1, suggesting that these plasmids form a distinct corynebacterial plasmid family, which was designated as the pBL1 family according to its representative replicon [67]. The two His and Yuxk motifs are characteristic features for replication initiator proteins of the pBL1 plasmid family [67]. In addition to the complete nucleotide sequences of pAG3, pCG2, pCC1, and pXZ608, only limited information is available on molecular genetic features of the newly characterized plasmids. The minimal replicon of pXZ608 was identified to reside on a 2.14-kbp SacI-BstEII DNA fragment by means of bifunctional plasmids, which were transferred from an E. coli donor strain to C. glutamicum by conjugation. The respective DNA region of pXZ608 contains only the repA gene, which is essential for replication in C. glutamicum [29]. Plasmid pCC1 from C. callunae was solely used for the construction of hybrid plasmids for gene cloning [45,52,71]. The copy number of this plasmid was estimated to be about 30 copies per chromosome [45]. Experiments regarding the compatibility of pCC1 with pGA1 from C. glutamicum LP-6 (Table 4.1) are somehow contradictory since compatibility and incompatibility between both replicons was observed [56,71]. Furthermore, it was found that pBL1 derivatives are compatible with pCC1 in C. callunae and in C. glutamicum [48,71].

4.4 STRUCTURAL ORGANIZATION OF THE pCG1 FAMILY OF CORYNEBACTERIAL PLASMIDS 4.4.1 THE ARCHETYPE PLASMID PCG1 ATCC 31808

FROM

C.

GLUTAMICUM

The plasmid pCG1 is a small cryptic plasmid, originally isolated from C. glutamicum ATCC 31808 [42]. Plasmids with virtually identical restriction maps and genetic organization were identified in other C. glutamicum strains and designated pHM1519 [33], pCG100 [53,69], and pSR1 [76]. These plasmids can therefore be considered as closely related to pCG1, which is also obvious when comparing the almost identical nucleotide sequences of pCG1 (GenBank Acc. No. AB027714) and pSR1 [2]. The copy number of pCG100 and pSR1 is about 30 copies per chromosome in C. glutamicum [38,69]. Bioinformatics reannotation of the complete pCG1 plasmid sequence revealed the presence of three coding regions [67]. A physical and genetic map of the re-annotated pCG1 sequence is shown in Figure 4.4. Mutagenesis data for Tn5 and deletion mutants of pSR1 have localized the minimal replicon within a 2.1-kbp NcoI-BclI DNA fragment containing a single coding region [2]. The predicted protein exhibits significant amino acid sequence similarity to the replication protein of pNG2 from C. diphtheriae, indicating that both plasmids are members of the same family of corynebacterial plasmids [2,66]. A derivative of pNG2 was already shown to replicate by the rolling circle mechanism [77], suggesting the same mode of DNA replication for pSR1 from C. glutamicum. Likewise, the minimal region for autonomous replication of pCG100 in C. glutamicum was localized on a 1.9-kbp NcoI-BglII restriction fragment [69]. A 380-bp HindIII-SphI DNA fragment is able to replicate in the presence of the parental plasmid, which presumably provides a necessary trans-acting replication factor.

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Using a T7 expression system, five pCG100-encoded proteins were synthesized in E. coli when a number of overlapping DNA fragments was cloned in both orientations downstream of the strong Φ10 promoter [69]. This result is somewhat surprising when one considers that only three coding regions were identified by complete nucleotide sequence analysis. Comparative genomics studies with the replication initiator protein encoded by pCG1 identified homology only among respective proteins from corynebacterial plasmids, indicating that the corresponding replicons represent a novel family of plasmids using the rolling circle mode of DNA replication [38,66,77]. A group of such plasmids from amino acid–producing corynebacteria was recently named the pCG1 family (Table 4.1) according to its representative replicon [67]. The pCG1 family of corynebacterial plasmids is thus part of a much larger family (the pNG2 family) that also includes replicons from pathogenic Corynebacterium species [66]. Members of the pCG1 family are characterized by a conserved Yuxk/Yux3k motif within the predicted replication proteins [67]. The conserved Yux3k amino acid sequence strongly resembles the Yuxk motif of replication initiator proteins for rolling circle DNA replication [20]. The original Yuxk motif is only present in the replication proteins of pCG1and pGA1 from C. glutamicum and of pYM2 from C. efficiens, suggesting that the pCG1 family of corynebacterial plasmids can be divided into two subfamilies. The pCG1 family is also characterized by a typical noncoding feature, termed a 22-bp box of corynebacterial plasmids. This highly conserved nucleotide sequence motif (5′-CrTAAGCArwAhACGGTTCCCC-3′) is located downstream of the repA gene and present in one or two copies per plasmid genome [66].

4.4.2 THE SMALL CRYPTIC PLASMID PGA1 FROM C. GLUTAMICUM LP-6 The cryptic plasmid pGA1 from C. glutamicum LP-6 is the best-studied member of the pCG1 family of corynebacterial plasmids (Table 4.1; Figure 4.4). Plasmid pGA1 is compatible with pCG1 [56,60] and present in approximately 34 copies per C. glutamicum chromosome [38]. The function of the repA gene of pGA1 was confirmed by deletion mapping of a minimal replicating fragment [38]. Additionally, the double-strand origin of replication was precisely localized in the distal part of the repA gene, which differs from origin positions identified in other plasmids using the rolling circle mode of DNA replication [1]. The site- and strand-specific breakage of double-stranded pGA1 DNA occurs within the nucleotide sequence 5′-CTGG↓AT-3′ (nic site). Furthermore, a small countertranscribed RNA (ctRNA) of approximately 89 nucleotides in length is encoded in the upstream region of the repA gene [72]. Inactivation of the ctRNA promoter causes a dramatic increase of the copy number of the respective plasmid, indicating a negative role of the ctRNA in copy number control of pGA1. In addition to the ctRNA, the repA upstream region itself acts as a novel regulatory element negatively influencing repA gene expression [72]. Plasmid pGA1 contains two genes, per and aes, whose products positively influence stable maintenance of the plasmid (Figure 4.4). Derivatives of pGA1 devoid of the per gene, encoding a positive effector of replication, exhibit significant effects

Native Plasmids of Amino Acid–Producing Corynebacteria

69

on plasmid copy number and on segregational stability [38]. Deletion of the per gene results in unstable low-copy-number derivatives of pGA1, whereas the presence of per in trans causes a remarkable increase in copy number of the deletion derivatives and at the same time ensures their stable maintenance [38]. A similar positive effector of replication was identified on pCG1/pSR1 by comparative analysis and shown to act in trans on unstable pGA1 derivatives [38]. The small aes gene, encoding an accessory effector of stable maintenance, was shown to increase the segregational stability of pGA1 derivatives in the presence of the main stability determinant per [70].

4.4.3 LARGE (ANTIBIOTIC RESISTANCE) PLASMIDS OF THE PCG1 FAMILY Besides the small cryptic plasmids pCG1 and pGA1, the pCG1 family comprises a set of large plasmids, which were analyzed during systematic DNA-sequencing studies, such as pAG1 [62], pTET3 [63], pCG4 [67], and pGA2 [67], as well as two members, pCE2 and pCE3, which were analyzed in the course of the C. efficiens genome project [40]. Large plasmids appear to exist in low copy number in corynebacteria with approximately 5 to 10 copies per chromosome [26,27,60]. In contrast to the stability mechanisms encoded by pGA1 and pCG1, faithful segregation during cell division of the low-copy-number plasmids might be mediated by a class Ib partitioning system [15]. Characteristic parA and parB genes were identified on the completely sequenced plasmid genomes. In all cases known, the parA gene encodes an ATPase that is essential for the plasmid DNA segregation process, whereas parB encodes a protein that binds to a centromere-like region. Another type of partitioning function that is required for stable maintenance of plasmid DNA was identified on pBY503 from B. stationis (Table 4.1). A 673-bp HindIII-NspV DNA fragment located adjacent to the replication region of pBY503 is able to stabilize corynebacterial plasmids by acting in cis but not in trans [28]. A further remarkable feature deduced from nucleotide sequence annotation of large plasmids of the pCG1 family is the presence of proteins putatively involved in conjugation processes [17]. Putative conjugative relaxases are encoded by pCG4, pGA2, and pTET3 from C. glutamicum and by pCE2 and pCE3 from C. efficiens. Relaxases represent the key enzymes in the initiation of conjugative DNA transfer. Assuming functional similarities with relaxases from broad-host-range plasmids, one might speculate that conjugative transfer of the corynebacterial plasmids involves a nicking reaction at an oriT sequence and the subsequent transfer of a single-stranded DNA molecule [17]. Furthermore, comparative genomics provided insights into the global genetic organization and evolution of large plasmids from the pCG1 family. In particular, a comparison between the genetic maps of the cryptic plasmid pGA2 and the antibiotic resistance plasmid pTET3 is noteworthy (Figure 4.5) since these plasmids stably coexist in C. glutamicum LP-6 (Table 4.1). Both pGA2 and pTET3 are characterized by a unique replication region encoding an initiator protein of the pCG1 family [67]. Alternatively, pGA2 and pTET3 share a virtually identical DNA segment of 5.6 kbp encoding hypothetical proteins, a putative resolvase function and insertion sequences (Figure 4.5). A 2.5-kb DNA region, which is also present in the genome sequence

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FIGURE 4.5 (Color insert follows page 208.) Genetic maps of pGA2 and pTET3 present in C. glutamicum LP-6. Coding regions predicted from complete nucleotide sequences of pGA2 and pTET3 are shown by arrows indicating the direction of transcription. The positions of insertion sequences (IS) within the plasmid backbones are indicated by boxes. Detailed annotations of the plasmid genomes have been deposited in the GenBank database with accession numbers AY172687 and AJ420072, respectively. Green, plasmid replication (repA) and partitioning (parAB) functions; yellow, putative conjugative relaxase gene traA; gray, hypothetical coding regions and putative site-specific methyltransferase gene ssmT; blue, coding regions virtually identical at the nucleotide sequence level in both plasmids, including the resolvase gene res; orange; coding regions virtually identical at the nucleotide sequence level in both plasmids and in the C. glutamicum ATCC 13032 chromosome [63]; red; antibiotic resistance determinant (R-determinant) of pTET3 flanked by identical copies of IS6100.

of C. glutamicum ATCC 13032, is an integral part of this shared DNA segment, indicating that gene transfer occurred between the plasmid replicons and the chromosome [63]. Additionally, pTET3 carries a large resistance determinant, which is obviously flanked by identical copies of the widespread insertion sequence IS6100 (Figure 4.5). Consequently, the genomes of pGA2 and pTET3 can be divided into distinct DNA segments, reflecting the modular evolution of their plasmid backbones. Systematic searches for the presence of plasmid-encoded antibiotic resistances have been performed with C. glutamicum isolates, demonstrating that large plasmids of the pCG1 family (pAG1, pCG4, and pTET3) encode antibiotic resistance determinants against tetracycline, the aminoglycosides streptomycin and spectinomycin, as well as against sulfonamides [23,39,59,63]. Relevant genetic features of the identified antibiotic resistance determinants are listed in Table 4.2. Plasmid-encoded tetracycline resistance in C. glutamicum is mediated by the novel determinants Tet Z and Tet 33, showing homology to Gram-negative-regulated efflux systems [62,63]. Aminoglycoside resistance is mediated by the gene cassettes aadA2 and aadA9, which are part of typical class I integron structures on pCG4 and pTET3 [39,63]. Integrons are genetic elements characterized by their ability to integrate and excise gene cassettes by site-specific recombination [13,44]. Gene cassettes consist of one coding region that is transcribed from a specific promoter within the integron. In

a

Aminoglycoside adenyltransferase Sulfonamide insensitive dihydropteroate synthase Repressor-regulated efflux system Repressor-regulated efflux system 23S rRNA methyltransferase

aadA9 sulI tetA(Z), tetR(Z) tetA(33), tetR(33) rlmAII a

Inducible efflux system Aminoglycoside adenyltransferase

Resistance Mechanism

cmr aadA2

Resistance Determinant

pAG1 pTET3 pAG1

pTET3 pCG4; pTET3

pXZ10145 pCG4

Plasmid

— Flanked by IS6100 sequences —

Gene cassette of class I integron 3′-conserved segment of class I integron

Transposon Tn45 Gene cassette of class I integron

Genetic Element

Initially named orf9 [62]. Resistance is only mediated when acting in synergy with tlrD from Streptomyces fradiae [32].

Tylosin

Sulfafurazol, Sulfamethoxazole Tetracycline

Chloramphenicol Streptomycin, Spectinomycin

Antibiotic

TABLE 4.2 Plasmid-Encoded Antibiotic Resistance Determinants in Amino Acid–Producing Corynebacteria

[62] [63] [32,62]

[63] [39,60,63]

[54,61] [39,63]

Reference

Native Plasmids of Amino Acid–Producing Corynebacteria 71

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particular, the aadA9 resistance determinant represents a novel type of gene cassette, which is currently only known in C. glutamicum. In contrast, the sulI gene is a wellcharacterized genetic element of the 3′-conserved segment of class I integrons and encodes an insensitive dihydropteroate synthase, which confers resistance against sulfonamides [13]. Plasmid pAG1 possesses its own tylosin resistance determinant rlmAII but its host strain C. glutamicum 22243 shows no resistance against this macrolide antibiotic [32,62]. Interestingly, C. glutamicum 22243 becomes resistant against tylosin on transformation with the tlrD gene from Streptomyces fradiae, although tlrD alone is generally insufficient to confer tylosin resistance [32]. Both rlmAII and tlrD encode 23S rRNA methyltransferases obviously acting by a synergistic mechanism to mediate tylosin resistance by methylating nucleotides G748 and A2058 of the 23S rRNA of C. glutamicum [32].

4.5 STRUCTURAL ORGANIZATION OF pXZ10142 AND pXZ10145 FROM C. GLUTAMICUM 1014 The cryptic plasmid pXZ10142 has a size of only 2,444 bp and is the smallest plasmid identified in amino acid–producing corynebacteria to date (Table 4.1). The deduced genetic map of pXZ10142 is shown in Figure 4.4. The plasmid pXZ10142 was originally isolated along with the chloramphenicol-resistance plasmid pXZ10145 from C. glutamicum 1014. Comparison of the nucleotide sequences of both plasmids revealed that pXZ10142 is a spontaneous deletion derivative of pXZ10145, which had lost the chloramphenicol-resistance transposon Tn45 by precise excision at the duplicated insertion site within orf1 [54]. Transposon Tn45 is a transposable element with an unusual genetic structure consisting of an insertion sequence–like transposase gene and the cmr resistance determinant, which encodes an inducible chloramphenicol efflux system (Table 4.2). Two overlapping coding regions, designated repA and repB, were detected in the pXZ10142 sequence and shown to be essential for autonomous replication in C. glutamicum [36]. The predicted RepA protein shows significant amino acid sequence similarities to replicases from pAL5000-related plasmids [67]. The RepA proteins of this group of plasmids are similar to their counterparts on theta replicating ColE2-type plasmids [10,19], suggesting that the replication protein of pXZ10142 may also act as a plasmid-specific primase, which synthesizes a specific primer RNA at the origin of replication [58]. A 15-bp DNA element (5′-AAATATCTGACTTGG-3′) conserved among pAL5000-related plasmids and resembling the core sequence of the origin region in ColE2-type plasmids was noticed upstream of the repA gene of pXZ10142 [8,19]. Additionally, two putative DNA-binding domains are present in the RepB protein sequences of pAL5000-related plasmids, whereas possible DNAbinding regions could not be identified in the respective RepA proteins [8,57]. Therefore, the primase RepA and the DNA-binding protein RepB of pXZ10142 may act together when initiating plasmid DNA replication. The conserved genetic arrangement of the repA and repB genes and the presence of theta motifs in the RepA protein of pXZ10142 strongly indicate that pXZ10142 is a member of the ColE2/ColE2-P9 family and replicates via a DNA polymerase I-dependent theta

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mechanism in C. glutamicum [10]. Plasmids pXZ10142 and pXZ10145 are thus very closely related members of a new corynebacterial plasmid family, named the pXZ10142 family [67].

4.6 GENETIC ORGANIZATION OF THE BASIC REPLICON OF pCRY4 FROM C. GLUTAMICUM LP-6 The large cryptic plasmid pCRY4 was identified in C. glutamicum LP-6 and represents the prototype of a fourth corynebacterial plasmid family [67]. Plasmid pCRY4 has a size of approximately 48 kbp and coexists in C. glutamicum LP-6 together with the cryptic plasmids pGA1 and pGA2 as well as with the antibiotic resistance plasmid pTET3, which are all members of the pCG1 family of corynebacterial plasmids (Tab. 4.1). A minimal DNA region capable of autonomous replication in C. glutamicum is present on a 1,856-bp SphI fragment and has been used for the construction of a cloning vector [60]. The copy number of the recombinant derivative pCRY4-Rep was calculated with three copies per chromosome in C. glutamicum, indicating that pCRY4 is of low copy number. Plasmid pCRY4 is compatible in C. glutamicum LP-6, with vector plasmids derived from pBL1, pCG1, and pAG3 [60]. Nucleotide sequence analysis of the minimal replicon from pCRY4 revealed only one coding region, which obviously encodes the putative replication protein RepA [60]. It is noteworthy that a cluster of five 22-bp direct repeats is present downstream of the repA gene of pCRY4. Clusters of direct repeats, termed iterons, occur in the replication origin region of several plasmids using theta replication, constituting binding sites of the replication protein and thereby playing an important role in the control of plasmid replication [7]. The deduced replication protein of pCRY4 exhibits different degrees of global amino acid sequence similarity to RepA proteins from various plasmids, including the theta-replicating IncW plasmid pSa [41,67] and conjugative broad-host-range plasmids isolated from uncultured bacteria, such as pSB102 [51] and pIPO2 [65]. Multiple amino acid sequence alignments identified the conserved amino acid sequence motif GLPYGxxPR within the replication proteins of these plasmids, which can be used as a characteristic feature for the molecular classification of corynebacterial plasmids into the pCRY4 family [67].

4.7 HOST RANGE OF PLASMIDS FROM AMINO ACID–PRODUCING CORYNEBACTERIA A number of studies have reported on the host range of cryptic plasmids from amino acid–producing corynebacteria (Table 4.3). These studies were mostly initiated to establish vector transfer systems for taxonomically related Gram-positive bacteria with a DNA of high G+C content. Therefore, plasmids were not only transferred to other Corynebacterium species but also to members of the genera Arthrobacter, Bifidobacterium, Brevibacterium, Clavibacter, and Rhodococcus. Derivatives of pBL1 and pCG1 were shown to replicate in a large number of heterologous hosts, including the pathogenic corynebacteria C. diphtheriae [64] and C. pilosum [50].

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TABLE 4.3 Host Range of Plasmids from Amino Acid–Producing Corynebacteria Plasmid

Native Host Species

pBL1

C. glutamicum

pBY503 pCC1

B. stationis C. callunae

pCG1

C. glutamicum

pGA1

C. glutamicum

a

Host Range

Reference

Arthrobacter sp. MIS38 Arthrobacter sp. B3728 Bifidobacterium animalis Brevibacterium linens a Brevibacterium methylicum Corynebacterium callunae Corynebacterium diphtheriae Rhodococcus sp. 312 Corynebacterium glutamicum Arthrobacter sp. B3728 Corynebacterium glutamicum Arthrobacter sp. B3728 Arthrobacter ilicis Bifidobacterium animalis Brevibacterium methylicum Brevibacterium stationis Corynebacterium ammoniagenes Corynebacterium callunae Corynebacterium diphtheriae Corynebacterium pilosum Rhodococcus erythropolis Rhodococcus fascians Rhodococcus sp. 312 Clavibacter michiganensis subsp. michiganensis Corynebacterium callunae Corynebacterium diphtheriae Rhodococcus erythropolis

[34] [53] [3] [46] [37] [46] [64] [9] [49] [52] [45] [53] [50] [3] [37] [50] [50] [50] [64] [50] [73] [50] [5] [24] [56] [64] [73]

Could not be reproduced in other laboratories [30].

4.8 CONCLUDING REMARKS AND PERSPECTIVES A very good overview now exists on the molecular biology of plasmids from amino acid–producing corynebacteria, mainly owing to the growing number of completely sequenced plasmid genomes. Subsequent nucleotide sequence annotation by bioinformatics provides not only a wealth of genetic information, but also allows the classification of corynebacterial plasmids into distinct plasmid families [67]. Most corynebacterial plasmids replicate by a rolling circle mechanism and belong to the pBL1 or pCG1 family, whereas plasmids pXZ10142 and pCRY4 are thought to replicate by a theta mechanism. Nevertheless, several large plasmids from corynebacteria are characterized insufficiently and it would be interesting to define and classify their basic replicons by nucleotide sequence analysis. Such efforts could complete the current view on the replicon types present in amino acid–producing

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corynebacteria. Plasmids of the pCG1 family are of special interest since they can be regarded as members of a fifth basic group of plasmids using rolling circle DNA replication, which was identified only in corynebacteria [38,66,77]. Investigating replication, stability, and maintenance functions of pNG2/pCG1 family plasmid is thus of general interest for plasmid biology. In particular, pGA1 from C. glutamicum LP-6 was analyzed intensively by Neˇsvera and co-workers during recent years and has become a model system for this new group of plasmids [1,38,70,72]. However, further studies are necessary to elucidate the interactions of the identified elements in pGA1 replication, stability, and copy number control. One should also keep in mind that large (low-copy-number) plasmids of the pCG1 family use a different mechanism for plasmid maintenance by encoding a class Ib partitioning system [15]. Identifying the genetic and physiological parameters critical for a proper functioning of the partitioning system would greatly extend our knowledge on this class of plasmids. A further interesting feature deduced from nucleotide sequence annotations of large corynebacterial plasmids is the presence of traA genes encoding putative conjugative relaxases [17]. Therefore, it would be interesting to investigate the capacity of conjugative DNA transfer in corynebacteria by means of plasmids belonging to the pCG1 family. This aspect of plasmid biology is closely linked to a more systematic analysis of the host range of corynebacterial plasmids since more information is required to determine their contribution to horizontal gene transfer in Gram-positive bacteria. In conclusion, it is obvious that future work with corynebacterial plasmids will be greatly facilitated by improved knowledge about their global genetic organization, which allows us to design experiments to elucidate specific plasmid functions in detail.

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27. Kronemeyer W, Peekhaus N, Krämer R, Sahm H, and Eggeling L. (1995) Structure of the gluABCD cluster encoding the glutamate uptake system of Corynebacterium glutamicum. J. Bacteriol. 177, 1152–1158. 28. Kurusu Y, Satoh Y, Inui M, Kohama K, Kobayashi M, Terasawa M, and Yukawa H. (1991) Identification of plasmid partition function in coryneform bacteria. Appl. Environ. Microbiol. 57:759–764. 29. Lei C, Ren Z, Yang W, Chen Y, Chen D, Liu M, Yan W, and Zheng Z. (2002) Characterization of a novel plasmid pXZ608 from Corynebacterium glutamicum. FEMS Microbiol. Lett. 216:71–75. 30. Leret V, Trautwetter A, Rincé A, and Blanco C. (1998) pBLA8, from Brevibacterium linens, belongs to a gram-positive subfamily of ColE2-related plasmids. Microbiology 144:2827–2836. 31. Lilley A, Young P, and Bailey M. (2000) Bacterial population genetics: do plasmids maintain bacterial diversity and adaptation? In The Horizontal Gene Pool: Bacterial Plasmids and Gene Spread, Thomas CM (Ed.), Harwood Academic Publishers, Amsterdam, pp. 287–300. 32. Liu M and Douthwaite S. (2002) Resistance to the macrolide antibiotic tylosin is conferred by single methylations at 23S rRNA nucleotides G748 and A2058 acting in synergy. Proc. Natl. Acad. Sci. USA 99:14658–14663. 33. Miwa K, Matsui H, Terabe M, Nakamori S, Sano K, and Momose H. (1984) Cryptic plasmids in glutamic acid-producing bacteria. Agric. Biol. Chem. 48:2901–2903. 34. Morikawa M, Daido H, Pongpobpibool S, and Imanaka T. (1994) Construction of a new host-vector system in Arthrobacter sp. and cloning of the lipase gene. Appl. Microbiol. Biotechnol. 42:300–303. 35. Mukherjee KJ, Deb JK, and Ramachandran KB. (1990) Construction of vector of Brevibacterium lactofermentum and study of its stability in continuous culture. J. Biotechnol. 16:109–122. 36. Na S, Shen T, Jia P, Men D, and Chen Q. (1991) Characterization of the natural deletion mutant of plasmid pXZ10145 in Corynebacterium glutamicum and construction of a recombinant plasmid. Chin. J. Biotechnol. 7:271–277. ˘ 37. Nesˇvera J, Hochmannová J, Pátek M, Sroglová A, and Bevá˘rova V. (1994) Transfer of the broad-host-range IncQ plasmid RSF1010 and other plasmid vectors to the gram-positive methylothroph Brevibacterium methylicum by electrotransformation. Appl. Microbiol. Biotechnol. 40:864–866. 38. Nesˇvera J, Pátek M, Hochmannová J, Abrhámová Z, Bevá˘rova V, Jelínková M, and Vohradsk´y J. (1997) Plasmid pGA1 from Corynebacterium glutamicum codes for a gene product that positively influences plasmid copy number. J. Bacteriol. 179:1525–1532. 39. Nesˇvera J, Hochmannová J, and Pátek M. (1998) An integron of class 1 is present on the plasmid pCG4 from gram-positive bacterium Corynebacterium glutamicum. FEMS Microbiol. Lett. 169:391–395. 40. Nishio Y, Nakamura Y, Kawarabayasi Y, Usuda Y, Kimura E, Sugimoto S, Matsui K, Yamagishi A, Kikuchi H, Ikeo K, and Gojobori T. (2003) Comparative complete genome sequence analysis of the amino acid replacements responsible for the thermostability of Corynebacterium efficiens. Genome Res. 13:1572–1579. 41. Okumura MS and Kado CI. (1992) The region essential for efficient autonomous replication of pSa in Escherichia coli. Mol. Gen. Genet. 235:55–63. 42. Ozaki A, Katsumata R, Oka T, and Furuya A. (1984) Functional expression of the genes of Escherichia coli in gram-positive Corynebacterium glutamicum. Mol. Gen. Genet. 196:175–178.

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Handbook of Corynebacterium glutamicum 43. Pátek M, Nesˇvera J, and Hochmannová J. (1989) Plasmid cloning vectors replicating in Escherichia coli, amino acid-producing coryneform bacteria and Methylobacillus sp. Appl. Microbiol. Biotechnol. 31:65–69. 44. Recchia GD and Hall RM. (1995) Gene cassettes: a new class of mobile element. Microbiology 141:3015–3027. 45. Sandoval H, Aguilar A, Paniagua C, and Martín JF. (1984) Isolation and physical characterization of plasmid pCC1 from Corynebacterium callunae and construction of hybrid derivatives. Appl. Microbiol. Biotechnol. 19:409–413. 46. Sandoval H, del Real G, Mateos LM, Aguilar A, and Martín JF. (1985) Screening of plasmids in non-pathogenic corynebacteria. FEMS Microbiol. Lett. 27:93–98. 47. Santamaría R, Gil JA, Mesas JM, and Martín JF. (1984) Characterization of an endogenous plasmid and development of cloning vectors and a transformation system in Brevibacterium lactofermentum. J. Gen. Microbiol. 130:2237–2246. 48. Santamaría RI, Gil JA, and Martín JF. (1985) High-frequency transformation of Brevibacterium lactofermentum protoplasts by plasmid DNA. J. Bacteriol. 162:463–467. 49. Satoh Y, Hatakeyama K, Kohama K, Kobayashi M, Kurusu Y, and Yukawa H. (1990) Electrotransformation of intact cells of Brevibacterium flavum MJ-233. J. Indust. Microbiol. 5:159–166. 50. Schäfer A, Kalinowski J, Simon R, Seep-Feldhaus A-H, and Pühler A. (1990) Highfrequency conjugal plasmid transfer from gram-negative Escherichia coli to various gram-positive coryneform bacteria. J. Bacteriol. 172:1663–1666. 51. Schneiker S, Keller M, Dröge M, Lanka E, Pühler A, and Selbitschka W. (2001) The genetic organization and evolution of the broad-host-range mercury resistance plasmid pSB102 isolated from a microbial population residing in the rhizosphere of alfalfa. Nucleic Acids Res. 29:5169–5181. 52. Shaw P-C. (1989) Transformation of a Corynebacterium callunae-Escherichia coli hybrid plasmid to an Arthrobacter species. Agric. Biol. Chem. 53:1717–1719. 53. Shaw P-C and Hartley BS. (1988) A host-vector system for an Arthrobacter species. J. Gen. Microbiol. 134:903–911. 54. Shen T, Jia P, Na S, and Men D. (1993) Determination of nucleotide sequence of Corynebacterium glutamicum plasmid pXZ10145. Chin. J. Biotechnol. 9:171–178. 55. Smith MD, Flickinger JL, Lineberger DW, and Schmidt B. (1986) Protoplast transformation in coryneform bacteria and introduction of an α-amylase gene from Bacillus amyloliquefaciens into Brevibacterium lactofermentum. Appl. Environ. Microbiol. 51:634–639. 56. Sonnen H, Thierbach G, Kautz S, Kalinowski J, Schneider J, Pühler A, and Kutzner HJ. (1991) Characterization of pGA1, a new plasmid from Corynebacterium glutamicum LP-6. Gene 107:69–74. 57. Stolt P and Stoker NG. (1996b) Protein-DNA interactions in the ori region of the Mycobacterium fortuitum plasmid pAL5000. J. Bacteriol. 178:6693–6700. 58. Takechi S, Matsui H, and Itoh T. (1995) Primer RNA synthesis by plasmid-specific Rep protein for initiation of ColE2 DNA replication. EMBO J. 14:5141–5147. 59. Takeda Y, Fujii M, Nakajyoh Y, Nishimura T, and Isshiki S. (1990) Isolation of a tetracycline resistance plasmid from a glutamate-producing corynebacterium, Corynebacterium melassecola. J. Ferment. Bioeng. 70:177–179. 60. Tauch A, Kalinowski J, Pühler A, and Thierbach G. (1999) Plasmide aus Corynebacterium glutamicum und deren Verwendung. German Patent Application No. DE 19953206, issued 11/5/1999.

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61. Tauch A, Krieft S, Kalinowski J, and Pühler A. (2000) The 51,409-bp R-plasmid pTP10 from the multiresistant clinical isolate Corynebacterium striatum M82B is composed of DNA segments initially identified in soil bacteria and in plant, animal, and human pathogens. Mol. Gen. Genet. 263:1–11. 62. Tauch A, Pühler A, Kalinowski J, and Thierbach G. (2000) TetZ, a new tetracycline resistance determinant discovered in gram-positive bacteria, shows high homology to gram-negative regulated efflux systems. Plasmid 44:285–291. 63. Tauch A, Götker S, Pühler A, Kalinowski J, and Thierbach G. (2002) The 27.8-kb R-plasmid pTET3 from Corynebacterium glutamicum encodes the aminoglycoside adenyltransferase gene cassette aadA9 and the regulated tetracycline efflux system Tet 33 flanked by active copies of the widespread insertion sequence IS6100. Plasmid 48:117–129. 64. Tauch A, Kirchner O, Löffler B, Götker S, Pühler A, and Kalinowski J. (2002) Efficient electrotransformation of Corynebacterium diphtheriae with a mini-replicon derived from the Corynebacterium glutamicum plasmid pGA1. Curr. Microbiol. 45:362–367. 65. Tauch A, Schneiker S, Selbitschka W, Pühler A, van Overbeek LS, Smalla K, Thomas CM, Bailey MJ, Forney LJ, Weightman A, Ceglowski P, Pembroke T, Tietze E, Schröder G, Lanka E, and van Elsas JD. (2002) The complete nucleotide sequence and environmental distribution of the cryptic, conjugative, broad-host-range plasmid pIPO2 isolated from bacteria of the wheat rhizosphere. Microbiology 148:1637–1653. 66. Tauch A, Bischoff N, Brune I, and Kalinowski J. (2003) Insights into the genetic organization of the Corynebacterium diphtheriae erythromycin resistance plasmid pNG2 deduced from its complete nucleotide sequence. Plasmid 49:63–74. 67. Tauch A, Pühler A, Kalinowski J, and Thierbach G. (2003) Plasmids in Corynebacterium glutamicum and their molecular classification by comparative genomics. J. Biotechnol. 104:27–40. 68. Top EM, Moenne-Loccoz Y, Pembroke T, and Thomas CM. (2000) Phenotypic traits conferred by plasmids. In The Horizontal Gene Pool: Bacterial Plasmids and Gene Spread, Thomas CM (Ed.), Harwood Academic Publishers, Amsterdam, pp. 249–285. 69. Trautwetter A and Blanco C. (1991) Structural organization of the Corynebacterium glutamicum plasmid pCG100. J. Gen. Microbiol. 137:2093–2101. 70. Venkova T, Pátek M, and Nesˇvera J. (2001) Identification of a novel gene involved in stable maintenance of plasmid pGA1 from Corynebacterium glutamicum. Plasmid 46:153–162. 71. Venkova T, Pátek M, and Nesˇvera J. (2002) Characterization of the cryptic plasmid pCC1 from Corynebacterium callunae and its use for vector construction. Plasmid 48:268. 72. Venkova-Canova T, Pátek M, and Nesˇvera J. (2003) Control of rep gene expression in plasmid pGA1 from Corynebacterium glutamicum. J. Bacteriol. 185:2402–2409. ˇ 73. Vesely´ M, Pátek M, Nesˇvera J, Cejková A, Masák J, and Jirkuº V. (2003) Host-vector system for phenol-degrading Rhodococcus erythropolis based on Corynebacterium plasmids. Appl. Microbiol. Biotechnol. 61:523–527. 74. Yamaguchi R, Terabe M, Miwa K, Tsuchiya M, Takagi H, Morinaga Y, Nakamori S, Sano K, Momose H, and Yamazaki A. (1986) Determination of the complete nucleotide sequence of Brevibacterium lactofermentum plasmid pAM330 and analysis of its genetic information. Agric. Biol. Chem. 50:2771–2778. 75. Yeh P, Oreglia J, Prévots F, and Sicard AM. (1986) A shuttle vector system for Brevibacterium lactofermentum. Gene 47:301–306.

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5

Regulation of Gene Expression M. Pátek

CONTENTS 5.1 Introduction ....................................................................................................81 5.2 Promoters .......................................................................................................81 5.3 RNA Polymerase and Sigma Factors ............................................................85 5.4 Regulation of Transcription Initiation ...........................................................87 5.5 Transcriptional Attenuation............................................................................90 5.6 Leaderless Transcripts....................................................................................90 5.7 Strategies of Modulation of Gene Expression ..............................................92 Acknowledgments....................................................................................................93 References................................................................................................................94

5.1 INTRODUCTION Corynebacterium glutamicum, like other organisms, must react to changes in its environment to ensure that nutrition sources will be utilized economically and that the cell will adapt to various conditions. To meet these demands, the cell is equipped with numerous types of control over its metabolic activities. The primary and major point of control of gene expression is transcription initiation, which represents the first step in the flow of genetic information. Since most examples of regulation of gene expression in C. glutamicum describe transcriptional control, the main focus of this chapter is this level of regulation.

5.2 PROMOTERS Transcription of the coding strand of DNA into mRNA starts with binding of RNA polymerase (RNAP) to a promoter. The sigma factor is the subunit of the RNAP holoenzyme that ensures the specific binding to the promoter and transcription initiation at the proper site. C. glutamicum encodes several sigma factors, which direct RNAP to different classes of promoters. The set of experimentally determined C. glutamicum promoter sequences that are supposed to be recognized by the primary sigma factor encoded by sigA [14,47] is compiled in Figure 5.1. The individual promoters differ in their sequence and consequently in their strength, i.e., the 81

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P-aceA AACCCAAACCGATTGACGCACCAATGCCCGATGGAGCAATGTGTGAACCACGCCACCACG P-aceB ACGTGATGTGCATCGGTTTGCGTGGTGGCGTGGTTCACACATTGCTCCATCGGGCATTGG P-amt AAACAGAAATCTATAGAACGATAGGTAAAAACTGGACTAGGTTTATCTATAGCGGAATAG P-argS AAGTATGGGTCGTATTCTGTGCGACGGGTGTACCTCGGCTAGAATTTCTCCCCATGACAC P-askP1 AAACACTCCTCTGGCTAGGTAGACACAGTTTATAAAGGTAGAGTTGAGCGGGTAACTGTC P-askP2 GAATACGCTCGTGCATTCAATGTGCCACTTCGCGTACGCTCGTCTTATAGTAATGATCCC P-brnE TGGGAAAGGCTGCTAAATGACAACTGATTTCTCCTGTATTCTCCTTGTTGTCGCAGTATG P-brnF TGCGCAAACTGGCAACAAAACTACCCGGCAATTGTGTGATGATTGTAGTGTGCAAAAAAC P-dapA GCGGGGTTGTTTAACCCCCAAATGAGGGAAGAAGGTAACCTTGAACTCTATGAGCACAGG P-dapB1 AGGTATGGATATCAGCACCTTCTGAACGGGTACGTCTAGACTGGTGGGCGTTTGAAAAAC P-dapB2 GTTTGAAAAACTCTTCGCCCCACGAAAATGAAGGAGCATAATGGGAATCAAGGTTGGCGT P-fda AGGAAATATCACACGACAAAAGTTGAGTGATGCAGGCATAATTGGCTATAGGCAACTGAA P-gap TGATTTTGCATCTGCTGCGAAATCTTTGTTTCCCCGCTAAAGTTGAGGACAGGTTGACAC P-gdh TTTTAATTCTTTGTGGTCATATCTGTGCGACACTGCCATAATTGAACGTGAGCATTTACC P-glnA TTTCAGAAAAGTTTTGATAGATCGACAGGTAATGCATTATACTGACAACGTCGCAAGGAC P-gltA CTCACTTCGGATATGGCTAAACCGCATTTATCGGTATAGCGTGTTAACCGGACCAGATTG P-gltB ACCCTTTTGTTACTTGCGTTGCAGGTAGTGTCCCTGATTTTCTTATTATCGAACGATTGA P-hom ATTTCGGGGCTTTAAAGCAAAAATGAACAGCTTGGTCTATAGTGGCTAGGTACCCTTTTT P-ilvA CCTATGCCAAAGTAGGTGCAATTCTAGGAGAAGATTACACTAGTCAACCATGAGTGAAAC P-ilvB TTTCGTGCGTTGTGGAATTGGAAACTCGATGTGTGTAGCATGACACACCATGACCATTAT P-ilvC GTGACTAGTCAAACACCGTCTAATTACATGTGTGTGGTAGAACAATAATGTAGTTGTCTG P-leuA CTCCCCCGAGTTGCTACCCACACCACAAAGTTGTTGTATGCTTCACCACATGACTTCGCG P-leuB TAGTGGGGGTTGTTGGGCAGGTACGAGCTGTGATCAATCAGCTACACTAGTGAAGTCCAT P-lpdA CGCACCCGGATGCACGACAATGACCCACTAAACACGTATCCTTGAATGCGTGACTGAACA P-lrp GTAGTTTTGTTGCCAGTTTGCGCACCTCAACTAGGCTATTGTGCAATATATGAAGCTAGA P-lysE TAAGAACCAATCATTTTACTTAAGTACTTCCATAGGTCACGATGGTGATCATGGAAATCT P-lysG CATCGTGACCTATGGAAGTACTTAAGTAAAATGATTGGTTCTTAACATGGTTTAATATAG P-out (P-45) TGGTCAGGGATTTTTTCCCGAGGGCACTAATTTTGCTAAAGTAAGTGACGAAGAAGTTCA P-pgk CAATTGAATACCGGTGCCAGCGCCACACAATGTGTGGCAATCTGGGACAGTGCATCACAT P-pta2 TGCAAAGGGTGCTTCGCAACTTGTAACCGCTCCGTATTGTTTTCTACGGCAATAAGCATT P-pta1 CTTTGCAAACCGGGCTGTACGCAAGGCGGACGAACGCTAAACTATGTAAGAAATCACAAC P-pyc AAACAAAAACCGATGTTTGATTGGGGGAATCGGGGGTTACGATACTAGGACGCAGTGACT P-secE (P-37) CCTAATAAATATTGCGAGGGTTCGCGGGATTAATGTACTCTCGAAGGTTGAACACAGGGC P-sigA AACAGGCCCCTTTGTGACATCGGCGCAGTTGTTCAACTATAATGGAACGCTGATCGTGGA P-thrB AAGCCTGTTGTTAAGGCAATCAACAGTGTGATCCGCCTCGAAAGGGACTAATTTTACTGA P-thrC ATATTTGAGACGGTGTGGGGGAGTATTGTGTCACCCCTTGGGATAGGGTTATATCCGTGG P-thrE CTTAGCGTATGTGTACATCACAATGGAATTCGGGGCTAGAGTATCTGGTGAACCGTGCAT P-orf3-aroP ACCAAACGTTGAGTAAGGACAATTGGGTCGCCGAGGAGATCTAATCCTGGTTTGAGTTC P-orfMP GACAGGTGCTACTTCGCGAGCAACTCTTTAGTCAACTACCCTGAATCAAGTGCAAAGCAA P-1A TAAGCGTCACGATAAGAACGAGGGGCAAGGCTGATGTACTCTGTCAACCATGGATAAACC P-2 TAATTGGGGCGCGTGGAGCCATTCGGCTTTCAGTAGTACTTTATTTACTAGCTGCTGTGT P-10 GTTCGTGAGACTGAAGCGAAAACCAATGATCCGCAGTACGGTTATTAATAGAAAATGATA P-13 CTGGTCAAGGATCCGTCCCCGGGGAAGTGGGAAATGCTAAAATGGAACGAGCATTCGCAT P-22A TGTGTAGCTTTGAATTGGCCTTGGTGAATCCAGGCTTATGGTTATCTCTGCAGCTATTTC P-34 AACGTGGCGACTTATGGGATTGGATGCAAACGGTGATGGGGTAGCGGACCCCAACCAAAT P-64 GATTCTGCGCGAGTTCCGCCCACACGTCATCATTACCTATGATGAGAACGGCGGTTACCC P-75 AGCGATTAGCGCGCGCTGAGCTTTAGTTTACAGCTAACATGAGGTGCATAAACAAAACGG P-aes1(pGA1) GTTGATAATCCATCTTGCCTTTCCGCTGCAGATAAATTACGCTGAAAAACATAATGATAA P-aes2(pGA1) TAGAGTTTTGTTCCTTGCCGATGCACTTTCTGCGAGCTACTGTGAAGAAGTGGGATGGCG P-ctRNA(pGA1) CACGCGCTAGCTGTGACTGTGTCCTGCGGATCGGCTAGAGTCATGTCTTGAGTGCTTTCT P-I(pGA1) CGGCCAAAAGTTTTTTGAGCAATCGCAGAAAAAGTTTCACAATAAACGAAAATAAAAAAT P-orfB(pGA1) TCTAACGCATACTTCTCAATACCTAACGCATACCGTATTATTAATTCTTAATACCATTTA P-per(pGA1) TCAGTAATTGGGGCTAGAATTTTTAACGAACGTTCGTTATAATGGTGTCATGACCTTCAG P-rep(pGA1) CACGTAAGGTAGTTAAGCGTTCATTTACGAAGAAAACACGATAAGCTGCACAAATACCTG -35 region

-10 region

[11] [11] [21] [34] [10] [10] [26] [26] [49] [49] [49] [71] [60] [5] [45] [9] [2] [53] [49] [40] [49] [52] [51] [61] [26] [3] [3] [49] [60] [11] [11] [54] [49] [14] [35] [16] [62] [72] [49] [49] [49] [49] [49] [49] [49] [49] [49] [69] [69] [70] [52] [70] [70] [70]

TSP

FIGURE 5.1 DNA sequences of 54 C. glutamicum promoters. Experimentally determined transcriptional start points (TSPs) and the presumed –10 and –35 hexamers are in bold. The sequences (60 bp) are aligned at –10 hexamers. The promoters P-1A to P-75 were shotguncloned from C. glutamicum chromosome [49]. Adapted from Pátek et al. [52], with permission. The references are given in brackets.

frequency at which RNAP initiates transcription at the promoter. The strength of a promoter is a basic factor that affects gene expression. Although the promoters differ, they share some sequence characteristics that enable RNAP to reliably recognize the transcriptional signal. The definition of a promoter consensus sequence may

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prove useful for the following practical purposes: (i) recognition of the promoter sequences in the upstream regions of the genes of interest; (ii) assessment of the strength of the known promoter (transcriptional efficiency of a particular promoter may be estimated from the level to which it conforms the consensus sequence); (iii) prediction whether a gene may be expressed from a heterologous promoter, based on the degree of its similarity to the C. glutamicum consensus promoter; and (iv) design of site-directed mutagenesis aimed at the modulation of the promoter strength. To find common sequence motifs, 54 C. glutamicum promoter sequences (Figure 5.1) were first aligned at the experimentally determined transcriptional start points (TSPs) and computer-aided analysis of this set of sequences (70 bp) was performed by the program PROMSCAN [49]. According to the motifs found and the base distribution at the most conserved positions (on average, –12 to –7 relative to TSP; position of TSP = +1), the consensus TA(T/C)AAT could be determined for the –10 region [52]. This hexamer is closely similar in location and sequence to the Escherichia coli [31] and Bacillus subtilis [18] –10 consensus sequence (TATAAT). However, the overall level of base conservation in C. glutamicum (particularly in the middle positions, –10T/C, –9A, and –8A) is lower in comparison with these bacteria. Outside the –10 hexamer some other bases are weakly conserved. The consensus sequence of the extended –10 region (–17 to –5) can thus be defined as TgtG(c/g)TAtAATGG (bases conserved to more than 40% are shown in capital letters; –10 core hexamer is underlined; Figure 5.2). The sequences were then aligned according to the found –10 motif (Figure 5.1) and another round of computer-aided search for conserved sequences was run. This second analysis using different parameters revealed the alternative motifs TTGTTG, TTTGCC, TTGGCA, and TTGCCA within the –35 region. The degree of base conservation in this region is lower than that in the –10 region of C. glutamicum and also lower than that in the –35 region of E. coli. The trimer TTG, which is conserved in E. coli, was identified in only 8 of 54 C. glutamicum promoters. The blurred definition of the –35 region in C. glutamicum was already apparent from the previous analysis of 33 promoters [49]. A similar situation was described in related species of G+C–rich actinomycetes, e.g., Streptomyces or Mycobacterium, in which the –35 regions differ widely, preventing any definition of their consensus sequences [1,6]. In contrast, the –35 hexamers in promoters of species belonging to A+T–rich Gram-positive bacteria are usually highly similar to the respective consensus (TTGACA) [18]. In addition to the statistical analysis of the aligned promoters, mutagenesis of their key sequence motifs provides information about the importance of particular bases at the conserved positions and their contribution to the strength of the promoters. An extensive mutational study of the C. glutamicum dapA promoter was performed [68] to verify the results of the statistical analysis. The dapA gene codes for dihydrodipicolinate synthase, the first enzyme of the lysine-specific biosynthesis pathway. Its promoter, which is of mediocre strength (20% of the activity of the strong promoter P-45), is probably not regulated, or regulation plays a minor role in its expression [68]. Mutagenesis showed that deletion of the –35 region led to a negligible change of promoter activity (less than 3% decrease). However, as shown

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FIGURE 5.2 Base conservation within the extended –10 regions of 54 C. glutamicum promoter sequences. Positions –17 to –5 relative to transcriptional start point (TSP = +1) and occurrence of the most frequent bases (above the histogram) at the respective positions are shown. Bases occurring in more than 80% of the promoters are capitalized and in bold, bases occurring in more than 40% of the promoters are in capital letters, and less-conserved bases are in small letters. For comparison, the consensus sequence of B. subtilis [18] and E. coli [37] –10 region is shown on top.

in Figure 5.3, base alterations introduced into the–10 sequence can strongly influence the promoter strength. Mutations at the most conserved positions within the –10 hexamer (–12T and –7T) confirmed that T’s at these positions are essential. In addition to these mutations within the core region, those located within the extended –10 region, including –14G (clone C13 ) and –5G (clone O1) are vital for promoter activity. The importance of G at position –14, indicated by its level of conservation in C. glutamicum promoters (48%), was also confirmed by a mutation in the promoter of the InCg integron [44] from the C. glutamicum plasmid pCG4. A transversion G→C at this position resulted in a five-fold decrease of the promoter activity in C. glutamicum [44]. In P-dapA, a single-base alteration –15A→T (clone C20) that created a TG sequence at positions –15 to –14, increased the promoter strength fourfold [68]. All other mutations introduced into the –10 region of P-dapA also confirmed the conclusions based on the statistical analysis of the promoter sequences (Figure 5.2). Whereas the –35 region, whose consensus sequence is still elusive, plays probably a marginal role in many C. glutamicum promoters, the consensus sequence of the extended –10 region (positions –17 to –5) is well defined. Based on the consensus sequence derived from the set of native promoters and on mutational studies, the extended –10 sequence TGTG(C/G)TATAATGG, which characterizes

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FIGURE 5.3 Mutagenesis of the –10 region of the dapA promoter [68]. The substituted bases are shown in bold letters. Chloramphenicol acetyltransferase (CAT) activity reporting the promoter strength of mutant clones is expressed relative to activity of wild-type promoter (P-dapA) set to 1. Standard deviations from three independent measurements are indicated with error bars. On top, the extended –10 region of P-dapA is compared with the consensus deduced from both statistical analysis and site-specific mutagenesis. In this consensus, the positions conserved to more than 40% and two less-conserved positions (–10T and –15T) at which T’s were proved to increase the strength of the promoter are shown in capital letters.

a hypothetical strong C. glutamicum promoter (initiating transcription with a high frequency), may be defined.

5.3 RNA POLYMERASE AND SIGMA FACTORS The subunits of C. glutamicum RNA polymerase core enzyme, α, β, β′, and ω encoded by the genes rpoA (NCgl0540), rpoB (NCgl0471), rpoC (NCgl0472), and rpoZ (NCgl1543), respectively, are according to the analysis of their amino acid (aa) sequences most similar to the respective subunits from C. efficiens (86 to 96%) and C. diphtheriae (76 to 88%). Less similarity can be found to the RNAP subunits from mycobacteria (67 to 77%), streptomycetes (63 to 71%), and bacilli (about 50%). According to the similarity search, seven different sigma subunits (factors) of RNAP holoenzyme are encoded by C. glutamicum genome. Based on the homology with Mycobacterium tuberculosis sigma factors [33], which are most similar in their aa sequences to corynebacterial sigma factors, the respective C. glutamicum genes can

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be designated sigA (NCgl1836), sigB (NCgl1844), sigC (NCgl0250), sigD (NCgl0575), sigE (NCgl1075), sigH (NCgl0733), and sigM (NCgl2983). No sigma factor homologous to sigma-54, which is involved, e.g., in the transcription of genes governing nitrogen metabolism in E. coli or B. subtilis, is encoded by the C. glutamicum genome, similarly as in related bacteria such as C. diphtheriae and M. tuberculosis [66]. SigA is a primary sigma factor, whereas SigB represents a nonessential primary-like sigma factor. The other sigma factors belong to the group of environmentally responsive transcriptional regulators called ECF (extracytoplasmic function) [20]. These C. glutamicum sigma factors are probably involved in regulation of gene expression under starvation conditions and in response to various kinds of environmental stresses. C. efficiens codes for the same seven sigma factors, while C. diphtheriae codes for nine sigma factors (SigA, SigB, and seven ECF-type sigma factors). The related pathogen M. tuberculosis codes for 13 sigma factors [33]. Seven different sigma factors have also been found in E. coli, whereas many more have been detected in the sporulating Gram-positive bacterium Streptomyces coelicolor (63 sigma factors) [12]. The function of sigma factors is to navigate RNAP so that it binds exclusively to specific types of promoters. A sigma factor thus functions as a global regulator of transcription that ensures a coordinated expression of a class of genes forming a regulon. The primary sigma factor of C. glutamicum encoded by sigA directs the RNAP most probably to the promoters of a majority of genes, called housekeeping genes [14,47]. The sigA gene is transcribed constitutively during batch cultivation. The sigma factor encoded by sigB plays a role in the transcription of genes involved in response to environmental stress factors such as cold and heat shock, acidic pH, salt, and ethanol [15]. However, the aa sequence of subregion 2.4 of SigB, which is involved in recognition of the –10 promoter region, is highly similar to that of SigA. This suggests that SigA and SigB recognize similar promoter motifs and that their respective regulons therefore partially overlap. Similarly, it is assumed that SigB of M. tuberculosis functions as an alternative to SigA under the various stress conditions, when SigA is inactive [33]. Activity of the C. glutamicum sigB promoter itself is induced by acids, ethanol, and cold shock. The sequence upstream of the mapped TSP of P-sigB (TGGGAACTT-N15-CGTTGAA-N6-G) is almost identical to the motif of the M. tuberculosis sigB promoter, which is under the control of two ECF sigma factors, SigH and SigE [33]. The P-sigB from C. glutamicum might therefore also be recognized by an ECF sigma factor [15]. The transcript of C. glutamicum sigE was identified as one of the heat-shock–induced mRNAs in the genome-wide transcriptional study based on a DNA-microarray analysis [42]. In agreement with this result, the activity of the sigE promoter was found to be increased by a heat shock (author’s unpublished results). Activity of some sigma factors is regulated (inhibited) by anti-sigma factors via their reversible interaction [19]. According to the homology with anti-sigma factors from M. tuberculosis [65], three genes coding for putative anti-sigma factors have been identified within the C. glutamicum genome. Two of them (NCgl1076; no NCgl assigned to the other) are located downstream of sigE and sigH, respectively. A similar arrangement is present in C. efficiens and C. diphtheriae. In the case of sigH, the gene coding for the anti-sigma factor follows immediately downstream in all three corynebacteria, which suggests that the two genes are co-transcribed. The third

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gene coding for a putative anti-sigma factor (NCgl0852) is not associated with any gene for a sigma factor in C. glutamicum. A homologous anti-sigma factor also occurs in other corynebacteria and in mycobacteria. Identification of the regulons controlled by individual C. glutamicum sigma factors, description of the sequence features of the promoters recognized by these sigma factors, and elucidation of the functions of the proteins encoded by the respective genes are goals of future investigation.

5.4 REGULATION OF TRANSCRIPTION INITIATION The activity of promoters may be negatively or positively affected by changes of DNA conformation and by regulatory proteins interacting with the DNA within or near the promoter region or with RNAP. Although directed alterations of expression of the genes involved in amino acid biosynthesis pathways are of exceptional practical importance, little is known about the specific mechanisms of transcriptional control in C. glutamicum. According to the C. glutamicum genome analysis (Regulatory functions gene occurrence for each genome, http://www.tigr.org/tigr-scripts/ CMR2/genechart.spl?types=Regulatory_functions), 176 genes code for the regulatory proteins. Considering the total number of 3,099 genes [22] or 2,993 genes found in a revised version of the C. glutamicum genome sequence (http://www.ncbi.nih. gov/genomes/lproks.cgi), it makes 5.68% or 5.88%, respectively, of all genes. This is a similar percentage as in E. coli (4.28%) and in B. subtilis (5.64%), but less than in S. coelicolor (9.46%). The global repressor protein AmtR that controls expression of the genes involved in uptake and assimilation of nitrogen sources on the level of transcription is one of a few regulatory proteins studied so far [21]. Using gel retardation experiments and deletion analysis, the exact motif for binding of AmtR within the promoter regions of several genes (amt, amtB, gltBD, glnA, glnD, and glnK) was identified. A unique control of the genes expressed in response to nitrogen starvation was described in C. glutamicum [46]. Recently, isolation of the transcriptional repressor McbR of the TetR family allowed identification of six genes (hom, metY, ssuD, cysI, cysK, and metK) that are involved in metabolic pathways leading to methionine and cysteine synthesis [58]. To reveal these genes, proteome analysis of the wild-type strain and the mcbRdeletion strain was applied. The abundant proteins of the mutant strain were identified by mass spectrometry fingerprint analysis utilizing the C. glutamicum genome sequence information. This study provides an example of the efficient use of global approaches based on data from proteome and genome analysis. In two cases, potential transcriptional activators, participating in expression control of genes for amino acid export carriers, were described in C. glutamicum. Expression of the lysE gene that codes for the lysine-exporting carrier, depends on the LysG regulator encoded by the gene transcribed divergently from lysE [3]. A similar arrangement was found for the genes brnFE and lrp (coding for a homologue of the global regulator found in many bacteria). The brnFE genes code for the export carrier of isoleucine, valine, and leucine. The divergently transcribed lrp gene is essential for the expression of brnFE [26].

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Structural genes coding for enzymes of metabolic pathways or for proteins with related functions may form operons with coordinately controlled transcription. Many genes in C. glutamicum differ in their organization in operons or their separate location on the chromosome from the homologous genes in E. coli or B. subtilis. Thus leuA and leuB genes are separated from the leuC-leuD cluster in C. glutamicum [48] whereas all leu genes form an operon in E. coli [64]. Conversely, trp genes provide an example of a similar organization in these species (see Chapter 21). The C. glutamicum trp genes are organized in an operon [59] resembling the trp operon of E. coli [73]. The effects of mutations within the 14-bp palindromic trp operator located upstream of the promoter of the trp operon were studied [13]. Expression of the amy reporter gene cloned behind the trp promoter with the mutated operator increased five-fold in tryptophan-rich medium in comparison with the wild-type promoter-operator, indicating that the repression by the supposed TrpR repressor was relieved. However, in minimal medium, tryptophan stimulated the expression from mutated operator-promoter. This might be explained by growth rate–dependent expression of the trp operon [13]. Examples of C. glutamicum operons for which the transcripts have been experimentally determined are shown in Figure 5.4. Although promoters and transcripts have been mapped in these gene clusters, experimental data on the regulation of their transcription are in most cases still scarce. The hom-thrB transcription unit of C. glutamicum [35] codes for two of the five enzymes required for synthesis of threonine from aspartate (Figure 5.4A). The other enzymes are encoded by the gene cluster ask-asd [10] and the separated thrC gene [16]. Expression of the hom gene is controlled by transcriptional repressor McbR [58]. The hom gene codes for homoserine dehydrogenase, which is also involved in the biosynthesis of methionine. In the presence of methionine, the abundance of the Hom protein is reduced [58]. Another type of transcriptional regulation of the hom-thrB operon was described by Jetten and co-workers [24]. A sequence capable of forming a hairpin structure with a methionine codon followed by a stop codon at the loop is present immediately upstream of the translation start of hom. It is therefore supposed that the hom-thrB operon expression is regulated by a single stem-loop attenuator structure [24]. However, only a two-fold derepression was demonstrated in the strain, in which this structure was deleted. The ilvBNC operon represents the best-described example of regulation by transcriptional attenuation in C. glutamicum [40]. The transcription of ilvBNC is increased about twice under the conditions of growth limitation by any of the branched-chain amino acids (Val, Leu, and Ile). At the same time, expression of the operon is induced in the presence of α-ketobutyrate, one the substrates of acetohydroxy acid synthase encoded by ilvB and ilvN. Due to internal promoters, two further shorter transcripts covering either ilvN-ilvC or ilvC alone are formed (Figure 5.4B) [40]. Expression of ilvN-ilvC is also increased by α-ketobutyrate in the medium [25]. Moreover, mRNA level of ilvN, but not of ilvB and ilvC was reduced two- to three-fold by addition of 50 to 100 mM valine into glucose minimal medium according to transcriptome analysis [30]. The genes dapA and dapB, which encode two enzymes of lysine and diaminopimelate synthesis, are clustered with two other genes, orf2 (NCg1897) and orf4

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hom

89

thrB

A

ilvB

ilvN

ilvC

B

dapB

orf2

dapA

orf4

C

gap

pgk

tpi

ppc

D

1 kb

FIGURE 5.4 Transcriptional pattern of C. glutamicum operons. Coding regions of the genes are shown as thick arrows, their transcripts as arrows below. Thickness of the arrows indicates the relative quantity of the transcripts. Promoters are indicated by short open arrows. Transcriptional terminators are depicted as stem-and-loop structures. (A) The hom-thrB operon coding for enzymes of threonine biosynthesis pathway [35]. (B) The ilvBNC operon coding for the enzymes of the valine and isoleucine biosynthesis pathway [25]. (C) The dapB-orf2dapA-orf4 operon coding for the enzymes of the lysine biosynthesis pathway [50]. (D) The gap-pgk-tpi-ppc operon coding for the enzymes of glycolysis [60]. Adapted from Pátek et al. [52], with permission.

(NCg1895) [50]. The protein encoded by the nonessential orf2 is similar to the thymidylate synthase complementing protein, whereas the essential protein encoded by orf4 is homologous to metallo-beta-lactamases. There is the same organization of these genes in the genomes of C. diphtheriae and C. efficiens. The transcriptional unit also includes two internal promoters; however, no regulation of this operon was described (Figure 5.4C). A sophisticated transcriptional pattern of the gap-pgk-tpi-ppc cluster (Figure 5.4D), coding for enzymes of glycolysis, is controlled by two promoters and three terminators [60]. Such a complex transcriptional pattern probably ensures fine-tuning of the transcription level of the respective genes.

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5.5 TRANSCRIPTIONAL ATTENUATION Expression of several operons coding for enzymes involved in amino acid biosynthesis in bacteria is regulated by attenuation, a translation-mediated transcription control that monitors the level of the respective amino acids in the cells [28]. In C. glutamicum three transcription units that seem to be regulated by such a mechanism have been studied so far: the ilvBNC operon, the trp operon, and the leuA gene. The genes ilvBN and ilvC code for enzymes involved in the biosynthesis of branched-chain amino acids. The leader region of the ilvBNC operon, coding for a leader peptide of 15 aa, contains seven codons for three branched-chain amino acids (two Ile codons, three Val codons, and two Leu codons). Under starvation conditions for one of the amino acids or for all three, expression of the operon was enhanced about two-fold [40]. However, expression of the cat reporter gene (chloramphenicol acetyltransferase) driven by the cloned ilvB promoter with its regulatory region was enhanced more that 10-fold during valine starvation (author’s unpublished results). Mutagenesis and deletion studies of the region coding for the leader peptide adjacent to the promoter in the ilvBNC operon of C. glutamicum confirmed that transcription of the operon is controlled by a translation-coupled attenuation mechanism [40]. The sequence features between the trp promoter and the translation initiation codon of the trpE gene suggest that expression of the trp operon is also regulated by transcriptional attenuation. The short ORF (trpL) coding for the leader peptide contains three consecutive Trp codons. Mutations in the potential trpL terminator [36] and in the trpL gene [17] in tryptophan-producing strains support the attenuation model. The supposed alternative structures of the transcriptional terminator formed under the conditions of tryptophan surplus and of the antiterminator occurring at tryptophan limitation are shown in Figure 5.5. In the leader region of the leuA gene, a short ORF coding for a potential leader peptide of 18 aa was found to contain four consecutive Leu codons (Figure 5.6) [48]. Mutagenesis of the leader peptide showed that regulation of the leuA transcription is mediated by translation of this coding region and that the Leu codons are essential for the regulation (author’s unpublished results). Under the conditions of leucine starvation of the leu auxotroph, six-fold derepression of the wild-type gene was determined. No derepression was observed when the ATG start codon of the leader peptide was altered to AAG. The structure of the leader region and the results of the mutagenesis of the leader peptide indicate that transcription of the leuA gene is controlled by transcriptional attenuation. The leuA gene of C. glutamicum represents a rare case in which not a multicistronic operon but just a single gene is controlled by attenuation.

5.6 LEADERLESS TRANSCRIPTS The translation initiation rate significantly affects the level of gene expression. In most cases, translation in bacteria is initiated by binding of the Shine-Dalgarno (SD) sequence located at the 5′ end of the mRNA to the complementary 3′ end of 16S rRNA. The SD sequences in most bacteria are similar to the core motif GGAGG with a distance of 5 to 13 bp from the initiation codon [32]. The sequence of a

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FIGURE 5.5 Computer-generated models of alternative secondary structures of trp attenuator [59]. Nucleotides coding for the leader peptide are shown in bold face. (A) Terminator structure. It is supposed to be formed under the conditions of tryptophan excess. A substitution G→A destabilizing the terminator [36] in the mutant strain is underlined. (B) Antiterminator structure. It may be formed at tryptophan starvation. Adapted from Sano and Matsui [59], with permission.

FIGURE 5.6 Mutation in the leader peptide of the leuA attenuator. The amino acid sequence of the putative leader peptide is shown below the respective coding DNA sequence (leuL). The four regulatory Leu codons are in bold face. The mutation T→A within the translation initiation codon ATG (underlined) abolished the translation of leuL.

particular SD site and its spacing to the start codon may considerably influence the efficiency of translational initiation. Neither of these two features has been systematically studied in C. glutamicum. A complete lack of an SD sequence has been found as a striking feature of several C. glutamicum genes. In these cases, the transcriptional start points occur near the translation initiation codons or are even identical with the first nucleotide (nt) of the start codons, which inevitably means that leaderless transcripts are formed. In the genes brnF, lrp, lpdA, ilvA, and leuA (the leader peptide) the transcriptional start and the first base of the translation initiation codon are identical, whereas transcripts of ilvB (the leader peptide) and

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lysE start just 1 nt upstream of the respective initiation codon. The transcript of thrC includes only a 7-nt untranslated sequence. Such transcripts lacking an SD sequence occur infrequently in other actinomycetes and in archaea and rarely in other bacteria. It seems that in C. glutamicum such arrangement is relatively frequent. It has been suggested that the translation of leaderless transcripts is permitted by the complementarity of a downstream box located on the mRNA and a sequence within the 16S rRNA. However, no evidence of such an interaction has so far been produced and probably only the 5′-terminal AUG codon is necessary for initiating translation [39]. In the C. glutamicum genes thrC, brnF, and lpdA, the GUG triplet was identified as the initiation codon. However, it was found that only the AUG start codon supports a high-level expression in E. coli, whereas the GUG codon provides poor expression [39]. In E. coli the leaderless mRNAs are translated more efficiently at low temperature (25˚C) than mRNAs with a Shine-Dalgarno sequence. The mechanism of translation initiation from leaderless transcripts may represent still another regulation of gene expression in C. glutamicum.

5.7 STRATEGIES OF MODULATION OF GENE EXPRESSION Methods of gene engineering have been utilized in C. glutamicum to alter (mostly to increase) expression of the chosen genes with the aim to construct strains excreting commercially important compounds, chiefly amino acids. Many genes involved in amino acid biosynthesis have been cloned on multicopy plasmids, to achieve their amplification in the cells and to remove the supposed bottlenecks in biosynthesis pathways. To overcome repression of genes and inhibition of enzyme activity by the accumulated end product, derepressed genes or genes coding for deregulated enzymes from mutant strains isolated during the era of classical breeding have usually been used for cloning. Although this simple brutal-force approach is not considered a general strategy to obtain a producer strain [29], it has often been successful. A strain producing histidine was thus constructed by cloning the hisG gene encoding feedback-resistant ATP phosphoribosyl transferase [38]. However, severe interventions into regulation of gene expression may lead to metabolic imbalances. Thus, in some cases, cloning of the genes coding for feedback-resistant enzymes on multicopy plasmids resulted in genetically unstable strains with a poor overexpression [41]. A moderate gene amplification, which permits stable expression of deregulated genes, may be achieved by cloning the genes on low-copy-number plasmids. By this approach, the hom gene encoding feedback-resistant homoserine dehydrogenase was cloned together with thrB [57].The plasmid pWK-hom1 based on a low-copy-number replicon with three to four copies per chromosome was introduced into a lysine producer, which resulted in switching to threonine production. Integration of one to three copies of the hom and thrB genes into the chromosome leading to their stable expression represents another strategy to precisely achieve adjusted expression [57]. However, the construction of producer strains may not only require overexpression of amino-acid–specific biosynthesis genes, but also

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of genes determining supply of the precursors. Cloning of the pyc gene coding for pyruvate carboxylase, an anaplerotic enzyme for growth on carbohydrates, improved production of the strains excreting threonine, lysine, and glutamate to various extents (10 to 700%). Low expression of pyruvate carboxylase thus proved to be a major bottleneck for amino acid production by C. glutamicum [55]. The simultaneous overexpression of pyc (pyruvate carboxylase) and ask (aspartate kinase) cloned together on a plasmid led to a significant increase in lysine formation without impairing cell growth [27]. A coordinated, fine-tuned expression of multiple genes thus seems an ideal manner in which to overcome natural regulation of gene expression. The introduction of heterologous genes, which can extend the pathways existing in the cell and widen the cell’s abilities, belongs to the prime procedures of metabolic engineering. In C. glutamicum, introduction of the whole lactose operon from E. coli resulted in a strain capable of growing on lactose [7]. In addition to gene amplification, manipulation of the signals regulating the gene expression may be used in molecular breeding. The cloned lac operon contained the E. coli promoter with a mutation in the –10 hexamer (TATGTT→TATATT) which made it more efficient in C. glutamicum [7]. Strong mutant promoters P-dapA A16 or P-dapA C20 (Figure 5.3) were introduced with a second copy of the dapA gene into the chromosome. The resulting strains displayed three- to four-fold higher dihydrodipicolinate synthase activity and substantially increased lysine accumulation [8]. Apart from native C. glutamicum promoters, alien regulated promoters (e.g., from E. coli) seem to be attractive tools for controlled gene expression. The activity of lacUV5 promoter with negative regulation [67] and of the araBAD promoter with positive control [4] was demonstrated in C. glutamicum. The LacI-repressed promoter P-tac was used to reduce expression of the glyA gene coding for serine hydroxymethyltransferase (involved in threonine degradation) and to increase threonine production [63]. The technique of gene disruption and replacement may be used not only for introduction of DNA sequences into the chromosome but also for gene-directed deletions. Using this method, the genes ilvA and panBC were inactivated, to construct a valine-producing strain [56]. With the introduction of methods for global transcriptional profiling using DNA microarrays [23,30,42] new targets for modulating gene expression might be rapidly identified for the further development of producer strains.

ACKNOWLEDGMENTS I am particularly grateful to B. Eikmanns (Ulm), who initiated the studies on C. glutamicum promoters, to L. Eggeling (Jülich) for the lasting cooperation, to H. Sahm (Jülich) for the generous support, and to J. Nesˇ vera (Praha) for fruitful discussions and for the help during the preparation of the manuscript. Work in the author’s laboratory has been supported by grant 525/04/0548 from the Grant Agency of the Czech Republic, by the grant from EU (VALPAN, QLK3-2000-00497), and by Institutional Research Concept no. AV0Z5020903.

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Handbook of Corynebacterium glutamicum Mitchell JE, Zheng D, Busby SJ, and Minchin SD. (2003) Identification and analysis of ‘extended –10’ promoters in Escherichia coli. Nucleic Acids Res. 31:4689–4695. Mizukami T, Hamu A, Ikeda M, Oka T, and Katsumata R. (1994) Cloning of the ATP phosphoribosyl transferase gene of Corynebacterium glutamicum and application of the gene to L-histidine production. Biosci. Biotechnol. Biochem. 58:635–638. Moll I, Grill S, Gualerzi CO, and Blasi U. (2002) Leaderless mRNAs in bacteria: surprises in ribosomal recruitment and translational control. Mol. Microbiol. 43:239–246. Morbach S, Junger C, Sahm H, and Eggeling L. (2000) Attenuation control of ilvBNC in Corynebacterium glutamicum: evidence of leader peptide formation without the presence of a ribosome binding site. J. Biosci. Bioeng. 90:501–507. Morinaga Y, Takagi H, Ishii Y, Miwa K, Sato K, Nakamori S, and Sano K. (1987) Threonine production by co-existence of cloned genes coding homoserine dehydrogenase and homoserine kinase in Brevibacterium lactofermentum. Agric. Biol. Chem. 51:93–100. Muffler A, Bettermann S, Haushalter M, Horlein A, Neveling U, Schramm M, and Sorgenfrei O. (2002) Genome-wide transcription profiling of Corynebacterium glutamicum after heat shock and during growth on acetate and glucose. J. Biotechnol. 98:255–268. Nesˇvera J, Pátek M, Hochmannová J, Abrhámová Z, Beˇcváová V, Jelínková M, and Vohradsk´y J. (1997) Plasmid pGA1 from Corynebacterium glutamicum codes for a gene product that positively influences plasmid copy number. J. Bacteriol. 179:1525–1532. Nesˇvera J, Hochmannová J, and Pátek M. (1998) An integron of class 1 is present on the plasmid pCG4 from gram-positive bacterium Corynebacterium glutamicum. FEMS Microbiol. Lett. 169:391–395. Nolden L, Farwick M, Krämer R, and Burkovski A. (2001) Glutamine synthetases of Corynebacterium glutamicum: transcriptional control and regulation of activity. FEMS Microbiol. Lett. 201:91–98. Nolden L, Ngouoto-Nkili CE, Bendt AK, Krämer R, and Burkovski A. (2001) Sensing nitrogen limitation in Corynebacterium glutamicum: the role of glnK and glnD. Mol. Microbiol. 42:1281–1295. Oguiza JA, Marcos AT, and Martin JF. (1997) Transcriptional analysis of the sigA and sigB genes of Brevibacterium lactofermentum. FEMS Microbiol. Lett. 153:111–117. Pátek M, Krumbach K, Eggeling L, and Sahm H. (1994) Leucine synthesis in Corynebacterium glutamicum: enzyme activities, structure of leuA, and effect of leuA inactivation on lysine synthesis. Appl. Environ. Microbiol. 60:133–140. Pátek M, Eikmanns BJ, Pátek J, and Sahm H. (1996) Promoters from Corynebacterium glutamicum: cloning, molecular analysis and search for a consensus motif. Microbiology 142:1297–1309. Pátek M, Bilic M, Krumbach K, Eikmanns B, Sahm H, and Eggeling L. (1997) Identification and transcriptional analysis of the dapB-ORF2-dapA-ORF4 operon of Corynebacterium glutamicum, encoding two enzymes involved in L-lysine synthesis. Biotechnol. Lett. 19:1113–1117. Pátek M, Hochmannová J, Jelínková M, Nesˇvera J, and Eggeling L. (1998) Analysis of the leuB gene from Corynebacterium glutamicum. Appl. Microbiol. Biotechnol. 50:42–47. Pátek M, Nesˇvera J, Guyonvarch A, Reyes O, and Leblon G. (2003) Promoters of Corynebacterium glutamicum. J. Biotechnol. 104:311–323.

Regulation of Gene Expression [53]

[54]

[55]

[56]

[57]

[58]

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[60]

[61]

[62]

[63]

[64] [65]

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Peoples OP, Liebl W, Bodis M, Maeng PJ, Follettie MT, Archer JA, and Sinskey AJ. (1988) Nucleotide sequence and fine structural analysis of the Corynebacterium glutamicum hom-thrB operon. Mol. Microbiol. 2:63–72. Peters-Wendisch PG, Kreutzer C, Kalinowski J, Pátek M, Sahm H, and Eikmanns BJ. (1998) Pyruvate carboxylase from Corynebacterium glutamicum: characterization, expression and inactivation of the pyc gene. Microbiology 144:915–927. Peters-Wendisch PG, Schiel B, Wendisch VF, Katsoulidis E, Möckel B, Sahm H, and Eikmanns BJ. (2001) Pyruvate carboxylase is a major bottleneck for glutamate and lysine production by Corynebacterium glutamicum. J. Mol. Microbiol. Biotechnol. 3:295–300. Radmacher E, Vaitsikova A, Burger U, Krumbach K, Sahm H, and Eggeling L. (2002) Linking central metabolism with increased pathway flux: L-valine accumulation by Corynebacterium glutamicum. Appl. Environ. Microbiol. 68:2246–2250. Reinscheid DJ, Kronemeyer W, Eggeling L, Eikmanns BJ, and Sahm H. (1994) Stable expression of hom-1-thrB in Corynebacterium glutamicum and its effect on the carbon flux to threonine and related amino acids. Appl. Environ. Microbiol. 60:126–132. Rey DA, Puhler A, and Kalinowski J. (2003) The putative transcriptional repressor McbR, member of the TetR-family, is involved in the regulation of the metabolic network directing the synthesis of sulfur containing amino acids in Corynebacterium glutamicum. J. Biotechnol. 103:51–65. Sano K and Matsui K. (1987) Structure and function of the trp operon control regions of Brevibacterium lactofermentum, a glutamic-acid-producing bacterium. Gene 53:191–200. Schwinde JW, Thum-Schmitz N, Eikmanns BJ, and Sahm H. (1993) Transcriptional analysis of the gap-pgk-tpi-ppc gene cluster of Corynebacterium glutamicum. J. Bacteriol. 175:3905–3908. Schwinde JW, Hertz PF, Sahm H, Eikmanns BJ, and Guyonvarch A. (2001) Lipoamide dehydrogenase from Corynebacterium glutamicum: molecular and physiological analysis of the lpd gene and characterization of the enzyme. Microbiology 147:2223–2231. Simic P, Sahm H, and Eggeling L. (2001) L-Threonine export: use of peptides to identify a new translocator from Corynebacterium glutamicum. J. Bacteriol. 183:5317–5324. Simic P, Willuhn J, Sahm H, and Eggeling L. (2002) Identification of glyA (encoding serine hydroxymethyltransferase) and its use together with the exporter ThrE to increase L-threonine accumulation by Corynebacterium glutamicum. Appl. Environ. Microbiol. 68:3321–3327. Somers JM, Amzallag A, and Middleton RB. (1973) Genetic fine structure of the leucine operon of Escherichia coli K-12. J. Bacteriol. 113:1268–1272. Song T, Dove SL, Lee KH, and Husson RN. (2003) RshA, an anti-sigma factor that regulates the activity of the mycobacterial stress response sigma factor SigH. Mol. Microbiol. 50:949–959. Studholme DJ and Buck M. (2000) The biology of enhancer-dependent transcriptional regulation in bacteria: insights from genome sequences. FEMS Microbiol. Lett. 186:1–9. Tsuchiya M and Morinaga Y. (1988) Genetic control systems of Escherichia coli can confer inducible expression of cloned genes in coryneform bacteria. Bio/Technology 6:428– 430. Vaˇsicová P, Pátek M, Nesˇvera J, Sahm H, and Eikmanns B. (1999) Analysis of the Corynebacterium glutamicum dapA promoter. J. Bacteriol. 181:6188–6191.

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Handbook of Corynebacterium glutamicum Venkova T, Pátek M, and Nesˇvera J. (2001) Identification of a novel gene involved in stable maintenance of plasmid pGA1 from Corynebacterium glutamicum. Plasmid 46:153–162. Venkova-Canova T, Pátek M, and Nesˇvera J. (2003) Control of rep gene expression in the plasmid pGA1 from Corynebacterium glutamicum. J. Bacteriol. 185:2402–2409. von der Osten CH, Barbas CF III, Wong CH, and Sinskey AJ. (1989) Molecular cloning, nucleotide sequence and fine-structural analysis of the Corynebacterium glutamicum fda gene: structural comparison of C. glutamicum fructose-1,6-biphosphate aldolase to class I and class II aldolases. Mol. Microbiol. 3:1625–1637. Wehrmann A, Phillipp B, Sahm H, and Eggeling L. (1998) Different modes of diaminopimelate synthesis and their role in cell wall integrity: a study with Corynebacterium glutamicum. J. Bacteriol. 180:3159–3165. Yanofsky C. (1981) Attenuation in the control of expression of bacterial operons. Nature 289:751–758.

6

Proteomics S. Schaffer and A. Burkovski

CONTENTS 6.1 6.2

Introduction ....................................................................................................99 Two-Dimensional PAGE of Corynebacterium glutamicum ........................100 6.2.1 Analysis of Protein Modifications ...................................................100 6.2.2 Limitations of the Current Methods ................................................109 6.3 N-Terminal Processing of C. glutamicum Proteins.....................................111 6.4 Applications of Proteome Techniques .........................................................113 6.5 Recent Developments and Outlook .............................................................115 References..............................................................................................................116

6.1 INTRODUCTION The proteome — the sum of all proteins present in a cell at a certain time — in contrast to the static genome, is dynamic and is the final result of transcription and translation regulation processes as well as post-translational regulatory mechanisms, such as modification/demodification and proteolysis. Owing to this feature, proteome analysis is a potent tool for monitoring the adaptation processes of cells in response to changing environmental conditions. Important prerequisites for this approach, among others, are the availability of sufficient amounts of homogeneous sample material; high-resolution protein separation techniques that ideally are unbiased with respect to such physicochemical protein properties as isoelectric point or hydrophobicity; detection methods able to cope with the vast dynamic range of proteins in a single cell (106 up to 109-fold for plasma and serum proteins [27]); and fast and reliable means of protein identification. A number of recent reviews detail these challenges and summarize the current methodological state-of-the-art [25,37]. A basic component for the analysis of cellular proteins is two-dimensional polyacrylamide gel electrophoresis (2D PAGE), which separates individual proteins according to their isoelectric point (isoelectric focusing) and their apparent molecular mass (sodium dodecylsulfate–polyacrylamide gel electrophoresis, SDS-PAGE). Owing to its high resolution and sensitivity, up to 10,000 proteins can be resolved in a given sample [16]. After separation by 2D PAGE, distinct spots can be analyzed via microsequencing or by faster and more sensitive mass spectrometry methods like electrospray ionization (ESI) or matrix-assisted laser desorption/ionization– time-of-flight (MALDI-TOF) mass spectrometry (MS).

99

100

Handbook of Corynebacterium glutamicum

6.2 TWO-DIMENSIONAL PAGE OF CORYNEBACTERIUM GLUTAMICUM Various protocols for 2D PAGE of C. glutamicum proteins have been established over the last few years [9–11,31]. In contrast to other proteome projects, for C. glutamicum, a fractionation protocol according to cellular compartments was established from the beginning and submaps of cytoplasmic proteins, membrane fraction proteins, cell wall–associated proteins, and secreted proteins are now available. Although initially protein spots had to be identified by amino-terminal (N-terminal) microsequencing and subsequent comparison with public databases because of the absence of genomic sequence data [9,10], the sequencing of the C. glutamicum genome by several private companies and their providing this information in the public domain [14] made the fast, sensitive, and reliable identification of proteins by MALDI-TOF-MS fingerprint analyses possible [11,31]. Recently, a high-resolution reference map of cytoplasmic and membrane-associated proteins from C. glutamicum cells grown in minimal medium with glucose as carbon source has been published [31]. Table 6.1 lists all C. glutamicum proteins that have been identified on 2D gels in the course of the studies mentioned above with their putative functions and conserved domains indicated. As has been done previously for a number of other bacterial proteomes (for an updated list, see http://us.expasy.org/ch2d/2d-index.html#db) the data were integrated into a public database. The protein table and the corresponding master gel images, showing the protein patterns upon separation of cytosolic, membrane-associated, and secreted proteins are available online at www.fz-juelich.de/ibt/ biochem/biochem.html. The master gel images show about 970, 660, and 40 spots, respectively. Both protein table and images are clickable, allowing easy crossreferencing for a given spot or protein entry. As of December 2004, the database contained entries concerning 197 spots representing 164 proteins.

6.2.1 ANALYSIS

OF

PROTEIN MODIFICATIONS

As is true for eukaryotes, post-translational modification of proteins — most notably glycosylation and phosphorylation — is also found in bacterial systems. Phosphorylation is the most abundant covalent modification of proteins [15] and plays a pivotal role in the modulation of protein activity, especially of proteins acting in signal transduction processes. Phosphorus-containing C. glutamicum proteins were identified by two approaches: by immunostaining with phosphoamino acid–specific monoclonal antibodies and by in vivo radiolabeling using [33P]-phosphoric acid and subsequent autoradiography [1]. Whereas the first method is specific for phosphorylated amino acid residues, protein spots detected with the second method may also include proteins modified by other phosphorus-containing groups, such as adenylyl and uridylyl residues. After 2D PAGE, approximately 90 immunostained protein spots and approximately 60 [33P]-labeled protein spots were detected. Thirty-one of these protein spots were detected with both methods. By peptide mass fingerprinting, 41 proteins were identified and a first phosphoproteome map was established based on the combined results of both methods [1]. To our knowledge, this is the first

Proteomics

101

TABLE 6.1 Corynebacterium glutamicum Proteins Identified on 2D Gelsa No.b I

NCgl

Functionc

Small molecule metabolism A Energy metabolism 1 Glycolysis 5 1525 Phosphoglycerate kinase Pgk 28 0935 Enolase Eno 65 1860 Phosphofructokinase Pfk 71 1524 Triosephosphate isomerase Tpi 87 0390 Phosphoglycerate mutase Gpm 110 2673 Fructose-bisphosphate aldolase Fda 116 2673 Fructose-bisphosphate aldolase Fda 132 1526 Glyceraldehyde-3-phosphate dehydrogenase Gap

15 17 58 94 124

101

34 103 107 166 139 164 169 M05 M06 M07 M08 M09

2 Pentose phosphate cycle 1516 Glucose-6-phosphate dehydrogenase DevB 1536 Ribulose-phosphate 3-epimerase Rpe 1512 Transketolase Tkt 1512 Transketolase Tkt (338-695) 2337 Ribose 5-phosphate isomerase Rpi 3 Pyruvate dehydrogenase 2167 Pyruvate dehydrogenase subunit E1 AceE (647–924) 4 TCA cycle and glyoxylate shunt 0634 Isocitrate dehydrogenase Icd 0967 Fumarase FumC 0967 Fumarase FumC 0967 Fumarase FumC 1926 Malate:quinone oxidoreductase Mqo 2247 Malate synthase AceB 1482 Aconitase Acn 0360 Succinate dehydrogenase subunit A SdhA 0360 Succinate dehydrogenase subunit A SdhA 0360 Succinate dehydrogenase subunit A SdhA 0360 Succinate dehydrogenase subunit A SdhA 0360 Succinate dehydrogenase subunit A SdhA

Ed

Pe

Lf

Ag

PGK Enolase PfkB TIM PGAM FbP aldolase FbP aldolase Gpdh

D B B D B B B D

ND +/+ ND ND ND ND +/+ +/+

C C C C C C C C

1.43 2.76 0.05 3.79 3.08 1.09 6.46 3.34

Glucosamin iso

D

ND

C

0.32

Ribul P3 epim Transketolase Transketolase LacAB rpiB

B D D B

ND +/+ ND ND

C C C C

1.08 1.13 0.07 2.46

Transketolase

D

ND

C

0.18

IDH Lyase 1 Lyase 1 Lyase 1 — Malatesynthase Aconitase Succ DH flav C

D B B B D D B B

+/+ ND +/? ND ND ND +/+ ND

C C C C C C C M

0.89 0.44 1.98 0.47 0.42 0.07 0.21 ND

Succ DH flav C

B

ND

M

ND

Succ DH flav C

B

ND

M

ND

Succ DH flav C

B

ND

M

ND

Succ DH flav C

B

ND

M

ND

PFAM

102

Handbook of Corynebacterium glutamicum

TABLE 6.1 (continued) Corynebacterium glutamicum Proteins Identified on 2D Gelsa No.b

NCgl

M19

0360

M20

0360

M21

0360

M13

0361

M14

0361

8 20 48 60 153

157 M02

Functionc Succinate dehydrogenase SdhA (450–625) Succinate dehydrogenase SdhA (450-625) Succinate dehydrogenase SdhA (450–625) Succinate dehydrogenase SdhB Succinate dehydrogenase SdhB

PFAM

Ed

Pe

Lf

Ag

subunit A

Succ DH flav C

B

ND

M

ND

subunit A

Succ DH flav C

B

ND

M

ND

subunit A

Succ DH flav C

B

ND

M

ND

subunit B

Fer4

B

ND

M

ND

subunit B

Fer4

B

ND

M

ND

ETF alpha

B

ND

C

0.39

ETF alpha

B

ND

C

0.26

ETF beta

B

ND

C

0.58

5 Respiration 1183 α Subunit of electron-transferring flavoprotein EtfA 1182 β Subunit of electron-transferring flavoprotein EtfB 1182 β Subunit of electron-transferring flavoprotein EtfB 2719 Ferredoxin-NADP(+) oxidoreductase Fnr 0865 D-Lactate dehydrogenase Dld

Pyr redox

B

ND

C

0.93

FAD binding 4

B

ND

C

0.03

6 ATP synthase 1163 α Subunit of F0F1-ATPase AtpA 1163 α Subunit of F0F1-ATPase AtpA

ATP-synt ab ATP-synt ab

D D

+/− ND

C C

0.20 ND

PEPCK PEPCK FBPase glpX FBPase glpX

D D D D

ND ND ND ND

C C C C

0.04 0.36 1.24 0.05

PGM PMM PGM PMM dTDP sugar

B B B

ND ND ND

C C C

0.02 0.52 0.17

Alpha amylase

B

ND

C

0.63

Epimerase

B

ND

C

0.50

B Central intermediary metabolism 1 Gluconeogenesis 32 2765 PEP carboxykinase PckA 33 2765 PEP carboxykinase PckA 109 0976 Fructose-1,6-bisphosphatase Fbp 135 0976 Fructose-1,6-bisphosphatase Fbp 2 Sugars 0714 Phosphomannomutase PmmA 2453 Phosphoglucomutase Pgm 0326 dTDP-4-keto-6-deoxyglucose-3,5epimerase DexB 105 2045 Maltooligosyltrehalose trehalohydrolase GlgZ 117 0317 dTDP-glucose-4,6-dehydratase 4 55 95

Proteomics

103

TABLE 6.1 (continued) Corynebacterium glutamicum Proteins Identified on 2D Gelsa No.b

NCgl

Functionc

C Amino acid biosynthesis 1 Glutamate family 45 1341 Glutamate N-acetyltransferase ArgJ (60–146) 63 1344 Ornithine carbamoyltransferase ArgF 93 1346 Argininosuccinate synthetase ArgG 111 1340 N-acetyl-γ-glutamyl-phosphate reductase ArgC 168 1999 Glutamate dehydrogenase (NADP+dependent) Gdh 2 Aspartate family 1896 Dihydrodipicolinate synthase DapA 1896 Dihydrodipicolinate synthase DapA 0625 O-Acetylhomoserine sulfhydrylase MetY 145 2528 D,L-Diaminopimelate dehydrogenase Ddh 152 1446 Aspartate ammonia-lyase AspA 156 2048 Methionine synthase MetE 89 90 138

26 66 73 82 92 144

3 Serine 0794 2473 2473 2473 2473 0954

family Phosphoserine aminotransferase SerC Cysteine synthase CysK Cysteine synthase CysK (48–299) Cysteine synthase CysK (35–153) Cysteine synthase CysK Serine hydroxymethyltransferase GlyA

4 Aromatic amino acid family 0730 5-Enolpyruvylshikimate-3-phosphate synthase AroA 99 2098 2-Dehydro-3-deoxyphosphoheptonate synthase AroF 130 1087 3-Dehydroquinate dehydratase AroD 133 1559 3-Dehydroquinate synthase AroB 61

6 29 114 117 128

5 Histidine family 0215 Histidinol phosphate aminotransferase HisC 2021 Histidinol dehydrogenase HisD 2013 Cyclase HisF 1447 ATP phosphoribosyltransferase HisG 2016 Amidotransferase HisH

Ed

Pe

Lf

Ag

ArgJ

D

ND

C

1.92

OTCace Arginosu synth Semialdhyd dh

D D D

ND ND ND

C C C

0.68 0.12 0.58

GLFV dehydrog

D

ND

C

0.12

DHDPS DHDPS —

D D D

ND ND +/−

C C C

0.19 0.20 0.02

D

ND

C

1.55

Lyase 1 —

B D

ND ND

C C

0.17 0.52

— — — — — SHMT

D B B B B D

ND +/? ND ND ND +/+

C C C C C C

1.10 2.41 1.61 0.61 1.10 1.13

B

+/+

C

1.73

DAHP synth2

B

ND

C

0.26

Shikimate DH DHQ synthase

B B

ND ND

C C

0.75 0.05

Aminotran 1 2

B

ND

C

0.03

Histidinol DH His biosynth HisG GATase

B B D B

ND ND ND ND

C C C C

0.29 0.51 0.50 0.45

PFAM

—

—

104

Handbook of Corynebacterium glutamicum

TABLE 6.1 (continued) Corynebacterium glutamicum Proteins Identified on 2D Gelsa No.b

NCgl

Functionc

6 Branched chain amino acid family 1237 3-Isopropylmalate dehydrogenase LeuB 102 1262 3-Isopropylmalate isomerase subunit LeuC 106 1262 3-Isopropylmalate isomerase subunit LeuC 67

Ed

Pe

Lf

Ag

Isodh

D

ND

C

0.31

Aconitase

B

ND

C

0.87

Aconitase

B

ND

C

0.48

B B B B

ND ND ND ND

C C C C

0.86 0.55 0.67 0.22

B

ND

C

0.09

B

ND

C

0.23

B B

ND ND

C C

0.35 1.31

PFAM

D Purines, pyrimidines, nucleosides and nucleotides 1 Purines 96 0579 IMP dehydrogenase GuaB — 100 0582 GMP synthetase GuaA GMP synth 141 0827 IMP cyclohydrolase PurH — 149 0905 Ribose-phosphate pyrophosphokinase Pribosyltran PrsA 151 0371 Formyltetrahydrofolate deformylase Formyl transf PurU 155 2508 SAICAR synthase PurC SAICAR synth

84 118

44 72 78

9 136

2 Pyrimidines 2025 Uridylate kinase PyrH 1948 Uridylate kinase PyrH

— —

3 Nucleotides 0533 Adenylate kinase Adk 0720 Thymidylate kinase Tmk 2287 Nucleoside diphosphate kinase Ndk

ADK Thymidylate kin NDK

B B B

ND ND ND

C C C

2.45 0.47 8.51

4 Salvage of nucleotides, miscellaneous 0654 Uracil phosphoribosyltransferase Upp 0075 Cytosine deaminase CodA

Pribosyltran —

B B

ND ND

C C

0.47 0.05

B

ND

C

0.56

B B D B D B

ND ND ND ND ND ND

C C C C C C

0.85 3.83 1.23 1.58 0.24 0.19

B

ND

C

0.91

E Biosynthesis of prosthetic groups, cofactors and carriers 12 0888 2-Demethylmenaquinone — methyltransferase MenG 23 2984 Thioredoxin reductase TrxB Pyr redox 76 2985 Thioredoxin Txr (55–113) Thiored Aldo ket red 88 2277 2,5-diketo-D-gluconic acid reductase 97 0754 Pyridoxine biosynthesis protein SOR SNZ 113 0112 Pantoate-β-alanine ligase PanC Pantoate ligase 147 1023 Nicotinate-nucleotide QRPTase pyrophosphorylase NadC 150 0620 Methylene tetrahydrofolate THF-DHG-CYH dehydrogenase FolD

Proteomics

105

TABLE 6.1 (continued) Corynebacterium glutamicum Proteins Identified on 2D Gelsa No.b

NCgl

Functionc

F Degradation 30 0049 Succinate semialdehyde dehydrogenase GabD 36 2607 Inorganic pyrophosphatase Ppa 49 2919 2-Hydroxyhepta-2,4-diene-1,7dioateisomerase II Macromolecule metabolism A Macromolecule synthesis 1 Aminoacyl-tRNA synthetases 167 1335 Phenylanalyl-tRNA synthetase PheS, α-subunit

37

59 83 73 85 98 104 91 119 121

53

Ed

Pe

Lf

Ag

Aldedh

B

ND

C

0.32

Pyropphatase FAA hydrolase

B B

+/− ND

C C

3.20 0.87

B

ND

C

9.77

PFAM

—

2 Ribosomal proteins 0469 50S Ribosomal protein L12 RplL

Ribosomal L12

B

ND

C

9.77

3 Translation 0478 Elongation factor G Efg 0478 Elongation factor G Efg (32–385) 0480 Elongation factor Tu Tuf (253–336) 0480 Elongation factor Tu Tuf (236–336) 0480 Elongation factor Tu Tuf 0480 Elongation factor Tu Tuf 1949 Elongation factor Ts Tsf 1557 Elongation factor P Efp 1947 Ribosome recycling factor Frr

— — GTP EFTU GTP EFTU GTP EFTU GTP EFTU EF Ts EFP RRF

B B B B B B B B B

+/+ ND ND ND +/+ ND +/− +/+ ND

C C C C C C C C C

0.30 1.40 1.61 1.95 5.57 0.83 2.84 1.72 2.43

4 DNA replication 0002 βˆ Subunit of DNA polymerase III DnaN

Pol3Bc

B

ND

C

0.36

Esterase Esterase Esterase EPSP synthase

D D D B

ND ND ND ND

C S S C

0.10 ND ND 0.29

D

ND

M

ND

Esterase

B

ND

M

ND

CLP protease

B D

ND ND

C C

0.54 1.58

5 Cell wall synthesis 2777 Trehalose mycoloyltransferase PS1 2777 Trehalose mycoloyltransferase PS1 2777 Trehalose mycoloyltransferase PS1 0345 UDP-N-acetylglucosamine 1transferase MurA M10 0184 Arabinosyltransferase EmbB (867–1146) M18 2779 Surface layer protein 31 S05 S06 134

B Degradation of macromolecules 1 Degradation of proteins and peptides 2 2631 Peptidase 46 2327 ATP-dependent protease Clp subunit ClpP2

—

106

Handbook of Corynebacterium glutamicum

TABLE 6.1 (continued) Corynebacterium glutamicum Proteins Identified on 2D Gelsa No.b

NCgl

62 74

1558 2328

134 138 158 159

0566 1442 0333 1430

112 161

Functionc Cytoplasmic peptidase PepQ ATP-dependent protease Clp subunit ClpP1 Proline iminopeptidase Pip Aspartyl aminopeptidase PepC Prolyl oligopeptidase X-Pro dipeptidase PepQ

2 Degradation of nucleic acids 0641 Exodeoxyribonuclease ExoA 2645 Exodeoxyribonuclease III XthA

III Cellular processes A Transport 1 2375 Maltose-binding protein AmyE S02 2375 Maltose-binding protein AmyE S11 2375 Maltose-binding protein AmyE 69 1858 Enzyme I of the PEP-PTS PtsI (48–252) 86 1501 ATP-binding subunit of ATP transporter 137 1502 ABC transporter ATP-binding protein M01 2375 Maltose-binding protein AmyE M03 2732 ATP-binding subunit of ATP transporter M11 1875 ATP-binding subunit of glutamate transporter GluB M12 1276 ATP-binding subunit of glutamine transporter GlnQ M15 1577 ABC transporter M16 1305 Mannose-specific PTS enzyme II PtsM (585–683) M17 1305 Mannose-specific PTS enzyme II PtsM (526–683) B Chaperones 3 2702 DnaK 52 2702 DnaK (35–337) 54 2702 DnaK 42 0572 GroES 56 2621 GroEL2 57 0573 GroEL 135 2210 DnaJ2 165 2682 ClpB

Ed

Pe

Lf

Ag

Peptidase M24 CLP protease

B D

ND ND

C C

0.60 2.75

-

B B B B

ND ND ND ND

C C C C

0.29 0.05 0.05 0.07

Exo endo phos

B B

ND ND

C C

0.45 0.39

SBP bac 1 SBP bac 1 SBP bac 1 —

B B B D

ND ND ND ND

C S S C

1.50 ND ND 0.52

AAA

B

ND

C

1.66

UPF0051 — —

B B B

ND ND ND

C C C

0.52 ND ND

—

D

ND

C

ND

—

B

ND

C

ND

— PTS EIIABC

B D

ND ND

C C

ND ND

PTS EIIABC

D

ND

C

ND

HSP70 HSP70 HSP70 — — — DnaJ Clp/AAA/UBA

B B B B B B B B

+/− ND ND ND +/− ND +/− ND

C C C C C C C C

1.56 0.57 0.32 9.02 0.82 0.91 0.05 0.31

PFAM

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TABLE 6.1 (continued) Corynebacterium glutamicum Proteins Identified on 2D Gelsa No.b

NCgl

Functionc

C Detoxification 10 1041 Thioredoxin peroxidase Tpx 64 0313 Formaldehyde dehydrogenase FadH 120 2826 Superoxide dismutase SodA 140 0251 Catalase KatA IV Miscellaneous 7 2449 Sugar phosphatase 11 1316 Universal stress protein 13 1316 Universal stress protein 14 1316 Universal stress protein 19 2339 Oxidoreductase 21 2582 L-2,3-Butanediol dehydrogenase BudC 22 1318 Nucleoside-diphosphate-sugar epimerase 38 2897 DNA protection during starvation protein PexB 40 2659 Acetyltransferase 47 0314 Hydrolase 51 2894 myo-Inositol-1-phosphate synthase 50 1510 Quinone oxidoreductase Qor 69 0285 Hydrolase 70 2530 Hydrolase 80 2365 Esterase 81 2136 bis(5′-Nucleosyl)-tetraphosphatase 102 2698 Aldehyde dehydrogenase AldA 108 2480 Succinyl-CoA:CoA transferase Cat1 131 2709 Alcohol dehydrogenase Adh 143 1500 Cysteine desulfhydrase/selenocysteine lyase 146 2449 Oxidoreductase 148 2358 Oxidoreductase 152 0187 Oxidoreductase 154 2487 Acetyltransferase S12 0872 Resuscitation factor V Proteins with unknown function 16 1466 Unknown 18 1485 Unknown 24 1996 Unknown 25 0148 Unknown 27 2349 Unknown 35 1385 Unknown 125 1385 Unknown (70-87) 39 1123 Unknown

PFAM

Ed

Pe

Lf

Ag

AhpC-TSA Adh-zinc Sodfe Catalase

B B B D

ND ND ND ND

C C C C

5.11 1.21 2.41 0.72

HAD Usp Usp Usp DSBA Adh short —

B B B B B B B

+/− ND ND ND ND ND ND

C C C C C C C

0.26 4.48 0.28 1.11 0.35 2.57 0.18

Ferritin

B

ND

C

0.83

Acetyltransf Lactamase B Inos-1-P synth Adh zinc Lactamase B Lactamase B 4HBT NUDIX Aldedh AcetylCoA hyd Adh zinc Aminotran_5

B B B B B B B B B D B B

ND ND ND ND ND ND ND ND ND ND ND ND

C C C C C C C C C C C C

0.66 0.38 0.86 0.33 0.52 0.50 1.67 0.10 0.87 1.76 0.30 1.78

ADH zinc N Adh short HAD Acetyltransf —

B B B B B

ND ND ND ND ND

C C C C S

1.15 0.39 0.17 0.10 ND

PBP — — — — FHA FHA YceI

— — — — — — — —

ND ND ND ND ND ND ND ND

C C C C C C C C

0.97 0.87 1.03 0.47 0.18 1.61 0.10 1.98

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TABLE 6.1 (continued) Corynebacterium glutamicum Proteins Identified on 2D Gelsa No.b

NCgl

41 77 43 68 75 79 115 122 123 126 127 129 142 160 162 163 M04 M22 S01 S04 S07 S08 S09 S11

0738 0738 2512 0948 2531 2501 2806 0673 0673 1170 1155 2073 0627 1829 2490 1599 2070 0651 1480 1289 1289 1289 0535 2664

a

Functionc Unknown Unknown Unknown Unknown Unknown Unknown (12–38) Unknown Unknown Unknown Unknown Unknown Unknown Unknown Unknown Unknown Unknown Unknown Unknown Unknown Unknown Unknown Unknown Unknown Unknown

PFAM

Ed

Pe

Lf

Ag

— — HIT DUF158 — PurC — — — Glyoxalase Sua5 yciO yrdC — UPF0001 — — DUF28 DivIVA — NLPC-P60 — — — ErfKYbiS YnhG —

— — — — — — — — — — — — — — — — — — — — — — — —

ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND

C C C C C C C C C C C C C C C C M M S S S S S S

3.19 0.63 1.70 0.26 1.10 4.13 0.10 1.59 1.46 0.62 0.37 0.28 0.12 0.31 0.82 0.03 ND ND ND ND ND ND ND ND

The spot numbers (No.) refer to protein spots on 2D images of the soluble, membrane-associated (spot number preceded by M) and secreted (spot number preceded by S) protein fractions available at www.fzjuelich.de/ibt/biochem/biochem.html. The GenBank accession number (NCgl) is followed by the putative function of the protein and the PFAM domains present in the protein. Moreover, the kind of evidence used for function assignment, available evidence for protein phosphorylation [1], and subcellular localization as well as relative abundance of the protein are given in the columns labeled E, P, L, and A, respectively. b Spot numbers in italics indicate that the protein is found in more than one spot; underlined spot numbers indicate that the spot represents more than one protein. c In some cases, a protein’s molecular weight is predicted to be considerably higher than observed, indicating that the spot represents only a fragment of the protein. Here, numbers in parentheses indicate the amino acid residues that are covered by peptides detected during mass spectrometry. d Function assignment is based on experimental evidence (D) or bioinformatics analysis (B). e Proteins were detected upon metabolic labeling with [33P]-phosphoric acid, two-dimensional electrophoresis of proteins, and subsequent autoradiography and/or upon two-dimensional electrophoresis of proteins and subsequent Western blotting with monoclonal anti-phosphoserine antibodies [1]. +: detected; -: not detected; ?: detection uncertain; ND: not determined. f Assignment of subcellular localization is based on detection of the proteins upon two-dimensional electrophoresis of soluble and membrane-associated protein fractions [31] as well as of proteins secreted into the culture medium [11]. C: cytosolic; M: membrane-associated; S: secreted. g Relative abundance of cytoplasmic proteins was calculated as described in [31]. ND: not determined.

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comprehensive study of protein phosphorylation in bacteria. The results show that this type of covalent modification is more widespread in bacterial proteins than previously thought.

6.2.2 LIMITATIONS

OF THE

CURRENT METHODS

A graphical representation of the C. glutamicum proteome calculated from the genomic DNA sequence shows a bimodal isoelectric distribution (Figure 6.1A) similar to the theoretical proteome map of other bacteria [12,20,33]. Of these proteins, 741 (23.9%) exhibit pI values higher than 7, and there are no proteins with calculated pI values between 7.27 and 7.58. When the actual 2D protein pattern obtained for C. glutamicum is compared with this calculated map, it is characterized by an almost complete loss of basic proteins (Figure 6.1B). This in accordance with other studies on bacterial proteomes, where only few basic proteins apart from the

A

FIGURE 6.1 Comparison of theoretical and experimental two-dimensional protein pattern of C. glutamicum. Panel A shows the conceptual C. glutamicum proteome as predicted by calculation of pI and molecular mass for 3,103 protein entries at http://www.ebi.ac.uk/proteome/index.html. The proteins in the large and small rectangles are theoretically amenable to 2D analysis when using Immobiline DryStrips of pH 3 to 10 or 4 to 7, respectively, for isoelectric focusing and Excel SDS gradient gels of 12 to 14% for SDS-PAGE. Panel B shows the 2D image of C. glutamicum proteins upon separation of 300 μg of protein using Immobiline DryStrips, pH 3 to 10, for isoelectric focusing and Excel SDS gradient gels 12 to 14% for SDS-PAGE. The gels were subsequently stained with colloidal Coomassie G-250.

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pl 3

7

10

Molecular mass (KDa)

180 130 100 70 55 45 35 25 15

10 B

FIGURE 6.1 (continued).

highly abundant ribosomal proteins could be detected [4,20,24,33]. Several technical reasons account for this, including active water transport toward the anode of immobilized pH gradient (IPG) gels (reverse electroendosmosis) caused by the strong positive charge of basic acrylamido buffers, as well as the hydrolysis of acrylamide to acrylic acid at alkaline pH and the migration of reducing agents such as dithiothreitol. Moreover, there is a positive correlation between high isoelectric point and hydrophobicity (see below). Another group of proteins significantly underrepresented in the current protein maps are membrane proteins. Although one-third of all genes in a typical bacterial genome encode integral membrane proteins (see Chapter 8), they are vastly underrepresented on 2D gels, when unfractionated protein extracts are analyzed. The membrane-associated protein fraction represents only about 5 to 10% of the total protein content of a C. glutamicum cell (Schaffer, unpublished results). Therefore, preparation of washed membrane fractions prior to 2D separation should allow for a significantly better representation of membrane-associated proteins on 2D gels. In fact, all proteins identified on such gels were predicted to be membrane associated. However, membrane proteins with more than one transmembrane helix or with grand-average-of-hydropathy (GRAVY) values higher than 0.013 could not be identified up until now with the exception of two proteins, EmbB and PtsM. However, the corresponding proteins spots exhibit pI values and molecular masses differing considerably from those calculated. Moreover, MALDI-TOF analysis of tryptic digests of these spots revealed that matching peptides all mapped to hydrophilic regions of these proteins. Consequently, the spots represent EmbB and PtsM fragments that either arose by in vivo proteolytic processing or by degradation during sample preparation [31].

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As already mentioned, the difference in abundance of individual proteins within a given cell spans at least six orders of magnitude and at least nine orders of magnitude in the case of human serum or plasma proteins [27]. In consequence, low-abundance proteins are underrepresented in the current proteome maps, as is indicated by the complete failure to identify regulatory proteins, such as transcriptional regulators or protein kinases. The use of narrow-IPG strips for isoelectric focusing and of colloidal Coomassie for protein detection proteins allowed the display of proteins with differences in abundance of only about two orders of magnitude [31]. The problem might be partially solved by using more sensitive detection methods, because the transcriptional regulator ClgR could be visualized using a silver-staining protocol (Engels, unpublished results). However, this protein became visible only upon long exposure to the development reagent, leading to the complete loss of resolution in those areas of the gel with higher spot densities. As fluorescent dyes are reported to allow protein detection with sensitivity comparable to silver staining and a dynamic range of up to five orders of magnitude [26], they might be the detection tools of choice in future. First experiments using these dyes for the analysis of C. glutamicum proteins have already been carried out [8].

6.3 N-TERMINAL PROCESSING OF C. GLUTAMICUM PROTEINS A number of C. glutamicum proteins were analyzed by N-terminal microsequencing [5,9–11,19,21,28,29] and additionally the identity of N-terminal peptides was determined by MALDI-TOF-MS-based post-source decay (PSD) analysis [31]. When these sequences were used for database searches at the European Bioinformatics Institute (www.ebi.ac.uk), 33 proteins could be identified (summarized in Table 6.2). An astonishing number of six database entries (18%) suggested translation initiation at start codons different from those determined experimentally, indicating incorrect annotations or alternative translational start sites. Twenty-three proteins showed methionine aminopeptidase-dependent processing of their N-termini (70%). The N-terminal methionine was always removed when the following amino acid residue was serine (four cases), glycine (one), glutamine (one), leucine (one), or proline (one) and cleaved off in most cases when it was followed by alanine (eight out of nine proteins) or threonine (six out of seven). Other amino acid residues revealed a different behavior. In the case of lysine following the initiator methionine, one protein was processed and one N-terminus was unprocessed, whereas arginine (two cases), histidine (one) aspartate (one), and glutamate residues (one) never promoted processing of the N-terminal methionine. These data indicate that N-terminal processing in C. glutamicum follows rules similar to those for E. coli, in which the N-terminal methionine is always cleaved when the penultimate amino acid residue is either serine or alanine. Occurrence of cleavage is variable if either threonine, glycine, or proline is the penultimate residue [20]. In contrast to E. coli, methionine aminopeptidase-dependent processing appears to occur in C. glutamicum also in proteins with glutamine, lysine, and leucine following the initiator methionine.

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TABLE 6.2 List of N-Terminal Sequences Determined for Protein Spots Predicted and Observed N-terminal Sequence MGHV VGISLDVVMMGVMTSKTATAILHTNR-GDITID TSKTATAILHTNRRGDITID MKLTLEIWRQAGPTAEGKF MKLTLEIXRQAGPTAEGKF MAKLTKDELIEAFKEMTLIELSEFV AKLTKDELIEAFKEMTLIELSEFV MAQEVLKDLNKVRNIGIMAH KQEVLKDLNKVRNIGIMAT MRDHVEIGIGR MRDHVEIGIGR MSFHITSVN MTSMSFHITSVN MATVTFDKVTIRYPGAERATVHE TVFFDKVTIRYPLAERATVXE MQVTESGSTASPLCGVGSSVM TETQETYQATTRVKRGLADMLKGGVIMDVV TETQETYQATTRVLRGLADMLKGGVIMDVV MAEIMHVFAREILDSRGNPT AEIMHVFAREILDSRGNPT MTEQEFRIEHD QEQEFRIEHD MGSMAKTHFQGNETATSGELPQVG AKTHFQGNETATSGELLQVG MHAASREALAKVSSDLDAALAADN MHAASREALAKVSSDLDAALAADN MLE ETTESRKNMAELTISSDEIRSAIANYTSSY AELTISSDEIRSAIANYTSXY MATIRELRDRIRSVNS XTIRELRDRIRSVNT MTSPVENSTSTEK MTSPVENSTSTEK MSDNNGTPEPQVETTSV M-DNNGTPETQVETTLV MSVNPTRPEGGR SVNPTRPEGGR MAVKTLKDLLDEG AVKTLKDLLDEG MTIRVGINGFGRIGRNFF TIRVGINGFGRIGRNFF MANPFSKAWKYLMALFDSKIE ANPFSKAXKYLMALFDSKIE

Protein (Gene Name) Peptidyl-prolyl cis-trans isomerase

NCgl Spot No. 0033

Succinate dehydrogenase/ 0361 M13 fumarate reductase Fe-S protein M14 50S ribosomal protein L7/L12 0469 37 (rplL) Elongation factor G (fusA) 0478 59, 83 IMP dehydrogenase/GMP reductase Exonuclease III

0579 96 0641 112

ABC-type transporter, ATPase 0698 component Pyridoxine biosynthesis enzyme 0754 97

Enolase (eno)

0935 28

Fumarase (fum) Peroxiredoxin

0967 103, 107, 166 1041 10

H+-ATPase δ-subunit (atpH)

1162

H+-ATPase α-subunit (atpA)

1163 157, M02

H+-ATPase γ-subunit (atpG)

1164

3-isopropylmalate dehydratase large subunit (leuC) FHA-domain-containing protein NADH dehydrogenase, FADcontaining subunit (ndh) 3-Phosphoglycerate kinase (pgk) Glyceraldehyde 3-phosphate dehydrogenase (gap) Phage shock protein A (IM30)

1262 102, 106 1385 35, 125 1409 1525 5 1526 132 1886

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TABLE 6.2 (continued) List of N-Terminal Sequences Determined for Protein Spots Predicted and Observed N-terminal Sequence MANYTAADVKKLRELTGSGMLDCKK ANYTAADVKKLRELTGAGMLDCKK MPLTPADVHNVAFNKP LTPADVLNVAFNKP MTEQELLSAQT TEQELLSAQT MSNVGKPRTAQ SNVGKPRTAQ MTERTLILIKPDGVTNGHV TERTLILIKPDGVTNGAV MRVYLGADHAGFETK MRVYLGADHAGPETK MAANNVNDDTQNNLHVTEVDLR MAANNVND-TQNNLHVTEVDLR MIGAPPDMGNVYNNITETIGHTPLVKLNKL GNVYNNITETIGHTPLVKLNKL MSDRIASEKLRSKLM SDYIASEKLYGKLM MSKVAMVTGGAQGIGRG XKVAMVTGGAXGIRMG MPIATPEVYNEMLDRAKEGGF PIATPEVYNEMLDRAKEGGF MENVYEFLGNLDVL MENVYEFLGNLDVL SGSGLIGYVFDFLGASSKWAGAVADLIGLL XGSGLIGYVFDFLGASSKWAGAVADLIGLL MANPLSKGWKYLMA ANPLSKGITYLXA

Protein (Gene Name)

NCgl Spot No.

Elongation factor Ts (tsf)

1949 91

Cell division initiation protein, Antigen 84 Malate synthase G (aceB)

2070 M04 2247 164

Isocitrate lyase (aceA)

2248

Nucleoside diphosphate kinase (ndk) Ribose 5-phosphate isomerase (rpi) Predicted thioesterase

2287 78

Cysteine synthase (cysK) Acetyl-CoA hydrolase L-2,3-butanediol

2337 124 2365 80 2473 66, 73, 82, 92 2480 108

dehydrogenase 2582 21

Fructose-bisphosphate aldolase (fda) Porin (porA)

2673 110, 116

Hypothetical protein

2848

2715

a

For protein identification, experimentally determined N-terminal sequences [5,9–11,19,21,28,29,31] were compared with publicly available sequence data using the fasta3 program at the European Bioinformatics Institute (www.ebi.ac.uk). The single-letter code is used to indicate the different amino acids (X = unknown amino acid). Experimentally determined N-terminal sequences are given in bold with residues deviating from the prediction underlined. Accession numbers (NCgl) correspond to those in the NCBI nonredundant protein database, spot numbers to those in Table 6.1 and in clickable gel images available at www.fzjuelich.de/ibt/biochem/biochem.html.

6.4 APPLICATIONS OF PROTEOME TECHNIQUES As mentioned in the introductory section of this chapter, proteome analyses are especially suitable for the comparison of a cell’s protein pattern under different physiological conditions. The first studies in this respect were carried out to investigate the effect of nitrogen limitation [23,32]. Nolden and co-workers [23] found

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four proteins, the synthesis rate of which varies with the nitrogen status of the cell. Although synthesis of the ribosomal protein L12 ceases in response to nitrogen starvation, formation of an unknown protein, thioredoxin, and the dapD gene product, a component of the split diaminopimelate pathway, increase. Additionally, [35S]methionine in vivo labeling followed by 2D PAGE and autoradiography revealed first indications for cross-talk between nitrogen control and energy metabolism [32]. Because phosphorylation cascades are often involved in the regulation of global regulatory networks, the effect of nitrogen limitation on the phosphorylation pattern of C. glutamicum proteins was investigated. Differences between cultures grown with high nitrogen supply and nitrogen-starved cultures were not observed in this study, either by [33P]-phosphoric acid in vivo labeling and autoradiography or by Western blotting analyses with phosphoamino acid–specific monoclonal antibodies [1]. This result was unexpected because at least the adenylylation or uridylylation of proteins, depending on the cellular nitrogen supply, has already been shown (see chapter on nitrogen control in this book), and these modifications should be detectable after [33P]-phosphoric acid in vivo labeling and autoradiography. A possible explanation of the negative result might be the low abundance of phosphoruscontaining regulatory components [1]. The investigation of propionate metabolism provides another example of the application of proteomics techniques on C. glutamicum [2]. In a recent study, 2D PAGE combined with mass spectrometry revealed a strong induction of the prpD2B2C2 gene products 2-methylcitrate dehydratase, 2-methylisocitrate lyase, and 2-methylcitrate synthase, when propionate was added as an additional carbon source to acetate-grown cells. Genetic studies confirmed that the prpD2B2C2 operon is indeed required by C. glutamicum in order to grow on propionate as the sole carbon source [2]. The PrpD2 protein was also shown to be eightfold more abundant in the C. glutamicum wild-type when grown in the presence of 300 mM L-valine [18]. Other proteins more abundant under these conditions are the arginine repressor (ArgR; fivefold) and N-acetylglutamate semialdehyde dehydrogenase (ArgC; fourfold). In all cases, the increased abundance of the proteins correlates with increased mRNA levels of the respective genes (sixfold, twofold, and twofold, respectively). In contrast, the mRNA levels of structural genes coding for 11 proteins with decreased abundance in the presence of L-valine were shown to be unaffected by L-valine presence [18]. However, the physiological significance of the observed L-valine-induced changes in the protein expression profile is unclear. [35S]-Methionine in vivo labeling, in combination with 2D PAGE and peptide mass fingerprint analysis, was also used to study differences in the protein profile of glucose- and acetate-grown C. glutamicum cells [6]. Of about 500 protein spots detectable upon the analysis of cell lysates from glucose- and acetate-grown cultures after autoradiography, 54 were present in higher amounts and 26 in lower amounts in the lysate of acetate-grown cells. Ten acetate-induced proteins are identified, namely, isocitrate lyase, malate synthase, citrate synthase, fumarase, malate:chinone oxidoreductase, cysteine synthase, glycine-tRNA ligase, butyryl-CoA transferase, a putative ABC transporter, and an aminotransferase. Increased synthesis of the first four proteins is in agreement with data obtained by transcriptome analyses and also

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in accordance with higher activities of TCA and glyoxylate cycle enzymes in acetategrown cells. The physiological connection of the other proteins to acetate metabolism is not clear [6]. Engels et al. [3] made use of 2D PAGE and MALDI-TOF mass spectrometry in order to characterize C. glutamicum mutants with deletions in the clpC and clpX genes, coding for regulatory subunits of the ATP-dependent protease Clp. Deletion of clpC led to the drastically increased abundance of two proteins, ClpP1 and ClpP2, representing the proteolytic subunits of the Clp protease, accompanied by significantly higher clpP1 and clpP2 mRNA levels. Subsequent studies led to the identification of the transcriptional activator, ClgR, which controls clpC and clpP1P2 gene expression in C. glutamicum. Using 2D PAGE, it could further be demonstrated that the M. tuberculosis ortholog of ClgR is able to functionally replace C. glutamicum ClgR, supporting the hypothesis that the identified autoregulatory loop controlling clp gene expression and involving ClgR, as well as the ClpCP protease, is conserved in the order Actinomycetales. This had been suggested initially by the observation that clgR-like genes and ClgR operator sequences in front of clpC and clpP1P2 genes are present in nearly all sequenced Actinomycetales genomes [3]. When setting up a proteome map of the C. glutamicum cytosol, the presence of a highly abundant protein with similarity to GlpX-like fructose-1,6-bisphosphatases from other organisms was observed [31]. The constitutive high abundance in C. glutamicum was in sharp contrast to the tight regulation of the corresponding gene in other organisms, suggesting a housekeeping function of GlpX in C. glutamicum. In fact, this protein was shown to have fructose-1,6-bisphosphatase activity and is required for growth of C. glutamicum on gluconeogenic substrates, therefore, probably representing the only fructose-1,6-bisphosphatase in C. glutamicum [30]. This example shows that mere proteome mapping can provide initial information on individual proteins and stimulate subsequent research that ultimately leads to functional characterization of these proteins.

6.5 RECENT DEVELOPMENTS AND OUTLOOK The aim of future approaches must be the development of protocols to overcome the current limitations of 2D PAGE, the analysis of alkaline, hydrophobic, and lowabundance proteins, either by improvement of 2D PAGE or by using alternatives, such as isotope-coded affinity tags (ICAT) or multidimensional protein identification technology (MudPIT) [7,35]. Another focus is the analysis of protein–protein and protein–DNA interactions on the proteomic scale, thereby extending the scope of proteomics beyond the analysis of isolated proteins, which often does not reflect the physiological conditions. Moreover, these approaches are suitable both for the elucidation of regulatory pathways and networks as well as for the functional characterization of proteins. Both microarrays and in vivo or in vitro tagging techniques have been successfully used [13,17]. In the case of C. glutamicum, the in vivo tagging of proteins has already been employed for the purification and characterization of the cytochrome bc1-aa3 supercomplex [22]. Similarly, immobilization of the regulatory DNA regions in front of the C. glutamicum clpC and clpP operons on paramagnetic beads was successfully employed for the enrichment and subsequent

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identification of the transcriptional regulator ClgR [3]. Finally, a combination of global approaches, such as genomics, proteomics, transcriptomics, and metabolomics with bioinformatics, will be desirable for providing a global view on cellular metabolism.

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15. Kaufmann H, Bailey JE, and Fussenegger M. (2001) Use of antibodies for detection of phosphorylated proteins separated by two-dimensional gel electrophoresis. Proteomics 1:194–199. 16. Klose J and Kobalz U. (1995) Two-dimensional electrophoresis of proteins: an updated protocol and implications for a functional analysis of the genome. Electrophoresis 16:1034–1059. 17. Krylov AS, Zasedateleva OA, Prokopenko DV, Rouviere-Yaniv J, and Mirzabekov AD. (2001) Massive parallel analysis of the binding specificity of histone-like protein HU to single- and double-stranded DNA with generic oligodeoxyribonucleotide microchips. Nucleic Acids Res. 29:2654–2660. 18. Lange C, Rittmann D, Wendisch VF, Bott M, and Sahm H. (2003) Global expression profiling and physiological characterization of Corynebacterium glutamicum grown in the presence of L-valine. Appl. Environ. Microbiol. 69:2521–2532. 19. Lichtinger T, Riess FG, Burkovski A, Engelbrecht F, Hesse D, Kratzin HD, Krämer R, and Benz R. (2001) The low-molecular-mass subunit of the cell wall channel of the Gram-positive Corynebacterium glutamicum. Immunological localization, cloning and sequencing of its gene porA. Eur. J. Biochem. 268:462–469. 20. Link AJ, Robison K, and Church GM. (1997) Comparing the predicted and observed properties of proteins encoded in the genome of Escherichia coli K-12. Electrophoresis 18:1259–1313. 21. Matsushita K, Otofuji A, Iwahashi M, Toyama H, and Adachi O. (2001) NADH dehydrogenase of Corynebacterium glutamicum. Purification of an NADH dehydrogenase II homolog able to oxidize NADPH. FEMS Microbiol. Lett. 204:271–276. 22. Niebisch A and Bott M. (2003) Purification of a cytochrome bc-aa3 supercomplex with quinol oxidase activity from Corynebacterium glutamicum. Identification of a fourth subunit of cytochrome aa3 oxidase and mutational analysis of diheme cytochrome c1. J. Biol. Chem. 278:4339–4346. 23. Nolden L, Ngouoto-Nkili CE, Bendt AK, Krämer R, and Burkovski A. (2001) Sensing nitrogen limitation in Corynebacterium glutamicum: the role of glnK and glnD. Mol. Microbiol. 42:1281–1295. 24. Ohlmeier S, Scharf C, and Hecker M. (2000) Alkaline proteins of Bacillus subtilis: first steps towards a two-dimensional alkaline master gel. Electrophoresis 21:3701–3709. 25. Patterson SD and Aebersold RH. (2003) Proteomics: the first decade and beyond. Nat. Genet. 33 Suppl:311–323. 26. Patton WF. (2000) A thousand points of light: the application of fluorescence detection technologies to two-dimensional gel electrophoresis and proteomics. Electrophoresis 21:1123–1144. 27. Pieper R et al. (2003) The human serum proteome: Display of nearly 3700 chromatographically separated protein spots on two-dimensional electrophoresis gels and identification of 325 distinct proteins. Proteomics 3:1345–1364. 28. Reinscheid DJ. (1994) Physiologische und genetische Untersuchungen des AcetatStoffwechsels in Corynebacterium glutamicum. Ph. D. Thesis. Universität Düsseldorf. 29. Reinscheid DJ, Eikmanns BJ, and Sahm H. (1994) Malate synthase from Corynebacterium glutamicum: sequence analysis of the gene and biochemical characterization of the enzyme. Microbiology 140:3099–3108. 30. Rittmann D, Schaffer S, Wendisch VF, and Sahm H. (2003) Fructose-1,6-bisphosphatase from Corynebacterium glutamicum: expression and inactivation of the fbp gene and biochemical characterization of the enzyme. Arch. Microbiol. 180:285–292.

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31. Schaffer S, Weil B, Nguyen VD, Dongmann G, Günther K, Nickolaus M, Hermann T, and Bott M. (2001) A high-resolution reference map for cytoplasmic and membraneassociated proteins of Corynebacterium glutamicum. Electrophoresis 22:4404–4422. 32. Schmid R, Uhlemann EM, Nolden L, Wersch G, Hecker R, Hermann T, Marx A, and Burkovski A. (2000) Response to nitrogen starvation in Corynebacterium glutamicum. FEMS Microbiol. Lett. 187:83–88. 33. Urquhart BL, Cordwell SJ, and Humphery-Smith I. (1998) Comparison of predicted and observed properties of proteins encoded in the genome of Mycobacterium tuberculosis H37Rv. Biochem. Biophys. Res. Commun. 253:70–79. 34. VanBogelen RA, Schiller EE, Thomas JD, and Neidhardt FC. (1999) Diagnosis of cellular states of microbial organisms using proteomics. Electrophoresis 20:2149–2159. 35. Washburn MP, Wolters D, and Yates JR, III. (2001) Large-scale analysis of the yeast proteome by multidimensional protein identification technology. Nat. Biotechnol. 19:242–247. 36. Wasinger VC, Pollack JD, and Humphery-Smith I. (2000) The proteome of Mycoplasma genitalium. CHAPS-soluble component. Eur. J. Biochem. 267:1571–1582. 37. Zhu H, Bilgin M, and Snyder M. (2003) Proteomics. Annu. Rev. Biochem. 72:783–812.

Part IV Transport

7

The Cell Envelope of Corynebacteria M. Daffé

CONTENTS 7.1 7.2

Introduction ..................................................................................................121 Ultrastructural Appearance of the Cell Envelope........................................122 7.2.1 Transmission Electron Microscopy .................................................122 7.2.2 Freeze-Etch Electron Microscopy ...................................................123 7.3 Chemical Nature of the Cell Envelope Layers............................................127 7.3.1 The Plasma Membrane ....................................................................127 7.3.2 Cell Wall Skeleton ...........................................................................129 7.3.2.1 Peptidoglycan....................................................................129 7.3.2.2 Arabinogalactan: Glycosyl Linkage Composition and Structural Features ............................................................129 7.3.2.3 Mycolic Acids: Structure and Biosynthesis .....................131 7.3.3 Cell Wall Proteins ............................................................................134 7.3.3.1 Mycoloyltransferases ........................................................135 7.3.3.2 Porins ................................................................................137 7.3.4 Noncovalently Bound Cell Wall Lipids...........................................138 7.3.5 Outer Layer ......................................................................................138 7.4 Features of the Cell Wall Lipid Layer.........................................................139 7.5 Future Prospects...........................................................................................139 Acknowledgments..................................................................................................140 References..............................................................................................................141

7.1 INTRODUCTION Corynebacteria belong to a suprageneric group of Gram-positive microorganisms called Corynebacterianeae, which includes mycobacteria, nocardia, rhodococci, and other phylogenetically related bacteria. This suborder within the Actinomycetales is typified by the presence of characteristic cell wall components, which give these bacteria a distinctive cell wall architecture. A unique feature of all members of this group is the presence of long-chain α-alkyl, β-hydroxy fatty acids, the so-called mycolic acids. These fatty acids are thought to form, in addition to the plasma membrane, a second lipid bilayer close to the cell surface [17,27]. Evidence has 121

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been presented that mycolic acids play a role in determining the permeability of the cell walls of Corynebacterianeae [33,40,90]. Therefore, although Corynebacterianeae are Gram-positive bacteria whose cell envelope is composed of a plasma membrane and a peptidoglycan-based cell wall [81], they share with Gram-negative bacteria the property of forming in their cell envelope an additional outer barrier that is distinct from the plasma membrane. The existence in mycobacteria and corynebacteria of this outer membrane diffusion barrier is reinforced by the characterization of cell envelope proteins with pore-forming ability [56,64–66,76,79,96,100,115,116]. According to current models [68,72,93] the outer permeability barrier in mycobacteria is composed of a bilayer that consists of mycoloyl residues covalently linked to the cell wall arabinogalactan and other lipids extractable with organic solvents, which are arranged to form the other leaflet of the lipid bilayer. In corynebacteria, however, the number of covalently linked mycoloyl residues is reduced and the extractable trehalose mycolates are likely present in both leaflets of the outer bilayer [91]. The cell wall–linked mycolates certainly participate in this barrier since the disruption of genes that encode mycoloyltransferases in both Mycobacterium tuberculosis and Corynebacterium glutamicum causes a decrease in the amount of cell wall–bound mycolates [30,52,90], and affects the permeability of the envelope of the mutants [52,90]. Additionally, the outermost part of the cell envelope of mycobacteria and corynebacteria consists of carbohydrate and protein with some lipid [63,83,84,91]. Most of the work on the cell envelope of Corynebacterianeae has been done on different Mycobacterium species, due to the medical importance of several of these bacteria, notably M. tuberculosis, the causative agent of tuberculosis, which still kills two to three million people each year worldwide. However, other members of Corynebacterianeae have also been studied, and there is now increasing information available on C. glutamicum since it has been recognized that features of the cell envelope characteristic of corynebacteria might also be important for amino acid excretion [35]. This chapter will review the development of ideas about the way the corynebacterial envelope, in particular that of C. glutamicum, is arranged to control permeability, along with the experimental data that support these ideas.

7.2 ULTRASTRUCTURAL APPEARANCE OF THE CELL ENVELOPE 7.2.1 TRANSMISSION ELECTRON MICROSCOPY Examination of ultrathin sections from conventionally fixed cells of C. glutamicum (Figure 7.1A) reveals a layered cell envelope structure: (i) a plasma membrane (PM) of 6 to 7 nm composed of two leaflets; (ii) a thick electron-dense layer (EDL) of 15 to 20 nm; (iii) an electron-transparent layer (ETL) of 7 to 8 nm; and (iv) a thin outer layer (OL) of 2 to 3 nm. This ultrastructural appearance is similar to that found in mycobacteria [27,31,86,94], except that in mycobacteria a space is usually observed between the PM and the EDL corresponding to a hypothetical periplasmic space [27]. In corynebacteria [69,91], the PM is tightly associated to the EDL. In

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FIGURE 7.1 Thin sections of C. glutamicum cells prepared by conventional embedding. Bacterial pellets were fixed 1 h at 4°C with 2.5% (w/v) glutaraldehyde and 0.05 M lysine in a 0.1 M cacodylate buffer containing (B, C) or not containing (A) 0.075% (w/v) ruthenium red. OL, outer layer; ETL, electron-transparent layer; EDL, electron-dense layer; PM, plasma membrane. Bars represent 200 nm (A, B) and 100 nm (C), respectively [91].

thin sections of corynebacterial cells an asymmetrical appearance of the PM is often visible [91], as similarly observed for mycobacteria [107,108], in that the outer leaflet of PM is thicker than the inner one (Figure 7.1C). This is attributed to the presence of excess glycoconjugates, possibly lipopolysaccharides in the outer leaflet. The electron density of the EDL makes it likely that it contains the cell wall peptidoglycan, which possesses charged groups able to bind metallic stains used in electron microscopy, and also contains part of the covalently linked arabinogalactan [27]. The ETL is traditionally attributed to the mycolic acid layer, based on the transparency of this layer to electrons; however, this interpretation is questionable since an ETL of similar width is also observed in thin sections of C. amycolatum [91], a species devoid of mycolic acids [9,23]. The OL has a width of 2 to 3 nm in thin sections conventionally stained with lead citrate. However, when ultrathin sections are stained with ruthenium red, a stain that has been previously shown to strongly react with the surface of mycobacteria [94,95], a much thicker OL [35 to 40 nm] is seen around C. glutamicum (Figure 7.1B and Figure 7.1C) and other corynebacteria such as C. amycolatum [91]. This observation indicates that the thin margin observed for OL in conventionally fixed cells and freeze-substituted samples of both corynebacteria [69] and mycobacteria [86] is the result of the shrinkage and distortion caused by dehydration of formerly hydrated structures during processing and does not reflect the original thickness of the layer [27]. In addition, the OL is not synonymous with the S-layer, as previously suggested [69], since no differences, in terms of either staining properties or thickness, have been observed between ultrathin sections of C. glutamicum possessing an S-layer and its isogenic mutant devoid of the S-layer [91].

7.2.2 FREEZE-ETCH ELECTRON MICROSCOPY One of the most decisive indications of the occurrence of an outer lipid bilayer in the cell envelope of Corynebacterianeae comes from electron microscopy of freezefractured preparations of whole cells. A characteristic of this technique is that it displays “fracture planes” in the specimen in regions where the fracture is diverted from its direct course through the specimen by planes of weakness in the biological structure. Such planes are produced by hydrophobic structures at low temperature

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(–125˚C). The classic example is provided by the ubiquitous plasma membrane: during preparation, when the fracture encounters a membrane, it tends to follow the hydrophobic interface between the two monolayers (leaflets) of fatty acid side-chains of the membrane [15]. This predictably happens in the case of the plasma membrane. Interestingly, in C. glutamicum (Figure 7.2A), as well as in most Corynebacterianeae [8,12,19,20,113], an additional fracture plane of weakness is observed, within the structure of the cell envelope, near the cell surface. This is in accordance with an

CW

500 nm A

PM

500 nm B

FIGURE 7.2 Freeze-fracture and deep-etch electron microscopy of corynebacterial strains grown on BHI-containing agar plates. Freeze-fractured specimen show that while the main fracture plane is seen in the cell wall (CW) of C. glutamicum CGL2005 close to the bacterial surface (A), it occurs in the plasma membrane (PM) of C. amycolatum (B). Deep-etched preparations of C glutamicum CGL2005 show the ordered surface layer (SL) of the bacterium (C; next page). (Courteously, M. Chami)

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SL

500 nm C

FIGURE 7.2 (continued).

outer lipid bilayer, and also agrees with the demonstration that C. amycolatum, which is devoid of mycolic acids, does not exhibit this second fracture plane, but only the fracture due to the PM (Figure 7.2B). However, as an exhaustive analysis has shown [91], the relative amounts of lipids contributing to the outer bilayer might be different in various Corynebacterianeae. These lipids consist mainly of mycolic acids that are either esterified to the arabinan termini of the cell wall arabinogalactan (see Section 7.3.2) or occur as trehalose mono- and di-mycolates loosely attached to the cell wall and extractable with organic solvents. Although in M. bovis (Bacillus Calmette-Guérin) the amount of arabinogalactan-bound mycolic acid is sufficient to form a monolayer with an area similar to that of a mycobacterial cell [82], and this is also the case for M. tuberculosis, in which the arabinogalactan-bound mycolic acid constitutes about 10% of the cellular dry weight [52], the amount of arabinogalactan-bound mycolic acid in Corynebacterium is considerably less [91]. In C. glutamicum, C. pseudodiphtheriticum, and C. diphtheriae it is 1.0 to 2.5%. Nevertheless, the second fracture plane is visible. Together with the rather high amount of loosely bound lipids of 5.8 to 8.0%, which is almost twice that found in C. amycolatum, this quantitative analysis is clear evidence that in corynebacteria such as C. glutamicum, the outer lipid bilayer consists in a large part of trehalose mycolates (Figure 7.3). Additional support for this model comes from the fact that the fracture plane is not visible in the type strain of C. xerosis. Although this strain possesses an amount of arabinogalactan-bound mycolic acid that is in the range of that of C. glutamicum and C. diphtheriae, it has only a low amount of noncovalently bound lipids (3.2%), which is apparently too little to consistently establish an outer lipid bilayer [91]. In contrast, inactivation of the mycoloyltransferase csp1 gene in C. glutamicum, resulting in a reduction of the arabinogalactan-bound mycolic acid content from 1.0 to 0.5%, does not affect the formation of the second fracture plane, suggesting that in this strain even in this situation, the amount of noncovalently bound lipids is sufficient

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ETL

OL

126

FP

EDL

AG

PM

PG

FP

FIGURE 7.3 A model for the cell envelope of C. glutamicum. From the cytoplasm to the external side of the bacteria the cell envelope is composed of a plasma membrane (PM), a complex wall that is seen in thin sections (Figure 7.1) as an electron-dense layer (EDL), an electron-transparent layer (ETL), and an outer layer (OL). The PM is a typical bilayer of proteins (rectangles and ovals) and phospholipids. The EDL consists of a thick peptidoglycan (PG) covalently linked to the heteropolysaccharide arabinogalactan (AG) with some of the arabinosyl termini of this polysaccharide esterified by C32-36 mycolic acids (thin parallel bars). These together with other noncovalently linked lipids, e.g., trehalose mono- and dicorynomycolates (a pair of empty squares with one or two pairs of thin parallel bars) forms the inner leaflet of a symmetric bilayer. The outer leaflet of this lipid layer is composed of noncovalently linked lipids. In addition, as with Gram-negative bacteria, the cell envelope of corynebacteria contains proteins (grey squares and circles), including those with pore-forming ability (grey squares forming a channel). In freeze-fractured and deep-etched preparations of C. glutamicum (see Figure 7.2) the major fracture plane (FP) is seen within the cell wall, presumably located between the two leaflets of the cell wall lipid layer (arrow). In strains devoid of corynomycolates, e.g., C. amycolatum (see Figure 7.2), the FP occurs within the PM (arrow). Different noncovalently linked lipids and proteins are also present in the OL which consists of a polysaccharide matrix (dotted). The S-layer present on the surface of some C. glutamicum strains is not represented in this model.

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to replace arabinogalactan-bound mycolic acid residues at the inner monolayer to form an outer lipid bilayer superimposable to that of its parent. Freeze-etch electron microscopy also shows the presence of highly ordered arrays on the surface of C. glutamicum CGL2005, previously named Brevibacterium lactofermentum15 [13] (Figure 7.2C). This surface layer (S-layer) is also visible in freeze-fractured micrographs of C. glutamicum ATCC17965 (formerly C. melassecola [87]) and ATCC14752, and of Corynebacterium sp. 2262 [109], but is not visible in C. glutamicum ATCC13032. In the case of strain CGL2005, the S-layer consists of a single protein of 52 kDa, PS2 [19,20,87]. The protein is encoded by the cspB gene, whose disruption causes the disappearance of the S-layer [87], and the formation of the S-layer depends on the growth conditions and carbon source. For instance, the surface of C. glutamicum ATCC 14752 is entirely covered with crystalline arrays of PS2 when grown on solid medium in the presence of glucose, whereas cells are only partly covered when grown in liquid medium, an observation that correlates with the amounts of PS2 extracted from cells grown under these conditions. Replacement of glucose by lactate as a carbon source increases the amount of the protein [109].

7.3 CHEMICAL NATURE OF THE CELL ENVELOPE LAYERS One working model of the cell envelope derived from studies of different Corynebacterium species is shown in Figure 7.3 [91]. The innermost layer of the envelope is the plasma membrane. One of the key functions of the rest of the envelope constituents is to protect this vital and sensitive structure from external influences. Adjacent to the plasma membrane is the cell wall skeleton, a giant macromolecule entirely surrounding the cell consisting of peptidoglycan covalently linked to arabinogalactan, a complex branched polysaccharide, which in turn is esterified by mycolic acids. Associated with the cell wall skeleton, but not covalently attached to it, are a variety of other lipids, of which a majority are phospholipids and trehalose mycolates. The surface of the envelope consists of a more-or-less loosely attached outer layer of polysaccharides, proteins, and lipids.

7.3.1 THE PLASMA MEMBRANE The basic structure of the plasma membrane of the corynebacterial cell envelope does not differ from that of the plasma membrane of other organisms. Polar lipids, mainly phospholipids, are assembled, in association with proteins, into a lipid bilayer. The polar lipids are composed of hydrophilic head groups and fatty acid chains that consist of mixtures of straight-chain, saturated, and unsaturated fatty acid residues having less than 20 carbons. In C. glutamicum palmitic (C16:0) and octadecenoic (C18:1) acids are the major fatty acids present [16,24,75]. 10-Methyloctadecanoic acid (tuberculostearic acid) is also found in small quantities [91]. The main phospholipid of C. glutamicum is phosphatidylglycerol, representing up to 80% of

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CH3OH

O

CH3OH O

O O

O O

O P

O

O

C

C

O

O O

O

CH3

CH3

OH

CH3 CH CH3 O

O

O

O

C OH CH

CH

C OH CH

O

CH

FIGURE 7.4 Examples of representative lipids of the corynebacterial envelope: phosphatidylmyo-inositol dimannoside (PIM2, left) and trehalose dicorynomycolate (TDCM, right). The chains of the mycolic acid, i.e., α-branch and meromycolic chains, are shown aligned as in a lipid layer.

the lipids present, followed by diphosphatidylglycerol (cardiolipin), phosphatidylinositol, and phosphatidylinositol dimannosides (PIM2, Figure 7.4), which occur in small amounts [44,77,91,102]. The PIM2 apparently occurs in the diacylated and triacylated forms and not in the tetraacylated component as prominent for mycobacteria [16]. While PIM6 and phosphatidyl ethanolamine are also present in mycobacteria, these compounds are absent from corynebacteria. The asymmetrical appearance of the corynebacterial plasma membrane in thin sections (Figure 7.1), similar to that observed in mycobacteria [27], is thought to be due to the presence of excess glycoconjugates in the thicker outer leaflet as compared to the inner one. These glycoconjugates are in Mycobacterium species the extended PIM derivatives lipoarabinomannan (LAM) and lipomannan (LM) [47,48]. They possess the lipophilic phoshatidylinositol tail presumably anchoring LAM and LM in mycobacteria in the plasma membrane, as does the lipid part of lipoteichoic acid in Gram-positive bacteria. Therefore, LAM and LM might be regarded as membrane components [17,27], though in fact their precise localization in the envelope is unknown. In Corynebacterium species, two types of lipopolysaccharides are present in phenol/water extracts [91]. However, they exhibit significantly faster mobilities in SDS-PAGE gels than the corresponding mycobacterial lipoglycans, indicating their smaller mass. Further characterization of these substances in C. glutamicum showed that they correspond to LM-like materials and to a lesser

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extent to LAM-like molecules. Both lipoglycans contain C16:0 and C18:1 fatty acid substituents and mannose; LAM-like substances also contain arabinose. Some strains of corynebacteria possess two types of LAMs that differ from each other by the size of their carbohydrate moieties, which are much shorter than those of mycobacterial LAM [41,91], consistent with the occurrence of two types of arabinomannans in the extracellular and surface-exposed materials of the species (see Section 7.3.5.).

7.3.2 CELL WALL SKELETON The chemical structure of the cell wall skeleton of Corynebacterianeae has been extensively studied (for reviews see [27,73–75,111]). It can be described as meso-diaminopimelic acid (DAP)–containing peptidoglycan to which is linked arabinogalactan, which in turn is esterified by mycolic acids, thus forming a large mycoloyl-arabinogalactan-peptidoglycan complex. 7.3.2.1 Peptidoglycan The peptidoglycan is similar to one of the most common types present in bacteria, as found for example in Escherichia coli. The glycan moiety is made up of alternating β-1,4-linked N-acetylglucosamine and N-acetyl muramic acid residues [99]. However, in all mycobacteria examined the muramic acid is N-acylated with a glycoloyl residue rather than the usual acetyl residue [2]. Although a similar variation may occur in corynebacterial peptidoglycan, there is lack of direct proof of such a modification. The carboxyl groups of muramic acid are linked to tri- or tetrapeptides; the major peptide units substituted on the muramic acid residues of the peptidoglycan of C. diphtheriae are the tetrapeptide L-Ala-D-Glu-meso-DAP-D-Ala and the tripeptide L-Ala-D-Glu-meso-DAP [57]. Peptides attached to muramic acid residues of different glycan chains may form interpeptide linkages, resulting in a rigid insoluble network surrounding the plasma membrane. Only a portion of the tetrapeptide and tripeptide subunits is reported to be cross-linked through D-Ala-meso-DAP bridges [57]; the remaining portion of the cross-linkages may consist of bonds involving two residues of DAP, as is typical in mycobacteria [121]. 7.3.2.2 Arabinogalactan: Glycosyl Linkage Composition and Structural Features Arabinogalactan is a heteropolysaccharide, composed mainly of D-arabinofuranosyl and D-galactosyl residues. In mycobacterial species, it is covalently attached to the peptidoglycan through a phosphodiester linkage via rhamnose and glucosamine [70]. It is likely that in the closely related genera a similar arrangement occurs since the constitutive sugars of the linker arm that terminates the galactan chain of arabinogalactan are present in rhodococci and nocardia [29]. Detailed glycosyl linkage composition analysis of arabinogalactan from C. glutamicum through the characterization of the different partially O-methylated, partially O-acetylated alditols [91] showed that the polysaccharide consists of the same types of glycosyl linkages found in mycobacterial arabinogalactans [28,29]. This shows that the arabinogalactan of C. glutamicum shares with the mycobacterial

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Rha GlcNAc P Mur

Corynomycolate T-β-D-Araf

T-β-D-Galf

2-α-D-Araf

5, 6-β-D-Galf

5-α-D-Araf

6-β-D-Galf

3, 5-α-D-Araf

5-β-D-Galf

FIGURE 7.5 A possible arrangement of the structural motifs of the cell wall arabinogalactan of C. glutamicum.

polysaccharides basic structural features, probably including the linear alternating 5- and 6-substituted β-D-galactofuranosyl of the homogalactan and the linear α-Darabinofuranosyl residues with branching produced by 3,5-substituted α-D-arabinofuranosyl units substituted at both positions by α-D-arabinofuranosyl residues (Figure 7.5). Interestingly, however, analysis of the 13C nuclear magnetic resonance (NMR) spectrum of the purified arabinogalactan of C. glutamicum (Tropis, M. and Daffé, M., unpublished data) showed that the polysaccharide is devoid of the nonreducing penta-arabinosyl termini that typify mycobacterial arabinogalactans

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[28,29]. Two carbon signals attributable to terminal β-arabinosyl and 2-substitutedα-arabinosyl units are present in the spectrum of the arabinogalactan of C. glutamicum, whereas four signals are seen in those of mycobacteria [28,29]. In mycobacteria, portions of these two arabinosyl residues are known to be mycoloylated [71]. Although a mycoloylarabinoside has been isolated from a Corynebacterium species [59], the precise location of the mycoloyl residue on the polysaccharide remains to be established. Based on the relatively small amount of cell wall–linked mycolate in corynebacteria, we arbitrary locate the corynomycoloyl residue on position 5 of the nonreducing terminal β-arabinosyl unit in the tentative structural model of the arabinogalactan of C. glutamicum shown in Figure 7.5. In contrast to C. glutamicum, the purified arabinogalactans of C. diphtheriae, C. xerosis, C. amycolatum [91], and C. hoffmanii, as well as that of leprosy-derived coryneforms [1], contain in addition to arabinose and galactose either glucose or mannose. A similar situation is also observed for Rhodococcus and Nocardia species [29]. 7.3.2.3 Mycolic Acids: Structure and Biosynthesis The presence of mycolic acids is a phylogenetic trait of Corynebacterianeae. A notable exception is found in the genus Corynebacterium, in which a few species, e.g., C. amycolatum [9,23], are devoid of mycolic acids. Fatty acids with 70 to 90 carbon atoms (eumycolates) are associated with mycobacterial species, those with about 50 carbon atoms (nocardomycolates) are characteristic of nocardia, and those with around 30 carbon atoms (corynomycolates) are found in corynebacteria. This criterion has been of great taxonomic interest and is of use in the identification and classification of a number of strains. Mycolic acids are found esterifying the mycoloyl arabinogalactan complex or trehalose or glycerol [49,50,59,101,123]. Small amounts of free mycolic acids may also be found. The first characterized mycolic acid in Corynebacterium was that from C. diphtheriae [60], a C32H64O3 acid (Figure 7.6). The key structural feature of mycolic acids is the occurrence of the α-alkyl, β-hydroxy signature, the “mycolic unit” that is found in all mycolic acids and confers to these fatty acids the property to be cleaved at high temperature to yield an aldehyde and an acid (Figure 7.6), a reaction similar to a reverse Claisen-type condensation. The resulting fragments of the C32 mycolic acid each contain 16 carbon atoms. The stereochemistry of the asymmetric carbon atoms at positions 2 and 3 of mycolic acids has been established and corresponds to 2R-tetradecyl-3R-hydroxyoctadecanoic acid [5]; this stereochemistry is conserved in all mycolic acids examined [6]. Corynomycolic acids represent the simplest forms of the mycolate family, since they possess the shortest chain lengths. In all mycolic acid–containing Corynebacterianeae investigated so far, there is a spectrum of mycolic acids with either saturated or unsaturated chains. The major species of the mycolic acids in C. glutamicum has a total carbon number varying from 30 to 34, though smaller and larger mycolic acids are also present [24,46,53,91]. As evident from a variety of analytical techniques [36,37,75,120,124], in C. glutamicum and other Corynebacterium species the total carbon number can be as small as 16 atoms [124; M. Daffé

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OH

R

OH

R O

R1

O

O

R1 H

aldehyde

pyrolysis

O

H

R2 O

OH

acid

R2 OH

FIGURE 7.6 Structure of the C32 mycolic acid (2R-tetradecyl-3R-hydroxy-octadecanoic acid) of Corynebacterium diphtheriae (top) and scheme of the pyrolytical cleavage that typifies mycolic acids (bottom). R corresponds to the absolute configuration of the asymmetric carbon whereas R1 and R2 represent the acyl long-chains shown on the top.

and M.-A. Lanéelle, unpublished data]. Indeed, the C24 form, composed of C16 aldehyde and C8 acid fragments, also represents a significant molecular species in C. glutamicum [M. Daffé and M.-A. Lanéelle, unpublished data], and in other organisms such as C. ovis [49] and C. diphtheriae [114] as well. The composition of the mycolic acids from a given Corynebacterium species might depend on the growth conditions. For instance, when palmitic and oleic acids are supplied in the growth medium, these fatty acids may be incorporated in both parts of the mycolic acid molecules, yielding C32:0, C34:1, and C36:2 mycolic acids [24,75]. Several lipophilic corynebacteria grown on Tween 80 exhibit a high content of unsaturated mycolic acids, presumably due to incorporation of oleic acid derived from the detergent [7,21,74]. Similarly, C24 to C36 bearing zero to four double bonds have been characterized in mycolic acids of some difficult-to-grow corynebacteria, the so-called leprosy-derived corynebacteria, when they are grown on media containing serum [38]. Also, as analyses of C. lepus [25] and Rhodococcus erythropolis [58] grown on hydrocarbons have shown, these strains appear to synthesize mycolic acids by incorporating degradation products of the hydrocarbons. Surprisingly, the detailed mechanism of mycolic acid synthesis is not yet known. However, due to the fact that corynebacteria possess the simplest forms of mycolic acids, studies with these bacteria have attracted much attention. Based on structural considerations, it has been postulated (Figure 7.7) that C32 mycolic acid could result from the condensation of two C16 fatty acids synthesized by the multienzyme complex

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O

R1

C16 fatty acid

C16 fatty acid OH activation

O

R1

O

R1

OH

X2 carboxylation

activation O

R1

O

R1

X1

X3 COOH

O

R1

O

R1

S Enz

X3 COO−

condensation/decarboxylation

R1

O

O X3

R1

O

O O-Trehalose

R1

R1

2-tetradecy-1, 3-keto-octadecanoate reduction R1

OH O

R1

OH O

X3 R1 C32 mycolate

O-Trehalose R1

FIGURE 7.7 Postulated biosynthetic pathway for the formation of 2-tetradecyl-3-keto-octadecanoate and C32 mycolate in the cell-free extracts of C. diphtheriae and C. matruchotti, respectively, found as esters of trehalose. This does not exclude the possibility that parallel biosynthetic mechanisms occur in whole cells. X1, X2, and X3 represent activated forms of acyl groups and can be different or identical; R1, tetradecyl; * labeled carbons from (1-14C)palmitic acid.

for fatty acid synthesis, the so-called fatty acid synthase I (FAS-I). C. glutamicum possesses two fas-I genes, each encoding a single huge polypeptide, whereas mycobacteria contain only one fas-I gene [22]. Inactivation of the individual corynebacterial fas genes in Brevibacterium ammoniagenes, which might be a C. glutamicum strain (see Chapter 2) demonstrated that only one FAS-IA is essential for growth [110]. Unlike mycobacteria, the genome of C. glutamicum contains no gene encoding

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enzymes involved in the FAS-II system. The intervention of two molecules of palmitic acid in the synthesis of mycolic acid has been proved by incubating either whole cells with palmitic acid [39] or use of cell-free preparations [104, 119]. In C. diphtheriae cell-free extracts incubated with 1-14C-palmitic acid, the labeling was specifically found in carbons 1 and 3 of the putative precursor of mycolic acid, a 2tetradecyl, 3-keto-octadecanoic acid (Figure 7.7). The condensation reaction was determined to involve a carboxylation step since it was inhibited by avidin, an inhibitor of biotin-dependent enzymes [3,89,119]. In contrast, avidin had no effect on the condensation reaction in a cell-free extract of C. matruchotii that synthesizes the mature mycolic acid [104]. Evidence that seems to support this last observation came from the incorporation of 2,2-2H palmitic acid in whole cells of C. matruchotii. Based on the fact that the deuterium atom was found at position C-2 of mycolic acid, it was concluded that the condensation reaction does not implicate an intermediate carboxylation step [61]. In fact, this result can be also explained as resulting from a concerted process of decarboxylation and condensation of the malonyl thioester, as known in fatty acid synthesis [4]. Therefore, it is not yet known whether or not the synthesis of mycolic acid involves a carboxylation step. The condensation products are found in both C. diphtheriae and C. matruchotii cell-free systems as esters of trehalose (Figure 7.7). In the cell-free extract of C. diphtheriae, a 6-(2-tetradecyl, 3-keto-octadecanoyl)-α-D-trehalose was detected in the first seconds of incubation after a pulse-labeling experiment with 1-14Cpalmitic acid [89]. With the cell-free extract of C. matruchotii incubated under the same conditions, the predominant lipid identified was trehalose 6-monomycolate [106]. Glucose and trehalose 6-phosphate, but not trehalose, have been shown to stimulate corynomycolate synthesis from palmitate in the presence of ATP into trehalose monocorynomycolate (TMCM), which is believed to serve as the mycoloyl donor to the cell wall arabinogalactan and yields trehalose dicorynomycolate (TDCM) [103,105]. The question of the importance of trehalose in corynomycolate metabolism has been recently addressed by the analysis of mutant strains of C. glutamicum in which the three pathways leading to the production of trehalose are impaired by deleting the corresponding genes [117,122]. The triple mutant is unable to synthesize mycolic acids when grown on sucrose; the defect is complemented by the addition of either glucose, maltose, or trehalose in the culture medium ([117], M. Tropis, A. Wolf, S. Morbach, R. Krämer, and M. Daffé, unpublished data).

7.3.3 CELL WALL PROTEINS The analysis of cell wall proteins of C. glutamicum revealed the presence of more than 100 individual polypeptides [42,43]. Among them are the two major extracytoplasmic proteins, namely the PS2 protein that constitutes the S-layer of some strains of C. glutamicum [19,20,54], and the PS1 protein that exhibits the property of transferring mycoloyl residues on cell envelope acceptors [90]. While PS2 is clearly surface-located, as already discussed (see Section 7.2.2), PS1 is not surface-exposed and may be cell wall–associated. In addition, porins have been recently characterized in C. glutamicum that are known to be, at least partly, located in the outer lipid bilayer of bacteria.

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7.3.3.1 Mycoloyltransferases Proteins whose deduced sequences are similar to those of mycobacterial antigen 85 complex, a family of 30 to 32 kDa proteins that exhibit mycoloyltransferase activity in vitro [11] and also known as fibronectin-binding proteins (Fbps), have been identified in C. glutamicum [14,30,54]. The first described Fbp-like corynebacterial protein, called PS1, is a polypeptide with a calculated molecular mass of 70,874 kDa, with about half of the polypeptide sequence located at the N-terminus exhibiting high sequence identity to Fbps. PS1 is encoded by the csp1 gene and contains a putative signal peptide, and is therefore predicted to be exported through the plasma membrane [54]. Indeed, pulse-chase studies showed that PS1 is rapidly exported to the cell wall [78] but the protein is definitely not exposed at the cellular surface since it is protected from protease digestion [45]. Nevertheless, small amounts of PS1 are invariably recovered from the stationary culture broth, presumably due to cell lysis or leakage during septation [10,78]. Consistent with its cell wall location, extraction with detergents proved that PS1 is found mainly associated with the outermost cell fraction [54,90]. Inactivation of the csp1 gene and biochemical characterization of the resulting mutant have established the mycolyltransferase activity of PS1 [90]. The csp1-mutant accumulates TMCM, elaborates less TDCM (Figure 7.4) and covalently linked corynomycolates. Furthermore, complementation of the csp1-mutant with truncated PS1 proteins demonstrated that the N-terminal part of PS1 is the portion of the protein that is required for the enzyme activity. The C-terminal part has virtually no function in the mycoloyl activity of the enzyme [90]. The production of TDCM, TMCM, and cell wall–linked corynomycolates by the csp1-mutant indicates the presence of other mycoloyltransferases. Thanks to the recent completion of the genome sequence of C. glutamicum (see Chapter 3), a total of six fbp-like genes have been identified [14,30]. Five of these are present in C. efficiens whereas four are found in C. diphtheriae (Table 7.1). The sizes of the identified proteins of 341 to 483 amino acid residues are similar to those of Fbps but smaller than that of PS1, which consists of 657 amino acids [30]. The proteins are equipped with signal peptides [14,30] and they are located in the cell wall. They all contain the esterase domain, including the three key amino acids (Ser, His, and Glu) necessary for catalysis [11,97]. Accordingly, the fbp-like genes, including csp1, were renamed cmyt (for corynebacterial mycoloyltransferase) by De Sousa- D’Auria et al. [30]. In a parallel work by Brand et al. [14] these genes were named cmt (for corynebacterium mycolyltransferase). One of the cmyts (cmytE) is a pseudogene in C. glutamicum CGL2005 [30] but is functional in C. glutamicum ATCC13032 [14]. This gene is absent from C. efficiens (Table 7.1). A functional analysis of the genes, by biochemical characterization of recombinant mutant strains, revealed that the inactivation of either cmytA, cmytB, cmytD, or cmytF results in the accumulation of TMCM with the concomitant decrease of TDCM and, as expected, complementation of the single mutant strains with the wild-type gene restores the balance between the two glycolipids [30,90]. Furthermore, the wild-type phenotype of the cmytA-inactivated mutant could be restored by complementation with any of the functionally active cmyt genes [30]. The inactivation of cmytA led to the accumulation of a glucose monocorynomycolate, indicating

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TABLE 7.1 Mycoloyltransferases in Corynebacterium Speciesa C. glutamicum ATCC13032

C. glutamicum CGL2005

Cop1, Csp1 657 aa NP_602067

cMytA 657 aa

Cmt2 341 aa NP_602069

cMytB 340 aa

Cmt1 365 aa NP_599594

cMytC 368 aa

Cmt5 411 aa NP_600254

cMytD 411 aa

Cmt3 483 aa NP_600150

cMytE truncatedb

Cmt4 483 aa NP_601385

cMytF 492 aa

C. efficiens JCM44549

C. diphtheriae NCTC13129

669 aa CE2709

638 aa

360 aa CE2710

355 aa

381 aa CE0356

355 aa

390 aa CE1488

406 aa

Absent

Absent

Other Strains C. glutamicum ATCC17965 PS1, 657 aa Q01377

C. ammoniagenes ATCC6872 Protein A, 358 aa BAB62413

Absent 484 aa CE0984

a

When available the accession number of proteins in databases is given. For C. diphtheriae, cMyt ORFs were identified from the chromosome sequencing data at the Sanger center. cMyt, corynebacterial mycoloyltransferase; Cmt, Corynebacterium mycolyltransferase. b Stop after amino acid 180.

that this glycolipid may also be involved in the biosynthesis of TDCM. In addition, the corresponding mutants were influenced by the transfer of corynomycoloyl residues to the cell wall arabinogalactan [30]. Cross-complementation experiments have also shown that while cMytA and cMytB are fully redundant and can replace each other, cMytD and cMytF can only complement the cmytA-inactivated strain for the transfer of mycoloyl residues on TMCM but not on arabinogalactan [30], pointing to the existence of two classes of cMyt proteins (Figure 7.8). Although highly similar to the other cmyts and expressed in both E. coli and C. glutamicum, the inactivation of cmytC has no effect on the transfer of corynomycolates [30]. Thus, C. glutamicum contains plethoric cmyt genes that encode mycoloyltransferase enzymes involved in the transfer of corynomycoloyl residues on both trehalose and the cell wall arabinogalactan. The enzymes are fully redundant for the transfer of mycoloyl residues

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FIGURE 7.8 Substrate specificities of mycoloyltransferases of C. glutamicum (cMyts). While cMytA and cMytB can transfer a mycoloyl residue from trehalose monocorynomycolate (TMCM) on another TMCM to yield trehlose dicorynomycolate (TDCM) and on the cell wall arabinogalactan (AG), cMytD and cMytF are able to synthesize TDCM from TMCM.

on trehalose, but they are only partially redundant for the transfer of these fatty acid residues on arabinogalactan. The simultaneous inactivation of three cmyt genes is required to cause a lack of production of TDCM [14]; TMCM, and presumably mycoloylated cell walls, are still produced by the triple mutant. While complementation of the cmytA-disrupted mutant with fbpA, fbpB, or fbpC restores the defect in the cell wall–linked corynomycolates of the mutant, it has no effect on the balance of trehalose glycolipids [90]. Consistent with this observation, the disruption of fbpC in M. tuberculosis results in a defect in cell wall–linked mycolates but has no impact on the balance of trehalose lipids [52]. Thus, in apparent conflict with the in vitro data showing that Fbps catalyzes the transfer of mycoloyl residues onto trehalose lipids [11], these proteins in vivo transfer mycoloyl residues onto the cell wall arabinogalactan but not on trehalose lipids, and are partially redundant [92]. 7.3.3.2 Porins Like Gram-negative bacteria, cell envelopes of some mycobacteria, corynebacteria, and nocardia contain specialized pore-forming proteins (porins) that might facilitate the passage of small hydrophilic molecules through their outer membranes [56,64–66,76,79,96,100,115,116]. The channel forming activity obtained with organic solvent extracts of C. glutamicum exhibits a conductance of 5.5 nS [64,80]. The major cell wall channel, called PorA, is an unusual hydrophobic small-molecular-mass acidic polypeptide of 45 amino acid residues. The channel formed by PorA is highly cation-selective with a purported diameter of about 2.2 nm. It is assumed to consist of a number of PorA polypeptides that are oligomerized to form the transmembrane channel. A mutant deleted of porA no longer shows the typical 5.5-nS channels and is less susceptible to a number of antibiotics [26]. However, channels with a conductance of about 0.7 nS were still observed with the deletion mutant. Functionally active small-molecular-mass porins also occur in C. diphtheriae and some strains of C. xeroxis, but not in all corynebacteria, e.g., C. amycolatum

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and the type strain of C. xeroxis (ATCC 373). As expected, these peptides are found mainly in the cell wall fractions. They are also present in culture filtrates and on the cell surface, but not in the plasma membranes of the bacteria [91]. A correlation exists between the possession of porins and the existence of an outer permeability barrier (as judged from the existence of a cell wall fracture plane, see Section 7.2.2). Indeed, strains devoid of this barrier would not need these proteins to facilitate the entry of nutrients via the cell wall.

7.3.4 NONCOVALENTLY BOUND CELL WALL LIPIDS Trehalose-containing corynomycolates, i.e., TMCM and TDCM (Figure 7.4), are the major extractable lipids of corynebacteria that produce mycolates. Corynebacteria and related genera produce other acylated sugars when grown in the presence of glucose, fructose, or sucrose [18,51,112]. The other constituents of the lipid extract of corynebacteria include the different classes of phospholipids and lipopolysaccharides that are typical for the plasma membrane (see Section 7.3.1). A tiny amount of phospholipids is found associated to the cell surface of both corynebacteria [91] and mycobacteria [63,84].

7.3.5 OUTER LAYER The cell surface macromolecules extracted from Corynebacterianeae by gentle shaking with glass beads consists primarily of polysaccharides ( 90%) with arabinose, mannose, and glucose being the major sugar constituents [62,63,83,91]. The major polysaccharide of C. glutamicum and other corynebacterial species examined so far is a neutral glucan that is eluted from a size-exclusion column at a position corresponding to an apparent molecular mass of 110 kDa [91]. Arabinomannans of apparent molecular masses of 13 kDa and 1.7 kDa are also found in the outermost constituents of corynebacteria while only the larger type of arabinomannan is present in mycobacteria [62,63,83]. The only documented protein present at the outermost bacterial surface of some corynebacterial strains is the S-layer protein, PS2 (see Section 7.2.2). The protein is tightly associated to the cell envelope and resistant to treatment with proteases and detergent [20]. Incubation of whole cells with proteases releases large patches of S-layer composed of PS2 protein truncated at its C-terminus. Lipids extracted both from the culture filtrates and the surface of Corynebacterium species consist mainly of TDCM and TMCM, and small amounts of phospholipids, notably PIM2 and phosphatidyl glycerol [91]. Corynebacteria differ from mycobacteria by the presence of large amounts of lipids in their culture filtrates (10 to 30% cell dry mass [91]), while only traces of lipids are found in the case of mycobacteria [62,63,84]. All the classes of lipids found for whole bacteria are also exposed on the cell surface of corynebacteria, in sharp contrast to what has been observed for mycobacteria, in which only selective classes of lipids were found exposed on the cell surface [84]. Thus, although the cell envelopes of corynebacteria and mycobacteria have in common several chemical and physical properties, the two groups may be different in terms of intimate arrangement of the constituents.

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7.4 FEATURES OF THE CELL WALL LIPID LAYER Based on the accumulated data presented in this chapter it appears that Corynebacterium species contain a specific outer lipid layer, different from the plasma membrane, in their cell envelope. The peculiarities of this lipid layer are (i) the existence of specialized pore-forming proteins (porins), (ii) the formation of a major fracture plane in addition to the expected plasma membrane fracture plane, and (iii) the likely occurrence of noncovalently linked lipids in both leaflets of the outer lipid layer in addition to the contribution of cell wall–bound lipids (mycolic acids) to the inner leaflet. This latter point is evident from quantitative analysis of bound and noncovalently linked lipids and from the fact that suspensions of synthetic TDCM exhibit well-defined transition phases [34]. Interestingly, C32-C36-corynomycolates that possess two parallel C16-C18 chains (see Figure 7.4) are clearly different from the mycobacterial mycolic acids (eumycolates) in which both fatty acid chains are different in length. Therefore, corynomycolates and phospholipids could participate together in the formation of lipid bilayers. Accordingly, we have postulated that noncovalently linked lipids would also participate in the structure of both leaflets of the outer membrane of corynebacteria and are probably arranged to form a symmetrical bilayer [91], as opposed to the asymmetric bilayer in mycobacteria [67,72,93]. The presence of the outer lipid layer in Corynebacterium species may be crucial in some physiological conditions. For instance, the cmytA-mutant strain derived from C. glutamicum ATCC13032 exhibits a growth defect when grown on minimal medium [14] but not on rich medium [55]. Although the mycolate content of this strain has not been investigated, mutation in the same gene results in a 50% defect in covalently linked corynomycolates in strain CGL2022 derived from C. glutamicum CGL2005 [90], a parent strain that does not grow on minimal media. Even on rich media, a double mutant with a deletion of cmytA and cmytB exhibits virtually no growth at 34˚C, and at a lower temperature (30˚C) the double mutant is impaired in its growth and morphology. This phenotype is certainly related to the impact of the two mutations on the outer membrane since the mutant is severely affected in its corynomycolate content, with 70% less cell wall–linked mycolates [55]. When the outer lipid barrier is less disturbed, for instance in the case of the cmytA-mutant with a 50% decrease of the amount of cell wall–bound corynomycolates, the initial rate of uptake of acetate and glycerol by the mutant strain is 2- and 10-fold higher, respectively, than that of the parent strain [90].

7.5 FUTURE PROSPECTS It is now realized that in Corynebacterianeae, cell envelope components other than the plasma membrane also influence the transport and access of small molecules, such as substrates, products, and antibiotics. This is particularly true for the outer mycolic acid layer, which has been shown for Mycobacterium species to represent a permeability barrier for the influx of antibiotics [17,32]. In this case major interest is attached to the influx of antibiotics; for C. glutamicum interest is focused rather on the efflux of amino acids. Although up to now only limited information on the efflux properties of corynebacteria is available, a recent study with a porA mutant

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of C. glutamicum clearly shows the influence of this porin on the diffusion properties of the cell [26]. The understanding of the organization and function of the cell envelope, and in particular the synthesis of mycolic acids, remains a major challenge. As the first studies have shown, C. glutamicum might represent a suitable model to deepen knowledge on these subjects, for several reasons: (i) Corynebacterium species possess the simplest cell wall structure, but their cell walls are functionally and structurally close to those of mycobacteria; (ii) they are rapid growers and do not form clumps; (iii) the sizes of the available genomes of the genus Corynebacterium are relatively small (3.3 Mbp for C. glutamicum and 2.5 Mbp for C. diphtheriae) when compared to those of mycobacteria (e.g., 4.4 Mbp for M. tuberculosis [22]; (iv) heterologous expression of mycobacterial antigens in C. glutamicum has proved to be effective [41,90,98]; and (v) C. glutamicum exhibits a large tolerance for mutations in genes involved in the biogenesis of important cell envelope constituents. While enzymes involved in cell wall synthesis are usually essential for the mycobacterial physiology [85,88,118], their absence may be tolerated by corynebacteria. This might be the case, for instance, for enzymes involved in the biosynthesis of mycolic acids as evidenced by the existence of C. amycolatum, a species devoid of these acids [9,23]. Accordingly, these properties of corynebacteria have been recently used for the identification of the key enzyme that condenses two long-chain fatty acids to yield mycolic acids, which may represent a good and specific target for antituberculosis drug development [88]. Similarly, although plethoric in both mycobacteria and corynebacteria the question of the importance of mycoloyltransferases could not yet be addressed in the former genus but was demonstrated in the latter [55], indicating that these enzymes represent putative targets for the development of new antituberculosis drugs. The cell envelope and its constituents control the efflux of amino acids such as L-glutamate, for whose production C. glutamicum is the outstanding organism. It is thus expected that the knowledge of the arrangement of the cell envelope constituents of C. glutamicum will help in the engineering of these bacteria for the cost-effective production of a number of amino acids. It has been demonstrated that overexpression and inactivation of genes involved in lipid biosynthesis have an impact on glutamate efflux [77]. Likewise, construction of mutants derived from glutamate producing strains and impaired in the production of defined cell envelope constituents such as mycolic acids, arabinogalactan and porins, and analysis of their cell envelope composition and their ability to produce amino acids will certainly clarify the contribution of these compounds in the export of molecules of interest. The recent availability of the sequence of C. glutamicum should greatly help the design of key experiments, which in turn would be of potential application in biotechnology.

ACKNOWLEDGMENTS This chapter is dedicated to Dr. Marie-Antoinette Lanéelle, who just retired after her excellent contribution to the chemistry of Corynebacterianeae for the last 40-years. I am indebted to Dr. Marie-Antoinette Lanéelle for initiating me to the chemistry of Corynebacterianeae and for her continuous support and stimulating discussion.

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I am also very grateful to all my collaborators who have contributed to the experimental work on biochemistry and molecular biology of corynebacteria quoted in this review. These include Virginie Puech, Raoudha Kacem, and Marielle Tropis (IPBS, Toulouse); Célia De Sousa-D’Auria, Christine Houssin, Nicolas Bayan, and Gérard Leblon (University of Orsay); Mohamed Chami (Bale); and Pierre Gounon (Nice). I thank Pr. Gilbert Lanéelle for the critical reading of the manuscript.

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15. Branton D. (1966) Fracture faces of frozen membranes. Proc. Natl. Acad. Sci. USA 55:1048. 16. Brennan PJ and Lehane DP. (1971) The phospholipids of corynebacteria. Lipids 6:401. 17. Brennan PJ and Nikaido H. (1995) The envelope of mycobacteria. Annu. Rev. Biochem. 64:29. 18. Brennan PJ, Lehane DP, and Thomas DW. (1970) Acylglucose of the corynebacteria and mycobacteria. Eur. J. Biochem. 13:117. 19. Chami M, Bayan N, Dedieu J-C, Leblon G, Shechter E, and Gulik-Krzywicki T. (1995) Organisation of the outer layers of the cell envelope of Corynebacterium glutamicum: a combined freeze-etch electron microscopy and biochemical study. Biol. Cell 83:219. 20. Chami M, Bayan N, Peyret J-L, Gulik-Krzywicki T, Leblon G, and Shechter E. (1997) The S-layer protein of Corynebacterium glutamicum is anchored to the cell wall by its C-terminal hydrophobic domain. Mol. Microbiol., 23:483. 21. Chevalier J, Pommier MT, Crémieux A, and Michel G. (1988) Influence of Tween 80 on the mycolic acid composition of three cutane corynebacteria. J. Gen. Microbiol. 134:2457. 22. Cole ST et al. (1998) Deciphering the biology of Mycobacterium tuberculosis from the complete genome sequence. Nature 393:537. 23. Collins MD, Burton RA, and Jones D. (1988) Corynebacterium amycolatum sp. nov., a new mycolic acid-less Corynebacterium species from human skin. FEMS Microbiol. Lett. 49:349. 24. Collins MD, Goodfellow M, and Minnikin DE. (1982) A survey of the structures of mycolic acids in Corynebacterium and related taxa. J. Gen. Microbiol. 128:129. 25. Cooper DG, Zajic JE, and Fergus Gracey DE. (1979) Analysis of mycolic acids and other fatty acids produced by Corynebacterium lepus grown on kerosene. J. Bacteriol. 137:795. 26. Costa-Riu N, Burkovski A, Krämer R, and Benz R. (2003) PorA represents the major cell wall channel of the Gram-positive bacterium Corynebacterium glutamicum. J. Bacteriol. 185:4779. 27. Daffé M and Draper P. (1998) The envelope layers of mycobacteria with reference to their pathogenicity. Adv. Microb. Phys. 39:131. 28. Daffé M, Brennan PJ, and McNeil M. (1990) Predominant structural features of the cell wall arabinogalactan of Mycobacterium tuberculosis as revealed through characterization of oligoglycosyl alditol fragments by gas chromatography/mass spectrometry and by 1H and 13C NMR analyses. J. Biol. Chem. 265:6734. 29. Daffé M, McNeil M, and Brennan PJ. (1993) Major structural features of the cell wall arabinogalactans of Mycobacterium, Rhodococcus and Nocardia. Carbohydr. Res. 249:383. 30. De Sousa-D’Auria C, Kacem R, Puech V, Tropis M, Leblon G, Houssin C, and Daffé M. (2003) New insights into the biogenesis of the cell envelope of corynebacteria: identification and functional characterization of five new mycoloyltransferase genes in Corynebacterium glutamicum. FEMS Microbiol. Lett. 224:35. 31. Draper P. (1982) The anatomy of Mycobacteria. In Ratledge C and Stanford JL (Eds.), The Biology of The Mycobacteria, Vol. 1, Academic Press, London, p. 9. 32. Draper P. (1998) The outer parts of the mycobacterial envelope as permeability barriers. Frontiers Biosci. 3:d1253. 33. Dubnau E, Chan J, Raynaud C, Mohan VP, Lanéelle M-A, Yu K, Quémard A, Smith I, and Daffé M. (2000) Oxygenated mycolic acids are necessary for virulence of M. tuberculosis in mice. Mol. Microbiol. 36:630.

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68. Liu J, Rosenberg EY, and Nikaido H. (1995) Fluidity of the lipid domain of cell wall from Mycobacterium chelonae. Proc. Natl. Acad. Sci. USA 92:11254. 69. Marienfeld S, Uhlemann E-M, Schmid R, Krämer R, and Burkovski A. (1997) Ultrastructure of the Corynebacterium glutamicum cell wall. Antonie van Leeuwenhoek 72:291. 70. McNeil M, Daffé M, and Brennan PJ. (1990) Evidence for the nature of the link between the arabinogalactan and peptidoglycan of mycobacterial cell walls. J. Biol. Chem. 265:18200. 71. McNeil M, Daffé M, and Brennan PJ. (1991) Location of the mycoloyl ester substituents in the cell walls of mycobacteria. J. Biol. Chem. 266:13217. 72. Minnikin DE. (1982) Lipids:complex lipids, their chemistry, biosynthesis and roles. In Ratledge C and Stanford JL (Eds.), The Biology of the Mycobacteria, Vol. 1, Academic Press, London, p. 95. 73. Minnikin DE and Goodfellow M. (1980) Lipid composition in the classification and identification of acid-fast bacteria. In Goodfellow M and Board RG (Eds.), Microbiological Classification and Identification, Academic Press, London, p. 189. 74. Minnikin DE and O’Donnell AG. (1984) Actinomycete envelope lipid and peptidoglycan composition. In Goodfellow M, Mordarski M, and Williams ST (Eds.), The Biology of Actinomycetes, Academic Press, London, p. 337. 75 Minnikin DE, Goodfellow M, and Collins MD. (1978) Lipid composition in the classification and identification of coryneform and related taxa. In Bousfield IJ and Galley G (Eds.), Coryneform Bacteria, Academic press, London, p. 85. 76. Mukhopadhyay S, Basu D, and Chakrabarti P. (1977) Characterization of a porin from Mycobacterium smegmatis. J. Bacteriol. 179:6205. 77. Nampoothiri KM, Krumbach K, Möckel B, Sahm H, and Eggeling L. (2002) Expression of genes of lipid synthesis and altered lipid composition modulates L-glutamate exflux of Corynebacterium glutamicum. Appl. Microbiol. Biotechnol. 58:89. 78. Nguyen DT, Houssin C, and Bayan N. (2001) Study of mycoloyltransferase transport across the cell envelope of Corynebacterium glutamicum. FEMS Microbiol. Lett. 201:145. 79. Niederweis M, Ehrt S, Heinz C, Klocker U, Karosi S, Swiderek KM, Riley LW, and Benz R. (1999) Cloning of the mspA gene encoding a porin from Mycobacterium smegmatis. Mol. Microbiol. 33:933. 80. Niederweis M, Maier E, Lichtinger T, Benz R, and Krämer R. (1995) Identification of channel-forming activity in the cell wall of Corynebacterium glutamicum. J. Bacteriol. 177:5716. 81. Nikaido H. (1994) Prevention of drug access to bacterial targets: permeability barriers and active efflux. Science 264:382. 82. Nikaido H, Kim S-H, and Rosenberg EY. (1995) Physical organization of lipids in the cell wall of Mycobacterium chelonae. Mol. Microbiol. 8:1025. 83. Ortalo-Magné A, Dupont M-A, Lemassu A, Andersen ÅB, Gounon P, and Daffé M. (1995) Molecular composition of the outermost capsular material of the tubercle bacillus. Microbiology 141:1609. 84. Ortalo-Magné A, Lemassu A, Lanéelle M-A, Bardou F, Silve G, Gounon P, Marchal G, and Daffé M. (1996) Identification of the surface-exposed lipids on the cell envelopes of Mycobacterium tuberculosis and other mycobacterial species. J. Bacteriol. 178:456. 85. Pan F, Jackson M, Ma Y, and McNeil M. (2001) Cell wall core galactofuran synthesis is essential for growth of mycobacteria. J. Bacteriol. 183:3991. 86. Paul TR and Beveridge TJ. (1992) Reevaluation of envelope profiles and cytoplasmic ultrastructure of mycobacteria processed by conventional embedding and freezesubstitution protocols. J. Bacteriol. 174:6508.

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87. Peyret J-L, Bayan N, Joliff G, Gulik-Krzywicki T, Mathieu L, Shechter E, and Leblon G. (1993) Characterization of the cspB gene encoding PS2, an ordered surface-layer protein in Corynebacterium glutamicum. Mol. Microbiol. 9:97. 88. Portevin D, de Sousa-D’Auria C, Houssin C, Grimaldi C, Daffé M, and Guilhot C. (2004) A polyketide synthase catalyzes the last condensation step of mycolic acid biosynthesis in mycobacteria and related organisms. Proc. Natl. Acad. Sci. USA 101:314. 89. Promé JC, Walker RW, and Lacave C. (1974) Condensation de deux molécules d’acide palmitique chez Corynebacterium diphtheriae: formation d’un β-céto-ester de tréhalose. C. R. Acad. Sc Paris 278:1065. 90. Puech V, Bayan N, Salim K, Leblon G, and Daffé M. (2000) Characterization of the in vivo acceptors of the mycoloyl residues transferred by the corynebacterial PS1 and the related mycobacterial antigens 85. Mol. Microbiol. 35:1026. 91. Puech V, Chami M, Lemassu A, Lanéelle M-A, Schiffler B, Gounon P, Bayan N, Benz R, and Daffé M. (2001) Structure of the cell envelope of corynebacteria: importance of the non-covalently bound lipids in the formation of the cell wall permeability barrier and fracture plane. Microbiology 147:1365. 92. Puech V, Guilhot C, Perez E, Tropis M, Armitige LY, Gicquel B, and Daffé M. (2002) Evidence for a partial redundancy of the fibronectin-binding proteins for the transfer of mycoloyl residues onto the cell wall arabinogalactan termini of Mycobacterium tuberculosis. Mol. Microbiol. 44:1109. 93. Rastogi N. (1991) Recent observations concerning structure and function relationships in the mycobacterial cell envelope:elaboration of a model in terms of mycobacterial pathogenicity, virulence and drug-resistance. Res. Microbiol. 142:464. 94. Rastogi N, Fréhel C, and David HL. (1986) Cell envelope architectures of leprosyderived corynebacteria, Mycobacterium leprae, and related organisms: a comparative study. Curr. Microbiol. 11:23. 95. Rastogi N, Fréhel C, and David HL. (1986) Triple-layered structure of mycobacterial cell wall: evidence for the existence of a polysaccharide-rich outer layer in 18 mycobacterial species. Curr. Microbiol. 13:237. 96. Rieβ FG, Lichtinger T, Cseh R, Yassin AF, Schaal KP, and Benz R. (1998) The cell wall porin of Nocardia farcinica: biochemical identification of the channel-forming protein and biophysical characterization of the channel properties. Mol. Microbiol. 29:139. 97. Ronning DR, Klabunde T, Besra GS, Vissa VD, Belisle JT, and Sacchettini JC. (2000) Crystal structure of the secreted form of antigen 85C reveals potential targets for mycobacterial drugs and vaccines. Nature Structural Biol. 7:141. 98. Salim K, Haedens V, Content J, Leblon G, and Huygen K. (1997) Heterologous expression of the Mycobacterium tuberculosis gene encoding antigen 85A in Corynebacterium glutamicum. Appl. Environ. Microbiol. 63:4392. 99. Schleifer KH and Kandler O. (1972) Peptidoglycan types of bacterial cell walls and their taxonomic implications. Bacteriol. Rev. 36:407. 100. Senaratne RH, Mobasheri H, Papavinasasundaram KG, Jenner P, Lea EJA, and Draper P. (1998) Expression of gene for a porin-like protein of the OmpA family from Mycobacterium tuberculosis H37Rv. J. Bacteriol. 180:3541. 101. Senn M, Ioneda T, Pudles J, and Lederer E. (1967) Spectrométrie de masse de glycolipides. I. Structure du cord factor de Corynebacterium diphtheriae. Eur. J. Biochem. 1:353.

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8

Genomic Analyses of Transporter Proteins in Corynebacterium glutamicum and Corynebacterium efficiens B. Winnen, J. Felce, and M.H. Saier Jr.

CONTENTS 8.1 Introduction ..................................................................................................149 8.2 Computer Methods.......................................................................................150 8.3 Topological Predictions for Membrane Proteins of Corynebacteria...........150 8.4 Classes of Transporters Found in C. glutamicum and C. efficiens .............150 8.5 Classes of Substrates Transported ...............................................................151 8.6 Global Analysis of Transporters and Their Family Associations ...............152 8.7 Channels .......................................................................................................152 8.8 Secondary Carriers.......................................................................................179 8.9 Primary Active Transporters ........................................................................181 8.10 Proton-Pumping Electron Carriers...............................................................182 8.11 Group Translocators.....................................................................................182 8.12 Transmembrane Electron Flow Carriers......................................................182 8.13 Poorly Defined Transporters ........................................................................182 8.14 Perspectives and Conclusions ......................................................................182 Acknowledgments..................................................................................................185 References..............................................................................................................185

8.1 INTRODUCTION Corynebacteria are of tremendous importance both to human health and for industrial purposes. Corynebacterium glutamicum (Cgl) and C. efficiens (Cef), for example, 149

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are two closely related producers of amino acids for commercial purposes [8,15]. Whereas Cef can grow above 40ºC, Cgl cannot, possibly because of differences in the amino acid compositions of the proteins present in the two organisms [15]. Both organisms have genome sizes of about 3.2 Mbp (3.31 for Cgl, 3.15 for Cef), and both genomes have been fully sequenced [9,15,22]. Specific transport systems responsible for the uptake and export of various metabolites related to amino acid metabolism have been better characterized in C. glutamicum than in any other bacterium [3,10,13]. However, comprehensive genome analyses of transporters in these bacteria have not previously been reported. In this chapter, we describe comparative studies of the transporters encoded within the genomes of Cgl and Cef.

8.2 COMPUTER METHODS The complete protein sequences of C. glutamicum and C. efficiens were extracted from the NCBI nonredundant database. Computer-aided analyses were conducted to retrieve all proteins encoded within the genomes of C. glutamicum and C. efficiens that are recognizably homologous to transport system constituents included in the transporter classification database (TCDB) [4,23]. Briefly, all proteins were blasted in an automated manner (using BLASTP) against the Transporter Classification and NCBI databases. Additional databases used for protein functional analysis were the nonredundant SWISSPROT and TrEMBL protein sequence databases. Several protein pattern databases (Conserved Domain Database at NCBI and Pfam) were also used. Charge-bias analyses of membrane protein topology were performed using the TMHMM [12] and WHAT [28] programs.

8.3 TOPOLOGICAL PREDICTIONS FOR MEMBRANE PROTEINS OF CORYNEBACTERIA A protein topological prediction program determined that about 60% of the proteins in both corynebacteria analyzed are expected to be cytoplasmic while about 40% are expected to be integral membrane constituents. Of the latter, 18% were predicted to have 1 TMS (transmembrane segment), 8% had 2 to 3 TMSs, 6% had 4 to 6 TMSs, 4% had 7 to 10 TMSs, and 4% had 11 or more TMSs. Many of the oneTMS proteins may be secreted via the Sec and Tat export systems (see Section 8.8). About 10% of all recognized proteins encoded within the two genomes were predicted to be homologs of recognized transport proteins. Since transporter families include proteins that are usually concerned exclusively with transport [21], it is probable that nearly all of these proteins function in transmembrane transport.

8.4 CLASSES OF TRANSPORTERS FOUND IN C. GLUTAMICUM AND C. EFFICIENS According to the transporter classification (TC) system, transporters are classified into five well-defined categories (classes 1 to 5) and two poorly defined categories

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TABLE 8.1 Categories of Recognized Transport Proteins Found in C. glutamicum and C. efficiens Number of Transport Proteins in TC Class 1 Channels 2 Secondary carriers 3 Primary transporters 4 Group translocators (PTS) 5 Transmembrane electron carriers 8 Auxiliary transport proteins 9 Poorly defined systems

C. glutamicum

C. efficiens

7 117 106 5 3 3 19

7 114 88 2 3 3 18

(classes 8 and 9). The well-defined categories are (1) channels, (2) secondary carriers, (3) primary transporters, (4) group translocators, and (5) transmembrane electron flow carriers [4,20]. The less-well-defined proteins include auxiliary transport proteins (class 8) and transporters or putative transporters of unknown mechanism of action or function (class 9) [13]. Table 8.1 summarizes the distribution of transporters in each of these categories. Only seven recognized channel proteins are found in each of the corynebacteria analyzed. The vast majority of transport proteins found in both organisms are secondary carriers (117 and 114 for Cgl and Cef, respectively) and constituents of primary active transporters (106 and 88 for Cgl and Cef, respectively). Because secondary carriers are usually single-component systems while primary active transporters are often multicomponent systems, the greater numbers of the former indicates that these predominate in corynebacteria. Only a few PTS proteins catalyze group translocation, and only a few transmembrane electron flow systems were identified. The latter fact may in part reflect the limited representation of transmembrane electron flow carriers in TCDB. Three auxiliary proteins of TC class 8 and either 18 or 19 putative transporters of TC class 9 were identified (Table 8.1). The probable functional identities of the individual proteins will be discussed in Sections 8.7–8.13.

8.5 CLASSES OF SUBSTRATES TRANSPORTED Table 8.2 summarizes the numbers of transporters involved in the transport of various types of substrates. About 50 systems for inorganic cations were identified in both organisms, and nearly 40 systems for drugs and hydrophobic substances were found. About 30 systems for amino acids and their derivatives were identified, but only 12 to 14 systems for sugars were detected. Minor differences between the two organisms were found in all of these categories. Surprisingly, Cgl has 14 transport systems for carboxylates, but Cef has only 8. The largest category includes the miscellaneous and unknown systems for which substrate identification was uncertain (140 to 150 proteins; see Table 8.2).

152

Handbook of Corynebacterium glutamicum

TABLE 8.2 Breakdown of Transport Proteins According to Predicted Substrate Types in C. glutamicum and C. efficiens Number of Transport Proteins in Substrate Type Inorganic cations Sugars and their derivatives Mono-, di-, and tricarboxylates Amino acids, amines, amides, and polyamines Drugs and toxic compounds Miscellaneous Unknown

C. glutamicum

C. efficiens

53 14 14 31 40 133 15

49 12 8 30 36 130 13

8.6 GLOBAL ANALYSIS OF TRANSPORTERS AND THEIR FAMILY ASSOCIATIONS Table 8.3 summarizes the results of our detailed analyses of transporters found in Cgl and Cef. Columns 1 through 3 show the TC family number, the name of the family, and its standard abbreviation. Column 4 presents the types of substrates known to be transported by members of the family. Columns 5 and 7 present the protein accession numbers, with the level of confidence for the corresponding functional assignment (1 = sure, 2 = probable, 3 = uncertain or unknown) in columns 6 and 8, and column 9 presents the protein TC number (X = unknown). When two proteins from the two organisms are adjacent to each other in the table, we consider it likely that they are orthologs serving the same function. They show the highest percent identity with each other as compared with any other potential ortholog. Column 10 presents the probable substrate(s) when known. Column 11 gives the percent identity between the two putative orthologs with the actual numbers of amino acids compared presented thereafter. Finally, columns 12 and 13 present the sizes of the proteins in numbers of amino acids for Cgl and Cef, respectively. If a protein is found in one organism, but no probable ortholog could be identified in the other organism, then the latter positions (columns 5 and 6 or columns 7 and 8) are left blank, and column 11 is of necessity also blank. The results of our analyses of the transporter types identified by analysis of both corynebacterial genomes (Table 8.3) revealed striking similarities between the two organisms, but also identified some interesting differences, as discussed in the sections that follow.

8.7 CHANNELS In category 1A (α-type channels), both organisms possess a single member of the VIC (1.A.1), MscL (1.A.22), MscS (1.A.23), and MIT (1.A.35) families. The VIC and MIT family members are probably monovalent- and divalent-cation transporters, respectively, while the MscL and MscS proteins are most likely nonspecific channels

Name of Family

Abbrev.

Typical Substratesd

1.A α-Type Channel-forming Proteins and Peptides 1.A.1 The Voltage-gated VIC Na+; K+; Ca2+; multiple Ion Channel cations (VIC) Superfamily 1.A.22 The Large MscL Proteins, ions Conductance (slightly catMechanosensitive ion selective) Ion Channel (MscL) Family 1.A.23 The Small MscS Ions (slightly Conductance anion selecMechanosensitive tive) Ion Channel (MscS) Family 1.A.29 The Urea/Amide UAC Urea, amides Channel (UAC) Family 1.A.33 The Cation HSP70 Ions, polyChannel-forming peptides Heat Shock Protein-70 (Hsp70) Family 1.A.35 The CorA Metal MIT Heavy metal Ion Transporter ions (MIT) Family

Family TC#a

3

3

3

2 3

3

BAB98272

BAB98663

BAB00194 BAB99771

BAB97454

Evidenced

BAB98170

Protein(s)b Cglc

BAC16888

BAC19094

3

3

2

3

BAC16834

BAC19439

3

3

3

d

Evidence

BAC18174

BAC17763

BAC17602

Protein(s)b Cefc

TABLE 8.3 Putative Transport Proteins Identified in C. glutamicum and C. efficiens

1.A.35.1.X

1.A.33.1.X

1.A.33.1.2

1.A.29.1.X

1.A.23.X.X

1.A.22.1.2

1.A.1.1.1

Protein TC#

373

68% 257/376

Mg2+, Co2+, and Ni2+

335

491

135

335

381

467

353

225

591

151

353

# of aae # of aac Cglc Cef

484

77% 259/334 63% 296/464

46% 204/441

71% 97/136

77% 259/334

% Identity

Ions

Ions

Urea or amides

Ions

K+

Probable Substratesd

Genomic Analyses of Transporter Proteins in C. glutamicum and C. efficiens 153

Name of Family

Abbrev.

DHA2 (14-Spanner)

DHA1 (12-Spanner)

SP

1.B Outer Membrane Porins (β-structure) 1.B.34 The PorA Corynebacterial Porin (PorA) Family 2.A Carrier-type Facilitators 2.A.1 The Major Facili- MFS tator Superfamily (MFS)

Family TC#a

Numerous small molecules (also can serve as re-ceptors)

Ions, small molecules

Typical Substratesd

1 1 2

3 2 3 3 3 1

3 3

BAC00452 BAB98603

BAC00400 BAB98895 BAB97715 BAB98530 BAC00137 AF237667

BAB97774 BAB97773

1

Evidenced

BAB97574

BAB00109 (PorA)

Protein(s)b Cglc

BAC16907 BAC16966 BAC17208

BAC18441 BAC17214 BAC18017 BAC19391

BAC17540

BAC17498

Protein(s)b Cefc

TABLE 8.3 (continued) Putative Transport Proteins Identified in C. glutamicum and C. efficiens

3 3 2

3 3 3 2

3

2

d

Evidence

2.A.1.3.X 2.A.1.3.X 2.A.1.3.X

2.A.1.2.18/ 15 2.A.1.2.X 2.A.1.2.X 2.A.1.2.X 2.A.1.2.X 2.A.1.3.9

2.A.1.1.26 2.A.1.2.14

2.A.1.1.26

1.B.34.1.1

Protein TC#

Lincomycin

Arabinose, lactose & melibiose

Myoinositol Myoinositol

Small molecules

Probable Substratesd

145/366 33/115 270/393 234/396

29% 77/263 66% 341/515

39% 28% 68% 59%

68% 292/424

% Identity

544 549

403 410 405 398 481

392

508 424

491

45

342 527

370 405 425 431 391

403

422

# of aae # of aac Cglc Cef

154 Handbook of Corynebacterium glutamicum

3 3 2

BAB98279 BAB97503

PP

2 3 2 2 2 2

BAB98243

BAB98583 BAB99615 BAC00419 BAB99778 BAB98469 BAB99802

2 3 2 2 2 3

2 2 2 2 2 2 3 3 3 3 2 1

CP

ACS AAHS

NNP

MHS

BAB98531 BAB98635 BAB98747 BAB99885 BAC00005 BAC00079 BAB97468 BAB97559 BAB98758 BAC00222 BAC00327 BAC00460 (ProP) BAB97814 BAB97676 BAB98860 BAC00311 BAC00031 BAC00468

3

3 2 2 2 3

BAC17941 BAC19120 BAC17446 BAC19525 BAC17736 BAC17770

3

3 3

2 2 2 2

2

2 2 2 2 2 2 3 3

BAC19672

BAC18226 BAC18107

BAC17250 BAC17472 BAC18405 BAC19571

BAC19499

BAC18018 BAC18147 BAC18774 BAC19200 BAC19305 BAC19341 BAC19660 BAC19701

2.A.1.17.1 2.A.1.18.X

2.A.1.17.1

2.A.1.8.9 2.A.1.14.X 2.A.1.15.5 2.A.1.15.6 2.A.1.15.X 2.A.1.15.X

2.A.1.6.X 2.A.1.6.X 2.A.1.6.X 2.A.1.6.X 2.A.1.6.X 2.A.1.6.X

2.A.1.3.X 2.A.1.3.X 2.A.1.3.X 2.A.1.3.X 2.A.1.3.X 2.A.1.3.X 2.A.1.3.X 2.A.1.3.X 2.A.1.3.X 2.A.1.3.X 2.A.1.3.X 2.A.1.6.4

Cyanate D-Arabinitol or ribitol

Cyanate

Benzoate

NO3–/NO2– NO3–/NO2–

Prolinebetaine

305/447 331/443 130/445 289/453 285/483 325/476 285/438 48/188

332/430 131/422 377/465 322/419

58% 227/388 64% 241/375

72% 240/330 73% 332/453

76% 340/446

79% 359/449

77% 31% 81% 76%

70% 349/493

68% 74% 29% 63% 59% 68% 65% 25%

400 445

391

450 441 452 475 431 460

467 451 462 433 285 279

449 487 474 475 494 481 483 458 483 403 497 504

417

338 470 449 442 408

446

433 477

533 466 468 465

501

454 512 481 486 531 513 500 414

Genomic Analyses of Transporter Proteins in C. glutamicum and C. efficiens 155

2.A.3

2.A.2

Family TC#a

3 3

BAB99827 BAB99491

AAT

1

2

BAC00372

BAB98500 (AroP)

2 2

BAB98458 BAB99616

Amino acids, polyamines, organocat-ions, (also serves as receptors)

2

BAB97515

Sugars (glycosides)

2

BAC00323

1

Evidenced

YnfM

Protein(s)b Cglc BAC00171

Typical Substratesd

DHA3 (12-Spanner)

Abbrev.

The GlycosideGPH PentosideHexuro-nide (GPH):Cation Symporter Family The Amino AcidAPC PolyamineOrganocation (APC) Family

Name of Family

BAC17974

BAC19192 BAC19191

BAC17724

BAC19624

BAC17142 BAC18934

BAC16934

BAC19579

BAC19418

Protein(s)b Cefc

TABLE 8.3 (continued) Putative Transport Proteins Identified in C. glutamicum and C. efficiens

2

3 2

3

2

3 3

3

2

3

d

Evidence

2.A.3.1.5

2.A.1.X.X 2.A.1.X.X

2.A.1.X.X

2.A.1.X.X 2.A.1.X.X

2.A.1.X.X

2.A.1.36.1

2.A.1.21.2

Protein TC#

% Identity

Tyrosine

Glucuronide

75% 340/451

Erythromy- 63% 294/ cin, tetra465 cycline, puromycin, bleomycin Acriflavin 69% 259/ 375 74% 313/ 422 31% 29/91 53% 84/157 79% 351/ 444

Probable Substratesd

463

373 422

448

406 187

424

378

459

486

393 315

447

480

407 445

433

413

468

# of aae # of aac Cglc Cef

156 Handbook of Corynebacterium glutamicum

The Drug/Metabolite Transporter (DMT) Superfamily

2.A.7

2.A.6

The Cation Diffusion Facilitator (CDF) Family The ResistanceNodulation-Cell Division (RND) Superfamily

2.A.4

DMT

HAE2

SecDF

RND

CDF

APA

Multiple drugs?

Heavy metal ions; multiple drugs; oligosaccharides; organic solvents, fatty acids; phospholipids, cholesterol

Cd2+, Co2+, Ni2+

2 1

BAB98548 BAB98361 (LysI) BAB98674 BAC00177

3 3 3

BAC00261 BAB98317

3

BAB99051

BAB97624

3

BAB99050

2 3

2

BAB97863

BAC18930

BAC19508

2

3

3

2

BAC17452 BAC17796

3

3

3

BAC17010

BAC18581

BAC18580

2 3 2

2

BAC18068 BAC17845 BAC17521 BAC18184

2

BAC18296

2.A.6.X.X

2.A.6.5.X

2.A.6.5.X

2.A.6.4.1

2.A.6.4.1

2.A.4.1.X 2.A.4.1.X

2.A.3.1.X 2.A.3.2.4

2.A.3.1.7

Blue pigment (indigoidine)

Protein secretion Protein secretion Lipid/Actinorhodin Lipid/Actinorhodin Lipid/Actinorhodin Lipid/Actinorhodin

Cd2+, Co2+, Ni2+

Lysine

D-Serine/ D-alanine/ glycine Histidine

37% 310/820 73% 573/775

70% 275/388 75% 461/611

70% 176/251

75% 358/472

72% 315/432

791

772

801

644

403

318 286

468 501

453

323

774

797

754

899

639

378

516 587 251

465

446

Genomic Analyses of Transporter Proteins in C. glutamicum and C. efficiens 157

2.A.11

2.A.10

2.A.9

2.A.8

Family TC#a

The Gluconate:H+ Symporter (GntP) Family The Cytochrome Oxidase Biogenesis (Oxa1) Family The 2-Keto-3deoxygluconate Transporter (KDGT) Family The CitrateMg2+:H+ (CitM) Citrate-Ca2+:H+ (CitH) Symporter (CitMHS) Family

Name of Family

BAC16870

BAC19716

Citrate, Me2+

CitMHS

BAC19754

BAC19597

2-Keto-3deoxygluconate 2

3

BAC00491

BAB97460

3

BAC00334

BAC17421

BAC18840

BAC18851

Protein(s)b Cefc

KdgT

Oxa1

Proteins

3 2

BAB99538 BAC00302

Gluconate, idonate

3

BAB97525

GntP

3

BAB99524

3

Evidenced

RarD

Protein(s)b Cglc BAB97998

Typical Substratesd

DME

Abbrev.

TABLE 8.3 (continued) Putative Transport Proteins Identified in C. glutamicum and C. efficiens

2

2

3

3

2

3

3

d

Evidence

2.A.11.1.2

2.A.9.3.X

2.A.9.3.X

2.A.7.X.X 2.A.8.1.X

2.A.7.7.X

2.A.7.7.1

2.A.7.3.6

Protein TC#

Citrate or D-isocitrate•M2+

2-keto-3deoxygluco -nate

Proteins

Proteins

Gluconate

Threonine/ Homoserine (possibly other amino acids) Chloramphenicol Chloramphenicol

Probable Substratesd

74% 363/489

64% 218/337 77% 246/317

62% 286/461

57% 162/280

30% 71/234

% Identity

489

317

434

286 463

293

280

307

533

362

316

397

464

293

299

# of aae # of aac Cglc Cef

158 Handbook of Corynebacterium glutamicum

The Neurotransmitter: Sodium Symporter (NSS) Family NSS

SSS

PiT

CaCA

2.A.22

2.A.21

2.A.20

2.A.19

The Protondependent Oligopeptide Transporter (POT) Family The Ca2+:Cation Antiporter (CaCA) Family The Inorganic Phosphate Transporter (PiT) Family The Solute:Sodium Symporter (SSS) Family

2.A.17 POT

The Betaine/Carni- BCCT tine/Choline Transporter (BCCT) Family

2.A.15

Sugars; amino BAB98556 acids; vita(PutP) mins; nucleosides; inositols; iodide, BAB98226 organic and inorganic anions; urea; organocations (also can serve as receptors) NeurotransBAB98423 mitters (often amino acids); osmolytes; taurine; creatine

BAB97853

BAB98475

Ca2+

Inorganic phosphate; sulfate

2

BAC00447

2

BAC17893

2

2.A.22.4.1

650

579

551

High affinity 70% 387/546 tryptophan: Na+

551

535

381 445

358

535

479

641

602

615

598

537 2

BAC17902

74% 409/551

524

425

350

453

452

630

615

595

Phenylacetate

Phenylacetate

2

2.A.21.7.1

76% 400/523

2

BAC18072

BAC17720

76% 289/376

54% 191/353

48% 219/448 67% 305/453

64% 389/599

69% 399/571 78% 482/613

Ca :H Low affinity Pi

+

Proline:Na+

2.A.20.1.X

2+

Ca2+:H+

Glycine betaine Ectosine/ glycine betaine/ pro-line Ectosine/ glycine betaine/ pro-line Di- or tripeptide Di- or tripeptide

2.A.21.2.1

2

BAC17261 BAC17284

2.A.19.1.1

2.A.17.1.1

2.A.17.1.1

2.A.15.X.X

2.A.15.1.2

2.A.15.1.1

2

2

2

2

BAC17948

BAC19713

1

2

2

2

BAC18233

2 2

BAC19474 BAC19576

2

2

BAC19027

1

2

BAC17776

1

BAB99996

BAB99727

BAB98285 (BetP) BAB99705 (EctP)

Peptides; nitrates; amino acids

Glycine; betaine; carnitine; choline; multiple organocations

Genomic Analyses of Transporter Proteins in C. glutamicum and C. efficiens 159

2.A.36

2.A.28

2.A.26

3 2

BAB98651 BAB98828

Na+/H+; Na+ or K+/H+

CPA1

3

BAB99720

Bile acids

BASS

1

2

2 2

BAB99944 BAC00421

BAB97599

2

Evidenced

BAB99988

Protein(s)b Cglc

Branched chain BAB99703 amino acids, (BrnQ) Leu, Ile, Val

Alanine, glycine

C4-dicarboxylates; acidic and neutral amino acids

Typical Substratesd

LIVCS

AGCS

The Alanine or Glycine: Cation Symporter (AGCS) Family The Branched Chain Amino Acid:Cation Symporter (LIVCS) Family The Bile Acid:Na+ Symporter (BASS) Family The Monovalent Cation: Proton Antiporter-1 (CPA1) Family

2.A.25

Abbrev.

The DAACS Dicarboxylate/A mino Acid:Cation (Na+ or H+) Symporter (DAACS) Family

Name of Family

2.A.23

Family TC#a

BAC19039

BAC19022

BAC18784

BAC19674

Protein(s)b Cefc

TABLE 8.3 (continued) Putative Transport Proteins Identified in C. glutamicum and C. efficiens

3

2

2

3

d

Evidence

2.A.28.X.X 2.A.36.X.X

2.A.28.X.X

2.A.26.1.3

2.A.25.1.2

2.A.23.1.X 2.A.23.X.X

2.A.23.1.3

Protein TC#

Branched chain amino acids

78% 238/304

63% 268/423

335 516

324

426

492

51% 250/487

Alanine:Na+

446

345

425

490

461

# of aae # of aac Cglc Cef

446 417

% Identity

Fumarate, D- and L-malate, succinate, succinamide, orotate, iticonate, mesaconate 77% 321/415

Probable Substratesd

160 Handbook of Corynebacterium glutamicum

The Hydroxy/Aromatic Amino Acid Permease (HAAAP) Family The Arsenite-Antimonite (ArsB) Efflux Family The Benzoate:H+ Symporter (BenE) Family The Divalent Anion:Na+ Symporter (DASS) Family

2.A.42

2.A.53 SulP

CHR

The Chromate Ion Transporter (CHR) Family The Sulfate Permease (SulP) Family

2.A.51

Di- and tricarboxylates; phosphate; sulfate

DASS

1

BAB99454 (AmtP) BAB99842

BAB98444

BAC17754

2

BAC17903

BAC19010

BAC19156

2

2

2

3

2

2 2

BAC19164 BAC18511 BAC18778

2

2

3

2

2

BAC17004

BAC19312

BAC16835

BAC19077

BAC18642

2

3

1

2

BAB97618

BAB98976 (Amt)

2

3

BAB99438

BAB99803

3

2

BAB99751 BAB97874

2

BAB99332

Chromate; sulfate (uptake or efflux) Sulfate; sulBAB98866 fate, bicarbonate; anions BAB98263

Ammonium

Benzoate

BenE

The Ammonium Amt Transporter (Amt) Family

Arsenite, antimonite

Hydroxy and aromatic amino acids

Nucleobases; urate

ArsB

HAAAP

NCS2

2.A.49

2.A.47

2.A.46

2.A.45

The Nucleobase: Cation Symporter-2 (NCS2) Family

2.A.40

2.A.53.X.X

2.A.53.X.X

2.A.53.3.1

2.A.51.1.X

2.A.49.1.3

2.A.49.1.2

2.A.47.X.X

2.A.47.3.X

2.A.46.1.1

2.A.42.X.X

2.A.40.3.X

2.A.40.1.1

77% 395/510

72% 293/403

75% 472/628

78% 337/429

Sulfate

Chromate

68% 335/492 60% 326/535 62% 358/573

47% 180/377

Ammonium/ 56% 249/442 methylammonium Ammonium 68% 299/438

Benzoate

Arsenite

Xanthine or Uric acid

Uracil

579

537

485

376

438

452

510

476

414

397

629

429

582

590

555

377

441

450 439

587

430

381

644

431

Genomic Analyses of Transporter Proteins in C. glutamicum and C. efficiens 161

2.A.63

2.A.59

Inorganic phosphate 2 2 2 3 3 3 3 3

BAB98903 BAB97655 BAC00123 BAC00124 BAC00125 BAC00126 BAC00128 BAC00127

K+ or Na+/H+

CPA3

3

Arsenite

BAC00138

BAC17040 BAC17038 BAC17037

BAC19382

BAC19380

BAC19379

BAC19378

BAC19377

BAC17685

3 3 3

3

3

3

3

2

2

2.A.63.1.X

2.A.63.1.X

2.A.63.1.X

2.A.63.1.X

2.A.63.1.X

2.A.59.1.1 2.A.63.1.X

2.A.59.1.1

2.A.58.1.2

Phosphate

449

249

160

437

91

147

556

163

Na+:H+ Na+:H+ Na+:H+

108/163 433/536 92/146 97/126

360 1019

392

126

66% 80% 63% 76%

68% 681/995

82% 306/370

388

85 547 151

127

169

592

163

1018

377

474

676

384

192

438

# of aae # of aac Cglc Cef

Na+:H+

Na+:H+

Na+:H+

Na+:H+

Arsenite

Arsenite

42% 165/387

22% 83/363

75% 331/436 68% 100/145 83% 181/217

% Identity

94 3

Fumarate, D- and L-malate, succinate, succinamide, orotate, iticonate and mesaconate

Probable Substratesd

BAB99710(R) BAC19392

2.A.56.X.X

2.A.56.1.1

Protein TC#

141

3

2

d

Evidence

BAB99712(M)

BAC17131

BAB99711(M)

3

BAC19051(M)

BAB99733(R)

BAC19049(M)

Protein(s)b Cefc

BAC19050(R)

3

Evidenced

BAB99732(M)

BAB99731(M)

Protein(s)b Cglc

ACR3

PNaS

The Phosphate:Na+ Symporter (PNaS) Family The Arsenical Resistance-3 (ACR3) Family The Monovalent Cation (K+ or Na+): Proton Antiporter-3 (CPA3) Family

2.A.58

Typical Substratesd C4-dicarboxylates; acidic amino acids; sugars (?)

Abbrev.

The Tripartite ATP- TRAP-T independent Periplasmic Transporter (TRAP-T) Family

Name of Family

2.A.56

Family TC#a

TABLE 8.3 (continued) Putative Transport Proteins Identified in C. glutamicum and C. efficiens

162 Handbook of Corynebacterium glutamicum

2.A.77

2.A.76

2.A.75

2.A.72

2.A.69

2.A.68

CadD

RhtB

BAB98655 (LysE)

Neutral amino BAB99737 acids and their deriva-tives BAB97539 BAC00050 BAB98300 Cd2+; cations

Basic amino acids

3 3 3

3

1

3 2

BAB99779 BAB98105

K+ (uptake)

KUP

LysE

3

BAB99707

Auxin (efflux)

3 2

AEC

Aminobenzoylglutamate

BAC00482 BAB97494

The p-Aminobenzoylglutamate Transporter (AbgT) Family The Auxin Efflux Carrier (AEC) Family The K+ Uptake Permease (KUP) Family The L-Lysine Exporter (LysE) Family The Resistance to Homoserine/Thre o-nine (RhtB) Family The Cadmium Resistance (CadD) Family

MVF AbgT

3

19552705

3

3

19552348

BAB97744

3

BAB98883

3

Drugs, dyes; nucleotides?

Proteins, mostly redox proteins

BAB99375

2.A.66

Tat

The Multidrug/ MOP Oligosaccharidyllipid/Polysaccharide (MOP) Flippase MATE Superfamily PST

The Twin Arginine Targeting (Tat) Family

2.A.64

BAC19055 BAC16955

BAC18167

BAC19098 BAC19201

BAC19031

BAC19741 BAC16915

BAC17188

BAC18685

BAC18430

BAC17990

BAC18492

3 3

2

3 3

3

3 2

3

3

3

3

3

2.A.76.X.X 2.A.76.X.X 2.A.77.X.X

2.A.76.1.2

2.A.75.1.1

2.A.69.X.X 2.A.72.1.1

2.A.69.X.X

2.A.66.4.1 2.A.68.1.1

2.A.66.2.X

2.A.66.1.X

2.A.64.1.1

2.A.64.1.1

2.A.64.1.1

Threonine

L-lysine

K+

Polysaccharides ? p-Aminobenzoylglutamate

Drugs

Proteins

Proteins

Proteins

60% 135/222

62% 145/233

44% 111/250

70% 202/286 71% 221/310

75% 391/517

63% 677/1069

57% 245/427

60% 191/316 49% 82/165 49% 52/105

226 207 207

223

236

310 624

309

1083 538

497

435

105

156

334

224 234

235

310 488

308

1259 563

404

458

103

165

313

Genomic Analyses of Transporter Proteins in C. glutamicum and C. efficiens 163

3.A. P-P-Bond Hydrolysis-driven Transporters 3.A.1 The ATP-binding ABC Cassette (ABC) Superfamily

2.A.80

The Threonine/ ThrE Serine Exporter (ThrE) Family The Tripartite TTT Transporter (TTT) Family

2.A.79

LIV-E

Abbrev.

The Branched Chain Amino Acid Exporter (LIV-E) Family

Name of Family

2.A.78

Family TC#a

All sorts of inorganic and organic molecules of small, intermediate and large sizes, from simple ions to macromolecules

Tricarboxylates

Thr, Ser

Leu, Ile, Val

Typical Substratesd

3 3

BAC00476 BAC00477

3 3 3

19554012(M) BAC00218(M) BAC00219(R)

1

1

BAB97651 (BrnF)

BAC00016 (ThrE)

1

Evidenced

BAB97652 (BrnE)

Protein(s)b Cglc

BAC19316

BAC19736

BAC19735

Protein(s)b Cefc

TABLE 8.3 (continued) Putative Transport Proteins Identified in C. glutamicum and C. efficiens

2

3

3

d

Evidence

2.A.80.1.1

2.A.80.1.1

2.A.80.1.1

2.A.79.1.1

2.A.78.X.X

2.A.78.X.X

2.A.78.1.2

2.A.78.1.2

Protein TC#

Tricarboxylates Tricarboxylates Tricarboxylates

Branched chain amino acids Branched chain amino acid Branched chain amino acid Branched chain amino acid Threonine/ Serine

Probable Substratesd

58% 279/475

65% 153/233

73% 84/115

34% 75/215

30% 30/100

% Identity

334

188

510

489

237

115

251

108

486

238

116

# of aae # of aac Cglc Cef

164 Handbook of Corynebacterium glutamicum

PAAT

CUT2

CUT1

BAB97609(C) BAB98117(M) BAB98118(M) BAB98120(R) BAB98121(C) BAB99852(M) BAB99853(M) BAB99854(R) BAB99856(C) BAB98776(M) BAB98777(M) BAB98778(R) BAB98779(C) BAB97423(R) BAB97424(M) BAB97425(C) BAB98646(M) BAB98645(C) BAB98647(R) BAB98648(M) BAB99343(C) (GluA) BAB99344(R) (GluB) BAB99345(M) (GluC) BAB99346(M) (GluD) BAB98723(C) BAB98724(M) BAB98725(R) 2

1

2

3

2

2

2 2

BAC18253(C) BAC18254(M) BAC18256(R) BAC18389(R) BAC19398(R) BAC19475(R)

3 3 3

2

3.A.1.3.X

88% 220/250 91% 218/239 66% 206/309

77% 211/271

BAC18657(M)

85% 206/242

172/321 235/296 237/278 349/436 331/394 231/322 264/340 197/253 48/195

53% 79% 85% 80% 84% 71% 77% 77% 24%

91% 209/228

Glutamate

179/302 236/278 261/314 323/434 279/334 67/276 63/258

59% 84% 83% 74% 83% 24% 24%

BAC18656(M)

3.A.1.3.9

3.A.1.2.X

3.A.1.2.X

3.A.1.1.X

3.A.1.1.X

3.A.1.1.X 3.A.1.1.X

85% 251/293

3

3

3

2

2

2 2

BAC18655(R)

BAC18654(C)

BAC17001(C) BAC17558(M) BAC17559(M) BAC17561(R) BAC17562(C) BAC17678(M) BAC17679(M) BAC17680(R) BAC17681(C) BAC18324(M) BAC18325(M) BAC18326(R) BAC18327(C) BAC16829(R) BAC16830(M) BAC16831(C) BAC18955(M)

250 316 334

273

228

295

304 278 344 424 332 304 281 443 376 301 278 438 408 327 341 253 324 524 314 123 242

254 240 348 292 368 178

316

249

294

269

306 278 348 434 334 296 526 420 393 309 278 441 402 322 346 253 313

Genomic Analyses of Transporter Proteins in C. glutamicum and C. efficiens 165

Family TC#a

Name of Family

PhoT

PepT

HAAT

Abbrev.

Typical Substratesd BAB98323(R) BAB98324(M) BAB98325(M) BAB98326(C) BAB98327(C) BAB99713(R) BAB99714(M) BAB99715(M) BAB99716 (C-C) BAB99383(R) BAB99384(M) BAB99385(M) BAB99386 (C-C) BAB99829 (C-C) BAB99830(M) BAB99831(M) BAB99832(R) BAB99770 BAC00046(R) BAC00047(M) BAC00048(C) BAC00049(C) BAB99965(C) BAB99966(M)

Protein(s)b Cglc

2

2 2

BAC19141(C)

2

BAC19274(C) BAC19275(M)

BAC19142(M) BAC19143(M) BAC19144(R) BAC17967(R)

BAC18694(R) BAC18695(M) BAC18696(M) BAC18697(C)

BAC17810(R) BAC17811(M) BAC17813(M) BAC17814(C) BAC17815(C) BAC19034(R) BAC19035(M) BAC19036(M) BAC19037(C)

Protein(s)b Cefc

3

2

2

Evidenced

TABLE 8.3 (continued) Putative Transport Proteins Identified in C. glutamicum and C. efficiens

2

3

2

2

2

d

Evidence

3.A.1.7.1

3.A.1.5.X 3.A.1.5.X

3.A.1.5.X

3.A.1.5.X

3.A.1.5.X

3.A.1.4.4

Protein TC#

Phosphate

Urea

Probable Substratesd

418/496 253/308 278/333 487/571

331/423 216/294 189/299 195/239 155/233 374/493 226/307 177/273 307/477

85% 259/302 78% 243/310

67% 199/294 62% 195/313 73% 388/525

81% 446/546

90% 82% 83% 85%

78% 73% 63% 81% 66% 75% 73% 64% 64%

% Identity

294 313 569 525 536 317 356 272 299 307

547

534 308 333 577

423 294 359 242 233 503 322 276 479

302 310

305 313 615 553

561

535 337 345 579

417 294 328 247 234 550 311 290 491

# of aae # of aac Cglc Cef

166 Handbook of Corynebacterium glutamicum

FeCT

QAT

PhnT

MolT

3

3

3(2)

2

2

BAB99853(C) BAB99854(R) BAB98851(M)

BAB98059(C) BAB98060(M) BAB98061(M) BAB98062(R) BAB98067(R) BAB98068(C) BAB98069(M) BAB98203(R) BAB98204(M) BAB98205(M) BAB98206(C) BAB99505(C) BAB99506(M)

3

2

BAB9967(M) BAB99968(R) BAB97606(M) BAB97607(R) BAB98852(M)

BAC16976(M) BAC16977(M) BAC16978(C) BAC17494(C) BAC17495(M) BAC17496(M) BAC17497(R) BAC17502(R) BAC17503(C) BAC17504(M) BAC17691(R) BAC17692(M) BAC17693(M) BAC17694(C) BAC18821(C) BAC18822(M)

BAC16975(R)

BAC19276(M) BAC19277(R) BAC16998(M) BAC16999(R)

2

3

2

2

3

2

3

3.A.1.14.X

3.A.1.14.X

3.A.1.14.X

3.A.1.14.X

3.A.1.9.X

3.A.1.9.1

3.A.1.8.1

Iron chelates 71% 49% 56% 66% Iron chelates 66% 65% 68% Iron chelates 74% 80% 76% 90% Iron chelates 64% 63%

Phosphonate/organ ophosphate ester Quaternary ammonium compounds

Phosphonate/organophosphate ester

Molybdate

75% 78% 75% 46%

191/266 153/308 177/316 230/346 184/277 168/258 233/339 251/338 261/326 231/303 228/251 172/267 202/318

254/337 251/319 175/233 121/259

269 353 332 350 325 260 339 338 333 314 251 266 346

268 350 282

338 375 241 268 268

222 232 276 272 340 332 348 341 264 335 384 469 371 251 271 348

303

355 373 283 265

Genomic Analyses of Transporter Proteins in C. glutamicum and C. efficiens 167

Family TC#a

Name of Family

NitT

MZT

Abbrev.

Typical Substratesd BAB99507(R) BAB99019(M) BAB99020(R) BAB99021(C) BAB97430(M) BAB97431(C) BAB97729(R) BAB97782(R) BAB97783(M) BAB97784(C) BAB97893(C) BAB97894(M) BAB97895(M) BAB98039(R) BAB98201(R) BAB98650(R) BAB99428(R) BAC00469(R) BAC00024(M) BAC00022(R) BAC00023(C) BAB97420(R) BAB97421(M) BAB97422(C) BAB98667(C) BAB98669(C)

Protein(s)b Cglc

3(2)

3

BAC18178(C) BAC18179(M)

2

3.A.1.16.X

3.A.1.15.X

3.A.1.14.X 3.A.1.14.X 3.A.1.14.X 3.A.1.14.X 3.A.1.14.X 3.A.1.15.X

3 3 3 3 2 3

3

3.A.1.14.X

2

BAC19324(M)

3.A.1.14.X 3.A.1.14.X

3.A.1.14.X

Protein TC#

3 3

2

d

Evidence

3.A.1.14.X

BAC18823(R) BAC19487(M) BAC19486(R) BAC19485(C)

Protein(s)b Cefc

2

3

Evidenced

TABLE 8.3 (continued) Putative Transport Proteins Identified in C. glutamicum and C. efficiens

Iron Iron Iron Iron Iron

chelates chelates chelates chelates chelates

Iron chelates

Iron chelates Iron chelates

66% 101/151

67% 121/178

165/269 194/346 155/308 152/249

% Identity

61% Iron chelates 56% 50% 61% Iron chelates

Probable Substratesd

329 358 319 251 326 259 315 359 356 271 264 345 345 332 306 338 312 319 185 314 230 318 283 219 166 89

238 261

290

343 366 330 258

# of aae # of aac Cglc Cef

168 Handbook of Corynebacterium glutamicum

DrugE1

Drug RA1/RA2

Drug RA1

DrugE2/ Pep4E DrugE3

BAB98825 (C-M) BAB98826 (C-M) BAB98917 (C-M) BAB98345 (C-M) BAB98346 (C-M) BAB98813 (C-C) BAB98305 (C-C) 2

BAC16827 (C-C) BAC17784 (C-C)

2

2

3.A.1.119.X

3.A.1.119.X

3.A.1.119.X

3.A.1.117/ 123.X 3.A.1.119.2

3.A.1.117.X

3.A.1.120/ 121

3.A.1.120.X

Tetracycline/oxytetracycline/oxacillin

3.A.1.105.X Daunorubicin; doxorubicin or oleandomycin

3.A.1.119.X

2

Aromatic sulfonate

3.A.1.104.X Teichoic acid

2

2

2

2

2

2

2

3.A.1.17.2

3.A.1.119.X

BAC18374 (C-M) BAC18375 (C-M) BAC18456 (C-M)

BAC18457 (C-M)

BAC19007 (C-M)

BAC16988(M)

BAC16987(C)

BAC18180(R)

2

2

2

3

2

2

3

2(3)

BAB97594(M) BAB98960(C)

Tae

BAB98961(M) BAB98357 (C-M) BAB98358 (C-M) BAB98918 (C-M)

2

BAB98615(C) BAB98616(R) BAB97593(C)

DrugE2

2

BAB98670(R) BAB98614(M)

TauT

84% 519/615

36% 194/530

62% 368/593

59% 349/582

68% 341/500

61% 327/350

28% 178/631

91% 245/268

96% 243/252

56% 144/257

611

510

578

480

599

589

518

529

577

268 656

268 340

243 319 263

294 256

615

590

612

586

537

570

621

323

274

327

Genomic Analyses of Transporter Proteins in C. glutamicum and C. efficiens 169

Family TC#a

Name of Family

MDR

Drug RA1

Abbrev.

Typical Substratesd

BAC17355(M) BAC18135(C) BAC17439(M) BAC17440(C) BAC17742(C) BAC17829(C) BAC18164(M)

2 2 3

2 2

3

BAB97819(C) BAB97820(M) BAB97821(R) BAB97925 (C-C) BAB97926(M) BAB98620(C) BAB98028(M) BAB98029(C) BAB98030(R) BAB98249(C) BAB98339(C) BAB98340(M) 3

3

2

2

2

2

BAC17911 (C-C) BAC18488 (C-C) BAC18845 (C-C) BAC19159 (C-C) BAC16827 (C-C) BAC18001 (C-C) BAC19685(M) BAC18163(C) BAC19684(C) BAC17256(C) BAC17257(M) BAC17258(R) BAC17354(C)

Protein(s)b Cefc

3

Evidenced

BAB99411(M) BAB99412(C)

BAB98454 (C-C) BAB99847 (C-C) BAB99534 (C-C) BAB98947 (C-C) BAB98813 (C-C)

Protein(s)b Cglc

TABLE 8.3 (continued) Putative Transport Proteins Identified in C. glutamicum and C. efficiens

2(3) 2 2(3)

2 3?

2(3)

2(3) 2 3

2

2

2

2

2

2

d

Evidence

3.A.1.X.X 3.A.1.X.X

3.A.1.X.X 3.A.1.X.X 3.A.1.X.X

3.A.1.X.X

3.A.1.X.X

3.A.1.X.X

3.A.1.120.X

3.A.1.120.X

3.A.1.120/ 121 3.A.1.120.X

3.A.1.120.X

Protein TC#

Probable Substratesd

253/371 236/274 170/222 290/347

219/349 359/529 277/354 438/568

78% 171/218 72% 216/297 27% 17/61

68% 86% 76% 83%

62% 67% 78% 77%

26% 29/111 35% 77/217

36% 194/530

85% 477/556

68% 373/548

87% 473/543

66% 367/550

% Identity

382 293 222 360 299 222 306 277

350 548 353 554

526 285

510

556

550

543

547

228 325 277

431 287 225 359

532 302 297 349 534 379 607

1242

590

556

544

543

565

# of aae # of aac Cglc Cef

170 Handbook of Corynebacterium glutamicum

BAB98472 (C-C) BAB98473(M) BAB98523 (C-M) BAB98540 (C-M) BAB98541 (C-M) BAB98848(C) BAB98849(M) BAB98955(C) BAB98956(M) BAB98957(M) BAB99352(C) BAB99353(M) BAB99449(M) BAB99450(C) BAB99470 (C-M) BAB99471 (C-M) BAB99642(C) BAB99643(M) BAB99644(R) BAB99840(C) BAB99945(M) BAB99946(C) BAB97799(M) BAC00140(M) BAC00141(C) BAC00144(R) BAC00432(C) BAC18662(C) BAC18663(M)

2(3)

2

3 2

2 3

3

2

3

2

3

2

BAC19684(C),

3 3 2

BAC18956(R) BAC19154(C) BAC19246(M) BAC19247(C) BAC17236(M) BAC19393(M) BAC19394(C)

2

3 2(3)

2

2 3

3

2 2(3) 2

2

2

2

2(3)

BAC18954(C)

BAC18791(C)

BAC18773(C) BAC18790(C)

BAC17946(M) BAC18737 (C-M) BAC18060 (C-M) BAC18061 (C-M) BAC18847(C) BAC17741(M) BAC18494(C)

3 2 3

BAC17945(C)

2

3.A.1.X.X

3.A.1.X.X 3.A.1.X.X

3.A.1.X.X 3.A.1.X.X

3.A.1.X.X

3.A.1.X.X

3.A.1.X.X

3.A.1.X.X

3.A.1.X.X

3.A.1.X.X

3.A.1.X.X

3.A.1.X.X

3.A.1.X.X 3.A.1.X.X

3.A.1.X.X

241/314 174/258 511/850 192/235 516/862 173/251 215/305 71% 210/294

76% 67% 60% 81% 59% 68% 70%

79% 209/263

50% 255/509

70% 202/286 51% 284/550

79% 181/227 66% 135/203

32% 65/197 30% 16/52 93% 235/252

64% 323/500

51% 267/516

65% 164/251 30% 144/466

66% 301/453

263 296 313 264 847 236 862 256 312 344 302

510

247 247 252 392 481 233 203 289 292 548

511

518

251 1247

446

297

331 271 854 236 880 273 312

267

508

337 548

232 203

304 852 252

510

522

251 581

456

Genomic Analyses of Transporter Proteins in C. glutamicum and C. efficiens 171

3.A.2

Family TC#a

The H+- or Na+translocating Ftype, V-type and A-type ATPase (F-ATPase) Superfamily

Name of Family

F-ATPase

Abbrev.

H+; Na+

Typical Substratesd

BAB98600 (C chain) BAB98601 (B chain)

BAB99404(M) BAB99405(C) BAB97924(M) BAB98026(C) BAB98680(R) BAB99691 (C-M) BAB97628 (C-M) BAB99080(C) BAB99081(M) BAC00225(C) BAC00237(C) BAC00387(M) BAC00388(C) BAB99080(C) BAB99081(M) BAB98599 (A chain)

Protein(s)b Cglc

3(2)

BAC18120 (C chain) BAC18121 (B chain)

BAC18119 (A chain)

3(2)

3

3.A.1.X.X

3.A.2.1.2

3.A.1.X.X

BAC16836(R)

3

2(3)

2(3)

3.A.1.X.X

3.A.1.X.X 3.A.1.X.X 3.A.1.X.X 3.A.1.X.X

3.A.1.X.X

Protein TC#

3.A.1.X.X 3.A.1.X.X 3.A.1.X.X

BAC16836(R)

3

3

3 2 3

d

Evidence

3 3 2

BAC19460(C)

3 2 3 2

Protein(s)b Cefc

BAC17300(C) BAC18499(C) BAC19322(R) BAC19323(C) BAC19459(C)

3

Evidenced

TABLE 8.3 (continued) Putative Transport Proteins Identified in C. glutamicum and C. efficiens

Na+

Probable Substratesd

80 188

74% 141/190

199 333 356 127 421 230 199 333 270

479

319 230 268 239 354 621

190

81

310

373

373

242

213 337 302 229 239

# of aae # of aac Cglc Cef

67% 54/80

85% 231/270

19% 23/116

36% 54/149

% Identity

172 Handbook of Corynebacterium glutamicum

3.A.3

The P-type ATPase P-ATPase (P-ATPase) Superfamily Cations (uptake and/or efflux): Na+, K+; H+, K+; Ca2+, K+; Na+; H+; K+; Ca+; Ca2+, Mn2+; Mg2+; Mn2+; Cu2+; Cu+, Ag+; Ag+; Zn2+, Cd2+, Co2+, Ni2+, Pb2+ (some systems may be specific for one or only a few of these metal cations); phospholipids (flipping) 2

2 2 2 2 2 2

BAB98602 (delta) BAB98603 (alpha) BAB98604 (gamma) BAB98605 (beta) BAB98606 (epsilon) BAB98600 (C chain) BAB98939

BAB97779 BAB97827 BAB97875 BAC00356 BAB98569 BAC00368

BAC17217 BAC17267 BAC18750 BAC17091 BAC16820 BAC16887

BAC18122 (delta) BAC18123 (alpha) BAC18124 (gamma) BAC18125 (beta) BAC18126 (epsilon) BAC18120 (C chain) BAC18480

2 2 2 2 2 2

2

3.A.3.5.X 3.A.3.5.X 3.A.3.5.X 3.A.3.5.X 3.A.3.6.X 3.A.3.6.X

3.A.3.2.4

80

67% 54/80

529/750 494/841 381/654 453/655 191/552 460/541

70% 58% 58% 69% 34% 85%

or Ag+ or Ag+ or Ag+ or Ag+

124

91% 113/124

Cu+ Cu+ Cu+ Cu+

483

88% 429/483

755 848 650 739 625 625

892

325

81% 325/326

62% 533/854

547

93% 512/546

Ca2+

271

80% 217/271

757 900 645 659 650 578

976

81

124

481

326

557

274

Genomic Analyses of Transporter Proteins in C. glutamicum and C. efficiens 173

The Type IV (Conjugal DNAProtein Transfer or VirB) Secretory Pathway (IVSP) Family The Bacterial Competencerelated DNA Transformation Transporter (DNA-T) Family

3.A.7

3.A.11

The Type II (General) Secretory Pathway (IISP) Family

Name of Family

3.A.5

Family TC#a

DNA-T

IVSP

IISP

Abbrev.

BAB97948 (SecY) BAB98153 (SecA) BAB98195 (FtsE) BAB98833 (SecA) BAB99451 (Fth) BAB99455 (FtsY) BAB98977 (SecG)

BAB97867 (SecE)

Protein(s)b Cglc

Single-stranded BAB99741 DNA

Proteins, protein-DNA complexes

Proteins

Typical Substratesd

3

3

Evidenced

BAC19059

BAC17374 (SecY) BAC17584 (SecA) BAC17622 (FtsE) BAC18382 (SecA) BAC18775 (Fth) BAC18780 (FtsY) BAC18512 (SecG) BAC17108

BAC17295 (SecE)

Protein(s)b Cefc

TABLE 8.3 (continued) Putative Transport Proteins Identified in C. glutamicum and C. efficiens

3

2

2

d

Evidence

3.A.11.X.X

3.A.5.1.1

Protein TC#

DNA

Proteins

Probable Substratesd

229 763 547 510 77

92% 212/229 77% 590/763 79% 436/547 76% 231/302 64% 50/77

194

845

86% 730/842

45% 96/213

440

111

255

602

77

636

540

775

229

845

440

109

# of aae # of aac Cglc Cef

87% 386/440

66% 75/112

% Identity

174 Handbook of Corynebacterium glutamicum

The Proton-translocating Cytochrome Oxidase (COX) Superfamily

3.D.4

4.A. Phosphotransferase Systems 4.A.1 The PTS Glucose- Glc Glucoside (Glc) Family

COX

The Proton-transQCR locating Quinol: Cytochrome c Reductase (QCR)

3.D.3

The Septal DNA S-DNA-T Translocator (S-DNA-T) Family 3.D. Oxidoreduction-driven Active Transporters 3.D.1 The Proton-transNDH locating NADH Dehydrogenase (NDH) Family

3.A.12

2 3

19552375 BAB99585 BAB99588 BAB99916

2

2

2

19552374

Glucose; NBAB98753 acetylglucosamine; α- and β-glucosides BAC00036 (i.e., maltose; trehalose; sucrose; arbutin; arbutin, cellobiose, salicin)

H+ (efflux)

BAB99584

3

BAB99583

3 2

BAB97662 BAB98858

3

3

Na+; H+ (efflux) BAB97660

BAB99582

2

BAB99361

H+ (efflux)

2

BAB97970

DNA, DNAprotein complexes

BAC18268

BAC18895 BAC18897 BAC19228

BAC18894

BAC18893

BAC17036 BAC18403 BAC18207 BAC18208 BAC18892

BAC18206

BAC18671

BAC17390

2

3

2

3

2 2 2 3 2

3

2

2

4.A.1.2.X

4.A.1.2.X

3.D.4.X.X

3.D.4.5.1

3.D.4.5.1

3.D.3.X.X

3.D.3.X.X

3.D.1.X.X 3.D.1.X.X

3.D.1.X.X

3.A.12.X.X

3.A.12.1.X

Sucrose

Glucose

H+

DNA

DNA

78% 538/686

93% 192/205 76% 273/359 92% 520/565

82% 242/295

82% 336/408

84% 451/536

32% 278/85 85% 396/464

22% 101/442

66% 607/907

71% 809/1139

661

683

205 359 584

513

333

295

408

539

962 467

510

921

1189

704

217 368 580

296

427

949 471 516 603 541

540

984

1269

Genomic Analyses of Transporter Proteins in C. glutamicum and C. efficiens 175

The PTS FructoseMannitol (Fru) Family The PTS L-Ascorbate (L-Asc) Family

Name of Family

L-Asc

Fru

Abbrev.

8.A. Auxiliary Transport Proteins 8.A.3 The Cytoplasmic MPA1 Membrane-Periplasmic Auxiliary-1 (MPA1) Protein with Cytoplasmic (C) Domain (MPA1-C or MPA1+C) Family

5.A. Transmembrane Electron Transfer Carriers 5.A.1 The Disulfide Bond DsbD Oxidoreductase D (DsbD) Family 5.A.3 The Prokaryotic PMO Molybdopterincontaining Oxidoreductase (PMO) Family

4.A.7

4.A.2

Family TC#a

Complex polysaccharides BAB97737

BAB98581 NarH BAB98582 NarG BAB97922

BAB98579 NarI

2e-

3

3

2(3)

3

3

BAC00431 (IIA) BAB97833

2

2

Evidenced

BAC00430 (IIC)

BAB99329

Protein(s)b Cglc

2e-

L-ascorbate

Fructose; mannitol

Typical Substratesd

BAC17167

BAC18105 NarH BAC18106 NarG BAC17350

BAC18103 NarI

BAC17273

BAC18639

Protein(s)b Cefc

TABLE 8.3 (continued) Putative Transport Proteins Identified in C. glutamicum and C. efficiens

2

2

2

3

2

d

Evidence

8.A.3.X.X

5.A.3.X.X

5.A.3.1.1.

5.A.1.2.1

4.A.7.1.X

4.A.7.1.X

4.A.2.1.1

Protein TC#

2e-

2e-

L-ascorbate

L-ascorbate

Fructose

Probable Substratesd

75% 327/435

478

711

1248

85% 1069/1252 84% 598/707

531

259

264

270

513

612

465

765

1249

533

259

283

688

# of aae # of aac Cglc Cef

89% 477/532

81% 212/259

65% 173/264

77% 463/597

% Identity

176 Handbook of Corynebacterium glutamicum

The Putative Permease (PerM) Family The Hly III (Hly III) Family

9.B.22

9.B.30

The Putative Fatty Acid Transporter (FAT) Family

9.B.17

Hly III

PerM

FAT

The Phosphotrans- EI ferase System Enzyme I (EI) Family 8.A.8 The Phosphotrans- HPr ferase System HPr (HPr) Family 9.A. Transporters of Unknown Classification 9.A.4 The Nicotinamide PnuC Mononucleotide (NMN) Uptake Permease (PnuC) Family MgtE 9.A.19 The Mg2+ Transporter-E (MgtE) Family 9.B. Putative Uncharacterized Transporters 9.B.3 The Putative MPE Bacterial Murein Precursor Exporter (MPE) Family

8.A.7

2

BAB97617

BAB97437

BAB99552

Mg2+, Co2+ (uptake)

Murein precursor

Ions? small molecules?

?

Fatty acids?

2

BAB97457

Nicotinamide mononucleotide (uptake)

2

3 3 3 3 2 3 3 3

BAB97793

BAB97843 BAB98591 BAB99689 BAC00266 BAB97677 BAB98421 BAC00195 BAB99980

2

3

3

BAB99330

Sugars

2

BAB99326 (ptsI)

Sugars

BAC19289

BAC17891

BAC19587 BAC18112 BAC19005 BAC19516

BAC17231

BAC18864

3

3

2 2 2 2

2

2

2

3

BAC17994 BAC16846

2

3

3

2

BAC16833

BAC16891

BAC18640

BAC18636

9.B.22.1.X 9.B.30.1.1

9.B.17.1.X 9.B.17.1.X 9.B.17.1.X 9.B.17.1.X 9.B.17.1.X 9.B.22.1.X

9.B.17.1.4

9.B.3.1.X

9.B.3.1.X

9.A.19.X.X

9.A.4.X.X

8.A.8.1.1

8.A.7.1.1

2+

48% 212/434

76% 165/217

62% 56/89

77% 421/542

Fatty Fatty Fatty Fatty Fatty

acids acids acids acids acids

75/298 464/575 464/613 504/593

78% 179/227

56% 249/442

25% 80% 75% 84%

Lipid-linked 61% 311/509 murein precursor Fatty acids 87% 492/565

Lipid-linked 72% 267/366 murein precursor

Mg , Co

2+

Mg2+, Co2+

NMN

Sugars

Sugars

444 247

441 575 615 596 512 440

568

550

381

450

230

89

568

260

488

980 581 682 617

579

560

452

431

473

303

112

570

Genomic Analyses of Transporter Proteins in C. glutamicum and C. efficiens 177

The Putative Vectorial Glycosyl Polymerization (VGP) Family The HlyC/CorC (HCC) Family

Name of Family

HCC

VGP

Abbrev.

Ions?

Polysaccharides (export)

Typical Substratesd

3 3 3 3 3

BAB98586 BAB98587 BAB98842 BAB99679

3

Evidenced

BAB98841

BAB97520

Protein(s)b Cglc

BAC18740 BAC18739 BAC18395 BAC18995 BAC17014 BAC18394

Protein(s)b Cefc

3 3 2 2 2 3

d

Evidence

9.B.37.X.X 9.B.37.X.X 9.B.37.X.X 9.B.37.X.X

9.B.37.2.1

9.B.32.1.X

Protein TC# Polysaccharides

Probable Substratesd

62% 64% 79% 75%

211/339 281/436 366/460 327/431

% Identity

336 467 460 440

354

487

348 469 460 445 445 384

# of aae # of aac Cglc Cef

c

b

TC#, number of the family according to the transporter classification system; for more detailed information about the TC system, see our website at www-biology.ucsd.edu/~msaier/transport/ Protein components of a single system are separated by commas; distinct systems, when presented on a single line, are separated by semicolons. Cgl, Corynebacterium glutamicum; Cef, Corynebacterium efficiens. d Substrates of single transporters within a family are separated by commas; substrates transported by different protein members of the family are separated by semicolons. d Evidence: 1, certain, based on direct experimental data; 2, probable, based on close sequence similarity; 3, possible, based on distant sequence similarities. e aa, amino acids. Corynebacterium glutamicum: 35 N/A. Corynebacterium efficiens: 29 N/A.

a

9.B.37

9.B.32

Family TC#a

TABLE 8.3 (continued) Putative Transport Proteins Identified in C. glutamicum and C. efficiens

178 Handbook of Corynebacterium glutamicum

Genomic Analyses of Transporter Proteins in C. glutamicum and C. efficiens

179

Porin PorA Outer lipid layer

Primary Transporter

Group Translocation Glucose

GluABCD

IICBAGluc

Secondary Transporter BrnQ

Channel

MscL Cytoplasmic membrane

ATP ADP Glutamate

PEP Pyruvate Glucose-P

Valine

FIGURE 8.1 Types of transporters found in Corynebacteria. In the outer member (OM), the PorA porin is found in Cgl but not in Cef (see Section 8.7). In the lower panel, the representative transporters depicted are present in both Cgl and Cef. These are (from left to right): (1) the ABC glutamate transporter, GluABCD (TC 3.A.1.3), (2) the glucose phosphotransferase system (PTS), IIBCAGlc (TC 4.A.1.1.1), (3) the BrnQ branched chain amino acid efflux system, and (4) the MscL channel (TC 1.A.22.1.1) involved in osmotic stress adaptation. Single-headed arrows indicate directionality in an energized process while double-headed arrows indicate free movement (diffusion limiting) in an energy-independent process.

for protection against osmotic stress [4,16,19]. Interestingly, Nottebrock et al. [16] have provided evidence that Cgl has a mechanosensitive channel activity not attributable to either the MscL or the MscS homolog. The only recognizable protein that might serve this function is the VIC family member. However, the activity Nottebrock et al. detected could also be due to the presence of a novel type of channel protein not yet recognized in any organism. Both organisms also exhibit two paralogs of the Hsp70 family of chaperones that have been shown to be capable of transmembrane channel formation [1]. However, only Cef has a UAC (urea/amide) channel [25] while only Cgl has PorA, the major outer membrane porin of this organism [14] (see Figure 8.1). PorA is presumed to be a β-barrel-type porin (TC subclass 1.B). Both Cef and Cgl possess two anionselective porins of the nonhomologous PorB family (TC #1.B.41) [5a].

8.8 SECONDARY CARRIERS By far the largest family of secondary carriers is the major facilitator superfamily (MFS). Cgl has 45 MFS carriers and Cef has 38. As shown in Table 8.3, over half of these MFS permeases are putative drug/amphiphile/hydrophobe transporters of MFS subfamilies DHA1, 2, and 3 [4]. Some of these are likely to serve as lipid exporters, but others undoubtedly play primary roles in defense, in toxic substance export, and in metabolite export.

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Handbook of Corynebacterium glutamicum

Two sugar transporters (SP subfamily), several metabolite transporters (MHS subfamily), and either one or two nitrate/nitrite transporters (NNP subfamily) of the MFS allow uptake of essential nutrients. Additionally, one anion transporter (ACS subfamily), several aromatic acid transporters (AAHS subfamily), and two putative cyanate transporters (CP subfamily) were identified. One putative polyol transporter was found in Cgl but not in Cef. Two putative glycoside transporters of the GPH family, distantly related to the MFS, were found in Cef but not in Cgl. Two of the subfamilies in the APC superfamily are represented in both organisms. These are predicted to transport zwitterionic and basic amino acids. The CDF family of heavy-metal divalent-cation transporters is also represented, with two members in Cgl and one in Cef. A pair of SecD/F proteins in the RND superfamily together facilitates protein secretion via the Sec pathway (TC 3.A.5; see next section). Moreover, a single member of the Oxa1 family (TC 2.A.9) is found in both Cgl and Cef. Bacterial Oxa1 family members facilitate insertion of proteins into the cytoplasm membrane either together with or independently of the Sec system, depending on the substrate protein to be transported. Three or four RND-type systems that probably function in drug or lipid export are also present. Finally, three or four members of the DMT superfamily are found in both organisms. These proteins may catalyze export of various metabolites and drugs. A single putative gluconate uptake system (GntP family) was found in both corynebacteria, but a putative 2-keto-3-deoxygluconate uptake permease is present in Cef but not in Cgl. Additionally, one member of the CitMHS (citrate uptake) family and two members each of the BCCT (organocations) and POT (peptide) families are found in both species. Several putative acidic amino acid/dicarboxylate uptake systems of the SSS, NSS, DAACS, AGCS, LIVCS, and HAAP families were found. Specificity could be assigned to several of these transporters. Two orthologous NCS2 family members were found in the two species; these probably transport pyrimidines and purines, respectively. All of these systems most likely function in nutrient uptake, providing sources of carbon and nitrogen. One and two putative Ca2+:H+ antiporters are present in Cgl and Cef, respectively, and one putative phosphate uptake permease of the Pit family was found in each organism. A single monovalent-cation exchanger of the CPA1 family was identified in Cgl but not in Cef, while a single arsenite efflux system of the ArsB family was identified in Cef but not in Cgl. These cation and anion facilitators probably function primarily in the maintenance of ionic homeostasis, but they may also play a secondary role in adaptation to various types of stress. Proteins of the next nine families represented in Table 8.3 are probably all involved in organic and inorganic cation and anion transport. These include a benzoate uptake permease; sulfate, chromate, phosphate, and arsenite transporters; two ammonia/ammonium transport systems; one or two tripartite TRAP-T family members of uncertain specificities; and either five (Cgl) or seven (Cef) components of putative multicomponent, orthologous, monovalent-cation antiporters of the CPA3 family. Most of these transporters probably function in nutrient acquisition, except for the CPA3 monovalent-cation antiporters, which presumably function in ionic homeostatic control.

Genomic Analyses of Transporter Proteins in C. glutamicum and C. efficiens

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A single TatC homolog of the twin arginine targeting (Tat) family is found in each corynebacterial species as well as a pair of TatA paralogs. Streptomyces coelicolor has the same combination of Tat protein constituents [27]. These systems function in the secretion of a subset of extracellular proteins including redox enzymes in Gram-negative bacteria [27]. Genome analyses of the leader sequences of potential secretory proteins should reveal which are substrates of the Tat system and which are exported via the Sec system. The multidrug/oligosaccharidyl-lipid/polysaccharide (MOP) superfamily is represented by some members that probably export drugs and others that export polysaccharides. Additional families of efflux pumps listed in Table 8.3 probably export amino acids and their derivatives. A member of the tripartite tricarboxylate uptake transporter (TTT) family is found in Cgl but not in Cef. In sum, it is clear that the two corynebacteria analyzed exhibit large numbers of secondary carrier orthologs, but several systems are specific to one or the other species.

8.9 PRIMARY ACTIVE TRANSPORTERS The vast majority of protein constituents of primary active transporters are members of the ABC superfamily: 153 proteins in Cgl and 133 proteins in Cef belong to this superfamily (see Figure 8.1). Since a single system can consist of one or two ATPhydrolyzing subunits, one or two membrane subunits, and anywhere from zero to several extracytoplasmic receptors per system, it is not possible to estimate accurately the number of intact ABC transporters present in each organism. This is especially true of ABC transporters, because the constituents of a system are not always encoded within a single operon. ABC transport systems can function in either the uptake or the efflux of a huge variety of substrates, including macromolecules. Careful examination of Table 8.3 reveals some probable substrates of these systems, but the specificities of most of them could not be reliably estimated. Each species of Corynebacterium has a single multicomponent F-type ATPase for the interconversion of chemical and chemiosmotic energy. Each of these bacteria also possesses seven paralogous cation transporting P-type ATPases. Complete multicomponent general protein secretory (Sec) systems (TC 3.A.5) were found in both Cgl and Cef, and these systems undoubtedly serve as the primary protein export system in both bacteria [5]. An ATPase homologous to those of type-IV conjugation systems was identified in Cef but not in Cgl, but since the other constituents of these multicomponent systems were not found, it can be presumed that this ATPase serves an unrelated function. Potential DNA translocation proteins of the DNA-T and S-DNA-T families were also identified (Table 8.3), but assignment of their specific functional roles must await experimental studies. Na+-transporting carboxylate decarboxylases (TC 3.B.1) are multicomponent systems in which the β subunit catalyzes Na+ export in response to cytoplasmic substrate decarboxylation catalyzed by the α subunit. These systems minimally require the presence of α, β, and γ subunits [7]. While several α-subunit homologs were identified, no β- or γ-subunit homologs were found. Cytoplasmic decarboxylation/carboxylation reactions catalyzed by α-subunit homologs in corynebacteria are therefore probably not coupled to Na+ export [2].

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Handbook of Corynebacterium glutamicum

8.10 PROTON-PUMPING ELECTRON CARRIERS Both Corynebacterium species have homologs of specific subunits of the NADH dehydrogenase (NDH) (see Table 8.3), but the complete system is absent. The homologs listed may be components of a monovalent-cation antiporter (TC 2.A.63). This suggests there is no proton pumping coupled to NADH oxidation via NADH dehydrogenase. However, both the quinol:cytochrome c reductase (QCR) and two members of the cytochrome oxidase (COX) family were found in Cgl, and one of the COX family members was also present in Cef. The cytochrome bd complex was found only in Cgl, not in Cef. These bacteria therefore have complete electron transfer chains for oxidizing various substrates using molecular oxygen as electron acceptor. These electron carrier complexes have the potential to generate an ionmotive force as a primary source of energy. We suggest that these coupled systems probably function together under aerobic conditions.

8.11 GROUP TRANSLOCATORS The complete phosphoenolpyruvate:sugar phosphotransferase system (PTS) is present in the two corynebacteria analyzed. Both species have at least one glucose (Glc)-type PTS permease (see Figure 8.1), and both have a single fructose (Fru)type PTS permease. Cgl but not Cef has a second glucose-type system as well as an L-ascorbate (L-Asc)-type system [29]. The sugar specificities of a glucose-specific permease and a β-glucoside–specific PTS permease in Cgl (both of the Glc type), as well as a fructose-specific system (Fru type), have been determined [11,18].

8.12 TRANSMEMBRANE ELECTRON FLOW CARRIERS Corynebacteria have several systems that catalyze transmembrane electron flow (TC categories 5A and 5B). Each organism analyzed has a single disulfide bond oxidoreductase D homolog as well as four members of the molybopterin-containing oxidoreductase family. Other families of transmembrane electron flow carriers may also be present in these corynebacteria, but we could not readily identify them because TC categories 5A and 5B are not yet complete [4].

8.13 POORLY DEFINED TRANSPORTERS Among the poorly characterized permeases of TC class 9.A, both corynebacteria analyzed have a putative nicotinamide mononucleotide (NMN) uptake permease. They have either one (Cgl) or two (Cef) putative Mg2+/Co2+ uptake permeases of the MgtE family. Several putative permeases of TC class 9.B were also identified (Table 8.3), but their functions are poorly defined.

8.14 PERSPECTIVES AND CONCLUSIONS We have analyzed transporters in two corynebacteria, C. glutamicum (Cgl) and C. efficiens (Cef). The vast majority of transport proteins found in either of these

Genomic Analyses of Transporter Proteins in C. glutamicum and C. efficiens

183

organisms is found in both, but there are some notable exceptions. These organisms possess few channel proteins, but of them, the outer membrane porin, PorA, known to be the major outer membrane porin in this organism [14], is not encoded within the genome of Cef. Both Cef and Ggl possess two porins of the PorB family (TC #1.B.41) [5a]. Cgl lacks a urea/amide channel protein found in the cytoplasmic membrane of Cef. Cef presumably utilizes this system for the metabolism of urea and short-chain aliphatic amides such as acetamide [24]. Regarding secondary carriers for sugars, corynebacteria seem to have few such systems relative to other Gram-positive bacteria such as Streptomyces, Bacillus, and most lactic acid bacteria. Thus, Cgl has only three MFS carbohydrate transporters: two in the sugar porter family and one in the polyol porter family. Surprisingly, none of these carriers were found in Cef. On the other hand, Cef has two putative glycoside transporters of the GPH family that are lacking in Cgl. Both organisms also have a putative gluconate permease, but only Cef has a member of the 2-keto-3-deoxygluconate transporter (KDGT) family. Both organisms also have several ABC uptake transporters specific for monosaccharides and small oligosaccharides of the CUT1 and CUT2 subfamilies. They also possess complete phosphotransferase systems although Cgl can probably transport glucosides and L-ascorbate, which Cef cannot, in addition to glucose and fructose, which both can transport (Table 8.3) [18]. Interestingly, C. diphtheriae has the glucose, fructose, and putative L-ascorbate PTS permeases but lacks the glucoside system [17,18]. It appears that glucose and fructose are utilized exclusively via the PTS [18], and based on growth studies it is possible that the glucoside PTS permease transports sucrose and maltose. Both of these sugars could not be utilized by a ptsI mutant lacking Enzyme I of the PTS but could be utilized by a double mutant lacking both the fructose and glucose PTS permeases [18]. The capacity of corynebacteria to transport carboxylic acids and their derivatives as sources of carbon appears to be fairly extensive. Thus, three or four families of secondary dicarboxylate carriers (MFS, DAACS, DASS, and TRAP-T) are present in both species of corynebacteria studied. Both possess a member of the citrate transporting CitMHS family, and Cgl but not Cef also has a putative tricarboxylate carrier of the TTT family [26]. Our genome analyses revealed a huge number of transporters that are probably specific for amino acids and their derivatives. Thus, for the uptake of amino acids, six families of secondary carriers were represented (MHS of the MFS, SSS, NSS, AGCS, LIVCS, and HAAAP), and members of two ABC families with this specificity (PAAT and HAAT) were found. For the uptake of peptides, three potential families of secondary carriers (POT, AbgT, and MPE) as well as one ABC family (PepT) were represented. Finally, for amino acid efflux, members of six potential families were identified (DMT, AEC, LysE, RhtB, LIV-E, and ThrE). Several of these families were first defined by studies conducted with Cgl. It seems clear that the transport and metabolism of amino acids and their derivatives is exceptionally important to the lifestyles of these bacteria [3,13]. Nevertheless, nonorthologous occurrences between Cgl and Cef with respect to possible amino acid transporters can be found in the APC, DAACS, HAAAP, DASS, RhtB, and LIV-E families (see Table 8.3). These observations suggest major differences in amino acid metabolic capabilities in these two organisms.

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Handbook of Corynebacterium glutamicum

Our analyses revealed a large number of potential drug/hydrophobe/amphiphile export systems. Many of these belong to the DHA1, 2, and 3 families of the MFS. In fact, of the nearly 50 MFS paralogs identified in these two bacteria, six were in the DHA1 family, fourteen were in the DHA2 family, and one was in the DHA3 family. Thus, over 40% of all MFS permeases in these bacteria are involved in the export of hydrophobic and amphipathic substances. While a few of these efflux pumps may be involved in sugar export (Table 8.3) [20], it is possible that some export amino acids and their derivatives, particularly those of a hydrophobic nature. It should be noted, however, that this has not yet been established for any member of the three DHA families in the MFS. Other families including transporters that probably export hydrophobic substances include the HAE2 family in the RND superfamily, which may be involved in lipid export [6], and the DME and RarD families of the DMT superfamily, which may be concerned with drug export. Members of the MATE family within the MOP superfamily and several putative drug exporters of the ABC superfamily may serve similar functions. All of these families are represented in the corynebacteria. The diversity of substrates exported by these systems has yet to be studied. As noted in Table 8.2, about 50 transporters in both corynebacteria analyzed are probably concerned with inorganic ion transport. The following families are represented (see Table 8.3): (1) for monovalent cations (10 families): VIC, CPA1, Amt, CPA3, KUP, F-ATPase, P-ATPase, and two or three proton- or sodium-translocating electron carriers (QCR and COX families); (2) for di- or trivalent cations (10 families): MIT, NNP(MFS), CDF, CaCA, CadD, FeCT(ABC), MZT(ABC), Nit(ABC), P-ATPase, and MgtE; and (3) for anions (10 families): Pit, ArsB, DASS, CHR, SulP, PNaS, ACR3, PhoT(ABC), MolT(ABC), and NitT(ABC). Inspection of Table 8.3 reveals possible transporters for a variety of additional interesting metabolites such as organic anions (benzoate, phenylacetate, cyanate, phosphonates, and sulfonates). Transporters specific for osmolytes, both purine and pyrimidine bases and nucleosides, quaternary ammonium compounds, and possibly a nucleotide (nicotinamide mononucleotide, NMN) were identified. Macromolecular exporters were also found. Protein secretion and membrane protein insertion systems include the Sec, Tat, and Oxa1 systems, while carbohydrates can be exported via MOP, ABC, and VGP family transporters. Putative lipid exporters of the RND superfamily have been identified, and many MFS and ABC systems may similarly catalyze lipid “flip-flop,” which is equivalent to export from the inner leaflet of the cytoplasmic membrane bilayer to the outer leaflet. Some of these transporters may also export lipids from the inner membrane to the outer membrane in these bilayered enveloped bacteria. Finally, a large percentage of the identified transporters could not be assigned even a tentative function. It should also be kept in mind that transporters that belong to functionally uncharacterized families may not be included in the TC system and therefore may not be identified using the computer approaches used here. It is clear that we are only at the beginning of an understanding of molecular transport processes in corynebacteria.

Genomic Analyses of Transporter Proteins in C. glutamicum and C. efficiens

185

ACKNOWLEDGMENTS Work in our laboratory was supported by NIH grant GM55434. We thank Mary Beth Hiller for her assistance in the preparation of this chapter.

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[6] [7] [8]

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[13]

[14]

Arispe N and De Maio A. (2000) ATP and ADP modulate a cation channel formed by Hsc70 in acidic phospholipid membranes. J. Biol. Chem. 275:30839–30843. Buckel W. (2001) Sodium ion-translocating decarboxylases. Biochim. Biophys. Acta 1505:15–27. Burkovski A and Kramer R. (2002) Bacterial amino acid transport proteins: occurrence, functions, and significance for biotechnological applications. Appl. Microbiol. Biotechnol. 58:265–274. Busch W and Saier MH Jr. (2002) The Transporter Classification (TC) System, 2002. CRC Crit. Rev. Biochem. Mol. Biol. 37:287–337. Cao TB and Saier MH Jr. (2003) The general protein secretory pathway: Phylogenetic analyses leading to evolutionary conclusions. Biochim. Biophys. Acta 1609:115–125. Costa-Riu N, Maier E, Burkovski A, Krämer R, Lottspeich F, and Benz R. (2003). Identification of an anion-specific channel in the cell wall of the Gram-positive bacterium Cornyebacterium glutamicum. Mol. Microbiol. 50:1295–1308. Cox JS, Chen B, McNeil M, and Jacobs WR Jr. (1999) Complex lipid determines tissue-specific replication of Mycobacterium tuberculosis in mice. Nature 402:79–83. Dimroth P, Jockel P, and Schmid M. (2001) Coupling mechanism of the oxaloacetate decarboxylase Na+ pump. Biochim. Biophys. Acta 1505:1–14. Fudou R, Jojima Y, Seto A, Yamada K, Kimura E, Nakamatsu T, Hiraishi A, and Yamanaka S. (2002) Corynebacterium efficiens sp. nov., a glutamic-acid-producing species from soil and vegetables. Int. J. Syst. Evol. Microbiol. 52:1127–1131. Ikeda M and Nakagawa S. (2003) The Corynebacterium glutamicum genome: features and impacts on biotechnological processes. Appl. Microbiol. Biotechnol. 62:99–109. Kennerknecht N, Sahm H, Yen MR, Patek M, Saier MH Jr, and Eggeling L. (2002) Export of L-isoleucine from Corynebacterium glutamicum: a two-gene-encoded member of a new translocator family. J. Bacteriol. 184:3947–3956. Kotrba P, Inui M, and Yukawa H. (2003) A single V317A or V317M substitution in Enzyme II of a newly identified beta-glucoside phosphotransferase and utilization system of Corynebacterium glutamicum R extends its specificity towards cellobiose. Microbiology 149:1569–1580. Krogh A, Larsson B, von Heijne G, and Sonnhammer EL. (2001) Predicting transmembrane protein topology with a hidden Markov model. Application to complete genomes. J. Mol. Biol. 305:567–580. Kruse D, Kramer R, Eggeling L, Rieping M, Pfefferle W, Tchieu JH, Chung YJ, Saier MH Jr, and Burkovski A. (2002) Influence of threonine exporters on threonine production in Escherichia coli. Appl. Microbiol. Biotechnol. 59:205–210. Lichtinger T, Riess FG, Burkovski A, Engelbrecht F, Hesse D, Kratzin HD, Kramer R, and Benz R. (2001) The low-molecular-mass subunit of the cell wall channel of the Gram-positive Corynebacterium glutamicum: immunological localization, cloning and sequencing of its gene porA. Eur. J. Biochem. 268:462–469.

186 [15]

[16]

[17] [18]

[19]

[20] [21] [22]

[23]

[24] [25]

[26] [27]

[28]

[29]

Handbook of Corynebacterium glutamicum Nishio Y, Nakamura Y, Kawarabayasi Y, Usuda Y, Kimura E, Sugimoto S, Matsui K, Yamagishi A, Kikuchi H, Ikeo K, and Gojobori T. (2003) Comparative complete genome sequence analysis of the amino acid replacements responsible for the thermostability of Corynebacterium efficiens. Genome Res. 13:1572–1579. Nottebrock D, Meyer U, Kramer R, and Morbach S. (2003) Molecular and biochemical characterization of mechanosensitive channels in Corynebacterium glutamicum. FEMS Microbiol. Lett. 218:305–309. Parche S, Thomae AW, Schlicht M, and Titgemeyer F. (2001) Corynebacterium diphtheriae: a PTS view to the genome. J. Mol. Microbiol. Biotechnol. 3:415–422. Parche S, Burkovski A, Sprenger GA, Weil B, Kramer R, and Titgemeyer F. (2001) Corynebacterium glutamicum: a dissection of the PTS. J. Mol. Microbiol. Biotechnol. 3:423–428. Pivetti CD, Yen MR, Miller S, Busch W, Tseng YH, Booth IR, and Saier MH Jr. (2003) Two families of prokaryotic mechanosensitive channel proteins. Microbiol. Mol. Biol. Rev. 67:66–85. Saier MH Jr. (2000) A functional-phylogenetic classification system for transmembrane solute transporters. Microbiol. Mol. Biol. Rev. 64:354–411. Saier MH Jr. (2003) Answering fundamental questions in biology with bioinformatics. ASM News 69:175–181. Tauch A, Homann I, Mormann S, Ruberg S, Billault A, Bathe B, Brand S, BrockmannGretza O, Ruckert C, Schischka N, Wrenger C, Hoheisel J, Mockel B, Huthmacher K, Pfefferle W, Pühler A, and Kalinowski J. (2002) Strategy to sequence the genome of Corynebacterium glutamicum ATCC 13022: use of a cosmid and a bacterial artificial chromosome library. J. Biotechnol. 95:25–38. Tran CV, Yang NM, and Saier MH Jr. (2003) TC-DB: An architecture for membrane transport protein analysis. Proc. 2nd Intl. IEEE Computer Society Computational Systems Bioinformatic Conference, p. 658. Weeks DL and Sachs G. (2001) Sites of pH regulation of the urea channel of Helicobacter pylori. Mol. Microbiol. 40:1249–1259. Weeks DL, Eskandari S, Scott DR, and Sachs G. (2000) A H+-gated urea channel: the link between Helicobacter pylori urease and gastric colonization. Science 287:482–485. Winnen B, Hvorup RN, and Saier MH Jr. (2003) The tripartite tricarboxylate transporter (TTT) family. Res. Microbiol. 154:457–465. Yen MR, Tseng YH, Nguyen EH, Wu LF, and Saier MH Jr. (2002) Sequence and phylogenetic analyses of the twin arginine targeting (Tat) protein export system. Arch. Microbiol. 177:441–450. Zhai Y and Saier MH Jr. (2001) A web-based program (WHAT) for the simultaneous prediction of hydropathy, amphipathicity, secondary structure and transmembrane topology for a single protein sequence. J. Mol. Microbiol. Biotechnol. 3:501–502. Zhang Z, Aboulwafa M, Smith M, and Saier MH Jr. (2003) The ascorbate transporter of Escherichia coli. J. Bacteriol. 185:2243–2250.

9

Export of Amino Acids and Other Solutes L. Eggeling

CONTENTS 9.1 9.2

Introduction ..................................................................................................187 Export of L-Lysine .......................................................................................189 9.2.1 Identification of the L-Lysine Exporter LysE ..................................189 9.2.2 Functional Residues within LysE ....................................................190 9.2.3 Regulation of lysE Expression.........................................................192 9.2.4 Mechanism of L-Lysine Export........................................................192 9.2.5 Function of the Lysine Exporter......................................................193 9.2.6 Specificity of LysE...........................................................................194 9.2.7 Modulation of L-Lysine Export Activity..........................................194 9.3 The LysE Superfamily of Translocators......................................................195 9.3.1 The LysE and CadD Families..........................................................195 9.3.2 The RhtB Family .............................................................................196 9.4 Export of L-Threonine..................................................................................196 9.4.1 The ThrE Family of Exporters ........................................................198 9.5 Export of Branched-Chain Amino Acids ....................................................198 9.5.1 The LIV-E Family of Exporters ......................................................200 9.6 Export of L-Glutamate .................................................................................200 9.7 Contribution of Cell Wall to Amino Acid Export .......................................201 9.8 Further Exporters of C. glutamicum............................................................202 9.9 Outlook.........................................................................................................203 Acknowledgments..................................................................................................203 References..............................................................................................................204

9.1 INTRODUCTION After the discovery that Corynebacterium glutamicum can easily be influenced to excrete L-glutamate, powerful producers were rapidly developed for this amino acid and for L-lysine, too. Mutants of C. glutamicum producing these biotechnologically important compounds are also available for several other amino acids [2,47,66,126]. Whereas the key features for the regulation and individual steps of amino acid biosynthesis have been intensively studied, the analysis and understanding of export 187

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has not initially kept pace. One reason is undoubtedly that it was simply not conceivable that specific export carriers exist for this purpose. It is therefore not surprising that as the first idea for L-glutamate excretion it was assumed that some leakage (diffusion) across the cell membrane might occur [76,108]. Later, the idea of the functional inversion of a glutamate-uptake system mediating L-glutamate transport in the opposite direction was put forward [17]. In the case of L-lysine excretion, another idea was also discussed. Here it was suggested that efflux might be mediated by some osmotically controlled pores [69]. The functional inversion of uptake systems is still under discussion, at least for the ABC transporter HisF of Salmonella typhimurium mediating the ATP-coupled histidine uptake, which is thought to additionally enable excretion of its substrate [45]. This speculative model, however, does not hold for L-glutamate export in C. glutamicum, since a mutant deleted of the ATP-dependent GluABCD uptake system is unaltered in its L-glutamate export activity [62]. Based on the diffusion properties of L-glutamate and L-lysine — both amino acids that carry a net charge — it is obvious that passive diffusion of these amino acids must be rejected [71]. Moreover, both are translocated against a concentration gradient, thus excluding the involvement of a pore [60]. It is therefore now generally accepted that specific export proteins are present in C. glutamicum to translocate amino acids from the cytosol to the surrounding medium. The final confirmation for this was given by the molecular identification of such exporters. They represent in part completely novel types of translocators, which, however, based on genome information, apparently also occur in other microorganisms. Active export for amino acids is demonstrated in C. glutamicum for L-glutamate [43], L-lysine [13], L-isoleucine [129], and L-threonine [91]. In E. coli, active export is demonstrated for L-threonine [63], as well as L-cysteine [20]. Some amino acids are released from the cell in addition to active export by diffusion (Figure 9.1). This is demonstrated with C. glutamicum for the hydrophobic branched-chain amino acids [129], or L-threonine [91]. Diffusion might be the only mechanism for release of L-tyrosine and L-phenylalanine from the cell [14]. In order to express the basically different possibilities of passage of the amino acid over the entire cell wall barrier into the surrounding medium, in this summarizing overview the term export, or transport, is used for carrier-protein–catalyzed, energy-dependent translocation (Figure 9.1), whereas efflux additionally includes the passage of membrane barriers by diffusion as a consequence of concentration gradients. At present, consolidated findings mainly relate to carriers that catalyze export over the cytoplasmic membrane. Since the amino acids naturally also have to pass through the various layers of the cell wall (see Chapter 7), they could in principle also play a part in determining the efflux (and influx) properties of the cell. In particular, this could be the case for the second lipid layer of C. glutamicum. This outer lipid layer is characteristic of Corynebacterianeae [9,107] and consists of mycolic acids covalently bound to arabinogalactan together with soluble trehalose mycolates [93]. The possible involvement of the second outer lipid layer will be discussed in the context of L-glutamate export. Some aspects of amino acid export through the cell wall of C. glutamicum have been treated in recent reviews [14,27,28,60].

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FIGURE 9.1 Fluxes over the cell wall of C. glutamicum. Components of the cell wall are the cytoplasmic membrane (phospholipid layer), peptidoglycan, arabinogalactan, and the mycolic acid layer. Export is catalyzed by carriers such as LysE, BrnFE, or ThrE. In addition, undirected diffusion may take place as in the case of the hydrophobic isoleucine. Glutamate export is a special case because export can take place only by influencing cell wall components, the details of which are not yet known and are expressed by a question mark. Import might counteract export and negatively influence the efflux rate. In addition, porins might control fluxes of solutes via the outer mycolic acid layer.

9.2 EXPORT OF L-LYSINE 9.2.1 IDENTIFICATION

OF THE L-LYSINE

EXPORTER LYSE

The gene of the lysine exporter lysE was isolated by complementation of a mutant of C. glutamicum defective in export [120]. To this end, two specific features were used. First, detection for absence or presence of export from individual colonies in a plate assay and, second, inducible lysine excretion to ensure viability of the mutant devoid of exporter since L-lysine cannot be degraded by C. glutamicum [82]. Simply the addition of 5 mM L-methionine to C. glutamicum served to increase L-lysine synthesis and excretion [120]. Methionine addition reduces the hom gene expression (homoserine dehydrogenase) [77], therefore making available an increased aspartate semialdehyde concentration for the dihydrodipicolinate synthase. This results in an increased flux toward L-lysine [24]. The chromosomal fragment complementing the export-deficient mutant R127-NA8 contained the exporter gene lysE and the divergently transcribed transcriptional regulator gene lysG. The LysE polypeptide consists of 233 amino acyl residues. It is characterized by six hydrophobic segments [118]. As shown by a set of PhoA and LacZ fusions. five hydrophobic segments each traverse the membrane once, while the remaining hydrophobic segment (II in Figure 9.2) is not membrane-spanning, but either located peripherally on the periplasmic side of the membrane or possibly dipping into the membrane. This segment exhibits the highest hydrophobicity within LysE. It is noteworthy that in the recently obtained three-dimensional (3-D) structures of aquaporin

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FIGURE 9.2 Topology of the LysE carrier together with residues conserved or mutated. The gray boxes mark the six hydrophobic segments, which are numbered I through VI. Conserved residues are circled. Amino acyl residues for which muteins were created and analyzed are numbered according to their position in the polypeptide sequence.

[122] or K+ channels [85] hydrophobic regions are present that do not simply form transmembrane spanning α-helices and that are mechanistically of great significance. There is no information on the oligomeric structure of the L-lysine exporter. However, in agreement with the notion that solute translocation usually requires a protein made up of 10 to 14 transmembrane spanning helices, and by analogy with other small transporter proteins, such as Emr or AQP1 [78,106], LysE might be active as a homo-dimer.

9.2.2 FUNCTIONAL RESIDUES

WITHIN

LYSE

To obtain information on the path of L-lysine through the carrier and on the localization of substrate binding sites LysE was mutated and the translocation velocity analyzed in 13032ΔlysEG [37]. First, the long loop connecting hydrophobic segments III and IV was studied. Although with large deletions encompassing the entire loop, like Δ96–137, Δ88–123, or Δ88–137, no export occurs, with small deletions in the loop, irrespective of where they are located, like Δ88–95, Δ96–112, Δ112–123, or Δ124–137 the export rate was maximally reduced from 9.6 to 6.9 nmol⋅min–1 ⋅(mg dry weight)–1. Therefore, the loop is not really essential. Instead, it appears that it must have a minimal length of 24 amino acids for a proper positioning of the helices and

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191

12 10 8 6 4 2 0 p UMB

46 D-N

46 D-K

46 D-A

46 D-E

150 T-A

152 L-A

153 N-A

153 N-Q

154 P-A

155 N-A

155 N-H

159 D-E

159 D-A

159 D-N

159 D-K

FIGURE 9.3 Export activities obtained with LysE mutant proteins. L-Lysine export is given in nmol⋅min-1⋅(mg dry weight)-1 on the y-axis. The bar labeled with pUMB represents the activity obtained with wild-type LysE.

functional residues in their 3-D organization. The minimal length approximately corresponds to the length of the loop in LysE proteins of other bacteria. The relevance of charged residues was similarily studied with specific muteins. In position 58, mutations D58K or D58E did not influence the export activity, which is in accord with the fact that this residue is not conserved within LysE proteins present in other bacteria. In the L-lysine producer C. efficiens, a glycine residue is present at this position. Similarly, mutations in the nonconserved positions 69 and 72, like D69A and D69E, as well as R72E and R72K, did not influence transport. These studies also exclude involvement in intrahelical salt bridge formation of the residues assayed. However, mutating the conserved aspartate residue in position 46 strongly affects export (Figure 9.3). Apparently, the negative charge in position 46 is essential for translocation since with D46E L-lysine is still excreted, whereas upon introduction of a positive charge or a neutral residue at this position virtually no export occurs. Of particular interest is the conserved motif LNPNAYLD (aa 152–159), which is located in the center of hydrophobic segment IV. Whereas introducing the L152A mutation did not influence export, the following asparagine residue is sensitive to mutation (Figure 9.3). With N153A, less than half of the export rate is observed, and the replacement of the asparagine residue by glutamine, which is only one methylene group longer, reduces the export even further. This points to a steric hindrance of the transport of L-lysine. The adjacent prolyl residue is in accordance with the significance of N153 for translocation. Prolyl residues are known to introduce kink angles of about 20˚ into transmembrane helices to position functionally important residues within the three-dimensional structure of carriers [36]. Accordingly, the P154A mutation results in almost inactive export. The aspartate residue in position 159 seems to be of special significance (Figure 9.3). Extending the side

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chain by one methylene group through D159E reduces the activity to less than onethird and the exchange of charge in D159K leads to an inactive protein. Owing to these structural and functional characteristics, D159 may possibly be directly involved in binding the substrate L-lysine. Furthermore, the fact that D159 is two turns away from N155 indicates that these residues might line the translocation channel.

9.2.3 REGULATION

OF LYSE

EXPRESSION

The transcription of lysE is tightly controlled by the lysG encoded regulator LysG. This regulator belongs to the large group of LysR-type transcriptional regulators (LTTR), which probably make up the largest family of prokaryotic regulatory proteins [40]. Corynebacterium glutamicum possesses ten regulators of this type. LysG enables the up-to-20-fold induction of lysE in the presence of L-lysine as quantified by β-galactosidase activities of single-copy transcriptional fusions [7]. Further in vivo studies have shown that LysG interacts with the lysE–lysG intergenic region. In addition, in vitro gel-shift experiments with purified LysG and DIG-labeled lysE–lysG fragments demonstrated the direct interaction of the regulator with the lysE promoter [6]. According to these data and the strong homology with other members of the LTTR family, L-lysine is probably recognized by LysG in a cavity made up of residues within the center of the regulatory protein [114]. As typical of LTTRs, this might result in further cooperative interactions between a LysG multimer and the promoter region to increase the affinity for RNA polymerase, thus causing increased lysE transcript initiation. As deduced from DNA-chip-based expression studies, LysG most likely does not have any other target gene [61]. As mentioned, the co-inducer of lysE expression is L-lysine. In addition, also the basic amino acids L-arginine, L-citrulline, and L-histidine are recognized by LysG to promote activation of lysE transcription [7]. However, L-citrulline and L-histidine do not serve as carrier substrates. Using a LysE–LacZ fusion, the intracellular Llysine concentration required for maximal lysE expression was determined as 35 mM to 42 mM [7]. These data fit with results obtained by methionine-feeding-induced L-lysine excretion, where significant excretion was only obtained above an intracellular L-lysine concentration of about 20 mM [120]. The only other LTTR regulator for which an attempt has been made to quantify the intracellular co-inducer concentration required for activation is NhaR, which controls the synthesis of the Na+/H+ antiporter NhaA of E. coli. Full induction of nhaA by NhaR is obtained when the intracellular Na+ concentration is around 60 mM [38].

9.2.4 MECHANISM

OF L-LYSINE

EXPORT

The components driving L-lysine translocation are the electrochemical proton potential (inside negative) and the chemical concentration gradient of the solute S (RTlog[S]in /[S]out) [12]. L-Lysine excretion is thus characterized as a secondary transport system. A formal description of the energetic steps of the carrier during translocation is given in Figure 9.4. The positively charged L-lysine is presumably exported in symport with two OH- ions (which cannot be discriminated from an

Export of Amino Acids and Other Solutes

Δ Lys Δ pH

Medium

+ 2 OHLys+

OHOH+ Lys

193

C+

C+

C+

C+

Cytosol

OHOH+ Lys

− 2 OHLys+

ΔΨ FIGURE 9.4 Kinetic model of L-lysine export. The deduced steps of loading the carrier at the cytosolic side, translocation, release of substrate, and reconstitution of carrier are given.

antiport of two H+). For the substrate translocation step, S and pH are decisive, whereas for reorientation of the carrier ΔΨ is required. At high external L-lysine concentrations, the substrate gradient might become more important as a limiting step within the catalytic cycle [53]. The estimated Km of the export carrier is about 20 mM [12]. This ensures that under normal physiological situations, in which such a high concentration is rarely reached, L-lysine is not excreted in substantial amounts. The export of the cellular building block L-lysine is therefore subject to a double safety control: both the biochemical properties of the carrier and the expression control of the gene are ideally suited for an excretion system of a primary cellular building block. The Vmax of L-lysine export in the wild-type is 1.8 nmol⋅min–1⋅(mg dry weight)–1 [10]. The kinetic parameters of L-lysine export (in particular, the low affinity of the export system) are in sharp contrast to those of the L-lysine uptake system. The L-lysine importer LysI is characterized by a high affinity with a Km of 10 μM and a Vmax of 0.23 nmol⋅min–1⋅(mg dry weight)–1 [11]. LysI imports L-lysine or its analogue S-(2aminoethyl)-cysteine [99] in exchange for L-alanine, L-valine, or L-leucine. In the presence of significant internal L-lysine concentrations, lysine/lysine counterexchange without net transport occurs. LysI is not relevant for L-lysine production.

9.2.5 FUNCTION

OF THE

LYSINE EXPORTER

The physiological function of the exporter is to excrete excess L-lysine or L-arginine as a result of natural flux imbalances when increased intracellular basic amino acid concentrations are present. These flux imbalances are due to poor regulation as well as limited degradation capabilities. An example of such an imbalance is the surplus of L-lysine formation upon external L-methionine supply owing to a weak control of aspartate semialdehyde distribution in C. glutamicum [24]. Another situation in which an excess of L-lysine or L-arginine can be formed intracellularly is growth on

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complex media [118] or, more directly, growth on salt media in the presence of peptides [7]. L-Lysine-containing peptides, such as Lys-Lys-Lys, Lys-Glu, Ile-Lys, Ala-Lys, or Lys-Ala, are taken up with velocities of up to 11.4 nmol⋅min–1⋅(mg dry weight)–1 [32,129]. As deduced from the genome, C. glutamicum might have at its disposal at least three putative ABC-carriers for the uptake of peptides. These peptides are then hydrolyzed to obtain access to the amino acids, as is the case with other bacteria, too. However, C. glutamicum is unable to degrade L-lysine [82], forcing the cell to excrete a surplus of amino acid not used for growth. The significance of the exporter for the wild-type becomes immediately apparent upon chromosomal deletion of lysE. Supply of 3 mM Lys-Ala to a salt medium containing glucose as a carbon and energy source results in the intracellular accumulation of 1,100 mM of L-lysine with bacteriostasis as a consequence [118]. Similarily, with 2 mM Arg-Ala, an accumulation of 900 mM L-arginine is the result in the deletion mutant, with growth strongly reduced [7]. In addition, on complex media like LB or BHI the exporter is necessary, since growth of the deletion mutant is retarded [118], and lysE expression is induced on such media [7].

9.2.6 SPECIFICITY

OF

LYSE

LysE exports the basic amino acids L-lysine and L-arginine at comparable rates of 0.75 nmol⋅min–1⋅(mg dry weight)–1. Although L-histidine and L-citrulline also act as co-inducers of lysE expression, these two amino acids are not exported by LysE. However, C. glutamicum has additional export activities, other than those of LysE at its disposal for exporting L-histidine, L-citrulline, and L-ornithine [7]. In agreement with these observations, producer strains of C. glutamicum and Brevibacterium ketoglutamicum are known for L-ornithine and L-histidine, respectively [16,72]. The basic amino acid D,L-diaminopimelate is not accepted as a substrate by any of the exporters. Instead, a lysA deletion mutant devoid of diaminopimelate decarboxylase accumulates more than 1 M of this particular amino acid intracellularly, with negligible concentrations found extracellularly (5 mM), illustrating again that any diffusive passage for charged amino acids over the cytoplasmic membrane is insignificant [7].

9.2.7 MODULATION

OF L-LYSINE

EXPORT ACTIVITY

As known from many examples, the lipid environment of carriers can massively influence their kinetic properties [1]. The reason is that the carrier is a constituent of a larger structure, the lipid bilayer, and therefore many interactions with this structure are possible. Even the location of subdomains in carriers can be affected by the lipid composition of the membrane as demonstrated for the lactose permease LacY of E. coli [123]. This could be put forward as an explanation for the wide range of differences in the lysine export properties of specific C. glutamicum strains. Subtle differences regarding the export activity in pH and the membrane potential were detected between strains 52-5 and MH20-22B [10]. Moreover, export with the wild-type does not proceed according to Michaelis-Menten kinetics [32] and occurs only above a threshold level of 20 mM, whereas this is not the case with producer

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195

strains [33]. Also, a cooperative interaction of internal L-lysine with its excretion system has been assumed [34]. Another observation is that the choice of the carbon and energy source present in efflux assays results in different export rates. Thus in the wild-type and MH20-22B as well, the export in the presence of glucose is 2.6 nmol⋅min–1⋅(mg dry weight)–1, but it is 5.6 nmol⋅min–1⋅(mg dry weight)–1 in the presence of glutamate [33,34]. Sequencing of a 2.374-kbp chromosomal BamHI fragment encompassing lysEG from strains 52-5 and MH20-22B did not reveal any mutation as compared with the wild-type sequence (unpublished data). This fact, together with the differences in export activity noted, is consistent with a likely modulation of the carrier activity by the lipid environment. In addition, the high-level L-lysine producer B6 is not mutated in its L-lysine export carrier [88]. A particularly interesting case of an exportnegative mutant is strain 35/48 obtained by undirected mutagenesis [120]. This strain is again not mutated within the 2.374-kbp chromosomal BamHI fragment but is nevertheless unable to excrete L-lysine and accumulates up to 350 mM of this amino acid in its cytosol. One possibility could be that in this strain the carrier is unincorporated or misincorporated into the membrane because the strain also displays greatly modified transformation properties with plasmid DNA.

9.3 THE LYSE SUPERFAMILY OF TRANSLOCATORS LysE of C. glutamicum is the paradigm of a new superfamily of translocators, most of which are known only from genome sequencing. The superfamily is restricted to bacteria and archaea and consists of the LysE, RhtB, and CadD families [119]. All members of these three families consist of proteins of similar sizes (about 200 residues) and exhibit six hydrophobic α-helical segments. Since members of a single transporter family seldom catalyze the transport of structurally divergent types of compounds (i.e., sugars versus amino acids), and moreover function with strongly preferential polarity of transport direction (i.e., inward versus outward) [97], it is very probable that the majority of members of the LysE superfamily are transporters that export small molecules out of the cell.

9.3.1 THE LYSE

AND

CADD FAMILIES

More than 70 members of the LysE family are currently known. It may be assumed that some of them also excrete basic amino acids or derivatives of these. In fact, it was recently demonstrated that the LysE homolog AttX, which is relevant for the virulence of Rhodococcus fasciens, is present in the att locus, which is essential for the synthesis of an unidentified compound [70]. Although the substance causing the virulence has not yet been identified, the attA, attB, and attH genes involved in its synthesis share identities with L-arginine biosynthesis genes. This scenario makes it possible that the LysE homolog AttX of R. fasciens extrudes an arginine derivative as the virulence-inducing factor. The CadD family is the smallest group within the LysE superfamily. Members of this family have been demonstrated to confer cadmium resistance to Staphylococcus species [15]. Another member is suggested to export quaternary ammonium

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ions from Bacillus pseudofirmus (data bank entry Q45153). The export of the positively charged solutes by these exporters, as well as the export of basic amino acids by LysE from C. glutamicum and the special features of the att locus in R. fasciens, provides a strong indication that many members of the LysE and CadD families probably export positively charged molecules.

9.3.2 THE RHTB FAMILY The RhtB family is the largest group within the LysE superfamily. Astonishingly, a number of organisms have paralogous genes. Thus, B. subtilis has two paralogs, E. coli as Bacillus anthracis has five [95], and Shewanella oneidensis has as many as seven [39]. The high number is an intriguing argument in favor of a novel function of these proteins. As one idea, it has been suggested that RhtB family members could be involved in quorum sensing [3]. However, for some of the five paralogs of E. coli their relatedness to amino acid export has been demonstrated. RhtB of E. coli participates in the export of L-homoserine and confers resistance to L-homoserine lactone and β-hydroxynorvaline [128], as does YeaS [28]. RhtC and YfiK confer resistance to L-threonine [3]. Recently, it has been demonstrated that YfiK mediates L-cysteine export from E. coli [35]. Although the unambiguous participation and the respective fraction of the five paralogous carriers of E. coli in threonine export has not been finally clarified and a further exporter for L-threonine may even exist [128], the carriers of the RhtB family are nevertheless relevant for the excellent properties of threonine-producing E. coli strains [22]. In accord with this view is the finding that upon rhtB or rhtC overexpression the specific productivities in producer strains are slightly increased [63]. Another producer of E. coli is impaired in its L-threonine uptake [89], illustrating in this case the relevance of reduced re-uptake of product (Figure 9.1), as is similar in the case for L-tryptophan production with C. glutamicum, where mutants impaired in uptake are more effective in L-tryptophan accumulation (see Chapter 21).

9.4 EXPORT OF L-THREONINE The export of L-threonine in C. glutamicum is driven by proton motive force [91]. The gene encoding the L-threonine exporter (thrE) was identified by analyzing transposon mutants exhibiting increased sensitivity to the tripeptide Thr-Thr-Thr [104]. The corresponding polypeptide is a membrane protein that is characterized at its C-terminus by 10 hydrophobic α-helical segments. In addition, an extended N-terminus of 166 amino acyl residues is present, although probably not directly required for export (Figure 9.5). When thrE is overexpressed, L-threonine is exported at an increased rate of 3.8 nmol⋅min–1⋅(mg dry weight)–1, whereas in a thrE deletion mutant the rate is 1.1 nmol⋅min–1⋅(mg dry weight)–1. L-Serine is also a substrate of ThrE, but glycine is not. The identification of ThrE enabled the different efflux routes of L-threonine to be quantified in detail. At an intracellular concentration of about 170 mM L-threonine, at least three separate components contribute to total L-threonine efflux. The major component of 59% is the export driven by ThrE. The efflux

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S. co.

197

C. gl. C. je. E. co. P. mu. V. ch. R. ch. X. fa. B. ha. C. ac. M. tu. M. th.

P. fa. S. ce. S. po. FIGURE 9.5 Domain structure of selected proteins of the ThrE family of exporters. Arrows indicate the C-termini. ThrE of C. glutamicum is given at the top showing the hydrophobic part as two hatched rectangles representing the carrier part with ten-transmembrane helices, and illustrating its origin by duplication. In Campylobacter jejeuni, two adjacent genes encode the exporter where the two polypeptides, each with five-transmembrane helices, are not fused. An amphiphilic domain probably not involved in export is indicated as a black box. Mycobacterium tuberculosis has two such hydrophilic domains. C. glu., Corynebacterium glutamicum; C. je., Campylobacter jejeuni; P. mu., Pasteurella multocida; V. ch., Vibrio cholerae; R. ch., Rhodobacter capsulatus; X. fa., Xylella fastidiosa; B. ha,. Bacillus halodurans; C. ac., Clostridium acetobutylicum; M. tu., Mycobacterium tuberculosis; M. th., Methanobacterium thermoautotrophicum; S. co., Streptomyces coelicolor; P. fa., Pichia farinosa; S. ce., Saccharomyces cerevisiae; S. po., Schizosaccharomyces pombe.

resulting from passive diffusion contributes 22%, and the remaining 19% is due to at least one other still unidentified carrier. To convert the wild-type of C. glutamicum into an L-threonine producer, one of the major obstacles is clearly the export. This was demonstrated by graded expression of homFbr encoding the feedback-resistant key enzyme homoserine dehydrogenase [96]. With increasing copy numbers of homFbr, only a moderate rise in extracellular L-threonine was obtained, but a dramatically increased intracellular concentration of up to 100 mM was obtained. The originally observed plasmid instabilities in the case of overexpressed homFbr also indicate that C. glutamicum does not easily tolerate high internal L-threonine concentrations and that export is limiting [4,48,77,79]. Thus, although ThrE and a high intracellular L-threonine concentration are present, this does not result in an intensely increased export activity. This clearly differentiates L-threonine export from L-lysine export, where all information available confirms that the LysE–LysG system is designed to control the intracellular concentration of

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L-lysine

or L-arginine by export. Nevertheless, in a model strain overexpression of thrE led to an approximately 40% increased extracellular L-threonine accumulation [105]. The extracellular accumulation of L-threonine is further increased if the intracellular unspecific degradation of L-threonine by serine hydroxymethyltransferase is reduced [105].

9.4.1 THE THRE FAMILY

OF

EXPORTERS

As of November 2004, more than 30 ThrE homologs are identified and constitute the ThrE family of translocators. The family is thus relatively small, with members present in all domains of life, but, within the eukarya, only in the fungal kingdom. It is suggested that this family is an ancient family that arose before the split that separated the three domains of life, and that genes have been selectively lost from many organisms [127]. There are two striking structural features of ThrE that are obvious when comparing the family members (Figure 9.5). The ThrE proteins have the ten hydrophobic segments that are typical of a polytopic membrane protein, but in addition they carry the extensive long N-terminus already mentioned, and which is predicted to be located at the cytoplasmic side. In the case of C. glutamicum this N-terminal part is 166 amino acyl residues (aa) long. The longest, consisting of 504 aa, is present in Schizosaccharomyces pombe. This part of the protein carries an amphipatic region (black box in Figure 9.5) that exhibits sequence identities with portions of hydrolases, such as peptidases and glycosidases. Another structural feature of the members of the ThrE family is that they exist either as a single polypeptide chain or as two gene products (Figure 9.5). Single peptides are present in M. tuberculosis and Streptomyces coelicolor, for instance [104], which are the closest homologs to ThrE of C. glutamicum, whereas "spliced" two-component systems are found among others in E. coli, Vibrio cholerae, Saccharomyces cerevisiae, or S. pombe [127]. The existence of the spliced homologs, together with weak sequence identities between both parts and the respective parts of the large polypeptide, is an indication that the proteins result from a gene duplication event. A similiar situation is present with other carriers, such as the drug/metabolite efflux (DME) family of efflux carriers or the Ca2+:cation antiporter (CaCA) family [98].

9.5 EXPORT OF BRANCHED-CHAIN AMINO ACIDS Efflux of the branched-chain amino acids L-isoleucine, L-leucine, and L-valine occurs in part via diffusion (Figure. 9.1). In addition, carrier-mediated export is driven by the proton motive force [41,129]. The L-isoleucine exporter is a two-component permease encoded by brnF and brnE [54]. The genes are organized as an operon. Divergently transcribed is lrp, which is required for exporter expression and thus encodes the positive regulator (Figure 9.6). When both brnF and brnE are overexpressed, export rates of 8 nmol⋅min–1⋅(mg dry weight)–1 are obtained, which is a twofold increase as compared with the wild-type. Upon deletion of both genes, no active export occurs, showing that BrnFE is the only relevant carrier exporting

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FIGURE 9.6 Overview of the brnE-brnF-lrp locus of C. glutamicum. The thick arrows represent the genes with their sizes in base pairs (bp). In the lower part, the intergenic region between the regulator gene lrp and brnFE is given. The transcriptional start sites are marked +1. The –10 and –35 promoter regions are underlined, and the initiation codons for lrp and brnF are shown in bold print. The average hydrophobicity (solid line) and similarity (dashed line) for the BrnE and BrnF families is indicated on top. L-isoleucine

from the cell. The carrier also exports L-leucine with comparable activity, but not L-valine, which is exported at a significantly reduced rate of about 50%. For the carrier, a low affinity of 21 mM was observed with L-isoleucine as substrate [41]. Since this is in the range of that known for LysE and L-lysine, low affinity could be a general feature of amino acid exporters. As mentioned, part of the total efflux of the branched-chain amino acids is due to diffusion. Whereas the cytoplasmic membrane is almost impermeable for L-lysine, with a diffusion rate constant of 0.002⋅min–1 [118], it is almost freely permeable for L-phenylalanine, with a rate constant of 0.45⋅min–1 [60]. An intermediate position is occupied by the branched-chain amino acids, with L-isoleucine diffusion exhibiting a first-order rate constant of 0.08⋅min–1 [129]. This means that at a given intracellular concentration of about 40 mM L-isoleucine and an observed efflux rate (export plus diffusion; see Figure 9.1) of about 8 nmol⋅min–1⋅(mg dry weight)–1, both components contribute almost equally to efflux [54]. The highest efflux rate obtained after deregulation of the L-isoleucine biosynthesis pathway [28] is 12.6 nmol⋅min–1⋅(mg dry weight)–1 [74,75]. Higher efflux rates have not yet been documented, which could be due to constant back flux into the cell via diffusion, thus preventing maintenance of a low intracelluar L-isoleucine concentration. In fact, the intracellular L-isoleucine concentration is always higher than the extracellular one after deregulation

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of the synthesis pathway, indicating that the flux capacity through the pathway exceeds the net efflux [73]. Since neiher overexpression of brnFE nor deletion of the L-isoleucine uptake carrier gene brnQ [111] lead to significantly increased L-isoleucine concentration (unpublished), the diffusion properties in the strains studied are probably decisive for L-isoleucine accumulation. In contrast, a product accumulation increase of more than 10% has been observed for tryptophan [46] and threonine [89] if the corresponding uptake systems are inactivated.

9.5.1 THE LIV-E FAMILY

OF

EXPORTERS

Corynebacterium glutamicum possesses a paralogous BrnFE system, which does not export branched-chain amino acids and whose function is unknown since a deletion mutant exhibits no phenotype (unpublished data). The BrnFE kind of exporter is present in diverse Gram-negative and Gram-positive bacteria as well as archaea but is lacking in eukaryotes, and the genes of the two-component carrier always map together in the same order constituting an operon. Both gene products together are the members of the novel LIV-E family of exporters [54]. The large polypeptide of the LIV-E exporter (BrnF) is predicted to span the membrane seven times, and that of the smaller protein four times. Interestingly, the last four transmembrane spanning helices of the BrnF homologs exhibit significant sequence similarity to the four transmembrane spanning helices of the BrnE homologs, which might indicate a partial extragenic duplication event that gave rise to the presentday members of the LIV-E family of efflux carriers [97]. The LIV-E system azlCD from B. subtilis gives rise to increased resistance to 4-azaleucine [5] and therefore presumably catalyzes export of the L-leucine analog. In addition to C. glutamicum, selected α-proteobacteria, such as Agrobacterium tumefaciens or Sinorhizobium meliloti, possess up to three paralogs of LIV-E exporters.

9.6 EXPORT OF L-GLUTAMATE The most striking finding with respect to amino acid production, and an incentive for biotechnology in general, was the discovery of C. glutamicum and its biotindependent L-glutamate excretion [59]. At a growth-limiting concentration of 2 μg of biotin per liter, L -glutamate is massively excreted at a rate of about 20 nmol⋅min–1⋅(mg dry weight)–1 [44]. Driven by the wish to use cheap substrates containing biotin, alternative processes have been developed, also leading to L-glutamate excretion (Table 9.1). The experimental section of this book describes two methods of easily acheiving L-glutamate excretion. In the actual large-scale process, temperature- and detergent-sensitive mutants are used to achieve L-glutamate excretion by temperature shift and, if required, by the addition of detergent. As can be seen from Table 9.1, the conditions for achieving L-glutamate efflux are surprisingly diversified. Correspondingly, there are also a wide range of models explaining export mechanistically [29,58,109]. A detailed model is given in Chapter 19 of this book. Molecular access has only recently become possible to individual genes, of in part unknown function, that influence export [42,56,57]. However, the exporter has not yet been described.

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TABLE 9.1 Overview of Steps to Achieve and Understand L-Glutamate Efflux By C. glutamicum. The selection and classification is limited and by no means complete. The individual observations are listed that lead to glutamate excretion (top), demonstrate active export (center) and permit molecular access to export (bottom). The development in the state of knowledge roughly coincides with the development over time. Observation or Knowledge Discovery of C. glutamicum Penicillin triggers efflux Lysozyme sensitivity triggers efflux Fatty acid esters (Tween-60) trigger efflux Oleic acid auxotrophy triggers efflux Glycerol auxotrophy triggers efflux Basic amine surfactants (dodecylammonium acetate) trigger efflux Efflux is an active process Local anesthetics trigger efflux Temperature increase triggers efflux dtsR expression influences efflux Penicillin-binding protein control causes temperature sensitivity and efflux ltsR inactivation causes lysozyme, temperature sensitivity and efflux alr, alanine racemase, influences efflux cma, acp, plsC, fadD15 genes of P-lipid synthesis alter efflux Ethambutol addition results in efflux Trehalose negative mutants exhibit increased accumulation 2-Oxoglutarate DH activity is reduced

Reference Biotin limitation triggers efflux Cell wall is involved

P-lipid composition is altered

59, 117, 101, 102 103, 84 100 103, 108, 109 90 81 23, 116

Exporter is necessary Importer is excluded

Link to lipid composition Link to peptidoglycan synthesis Link to cell wall synthesis

Link to P-lipid composition Link to arabinogalactan synthesis Link to mycolic acid content Link to central metabolism

43, 44 62 64 65 56, 57 121 42 29 83 94 80 52

9.7 CONTRIBUTION OF CELL WALL TO AMINO ACID EXPORT In addition to the cytoplasmic membrane and the carriers localized there, other cell wall components may be involved in the efflux. Initial findings are already available on the two amino acids L-lysine and L-glutamate produced with C. glutamicum. For these amino acids, the transport properties are also definitively determined by the outer lipid layer of the cell (Figure 9.1). This layer consists of mycolic acids, which

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are either bound to trehalose or directly to the arabinogalactan polymer (Chapter 7 details cell wall architecture). When the trehalose synthesis genes treY and otsA are inactivated, the L-glutamate accumulation rises from 40.2 to 45.6 g l–1 [80]. Also, in the case of L-lysine, the mycolic acid layer contributes to the efflux properties. A mutant that lacks treY and otsA is devoid of mycolic acids, either bound to trehalose or arabinogalactan [115,125]. It excretes L-lysine at an increased rate and accumulates 37% more of this amino acid (lysine per cell mass) than the original strain [124]. The significance of the physicochemical properties of the mycolic acid layer for the efflux can also be seen by the fact that upon inactivation of csp1 (synonym of cmytA or cop1 [8,21]), encoding a mycolyl transferase, which reduces the cell wall–bound mycolic acid content by 50%, the passage of glycerol is increased twofold [92]. From other Corynebacterianeae, like M. tuberculosis, it is known that the outer mycolic acid layer forms a highly ordered structure to produce cell walls of unusually low permeability [86]. For instance, the cell wall permeability to cephalosporins in M. chelonae is about three orders of magnitude lower than that of the E. coli outer membrane, and ten times lower than the permeability of the notoriously impermeable Pseudomonas aeruginosa outer membrane [50,51]. The diffusive flux over the mycolic acid layer also depends on the presence of porins and their properties. Recently, a porin of M. smegmatis has been identified [31], which, in contrast to that of Gram-negative bacteria, is a trimer, thus constituting the prototype of a new class of porins [67]. Porins are also present in C. glutamicum as concluded from black lipid bilayer experiments of cell wall fractions [67]. However, the proteins are unusually small. For instance, PorA of C. glutamicum is an acidic polypeptide of only 45 amino acyl residues [68,93]. A porA deletion mutant exhibits slower growth and is less susceptible to growth inhibition by antibiotics, but did not influence L-glutamate excretion [18]. The channel made up by PorA is thought to be the major hydrophilic pathway through the cell wall, whereas the second porin, PorB, present in C. glutamicum forms a small anion-specific channel [19]. The PorB protein contains a signal sequence, and the mature protein has 99 amino acyl residues. Whether the porins PorA, PorB, and the also detected PorC, influence amino acid excretion is not yet known. However, if import of a hydrophilic solute like a charged amino acid is limited by one of the porins, then also efflux of the same amino acid must be limited. It is therefore notable that the maximal import rate for L-glutamate is 1.5 nmol⋅min–1⋅(mg dry weight)–1 [62], whereas efflux upon induction by detergent addition, for instance, is at least one order of magnitude greater.

9.8 FURTHER EXPORTERS OF C. GLUTAMICUM In several C. glutamicum strains and related species, efflux carriers conferring resistance to cytotoxins have been identified. The majority of them are plasmidencoded (see Chaper 4). The tetracycline exporter TetZ is homologous to systems in Gram-negative species and regulated in its expression by the divergently oriented regulator gene (TetR) [110,113]. Whereas this exporter is substrate-specific, Cmr is

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a multidrug transporter exporting several unrelated antibiotics [49]. Interestingly, Cmr is chromosome-encoded, and it confers resistance only when expressed in E. coli, which could be due to a low expression level of cmr in C. glutamicum. Another exporter is TetAB, accepting as substrates selected tetracycline and β-lactam antibiotics [112]. This is a two-gene encoded ABC carrier where both genes exhibit the typical five membrane-spanning helices and the conserved ATP-binding motifs. The exporter LmrB belongs to the major facilitator family of transporters and mediates export of lincosamides, such as lincomycin and clindamycin [55]. Owing to the cloning procedure used, it is reasonable to assume that it also accepts as substrate 5-bromo-4-chloro-3-indolyl-β-D-galactopyranose (X-Gal). Further proteins mediating efflux are the osmoregulated channels MscL and YggB of C. glutamicum [87], which are described in detail in Chapter 18.

9.9 OUTLOOK The efflux of amino acids is both an important and a fascinating field of bacterial metabolite production. The fascination is in part due to the fact that in comparison to biosynthetic pathways, where the general principles of metabolic engineering have already been established [25,26], this is not the case for efflux. One reason is that the efflux mechanisms can be very different (Figure 9.1). A further reason is that the efflux behavior could be determined by the most varied structural components of the cell wall. In addition, each amino acid naturally displays its own special properties, thus preventing any generalization. The carriers are also of different significance in physiological terms. LysE is naturally designed to export basic amino acids, thus representing in addition to the regulation of amino acid synthesis a new mechanism for controlling the intracellular amino acids concentration. However, LysE probably represents a special case. ThrE and BrnFE, in contrast, could naturally serve other functions. They do not seem to have evolved specifically for the purpose of expelling endogenously synthesized amino acids, but they probably do so as a fortuitous side-effect. Even further functionally apart is the "cysteine" exporter YdeD from E. coli, which is decisive for extracellular cysteine accumulation [20]. It does not export cysteine, but rather an adduct of cysteine with ketones or aldehydes, such as pyruvate and glyoxylate, to produce thiazolidine derivatives, which are excreted. This illustrates that a great many novel carriers and also efflux-determining properties can still be expected for amino acid production with C. glutamicum.

ACKNOWLEDGMENTS I thank Degussa and H. Sahm for their constant support of the work, as well as the many co-workers involved in studies on bacterial amino acid synthesis and export.

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37. Haier B. (2001) Funktionelle Analyse des Lysin-Exportcarriers aus Corynebacterium glutamicum. Dissertation, Universität zu Köln. 38. Harel-Bronstein M, Dibrov P, Olami Y, Pinner E, Schuldiner S, and Padan E. (1995) MH1, a second-site revertant of an Escherichia coli mutant lacking Na+/H+ antiporters regains Na+ resistance and a capacity to excrete Na+ in a μH+-independent fashion. J. Biol. Chem. 270:3816–3822. 39. Heidelberg JF, Paulsen IT, Nelson KE, Gaidos EJ, Nelson WC, Read TD, Eisen JA, Seshadri R, Ward N, Methe B, Clayton RA, Meyer T et al. (2002) Genome sequence of the dissimilatory metal ion-reducing bacterium Shewanella oneidensis. Nat. Biotechnol. 20:1118–1123. 40. Henickoff S, Haughn GW, Calvo JM, and Wallace JC. (1988) A large family of activator proteins. Proc. Natl. Acad. Sci. USA 85:6602–6606. 41. Hermann T and Krämer R. (1996) Mechanism and regulation of isoleucine excretion in Corynebacterium glutamicum. Appl. Environ. Microbiol. 62:3238–3244. 42. Hirasawa T, Wachi M, and Nagai K. (2000) A mutation in the Corynebacterium glutamicum ltsA gene causes susceptibility to lysozyme, temperature-sensitive growth, and L-glutamate production. J. Bacteriol. 182:2696–2701. 43. Hoischen C and Krämer R. (1989) Evidence for an efflux carrier system involved in the secretion of glutamate by Corynebacterium glutamicum. Arch. Microbiol. 151:342–347. 44. Hoischen C and Krämer R. (1990) Membrane alteration is necessary but not sufficient for effective glutamate secretion in Corynebacterium glutamicum. J. Bacteriol. 172:3409–3416. 45. Hosie AHF, Allaway D, Jones MA, Walshaw DL, Johnston AWB, and Poole PS. (2001) Solute-binding protein-dependent ABC transporters are responsible for solute efflux in addition to solute uptake. Mol. Microbiol. 40:1449–1459. 46. Ikeda M and Katsumata R. (1995) Tryptophan production by transport mutants of Corynebacterium glutamicum. Biosci. Biotechnol. Biochem. 59:1600–1602. 47. Ikeda M. (2003) Amino acid production processes. Adv. Biochem. Eng. 79:2–35. 48. Ishida M, Kawashima H, Sato K, Hashiguchi K, Ito K, Enei H, and Nakamori S. (1994) Factors improving L-threonine production by a three L-threonine biosynthetic genes-amplified recombinant strain of Brevibacterium flavum. Agric. Biol. Chem. 58:768–770. 49. Jäger W, Kalinowski J, and Pühler A. (1997) A Corynebacterium glutamicum gene conferring multidrug resistance in the heterologous host Escherichia coli. J. Bacteriol. 179:2449–2451. 50. Jarlier V and Nikaido H. (1990) Permeability barrier to hydrophilic solutes in Mycobacterium chelonei. J. Bacteriol. 172:1418–1423. 51. Jarlier V and Nikaido H. (1994) Mycobacterial cell wall: Structure and role in natural resistance to antibiotics. FEMS Microbiol. Lett. 123:11–18. 52. Kawahara Y, Takahashi-Fuke K, Shimizu E, Nakamatsu T, and Nakamori S. (1997) Relationship between the glutamate production and the activity of 2-oxoglutarate dehydrogenase in Brevibacterium lactofermentum. Biosci. Biotechnol. Biochem. 61:1109–1112. 53. Kelle R, Laufer B, Brunzema C, WeusterBotz D, Kramer R, and Wandrey C. (1996) Reaction engineering analysis of L-lysine transport by Corynebacterium glutamicum. Biotechnol. Bioeng. 51:40–50. 54. Kennerknecht N, Sahm H, Yen MR, Patek M, Saier Jr MH Jr, and Eggeling L. (2002) Export of L-isoleucine from Corynebacterium glutamicum: a two-gene-encoded member of a new translocator family. J. Bacteriol. 184:3947–3956.

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55. Kim H-J, Kim Y, Lee M-S, and Lee H-S. (2001) Gene lmrB of Corynebacterium glutamicum confers efflux-mediated resistance to lincomycin. Mol. Cells 12:112–116. 56. Kimura E, Abe C, Kawahara Y, Nakamatsu T, and Tokuda H. (1997) A dtsR genedisrupted mutant Brevibacterium lactofermentum requires fatty-acids for growth and efficiently produces L-glutamate in the presence of an excess biotin. Biochem. Biophys. Res. Commun. 234:157–161. 57. Kimura E, Yagoshi C, Kawahara Y, Ohsumi T, Nakamatsu T, and Tokuda H. (1999) Glutamate overproduction in Corynebacterium glutamicum triggered by a decrease in the level of a complex comprising DtsR and a biotin-containing subunit. Biosc. Biotech. Biochem. 63:1274–1278. 58. Kimura E. (2003) Metabolic engineering of glutamate production. Adv. Biochem. Eng. Biotechnol. 79:37–57. 59. Kinoshita S, Udaka S, and Shimono M. (1957) Studies on the amino acid fermentation. Production of L-glutamate by various microorganisms. J. Gen. Appl. Microbiol. 3:193–205. 60. Krämer R. (1994) Secretion of amino acids by bacteria: physiology and mechanism. FEMS Microbiol. Rev. 13:75–94. 61. Krings E. (2003) Genomweite Effekte des Transkriptionsregulators LysG in Corynebacterium glutamicum. Dissertation, Universität Düsseldorf. 62. Kronemeyer W, Peekhaus N, Krämer R, Sahm H, and Eggeling L. (1995) Structure of the gluABCD cluster encoding the glutamate uptake system of Corynebacterium glutamicum. J. Bacteriol. 177:1152–1158. 63. Kruse D, Krämer R, Eggeling L, Rieping M, Pfefferle W, Tchieu JH, Chung YJ, Jr Saier MH, and Burkovski A. (2002) Influence of threonine exporters on threonine production in Escherichia coli. Appl. Microbiol. Biotechnol. 59:205–210. 64. Lambert C, Erdmann A, Eikmanns M, and Krämer R. (1995) Triggering glutamate excretion in Corynebacterium glutamicum by modulating the membrane state with local anesthetics and osmotic gradients. Appl. Env. Microbiol. 61:4334–4342. 65. Lapujade P, Goergen J, and Engasser J-M. (1999) Glutamate excretion as a major kinetic bottleneck for the thermally triggered production of glutamic acid by Corynebacterium glutamicum. Metabol. Eng. 1:255–261. 66. Leuchtenberger W. (1996). Amino acids — technical production and use. In Rehm HJ, Pühler A, Reed G, and Stadler PJW (Eds.), Biotechnology, Vol 6, VCH Verlagsgesellschaft, Weinheim, Germany, pp. 465–502. 67. Lichtinger T, Burkovski A, Niederweis M, Kramer R, and Benz R. (1998) Biochemical and biophysical characterization of the cell wall porin of Corynebacterium glutamicum: the channel is formed by a low molecular mass polypeptide. Biochemistry 37:15024–15032. 68. Lichtinger T, Riess FG, Burkovski A, Engelbrecht F, Hesse D, Kratzin HD, Krämer R, and Benz R. (2001) The low-molecular-mass subunit of the cell wall channel of the Gram-positive Corynebacterium glutamicum. Immunological localization, cloning and sequencing of its gene porA. Eur. J. Biochem. 268:462–469. 69. Luntz MG, Zhdanova NI, and Bourd GI. (1986) Transport and excretion of L-lysine in Corynebacterium glutamicum. J. Gen. Microbiol. 132:2137–2146. 70. Maes T, Vereecke D, Ritsema T, Cornelis K, Thu HN, Van Montagu M, Holsters M, and Goethals K. (2001) The att locus of Rhodococcus fascians strain D188 is essential for full virulence on tobacco through the production of an autoregulatory compound. Mol. Microbiol. 42:13–28. 71. Milner JL and Wood JM. (1987) Transmembrane amino acid fluxes in bacterial cells. CRC Crit. Rev. Biotechnol. 5:1–47.

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72. Mizukami T, Hamu A, Ikeda M, Oka T, and Katsumata R. (1994) Cloning of the ATP phosphoribosyl transferase gene of Corynebacterium glutamicum and application of the gene to L-histidine production. Biosci. Biotechnol. Biochem. 58:635–638. 73. Morbach S, Kelle R, Winkels S, Eggeling L, and Sahm H. (1996) Engineering the homoserine dehydrogenase and threonine dehydratase control points to analyse flux towards L-isoleucine in Corynebacterium glutamicum. Appl. Microbiol. Biotechnol. 45:612–620. 74. Morbach S, Sahm H, and Eggeling L. (1996) L-Isoleucine production with Corynebacterium glutamicum: further flux increase and limitation of export. Appl. Environ. Microbiol. 62:4345–4351. 75. Morbach S, Junger C, Sahm H, and Eggeling L. (2000) Attenuation control of ilvBNC in Corynebacterium glutamicum: evidence of leader peptide formation without the presence of a ribosome binding site. Biosci. Biotechnol. Biochem. 90:501–507. 76. Mori M and Shiio I. (1983) Glutamate transport and production in Brevibacterium flavum. Agric. Biol. Chem. (Japan) 47:983–990. 77. Morinaga Y, Takagi H, Ishida M, and Miwa K. (1987) Threonine production by coexistance of cloned genes-coding homoserine dehydrogenase and homoserine kinase in Brevibacterium lactofermentum. Agric. Biol. Chem. (Japan) 51:93–100. 78. Murata K, Mitsuoka K, Hirai T, Walz T, Agre P, Heymann JB, Engel A, and Fujiyoshi Y. (2000) Structural determinants of water permeation through aquaporin1. Nature 407:599–605. 79. Nakamori S, Ishida M, Takagi H, Ito K, Miwa K, and Sano K. (1987) Improved L-threonine production by the amplification of the gene encoding homoserine dehydrogenase in Brevibacterium lactofermentum. Agric. Biol. Chem. (Japan) 51:87–91. 80. Nakamura J, Izui H, and Nakamatsu T. (2001) Bacterium producing L-glutamic acid and method for producing L-glutamic acid. European Patent Application 1 174 508. 81. Nakao Y, Kikuchi M, Suzuki M, and Doi M. (1972) Microbial production of L-glutamate by glycerol auxotrophs. Part I. Induction of glycerol auxotrophs and production of L-glutamic acid from n-paraffins. Agric. Biol. Chem. 36:490–496. 82. Nakayama K. (1985) Lysine. In Moo-Young (Ed.), Comprehensive Biotechnology, Vol. 3, Pergamon Press, New York, pp. 607–620. 83. Nampoothiri KM, Hoischen C, Bathe B, Möckel B, Pfefferle W, Krumbach K, Sahm H, and Eggeling L. (2002) Expression of genes of lipid synthesis and altered lipid composition modulates L-glutamate efflux of Corynebacterium glutamicum. Appl. Microbiol. Biotechnol. 58:89–96. 84. Nara T, Sameljima H, and Kinoshita S. (1964) Effect og penicillin on amino acid fermentation. Agric. Biol. Chem. (Japan) 28:120–124. 85. Nelson RD, Kuan G, Saier MH Jr, and Montal M. (1999) Modular assembly of voltage-gated channel proteins: a sequence analysis and phylogenetic study. J. Mol. Microbiol. Biotechnol. 1:281–287. 86. Nikaido H. (1994) Prevention of drug access to bacterial targets: Permeability barriers and active efflux. Science 264:382–387. 87. Nottebrock D, Meyer U, Kramer R, and Morbach S. Molecular and biochemical characterization of mechanosensitive channels in Corynebacterium glutamicum (2003) FEMS Microbiol. Lett. 218:305–309. 88. Ohnishi J, Mitsuhashi S, Hayashi M, Ando S, Yokoi H, Ochiai K, Ikeda M. (2002) A novel methodology employing Corynebacterium glutamicum genome information to generate a new L-lysine-producing mutant. Appl. Microbiol. Biotechnol. 58:217–223.

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89. Okamoto K Kino K, Ikeda M. (1997) Hyperproduction of L-threonine by an Escherichia coli mutant with impaired L-threonine uptake. Biosci. Biotechnol. Biochem. 61:1877–1882. 90. Okazaki H, Kanzaki T, Doi M, Sumino Y, Fukuda H. (1967) L-Glutamic acid fermentation. II. The production of L-glutamic acid by an oleic-acid requiring mutant. Agric. Biol. Chem.Tokyo 31:1314–1317. 91. Palmieri L, Berns D, Kramer R, Eikmanns M. (1996) Threonine diffusion and threonine transport in Corynebacterium glutamicum and their role in threonine production. Arch. Microbiol. 165:48–54. 92. Puech V, Bayan N, Salim K, Leblon G, Daffe M. (2000) Characterization of the in vivo acceptors of the mycoloyl residues transferred by the corynebacterial PS1 and the related mycobacterial antigens 85. Mol. Microbiol. 35:1026–1041. 93. Puech V, Chami M, Lemassu A, Laneelle MA, Schiffler B, Gounon P, Bayan N, Benz R, Daffe M. (2001) Structure of the cell envelope of corynebacteria: importance of the non-covalently bound lipids in the formation of the cell wall permeability barrier and fracture plane. Microbiology 147:1365–1382. 94. Radmacher E, Stansen KC, Besra GS, Hollweg G, Wendisch VF, Sahm H, Eggeling L. (2003) Ethambutol, a cell wall inhibitor of Mycobacterium tuberculosis, causes glutamate efflux of Corynebacterium glutamicum and alters its genome-wide expressions. 95. Read TD, Peterson SN, Tourasse N, Baillie LW, Paulsen IT, Nelson KE, Tettelin H, Fouts DE, Eisen JA, Gill SR, Holtzapple EK, Økstad OA et al. (2003) The genome sequence of Bacillus anthracis Ames and comparison to closely related Nature 423:81–86. 96. Reinscheid DJ, Kronemeyer W, Eggeling L, Eikmanns BJ, Sahm H. (1994) Stable expression of hom-1-thrB in Corynebacterium glutamicum and its effect on the carbon flux to threonine and related amino acids. Appl. Environ. Microbiol. 60:126–132. 97. Saier MH Jr. (2000) Families of transmembrane transporters selective for amino acids and their derivatives. Microbiology 146:1775–1795. 98. Saier MH Jr. (2003) Tracing pathways of transport protein evolution. Mol. Microbiol. 48:1145–1156. 99. Seep-Feldhaus AH, Kalinowski J, Pühler A. (1991) Molecular analysis of the Corynebacterium glutamicum lysl gene involved in lysine uptake. Mol. Microbiol. 5:2995–3005. 100. Shibukwa M, Ohsawa T. (1966) L-Glutamic acid fermentation with molasses. part IV. Effect of saturated-unsaturated fatty acid ratio in the cell membrane fraction on the extracellular accumulation of L-glutamate. Agric. Biol. Chem. 30:750–758. 101. Shiio I, Mitsugi K, Tsunoda T. (1959) Bacterial formation of glutamic acid from acetic acid in the growing culture medium. J. Biochem. (Tokyo) 46:1665–1666. 102. Shiio I, Otsuko S, Takahasi M. (1962) Effect of biotin on the bacterial formation of glutamic acid. I: Glutamate formation and cellular permeability of amino acids. J. Biochem. (Tokyo) 51:56–62. 103. Shiio I, Otsuka S, Katsuya N. (1963) Cellular permeability and extracellar formation of glutamic acid in Brevibacterium flavum. J. Biochem. (Tokyo) 53:333–340. 104. Simic P, Sahm H, Eggeling L. (2001) L-threonine export: use of peptides to identify a new translocator from Corynebacterium glutamicum. J. Bacteriol. 183:5317–5324. 105. Simic P, Willuhn J, Sahm H, Eggeling L. (2002) Identification of glyA (encoding serine hydroxymethyltransferase) and its use together with the exporter ThrE to increase L-threonine accumulation by Corynebacterium glutamicum. Appl. Env. Microbiol. 68:3321–3327.

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106. Soskine M, Steiner-Mordoch S, Schuldiner S. (2002) Crosslinking of membraneembedded cysteines reveals contact points in the EmrE oligomer. Proc. Natl. Acad. Sci. USA, 99:12043–12048. 107. Stackebrandt E, Rainey FA, Ward-Rainey NL. (1997) Proposal for a new hierarchic classification system, Actinobacteria classis nov. Int. J. Syst. Bacteriol. 47:479–491. 108. Takinami K, Yoshii H, Tsura H, Okada H. (1965) Biochemical effects of fatty acid and its derivatives on L-glutamic acid fermentation. Agric. Biol. Chem. 29:351–359. 109. Takinami K, Yoshii H, Yamada Y, Okada H, Kinoshita K. (1968) Control of L-glutamic acid fermentation by biotin and fatty acid. Amino acid and Nucleic acid 18:120–160. 110. Tauch A, Götker S, Pühler A, Kalinowski J, Thierbach G. (2002) The 27.8-kb R-plasmid pTET3 from Corynebacterium glutamicum encodes the aminoglycoside adenyltransferase gene cassette aadA9 and the regulated tetracycline efflux system Tet 33 flanked by active copies of the widespread insertion sequence IS6100. Plasmid 48:117–129. 111. Tauch A, Hermann T, Burkovski A, Krämer R, Pühler A, Kalinowski J. (1998) Isoleucine uptake in Corynebacterium glutamicum ATCC 13032 is directed by the brnQ gene product. Arch. Microbiol. 169:303–312. 112. Tauch A, Krieft S, Pühler A, Kalinowski J. (1999) The tetAB genes of the Corynebacterium striatum R-plasmid pTP10 encode an ABC transporter and confer tetracycline, oxytetracycline and oxacillin resistance in Corynebacterium glutamicum. FEMS Microbiol. Lett. 173:203–209. 113. Tauch A, Pühler A, Kalinowski J, Thierbach G. (2000) TetZ, a new tetracycline resistance determinant discovered in gram-positive bacteria, shows high homology to gram-negative regulated efflux systems. Plasmid 44:285–291. 114. Tyrrell R, Verschueren KH, Dodson EJ, Murshudov GN, Addy C, Wilkinson AJ. (1997) The structure of the cofactor-binding fragment of the LysR family member, CysB: a familiar fold with a surprising subunit arrangement. Structure 5:1017–1032. 115. Tzvetkov M, Klopprogge C, Zelder O, Liebl W. (2003) Genetic dissection of trehalose biosynthesis in Corynebacterium glutamicum: inactivation of trehalose production leads to impaired growth and an altered cell wall lipid composition. Microbiology 149:1659–1673. 116. Udagawa K, Abe S, Kinoshita S. (1962) Effects of surface agents in L-glutamic acid fermentation. J. Biochem. Tokyo 40:614–619. 117. Udaka S. (1960) Screening method for microorganisms accumulating metabolites and its use in the isolation of Micrococcus glutamicus. J. Bacteriol. 79:754–755. 118. Vrljic M, Eggeling L, Sahm H. (1996) A new type of transporter with a new type of cellular function: L-lysine export from Corynebacterium glutamicum. Mol. Microbiol. 22:815–826. 119. Vrljic M, GargJ, Bellmann A, Wachu S, Freudl R, Malecki MJ, Sahm H, Kozina VJ, Eggeling L, Saier MH Jr. (1999) The LysE superfamily: Topology of the lysine exporter LysE of Corynebacterium glutamicum, a paradyme for a novel superfamily of transmembrane solute translocators. J. Mol. Microbiol. Biotechnol. 327–336. 120. Vrljic M, Kronemeyer W, Sahm H, Eggeling L. (1995) Unbalance of L-lysine flux in Corynebacterium glutamicum and its use for the isolation of excretion-defective mutants. J. Bacteriol. 177:4021–4027. 121. Wachi M, Nagai K. (1999) Penicillin binding protein gene and process for producing L-glutamic acid. European Patent Application EP 1 059 358 A1. 122. Walz T, Hirai T, Murata K, Heyman JB, Mitsuoka, K, Fujiyoshi Y, Smith BL, Agre P, Engel A. (1997) The three-dimensional structure of aquaporin-1. Nature 387:624–627.

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123. Wang X, Bogdanov M, Dowhan W. (2002) Topology of polytopic membrane protein subdomains is dictated by membrane phospholipid composition. EMBO J. 21:5673–5681. 124. Wolf A. (2002) Trehalosesynthese in Corynebacterium glutamicum. Dissertation University of Cologne 125. Wolf A, Kramer R, Morbach S. (2003) Three pathways for trehalose metabolism in Corynebacterium glutamicum ATCC13032 and their significance in response to osmotic stress. Mol. Microbiol. 49:1119–1134. 126. Yamada K, Kinoshita S, Tsunoda T, Aida K (Eds.). (1972) The microbial production of amino acids. Kodansha Ltd., Tokyo. 127. Yen MR, Tseng YH, Simic P, Sahm H, Eggeling L, Saier MH Jr. (2002) The ubiquitous ThrE family of putative transmembrane amino acid efflux transporters. Res. Microbiol. 153:19–25. 128. Zakataeva NP, Aleshin VV, Tokmakova LL, Troshin PV, Livshits VA. (1999) The novel transmembrane Escherichia coli proteins involved in the amino acid efflux. FEBS Let. 452:228–232. 129. Zittrich S, Krämer R. (1994) Quantitative discrimination of carrier-mediated excretion of isoleucine from uptake and diffusion in Corynebacterium glutamicum. J Bacteriol. 176:6892–6899.

Part V Physiology and Regulation

10

Central Metabolism: Sugar Uptake and Conversion A. Yokota and N.D. Lindley

CONTENTS 10.1 Introduction ..................................................................................................216 10.2 Sugar Uptake Systems .................................................................................216 10.2.1 Genome Analysis .............................................................................218 10.2.2 Metabolic Regulation .......................................................................219 10.3 Glycolysis.....................................................................................................219 10.3.1 Genetic Organization .......................................................................221 10.3.1.1 Fructose-1, 6-Bisphosphate Aldolase (fda) ......................221 10.3.1.2 Pyruvate Kinase (pyk).......................................................221 10.3.2 Enzyme Characterization .................................................................221 10.3.2.1 Glucose-6-Phosphate Isomerase.......................................221 10.3.2.2 6-Phosphofructokinase......................................................222 10.3.2.3 Fructose-1, 6-Bisphosphatase ...........................................222 10.3.2.4 Glyceraldehyde-3-Phosphate Dehydrogenase ..................222 10.3.2.5 Pyruvate Kinase ................................................................223 10.3.2.6 PEP Synthetase .................................................................223 10.4 The Pentose Phosphate Pathway .................................................................223 10.4.1 Genetic Organization .......................................................................225 10.4.2 Enzyme Characterization .................................................................228 10.4.2.1 Glucose-6-Phosphate Dehydrogenase ..............................228 10.4.2.2 6-Phosphogluconate Dehydrogenase................................229 10.4.2.3 Transketolase.....................................................................229 10.5 Functional Operation of Glycolysis and the Pentose Phosphate Pathway ....229 10.5.1 Effect of Carbon Sources on the Operation of the Glycolytic and Pentose Phosphate Pathways ....................................................229 10.5.2 Functional Operation of the Pentose Phosphate Pathway as Revealed by Mutant Analysis ..........................................................230 10.5.3 Flux Distribution in Glutamic Acid and Lysine Producers.............231 10.5.3.1 Flux Distribution in Glutamate Producers .......................231 10.5.3.2 Flux Distribution in Lysine Producers .............................231

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10.6 Control of Central Metabolism....................................................................232 10.6.1 Glycolysis.........................................................................................232 10.6.2 The Pentose Phosphate Pathway .....................................................233 10.7 Amino Acid and Nucleotide/Nucleoside Production and Central Metabolism...................................................................................................233 10.7.1 Engineering of Glycolysis ...............................................................233 10.7.1.1 Phosphoglucose Isomerase–Defective Mutant.................233 10.7.1.2 Pyruvate Kinase–Defective Mutants ................................234 10.7.2 Engineering of the Pentose Phosphate Pathway .............................234 10.7.2.1 Aromatic Amino Acid Production by C. glutamicum .....234 10.7.2.2 Purine Nucleotide/Nucleoside Production by C. ammoniagenes..............................................................235 10.8 Concluding Remarks....................................................................................236 Acknowledgment ...................................................................................................236 References..............................................................................................................236

10.1 INTRODUCTION Corynebacterium glutamicum and related species (e.g., C. efficiens and C. ammoniagenes) are used extensively by industry to produce a variety of amino acids and nucleotides at high yields from sugar substrates. These cellular building blocks draw carbon metabolites from central metabolism at different levels within the catabolic network and have an altered cofactor demand, both of which lead to carbon flux patterns explicitly different from those seen under pure growth conditions. Initial strain improvement programs for biotechnological exploitation of C. glutamicum were primarily aimed at genetic selection of strains with modified characteristics within the biosynthetic pathway of the specific product, notably by selection of gene products rendered less sensitive to feedback inhibition mechanisms [8]. This has led to efficient bioconversion processes, but it is now recognized that continued increases in performance also require fine-tuning of the central pathways to better balance the carbon precursor and cofactor requirements feeding the anabolic pathways. In this respect C. glutamicum has become a model organism in the study of metabolic flux analysis and many of the flux studies have gone some way toward overcoming an underlying lack of basic information concerning many of the catabolic reactions. Thus, in contrast with many microorganisms, pathway flux has been measured experimentally prior to the detailed study of individual reactions. In this chapter, the sugar uptake reactions and initial pathways involved in sugar catabolism (glycolysis and the pentose phosphate pathway) are reviewed.

10.2 SUGAR UPTAKE SYSTEMS Classical genetic and biochemical analysis of C. glutamicum indicated that this bacterium possesses at least three phosphotransferase systems (PTSs) enabling uptake of glucose, fructose, and sucrose (Figure 10.1). This was first established by Mori and Shiio [39] using a Brevibacterium flavum (C. glutamicum) ATCC14067 strain

Central Metabolism: Sugar Uptake and Conversion

PTS EII

Fru

Suc

?

PTS ptsS

Fru

PTS ptsF

HPr ptsH

Glc

Suc

Glc

Glc

PTS ptsG

Suc6P

P

glk

G6P

Fru

Pentose phosphate pathway

pgi

F6P

F1P pfkB

out membrane in

?

Glc

scrB EI pts1

PEP

217

fbp

pfk

FBP

Glycolysis FIGURE 10.1 Sugar transport systems of C. glutamicum. PTSGlc, glucose PTS; PTSFru, fructose PTS; PTSSuc, sucrose PTS; ?, unidentified transport system; Fru, fructose; Suc, sucrose; Glc, glucose; G6P, glucose-6-phosphate; F6P, fructose-6-phosphate; FBP, fructose-1,6-bisphosphate; Suc6P, sucrose-6-phosphate. Gene symbols are described in the text and in Table 10.1. The inset shows the phosphoryltransfer derived from PEP via the general PTS phosphotransferases I (EI) and HPr proteins shared by the three substrate specific EII PTS components.

that was shown to have both glucose and fructose PTS activities. They later obtained mutants lacking one or both of these activities [40]. In similar work with C. glutamicum evidence for at least three independent PTSs was obtained [4,6,35], as well as for a probable further permease transport system, at least as concerns glucose [2,3] (Figure 10.1). No biochemical data were available to substantiate this claim, though continued uptake of glucose in genetic backgrounds lacking PTS activity lends circumstantial support to this claim. Recently, this permease activity has been shown to have some importance in biotechnological applications, as it does not appear to be sensitive to high osmotic potential, known to inhibit PTS activity under industrial fermentation conditions [14]. Interestingly, glucose kinase activity was detected [40] and the gene (glk) encoding this activity is described [45] (Figure 10.1), giving further support for entry, or at least occurrence, of free glucose in the cell. Sucrose uptake appears to be somewhat unusual as mutants lacking PTSFru activity accumulate fructose extracellularly [4] (Figure 10.1), and this together with the absence of any fructokinase activity indicates that fructose is exported out of the cell by an unidentified sugar carrier system prior to fructose phosphorylation

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coupled to uptake via the PTS (Figure 10.1). Again definitive molecular data to substantiate this hypothesis are not yet available. Prior to the publication of the C. glutamicum ATCC13032 genome sequence [20,28] only a mannose-specific EII gene [33], postulated and later confirmed to be involved in glucose uptake [34], and the gene encoding the EI component of the PTS (Figure 10.1) had been described [32]. In parallel, biochemical characterization of EI and HPr [44] has indicated molecular masses of 65,000 and 14,000, respectively.

10.2.1 GENOME ANALYSIS Analysis of the genome sequence has confirmed that genes having high similarity for the three PTSs (glucose, fructose, and sucrose) are present together with an additional PTS permease and genes of as yet unknown function (Table 10.1). The search for the general PTS components EI and HPr, have revealed ptsI and, close by but transcribed in the opposite direction, a ptsH-like ORF (Figure 10.2). These genes are separated by three other ORFs encoding a putative transcriptional regulator, a 1-phosphofructokinase (pfkB) and a putative fructose-specific enzyme II (ptsF). This clustering of pfkB and ptsF is classical and also present in the other Corynebacterium species, although ptsF in C. diphtheriae seems to be mutated (Figure 10.2). The conserved locus is explained by the fact that the PTSFru generates fructose1-phosphate (F1P), which enters glycolysis at the level of fructose-1,6-bisphosphate (FBP) after further phosphorylation via 1-phosphofructokinase [6] (Figure 10.1), as is frequently observed in other organisms taking up fructose by the PTSFru. Similarly, a gene showing virtual homology with the ptsM gene described by Lee et al. [34] has been found within the genome sequence though this gene has been renamed

TABLE 10.1 Identified and Annotated Phosphotransferase System (PTS) Components and Sugar-Activating Kinases in C. glutamicum Enzyme PTS, glucose-specific IIABC component, PTSGlu PTS, fructose-specific IIABC component, PTSFru PTS, sucrose-specific IIABC component, PTSSuc PTS, EI component (enzyme I) PTS, HPr-related protein Fragment of Phosphotransferase system II component Fragment of Phosphotransferase system II component Phosphotransferase system II 1-Phosphofructokinase 1-Phosphofructokinase Glucokinase a

According to systematic gene name provided by NCBI.

Gene

Locus Taga

Length (Aa)

ptsG (M) ptsF ptsS ptsI ptsH

NCgl1305 NCgl1861 NCgl2553 NCgl1858 NCgl1862 NCgl2613 NCgl2614 NCgl2934 NCgl1860 NCgl1857 NCgl2105

684 689 662 569 90 84 89 271 331 321 324

pfkB glk

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FIGURE 10.2 Genomic organization of the conserved locus responsible for fructose uptake. ptsI encodes the general EI enzyme, ptsF the specific EII enzyme of PTSFru, pfkB the 1-phosphofructokinase, and ptsH the general HPr protein. Upstream of ptsH in C. glutamicum and C. efficiens a gene of unknown function is located, but which is probably also a fructokinase. The genes marked grey encode regulators.

ptsG because it encodes the principle uptake system for glucose. A third EII gene (ptsS) has been found with high similarity to sucrose-specific EII and located next in the genome to a sucrose-6-phosphate hydrolase (scrB). In addition to these specific systems, which could be anticipated from previous experimental evidence, two additional EII-like genes have been detected, encoding putative EIIC and EIIA proteins. These genes are clustered with an ABC-type transporter whose substrate specificity has yet to be determined.

10.2.2 METABOLIC REGULATION In many bacteria, PTS uptake is clearly linked directly to catabolite repression [56] and in low-G+C Gram-positive bacteria this mechanism involves phosphorylation of the HPr protein [48]. However, it is now clear that this mechanism is not functional in high-G+C Gram-positive bacteria, though as yet no plausible alternative mechanism explaining the catabolite repression mechanism in these bacteria has been proposed. Unlike the carefully controlled expression of sugar uptake systems of many bacteria, the various PTSs of C. glutamicum are expressed constitutively [3] and enable simultaneous co-consumption of sugars from mixtures. The sugar-specific activity is, at best, twofold increased in the presence of the respective sugar substrate. Thus, C. glutamicum appears to have evolved as an opportunist organism able to transport a number of sugars rather than as a specialist with a metabolism fine-tuned to enable efficient transport of a single component of a sugar mixture.

10.3 GLYCOLYSIS After sugar uptake and phosphorylation, further metabolism of the sugar phosphate occurs via both the classical central metabolic pathways: glycolysis and the pentose phosphate pathway (Figure 10.3). Activities of several glycolytic enzymes of C. glutamicum ATCC14067 were first reported by Shiio and co-workers [52]. Although glycolysis is the most important sequence of fueling reactions, relatively few of the glycolytic enzymes have been characterized biochemically or genetically in C. glutamicum (Table 10.2).

fda tpi gapA gapB pgk pgm eno pyk pps

Fructose bisphosphate aldolase (EC 4.1.2.13) Triosephosphate isomerase (EC 5.3.1.1) Glyceraldehyde-3-phosphate dehydrogenase (EC 1.2.1.12) 3-Phosphoglycerate kinase (EC 2.7.2.3) Phosphoglycerate mutase (EC 5.4.2.1) Enolase (EC 4.2.1.11) Pyruvate kinase (EC 2.7.1.40) Phosphoenolpyruvate synthetase (EC 2.7.9.2)

e

d

c

b

NCgl2673 NCgl1524 NCgl1526 NCgl0900 NCgl1525 NCgl0390 NCgl0935 NCgl2008 NCgl0528

NCgl0817 NCgl1202 NCgl0976

Locus taga

According to systematic gene name provided by NCBI. Calculated relative molecular mass. Oligomeric structure of the enzyme was described when available. Positive(+) and negative(-) effectors. Not conducted or described.

pgi pfkB fbp

Glucose-6-phosphate isomerase (EC 5.3.1.9) 6-Phosphofructokinase (EC 2.7.1.11) Fructose-1,6-bisphosphatase (EC 3. 1. 3. 11)

a

Gene

Enzyme

Yes Yes Yes — Yes — — Yes — (59,912)

(42,654)

(37,092) (27,198) (36,204)

—e — Yes

Cloning (cal. Mr)b

Genetic Characterization

TABLE 10.2 Characterization of Glycolytic/Gluconeogenetic Enzymes

Partial Partial Yes (35,500x4) Partial Yes (35,500) — — — — — — Yes (58,000x4) —

Purification (Mr) c 1.4(G6P), 0.54(F6P) 2.4(F6P), 0.15(ATP) 0.014(FBP) 0.022(FBP) — — — — — — — 1.2(PEP), 0.07(ADP) 0.4(ATP)

Km (mM)

Effectord (–)E4P Not found (–)AMP, PEP (–)AMP, S7P — — — — — — — (+)AMP, (–)ATP (+)ATP, (–)AMP, PEP

Biochemical Characterization

26, 43 27

60 59 46 60 62 7 7,20 20 7

Ref.

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10.3.1 GENETIC ORGANIZATION After the complete genome sequencing of C. glutamicum, ORFs of all the enzymes for glycolysis/gluconeogenesis were allocated on the chromosome by cloning and expression or by sequence similarity search as summarized in Table 10.2. Unlike E. coli, C. glutamicum possesses single genes for 6-phosphofructokinase and pyruvate kinase but two genes for glyceraldehydes-3-phosphate dehydrogenase, presumably with different physiological functions, gapA for glycolysis and gapB for gluconeogenesis [16,20], as is also the case in Bacillus subtilis [9]. Investigation of the cofactor signatures of the two glyceraldehyde-3-phosphate dehydrogenases suggested that the gapA product is probably NAD-dependent, while the gapB product is possibly NADP-dependent [20]. This putative NADP-generating reaction within glycolysis could be a target for the strain improvement of the amino acid producers, although transcription is probably strongly repressed by sugars [16]. An operon structure regrouping gapA-pgk-tpi-ppc was described with complex transcriptional control [7,50] (see also Chapter 5). 10.3.1.1 Fructose-1, 6-Bisphosphate Aldolase (fda) The first glycolytic gene cloned and expressed in C. glutamicum [62] was fda, which encodes fructose-1,6-bisphosphate aldolase. Primary structure homology suggested that the C. glutamicum aldolase was a class II aldolase. Analysis of the upstream region by S1-nuclease mapping identified the transcriptional start site, and the promoter region (–35 and –10) and Shine-Dalgarno sequence were deduced. 10.3.1.2 Pyruvate Kinase (pyk) The pyk gene has been cloned and sequenced from C. glutamicum [15,26]. Genome sequencing has confirmed that a single pyk gene is present, unlike with certain other microorganisms. Although the sequence has been examined in detail, no clear indication is available as to the manner in which the expression of this gene is regulated.

10.3.2 ENZYME CHARACTERIZATION The principal biochemical studies have been centered upon the irreversible reactions and to date five enzymes have been purified or partially purified and six enzymes were characterized biochemically (Table 10.2 and Figure 10.3). Characterization of these enzymes revealed some distinct features of regulation of glycolysis in C. glutamicum. 10.3.2.1 Glucose-6-Phosphate Isomerase Glucose-6-phosophate isomerase was partially purified from C. glutamicum ATCC14067, and found to be inhibited strongly by erythrose-4-phosphate (E4P) [60]. The inhibition was 93% at 1 mM of E4P under the assay conditions, suggesting that this mechanism will have a functional in vivo role. This inhibition was interpreted as a feedback control to prevent overproduction of E4P. It has been postulated that the metabolic flow downstream of fructose-6-phosphate (F6P) would not be

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disturbed by this inhibition because of the equilibration reached between F6P and E4P via the transketolase reaction, thereby feeding carbon from the pentose pathway back into glycolysis. The affinity constants of 1.4 mM and 0.54 mM for glucose-6phosphate (G6P) and F6P, respectively, are similar to those reported for other glucose-6-phosphate isomerases. Metabolite pool concentrations quantified during growth on various sugars indicate that this reaction is operating close to the thermodynamic equilibrium of the reaction [6,13] and is therefore unlikely to have any significant control over glycolytic flux. 10.3.2.2 6-Phosphofructokinase The 6-phosphofructokinase of C. glutamicum ATCC14067, unlike that described in many other forms of life, is neither inhibited by ATP, citrate, or phosphoenolpyruvate (PEP) nor activated by ADP, AMP, or fructose-2,6-bisphosphate [59]. Somewhat surprisingly, the 6-phosphofructokinase of C. glutamicum has been reported to be inhibited by ADP [59]. The lack of the normal allosteric control mechanisms explains not only why this enzyme was found to have little control over glycolysis but also why considerable carbon flux can be maintained through glycolysis for amino acid overproduction. 10.3.2.3 Fructose-1, 6-Bisphosphatase This enzyme is essential during gluconeogenesis, catalyzing the conversion of FBP into F6P. Analysis of the partially purified preparation from C. glutamicum ATCC14067 [60] revealed not only AMP but also sedoheptulose-7-phosphate (S7P) as enzyme inhibitors. At 1 mM concentrations, these compounds inhibit the enzyme activity almost completely. Inhibition by AMP is to prevent gluconeogenesis under energy-deficient conditions, as is the case in many other organisms. Inhibition by S7P was demonstrated for the first time in C. glutamicum, and interpreted as the means to feedback-inhibit excess formation of E4P and intermediates of the pentose phosphate pathway when grown on fructose, acetate, or other organic acids as a carbon source. Characterization of the purified enzyme from C. glutamicum ATCC13032 [46] revealed the tetrameric structure with the predicted molecular mass for the monomer of 35,500. As was found in C. glutamicum ATCC14067, AMP strongly inhibited the enzyme activity. PEP was also found to be an inhibitor for the enzyme from C. glutamicum ATCC13032. However, the physiological role of this PEP inhibition is difficult to interprete since PEP is an intermediate for gluconeogenesis. 10.3.2.4 Glyceraldehyde-3-Phosphate Dehydrogenase Although neither the NAD-dependent enzyme nor the postulated NADP-dependent enzyme has been purified, the literature does contain some important information concerning the manner in which this reaction is regulated by the redox charge of the cell. Thus, glyceraldehyde-3-dehydrogenase is strongly regulated by the NADH/NAD ratio [6] and may under circumstances in which the redox charge of the cells increases (oxygen limitation, high glycolytic flux, or overproduction of glutamate) have an important control over pathway flux, as has been described for

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Lactococcus lactis [10]. This control can in certain circumstances, e.g., growth on fructose [4], lead to an important increase in the triose-phosphate pool concentration and provoke dihydroxyacetone overflow. 10.3.2.5 Pyruvate Kinase To date, the most important control point recognized in glycolysis is pyruvate kinase. This enzyme was first characterized in a partially purified preparation from C. glutamicum ATCC14067 [43], and found to be an allosteric enzyme, as is also the case for rat liver, yeast, and E. coli, showing a sigmoidal curve for the initial rate of reaction against PEP concentration with a Hill coefficient of 3.0. AMP activated the enzyme reaction as observed in the case of E. coli PYK-II, but FBP, known as a positive effector in rat liver, yeast, and E. coli PYK-I, did not. About fourfold activation was observed in the presence of 2 mM AMP at 0.1 mM PEP. ATP and GTP were identified as negative effectors. Strong inhibition by ATP (87% by 1 mM ATP at 0.5 mM PEP) was observed. The inhibition by ATP was noncompetitive with respect to ADP. This means that the interaction with ATP does not take place at the active site of the enzyme, suggesting feedback inhibition rather than competitive product inhibition. The fact that this enzyme is regulated by adenine nucleotides, AMP, and ATP, clearly demonstrates the importance of this enzyme in the control of energy metabolism in C. glutamicum. Characterization of the purified enzyme originating from C. glutamicum ATCC13059 [26] elucidated its tetrameric structure as reported for E. coli PYK-I, and the enzymes of B. stearothermophilus, and L. lactis. The activation by AMP and inhibition by ATP were also confirmed with this purified enzyme. The apparent Km for PEP observed at 2 mM ADP was 1.2 mM; it increased to 2.8 mM in the presence of 2 mM ATP, and decreased to 0.4 mM in the presence of 2 mM AMP. 10.3.2.6 PEP Synthetase The activity of PEP synthetase has been detected in C. glutamicum ATCC13032, ATCC14067, and ATCC13869, although in relatively low specific activities [27]. mRNA expression was also found at relatively low levels in metabolic array analysis [16]. The activity was inhibited by AMP or PEP and activated by ATP. Cells cultured with lactate as a carbon source showed two to three times higher enzyme activity than that observed with glucose as a carbon source [27]. These data clearly indicate the gluconeogenetic nature of this enzyme.

10.4 THE PENTOSE PHOSPHATE PATHWAY The pentose phosphate pathway (Figure 10.3) forms a bypass of glycolysis, branching at G6P and refueling glycolysis at the levels of F6P and glyceraldehyde-3phosphate. The pentose phosphate pathway involves seven enzymes. The first three enzymes, glucose-6-phosphate dehydrogenase, 6-phosphogluconolactonase, and 6-phosphogluconate dehydrogenase, constitute an oxidative route in which G6P is converted into ribulose-5-phosphate (Ru5P) with the formation of 2 moles of NADPH. Since the 6-phosphogluconate dehydrogenase reaction evolves CO2, the

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Glucose ptsG -

Acetate

(-)NADPH, FBP, PRPP, OAA (-)NADPH, ATP, Ru5P,FBP, GA3P, E4P, PRPP, OAA

- Acetate, Lactate, Citrate, Succinate zwf, opcA - Acetate pgl

G6P pgi

(-)E4P

6PGL

6PG NADPH CO2

NADPH

F6P

Xu5P ATP

fbp

(+)(-)None

(-)AMP, PEP, S7P

pfk

+

(+)(-)None

Acetate, Citrate

FBP

-

tkt - Acetate

Acetate

fda

DHAP

Ru5P rpe

rpi

R5P

tkt

Acetate

Acetate

S7P tal

E4P

gapA -

NADP+

Acetate, Lactate Gluconate - Acetate

GA3P -

GA3P

tpi

gnd +

F6P

Acetate

NADH

+ Acetate gapB

1,3BPG pgk

ATP

Shikimic acid

3PG pgm

Aromatic amino acids Aromatic vitamins

2PG eno (+)ATP (-)AMP, PEP

+

PEP

pps

Lactate

ATP

(+)AMP (-)ATP

pyk

ATP -

Acetate

Lactate

Pyruvate -

lct Acetate

FIGURE 10.3 Glycolysis and pentose phosphate pathway and their control in C. glutamicum. Activators (+) or inhibitors (–) of the enzyme activity are given in boxes. Enzyme activity is increased (⊕) or decreased () when grown on the indicated carbon source as compared to that measured with glucose as a carbon source. Transcriptome analysis revealed up-regulation + ) or down-regulation ( – ) when grown on acetate as a carbon source in comparison with ( transciption level obtained with glucose as a carbon source. Abbreviations: G6P, F6P; FBP as

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oxidative route is irreversible under physiological conditons. The rest of the enzymes including transketolase and transaldolase constitute a reversible nonoxidative route of isomerization, epimerization, and interconversion reactions involving triose, tetrose, pentose, and heptose phosphates as intermediates. The general role of the pentose phosphate pathway is to supply anabolic reducing power and precursor metabolites, i.e., NADPH, ribose-5-phosphate (R5P), and E4P, for the biosynthesis of building blocks. In C. glutamicum strains used for aromatic amino acid production, E4P is a major pathway precursor that needs to be produced in considerably higher amounts than during growth. A similar situation holds true in nucleotide- and nucleoside-producing C. ammoniagenes strains that have to replenish R5P as a starting material for nucleotide biosynthesis. Furthermore, the high NADPH requirement for production of many amino acids is such that considerable attention has been paid to the pentose phosphate pathway and the manner in which strain improvement programs can modify the repartition between glycolysis and the pentose-phosphate pathway.

10.4.1 GENETIC ORGANIZATION All the genes of the pentose phosphate pathway have been annotated on the genome of C. glutamicum as shown in Table 10.3. No other sequence(s) with significant similarity to these genes was found in the genome, suggesting the presence of each gene as a single copy in C. glutamicum. In all Corynebacterianeae with annotated genome sequences available, including C. glutamicum, C. diphteriae, C. efficiens, Mycobacterium tuberculosis, M. leprae, M. bovis, M. marinum, and also in C. ammoniagenes, the five genes tkt, tal, zwf, opcA, and pgl encoding transketolase, transaldolase, glucose-6-phosphate dehydrogenase, and 6-phosphogluconolactonase are clustered in this order on the chromosome and very likely form an operon [30]. Only transketolase from C. glutamicum ATCC31833 has been cloned, and its promoter region, ribosomal binding site, start codon, terminator region, and cofactorand substrate-binding sites were deduced from the sequences of transketolases from other microbes [18]. In Southern blot analysis, the cloned transketolase fragment hybridized with genomic DNA digest of ATCC31833 to give a single positive band [21]. This confirmed the presence of a single copy of the transketolase gene on the chromosome. Genetic organization of the two glucose-6-phosphate dehydrogenase genes seems interesting, although it is not yet conclusive. As will be described below, glucose-6-phosphate dehydrogenase from C. glutamicum was found to be a heteromultimeric protein consisting of a well-known zwf product and an additional opcA product [41]. Thus, this enzyme must be coded by zwf and opcA. Although the

FIGURE 10.3 (continued) in legend to Figure 10.2; GA3P, glyceraldehyde-3-phosphate; DHAP, dihydroxyacetonephosphate; 1,3BPG, 1,3-bisphosphoglycerate; 3PG, 3-phosphoglycerate; 2PG, 2-phosphoglycerate; PEP, phosphoenolpyruvate; 6PGL, 6-phosphogluconolactone; 6PG, 6-phosphogluconate; Ru5P, ribulose-5-phosphate; R5P, ribose-5-phosphate; Xu5P, xylulose-5-phosphate; S7P, sedoheptulose-7-phosphate; E4P, erythrose-4-phosphate; PRPP, phosphoribosyl pyrophosphate; OAA, oxaloacetate. Gene symbols are according to Hayashi et al. [16] and listed in Table 10.1 and Table 10.2.

C. glutamicum Glucose-6-phosphate dehydrogenase (EC 1.1.1.49) 6-Phosphogluconolactonase (EC 3.1.1.31) 6-Phosphogluconate dehydrogenase (EC 1.1.1.44)

Enzyme

NCgl1514 NCgl1515 NCgl1516 NCgl1396

zwf opcA pgl gnd

Gene

Locus Taga

—

—e — — —

Cloning (cal. Mr)b

Genetic Characterization

0.034(6PG), 0.017(NADP+) 0.071(6PG), 0.043(NADP+) 0.045(6PG), 0.036(NADP+)

Yes(52,500 × 2) Partial

0.15(G6P), 0.037(NADP+) 0.14(G6P), 0.024(NADP+) —

Km (mM)

(–)NADPH, ATP, Ru5P, FBP, GA3P, E4P — (–)NADPH, FBP, PRPP, Ru5P, GA3P, OAA, E4P

1 57

41

58

(–)NADPH, OAA, FBP, PRPP —

Ref.

41

Effector d

(–)NADPH

Biochemical Characterization

Yes

Yes(60,000) Yes(30,000) Partial —

Purification (Mr)c

TABLE 10.3 Genetic and Biochemical Characterization of Pentose Phosphate Pathway Enzymes of C. glutamicum and C. ammoninagenes

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e

d

c

b

a

— — —

zwf tkt tal

NCgl1512

tkt NCgl1513

NCgl2337

rpi

tal

NCgl1536

rpe

Yes Yes

Yes

—

Yes(75,000)

—

—

According to systematic gene name provided by NCBI. Calculated relative molecular mass. Oligomeric structure of the enzyme was described when available. Positive(+) and negative(-) effectors. Not conducted or described.

Transaldolase (EC 2.2.1.2) C. ammoniagenes Glucose-6-phosphate dehydrogenase Transketolase Transaldolase

Ribulose-5-phosphate epimerase (EC 5.1.3.1) Ribose-5-phosphate isomerase (EC 5.3.1.6) Transketolase (EC 2.2.1.1)

— —

—

—

—

—

—

— —

—

— 0.11(Xu5P), 1.2(R5P) —

—

—

— —

—

— Not found —

—

—

30 30

30

18 60

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opcA product appeares to be essential for glucose-6-phosphate dehydrogenase activity in cyanobacteria [61], its role in C. glutamicum still needs to be elucidated.

10.4.2 ENZYME CHARACTERIZATION As summarized in Table 10.3, so far, two enzymes in the oxidative route of the pentose phosphate pathway have been purified and characterized. 10.4.2.1 Glucose-6-Phosphate Dehydrogenase The activity of this enzyme was first reported in C. glutamicum ATCC14067 by Shiio and co-workers [52], suggesting the presence of the pentose phosphate pathway in this microorganism. However, it was necessary to wait more than 40 years for the successful purification and characterization of this enzyme [41]. Purification of glucose-6-phosphate dehydrogenase from C. glutamicum ATCC13032 disclosed the very unique structure of this enzyme. It appears that the enzyme forms a heteromultimeric complex consisting of two different proteins, i.e., a well-known zwf product and an additional opcA product. NADPH was demonstrated to be a physiologically important inhibitor with a Ki value of 0.02 mM. As a reaction product, NADPH inhibits competitively with respect to NADP+. In a previous study with a partially purified preparation from C. glutamicum ATCC14067, Sugimoto and Shiio [58] showed similar Km values for both substrates with those of C. glutamicum ATCC13032. However, regulatory properties of the glucose-6-phosphate dehydrogenase from ATCC14067 showed significant differences. Although NADPH showed the strongest inhibition among the metabolites tested, oxaloacetic acid (OAA) also served as a strong inhibitor with 50% inhibition obtained with 0.09 mM OAA. FBP and phosphoribosyl pyrophosphate (PRPP) were also reported as inhibitors but less effective, though it should be borne in mind that FBP intracellular concentrations have been shown to be extremely high in C. glutamicum [13]. The inhibition by OAA, FBP, and PRPP was lost by repeated ammonium sulfate precipitation, suggesting that the inhibitors bind not to the active center but to other site(s) labile to ammonium sulfate treatment. Sugimoto and Shiio also demonstrated the distribution of this OAA inhibition specifically in glutamateproducing coryneform bacteria including C. glutamicum ATCC13032, while inhibition by FBP and PRPP were found in various Gram-positive and -negative bacteria [58]. The discrepancy in OAA inhibition in these studies is not clear but might be related to the unique structure of this enzyme. Both glucose-6-phosphate dehydrogenases showed strict cofactor specificity for NADP+, and the reaction did not proceed with NAD+ at all. The inhibitory effect of FBP with partially purified glucose-6-phosphate dehydrogenase from an industrial glutamate-producing strain has recently been confirmed [12]. Kinetic constants were similar to those described by other workers, but a more detailed appraisal of the FBP inhibition enabled this to be characterized as a mixed inhibition with diminished substrate affinity and maximum velocity as a function of increasing FBP concentrations. Taking into account the intracellular metabolite concentrations, this effect was shown to be sufficient to account for the decreased

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flux observed during the glutamate production phase. In light of this, at least two distinct physiological control mechanisms are postulated to control enzyme activity, allowing direct control of the pentose phosphate pathway as a function of either overall carbon flux or anabolic reducing power availability. 10.4.2.2 6-Phosphogluconate Dehydrogenase Characterization with a partially purified preparation from C. glutamicum ATCC14067 [57], and that of purified enzyme from C. glutamicum ATCC13032 [41] and an industrial lysine-producer strain [1] showed similar properties for 6-phosphogluconate dehydrogenase. The studies revealed almost the same Km value for both 6-phosphogluconate (6PG) and NADP+, and have NADPH, FBP, Ru5P, GA3P, and E4P in common as inhibitors. As a reaction product, NADPH served as a potent competitive inhibitor of 6-phosphogluconate dehydrogenase and glucose-6-phosphate dehydrogenase as well. In the case of 6-phosphogluconate dehydrogenase from C. glutamicum ATCC14067, FBP also appeares to be as strong an inhibitor as NADPH. ATP, which often inhibits the enzyme reaction in other microorganisms, is an inhibitor in C. glutamicum ATCC13032 but not in C. glutamicum ATCC14067. Again OAA showed an inhibitory effect only in C. glutamicum ATCC14067. 10.4.2.3 Transketolase This enzyme was measured with crude extract from C. glutamicum ATCC14067, and some kinetic and regulatory properties were investigated [60]. No apparent inhibition of the enzyme activity was observed for the metabolites tested.

10.5 FUNCTIONAL OPERATION OF GLYCOLYSIS AND THE PENTOSE PHOSPHATE PATHWAY In this section, functional operation of these pathways in response to various carbon sources, roles of pentose phosphate pathway in catabolism and anabolism, and flux distribution in glutamic acid–producing wild-type strain and in lysine-producing mutants will be discussed.

10.5.1 EFFECT OF CARBON SOURCES ON THE OPERATION OF GLYCOLYTIC AND PENTOSE PHOSPHATE PATHWAYS

THE

Gene expression of enzymes in glycolysis and the pentose phosphate pathway (Figure 10.3) changes significantly when different carbon sources are supplied for growth. Recent transcriptome analysis of C. glutamicum ATCC13032 cells grown on acetate [11,16,42] clearly demonstrated down-regulation of glycolytic genes (pfk, gapA, pyk) and of pentose phosphate genes (zwf, gnd, tkt, tal) as compared to glucose-grown cells. Glucose-specific PTS EII gene (ptsG), pyruvate dehydrogenase gene (aceE), and lactate dehydrogenase gene (lct) were also down-regulated. In contrast, genes involved in acetate assimilation (ack and pta), glyoxylate shunt (aceA and aceB), and TCA cycle (gltA, acn, sdhA, sdhB, sdhCD, fumH, and mdh) were up-regulated.

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Gluconeogenic genes (pck and gapB) were also up-regulated. The surprising finding was the fourfold up-regulation of the newly identified putative gapB gene; in contrast, the already-known gapA was down-regulated (0.4-fold). These results strongly support the postulated role of these two isozymes, i.e., gapA works for glycolysis and gapB for gluconeogenesis. Enzyme activity measurements of the pentose phosphate pathway of C. glutamicum ATCC14067 showed decreased activity of glucose-6-phosphate dehydrogenases when grown on organic acids such as lactic, acetic, succinic, and citric acid, as compared to that of glucose-grown cells [58]. 6-Phosphogluconate dehydrogenase activity was twofold lower in acetate- and lactate-grown cells as compared to that of glucose-grown cells, while that of the gluconate-grown cells was approximately threefold higher, indicating the role of this pathway in glucose and gluconate assimilation [1,57]. As to glycolytic enzymes, no significant difference was observed in phosphoglucose isomerase (pgi) and phosphofructokinase (pfk) activities between the cells grown on glucose and the cells grown on acetate [60]. Gluconeogenic FBPase (fbp) activity was measured to be 2.6-fold higher in cells grown on acetate than that of glucose-grown cells in C. glutamicum ATCC14067 [60]. In C. glutamicum ATCC13032, the FBPase (fbp) of citrate-grown cells was up-regulated about twofold on the basis of activity measurement and relative abundance of protein in Coomassiestained 2D gels [46], confirming the rational response of this enzyme. Similarly, growth on fructose led to the enhanced expression of the 1-phosphofructokinase (pfkB) activity necessary to transform F1P to FBP [5], and to an increased level of fructose bisphosphate [53] (Figure 10.1). The diminished flux shown to pass through the pentose phosphate pathway under such conditions was however attributed to the sugar uptake characteristics and the extremely high FBP concentration, postulated to have an inhibitory effect on the glucose-6-phosphate dehydrogenase activity. It is interesting to note that the higher NADH concentration in fructose-grown cells provoked metabolite overflow with significant accumulation of both dihydroxyacetone and lactate, despite adequate aeration conditions [3].

10.5.2 FUNCTIONAL OPERATION OF THE PENTOSE PHOSPHATE PATHWAY AS REVEALED BY MUTANT ANALYSIS Analysis of a transketolase-defective mutant of C. glutamicum ATCC31833, RA60, that completely lost transketolase activity, defined several important roles of the pentose phosphate pathway in this organism [22]. Strain AR60 was successfully obtained by mutagenesis followed by selection for shikimic acid auxotrophy and nonassimilation of ribose. This mutant appeared to be unable to assimilate gluconic acid, and the growth was also complemented by the addition of three aromatic amino acids plus aromatic vitamins instead of shikimic acid. The spontaneous revertant that no longer showed shikimic acid auxotrophy recovered ability to assimilate ribose and gluconic acid as well as transketolase activity. These properties of the mutant defined at least the following properties of this pathway in C. glutamicum: (i) absolute dependence of formation of E4P on transketolase activity, (ii) existence of a single transketolase, and (iii) involvement of the oxidative pathway for ribose

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formation. The lack of gluconic acid assimilation also suggests the absence of a functional Entner-Doudoroff pathway for gluconic acid metabolism in C. glutamicum, which had already been speculated on the basis of the absence of enzyme activity of 2-keto-3-deoxygluconate-6-phosphate aldolase [62]. In C. ammoniagenes, transketolase-defective mutants also showed a requirement for shikimic acid for growth, confirming the dependency of E4P formation on transketolase activity [30].

10.5.3 FLUX DISTRIBUTION IN GLUTAMIC ACID AND LYSINE PRODUCERS Under normal growth conditions on glucose, the carbon flux distribution between glycolysis and the pentose phosphate pathway has been shown to be slightly in favor of the pentose phosphate pathway in C. glutamicum [2,36,47], unlike many microorganisms in which the glycolytic flux is significantly higher than that through the pentose phosphate pathway. In all industrial fermentation processes for amino acid production, the phase of exponential growth is blocked and gives way to a production phase in which amino acid overproduction is induced [17]. Thus one of the key factors studied over the last decade has been the manner in which central metabolic pathways readjust to these modified conditions. 10.5.3.1 Flux Distribution in Glutamate Producers During glutamate production, the NADPH demand can be met directly from the isocitrate dehydrogenase reaction, so it would appear logical that the flux through the pentose phosphate pathway should rapidly diminish as growth gives way to production. Data from 13C-labeling studies coupled to NMR analysis has given a variety of flux values [23,47,63] with no coherent consensus, until a recent study [12] that followed the pentose phosphate flux throughout the production phase enabled a clearer picture to be obtained. Labeling patterns clearly illustrated that the flux through the pentose pathway decreases throughout the production phase. This flux decrease was a direct function of the decreased NADPH demand, though the mechanism postulated to provoke this shift was linked to FBP concentrations, which increased as an inverse function of the pentose phosphate pathway flux. The discrepancies in previous reports were believed to be due to the duration of the incubation period. Thus, flux through the pentose phosphate pathway during the glutamate production phase is virtually null once stationary growth has been reached, and in this respect is similar to that observed under oxygen-limited growth conditions [5]. 10.5.3.2 Flux Distribution in Lysine Producers The situation is entirely different in lysine producers, in which the biosynthetic pathway has a considerably higher requirement for NADPH. In such strains various studies [23,36,55] have indicated that flux partitioning through the pentose phosphate pathway represents a majority of the flux available as G6P. This has led to the idea that lysine yields could be improved by further increasing this flux, or by decreasing

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the NADPH requirement. In an elegant experiment, this requirement was effectively diminished by replacing the NADPH-dependent glutamate dehydrogenase by a heterologous NADH-dependent enzyme [37]. However, no increase in lysine production was obtained and NMR analysis demonstrated that the flux through the pentose phosphate pathway decreased significantly. This has, in conjunction with the biochemical data [41], led to the concept that flux through the pentose phosphate pathway is determined by the NADPH demand, rather than the converse (in which flux through the pentose phosphate pathway would determine NADPH availability and hence directly affect lysine yields).

10.6 CONTROL OF CENTRAL METABOLISM Based on the available genetic and kinetic knowledge concerning enzymes of sugar uptake and metabolism, it is interesting to see how central metabolism could be optimized.

10.6.1 GLYCOLYSIS Enhancement of glycolysis is one of the most important issues of industrial microbiology for the improvement of productivity by fermentation. As mentioned in Section 10.3.2.3, the most important controlling point in glycolysis of C. glutamicum is considered to be at the level of pyruvate kinase, where the enzyme activity is inhibited by ATP and activated by AMP [26,43]. These findings suggest that glycolysis of C. glutamicum is controlled not directly by intermediary metabolites but rather by the energy status of the cell, and thus it might be expected that energy shortage would enhance glycolysis. An attempt to enhance glycolysis in C. glutamicum was reported by Sekine et al. [51] in which a mutant with reduced H+-ATPase activity (25% of the parent) was derived from C. glutamicum ATCC14067 by spontaneous mutation. The idea of this work was to provoke an energy shortage in C. glutamicum due to the impairment of the oxidative phosphorylation with a defective H+-ATPase. As expected, jarfermentation analyses revealed that the specific rate of glucose consumption during exponential growth of this mutant, F172-8, was 70% higher than that of the parent. The respiration rate of the mutant was also twofold higher, which seemed to be the indication of the enhanced reoxidation of NADH formed in excess along with the higher rate of glycolysis. Since the revertant of strain F172-8 with a comparable H+ATPase activity to the parent showed a similar fermentation profile to that of the original ATCC14067 strain, it was concluded that the enhanced glucose consumption rate was brought about by the defect in H+-ATPase. Sequence analysis of the atp operon genes coding for the H+-ATPase revealed a single base change in the gamma subunit of the H+-ATPase gene in strain F172-8, which was found to be changed to pseudo-wild-type in the revertant. Although the mechanism behind these physiological changes is still not clear, the fact that manipulation of energy metabolism by a H+-ATPase defect led to the enhancement of glycolysis is quite significant in light of the industrial importance of this microorganism. Similar effects of H+-ATPase

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defects have already been reported in E. coli [24,25,64,65], and in B. subtilis [48]. The cellular ATP demand as the determinant for the glycolytic flux has been demonstrated in E. coli [31]. It is interesting to note that many of the traditional mutationselection procedures used to optimize glutamate-producing strains use respiratory inhibitors and probably generate strains with increased rates of sugar catabolism due to modified energetic efficiency. Few if any of these mutants have been systematically analyzed, however.

10.6.2 THE PENTOSE PHOSPHATE PATHWAY As mentioned in Section 10.4.2, activities of enzymes in the oxidative route of this part of metabolism are regulated by various metabolites. Comparison of intracellular metabolite concentration with Ki of corresponding metabolites in C. glutamicum ATCC13032 revealed physiologically important inhibitors to be NADPH for both glucose-6-phosphate dehydrogenase and 6-phosphogluconate dehydrogenase, and ATP, FBP, and GA3P for 6-phosphogluconate dehydrogenase [41]. The important conclusion derived from the prediction of an in vivo activity of glucose-6-phosphate dehydrogenase by the use of the kinetic rate equation is that the enzyme activity is controlled more by the concentration ratio of NADP+ to NADPH ([NADP+]/[NADPH]) than by the concentration of G6P or NADPH itself. Since glucose-6-phosphate dehydrogenase is the key enzyme for the pentose phosphate pathway, it has been concluded that the flux of this pathway is determined by the [NADP+]/[NADPH] as well as by the specific activity of the enzyme [41].

10.7 AMINO ACID AND NUCLEOTIDE/NUCLEOSIDE PRODUCTION AND CENTRAL METABOLISM In this section, some examples of the manipulation of glycolytic and pentose phosphate pathways for improved productivity will be described.

10.7.1 ENGINEERING

OF

GLYCOLYSIS

10.7.1.1 Phosphoglucose Isomerase–Defective Mutant A mutant lacking phosphoglucose isomerase, which catalyzes interconversion between G6P and F6P, has been reported [38]. This mutant was derived from a C. glutamicum lysine-producer strain (DSM5715) by the disruption of the phosphoglucose isomerase gene by homologous recombination. The obtained mutant showed a lower level of growth and was retarded in glucose consumption, but produced lysine 1.7-fold higher than the parent strain. The by-product concentration was also dramatically reduced. The lysine yield (as HCl salt) of the mutant after 48-h culture in a medium containing 30 g/l of glucose was 7.19 g/l, while the parent produced only 4.76 g/l under the same conditions. In this mutant, G6P is fluxed exclusively into the pentose phosphate pathway, resulting in the abundant formation of intracellular NADPH. The mechanism enabling this improved yield of lysine remains to be elucidated.

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10.7.1.2 Pyruvate Kinase–Defective Mutants PEP is located at a branch point in metabolism, from which pyruvate kinase reaction (and PEP-dependent sugar uptake) can be seen as an energy-producing catabolic route, whereas PEP carboxylase replenishes the TCA cycle at the level of OAA, leading to the biosynthesis of aspartate-family amino acids. In lysine production by C. glutamicum, an increase in the supply of PEP as a precursor metabolite therefore is seen as being important for the enhancement of the lysine yield. In this context, pyruvate kinase–defective mutants of C. glutamicum lysine-producer strains have been derived and their capacities for lysine production have been evaluated. In the first trial conducted by Shiio and co-workers [54], three pyruvate kinase–defective mutants were obtained by mutagenesis from a lysine-producing mutant of C. glutamicum ATCC14067, strain No. 2-190, which has a feedback-resistant aspartokinase, a feedback-resistant PEP carboxylase, and low-level citrate synthase activity. Interestingly, all of the pyruvate kinase–defective mutants showed higher lysine productivity than that of the parent strain. The representative mutant, KL-18, showed comparable growth to the parent and produced 42.7 g/l of lysine as HCl salt, while the parent produced 38.0 g/l in shake-flask cultures. Another attempt was later made by Gubler et al. [15], in which pyruvate kinase-knockout mutants were elaborated by homologous recombination from an S-2-aminoethyl-L-cysteine-resistant lysineproducing C. glutamicum strain (B. lactofermentum ATCC21799). Although all the mutants obtained showed similar growth profiles and glucose consumption to those of the parent, their lysine productivities were 40% lower than that of the parent, with increased by-production of acetate. In addition, all the mutants were found to produce dihydroxyacetone and glyceraldehyde, which were not produced by the parent. The discrepancy between these two examples is not well understood, though it is probably linked to the genetic background of each strain. However, the presence of a feedback-resistant PEP carboxylase in strain No. 2-190 may be the key determinant, since the increased pool of PEP brought about by the pyruvate kinase defect would be replenished to OAA much more efficiently in this background than in a strain with a normal PEP carboxylase.

10.7.2 ENGINEERING

OF THE

PENTOSE PHOSPHATE PATHWAY

Since the pentose phosphate pathway supplies precursor metabolites for the biosynthesis of aromatic amino acids and nucleotides, appropriate manipulation of this pathway would be expected to improve the productivity of producers of these compounds. Such results were successfully achieved by Ikeda and co-workers through manipulation of transketolase activity in C. glutamicum and in C. ammoniagenes (see Chapter 21). 10.7.2.1 Aromatic Amino Acid Production by C. glutamicum In the case of aromatic amino acid production by C. glutamicum, increase in the activity of transketolase appeared to be effective for the production probably due to the increased supply of E4P, a precursor metabolite for aromatic amino acids biosynthesis [21]. In a tryptophan and lysine co-producer, overexpression of the transketolase

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gene by the introduction of recombinant plasmids of different copy number (low, medium, or high) resulted in a corresponding increase in the transketolase activity with concomitant increase and decrease in tryptophan yield and in lysine yield, respectively. The decreased yield of lysine was considered to be due to the decrease in the supply of PEP for aspartate biosynthesis. Introduction of the high–copynumber plasmid into three different strains producing phenylalanine, tyrosine, or tryptophan appeared to increase the transketolase activity about 10-fold, provoking increases in the yield of the respective aromatic amino acids by about 5 to 20%. The effects of transketolase overexpression were observed not only in the aforementioned laboratory-level producers, i.e., low-yield producers, but also in a hyperproducer capable of producing 50 g/l of tryptophan [19]. By the detailed analysis of fermentations, the defect of the hyperproducer KY9218 carrying a plasmid pKW9901 with desensitized tryptophan biosynthetic genes and a serine biosynthetic gene appeared to be the decrease in the tryptophan yield in the later stage of fermentation with concomitant increase in CO2 evolution. This seemed to be due to the changes in the carbon flow from the pentose phosphate pathway to the TCA cycle as described for a glutamate producer [14]. To fine-tune the carbon flow, a low–copy-number plasmid, pIK9960, carrying a transketolase gene and the same tryptophan and serine biosynthetic genes as pKW9901, was constructed. The strain KY9218 carrying plasmid pIK9960 showed a transketolase activity about three times higher than that of the host strain. An increase in the tryptophan yield and a decrease in the CO2 evolution were observed, especially in the late fermentation stage, and a tryptophan concentration as high as 58 g/l was attained. 10.7.2.2 Purine Nucleotide/Nucleoside Production by C. ammoniagenes In the case of purine nucleotide/nucleoside production by C. ammoniagenes mutants, transketolase activity was inversely related to nucleotide production [30]. When a plasmid containing the transketolase gene (tkt) from C. ammoniagenes was overexpressed in the inosine-producing strain KY13761 and in the 5′-xanthylic acid–producing strain KY13203, yields of these products were decreased by about 11% and 15%, respectively. On the other hand, disruption of the transketolase gene by homologous recombination in these strains resulted in the improvement of the production by about 11% and 28% respectively. These results suggests that in C. ammoniagenes, the oxidative route of the pentose phosphate pathway is important for the supply of R5P, a precursor metabolite for nucleotide biosynthesis, and that transketolase activity drains this pool back into glycolysis. Direct evidence for the significance of the oxidative route for the biosynthesis of purine nucleotides was obtained by the disruption of the glucose-6-phosphate dehydrogenase gene (zwf) by homologous recombination [29]. The disruptants from strain KY13761 and KY13203 produced about half the amounts of inosine and 5′-xanthylic acid, respectively. Amplification of zwf on a high–copy-number plasmid for enhanced production was tested in the transketolase-negative background, but was not successful and accompanied by growth retardation, suggesting the necessity for optimization of glucose-6-phosphate dehydrogenase activity [30].

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From these results, it can be concluded that elevation of transketolase activity in the nonoxidative route of the pentose phosphate pathway was effective for E4P supply for aromatic amino acid production in C. glutamicum, while the oxidative route of the pathway plays an important role in R5P supply for purine nucleotide/nucleoside production in C. ammoniagenes.

10.8 CONCLUDING REMARKS Understanding and manipulation of the two major sugar catabolism pathways (glycolysis and the pentose phosphate pathway) has been to some extent achieved and the gene library now available from the genome sequencing projects will undoubtedly see an increase in this metabolic engineering activity. Sugar uptake has received considerably less attention up to now, though this process is clearly of importance for rates of sugar conversion and, in certain cases, product yields. This is clearly the case for metabolites leaving central metabolism upstream of PEP, in which the essential flux for PTS sugar uptake will impose a major yield limitation. In a more general context, many of the details of gene expression are as yet descriptive; no essential control mechanism has been identified to explain the variations seen in gene expression on different substrates. This aspect will also benefit from the availability of modern post–genome-sequencing technology, which should facilitate the identification of regulatory control circuits. The combination of such information with existing biochemical data and the well-established flux analysis approaches developed around C. glutamicum should pave the way for metabolic modeling and in silico pathway design for a variety of products derived from the catabolic pathways. It will also help decipher some of the surprising metabolic reactions to strain development and hence increase our understanding of the complex control inherent to major central pathways. When such an integrated approach has been successfully initiated, the already significant biotechnological potential of C. glutamicum and related bacteria should be considerably enhanced.

ACKNOWLEDGMENT The authors thank Mr. Ryo Aoki for his excellent technical assistance in the preparation of this manuscript.

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35. Malin GM and Bourd GI. (1991) Phosphotransferase-dependent glucose transport in Corynebacterium glutamicum. J. Appl. Bacteriol. 71:517–523. 36. Marx A, de Graaf AA, Wiechert W, Eggeling L, and Sahm H. (1996) Determination of the fluxes in the central metabolism of Corynebacterium glutamicum by nuclear magnetic resonance spectroscopy combined with metabolite balancing. Biotechnol. Bioeng. 49:111–129. 37. Marx A, Eikmanns BJ, Sahm H, de Graaf AA, and Eggeling L. (1999) Response of the central metabolism in Corynebacterium glutamicum to the use of an NADHdependent glutamate dehydrogenase. Metabol. Eng. 1:35–48. 38. Marx A, Hans S, Möckel B, Bathe, B, and de Graaf AA. (2003) Metabolic phenotype of phosphoglucose isomerase mutants of Corynebacterium glutamicum. J. Biotechnol. 104:185–197. 39. Mori M and Shiio I. (1987) Pyruvate formation and sugar metabolism in an amino acid-producing bacterium, Brevibacterium flavum. Agric. Biol. Chem. 51:129–138. 40. Mori M and Shiio I. (1987) Phosphoenolpyruvate: sugar phosphotransferase systems and sugar metabolism in Brevibacterium flavum. Agric. Biol. Chem. 51:2671–2678. 41. Moritz B, Striegel K, De Graaf AA, and Sahm H. (2000) Kinetic properties of the glucose-6-phosphate and 6-phosphogluconate dehydrogenases from Corynebacterium glutamicum and their application for predicting pentose phosphate pathway flux in vivo. Eur. J. Biochem. 267:3442–3452. 42. Muffler A, Bettermann S, Haushalter M, Horlein A, Neveling U, Schramm M, and Sorgenfrei O. (2002) Genome-wide transcription profiling of Corynebacterium glutamicum after heat shock and during growth on acetate and glucose. J. Biotechnol. 98:255–268. 43. Ozaki H and Shiio I. (1969) Regulation of the TCA and glyoxylate cycles in Brevibacterium flavum. II. Regulation of phosphoenolpyruvate carboxylase and pyruvate kinase. J. Biochem. 66:297–311. 44. Parche S, Burkovski A, Sprenger GA, Weil B, Kramer R, and Titgemeyer F. (2001) Corynebacterium glutamicum: a dissection of the PTS. J. Mol. Microbiol. Biotechnol. 3:423–428. 45. Park SY, Kim HK, Yoo SK, Oh TK, and Lee JK. (2000) Characterization of glk, a gene coding for glucose kinase of Corynebacterium glutamicum. FEMS Microbiol. Lett. 188:209–215. 46. Rittmann D, Schaffer S, Wendisch VF, and Sahm H. (2003) Fructose-1,6-bisphosphatase from Corynebacterium glutamicum: expression and deletion of the fbp gene and biochemical characterization of the enzyme. Arch. Microbiol. 180:285–292. 47. Rollin C, Morgant V, Guyonvarch A, and Guerquin-Kern JL. (1995) 13C-NMR studies of Corynebacterium melassecola metabolic pathways. Eur. J. Biochem. 227:488–493. 48. Saier MH Jr, Chauvaux S, Cook GM, Deutscher J, Paulsen IT, Reizer J, and Ye JJ. (1996) Catabolite repression and inducer control in Gram-positive bacteria. Microbiology 142:217–230. 49. Santana M, Ionescu MS, Vertes A, Longin R, Kunst F, Danchin A, and Glaser P. (1994) Bacillus subtilis FoF1 ATPase: DNA sequence of the atp operon and characterization of atp mutants. J. Bacteriol. 176:6802–6811. 50. Schwinde JW, Thum-Schmitz N, Eikmanns BJ, and Sahm H. (1993) Transcriptional analysis of the gap-pgk-tpi-ppc gene cluster of Corynebacteium glutamicum. J. Bacteriol. 175:3905–3908. 51. Sekine H, Shimada T, Hayashi C, Ishiguro A, Tomita F, and Yokota A. (2001) H+ATPase defect in Corynebacterium glutamicum abolishes glutamic acid production with enhancement of glucose consumption rate. Appl. Microbiol. Biotechnol. 57:534–540.

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52. Shiio I, Otsuka S, and Tsunoda T. (1959) Glutamic acid formation from glucose by bacteria. I. Enzymes of the Embden-Myerhof-Parnas pathway, the Krebs cycle, and the glycoxylate bypass in cell extracts of Brevibacterium flavum No. 2247. J. Biochem. 46:1303–1311. 53. Shiio I, Sugimoto S, and Kawamura K. (1990) Effects of carbon source sugars on the yield of amino acid production and sucrose metabolism in Brevibacterium flavum. Agric. Biol. Chem. 54:1513–1519. 54. Shiio I, Yokota A, and Sugimoto S. (1987) Effect of pyruvate kinase deficiency on L-lysine productivities of mutants with feedback-resistant aspartokinases. Agric. Biol. Chem. 51:2485–2493. 55. Sonntag K, Schwinde J, de Graaf AA, Marx A, Eikmanns BJ, Wiechert W, and Sahm H. (1995) 13C NMR studies of the fluxes in the central metabolism of Corynebacterium glutamicum during growth and overproduction of amino acids in batch cultures. Appl. Microbiol. Biotechnol. 44:489–495. 56. Stülke J and Hillen W. (1999) Carbon catabolite repression in bacteria. Curr. Opin. Microbiol. 2:195–201. 57. Sugimoto S and Shiio I. (1987) Regulation of 6-phosphogluconate dehydrogenase in Brevibacterium flavum. Agric. Biol. Chem. 51:1257–1263. 58. Sugimoto S and Shiio I. (1987) Regulation of glucose-6-phosphate dehydrogenase in Brevibacterium flavum. Agric. Biol. Chem. 51:101–108. 59. Sugimoto S and Shiio I. (1989) Fructose metabolism and regulation of 1-phosphofructokinase and 6-phosphofructokinase in Brevibacterium flavum. Agric. Biol. Chem. 53:1261–1268. 60. Sugimoto S and Shiio I. (1989) Regulation of enzymes for erythrose 4-phosphate synthesis in Brevibacterium flavum. Agric. Biol. Chem. 53:2081. 61. Sundaram S, Karakaya H, Scanlan DJ, and Mann NH. (1998) Multiple oligomeric forms of glucose-6-phosphate dehydrogenase in cyanobacteria and the role of OpcA in the assembly process. Microbiology, 144: 1549–1556. 62. von der Osten CH, Barbas CF, Wong CH, Sinskey AJ. (1989) Molecular cloning, nucleotide sequence and fine-structural analysis of the Corynebacterium glutamicum fda gene: structural comparison of C. glutamicum fructose-1,6-bisphosphate adlolase to class I and class II adlolases. Mol. Microbiol. 3:1625–1637. 63. Walker TE, Han CH, Kollman VH, London RE, and Matwiyoff NA. (1982) 13C nuclear magnetic resonance studies of the biosynthesis by Microbacterium ammoniaphilum of L-glutamate selectively enriched with carbon-13. J. Biol. Chem. 257:1189–1195. 64. Yokota A, Amachi S, and Tomita F. (1999) Pyruvate, production using defective ATPase activity. In Flickinger MC and Drew SW (Eds.), Encyclopedia of Bioprocess Technology: Fermentation, Biocatalysis, and Bioseparation, John Wiley & Sons Inc. New York, pp. 2261–2268. 65. Yokota A, Terasawa Y, Takaoka N, Shimizu H, and Tomita F. (1994) Pyruvic acid production by an F1-ATPase-defective mutant of Escherichia coli W1485 lip2. Biosci. Biotechnol. Biochem. 58:2164–2167.

11

Central Metabolism: Tricarboxylic Acid Cycle and Anaplerotic Reactions B. Eikmanns

CONTENTS 11.1 Introduction ..................................................................................................242 11.2 Tricarboxylic Acid Cycle.............................................................................244 11.2.1 The Enzymes and Genes of the TCA Cycle and Their Regulation ........................................................................................245 11.2.1.1 Pyruvate Dehydrogenase Complex ..................................245 11.2.1.2 Citrate Synthase ................................................................249 11.2.1.3 Aconitase...........................................................................250 11.2.1.4 Isocitrate Dehydrogenase..................................................250 11.2.1.5 2-Oxoglutarate Dehydrogenase Complex ........................251 11.2.1.6 Succinyl-CoA Synthetase .................................................252 11.2.1.7 Succinate:Menaquinone Oxidoreductase..........................252 11.2.1.8 Fumarase ...........................................................................253 11.2.1.9 Malate Dehydrogenase and Malate:Quinone Oxidoreductase .................................................................253 11.2.2 Carbon Flux into and through the TCA Cycle ...............................254 11.2.3 Impact of TCA Cycle Reactions on Amino Acid Production ........256 11.3 Anaplerotic Reactions in Cells Growing on Carbohydrates .......................257 11.3.1 The Enzymes and Genes at the PEP-Pyruvate-Oxaloacetate Node and Their Regulation..............................................................258 11.3.1.1 PEP Carboxylase ..............................................................258 11.3.1.2 Pyruvate Carboxylase .......................................................258 11.3.1.3 PEP Carboxykinase ..........................................................259 11.3.1.4 Malic Enzyme...................................................................259 11.3.1.5 Oxaloacetate Decarboxylase.............................................260 11.3.2 Parallel and Bidirectional Carbon Fluxes at the PEP-PyruvateOxaloacetate Node ...........................................................................260 11.3.3 Impact of Anaplerotic Reactions on Amino Acid Production ........262 241

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11.4 Anaplerotic Reactions in Cells Growing on Substrates Other than Carbohydrates...............................................................................................264 11.4.1 The Enzymes and Genes of the Glyoxylate Cycle and Their Regulation ........................................................................................264 11.4.1.1 Isocitrate Lyase and Malate Synthase ..............................264 11.4.2 Carbon Fluxes through the Glyoxylate Cycle and Flux Control....265 11.5 Global Control of the TCA Cycle and Anaplerosis ....................................266 11.6 Concluding Remarks and Perspectives........................................................267 Acknowledgments..................................................................................................268 References..............................................................................................................268

11.1 INTRODUCTION Corynebacterium glutamicum is able to grow aerobically on a variety of carbohydrates, alcohols, and organic acids as single or combined sources of carbon and energy [53]. Invariably, independent of the carbon and energy sources used, the tricarboxylic acid (TCA) cycle (Figure 11.1), or at least parts of it, must be active [52]. One reason is that the TCA cycle serves catabolic and anabolic purposes as well [16,32]. On the one hand it is responsible for the complete oxidation of acetylCoA derived from the different substrates, it generates ATP (or GTP), and it provides reducing equivalents to membrane-bound respiratory systems. On the other hand it Glucose

Acetate

Glucose-6-P Glycolysis

Acetate

Gluconeo genesis

AK

Phosphoenolpyruvate CO2

PEPCk

Fum: fumarase

Pyruvate CO2

PEPCx

Acetyl-P

PDHC

ICD: isocitrate dehydrogenase ICL: isocitrate lyase

PTA

PCx

AK: acetate kinase CS: citrate synthase

PK

CO2

ACN: aconitase

Acetyl-CoA

MS: malate synthase

CS

MQO: malate: quinone oxidoreductase Aspartate

Oxaloacetate

OGDHC: 2-oxoglutarate dehydrogenase complex

Citrate ACN

MQO

Malate

Isocitrate

MS

Fum

Glyoxylate

PCx: pyruvate carboxylase PDHC: pyruvate dehydrogenase complex

Acetyl-CoA

ICD

Fumarate

PEPCk: PEP carboxykinase PEPCx: PEP carboxylase

ICL

PK: pyruvate kinase PTA: phosphotransacetylase

SQO

2-Oxoglutarate

Succinate SCS

SCS: succinyl-CoA synthetase SQO: succinate: menaquinone oxidoreductase

Succinyl-CoA

OGDHC

Glutamate

FIGURE 11.1 The anaplerotic reactions, the tricarboxylic acid cycle, glyoxylate bypass, and the reactions to activate acetate.

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243

provides precursor metabolites for biosynthetic processes such as 2-oxoglutarate and oxaloacetate. During growth on substrates entering the central metabolism at the level of acetyl-CoA, e.g., acetate, fatty acids, or ethanol, the glyoxylate cycle is active (Figure 11.1) [50]. This bypass of the TCA cycle avoids the oxidative decarboxylation steps of isocitrate dehydrogenase (ICD) and the 2-oxoglutarate dehydrogenase complex (OGDHC) and finally leads to the net formation of one molecule of malate from two molecules of acetyl-CoA. In order to replace the intermediates withdrawn for anabolism, the TCA cycle of growing cells has to be effectively and continuously replenished. This process is called anaplerosis and the reactions responsible are catalyzed by the so-called anaplerotic enzymes [49,116,129]. During growth on carbohydrates, anaplerosis in bacteria is accomplished by carboxylation of either phosphoenolpyruvate (PEP) or pyruvate to yield the TCA cycle C4-intermediate oxaloacetate. As will be outlined in Section 11.3.1, C. glutamicum possesses both carboxylating activities [61,78,80,99] and thus represents an exception. As opposed to the anaplerotic reactions active during growth on carbohydrates, during growth on substrates that enter central metabolism at the level of acetyl-CoA, the glyoxylate cycle functions also to provide oxaloacetate as an anaplerotic reaction [50]. It is therefore evident that the PEP-pyruvate-oxaloacetate node, or anaplerotic node, represents the link between glycolysis and the TCA cycle. The PEP-pyruvateoxaloacetate node is therefore highly relevant for the proper distribution of the carbon flux. At this node, the end products of glycolysis, PEP and pyruvate either (i) directly serve as precursor metabolites for anabolic purposes, (ii) enter the TCA cycle via acetyl-CoA, or (iii) enter the TCA cycle via the anaplerotic reactions. In addition, during growing on acetate, fatty acids, ethanol, or a TCA cycle intermediate, the PEP-pyruvate-oxaloacetate node is the starting point for gluconeogenesis. In C. glutamicum the initial reaction of gluconeogenesis is accomplished by a PEP carboxykinase, decarboxylating oxaloacetate to PEP (Figure 11.1) [39,77,87]. As will be outlined in Section 11.3.1, C. glutamicum possesses two further C4-decarboxylating enzymes [29,40,77]. Thus, alltogether at least five enzyme activities are present in C. glutamicum directly interconverting C3 with C4 units at the PEPpyruvate-oxaloacetate node. Since pyruvate kinase (PK) [30,38] and the pyruvate dehydrogenase complex (PDHC) are also present [104], C. glutamicum is characterized by a surprisingly diverse set of reactions at the anaplerotic node and the question for the significance of all these reactions for growth and amino acid production by C. glutamicum arises. Selected genes of the central metabolic pathways of C. glutamicum have been functionally characterized. Moreover, inspection of the recently establishment genome sequence (NC_003450, BX927147, [36,41,112]) reveals that in principle all genes encoding the enzymes for the TCA cycle, the glyoxylate cycle, and anaplerotic enzymes are present [41]. Whereas some of the genes are clustered, others are not, and clusters and genes are scattered throughout the genome (Figure 11.2). Furthermore, genome inspection revealed the existence of particular isoenzymes in central metabolism and also for an unexpected pyruvate:quinone oxidoreductase (also called pyruvate oxidase). This represents an additional enzyme at the PEP-pyruvate-oxaloacetate node.

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pck

malE lpd sdhCAB

pqo sucCD 3.0 Mb

0.5 Mb

mdh aceB, aceA aceE sucB

2.5 Mb

(3 282 708 bps)

pyk mqo

icd pyc

Genome map C. glutamicum 1.0 Mb

gltA

fum

2.0 Mb 1.5 Mb

odhA

ppc acn

FIGURE 11.2 Chromosomal localization of relevant genes encoding TCA cycle enzymes and enzymes at the PEP-pyruvate-oxaloacetate node.

Based on biochemical, genetic, and regulatory studies, as well as on quantitative determinations of metabolic fluxes and analysis of the genome sequence, this chapter summarizes what is known about the TCA cycle, the glyoxylate cycle, and the related pathways at the PEP-pyruvate-oxaloacetate node of C. glutamicum. This chapter will discuss organization and expression control of genes and the in vitro and in vivo activities and regulation of the enzymes involved. Furthermore, the specific contribution of enzymes and pathways to optimal growth and amino acid production will be discussed. It becomes evident that, although the central metabolic pathways follow the same theme as in many bacteria, characteristic features are present in C. glutamicum, representing fascinating and essential targets of metabolic engineering in order to achieve optimized amino acid production with this organism.

11.2 TRICARBOXYLIC ACID CYCLE In general, the fueling substrate for the TCA cycle is acetyl-CoA, which is derived from pyruvate when cells grow on C6 or C5 carbohydrates or on lactate. In aerobic organisms such as C. glutamicum, the pyruvate dehydrogenase complex (PDHC) catalyzes the oxidative decarboxylation of pyruvate, yielding acetyl-CoA, CO2 and reduced NAD. When C. glutamicum grows on acetate, acetyl-CoA is provided by the combined activities of acetate kinase (AK) and phosphotransacetylase (PTA) [86,97]. Acetyl-CoA is condensed with oxaloacetate to form citrate and CoA, and

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in a series of seven further reactions, the entire cycle is put into operation (Figure 11.1). The most important intermediates serving biosynthetic purposes are 2-oxoglutarate, the precursor for glutamate and its derivatives, and oxaloacetate, the precursor of aspartate and its derivatives.

11.2.1 THE ENZYMES AND GENES AND THEIR REGULATION

OF THE

TCA CYCLE

Already in 1959, Shiio et al. [98] proved most of the TCA cycle enzymes in cellfree extracts of C. glutamicum ssp. flavum. Later, selected enzymes have been studied in greater detail. The present state of knowledge on activity and regulation of each of the enzymes and on expression and regulation of the respective genes is discussed in this section, with the relevant characteristics summarized in Tables 11.1 and 11.2. 11.2.1.1 Pyruvate Dehydrogenase Complex The pyruvate dehydrogenase complex (PDHC) represents a member of a multienzyme complex family that also includes the 2-oxoglutarate dehydrogenase complex (OGDHC) and the branched-chain 2-oxoacid dehydrogenase complex. These complexes catalyze the oxidative decarboxylation of pyruvate, 2-oxoglutarate, and the 2-oxo acids of the branched-chain amino acids L-leucine, L-valine, and L-isoleucine, respectively. In general, the complexes consist of multiple copies of three different subunits, a thiamine pyrophosphate containing E1, a lipoic acid-containing E2, and a flavoprotein LPD (lipoamide dehydrogenase). Subunits E1 and E2 are specific for each of the three multienzyme complexes, whereas LPD is common to them in most organisms. Accordingly, E1 and E2 of the PDHC are denoted E1p (pyruvate dehydrogenase) and E2p (dihydrolipoamide acetyltransferase), and E1 and E2 of the OGDHC E1o (2-oxoglutarate dehydrogenase) and E2o (dihydrolipoamide succinyltransferase). In many organisms subunit E1p is split into two polypeptides, E1pα and E1pβ. Activity of the PDHC has been detected in various strains of C. glutamicum [10,11,93,104]. However, despite its crucial role, relatively little effort has been devoted to the study of the complex at the molecular and structural level. The activity of the PDHC depends on the substrates and the cofactors thiamine pyrophosphate and Mg2+, and addition of cysteine or DTT increases the activity of the complex of C. glutamicum [104]. From enzyme measurements with cells cultivated in a chemostat at various dilution rates on glucose or lactate, Lindley and co-workers concluded that, at least under their conditions, the complex is not subject to any significant regulation modulating its activity [10,11]. This seems surprising in view of the complex regulation of PDHCs in other bacteria as well as eukaryotes [35,96]. Since the C. glutamicum PDHC as a whole has not been purified and biochemically analyzed, the question of its control remains to be clarified. The annotation of the genome sequence of C. glutamicum revealed the presence of open reading frames with similarity to the E. coli aceE (E1p) and aceF (E2p) genes. However, these genes have not been studied and it remains to be proven whether they in fact represent the functional E1p and E2p genes. In contrast, a

aceE aceF lpd gltA acn icd odhA

sucB sucCD sdhCAB

1.2.4.1 2.3.1.12 1.8.1.4 4.1.3.7 4.2.1.3 1.1.1.42 1.2.4.2

2.3.1.61 6.2.1.5 1.3.5.1

Pyruvate dehydrogenase complex (PDHC), E1p subunit PDHC, E2p subunit PDHC and OGDHC, LPD subunit Citrate synthase (CS) Aconitase (ACN) Isocitrate dehydrogenase (ICD) OGDHC (oxoglutarate dehydrogenase complex) E1o subunit OGDHC E2o subunit Succinyl-CoA synthetase (SCS) Succinate: menaquinone oxidoreductase (SQO)

Gene(s)

EC No.

Protein 922 n.i.c 469 437 943 739 1257

676 402, 294 257, 673, 249

n.i.c n.i.c Monocistronic Monocistronic n.i.c Monocistronic n.i.c

n.i.c n.i.c n.i.c

cg2466 n.i.c cg0441 cg0949 cg1737 cg0766 cg1280

cg2421 cg2837, cg2836 cg0445, cg0446, cg0447

ID No.

Amino Acids per Monomera

Transcriptional Organization

70.9 42.2, 30.3 28.4, 74.7, 26.6

n.i.c 50.6 48.9 102 80.1 138.7

102.8

Molecular Mass of Monomer (kDa)b

41 41 41

93 24 41 23 115

41

Reference

TABLE 11.1 Enzymes and Genes of the TCA Cycle, Glyoxylate Cycle, and PEP-Pyruvate-Oxaloacetate Node in C. glutamicum

246 Handbook of Corynebacterium glutamicum

c

b

a

pyc pck malE (mez) n.i.c pyk pqo (poxB)

2.7.1.40

mdh aceA aceB ppc

1.1.1.37 4.1.3.1 4.1.3.2 4.1.1.31 6.4.1.1 4.1.1.32 1.1.1.40 4.1.1.3

fum mqo

4.2.1.2 1.1.99.16

cg2291 cg2891

cg0791 cg3169 cg3335 n.i.c

cg2613 cg2560 cg2559 cg1787

cg1145 cg2192

469 500 328 432 739 919 1140 610 392 n.i. 475 579

n.i.c n.i.c n.i.c Monocistronic Monocistronic In the operon pgk-tpi-ppc Monocistronic Monocistronic n.i.c n.i.c n.i.c n.i.c

59.2 62.0

123.1 66.9 40.9 31.7

34.9 47.2 82.4 103.2

49.8 54.8

Deduced from the nucleotide sequence BX927147. In some cases there are differences using the annotation of Nakagawa (NC_003450) Deduced from the amino acid sequence or from SDS PAGE given in the reference n.i., not identified/not known

Pyruvate carboxylase (PCx) PEP carboxykinase (PEPCk) Malic enzyme (ME) Oxaloacetate decarboxylase (OADCx) Pyruvate kinase (PK) Pyruvate: quinone oxidoreductase (PQO)

Fumarase (Fum) Malate: quinone oxidoreductase (MQO) Malate dehydrogenase (MDH) Isocitrate lyase (ICL) Malate synthase (MS) PEP Carboxylase (PEPCx)

38 41

78 87 29 40

59 84 85 22, 69

41 60

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TABLE 11.2 Specific Activities and Known Effectors of Enzymes of the TCA Cycle and the PEP-Pyruvate-Oxaloacetate Node Enzyme Pyruvate dehydrogenase complex (PDHC) Citrate synthase (CS) Aconitase (ACN) Isocitrate dehydrogenase (ICD) 2-Oxoglutarate dehydrogenase complex (OGDHC) Succinyl-CoA synthetase (SCS) Succinate: menaquinone oxidoreductase (SQO) Fumarase (Fum) Malate: quinone oxidoreductase (MQO) Malate dehydrogenase (MDH) Isocitrate lyase (ICL)

Malate synthase (MS) PEP carboxylase (PEPCx) Pyruvate carboxylase (PCx) PEP carboxykinase (PEPCk) Malic enzyme (ME)

Oxaloacetate decarboxylase (OADCx) Pyruvate kinase (PK) a

Spec. Activity a, in μmol min-1 (mg protein)-1

Effector(s)b

Reference

0.03 (MM + Lac or Glc)

n.k.c

10, 11

0.5 – 0.8 (MM + Glc, Lac, orAc) 0.25 (MM + glc) 0.9 – 1.1 (MM + Glc, Lac, orAc) 0.03 – 0.05 (MM + glc)

ATP (-); cis-aconitate (-); isocitrate (-) n.k.c oxaloacetate (-); glyoxylate (-)

24, 101

cis-aconitate (-); succinyl-CoA (-); NADPH (-); NADH (-); pyruvate (-); oxaloacetate (-); acetyl-CoA (+)

pd 0.03 (MM + glc); 0.05 (MM + Ac) pd 0.1 (MM + glc); 0.38 (MM + Lac) 0.2 (MM + Glc); 0.6 (MM + Ac) 0.01 (MM + Glc); 2.48 (MM + Ac)

0.04 (MM + glc); 2.21 (MM + Ac) 0.15 (MM + glc) 0.02 (MM + Glc); 0.05 (MM + Lac) 0.04 (MM + Glc); 0.14 (MM + Lac) 0.14 (MM + glc); 0.18 (MM + Lac); 0.03 (MM + Ac) 0.5 – 0.6 (CM) 0.96 (MM + glc)

23, 100 93, 106

98 oxaloacetate (-)

59, 92

n.k.c FAD + liposomes (+)

98 59, 60

n.k.c

59

3-phosphoglycerate (-); 6phoshogluconate (-); PEP (-); fructose-1,6-bisphosphate (-); succinate (-); glyoxylate (-) oxalate (-); glycolate (-); ATP (-) aspartate (-); acetyl-CoA (+); fructose-1,6-bisphoshate (+) ADP (-); AMP (-); Acetyl-CoA (-); aspartate (-) ATP (-)

70, 84

39, 87

oxaloacetate (-); glutamate (-)

29

ADP (-); CoA (-); succinate (-)

40

AMP (-); ATP (-)

30, 38

85 22, 71 47, 80

Measured in extracts from cells grown on complex medium (CM), minimal medium (MM) plus glucose (glc), acetate (Ac), or lactate (Lac) b (+) positive effector; (-) negative effector c Not known d p, present with significant activities

Central Metabolism: Tricarboxylic Acid Cycle and Anaplerotic Reactions

249

functional lpd gene encoding the active LPD protein of C. glutamicum has recently been characterized [93]. The lpd gene is monocistronic and not clustered with the genes for the other subunits of the PDHC or the OGDHC, as is generally the case in other bacteria. Another peculiarity of the lpd gene is that its transcription is initiated exactly at the nucleotide defined as its translational start (see Chapter 5). The LPD polypeptide shows up to 58% identity to LPD enzymes from other organisms and the purified protein was shown to catalyze the reversible reoxidation of dihydrolipoic acid and NADH:NAD+ transhydrogenation. Overexpression of the isolated lpd gene in C. glutamicum resulted in 12-fold higher LPD activity. However, the PDHC and OGDHC activities were unaffected [93]. Since lpd-deficient mutants have not yet been generated, it remains unclear whether the characterized LPD protein functions in C. glutamicum as the third subunit of only the PDHC, only the OGDHC, or both complexes. In this respect it is worth mentioning that in the C. glutamicum genome there is another gene (cg0790) coding for a protein with some similarity (up to 28% identity) to LPD proteins from other bacteria. 11.2.1.2 Citrate Synthase Citrate synthase (CS) catalyzes the initial reaction of the TCA cycle, i.e., the condensation of acetyl-CoA and oxaloacetate. The enzyme is considered to be ratecontrolling for the entry of substrates into the cycle and therefore provoked much interest in its structural, kinetic, regulatory, and molecular characteristics. Accordingly, CSs from a variety of organisms have been studied in detail, and reviews are available in the literature [35,44,122,125]. The CS from C. glutamicum ssp. flavum has been partially purified, and biochemical analysis revealed that the enzyme shows features typical for that of Gram-positive bacteria, i.e., a homodimeric organization, a molecular weight of about 92,000, weak sensitivity to ATP (Ki = 5 mM) and cisaconitate (Ki = 5 mM), and insensitivity toward NADH and 2-oxoglutarate [101]. Characterization of the C. glutamicum wild-type CS [24] revealed somewhat higher Ki values for both inhibitors (10 mM for ATP, 15 mM for cis-aconitate) and a specific activity varying at best twofold in dependence on the carbon source in the growth medium. These results suggest that in C. glutamicum neither the formation nor the activity of CS is subject to severe regulation. The C. glutamicum CS gene gltA has been studied at the molecular level [24]. The predicted gene product shows up to 50% identity to well-known CS polypeptides from other organisms. The gltA gene is monocistronic and flanked by a putative phosphoserine aminotransferase gene (upstream, opposite orientation) and by a peptidyl-prolyl cis-trans isomerase gene (downstream, same orientation). Inactivation of gltA in C. glutamicum led to citrate or glutamate auxotrophy and to the absence of detectable CS activity [24]. On the basis of these findings it was concluded that only one CS gene is present in C. glutamicum. Recent genome-wide and comparative transcriptome analyses of C. glutamicum cells grown on glucose and/or on acetate indicated an about twofold up-regulation of gltA expression when the cells grow on acetate [27,34,64], which is in agreement with the enzyme activity determination. These results are furthermore in agreement with results obtained by comparative carbon flux determinations with cells grown on glucose and/or acetate

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[123]. Here the authors showed a two- to fourfold higher carbon flux into the TCA cycle when the cells were grown in the presence of acetate. However, it remains to be investigated whether expression regulation of the CS gene is in fact responsible for control of the carbon flow into the cycle. Recently, two gltA homologous genes have been identified in C. glutamicum, designated prpC1 and prpC2 [8]. Due to similarity to the methylcitrate synthase genes from E. coli and Salmonella enterica, and due to functional analysis, it has been determined that the prpC1 and prpC2 genes encode methylcitrate synthases that catalyze the condensation of propionyl-CoA and oxaloacetate to 2-methylcitrate. This reaction is part of the methylcitrate cycle, in which propionate is oxidized to pyruvate [113]. Overexpression of the prpC1 and prpC2 genes in C. glutamicum revealed their products to be active as methylcitrate synthase (0.84 and 5.93 μmol min–1 (mg protein)–1, respectively) and as CS (1.99 and 2.79 μmol min–1 (mg protein)–1) [8]. However, as seen in the glutamate auxotrophic and CS-negative phenotype of the gltA mutant of C. glutamicum, the expression of the genomic copies of prpC1 and prpC2 is obviously too low under the conditions tested to allow substitution of the gltA encoded CS activity. 11.2.1.3 Aconitase Aconitase (cis-aconitate hydratase) catalyzes the isomerization of citrate to isocitrate with cis-aconitate as an enzyme-bound intermediate. Almost no data are available on this enzyme in C. glutamicum, although activity was shown by Shiio et al. in 1959 [98]. The annotated acn gene encodes a polypeptide of 943 amino acid residues. This is appoximately the same size as the aconitase monomers of B. subtilis (909 residues [19]) and E. coli (890 residues[81]), with which it shares more than 54% sequence identity. The three conserved cysteine residues required for formation of the 4Fe-4S center are located at positions 479, 545, and 548 within the C. glutamicum sequence. Recent studies revealed an aconitase activity of 0.25 μmol min–1 (mg protein)–1 in glucose-grown C. glutamicum cells, and plasmid-encoded expression confirmed the functional identity of the annotated acn gene (Krug, Wendisch, and Bott, unpublished data). 11.2.1.4 Isocitrate Dehydrogenase NADP-dependent isocitrate dehydrogenase (ICD) catalyzes the oxidative decarboxylation of D-isocitrate to give 2-oxoglutarate and CO2. 2-Oxoglutarate can then be further oxidized within the TCA cycle or reductively aminated to glutamate. Thus, ICD supplies the cell with a key intermediate of the energy metabolism as well as with precursors and reducing power for biosynthetic purposes. In aerobic organisms such as C. glutamicum able to grow on acetate or ethanol, the enzyme has the additional role to control the carbon flux at the branchpoint between the TCA cycle and the glyoxylate cycle [49,50]. Due to this key role, the ICDs of many organisms have been extensively studied [7]. Partially purified ICD of C. glutamicum ssp. flavum and homogeneously purified enzyme from C. glutamicum wild-type have been found to be specific for NADP,

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to be weakly inhibited by the TCA cycle intermediates citrate, 2-oxoglutarate, and oxaloacetate, and to be strongly inhibited by the simultaneous addition of glyoxylate and oxaloacetate [23,100]. With respect to these regulatory features, the C. glutamicum enzyme resembles the ICDs of B. subtilis and E. coli [7], thus confirming its key role in flux control. However, whereas in the latter two bacteria the ICD is a dimeric enzyme, consisting of two identical subunits, the native enzyme of C. glutamicum is a monomer of about 80 kDa [23], which is about twice the size of the polypeptide of the dimeric form. Monomeric ICDs have been found so far only in a few Gram-negative bacteria, such as Acinetobacter calcoaceticus, Azotobacter vinelandii, Rhodomicrobium vannielii, Rhizobium melilotii, Vibrio parahaemolyticus, and Vibrio sp. ABE-1 [6]. The C. glutamicum ICD gene (icd) has been isolated and characterized. Its gene product consists of 739 residues exhibiting 59% identity with the monomeric ICD from Vibrio sp. ABE-1 [23]. There is no overall sequence similarity between the C. glutamicum ICD and any known ICD of the dimeric type, although there are three confined regions in the monomeric enzyme matching three motifs of the dimeric form [6]. A direct biochemical comparison of the purified C. glutamicum ICD to the structurally distinct enzyme of E. coli revealed a 10-fold higher catalytic efficiency and a 7-fold higher NADP specificity of the C. glutamicum enzyme, favoring NADP over NAD by a factor of 50,000 [6]. In addition, the substrate specificity of the C. glutamicum enzyme turned out to be similar to the dimeric enzyme but significantly more strict. The specific activity of the C. glutamicum ICD is independent of the growth substrate and the growth phase, indicating that the enzyme is formed constitutively [23]. In contrast, the ICD activity of other organisms may vary when the cells grow on different carbon sources [2,37,65,83]. In Vibrio sp. ABE-1, the variation in specific activity was shown to be due to a transcriptional regulation of the icd gene [37]. In E. coli, the change in specific activity is based on the reversible covalent modification of ICD by phosphorylation and dephosphorylation [14]. In the latter respect, it is worth mentioning that Bendt et al. [1] recently found the C. glutamicum ICD protein to be phosphorylated when the cells were grown on minimal medium containing glucose. However, so far there is no experimental evidence for transcriptional control of icd expression or translational regulation of the C. glutamicum enzyme. 11.2.1.5 2-Oxoglutarate Dehydrogenase Complex As previously mentioned, the 2-oxoglutarate dehydrogenase complex (OGDHC) is composed of the three subunits Eo1, Eo2, and LPD. In early studies on OGDHC activity in C. glutamicum, no or only extremely low activities of this enzyme were found, which led to the hypothesis that the interruption of the TCA cycle at this step leads to an overflow of oxoglutarate and consequently, after reductive amination, to glutamate excretion [46]. However, Shiio and Ujigawa-Takeda [106], as well as other investigators, showed that OGDHC is present in C. glutamicum with activities, kinetic characteristics, and effectors comparable to those of other bacteria [43,93,115]. The main characteristics of the OGDHC are its high specificity and the high affinities for 2-oxoglutarate and NAD, with Km values of 80 μM and 86 μM,

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respectively. Also characteristic is the strong inhibition by cis-aconitate, succinylCoA, NADPH, NADH, pyruvate, and oxaloacetate and the activation by acetyl-CoA. As already described in the section on PDHC, a functional lpd gene probably coding for the LPD subunits of the PDHC and the OGDHC has been characterized [93]. Usuda et al. [115] isolated the C. glutamicum odhA gene encoding the 2-oxoglutarate dehydrogenase (E1o subunit) of the OGDHC. The deduced protein possesses an N-terminal extension of 370 amino acids not found in the corresponding proteins of other bacteria. Due to some similarity of the N-terminal extension to the C-terminal region of E2o subunits of OGDHCs from other organisms, the authors speculated the odhA gene to encode a novel bifunctional protein with both E1o and E2o activity [115]. Thus, the E1o and E2o subunits are possibly represented in C. glutamicum by a new type of gene structure that also implies an unusual quaternary structure of the whole OGDHC. However, Kalinowski et al. [41] recently annotated a gene (sucB) exhibiting some similarity to E2o genes from other organisms, including odhB in Gram-positives or sucB in E. coli. Therefore, further enzymological studies are necessary to confirm and identify the suggested function of the genes mentioned. 11.2.1.6 Succinyl-CoA Synthetase Succinyl-CoA synthetase (SCS, also known as succinate thiokinase) catalyzes the formation of succinate and CoA from succinyl-CoA with simultaneous ATP formation and in general consists of two subunits, α and β. Almost no data are available on this enzyme in C. glutamicum. The genes sucC and sucD have been identified in the C. glutamicum genome, based on sequence similarity of the deduced polypeptides to identified succinyl-CoA synthetase subunits [41]. The genes are located next to each other in the same orientation and due to their close proximity (21 bp intergenic region), they may constitute an operon. 11.2.1.7 Succinate:Menaquinone Oxidoreductase Succinate:menaquinone oxidoreductase (SQO, also known as succinate dehydrogenase) is a tightly membrane-bound enzyme catalyzing the oxidation of succinate to fumarate [33]. The electrons released are transferred directly from the enzyme to a quinone present in the membrane. The only quinone present in C. glutamicum is menaquinone, as is also the case with other aerobic Gram-positive bacteria [12,13,35]. Thus menaquinone is regarded as the electron acceptor of the C. glutamicum SQO reaction [5]. Although succinate oxidase activity has been shown to be present in corynebacteria [92,98] and the DCPIP-dependent succinate dehydrogenase activity has been shown to be effectively inhibited by oxaloacetate (Ki = 0.15 to 0.22 μM [59]), the SQO of C. glutamicum has not been purified and analyzed so far. The three genes putatively encoding the respective subunits of SQO, namely SdhC (membrane anchor), SdhA (flavoprotein), and SdhB (iron-sulfur protein), have been identified in the genome of C. glutamicum [41]. They are likely to form an operon since they are oriented in the same direction and coordinately up-regulated when

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C. glutamicum is grown on acetate [27]. Selected properties of the SQO enzyme have been deduced recently by comparison of the three subunit sequences with those of characterized SQO members [5], and these are also described in Chapter 13. 11.2.1.8 Fumarase Fumarase (also known as fumarate hydratase, Fum) catalyzes the interconversion of fumarate and malate. Fum activity has been demonstrated in cell extracts of C. glutamicum ssp. flavum. A putative Fum gene (fum) has been identified in the C. glutamicum genome [41]. The protein encoded by fum consists of 469 amino acids (49.8 kDa) and is thus similar in size to the respective protein of B. subtilis with 464 residues [58] and of E. coli with 465 residues [130]. 11.2.1.9 Malate Dehydrogenase and Malate:Quinone Oxidoreductase The last step of the TCA cycle consists of the regeneration of oxaloacetate by oxidation of malate. C. glutamicum possesses two types of L-malate dehydrogenase, as is the case in other bacteria: a cytoplasmic, NAD-dependent malate dehydrogenase (MDH) and a highly active membrane-associated malate:quinone oxidoreductase (MQO), which transfers the reducing equivalents to menaquinone [60]. Both enzymes have been purified and their biochemistry and in vivo function have been thoroughly studied [59,60]. Under standard conditions, oxaloacetate formation by MDH is highly unfavorable (ΔG o = +28.6 kJ mol –1 ) whereas that by the menaquinone-dependent MQO is favorable (ΔGo = –18.9 kJ mol–1). Thus, it was proposed that MQO is mainly responsible for the net flux from malate to oxaloacetate within the TCA cycle and the MDH reaction in vivo can be driven only by low oxaloacetate and/or high malate concentrations [60]. In fact, after having identified and isolated the mqo and mdh genes encoding MQO and MDH, respectively, Molenaar et al. [59] showed that a defined MQO-deficient mutant was unable to grow on minimal medium whereas an MDH-negative mutant had no obvious phenotype. These results indicate that MQO is essential for a functional TCA cycle in C. glutamicum and that MDH in the wild-type is not of relevance under the conditions tested. Since growth of the Δmqo mutant but not of the Δmqo-Δmdh double mutant on minimal medium could be partially restored by the addition of nicotinamide [59], MDH is obviously able to take over the function of MQO in the Δmqo mutant. Although the physiological function of MDH in the wild-type is not entirely clear, the backflux from oxaloacetate to fumarate, as detected under conditions of NMRflux quantifications [56,79,128], might be attributed to this enzyme. As deduced from its gene, MQO consists of 500 amino acids, and the polypeptide shows up to 49% identity to MQOs of other organisms [42,60]. The MDH gene mdh encodes a polypeptide of 328 residues showing up to 58% identity to corresponding polypeptides. Genome-wide expression profiling did not reveal any indication for carbon source–dependent regulation of mqo or mdh expression [27], although the specific activities of MQO and MDH were found to be threefold higher when the cells were grown on acetate as compared to glucose [59].

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Handbook of Corynebacterium glutamicum INTO AND THROUGH THE

TCA CYCLE

In 1982, Walker et al. [119] showed complete TCA cycle activity in glucose- and in acetate-grown cells of Microbacterium ammoniaphilum (later renamed C. glutamicum) using 13C-labeling and 13C-NMR analysis of isotopically enriched metabolites [119]. Thus, independent of the proof of the presence of all TCA cycle enzymes in C. glutamicum, the functioning of the cycle was demonstrated by this technique. Since then, metabolic fluxes of the entire central metabolism of C. glutamicum have been extensively studied by integrated approaches combining 13C-labeling and isotopomer analysis with metabolite balancing [54,56,74,126,127] (see also Chapter 12). As could be expected, and already indicated by the regulation of selected TCA cycle enzymes at the enzymatic and genetic level, the cycle flux varies in dependence of the growth phase, osmotic stress, growth substrate, different reducing power demands, and the amino acid production status. These studies are briefly outlined in the following paragraphs. Based on metabolite balancing, the central carbon fluxes of a homoserine- and leucine-auxotrophic lysine-producing strain of C. glutamicum (ATCC 21253) were determined in different stages during batch cultivation [73,117]. The relative carbon fluxes into the TCA cycle (normalized to glucose uptake) at the level of citrate synthase were about constant over the course of cultivation, however, glucose uptake and the absolute flux at the entry of the TCA cycle dropped to about 45% when the growth rate decreased and the lysine production started to increase [73]. In contrast, the relative TCA cycle activity in the same C. glutamicum strain was significantly higher when the cells faced hyperosmotic stress [118]. Since the rate of biomass production and thus the requirements of TCA cycle intermediates decreased with increasing osmotic pressure, this result indicates that the cells probably responded to the higher energy demand for the synthesis of compatible solutes. Dominguez et al. [20] investigated the carbon flux distribution in the central metabolic pathways of C. glutamicum (melassecola) during growth on fructose and compared the data with those obtained by Rollin et al. [88] for growth of the same strain on glucose. NMR analyses indicated that the flux distribution in and adjacent to the TCA cycle is virtually identical. However, this contradicts the finding that growth on fructose is associated with enhanced production of CO2 and a diminished pentose phosphate pathway flux, which consequently was interpreted to be due to higher TCA cycle activity [20]. This discrepancy probably can only be solved by performing the experiments under identical conditions in parallel. The in vivo activities for central metabolic pathways in C. glutamicum were quantitated for growth on glucose, acetate, and both carbon sources together [123]. Interestingly, glucose and acetate were co-utilized under the latter condition and the acetate consumption rate was decreased compared to that for growth on acetate alone, as was the glucose consumption rate compared to that for growth on glucose as the sole carbon source. Nevertheless, the consumption rate of total carbon was similar under the three conditions (Table 11.3). When comparing the TCA cycle fluxes, metabolization of glucose was characterized by a relatively low in vivo activity of the TCA cycle, whereas C. glutamicum grown on acetate exhibited an about fourfold increased TCA cycle flux and cells grown on the substrate mixture

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TABLE 11.3 In vivo Activities (Metabolic Fluxes) of Selected Pathways/Reactions in the Central Metabolism of C. glutamicum during Growth in Minimal Medium Containing Different Carbon Sourcesa μmol min-1 (mg protein)-1) During Growth on Net Flux (μ Glucose

Glucose + Acetate

Acetate

Glucose consumption Acetate consumption Carbon consumption

148 888

72 270 972

540 1080

TCA cycle (CS) TCA cycle (ICD) Glyoxylate cycle PDHC reaction PEP/pyruvate carboxylation Oxaloacetate decarboxylation

111 111 <1 161 30 <1

219 169 50 33 <1 15

413 314 99 <1 <1 72

Pathway/Reaction

a

Data from Wendisch et al. [123].

were characterized by an intermediate activity of the cycle. The higher TCA cycle activity in acetate-grown cells may compensate for the lower amount of energy the cells obtain by metabolization of acetate as compared with glucose. The lower TCA cycle activity in glucose-grown cells might be correlated with a repression of genes encoding TCA cycle enzymes and, in addition, activity control of the CS. As outlined above, this enzyme is inhibited by ATP and cis-aconitate. Moreover, the intracellular acetyl-CoA concentration may also control the in vivo activity of the CS since the substrate constant for acetyl-CoA (51 μM [23]) is higher than the intracellular acetylCoA concentration during growth on glucose but lower than that during growth on acetate (24 and 145 μM, respectively [124]). Marx et al. [55] focused on the response of the central metabolism of C. glutamicum to different reducing power demands by constructing and comparing the metabolic fluxes of two isogenic recombinant strains differing only in the possession of an NADH- or NADPH-dependent glutamate dehydrogenase. The difference is due to overexpression of the genes for a heterologous (NADH-dependent) or a homologous (NADPH-dependent) enzyme in a glutamate dehydrogenase-negative background. Flux distribution analysis revealed a significantly higher flux (about 150%) entering the TCA cycle in the heterologous than in the homologous recombinant strain [55]. This result was interpreted to be due to an increased demand for NADH for biosynthetic purposes based on the NADH-dependent glutamate dehydrogenase reaction. Comparative flux analysis in isogenic C. glutamicum strains either growing or producing glutamate or lysine have revealed that the flux entering the TCA cycle at the level of CS is highest under glutamate-producing conditions [56]. This appears plausible due the increased demand of α-ketoglutarate for glutamate production. In

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contrast, the flux into and through the TCA cycle decreased significantly under lysine-production conditions. In a genealogy of five C. glutamicum strains exhibiting an inceased flux toward lysine from 1% to about 25% [128], it was found that with increasing lysine flux, the flux around the anaplerotic node was drastically changed and a significantly reduced flux into the TCA cycle at the level of CS, from 83% down to 60%, occurred. These and similar studies (see also Chapter 12) show the flexible response of the carbon flux into and through the TCA cycle to fulfill the demand for energy, precursors, and reducing equivalents for biosynthetic purposes in C. glutamicum. Although the regulatory mechanisms behind the flux variations observed are not yet known in detail, it can be speculated that both enzymatic as well as global genetic devices are responsible for flux control.

11.2.3 IMPACT OF TCA CYCLE REACTIONS ACID PRODUCTION

ON

AMINO

Because the TCA cycle provides precursors for the synthesis of the glutamate and aspartate family of amino acids, it can be expected that the flux of the cycle and the control of the fluxes are of major importance for the fermentative production of these amino acids. Accordingly, the activity of some of the TCA cycle enzymes and of the PDHC, which provides acetyl-CoA as fueling substrate, were shown to have a severe impact on the production of selected amino acids by C. glutamicum. The prominent example is the OGDHC which has a strong impact on glutamate production (see Chapter 19). The OGDHC competes with glutamate dehydrogenase (GDH) for the common substrate 2-oxoglutarate (Figure 11.1). It has been shown that GDH activity as well as its synthesis is not significantly regulated [4,103] and that the specific activity of the enzyme of growing cells is more than 20-fold higher than that of the OGDHC [4,93,106]. This difference in the specific activities has been suggested to be at least partially responsible for glutamate production [45]. Interestingly, comparing OGDHC and GDH activities under diverse glutamateproducing conditions, a significant reduction of the OGDHC activity (down to 10%) was observed, whereas the GDH activity remained almost constant [43,108]. The inverse correlation of OGDHC activity to glutamate production was recently confirmed by Shimizu et al. [107]. These authors investigated the effect of changes in ICD, GDH, and OGDHC activities on metabolic flux redistribution at the 2-oxoglutarate branch of C. glutamicum and concluded that attenuation of the OGDHC activity is the factor with the greatest impact on glutamate production. A further TCA cycle enzyme probably affecting amino acid production by C. glutamicum is CS. A mutant with low CS activity has been reported to overproduce L-aspartate under biotin-limiting conditions [102]. The same authors showed that S-(2-aminoethyl)-L-cysteine (AEC)–resistant C. glutamicum strains derived from the mutant with low CS activity produced significantly more lysine with a higher yield than an AEC-resistant strain derived from the original wild-type. However, the CS-leaky and AEC-resistant mutant strains were obtained by random mutagenesis and since they were not further biochemically characterized, the conclusions concerning a direct relation between CS activity and amino acid production certainly deserve further investigation. Based on DNA array analysis of a lysine-producing

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mutant of C. glutamicum, Ohnishi et al. [67] discussed a decreased expression of the CS gene gltA as the potential cause of the increased lysine production. In order to test whether the supply of pyruvate is a rate-limiting step for lysine biosynthesis, Shiio et al. [104] generated mutants with low PDHC activities from a lysine-producing C. glutamicum strain. In fact, the PDHC-leaky mutants showed almost identical glucose consumption rates, slower growth, and higher lysine accumulation when compared to the parent strain. However, also in this case the mutants were obtained by random mutagenesis, requiring confirmatory analyses. In favor of a strong impact of the PDHC activity on the production of pyruvate-derived amino acids is the finding that coenzyme A shortening within the cell and thus reduction of the carbon flux via PDHC let to a dramatic increase in L-valine accumulation with a genetically defined valine-producing C. glutamicum strain [82].

11.3 ANAPLEROTIC REACTIONS IN CELLS GROWING ON CARBOHYDRATES It is obvious that during growth and under amino acid production conditions of C. glutamicum, the TCA cycle has to be replenished (Figure 11.1). Naturally great attention has been paid to these reactions of C. glutamicum. In contrast to most other organisms, C. glutamicum possesses both a PEP carboxylase and a pyruvate carboxylase [22,61,80,99] and both enzymes are present during growth and amino acid production from glucose [18,47,76,78]. In addition to the two C3-carboxylating enzymes, C. glutamicum possesses three C4-decarboxylating enzymes that convert the TCA cycle intermediates oxaloacetate or malate to PEP or pyruvate: PEP carboxykinase, malic enzyme, and oxaloacetate decarboxylase (Figure 11.3) [29,39,40,77]. Carboxylating activity of the latter three enzymes, and thus a participation in anaplerosis during growth on glucose, has been excluded [78]. However, since the decarboxylating activities would affect the net anaplerotic flux in C. glutamicum, they cannot be neglected when discussing the enzymes, reactions, and fluxes of the anaplerotic system.

Phosphoenolpyruvate CO 2

CO 2

PEPCk

PQO

PK

ME: malic enzyme

Pyruvate

PDHC

CO 2

PEPCx

Acetate

PCx: pyruvate carboxylase

CO 2

PCx

ODx: oxaloacetate decarboxylase PDHC: pyruvate dehydrogenase complex

ODx

Acetyl-CoA

PEPCk: PEP carboxykinase PEPCx: PEP carboxylase PK: pyruvate kinase

Oxaloacetate ME

Citrate

PQO: pyruvate: quinone oxidoreductase

Malate

FIGURE 11.3 The PEP-pyruvate-oxaloacetate node in detail.

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11.3.1 THE ENZYMES AND GENES AT THE PEP-PYRUVATEOXALOACETATE NODE AND THEIR REGULATION The present state of knowledge about activity and regulation of the enzymes at the PEP-pyruvate-oxaloacetate node of C. glutamicum and about expression and regulation of the respective genes are discussed in this section. The relevant characteristics and the in vitro and in vivo activities are also summarized in Tables 11.1 and 11.2. 11.3.1.1 PEP Carboxylase PEP carboxylase (PEPCx) catalyzes bicarbonate fixation on PEP to form oxaloacetate and inorganic phosphate. Since 1960, the enzyme has been known to be present with high specific activities of up to 0.6 μmol–1 (mg protein)–1 in cell-free extracts of C. glutamicum [22,31,38,61,71,105]. For decades, the anaplerotic function was attributed exclusively to PEPCx [45,53,117]. The enzyme from C. glutamicum ssp. flavum was purified and shown to be activated by acetyl-CoA and fructose-1,6-bisphosphate, and inhibited by aspartate and 2-oxoglutarate (Ki of 44 μM and 4.7 mM, respectively [61,62]). The same regulatory pattern was observed for the enzyme from C. glutamicum wild-type [22]. These regulatory properties, as well as carbon flux studies, suggested a key role of PEPCx in the carbon flow to amino acids derived from the TCA cycle and therefore the enzyme has been proposed to be a very important target in breeding C. glutamicum amino acid–producing strains [22,117]. The C. glutamicum PEPCx gene (ppc) was studied in detail [22,69]. It is organized in a glycolytic gene cluster [21] and transcribed in an operon together with the genes encoding 3-phosphoglycerate kinase and triosephosphate isomerase [94] (see also Chapter 5). Surprisingly, the analysis of defined PEPCx-negative mutants of C. glutamicum revealed that the enzyme is dispensable for growth and lysine production [31,77]. These studies indicated that at least one additional anaplerotic enzyme operates in C. glutamicum. Genetic experiments as well as in vivo 13C-labeling with subsequent 1H-NMR analyses identified the alternative anaplerotic reaction in C. glutamicum to be a PEP or pyruvate carboxylation [72,79], which subsequently led to the identification of pyruvate carboxylase and its gene [78,80]. 11.3.1.2 Pyruvate Carboxylase Pyruvate carboxylase (PCx) catalyzes the irreversible carboxylation of pyruvate to oxaloacetate with concomitant ATP hydrolysis in a two-step reaction [95]. Whereas this enzyme plays a major anaplerotic role in vertebrate tissues and in yeast, only a few prokaryotic organisms use PCx as the sole anaplerotic enzyme. Although there were early indications for the presence of this activity in coryneform bacteria [114], the enzyme could be detected reliably only in an in situ enzyme assay using permeabilized cells [80]. Characterization of PCx activity in such cells revealed that the enzyme is effectively inhibited by AMP, ADP, acetyl-CoA, and aspartate (Ki values of about 0.75 mM, 2.6 mM, 110 μM, and 10 mM, respectively [47,80]). Further characterization indicated that PCx is a biotin-containing enzyme of about

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125 kDa and that its synthesis in C. glutamicum is about threefold up-regulated when using lactate as the substrate [47,78]. The PCx gene product of pyc consists of 1,140 amino acids with ATP and pyruvate binding sites and a typical biotin-carrier domain [48,78]. Inactivation of the chromosomal pyc and/or ppc and growth studies of the mutants indicate that (i) PCx is essential for growth on lactate and pyruvate, (ii) no further anaplerotic enzymes for growth on carbohydrates exist apart from PCx and PEPCx, and (iii) the two enzymes can replace each other as anaplerotic enzyme for growth on glucose [78]. Studies with 13C-NMR revealed that both PCx and PEPCx simultaneously are active in glucose-grown cells of C. glutamicum with the PCx enzyme contributing by far the largest part, 90%, to the total oxaloacetate synthesis by carboxylation [72,74]. 11.3.1.3 PEP Carboxykinase PEP carboxykinase (PEPCk) catalyzes the reversible decarboxylation and simultaneous ATP- or GTP-dependent phosphorylation of oxaloacetate [116]. C. glutamicum has been shown to possess GTP-dependent PEPCk activity [39,77], which is an unusual feature since microbial enzymes in general use ATP as the phosphate donor. The C. glutamicum PEPCk has been purified and kinetic analysis revealed that 0.1 mM ATP inhibits the enzyme in the oxaloacetate-forming reaction 60% [39]. The latter finding already suggested that the enzyme under physiological conditions mainly functions in gluconeogenesis and not in anaplerosis. The PEPCk gene pck from C. glutamicum was isolated, characterized, and used for construction and analysis of mutants and overexpressing strains [87]. The deduced polypeptide consists of 610 amino acids showing no similarity to the ATPdependent PEPCks. The pck gene is monocistronic and its expression is regulated by the carbon source in the growth medium, resulting in an about two- to threefold higher PEPCk specific activity in acetate- or lactate-grown cells than in glucosegrown cells [87]. Consistent with this, acetate-dependent up-regulation of pck was also shown by DNA microarray and quantitative RT-PCR techniques [27]. A growth phase–dependent regulation, as known for other bacterial pck genes [28], has not been observed in C. glutamicum [87]. The gluconeogenic function of PEPCk in C. glutamicum was shown by inactivation of the chromosomal pck gene [87]. The defined mutant was not able to grow on substrates that require gluconeogenesis whereas growth on glucose was unaffected. The inability of the mutant to grow on acetate or lactate further indicates that PEPCk is the only enzyme responsible for PEP synthesis from TCA cycle intermediates. 11.3.1.4 Malic Enzyme Malic enzyme (ME) catalyzes the reversible decarboxylation of malate to pyruvate with simultaneous reduction of NADP [25]. ME activity has been detected in C. glutamicum under various growth conditions [10,11,117], and it has been suggested that the enzyme may play an important role in NADPH generation on

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substrates other than glucose [20]. Together with PCx and the NADH-dependent MDH, ME would catalyze a metabolic cycle generating NADPH from NADH without loss of carbon. Recently, the enzyme has been purified from C. glutamicum and its biochemical properties established [29]. The enzyme is strictly dependent on NADP, activated by NH4+, and slightly inhibited by oxaloacetate (33% inhibition at 10 mM) and glutamate (25% inhibition at 200 mM). Vmax for the decarboxylation activity is about fivefold higher than for the carboxylating activity. This result and the relatively low affinity for pyruvate (Km = 13.4 mM) led to the conclusion that the physiological function of ME is the decarboxylation of malate linked to NADPH generation rather than the decarboxylation of pyruvate [29]. The ME gene malE has been cloned and characterized and strains with different ME activities have been analyzed [29]. The gene product consists of 392 amino acids exhibiting high identities to MEs of other bacteria. Inactivation of the chromosomal malE gene in C. glutamicum had no effect on the growth on either glucose or acetate, however, led to diminished growth rates on lactate [29]. These results substantiate the hypothesis that ME might be involved in the generation of NADPH on substrates known to have a low flux through the pentose pathway. 11.3.1.5 Oxaloacetate Decarboxylase Oxaloacetate decarboxylase (ODx) catalyzes the irreversible decarboxylation of oxaloacetate [51]. Mori and Shiio [63] reported the absence of ODx in C. glutamicum ssp. flavum. However, Jetten and Sinskey [40] were able to purify and biochemically characterize the enzyme from C. glutamicum. The molecular mass of the native ODx was estimated to be about 118 kDa and SDS-PAGE revealed one subunit with a molecular mass of about 32 kDa, suggesting an α4 subunit structure for the native enzyme. It is highly specific for oxaloacetate and inhibited by ADP, coenzyme A and succinate (Ki = 1.2 mM, 2.4 mM, and 2.8 mM, respectively). The purified enzyme did not require sodium ions for activity and is not inhibited by avidin showing that it does not belong to the membrane-bound sodium-dependent ODxs. Instead, it was dependent on divalent cations and accordingly falls into the class of divalent-cation– dependent ODxs. Although the enzyme has been thoroughly characterized, a gene encoding ODx has not yet been identified. Since metabolic network analysis did not identify direct carbon fluxes from oxaloacetate to pyruvate, the identity and function of ODx for growth and amino acid production remains unclear.

11.3.2 PARALLEL AND BIDIRECTIONAL CARBON FLUXES AT THE PEP-PYRUVATE-OXALOACETATE NODE As outlined above and shown in Figure 11.3, C. glutamicum possesses a surprising diversity of enzymes at the PEP-pyruvate-oxaloacetate node and all of them are present with significant specific activities in extracts of cells grown on glucose (Table 11.2). This provokes questions regarding the actual in vivo activities of all the enzymes and the actual fluxes at this metabolic branchpoint. These questions

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TABLE 11.4 C3 Carboxylation and C4 Decarboxylation Fluxes and Anaplerotic Net Fluxes under Different Conditions as Determined by 13C-NMR Analysesa C. glutamicum Strain

Cultivation Condition

C3 Carboxylation%

C4 Decarboxylation%

Anaplerotic Net Flux%

Reference

Wild-type (ATCC13032)

Grown in batch culture Grown in chemostat; lysineproducing Grown in chemostat Grown in chemostat; glutamateproducing

89

69

20

123

69

31

38

54

96

72

24

56

47

18

29

56

MH20-22B (lysine producer) LE 4b LE4

a b

Data given as percentage of the molar glucose uptake rate; data from de Graaf et al. [17]. MH20-22B with wild-type aspartokinase.

have been answered by sophisticated metabolic flux analysis techniques [17,89] (see also Chapter 12). During growth and amino acid production of C. glutamicum on glucose, the two major routes of carbon flux at the PEP-pyruvate-oxaloacetate node are the anaplerotic C3-carboxylation and the oxidative decarboxylation of pyruvate to CO2 and acetylCoA fueling of the TCA cycle. The flux quantifications in a number of strains [56,110,128] revealed that an increased carbon flux into the lysine biosynthetic pathway is always accompanied by both an increase of the anaplerotic flux and a decrease in the flux toward the TCA cycle via acetyl-CoA. Also under glutamate production conditions, the anaplerotic flux is increased. As furthermore revealed from in vivo flux quantifications, in addition to the C3carboxylation activity, there is at the same time a strong C4-decarboxylating activity present in vivo (Table 11.4 [54,55,56,110,123]). Thus, the net anaplerotic flux in fact is the sum of the bidirectional interconversions of C3 and C4 metabolites by simultaneous carboxylation and decarboxylation reactions. It is now clear that the same is true for the anaplerotic fluxes within Bacillus subtilis [90] and E. coli [91]. Petersen et al. [74] recently succeeded to resolve and quantify the in vivo fluxes at the PEP-pyruvate-oxaloacetate node of C. glutamicum. The flux distribution at this node s is shown in Figure 11.4. The authors found that, although the in vitro specific activity of PCx is much lower than that of PEPCx (Table 11.2), the PCx reaction is the main anaplerotic route with a contribution of more than 90% to C3-carboxylation

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1.79

Phosphoenolpyruvate CO 2

CO 2

0.11

0.99 PEPCk

PEPCx

PK 2.61

Pyruvate CO 2 CO 2

1.17 PCx

<0.1 ODx

Acetyl-CoA

ODx

Oxaloacetate 0.98 MQO

1.08 <0.1 ODx

Citrate

ME

Malate

FIGURE 11.4 Anaplerotic fluxes in C. glutamicum during growth on glucose [74].

activity. PEPCx operates in parallel with a small contribution of the remaining 10%. Furthermore, the in vivo C4-decarboxylating flux at the PEP-pyruvate-oxaloacetate node is exclusively due to PEPCk activity and, although present in cell extracts with significant activities, neither ME nor ODx are involved. Thus it became clear that PCx, PEPCk and PK are responsible for an energy (ATP/GTP) consuming cycle in which pyruvate is carboxylated to oxaloacetate, oxaloacetate is decarboxylated to PEP, and PEP is converted to pyruvate again. The basical and essential advantage of the simultaneous carboxylation and decarboxylation reactions might allow a finetuning of the fluxes at the PEP-pyruvate-oxaloacetate node in order to balance anaplerosis with the catabolic reactions, and to rapidly respond to altering environmental conditions [74].

11.3.3 IMPACT OF ANAPLEROTIC REACTIONS ACID PRODUCTION

ON

AMINO

The importance of TCA cycle precursers for amino acid production by C. glutamicum was experimentally addressed in the 1980s with simple feeding experiments [57]. Addition of fumarate to the growth medium was found to lead to about 30% higher lysine formation. It was concluded that an increased oxaloacetate and aspartate availability was rate-limiting for lysine production. Since PEPCx was for a long time considered the only anaplerotic enzyme of C. glutamicum, this enzyme naturally came into focus as a prime candidate for the molecular breeding of strains having increased precurser supply [117]. However, overexpression and inactivation of the PEPCx gene resulted in only marginal effects on lysine and glutamate production (Table 11.5) [15,18,31,77], contrary to the suggested primary role of the enzyme. The relevance of PCx for amino acid production of C. glutamicum has recently been addressed using recombinant strains with different PCx activities [76]. Increasing the PCx activity resulted in higher accumulations of glutamate, lysine, or threonine with the respective strains (Table 11.5), and conversely, abolition of PCx

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263

TABLE 11.5 Enzyme Activities and Amino Acid Production by Isogenic Recombinant Strains of C. glutamicum and of the Lysine Producers DG52-5 or MH2022B. Data Were Taken from Ref. 15, 76, and 87

C. glutamicum Strain WT WTΔppc WT(pMF1014-ppc) MH20-22B MH20-22B-PP (ppc-) DG52-5 DG52-5 (pMF1014-ppc) WT WTΔpyc WT(pVWEx-pyc) DG52-5 DG52-5Δpyc DG52-5(pVWEx-pyc) WT WTΔpck WT(pEK-pckB) MH20-22B MH20-22BΔpck MH20-22B(pEK-pckB)

Relevant Enzyme a

Spec. Activity of Relevant Enzyme μmol min-1 (μ (mg protein)-1)b

Glutamate in Supernatant (mM)

Lysine in Supernatant (mM)d

PEPCx PEPCx PEPCx PEPCx PEPCx PEPCx PEPCx PCx PCx PCx PCx PCx PCx PEPCk PEPCk PEPCk PEPCk PEPCk PEPCk

150 <5 1900 145 <1 160 1200 20 <1 202 8 <1 88 44 <3 413 75 <3 680

9 8 9 n.d.e n.d.e n.d.e n.d.e 8 3 54 n.d.e n.d.e n.d.e 10 44 4 n.d.e n.d.e n.d.e

n.d.e n.d.e n.d.e 42 40 40 49 n.d.e n.d.e n.d.e 34 14 50 n.d.e n.d.e n.d.e 54 66 43

a

For enzyme abbreviations see Figure 11.1. Determined in extracts of cells grown on minimal medium containing glucose c Glutamate production was induced by the addition of Tween 60 and determined after 24 h of cultivation d Lysine was determined after 48 h of cultivation e n.d., not determined b

activity led to a significant decrease in production of the amino acids. These results unequivocally identify the PCx reaction as the main bottleneck for the production of amino acids derived from the TCA cycle. Recently, this conclusion has been further substantiated by use of a mutated PCx introduced into an lysine-producing strain and resulting in increased lysine accumulation [68]. Unfortunately, the authors did not specify the biochemical effect of the pyc mutation and thus it remains to be studied whether the effect is due to improved catalytic activity or due to a deregulation of the PCx enzyme used. The data obtained by metabolic flux analysis of C. glutamicum (see Section 11.3.2 and Chapter 12) suggested that elimination of the reverse anaplerotic flux,

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i.e., of the C4-decarboxylating PEPCk, may also result in increased production of amino acids. This assumption has recently been verified by characterization of defined recombinant strains with altered PEPCk activity [87]. Abolition of PEPCk activity led to an increase of glutamate and lysine production, whereas increasing the PEPCk activity led to reduced accumulation (Table 11.5). The positive effect of decreased PEPCk activity on the production of glutamate and lysine can be explained by an increase in the net carbon flux toward oxaloacetate and an increase in the intracellular oxaloacetate concentration, and thus by an increase in the precursor supply. This hypothesis has been confirmed by comparative carbon flux analyses and intracellular metabolite quantifications in the wild-type and in derivatives with modified PEPCk activities [75].

11.4 ANAPLEROTIC REACTIONS IN CELLS GROWING ON SUBSTRATES OTHER THAN CARBOHYDRATES The growth of C. glutamicum on acetate, ethanol, or fatty acids as sole carbon and energy source requires the operation of the glyoxylate cycle as anaplerotic pathway (Figure 11.1). It has been generally supposed that the key enzymes of the glyoxylate cycle, isocitrate lyase and malate synthase, are repressed via catabolite repression and the glyoxylate bypass is turned off when glucose or other easily metabolizable carbon sources are available [9,14]. However, the regulation of the synthesis of these enzymes in C. glutamicum is clearly different from that known of other organisms [27].

11.4.1 THE ENZYMES AND GENES AND THEIR REGULATION

OF THE

GLYOXYLATE CYCLE

The key enzymes of the glyoxylate cycle are isocitrate lyase and malate synthase. Both enzyme activities have been shown already in 1968 to be present in C. glutamicum [70], and it is also known that C. glutamicum is able to produce amino acids from acetate and from ethanol [45]. Relevant characteristics of enzymes and genes of the glyoxylate cycle are summarized in Tables 11.1 and 11.2, and discussed in the following sections. 11.4.1.1 Isocitrate Lyase and Malate Synthase Isocitrate lyase (ICL) catalyzes the cleavage of isocitrate to succinate and glyoxylate. The enzyme from C. glutamicum has been purified and found to have a moderate affinity for its substrate isocitrate (Km = 0.28 mM) and to be allosterically controlled by a number of intermediates of the central metabolism [84]. 3-Phosphoglycerate, 6-phosphogluconate, PEP, fructose-1,6-bisphosphate, succinate, and glyoxylate were shown to be effective ICL inhibitors with inhibition constants between 0.5 and 1.5 mM. The manifold regulation of ICL on the protein level indicates that this enzyme excerts a tight control on the carbon flux. Malate synthase (MS) catalyzes the irreversible aldol condensation of glyoxylate and acetyl-CoA to form malate and CoA. Malate synthase from C. glutamicum has

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265

also been purified and biochemically characterized [85]. The enzyme shows high affinities for its substrates (30 μM for glyoxylate; 12 μM for acetyl-CoA), but it is not inhibited by any of a variety of intermediates of glycolysis or the TCA cycle tested. Therefore, it has been speculated that MS is only of minor relevance in controlling flux through the glyoxylate cycle [85]. Surprisingly, biochemical analysis of the MS as well as characterization of the corresponding aceB gene revealed that the structure of the enzyme from C. glutamicum is completely different from that of MS enzymes occurring in other organisms as glyoxylate cycle enzymes [85]. Interestingly, the C. glutamicum MS corresponds to the malate synthase protein MS-G of E. coli that is specifically involved in glycolate and glyoxylate metabolism and is induced by glycolate rather than acetate in the growth medium. The ICL and MS genes (aceA and aceB, respectively) from C. glutamicum have been isolated and their structure, genomic organization, expression, and transcriptional regulation have been characterized [27,84,85,124]. The two genes are separated by 597 bp and oriented in opposite directions. They code for polypeptides of 432 amino acids (ICL) and 739 amino acids (MS). Gene inactivations revealed that ICL and MS are essential for growth of C. glutamicum on acetate. Further studies showed that the expression of both genes is coordinately and specifically up-regulated in the presence of acetate (or palmitate) in the growth medium, resulting in much higher specific activities of ICL and MS under these conditions [27,124]. This finding was corroborated by comparative genome-wide expression analyses of C. glutamicum [27,34,64]. These studies additionally revealed the aceA and aceB genes to be members of an acetate stimulon that comprises at least 60 genes. Furthermore, Gerstmeir et al. [26] recently succeeded in the identification of a novel transcriptional regulator, RamB, that negatively controls the expression of aceA and aceB in the presence of glucose in the growth medium.

11.4.2 CARBON FLUXES THROUGH AND FLUX CONTROL

THE

GLYOXYLATE CYCLE

Several studies on the carbon flux within the central metabolism of C. glutamicum revealed that the glyoxylate cycle is completely inactive and that anaplerosis is accomplished solely by C3-carboxylation when the organism grows on carbohydrates [54–56,110,117,126]. These results are in agreement with the general view that the glyoxylate cycle is functional only when acetate or substrates entering the metabolism downstream from pyruvate are the only carbon and energy sources. Wendisch et al. [123] quantified the in vivo activities of the central metabolic pathways of C. glutamicum during growth on acetate, on glucose, and on both carbon sources together (Table 11.3; see also Section 11.2.2). Although the specific activities of the C3-carboxylating enzymes PEPCx and PCx during growth on acetate were comparable to those during growth on glucose, the anaplerotic function on acetate was exclusively fulfilled by the glyoxylate cycle. About 24% of the isocitrate generated in the TCA cycle was channeled into the glyoxylate cycle, which is comparable to the situation in E. coli [120]. Surprisingly, the glyoxylate cycle was also active during growth of C. glutamicum on the mixture of acetate and glucose, and

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anaplerosis occurred exclusively via the glyoxylate cycle under these conditions. The glyoxylate cycle not only replenished the TCA cycle for the withdrawal of precursor metabolites for oxaloacetate- or 2-oxoglutarate–derived amino acids but additionally provided oxaloacetate for gluconeogenesis. This is in sharp contrast to the situation found in E. coli [121]. However, it is in accordance with the finding that in C. glutamicum all three enzymes (ICL, MS, and PEPCk) are present with high specific activity under these conditions [27,124]. The differences between the two organisms are most probably due to different transcriptional control of the respective genes in E. coli [9,14] and C. glutamicum [27]. In addition to the control by regulation of aceA and aceB expression and, thus, by the ICL and MS enzyme content, the carbon flux over the glyoxylate pathway is probably tightly controlled by inhibition of ICL and ICD, both competing for their common substrate isocitrate (Figure 11.1). As already described, both enzyme activities are regulated, and from the inhibition pattern it can be expected that ICL activity decreases when glycolysis is active and ICD activity decreases when the glyoxylate cycle is operative. Additionally, the carbon flux at the isocitrate branchpoint is probably governed by the intracellular concentration of isocitrate, i.e., by the ICL and ICD substrate affinities, which differ by a factor of more than 20 (Km values of 280 μM and 12 μM, respectively [84]).

11.5 GLOBAL CONTROL OF THE TCA CYCLE AND ANAPLEROSIS Although it is obvious that the carbon flux of the TCA cycle and anaplerosis is regulated in C. glutamicum, information on the global control, i.e., on appropriate and coordinated regulation of the enzymes and genes, is only recently emerging. A coordinated regulation of the TCA cycle enzymes and of the anaplerotic enzymes should be expected in order to ensure that energy and precursor generation matches the needs for growth under given conditions. In fact, recent genome-wide expression and proteome analyses revealed coordinated and specific expression of several of the TCA cycle and glyoxylate cycle genes that was dependent on the growth substrate [27,34,64]. The genes encoding the TCA cycle enzymes CS, ACN, SQO, and Fum; the glyoxylate cycle genes for ICL and MS; the genes for the acetate-activating enzymes AK and PTA; and the genes for the E1p subunit of the PDHC, as well as for ME and PEPCk are under transcriptional control in response to the presence or absence of acetate in the growth medium (Figure 11.5). In accordance with induction or repression of these genes by common regulatory devices, Gerstmeir et al. [26] identified a highly conserved 13-bp motif (AAAACTTTGCAAA) in the upstream region of selected genes mentioned above. This motif is a cis-regulatory element for the expression of the ICL and MS genes. Whereas in E. coli the regulatory systems mainly involved in expression of TCA and glyoxylate cycle enzymes are the Crp-dependent catabolite repression and the ArcAB two-component system [16,66,111], in B. subtilis genes for most of the TCA cycle enzymes are repressed either directly or indirectly by the CcpA, CcpC, AbrB, or CodY proteins [3,109]. However, no such system has been found so far in C. glutamicum.

Central Metabolism: Tricarboxylic Acid Cycle and Anaplerotic Reactions

Glucose

Acetate

Glucose-6-P Glycolysis

Acetate

Gluconeo genesis

+3.1 ack

Phosphoenolpyruvate -2.0 pyk

CO2

CO2 +3.8

pck

267

Pyruvate CO2

Acetyl-P

-3.1 aceE +6.2 pta

Acetyl-CoA +2.0 gltA

Aspartate

Oxaloacetate

Citrate +3.7

Malate fum

aceB +8.4

acn

Acetyl-CoA Isocitrate

+1.9

Glyoxylate

aceA +28.6

Fumarate +2.1 sdhCAB

2-Oxoglutarate

Succinate Succinyl-CoA

Glutamate

FIGURE 11.5 Expression changes of selected genes coding for enzymes of the central metabolism in acetate-grown cells of C. glutamicum compared with glucose-grown cells [27]. Positive numbers indicate increased expression on acetate; negative numbers indicate decreased expression. All genes indicated except those for acetate kinase (ack) and phosphotransacetylase (pta) are listed in Table 11.1.

11.6 CONCLUDING REMARKS AND PERSPECTIVES Substantial information has been accumulated on the biochemistry, physiology, and molecular biology of the anaplerotic reactions, the TCA cycle, and the glyoxylate cycle. The regulation of individual enzyme activities and control of gene expression is partly elucidated, and the relevance of the respective enzyme activity for growth and amino acid production is clarified. Due to the quantitative assessment of metabolic fluxes in central metabolism, detailed information is available not only for single reactions but also for pathways as a whole.

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Nevertheless, there is still a significant lack of knowledge regarding the features and relevance of enzymes such as PDHC, ACN, and SQO for overall control of the carbon flow. Moreover, the molecular mechanisms of transcriptional and post-transcriptional regulation, and of the signals modulating the function of regulatory proteins involved, remain to be determined. The recent studies on the phosphoproteome of C. glutamicum have indicated that several of the TCA cycle enzymes such as CS, ACN, ICD, LPD, and Fum, as well as PCx and PK, are present in a phosphorylated form [1], and it is a challenge remaining for future studies to reveal the relevance of phosphorylation/dephosphorylation. Since there are strong indications for an important role of TCA cycle enzymes and PDHC for amino acid production, further investigations are expected to be profitable both from the scientific point of view and for amino acid production with C. glutamicum.

ACKNOWLEDGMENTS I thank M. Eikmanns, and M. Bott for suggestions and valuable help in preparing this manuscript. The work in my lab is supported by the Ministry of Education and Science (BMBF); the Ministry of Consumer Protection, Nutrition and Agriculture (BMVEL); the EU; and Degussa.

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57. Menkel E, Thierbach G, Eggeling L, and Sahm H. (1989) Influence of increased aspartate availability on lysine formation by a recombinant strain of Corynebacterium glutamicum and utilization of fumarate. Appl. Environm. Microbiol. 55:684–688. 58. Miles JS and Guest JR. (1987) Molecular genetic aspects of the citric acid cycle of Escherichia coli. Biochem. Soc. Symp. 54:45–65. 59. Molenaar D, van der Rest ME, Drysch A, and Yucel R. (2000) Functions of the membrane-associated and cytoplasmic malate dehydrogenases in the citric acid cycle of Corynebacterium glutamicum. J. Bacteriol. 182:6884–6891. 60. Molenaar D, van der Rest ME, and Petrovic S. (1998) Biochemical and genetic characterization of the membrane associated malate dehydrogenase (acceptor) from Corynebacterium glutamicum. Eur. J. Biochem. 254:395–403. 61. Mori M and Shiio I. (1985) Purification and some properties of phosphoenolpyruvate carboxylase from Brevibacterium flavum and its aspartate-overproducing mutant. J. Biochem. 97:1119–1128. 62. Mori M and Shiio I. (1985) Synergistic inhibition of phosphoenolpyruvate carboxylase by aspartate and 2-oxoglutarate in Brevibacterium flavum. J. Biochem. 98:1621–1630. 63. Mori M and Shiio I. (1987) Pyruvate formation and sugar metabolism in an amino acid-producing bacterium, Brevibacterium flavum. Agric. Biol. Chem. 51:129–138. 64. Muffler A, Bettermann S, Haushalter M, Hörlein A, Neveling U, Schramm M, and Sorgenfrei O. (2002) Genome-wide transcription profiling of Corynebacterium glutamicum after heat shock and during growth on acetate and glucose. J. Biotechnol. 98:255–268. 65. Nimmo HG. (1987) The tricarboxylic acid cycle and anaplerotic reactions. In Neidhardt FC, Curtiss R, Ingraham JL, Lin ECC, Low KB, Magasanik B, Reznikoff WS, Riley M, Schaechte M, and Umbarger HE (Eds.), Escherichia coli and Salmonella typhimurium: cellular and molecular biology, Vol. 1, American Society for Microbiology, Washington, D.C., pp. 156–169. 66. Oh MK, Rohlin L, Kao KC, and Liao JC. (2002). Global expression profiling of acetate-grown Escherichia coli. J. Biol. Chem. 277:13175–13183. 67. Ohnishi J, Hayashi M, Mitsuhashi S, and Ikeda M. (2003) Efficient 40˚C fermentation of L-lysine by a new Corynebacterium glutamicum mutant developed by genome breeding. Appl. Microbiol. Biotechnol. 62:69–75. 68. Ohnishi J, Mitsuhashi S, Hayashi M, Ando S, Yokoi H, Ochiai K, and Ikeda M. (2002) A novel methodology employing Corynebacterium glutamicum genome information to generate a new L-lysine-producing mutant. Appl. Microbiol. Biotechnol 58:217–223. 69. O’Regan M, Thierbach G, Bachmann B, Villeval D, Lepage P, Viret JF, and Lemoine Y. (1989) Cloning and nucleotide sequence of the phosphoenolpyruvate carboxylase gene of Corynebacterium glutamicum ATCC13032. Gene 77:237–251. 70. Ozaki H and Shiio I. (1968) Regulation of the TCA and glyoxylate cycles in Brevibacterium flavum. J. Biochem. 64:355–363. 71. Ozaki H and Shiio I. (1969) Regulation of the TCA and glyoxylate cycles in Brevibacterium flavum. J. Biochem. 297–311. 72. Park SM, Shaw-Reid C, Sinskey AJ, and Stephanopoulos G. (1997) Elucidation of anaplerotic pathways in Corynebacterium glutamicum via 13C-NMR spectroscopy and GC-MS. Appl. Microbiol. Biotechnol. 47:430–440. 73. Park SM, Sinskey AJ, and Stephanopoulos G. (1997) Metabolic and physiological studies of Corynebacterium glutamicum mutants. Biotechnol. Bioeng. 55:864–879. 74. Petersen S, de Graaf AA, Eggeling L, Möllney M, Wiechert W, and Sahm H. (2000) In vivo quantification of parallel and bidirectional fluxes in the anaplerosis of Corynebacterium glutamicum. J. Biol. Chem. 275:35932–35941.

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75. Petersen S, Mack C, de Graaf AA, Riedel C, Eikmanns BJ, and Sahm H. (2001) Metabolic consequences of altered phosphoenolpyruvate carboxykinase activity in Corynebacterium glutamicum reveal anaplerotic regulation in vivo. Metabolic Engineering 3:344–361. 76. Peters-Wendisch P, Schiel B, Wendisch VF, Katsoulidis E, Möckel B, Sahm H, and Eikmanns BJ. (2001) Pyruvate carboxylase as a major bottleneck for glutamate and lysine production by Corynebacterium glutamicum. J. Mol. Microbiol. Biotechnol. 3:295–300. 77. Peters-Wendisch PG, Eikmanns BJ, Thierbach G, Bachmann B, and Sahm H. (1993) Phosphoenolpyruvate carboxylase in Corynebacterium glutamicum is dispensable for growth and lysine production. FEMS Microbiol. Lett. 112:269–274. 78. Peters-Wendisch PG, Kreutzer C, Kalinowski J, Patek M, Sahm H, and Eikmanns BJ. (1998) Pyruvate carboxylase from Corynebacterium glutamicum: characterization, expression and inactivation of the pyc gene. Microbiology 144:915–927. 79. Peters-Wendisch PG, Wendisch VF, de Graaf AA, Eikmanns BJ, and Sahm H. (1996) C3-Carboxylation as anaplerotic reaction in PEP carboxylase-deficient Corynebacterium glutamicum. Arch. Microbiol. 65:387–396. 80. Peters-Wendisch PG, Wendisch VF, Paul S, Eikmanns BJ, and Sahm H. (1997) Pyruvate carboxylase as anaplerotic enzyme in Corynebacterium glutamicum. Microbiology 143:1095–1103. 81. Prodromou C, Artymiuk PJ, and Guest JR. (1992) The aconitase of Escherichia coli. Nucleotide sequence of the aconitase gene and amino acid sequence similarity with mitochondrial aconitases, the iron-responsive element-binding protein and isopropylmalate isomerases. Eur. J. Biochem. 204:599–609. 82. Radmacher E, Vaitsikova A, Burger U, Krumbach K, Sahm H, and Eggeling L. (2002) Linking central metabolism with increased pathway flux: L-valine accumulation by Corynebacterium glutamicum. Appl. Environm. Microbiol. 68:2246–2250. 83. Reeves HC, O’Neil S, and Weitzman PDJ. (1986) Changes in NADP-isocitrate dehydrogenase isoenzyme levels in Acinetobacter calcoaceticus in response to acetate. FEMS Microbiol. Lett. 35:229–232. 84. Reinscheid DJ, Eikmanns BJ, and Sahm H. (1994). Characterization of the isocitrate lyase gene from Corynebacterium glutamicum and biochemical analysis of the enzyme. J. Bacteriol. 176:3474–3483. 85. Reinscheid DJ, Eikmanns BJ, and Sahm H. (1994) Malate synthase from Corynebacterium glutamicum: sequence analysis of the gene and biochemical characterization of the enzyme. Microbiology 140:3099–3108. 86. Reinscheid DJ, Schnicke S, Rittmann D, Zahnow U, Sahm H, and Eikmanns BJ. (1999) Cloning, sequence analysis, expression and inactivation of the Corynebacterium glutamicum pta-ack operon encoding phosphotransacetylase and acetate kinase. Microbiology 145:503–513. 87. Riedel C, Rittmann D, Dangel P, Möckel B, Sahm H, and Eikmanns BJ. (2001) Characterization, expression, and inactivation of the phosphoenolpyruvate carboxykinase gene from Corynebacterium glutamicum and significance of the enzyme for growth and amino acid production. J. Mol. Microbiol. Biotechnol. 3:573–583. 88. Rollin C, Morgant V, Guyonvarch A, and Guerquin-Kern JL. (1995) 13C-NMR studies of Corynebacterium melassecola metabolic pathways. Eur. J. Biochem. 227:488–493. 89. Sahm H, Eggeling L, and de Graaf AA. (2000) Pathway analysis and metabolic engineering in Corynebacterium glutamicum. Biol. Chem. 381:899–910. 90. Sauer U, Hatzimanikatis V, Bailey JE, Hochuli M, Szyperski T, and Wuthrich K. (1997) Metabolic fluxes in riboflavin-producing Bacillus subtilis. Nat. Biotechnol. 15:448–452.

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91. Sauer U, Lasko DR, Fiaux J, Hochuli M, Glaser R, Szyperski T, Wuthrich K, and Bailey JE. (1999) Metabolic flux ratio analysis of genetic and environmental modulations of Escherichia coli central carbon metabolism. J. Bacteriol. 181:6679–6688. 92. Schirawski J and Unden G. (1998) Menaquinone-dependent succinate dehydrogenase of bacteria catalyzes reversed electron transport driven by the proton potential. Eur. J. Biochem. 257:210–215. 93. Schwinde J, Hertz PF, Sahm H, Eikmanns BJ, and Guyonvarch A. (2001) Lipoamide dehydrogenase from Corynebacterium glutamicum: molecular and physiological analysis of the lpd gene and characterization of the enzyme. Microbiology 147:2223–2231. 94. Schwinde JW, Thum-Schmitz N, Eikmanns BJ, and Sahm H. (1993). Transcriptional analysis of the gap-pgk-tpi-ppc gene cluster of Corynebacterium glutamicum. J. Bacteriol. 175:3905–3908. 95. Scrutton MC and Young MR. (1972) Pyruvate carboxylase. In Boyer PD (Ed.), The Enzymes, Vol. 6, Academic Press, New York, pp. 1–35. 96. Shen LC and Atkinson DE. (1970) Regulation of pyruvate dehydrogenase from Escherichia coli. J. Biol. Chem. 245:5974–5978. 97. Shiio I, Momose H, and Oyama A. (1969) Genetic and biochemical studies on bacterial formation of L-glutamate I. Relationship between isocitrate lyase, acetate kinase, and phosphate acetyltranferase levels and glutamate production in Brevibacterium flavum. J. Gen. Appl. Microbiol. 15:27–40. 98. Shiio I, Otsuka S, and Tsunoda T. (1959) Glutamic acid formation from glucose by bacteria I. Enzymes of the Embden-Meyerhof-Parnas pathway, the Krebs cycle, and the glyoxylate bypass in cell extracts of Brevibacterium flavum No. 2247. J. Biol. Chem. 46:1303–1311. 99. Shiio I, Otsuka SI, and Tsunoda T. (1960) Glutamic acid formation from glucose by bacteria. J. Biochem. 47:414–421. 100. Shiio I and Ozaki H. (1968) Concerted inhibition of isocitrate dehydrogenase by glyoxylate plus oxalacetate. J. Biochem. 64:45–53. 101. Shiio I, Ozaki H, and Ujigawa K. (1977) Regulation of citrate synthase in Brevibacterium flavum, a glutamate-producing bacterium. J. Biochem. 82:395–405. 102. Shiio I, Ozaki H, and Ujigawa-Takeda K. (1982) Production of aspartic acid and lysine by citrate synthase mutants of Brevibacterium flavum. Agric. Biol. Chem. 46:101–107. 103. Shiio I and Ozaki H. (1970) Regulation of nicotinamide adenine dinucleotide phosphate-specific glutamate dehydrogenase from Brevibacterium flavum, a glutamateproducing bacterium. J. Biochem. 68:633–647. 104. Shiio I, Toride Y, and Sugimoto S. (1984) Production of lysine by pyruvate dehydrogenase mutants of Brevibacterium flavum. Agric. Biol. Chem. 48:3091–3098. 105. Shiio I and Ujigawa K. (1978) Enzymes of the glutamate and aspartate synthetic pathways in a glutamate-producing bacterium, Brevibacterium flavum. J. Biochem. 84:647–657. 106. Shiio I and Ujigawa-Takeda K. (1980) Presence and regulation of α-ketoglutarate dehydrogenase complex in a glutamate-producing bacterium, Brevibacterium flavum. Agric. Biol. Chem. 44:1897–1904. 107. Shimizu H, Tanaka H, Nakato A, Nagahisa K, Kimura E, and Shioya S. (2003) Effects of the changes in enzyme activities on metabolic flux redistribution around the 2-oxoglutarate branch in glutamate production by Corynebacterium glutamicum. Bioprocess Biosyst. Eng. 25:291–298.

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108. Shingu H and Terui G. (1971) Studies on the process of glutamic acid fermentation at the enzyme level: on the changes of α-ketoglutaric acid dehydrogenase in the course of culture. J. Ferment. Technol. 49:400–405. 109. Sonenshein AL. (2002) The Krebs citric acid cycle. In Sonenshein AL, Losick RM, and Hoch HA (Eds.), Bacillus subtilis and Its Closest Relatives: from Genes to Cell. ASM Press, Washington, D.C., pp. 151–162. 110. Sonntag K, Schwinde J, de Graaf A, Marx A, Eikmanns BJ, Wiechert W, and Sahm H. (1995) 13C NMR studies of the fluxes in the central metabolism of Corynebacterium glutamicum during growth and overproduction of amino acids in batch cultures. Appl. Microbiol. Biotechnol. 44:489–495. 111. Spencer ME and Guest JR. (1987) Regulation of citric acid cycle genes in facultative bacteria. Microbiol. Sci. 4:164–168. 112. Tauch A, Homann I, Mormann S, Rüberg S, Billault A, Bathe B, Brand S, BrockmannGretza O, Rückert C, Schischka N, Wrenger C, Hoheisel J, Möckel B, Huthmacher K, Pfefferle W, Pühler A, and Kalinowski J. (2002) Strategy to sequence the genome of Corynebacterium glutamicum ATCC 13032: use of a cosmid and a bacterial artificial chromosome library. J. Biotechnol. 95:25–38. 113. Textor S, Wendisch VF, de Graaf AA, Müller U, Linder I, Linder D, and Buckel W. (1997) Propionate oxidation in Escherichia coli: evidence for operation of a methylcitrate cycle in bacteria. Arch. Microbiol. 168:428–436. 114. Tosaka O, Morioka H, and Takinami K. (1979) The role of biotin-dependent pyruvate carboxylase in L-lysine production. Agric. Biol. Chem. 43:1513–1519. 115. Usuda Y, Tujimoto N, Abe C, Asakura Y, Kimura E, Kawahara Y, Kurahashi O, and Matsui H. (1996) Molecular cloning of the Corynebacterium glutamicum (Brevibacterium lactofermentum AJ12036) odhA gene encoding a novel type of 2-oxoglutarate dehydrogenase. Microbiology 142:3347–3354. 116. Utter MF and Kolenbrander HM. (1972) Formation of oxaloacetate by CO2 fixation on phosphoenolpyruvate. In The Enzymes, Vol. VI, Boyer, P.D. (Ed.), Academic Press, New York, pp. 117–170. 117. Vallino JJ and Stephanopoulos G. (1993) Metabolic flux distributions in Corynebacterium glutamicum during growth and lysine overproduction. Biotechnol. Bioeng. 41:633–646. 118. Varela C, Agosin E, Baez M, Klapa M, and Stephanopoulos G. (2003) Metabolic flux redistribution in Corynebacterium glutamicum in response to osmotic stress. Appl. Microbiol. Biotechnol. 60:547–555. 119. Walker TE, Han CH, Kollman VH, London RE, and Matwiyoff NA. (1982) 13C Nuclear magnetic resonance studies of the biosynthesis by Microbacterium ammoniaphilum of L-glutamate selectively enriched with 13C. J. Biol. Chem. 257:1189–1195. 120. Walsh K and Koshland DE Jr. (1984) Determination of flux through the branch point of two metabolic cycles. J. Biol. Chem. 259:9646–9654. 121. Walsh K and Koshland DE Jr. (1985) Branch point control by the phosphorylation state of isocitrate dehydrogenase. J. Biol. Chem. 260:8430–8437. 122. Weitzman PD. (1981) Unity and diversity in some bacterial citric acid cycle enzymes. Adv. Microb. Physiol. 22:185–243. 123. Wendisch VF, de Graaf AA, Sahm H, and Eikmanns BJ. (2000) Quantitative determination of metabolic fluxes during coutilization of two carbon sources: Comparative analyses with Corynebacterium glutamicum during growth on acetate and/or glucose. J. Bacteriol. 182:3088–3096.

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124. Wendisch VF, Spies M, Reinscheid DJ, Schnicke S, Sahm H, and Eikmanns BJ. (1997). Regulation of acetate metabolism in Corynebacterium glutamicum: Transcriptional control of the isocitrate lyase and malate synthase genes. Arch. Microbiol. 168:262–269. 125. Wiegand G and Remington SJ. (1986) Citrate synthase: Structure, control, and mechanisms. Annu. Rev. Biophys. Chem. 15:97–117. 126. Wittmann C and Heinzle E. (2001) Modeling and experimental design for metabolic flux analysis of lysine-producing Corynebacteria by mass spectrometry. Metabol. Eng. 3:171–193. 127. Wittmann C and Heinzle E. (2001) Application of MALDI-TOF MS to lysineproducing Corynebacterium glutamicum: a novel approach for metabolic flux analysis. Eur. J. Biochem. 268:2441–2455. 128. Wittmann C and Heinzle E. (2002) Genealogy profiling through strain improvement by using metabolic network analysis: metabolic flux genealogy of several generations of lysine-producing corynebacteria. Appl. Environ. Microbiol. 68:5843–5859. 129. Wood HD and Utter MF. (1965) The role of CO2 fixation in metabolism. In Campbell PN and Greville GD (Eds.), Essays in Biochemistry, Vol. 2. Academic Press, London and New York, pp. 1–27. 130. Woods SA, Miles JS, Roberts RE, and Guest JR. (1986) Structural and functional relationships between fumarase and aspartase. Biochem. J. 237:547–557.

12

Metabolic Flux Analysis in Corynebacterium glutamicum C. Wittmann and A.A. De Graaf

CONTENTS 12.1 Introduction ..................................................................................................277 12.2 Metabolic Flux Analysis: Concepts and Tools............................................278 12.2.1 Prerequisites: Network Topology and Cellular Composition .........278 12.2.2 Metabolite Balancing .......................................................................279 12.2.3 Isotope Labeling...............................................................................284 12.2.4 Isotopomer Modeling.......................................................................286 12.3 Metabolic Fluxes in C. glutamicum ............................................................286 12.3.1 Fluxes for the Generation of Reducing Power................................287 12.3.2 Anaplerotic Fluxes ...........................................................................289 12.3.3 Fluxes in a Genealogy of Strains ....................................................291 12.3.4 Fluxes on Different Carbon Sources ...............................................293 12.3.5 Response of Fluxes to Different Cellular Demands .......................296 12.3.6 Nitrogen Fluxes................................................................................298 12.4 Concluding Remarks....................................................................................300 References..............................................................................................................300

12.1 INTRODUCTION Initiated by the pioneering discovery of the glutamate-producing Corynebacterium glutamicum [20], extensive biochemical and genetical analysis of the metabolic network of this microorganism has been carried out [11,21,39]. This provided detailed knowledge on its enzymes and pathways and was therefore very useful for the optimization of strains and production processes. In recent years, novel approaches were developed that allow the in vivo quantification of fluxes in the metabolism of C. glutamicum. These approaches revealed fascinating insights into metabolic properties of C. glutamicum otherwise not accessible. In addition, they offered new possibilities as a rational basis for targeted strain improvement. The present chapter gives an overview of metabolic flux quantifications of C. glutamicum,

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whereby, with respect to metabolic fluxes, C. glutamicum is probably the most extensively studied organism so far.

12.2 METABOLIC FLUX ANALYSIS: CONCEPTS AND TOOLS Beyond a doubt, the targeted improvement of cellular processes requires a detailed understanding of the reactions supporting growth and product formation. Of central importance in this regard is quantitative knowledge about the in vivo activity of individual pathways and their contribution to the overall activity of the cell. Such knowledge on metabolic fluxes has proven especially useful for the understanding and optimization of C. glutamicum, characterized by a close link between central metabolism and amino acid biosynthesis. The two major approaches for metabolic flux analysis, metabolite balancing and isotope labeling, are introduced in the present chapter.

12.2.1 PREREQUISITES: NETWORK TOPOLOGY AND CELLULAR COMPOSITION The basic prerequisite for deriving in vivo fluxes is knowledge on the existing cellular reactions, on the network topology, and, furthermore, on the cellular composition. For C. glutamicum, the reactions of the central metabolism involving substrate uptake, glycolysis, pentose phosphate pathway, TCA cycle, anaplerotic carboxylation, glyoxylate cycle, and product synthesis are well established (Figure 12.1). It should be noticed that the knowledge about the metabolic network is continuously updated with new findings, such as the detection of pyruvate carboxylase as anaplerotic enzyme in C. glutamicum [35] or the consideration of new reactions for growth on specific substrates, such as ribose [53] or sucrose [8]. In the central metabolism of C. glutamicum, various anabolic reactions drain carbon via different precursor compounds from the intermediary metabolism. These anabolic reactions must be taken into account in all studies on metabolic fluxes, where growing cells are investigated. Owing to extensive biochemical work in the past, the knowledge on the biochemical pathways leading to the biomass constituents is well established. The precursors for the formation of biomass in C. glutamicum are glucose-6-phosphate, fructose-6-phosphate, ribose-5-phosphate, erythrose-4-phosphate, glyceraldehyde-3-phosphate, 3-phosphoglycerate, pyruvate, phosphoenolpyruvate, acetylCoA, oxoglutarate, and oxaloacetate as shown in Figure 12.1. For C. glutamicum, detailed analysis of cellular composition and resulting demand for anabolic precursors has been carried out in different studies [3,23]. The composition of C. glutamicum is given in Table 12.1. The data shown were obtained by detailed and thorough analysis of cellular composition [29], including a correction for the different cell wall composition on basis of the diaminopimelate content of the cell [5]. Overall about 16,400 μmol of NADPH is required for synthesis of 1 g of biomass. Considering a biomass yield of 0.5 (g biodrymass)/(g glucose), which is achieved by C. glutamicum under aerobic conditions, this results in 1.5 mol NADPH/(mol

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glucose) that have to be generated by the NADPH-forming reactions in the PPP and the TCA cycles. It is obvious that these NADPH requirements for cell material formation compete with the increased need for biosynthetic pathways when the corresponding products are overproduced, such as lysine, isoleucine, valine, threonine, histidine, aromatic amino acids, or panthotenate.

12.2.2 METABOLITE BALANCING Metabolite balancing is the classical and easy accessible approach for metabolic flux analysis because the measurement of a few fluxes, e.g., of substrate uptake, biomass, or product formation, is usually sufficient for intracellular flux calculation. It is based on a stoichiometric model of the metabolism, mass balances around intracellular metabolite pools, and a set of measured fluxes [43]. Assuming steady state, i.e., the sum of fluxes into a metabolite pool equals the sum of fluxes out, flux estimation is a problem of solving a set of linear equations. Intracellular fluxes are hereby calculated from measured extracellular fluxes, e.g., for substrate uptake or product secretion, together with the anabolic fluxes derived from the calculation of precursor requirements for anabolic purposes involving experimental quantification of the biomass yield, and detailed knowledge of cellular composition. The following examples give an overview on metabolite-balancing studies applied to C. glutamicum. A detailed analysis of metabolic fluxes employing metabolite balancing in different stages of a batch cultivation of a lysine-producing C. glutamicum ATCC 21253 was carried out by Vallino and Stephanopoulos [46]. The threonine and methionine auxotrophic strain they used exhibited an initial phase of growth in the presence of the essential amino acids, followed by a phase of lysine production after their depletion from the medium. During the switch from growth to lysine production, C. glutamicum ATCC 21253 showed a number of characteristic changes in intracellular fluxes. The observed increase of the flux into lysine biosynthesis was accompanied by increased fluxes through the PPP, anaplerotic carboxylation, and glutamate dehydrogenase, respectively, whereas the TCA cycle was affected only to a minor extent. Subsequently, studies on metabolic network rigidity in lysine-producing C. glutamicum were carried by the same authors. They revealed flexibility of the nodes of glucose-6-phosphate and pyruvate in response to different metabolic perturbations and indicated that lysine production was not limited by either pyruvate or NADPH availability [45,46]. Recently, metabolite balancing was applied by the same group to study the impact of osmotic stress, a major stress factor in industrial production processes, on lysine-producing C. glutamicum ATCC 21253 [48]. In response to high osmolarity, the strain exhibited increased fluxes toward the formation of compatible solutes, such as trehalose and proline, and through energygenerating pathways of glycolysis and TCA cycle resulting in higher specific ATP production rates. Dominguez et al. [7] investigated the influence of transient oxygen limitation during batch culture of C. melassecola ATCC 17965. Unfortunately, interpretation of the results is hampered by the complex response observed and the great variation of the determined fluxes. In additional studies of C. melassecola ATCC 17965 growing on lactate and on glucose, respectively, metabolite balancing

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FIGURE 12.1 Central metabolic network of C. glutamicum involving sucrose PTS system (1), fructose PTS system (2), mannose PTS system (3), glucose PTS system (4), sucrase (5), glucose-6-phosphate isomerase (6), phosphofructokinase (7), fructose 1,6-bisphosphatase (8), fructose-1,6-bisphosphate aldolase (9), triose phosphate isomerase (10), glucose-6-phosphate

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and enzyme measurements were combined to obtain additional constraints on the network topology [3,4]. Pons et al. investigated metabolic fluxes of C. melassecola ATCC 17965 during growth on fructose, glucose, and mixtures of both substrates [36]. Metabolite balancing was also applied to quantify selected fluxes in the metabolism of pantothenate-overproducing C. glutamicum strain ATCC 13032 ilvA pECM3ilvBNCD pEKEx2panBC [2]. The performed study involved cofactor balancing but could not resolve flux partitioning between PPP and glycolysis. It was found that about 85% of the flux arriving at pyruvate is redirected toward ketoisovalerate, from which only a minor fraction of 5% was converted into the desired product. The majority of carbon was channeled into by-products, whereby valine was the dominating compound. As shown by the given examples, metabolite balancing has proven to be very useful for metabolic flux quantification of C. glutamicum. Admittedly, this approach exhibits different limitations. In order to arrive at a fully determined network, additional constraints are required because the number of measurements and metabolite balances is usually lower than the degrees of freedom of the system. In most cases, additional constraints are obtained from balances over ATP, NADH, or NADPH. For example, Vallino and Stephanopoulos assumed the glyoxylate pathway to be inactive; glucose-6-phosphate dehydrogenase, 6-phosphogluconate dehydrogenase, and isocitrate dehydrogenase to be the sole sources for NADPH; and PEP carboxylase to be the sole anaplerotic enzyme [46]. Such constraints may not be valid: (1) unaccounted NADPH oxidase activity would cause an underestimation of the flux through the PPP and (2) unaccounted activity of the glyoxylate pathway would lead to an overestimated anaplerotic flux. The second drawback of metabolite balancing is the fact that important fine structures of the metabolic network of C. glutamicum cannot be resolved. Among these are parallel reactions with identical stoichiometry around the pyruvate node, such as (1) pyruvate carboxylase and PEP carboxylase for anaplerotic carboxylation and (2) malic enzyme, oxaloacetate carboxylase, and PEP carboxykinase catalyzing the conversion of C4 metabolites of the TCA cycle into glycolytic C3 metabolites. Especially for those amino acids derived from oxaloacetate, like lysine or threonine, it is important to know how these enzymes contribute to the overall supply of the precursor. Also, the relative activity of the two alternative pathways in lysine biosynthesis cannot be resolved by metabolite balancing.

FIGURE 12.1 (continued) dehydrogenase (11), 6-phosphogluconate dehydrogenase (12), phosphoribose isomerase (13), ribulose-5-phosphate epimerase (14), transketolase (15, 16), transaldolase (17), glyceraldehyde-3-phosphate dehydrogenase (18), phosphoglycerate kinase (19), phospoglycerate mutase (20), enolase (21), pyruvate kinase (22), pyruvate dehydrogenase (23), malic enzyme (24), oxaloacetate decarboxylase (25), pyruvate carboxylase (26), phosphoenolpyruvate carboxykinase (27), phosphoenolpyruvate carboxylase (28), citrate synthase (29), aconitase (30), isocitrate dehydrogenase (31), oxoglutarate dehydrogenase (32), succinate thiokinase (33), succinate dehydrogenase (34), fumarase (35), malate dehydrogenase (36), acetate kinase (37), phosphotransacetylase (38), isocitrate lyase (39), malate synthase (40), reactions consuming intermediary metabolites for anabolic purposes (41–51).

Ala Arg Asx Cys Glu Gln Gluintra Glnintra Gly His Ile Leu Lys Met Phe Pro Ser

Precursor 606 189 399 87 360 147 250 49 361 71 202 440 202 146 133 170 225

Demand (μmol/g) G6P

F6P

1

R5P

1

E4P

GAP

1

1

1

PGA

2

PEP

1 2 1

1

PYR

1

AcCoA

1 1

1

1

OAA

TABLE 12.1 Precursor Demand for Biomass Synthesis in C. glutamicum in μmol⋅⋅(g dry cell mass)–1

1

1 1 1 1

1

AKG

–1

–1 –2 –1

1

CO2

1 4 1 5 1 1 1 1 1 1 5 2 4 8 2 3 1

NADPH

282 Handbook of Corynebacterium glutamicum

275 54 81 284 146

205

154

51

0

308

16 292

0

879

24

630 100

125

1

268

268

1 1

129

129

0

1295

49

368 50 129 24

673

652

24 146

482

1 2

2604

2604

2 1

3177

2116 329 292

440

1680

262 50

1370

1

1

1224

59

1165

–3654

368 50 0 0 0

–1647

–1 –1 –1

16429

49 180

1152 272 3612 470 146

10548

3 2 2 2 4

Note: Amino acid composition was determined for C. glutamicum MH 20-22B in continuous culture [23]. The intracellular pools of glutamate and glutamine contributing significantly to the anabolic demand were additionally considered. Nucleotide contents were determined from the relative GC content of C. glutamicum of 56%. The peptidoglycan content was estimated from the diaminopimelate content. Other constituents are taken from the literature [29].

Total

RNA DNA Lipids LPS Peptidoglycan Glycogen C1-units Polyamines

Protein

Thr Trp Tyr Val Dap

Metabolic Flux Analysis in Corynebacterium glutamicum 283

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12.2.3 ISOTOPE LABELING In order to overcome limitations of metabolite balancing, metabolic flux analysis employing stable isotopes such as 13C or 15N has emerged in recent years and displays — in combination with metabolite balancing — the state of the art in metabolic flux analysis [17,44,55]. In such studies, isotope-labeled compounds are used as tracer substrates. Their utilization by the cultivated cells leads to distribution of labeled atoms within the metabolic network, whereby the resulting metabolic labeling patterns specifically depend on the carbon–carbon transition of the involved reactions and their fluxes. The metabolic labeling patterns in such tracer studies can be measured by mass spectrometry (MS) or nuclear magnetic resonance (NMR) spectroscopy, respectively. Because the carbon transfer within the metabolic network is well defined, the labeling data can be used to calculate metabolic fluxes and overcome the limitations of sole metabolite balancing. The high potential of isotope labeling is illustrated for the problem of quantifying the flux partitioning at the glucose-6-phosphate node between PPP and glycolysis. Flux estimation for these pathways by sole metabolite balancing requires NADPH balancing, which is related to high uncertainty as depicted above. Carbon 13 flux analysis with [1-13C] glucose as tracer substrate, however, provides labeling data that sensitively depend on the flux partitioning between the two pathways and can therefore be used for precise estimation of their activity. The key for the flux estimation is the specificity of 6-phosphogluconate dehydrogenase, which exclusively releases the C1, the only 13C atom of the applied tracer substrate in the example in Figure 12.2A as CO2. All carbon channeled through the PPP and reentering glycolysis via fructose-6-phosphate or glyceraldehyde-3-phosphate is therefore unlabeled. Accordingly, the amount of 13C labeling of pyruvate and in the pyruvatederived metabolites decreases with increasing relative flux through the PPP. As revealed by metabolic simulations with varied flux partitioning between PPP and glycolysis [56], this is reflected (1) by a decrease of the ratio of single-labeled to unlabeled pyruvate and (2) by a decrease of the labeling degree of the C3 of pyruvate (Figure 12.2B). It becomes obvious, that both, MS and NMR, can be applied for quantification of the flux partitioning between PPP and glycolysis (Figure 12.2C). Mass spectroscopy can resolve single mass isotopomers differing by the number of labeled atoms and thus distinguish between single-labeled and unlabeled pyruvate in the present example. With NMR, the specific labeling degree of single atoms (fractional enrichment) can be measured. The real application of MS for quantifying the flux split between PPP and glycolysis is illustrated in Figure 12.2D, depicting MALDI-TOF MS spectra of intracellular pyruvate-derived alanine extracted from S. cerevisiae and from lysine-producing C. glutamicum in tracer studies on [1-13C]glucose (Wittmann, unpublished results). In comparison with S. cerevisiae, the fraction of single-labeled alanine is markedly reduced in C. glutamicum, revealing a significantly higher flux through the PPP in this organism, which is related to the increased NADPH demand for lysine production. For C. glutamicum, this approach allowed very precise estimation of PPP and glycolytic flux [57,58]. In a similar way, isotope labeling allows the quantification of the flux partitioning between the dual

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FIGURE 12.2 Quantification of the flux partitioning between PPP and glycolysis by 13C flux analysis. Transition of the 13C labeling through PPP and glycolysis employing [1-13C] glucose as tracer substrate (A). Fractional enrichment of the carbon C3 of pyruvate (dotted line) and ratio between single and unlabeled pyruvate (solid line) as function of the relative flux through the PPP revealed by metabolic simulations (B). Dependence of MS and NMR accessible labeling patterns of pyruvate on the relative flux through PPP (C). MALDI-TOF MS spectra of intracellular alanine from S. cerevisiae and C. glutamicum from tracer experiments on [1-13C] glucose obtained by quick filtration and subsequent extraction in boiling water (D).

pathways in lysine biosynthesis [43] and the resolution of all fluxes through the complex network around the pyruvate node [33]. The technique of choice for the experimental setup involving the tracer cultivation and method for labeling measurement depends on the goal of the performed study. Selected studies aiming at high resolution of the metabolic network under completely defined conditions will surely justify a high experimental effort. In this regard, 13C chemostat experiments with extensive labeling analysis of amino acids from cell protein hydrolysates by NMR [10,23,24] or 2D NMR [6,31,47] have proven useful in the past and provided detailed data on metabolic fluxes in C. glutamicum and other organisms. Hereby the hydrolysis of the cell protein gives access to the

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complete range of amino acids in which the carbon skeletons of the corresponding precursors (Table 12.1) are preserved in a defined and well-known way. Applying 2D NMR, complete isotopomer sets of amino acids can be obtained via the analysis of 13C multiplet hyperfine structures, thus providing rich information content for flux analysis. In addition, novel approaches (1) applicable to industrially relevant batch or fed-batch cultures, (2) providing high precision of flux estimates, and (3) allowing comparative flux analysis on a broad level with relatively low experimental effort have been recently developed and applied. These involve tracer experiments in small volume, such as minifermenters, shaken flasks, or microtiter plates, combined with labeling analysis by MS techniques such as MALDI-TOF MS [57] or GC/MS [13,58]. It turns out now that MS is particular useful towing to its superior sensitivity, therefore requiring smallest sample volumes only, and at the same time resulting in highest precision. Therefore, MS is expected to play a superior role in the future for flux analysis involving high-throughput approaches to screen large strain numbers or cultivation conditions and involving flux analysis under dynamic conditions, where a number of analyses of free metabolite pools are required to follow the change of the labeling patterns and to derive actual flux states.

12.2.4 ISOTOPOMER MODELING The mathematical method of choice to calculate fluxes from labeling data is (1) general so that all metabolic features, e.g., bidirectional fluxes or metabolic channeling can be considered; (2) readily amenable to modifications of the metabolic network; and (3) based on positional isotopomers so that all types of labeling information, e.g., mass isotopomer distributions or NMR coupling patterns can be included. In this context, general modeling frameworks with different tools for model generation, experimental design, parameter estimation, and statistical analysis are state of the art in metabolic flux analysis of C. glutamicum [55,59]. In specific cases, analytical equations can be formulated for metabolic flux calculation as previously presented for the estimation of the flux partitioning between PPP and glycolysis or between succinylase and dehydrogenase pathway in lysine biosynthesis in C. glutamicum [5]. These equations are however based on simplified networks, e.g., assuming only unidirectional fluxes for all reactions, and often so complex that even slight changes in the metabolic network require extensive and difficult modifications.

12.3 METABOLIC FLUXES IN C. GLUTAMICUM The first isotopic labeling studies of C. glutamicum and its subspecies flavum and ammoniagenes were carried out shortly after the isolation of the first overproducers in 1957. Initially, radioactive tracer substrates were applied for investigations of the metabolic network [40]. In 1982, a comprehensive study of central metabolic fluxes, e.g., through PEP carboxylase, citric acid cycle, and glyoxylate cycle, was carried out for glutamate biosynthesis in C. glutamicum, at that time called Microbacterium ammoniaphilum [49]. In this pioneering work, stable isotopic tracers, isotopomer analysis by 13C NMR, and mathematical equations relating labeling patterns and intracellular fluxes were combined for the first time in C. glutamicum. Subsequent

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studies were performed elucidating aspects of the metabolic network of C. glutamicum, e.g., revealing the in vivo presence of the dual lysine biosynthetic pathway [15], characterizing fluxes through PPP and glycolysis in a histidineproducing strain, C. glutamicum N-730 [16], and estimating selected flux parameters for glucose-grown C. melassecola ATCC 17965 [38]. These initial labeling studies together with the important contribution by Vallino and Stephanopoulos [46] stimulated recent developments of fully integrated approaches combining metabolite balancing and 13C-labeling for metabolic flux analysis of C. glutamicum. The most important studies on metabolic fluxes in C. glutamicum applying isotopic tracer substrates are discussed in detail in the following chapter.

12.3.1 FLUXES

FOR THE

GENERATION

OF

REDUCING POWER

Metabolic flux analysis by 13C tracer experiments allows establishing NADPH balances without additional assumptions on the energy metabolism. This is of particular importance because the supply of NADPH plays an important role for biosynthesis and amino acid overproduction in C. glutamicum. This holds especially true for the synthesis of lysine and isoleucine, for instance, characterized by a high demand of four NADPH per synthesized amino acid molecule. Therefore, quantification of fluxes through NADPH generating and consuming reactions is relevant for understanding and the targeted optimization of C. glutamicum. Approaches based on NMR [23] as well as on MALDI-TOF MS [57] led to almost identical flux distributions for different strains of C. glutamicum under lysine-producing conditions. In both cases, C. glutamicum revealed a high-PPP flux of 66 and 71%, respectively, and a correspondingly minor glycolytic flux. About 70% of the total NADPH is generated via the PPP, which therefore is the major pathway for supply of biosynthetic reducing power. The remaining 30% of NADPH is supplied by isocitrate dehydrogenase. These and similar studies with various strains and cultivation conditions [22,24,25,30,42,57,58] give a fascinating insight into the NAPDH metabolism of C. glutamicum (Figure 12.3A–C). In all cases, flux quantifications revealed an apparent excess of NADPH, indicating an as yet unassigned flux for NADPH consumption that is not related to product formation or cell material formation (Figure 12.3A). A potential route for NADPH consumption was recently identified by Matsushita et al., who showed that the respiratory chain in C. glutamicum is capable of oxidizing NADPH [27]. The large NADPH potential of 80% apparent excess during growth of the wild-type strain C. glutamicum ATCC 13032 surely displays a key feature for efficient amino acid production achieved in mutants derived from this parent strain during the past decades. The apparent NADPH excess, however, shows a significant decrease with increasing lysine yield, so that NADPH limitation of industrial lysine producers appears likely (Figure 12.3A). Indeed, a phosphoglucose isomerase–null mutant with the glucose-degradative flux forced to occur via the PPP results in a 1.7-fold increase in lysine accumulation [26]. The further comparison of metabolic fluxes in various lysine-producing strains of C. glutamicum shows that, depending on the lysine yield, the relative contribution of PPP and TCA cycle to NADPH generation differs markedly. Whereas the NADPH supply by the PPP exhibits a significant increase with increasing lysine yield, the

FIGURE 12.3 NADPH metabolism of C. glutamicum under various cultivation conditions as revealed by 13C flux analysis. Calculated fluxes of NADPH supply as a function of NADPH demand (A). Relative contribution of the PPP to the total NADPH supply as a function of the molar lysine yield (B). Relative contribution of isocitrate dehydrogenase to the total NADPH supply as a function of the molar lysine yield (C). Studies involved flux analysis of (1) nonproducing C. glutamicum ATCC 13032 in batch culture (open diamond) [42], (2) nonproducing C. glutamicum ATCC 13032 and lysine-producing C. glutamicum ATCC 13287, ATCC 21253, ATCC 21526, and ATCC 21543 in batch culture (open circle) [58], (3) lysineproducing C. glutamicum ATCC 21253 during maximum production phase in batch culture (open square) [57], (4) nonproducing C. glutamicum LE4 and lysine-producing C. glutamicum MH 20-22B in continuous culture (closed circles) [24], and (5) lysine-producing C. glutamicum MH 20-22B in continuous (closed square) [25]. G6P dehydrogenase, 6PG dehydrogenase, and isocitrate dehydrogenase were considered for calculation of the NADPH supply. The NADPH demand for lysine was four NADPH per lysine. The anabolic requirement for NADPH was recalculated for all studies considering the actual data on cellular composition with a demand of 16,400 μmol NADPH (g cell dry mass)–1 (Table 12.1).

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opposite is found for NADPH supply by isocitrate dehydrogenase (Figure. 12.B, C). Under conditions of pure growth, isocitrate is the main source for NADPH, whereas the PPP is the dominating pathway under lysine overproduction. In this context, C. glutamicum typically exhibits reduced growth during lysine production and, as shown above, a decreased apparent NADPH excess. This is probably linked to a decreased intracellular concentration of NADPH, which causes the enhanced PPP flux observed, since NADPH is a strong inhibitor of glucose-6-phosphate dehydrogenase and 6-phosphogluconate dehydrogenase in C. glutamicum [28].

12.3.2 ANAPLEROTIC FLUXES Corynebacterium glutamicum possesses multiple enzymes for the interconversion between glycolytic C3 metabolites and C4 metabolites of the TCA cycle. The different anaplerotic reactions are of special importance in C. glutamicum because many industrially relevant products, such as glutamate, lysine, threonine, or isoleucine, require a sufficient supply of oxaloacetate as a precursor for product synthesis. Analyses of the complex network around the pyruvate node have consistently revealed excessive substrate cycling in C. glutamicum in various physiological stages (Table 12.2). This illustrates that C4 decarboxylating activity is constitutively present in C. glutamicum even during pure growth on glucose. Although the details for the existence of the decarboxylating activities are not yet known, the resulting metabolic cycle of C3 carboxylating and C4 decarboxylating reactions might contribute to the metabolic flexibility of C. glutamicum regarding the supply of energy or anabolic precursor compounds. The importance of anaplerosis for amino acid production in C. glutamicum becomes obvious from the relationship between anaplerotic net flux

TABLE 12.2 Metabolic Cycling in the Anaplerotic Carboxylation of C. glutamicum in Various Physiological States as Determined by 13C Flux Analysis

Strain

Cultivation

Product

Anaplerotic Net Flux (%)

ATCC 13032 MH20-22B LE4 LE4 LE4 ATCC 21253 ATCC 13032 ATCC 13287 ATCC 21253 ATCC 21526 ATCC 21543

Batch Chemostat Chemostat Chemostat Chemostat Batch Batch Batch Batch Batch Batch

— Lysine — Glutamate Lysine Lysine Lysine Lysine Lysine Lysine Lysine

23 38 24 29 44 37 22 31 33 32 40

Note: The values given are related to the glucose uptake.

Carboxylation Flux (%)

Decarboxylation Flux (%)

Ref.

72 69 96 47 55 55 52 56 63 57 59

49 31 72 18 10 18 29 25 30 25 20

[40] [21] [22] [22] [23] [55] [56] [56] [56] [56] [56]

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FIGURE 12.4 Anaplerotic net flux of C. glutamicum as a function of the molar lysine yield determined under various cultivation conditions by 13C flux analysis. Studies involved flux analysis of (1) nonproducing C. glutamicum ATCC 13032 in batch culture (open diamond) [42] (2) nonproducing C. glutamicum ATCC 13032 and lysine-producing C. glutamicum ATCC 13287, ATCC 21253, ATCC 21526, and ATCC 21543 in batch culture (open circle) [58], (3) lysine-producing C. glutamicum ATCC 21253 during maximum production phase in batch culture (open square) [57], (4) nonproducing C. glutamicum LE4 and lysine-producing C. glutamicum MH 20-22B in continuous culture (closed circles) [24], and (5) lysine-producing C. glutamicum MH 20-22B in continuous (closed square) [25]. The anaplerotic flux is given as molar percentage of the corresponding specific glucose uptake rate.

and lysine yield obtained by the performed studies (Figure 12.4). Clearly, improved lysine formation is based on an enhanced anaplerotic net flux providing the product precursor oxaloacetate. This holds true for both carbon-limited chemostat studies and batch cultivations with nonlimiting conditions concerning the carbon source. The activity of anaplerotic enzymes during lysine production with a yield in a range of 0.25 to 0.30 mol per mole of glucose is about twice as high as compared with pure growth. With approaches involving specifically designed labeling experiments together with extended isotopomer analysis, the anaplerotic fluxes in C. glutamicum could be completely resolved. In an initial study, Park et al. [30] demonstrated the in vivo activity of pyruvate carboxylase and showed that it is the dominating anaplerotic enzyme in C. glutamicum. In their study, the authors used mutants of C. glutamicum with deletion of pyruvate kinase and/or PEP carboxylase and isotope labeling with MS and 13C NMR analysis. Recently, all anaplerotic fluxes around the pyruvate node could be completely resolved by a labeling study in continuous culture employing analysis of single isotopomers of amino acids from the cell protein by 2D NMR [33]. The flux estimation was based on the use of 21 mM glucose and 4 mM L-lactate consisting of a defined mixture of [1-13C]-glucose, [6-13C]-glucose, unlabeled glucose, and [3-13C]-lactate, leading to a differential labeling of the PEP and the pyruvate pools. The flux distribution obtained is shown in Figure 12.5. Pyruvate carboxylase was identified as major enzyme of anaplerotic carboxylation. Only very little activity

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FIGURE 12.5 Metabolic flux distribution in the anaplerosis of C. glutamicum ATCC 13032 involving PEP carboxylase, pyruvate carboxylase, PEP carboxykinase, malic enzyme, and oxaloacetate decarboxylase [33]. All values are given in mmol⋅(g cell dry mass)–1⋅h–1.

was observed for PEP carboxylase, which originally was assumed to be the sole anaplerotic enzyme in C. glutamicum. By using mainly pyruvate for anaplerotic supply, the organism probably avoids a limitation of PEP for the uptake of substrate molecules via the corresponding PTS systems. Excess oxaloacetate was recycled back to PEP by PEP carboxykinase, whereas oxaloacetate decarboxylase flux was below the detection limit. The resulting metabolic cycling leads to net consumption of about 1 mmol ATP⋅(g biomass)-1⋅h-1. Related to an energy demand of about 3.5 mmol ATP⋅(g biomass)-1⋅h-1 for biosynthesis, it therefore significantly contributes to the maintenance requirement in this organism [5]. C4-decarboxylating enzymes furthermore display important targets for metabolic engineering, owing to the fact that they withdraw the lysine precursor oxaloacetate. Subsequent prevention of decarboxylating flux by deletion of PEP carboxykinase leads up to 20% increased lysine accumulation [34,37].

12.3.3 FLUXES

IN A

GENEALOGY

OF

STRAINS

A comprehensive approach of 13C tracer studies with MS labeling analysis, metabolite balancing, and isotopomer modeling was applied for comparative metabolic flux analysis of a genealogy of five successive generations of lysine-producing C. glutamicum in batch cultures [58]. The aim of this study was to identify the consequences of strain optimization on the activities of the pathways in the metabolism of C. glutamicum and establish characteristic correlations between improved production properties and key fluxes. Strain optimization was accompanied by significant changes of intracellular flux distributions (Figure 12.6). The relative flux toward lysine increased markedly from 1.2 to 24.9%, posing a significantly enhanced

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demand for NADPH in the different production strains. This was reflected by successive increase of the relative PPP flux from 51% in the nonproducing wild-type strain to 64% in the mutant with the highest lysine yield. Drastic differences in metabolic fluxes resulted also for the pyruvate node. The anaplerotic net flux increased almost twofold to cover the increased demand for lysine formation, whereby the overall increase was a consequence of concerted up-regulation of C3 carboxylation and down-regulation of C4 decarboxylation fluxes. This is expressed by a more than twofold reduction of the reversibility (ratio between backward and net flux) of the involved bidirectional reactions. The ratio between backward flux and anaplerotic net flux decreased from 1.3 to about 0.5 with increasing lysine yield. The relative flux through isocitrate dehydrogenase dropped from 83% in the wildtype to 60% in the lysine-producing mutants, which exhibited reduced growth in comparison to the wild-type. Relative contribution of PPP and TCA cycle to NADPH generation differed markedly, indicating that C. glutamicum is able to maintain a certain supply of NADPH under completely different flux conditions. The flux through glyoxylate pathway was almost zero, indicating only marginal activity of this pathway. This is in accordance with the fact that the glyoxylate pathway is significantly suppressed during growth on glucose as compared with acetate as the carbon source [52]. Both alternative pathways in lysine biosynthesis were shown to be active in vivo and thus confirmed previous findings of 13C flux analysis by NMR in lysine-producing strains of C. glutamicum [41]. Phosphoglucose isomerase was highly reversible in all strains, which is probably related to fine adjustment of the intracellular pools of glucose-6-phosphate and fructose-6-phosphate. In addition, the interconversion of TCA cycle metabolites oxaloacetate, malate, and fumarate was found reversible in all strains. To summarize, genealogy profiling allows us to identify detailed consequences of strain selection. Combined with increasingly powerful sequencing techniques, it will be possible in the future to attribute observed changes on the fluxome level to specific mutations in key enzymes involved. Because mutations introduced by random mutagenesis are mainly point mutations, they could then easily be applied for targeted strain improvement [31].

12.3.4 FLUXES

ON

DIFFERENT CARBON SOURCES

Selected 13C flux studies on the influence of the carbon source have been performed for C. glutamicum ATCC 13032 growing in batch culture on glucose, acetate, and a mixture of glucose and acetate [50] and for C. melassecola ATCC 17965 growing in batch mode on fructose [9]. The study of the wild-type strain on glucose and acetate by Wendisch et al. [52] was carried out in batch cultures with [1-13C]-acetate, FIGURE 12.6 (opposite page) Metabolic flux distributions in a strain genealogy of lysine producers comprising C. glutamicum ATCC 13032, ATCC 13287, ATCC 21253, ATCC 21526, and ATCC 21543 (displayed in this order from top to bottom for each reaction) during the phase of lysine production in batch culture. All fluxes are expressed as a molar percentage of the mean specific glucose uptake rate (1.08 mmol⋅g-1⋅h-1 for ATCC 13032, 1.13 mmol⋅g1⋅h-1 for ATCC 13287, 1.13 mmol⋅g-1⋅h-1 for ATCC 21253, 1.05 mmol⋅g-1⋅h-1 for ATCC 21526, and 1.19 mmol⋅g-1⋅h-1 for ATCC 21543). Data were taken from [58].

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[5-13C]-glucose, or mixtures of [1-13C]-acetate plus unlabeled glucose applying combined metabolite balancing, NMR labeling measurement of amino acids from protein hydrolysates, and isotopomer modeling. Interestingly, C. glutamicum reveals nondiauxic growth with simultaneous metabolization of both acetate and glucose. Completely different flux situations are present on the different carbon sources (Figure 12.7 A–C). Whereas on glucose the flux pattern agreed with previous findings and resulted with a high relative flux through the PPP and an inactive glyoxylate cycle (Figure 12.7A), a completely different flux distribution was observed for growth on acetate. Hereby the entry point of acetate at the level of acetyl-CoA resulted in a high flux of 18% through the glyoxylate cycle (Figure 12.7B). Under these conditions, C. glutamicum exhibits a strong gluconeogenetic net flux through PEP carboxykinase of 54% to supply anabolic precursors of upper glycolysis (G6P, F6P) and of the PPP (E4P, R5P). During cultivation of C. glutamicum on a mixture of the two substrates, the relative specific uptake rate for acetate was about threefold higher compared with glucose, leading to a higher relative entry of carbon at the level of acetate as compared with G6P (Figure 12.7B). Interestingly, the anaplerotic function was also completely fulfilled by the glyoxylate cycle under these conditions. This is a surprising result because the presence of glucose did not essentially require this flux orientation. The conversion between glycolytic C3 metabolites and C4 metabolites of the TCA cycle is highly dependent on the available substrate. On glucose a high anaplerotic net flux of 30% is present, whereas on acetate and on acetate plus glucose gluconeogenetic fluxes of 54 and 12% were observed. In all cases, the conversion between glycolytic C3 metabolites and C4 metabolites of the TCA cycle was found reversible. Drastic differences resulted also for pyruvate dehydrogenase, which was highly active on glucose (109%), weakly active (12%) during growth on acetate/glucose, and inactive during growth on acetate. Glucose-6-phosphate isomerase operated in opposite directions depending on the carbon source, underlining the importance of the reversibility of this enzyme for metabolic flexibility in C. glutamicum. The relative flux through the TCA cycle was rather similar in all cases. Taking the different absolute fluxes for substrate uptake into account, the absolute activity of the TCA cycle was high on glucose, medium on glucose plus acetate, and low on acetate. Interesting insights into the metabolism of C. melassecola ATCC 17965 were obtained for exponential growth on fructose via two parallel experiments on [1-13C]and [6-13C]-glucose and NMR measurement of the fractional enrichment of intracellular glutamate from both cultivations. It was observed that fructose is taken up by C. melassecola simultaneously via a fructose and a mannose PTS and as a result of this enters the glycolysis at the two locations of (1) fructose-1,6-bisphosphate via fructose-1-phosphate and (2) fructose-6-phosphate, respectively. Selected flux estimations were carried out by directly calculating the relative contribution of glycolysis to substrate catabolization from the ratio of specific enrichments of single glutamate carbons with [1-13C]-glucose and [6-13C]-glucose, respectively. This simplified mathematical approach neglecting, e.g., bidirectional or anabolic fluxes, allows only an approximate flux estimation. Glycolysis contributed to about 80% to the catabolism of fructose, whereas the PPP made up the remaining 20%. This is a remarkable difference as compared with studies of C. glutamicum on glucose revealing a high

FIGURE 12.7 Metabolic flux distributions of C. glutamicum ATCC 13032 growing on glucose (A), a mixture of acetate and glucose (B) and acetate as carbon source. All fluxes are expressed as a molar percentage of the specific uptake rate, which was 148 nmol glucose⋅(mg protein)-1⋅min-1 (A), 270 nmol acetate⋅(mg protein)-1⋅min-1 (B), and 540 nmol acetate⋅(mg protein)-1⋅min-1 (C). Data were taken from [52].

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activity of the PPP [23,57,58]. It should be mentioned that fructose as carbon source leads to a 30% reduced yield for L-lysine production by C. glutamicum in comparison with glucose [18], which is probably related to the different entry points of the substrates into the central metabolism and resulting consequences on the NADPH metabolism. To summarize, almost all studies on metabolic fluxes in C. glutamicum have been carried out with glucose as substrate, with the consequence that most of our knowledge about the physiology of C. glutamicum is based on the metabolism of this substrate. In contrast, little attention was paid to other carbon sources, which, however, are of high importance for C. glutamicum. Important substrates involve, e.g., fructose and sucrose concerning industrial production processes based on cane or beet molasses. In addition, flux studies on substrates, such as acetate or lactate, are of relevance since they might be major by-products of industrial amino acid production and are probably rapidly reutilized in fermentations. Moreover, the use of alternative substrates channeled into the central metabolism at different entry points is an important possibility to study the central metabolism of C. glutamicum under completely different flux scenarios that may contribute to the understanding of its function and regulatory properties.

12.3.5 RESPONSE

OF

FLUXES

TO

DIFFERENT CELLULAR DEMANDS

Knowledge about the response of C. glutamicum to metabolic burdens caused by overproduction of single products or by high anabolic demands is important for understanding metabolic flexibility and rigidity and the underlying regulatory mechanisms in this organism. The impact of cellular demands on metabolic fluxes was studied by tracer studies with isogenic strains of C. glutamicum in chemostat [23] as well as two strains grown in batch culture [42]. Depending on the growth phase, C. glutamicum exhibited tremendous differences in metabolic fluxes. The flux maps depicted in Figure 12.8A–C illustrate the enormous flexibility of the central metabolism in C. glutamicum. As an example, the relative flux into the PPP increased drastically from 25% for glutamate production to 66% for lysine production, with the glycolytic flux accordingly decreasing. This reflects that the flexibility of the flux partitioning between these two pathways in C. glutamicum is a general characteristic of this organism, probably serving to maintain a constant intracellular NADPH concentration. Marked differences between the isogenic strains were also found for the flux partitioning at the pyruvate node. The net flux over anaplerotic carboxylation increased successively from growth (24%) and glutamate production (29%) to lysine production (37%). Accordingly, relative flux through pyruvate dehydrogenase revealed a strong decrease from glutamate production (135%) to lysine production (80%). The reason for the latter is the simultaneous demand for pyruvate and oxaloacetate for lysine production, which results in a high direction of carbon into anaplerosis and lysine biosynthesis, respectively. Summarizing, these flux analyses revealed a tremendous flexibility in the central metabolism of C. glutamicum that is capable of adjusting the carbon fluxes through its central metabolism over a broad range depending on the cellular demands. However, this does not exclude that in extreme flux scenarios NADPH might become limiting, as is evident from a study with a phosphoglucose isomerase–null mutant, showing 1.7-fold increased lysine

FIGURE 12.8 Metabolic flux distributions of isogenic strains of C. glutamicum producing no amino acid (A), glutamate (B), and lysine (C). All fluxes are expressed as a molar percentage of the specific uptake rate. Data were taken from [24].

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formation [26] or a transketolase-overexpressing strain that accumulated up to 20% more aromatic amino acids [14]. A study by Wittmann and Heinzle investigated metabolic fluxes in C. glutamicum ATCC 21253 during a phase of maximum lysine production in batch mode [57], thus enabling an interesting comparison with strain MH 20-22B analyzed under similar conditions [42]. Several interesting flux differences arose, probably owing to different precursor demands for growth and lysine production of the two strains. Whereas the lysine flux was comparable for both strains, C. glutamicum ATCC 21253 exhibited a 200% increased biomass yield and an increased secretion of various by-products. Anaplerotic net flux and PPP flux were significantly higher for C. glutamicum ATCC 21253 as compared with strain MH 20-22B, whereas the TCA cycle flux was markedly decreased. The higher biomass yield and by-product formation of C. glutamicum ATCC 21253 lead to a higher withdrawal of carbon over the entire network. This results in a lower fraction of carbon available at the level of pyruvate dehydrogenase and TCA cycle and causes a 50% reduced NADPH formation in the TCA cycle. Obviously, this is compensated for by an enhanced flux through the PPP, a phenomenon already observed in other studies (Figure 12.7). The increased anaplerotic net flux of C. glutamicum ATCC 21253 reflects the high demand for oxaloacetate for both growth and lysine production.

12.3.6 NITROGEN FLUXES The assimilation of nitrogen, i.e., ammonium, in C. glutamicum plays a key role for amino acid overproduction. Moreover, it is important for various growth-supporting reactions. This accounts for the high interest in understanding the functioning and regulation of the ammonium assimilation system in C. glutamicum. Glutamate dehydrogenase (GDH) and glutamine synthetase (GS)-glutamine-2-oxoglutarate-aminotransferase (GOGAT) represent the two main pathways for ammonium assimilation in C. glutamicum. In an early study, 15N-labeling combined with in vivo NMR was successfully applied to study regulation of nitrogen assimilation and amino acid production in C. glutamicum [12]. It has long been assumed that GDH is a key enzyme for ammonium assimilation in this organism, but surprisingly a GDHnegative mutant of the wild-type C. glutamicum ATCC 13032 did not exhibit a different phenotype compared with the parent strain [1]. This raised questions for possible alternative assimilatory routes, e.g., via alanine dehydrogenase or an altered regulation. In order to investigate this in detail, ammonium assimilation fluxes of the wild-type and the GDH mutant were quantified by a dynamic labeling study employing a membrane-cyclone bioreactor coupled to in vivo NMR [45]. At metabolic steady state, the ammonium concentration was increased within 2 min from 40 mmol/L to 150 mmol/L by the feed of 15N-labeled ammonium sulfate. As displayed in Figure 12.9A, the position of the incorporated 15N during assimilation depends on the pathway involved. On-line monitoring of 15N enrichment in glutamate (Nα) and glutamine (Nα, Nδ) allows the differentiation between GDH and GS-GOGAT activity. Manual inspection of the time course of the labeling data revealed marked differences between the wild-type and the GDH mutant. In the wild-type, strong 15N

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FIGURE 12.9 Nitrogen fluxes of C. glutamicum. Strategy for the flux quantification by feeding of [15N] ammoniumsulfate and using in vivo NMR (A). Nitrogen fluxes of C. glutamicum ATCC 13032 (B) and a glutamate dehydrogenase negative mutant (C). All fluxes are given in μmol⋅(g cell dry mass)-1⋅min-1. GOGAT, glutamine:2-oxoglutarate aminotransferase; GDH, glutamate dehydrogenase; TA, transaminases; GS, glutamine synthetase; AT, aminotransferase. Data were taken from [45]. The white and gray shaded nitrogen atoms display 14N and 15N, respectively.

enrichment was found for the Nα of glutamate and glutamine, pointing at a dominating role of GDH. In contrast, the major fraction of 15N was incorporated in the Nδ of glutamine in the GDH mutant indicating that the GS-GOGAT system, which is linked to an integration of NH4+ in the side chain of glutamine, was mainly responsible for ammonium assimilation in the mutant. These findings point at completely different flux scenarios in the two strains. Quantitative flux distributions in the ammonium-assimilating network were obtained by fitting the measurement data to a nitrogen flux model (Figure 12.9B, C). Whereas in the wild-type, 72% of the ammonium was assimilated via GDH and 28% via GS, 100% was assimilated via GOGAT in the GDH mutant. In the wild-type, glutamate resulted from operation of GDH (74%) and glutamine transaminases (26%). In the GDH mutant, the task of GDH was completely taken over by GOGAT, via which 72% of the glutamate was produced. The absence of the flux through GOGAT in the wild-type showed excellent accordance with enzyme measurements, as did the increased activity of this enzyme in the GDH mutant [45].

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12.4 CONCLUDING REMARKS During recent years, metabolic flux analysis has become a powerful tool for providing detailed quantitative knowledge on the in vivo activity of the various enzymes and pathways involved in the central metabolic network. Owing to the industrial interest and the impact on further strain improvement, C. glutamicum certainly occupies a prominent position in applications and further developments of this tool. A number of unique achievements have already been obtained, which are described in this article and which include even the resolution of parallel, exchange, and cyclic fluxes, a result previously thought to be hardly achievable. Future perspectives of the rapidly developing field of metabolic flux analysis of C. glutamicum are seen in other areas. One direction will surely be further development and application of metabolic flux approaches allowing comparative flux studies on a broad level. These approaches will involve miniaturized cultivation methods in milliliter-to-micoliter scale combined with powerful MS measurement tools [60]. This will allow parallel metabolic flux screening of various mutants or cultivation conditions in reasonable time [19,61,62], thus establishing metabolic flux analysis as a routine profiling tool. A further important future perspective is seen in metabolic flux analysis under dynamic conditions aiming at the determination of the in vivo kinetics of enzymes and of metabolic regulation. Such approaches will be based on labeling analysis of free intracellular metabolites reflecting the actual flux state of the examined cells and applicable to industrially relevant batch or fed-batch processes. Finally, combination of metabolic flux analysis with other tools, such as transcriptome or proteome analysis, seems very promising for deeper investigation of C. glutamicum [22]. Such system-oriented approaches will probably yield a significantly improved understanding of metabolic regulation and targeted optimization of C. glutamicum to improve currently established processes and create novel ones.

REFERENCES 1. Börmann-El Kholy ER, Eikmanns BJ, Gutmann M, and Sahm H. (1993) Glutamate dehydrogenase is not essential for glutamate formation by Corynebacterium glutamicum. Appl. Environ. Microbiol. 59:2329–2331. 2. Chassagnole C, Létisse F, Diano A, and Lindley ND. (2002) Carbon flux analysis in a pantothenate overproducing Corynebacterium glutamicum strain. Mol. Biol. Rep. 29:129–134. 3. Cocaign-Bousquet M and Lindley ND. (1995) Pyruvate overflow and carbon flux within the central metabolic pathways of Corynebacterium glutamicum during growth on lactate. Enz. Microbiol. Technol. 17:260–267. 4. Cocaign-Bousquet M, Guyonvarch A, and Lindley ND. (1996) Growth rate-dependent modulation of carbon flux through central metabolism and the kinetic consequences for glucose-limited chemostat cultures of Corynebacterium glutamicum. Appl. Environ. Microbiol. 62:429–436. 5. de Graaf AA. (2000) Metabolic flux analysis of Corynebacterium glutamicum. In Schügerl K and Bellgard KH (Eds.), Bioreaction Engineering, Springer Verlag, Berlin, Heidelberg, p. 506–555.

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6. Dauner M, Bailey JE, and Sauer U. (2001) Metabolic flux analysis with a comprehensive isotopomer model in Bacillus subtilis. Biotechnol. Bioeng. 76:144–156. 7. Dominguez H, Nezondet C, Lindley ND, and Cocaign M. (1993) Modified carbon flux during oxygen limited growth of Corynebacterium glutamicum and the consequences for amino acid overproduction. Biotechnol. Lett. 15:449–454. 8. Dominguez H and Lindley ND. (1996) Complete sucrose metabolism requires fructose phosphotransferase activity in Corynebacterium glutamicum to ensure phosphorylation of liberated fructose. Appl. Environ. Microbiol. 62:3878–3880. 9. Dominguez H, Rollin C, Guyonarch A, Guerquin-Kern JL, Cocaign-Bousquet M, and Lindley ND. (1998) Carbon-flux distribution in the central metabolic pathways of Corynebacterium glutamicum during growth on fructose Eur. J. Biochem. 254:96–102. 10. Drysch A, El Massaoudi M, Mack C, Takors R, de Graaf AA, and Sahm H. (2003) Production process monitoring by serial mapping of microbial carbon flux distributions using a novel Sensor Reactor approach: 13C-labeling-based metabolic flux analysis and L-lysine production. Metabol. Eng. 5:96–107. 11. Eggeling L and Sahm H. (1999) L-Glutamate and L-lysine: traditional products with impetuous developments. Appl. Microbiol. Biotechnol. 52:46–53. 12. Haran N, Kahana, ZE, and Lapidot A. (1983) In vivo 15N NMR studies of regulation of nitrogen assimilation and amino acid production by Brevibacterium lactofermentum. J. Biol. Chem. 258:12929–12933. 13. Heinzle E and Wittmann C. (2002) Proceedings of 4th European Symposium on Biochemical Engineering Science. Delft, The Netherlands, August 28–31, p. 4. 14. Ikeda M, Okamoto K, and Katsumata R. (1999) Cloning of the transketolase gene and the effect of its dosage on aromatic amino acid production in Corynebacterium glutamicum. Appl. Microbiol. Biotechnol. 51:201–206. 15. Ishino S, Kuga T, Yamaguchi K, Shirahata K, and Araki K. (1984) Involvement of meso-diaminopimelate D-dehydrogenase in lysine biosynthesis in Corynebacterium glutamicum. Agric. Biol. Chem. 48:2557–2560. 16. Ishino S, Kuga T, Yamaguchi K, Shirahata K, and Araki K. (1986) 13C NMR studies of histidine fermentation with a Corynebacterium glutamicum mutant. Agric. Biol. Chem. 50:307–310. 17. Kelleher JK. (2001) Flux estimation using isotopic tracers: common ground for metabolic physiology and metabolic engineering. Metabol. Eng. 3:100–110. 18. Kiefer P, Heinzle E, and Wittmann C. (2002) Influence of glucose, fructose and sucrose as carbon sources on kinetics and stoichiometry of lysine production by Corynebacterium glutamicum. J. Ind. Microbiol Biotechnol. 28:338–343. 19. Kiefer P, Heinzle E, Zelder O, Wittmann C. (2004) Comparative metabolic flux analysis of lysine-producing Corynebacterium glutamicum cultured on glucose or fructose. Appl. Env. Microbiol. 70:229–239. 20. Kinoshita S, Udaka S, and Shimono M. (1957) Studies on the amino acid fermentation part I. production of L-glutamic acid by various microorganisms. J. Gen. Appl. Microbiol. 3:193–205. 21. Kinoshita S and Tanaka K. (1972) Glutamic acid. In Yamada K (Ed.), The Microbial Production of Amino Acids, Wiley, New York, pp. 263–324. 22. Krömer JO, Sorgenfrei O, Klopprogge K, Heinzle E, Wittmann C. (2004) In-depth profiling of lysine producing Corynebacterium glutamicum by combined analysis of transcriptome, metabolome and fluxome. J. Bacteriol. 186:1769–1784. 23. Marx A, de Graaf AA, Wiechert W, Eggeling L, and Sahm H. (1996) Determination of the fluxes in the central metabolism of Corynebacterium glutamicum by nuclear magnetic resonance spectroscopy combined with metabolite balancing. Biotechnol. Bioeng. 49:111–129.

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24. Marx A, Striegel K, de Graaf AA, Sahm H, and Eggeling L. (1997) Response of the central metabolism of Corynebacterium glutamicum to different flux burdens. Biotechnol. Bioeng. 56:168–180. 25. Marx A, Eikmanns B, Sahm H, de Graaf AA, and Eggeling L. (1999) Response of the central metabolism in Corynebacterium glutamicum to the use of an NADHdependent glutamate dehydrogenase. Metabol. Eng. 1:35–48. 26. Marx A, Hans S, Möckel B, Bathe B, and de Graaf AA. (2003) Metabolic phenotype of phosphoglucose isomerase mutants of Corynebacterium glutamicum. J. Biotechnol. 104:185–197. 27. Matsushita K, Otofuji A, Iwahashi M, Toyama H, and Adachi O. (2001) NADH dehydrogenase of Corynebacterium glutamicum. Purification of an NADH dehydrogenase II homolog able to oxidize NADPH. FEMS Microbiol. Lett. 204:271–276. 28. Moritz B, Striegel K, de Graaf AA, and Sahm H. (2000) Kinetic properties of the glucose-6-phosphate and 6-phosphogluconate dehydrogenases from Corynebacterium glutamicum and their application for predicting pentose phosphate pathway flux in vivo. Eur. J. Biochem. 267:3442–3452. 29. Neidhardt FC, Igraham JL, and Schaechter M. (1990) Physiology of the Bacterial Cell: A Molecular Approach. Sinauer Associates, Inc., Sunderland, Massachusetts. 30. Oishi K and Aida K. (1965) Studies on amino acid fermentation. Part XI. Effect of biotin on the Embden-Meyerhof-Parnas pathway and the hexose-monophosphate shunt in a glutamic acid-producing bacterium, Brevibacterium ammoniagenes. Agric. Biol. Chem. 29:83–89. 31. Ohnishi J, Mitsuhashi S, Hayashi M, Ando S, Yokoi H, Ochiai K, and Ikeda MA. (2002) A novel methodology employing Corynebacterium glutamicum genome information to generate a new L-lysine-producing mutant Appl. Microbiol. Biotechnol. 58:217–223. 32. Park SM, Shaw-Reid C, Sinskey AJ, and Stephanopoulos G. (1997) Elucidation of anaplerotic pathways in Corynebacterium glutamicum via 13C-NMR spectroscopy and GC-MS. Appl. Microbiol. Biotechnol. 47:430–440. 33. Petersen S, De Graaf AA, Eggeling L, Möllney M, Wiechert W, and Sahm H. (2000) In vivo quantification of parallel and bidirectional fluxes in the anaplerosis of Corynebacterium glutamicum. J. Biol. Chem. 275:35932–35941. 34. Petersen S, Mack C, de Graaf AA, Riedel C, Eikmanns BJ, and Sahm H. (2001) Metabolic consequences of altered phosphoenolpyruvate carboxykinase activity in Corynebacterium glutamicum reveal anaplerotic regulation mechanisms in vivo. Metabol. Eng. 3:344–361. 35. Peters-Wendisch PG, Wendisch VF, Paul S, Eikmanns BJ, and Sahm H. (1997) Pyruvate carboxylase as an anaplerotic enzyme in Corynebacterium glutamicum. Microbiol. 143:1095–1103. 36. Pons A, Dussap G, Péquignot C, and Gros JB. (1996) Metabolic flux distribution in Corynebacterium melassecola ATCC 17965 for various carbon sources. Biotechnol. Bioeng. 51:177–189. 37. Riedel C, Rittmann D, Dangel P, Möckel B, Petersen S, Sahm H, and Eikmanns BJ. (2001) Characterization of the phosphoenolpyruvate carboxykinase gene from Corynebacterium glutamicum and significance of the enzyme for growth and amino acid production. J. Mol. Microbiol. Biotechnol. 3:573–583. 38. Rollin C, Morgant V, Guyonvarch A, and Guerquin-Kern JL. (1995) 13C-NMR studies of Corynebacterium melassecola metabolic pathways. Eur. J. Biochem. 227:488–493.

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39. Sahm H, Eggeling L, Eikmanns BJ, and Krämer R. (1995) Metabolic design in amino acid producing bacterium Corynebacterium glutamicum. FEMS Microbiol. Rev. 16:243–252. 40. Shiio I, Otsuka SI, and Tsunoda T. (1960) Glutamic acid formation from glucose by bacteria. J. Biochem. 47:414–421. 41. Sonntag K, Eggeling L, de Graaf AA, and Sahm H. (1993) Flux partitioning in the split pathway of lysine synthesis in Corynebacterium glutamicum. Quantification by 13C- and 1H-NMR spectroscopy. Eur. J. Biochem. 213:1325–1331. 42. Sonntag K, Schwinde J, de Graaf AA, Marx A, Eikmanns BJ, Wiechert W, and Sahm H. (1995) 13C NMR studies of the fluxes in the central metabolism of Corynebacterium glutamicum during growth and overproduction of amino acids in batch cultures. Appl. Microbiol. Biotechnol. 44:489–495. 43. Stephanopoulos G, Aristidou AA, and Nielsen J. (1998) Metabolic Engineering. Academic Press, New York. 44. Szyperski T. (1998) 13C-NMR, MS and metabolic flux balancing in biotechnology research. Q. Rev. Biophys. 31:41–106. 45. Tesch M, de Graaf AA, and Sahm H. (1999) In vivo fluxes in the ammoniumassimilatory pathways in Corynebacterium glutamicum studied by 15N nuclear magnetic resonance. Appl. Environ. Microbiol. 65:1099–1109. 46. Vallino JJ and Stephanopoulos G. (1993) Metabolic flux distributions in Corynebacterium glutamicum during growth and lysine overproduction. Biotechnol. Bioeng. 41:633–646. 47. Vallino JJ and Stephanopoulos G. (1994) Carbon flux distributions at the pyruvate branch point in Corynebacterium glutamicum during lysine overproduction Biotechnol. Prog. 10:320–326. 48. Vallino JJ and Stephanopoulos G. (1994) Carbon flux distributions at the glucose 6-phosphate branch point in Corynebacterium glutamicum during lysine overproduction. Biotechnol. Prog. 10:327–334. 49. van Winden WA, van Gulik WM, Schipper D, Verheijen PJ, Krabben P, Vinke JL, and Heijnen JJ. (2003) Metabolic flux and metabolic network analysis of Penicillium chrysogenum using 2D [13C, 1H] COSY NMR measurements and cumulative bondomer simulation. Biotechnol Bioeng. 83:75–92. 50. Varela C, Agosin E, Baez M, Klapa M, and Stephanopoulos G. (2003) Metabolic flux redistribution in Corynebacterium glutamicum in response to osmotic stress. Appl. Microbiol. Biotechnol. 60:547–555. 51. Walker TE, Han CH, Kollman VH, London RE, and Matwiyoff NA. (1982) 13C Nuclear magnetic resonance studies of the biosynthesis by Microbacterium ammoniaphilum of L-glutamate selectively enriched with 13C. J. Biol. Chem. 257:1189–1195. 52. Wendisch VF, De Graaf AA, Sahm H, and Eikmanns BJ. (2000) Quantitative determination of metabolic fluxes during coutilization of two carbon sources: comparative analyses with Corynebacterium glutamicum during growth on acetate and/or glucose. J. Bacteriol. 182:3088–3096. 53. Wendisch VF. (2003) Genome-wide expression analysis in Corynebacterium glutamicum using DNA microarrays. J. Biotechnol. 104:273–285. 54. Wiechert W. (2001) 13C metabolic flux analysis. Metabol. Eng. 3:195–206. 55. Wiechert W, Möllney M, Petersen S, and de Graaf AA. (2001) A universal framework for 13C metabolic flux analysis. Metabol. Eng. 3:265–283.

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56. Wittmann C and Heinzle E. (2001) Modeling and experimental design for metabolic flux analysis of lysine-producing Corynebacteria by mass spectrometry. Metabol. Eng. 3:171–193. 57. Wittmann C and Heinzle E. (2001) Application of MALDI-TOF MS to lysineproducing Corynebacterium glutamicum: a novel approach for metabolic flux analysis. Eur. J. Biochem. 268:2441–2455. 58. Wittmann C and Heinzle E. (2002) Genealogy profiling through strain improvement by using metabolic network analysis: metabolic flux genealogy of several generations of lysine-producing corynebacteria. Appl. Environ. Microbiol. 68:5843–5859. 59. Wittmann C. (2002) Mass spectrometry for metabolic flux analysis. Adv. Biochem. Eng. Biotechnol. 74:39–64. 60. Wittmann C, Kim HM, and Heinzle E. (2004) Metabolic flux analysis of lysine producing Corynebacterium glutamicum at miniaturized scale. Biotechnol. Bioeng. 87:1–6. 61. Wittmann C, Kiefer P, and Zelder O. (2004) Metabolic fluxes in Corynebacterium glutamicum during lysine production on sucrose. Appl. Env. Microbiol. in press. 62. Wittmann C and Heinzle E. (2004) Metabolic activity profiling by 13C tracer experiments and mass spectrometry in Corynebacterium glutamicum. In Barredo JL (Ed.), Methods in Biotechnology, Vol. 18. Microbial Processes and Products, Humana Press, Totowa, NJ, pp. 151–163.

13

Respiratory Energy Metabolism M. Bott and A. Niebisch

CONTENTS 13.1 Introduction ..................................................................................................305 13.2 Electron Transfer from Substrates to Menaquinone ...................................306 13.2.1 NADH Dehydrogenase ....................................................................306 13.2.2 Succinate Dehydrogenase ................................................................311 13.2.3 Malate:Quinone Oxidoreductase......................................................312 13.2.4 Pyruvate:Quinone Oxidoreductase (Pyruvate Oxidase) ..................313 13.2.5 Lactate Dehydrogenases ..................................................................313 13.2.6 Glycerol-3-Phosphate Dehydrogenase.............................................314 13.2.7 Proline Dehydrogenase ....................................................................315 13.2.8 Electron-Transferring Flavoprotein..................................................315 13.3 Electron Transfer from Menaquinol to Oxygen..........................................316 13.3.1 Cytochrome bc1 Complex ................................................................317 13.3.2 Cytochrome aa3 Oxidase .................................................................318 13.3.3 Identification of a Cytochrome bc1-aa3 Supercomplex...................321 13.3.4 Cytochrome bd Menaquinol Oxidase ..............................................321 13.3.5 Alternative Oxidase Activities .........................................................322 13.4 Electron Transfer from Menaquinol to Nitrate ...........................................322 13.5 Heme Biosynthesis and Cytochrome c Maturation.....................................323 13.6 Impact of F1F0-ATP Synthase on Metabolism ............................................324 13.7 Influence of Respiratory Chain Composition on the ATP Yield.................325 13.8 Biotechnological Aspects.............................................................................326 13.9 Concluding Remarks....................................................................................327 Acknowledgments..................................................................................................327 References..............................................................................................................327

13.1 INTRODUCTION Corynebacterium glutamicum has a respiratory kind of energy metabolism, with oxygen as the terminal electron acceptor. Growth by fermentative catabolism or by anaerobic respiration has not been reported. The first studies on the respiratory chain of this organism were triggered by the observation that copper deficiency decreased 305

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glutamate formation from acetate by an oleic acid–requiring C. glutamicum strain (Brevibacterium thiogenitalis) owing to an increased oxidation of acetate to CO2 [28]. The effect of copper was due to a functional change of the respiratory chain. Evidence was obtained that under copper sufficiency electron transfer from NADH to oxygen involves a cytochrome bc1 complex and cytochrome aa3 oxidase, whereas under copper deficiency a- and c-type cytochromes are missing and instead cytochrome d was formed [69,70]. The studies made in the 1970s and 1980s [30,74] were continued only recently by several groups, leading to a much more detailed view of the respiratory chain in C. glutamicum. This chapter summarizes the current knowledge on this topic, including relevant information from the genome sequence [23,26] and on aspects that have not yet been studied experimentally. A compilation of the genes involved in respiratory energy metabolism is given in Table 13.1, which also accounts for the different annotations that are presently available for the C. glutamicum genome. A schematic overview of the components that constitute the respiratory chain is given in Figure 13.1, whereas Figure 13.2 shows in more detail their subunit composition, localization, cofactors, and involvement in the generation of a proton-motive force.

13.2 ELECTRON TRANSFER FROM SUBSTRATES TO MENAQUINONE Based on experimental data and genome information, at least eight dehydrogenases are present in C. glutamicum that transfer electrons into the respiratory chain using menaquinone as the primary acceptor. These are NADH dehydrogenase, succinate dehydrogenase, malate:quinone oxidoreductase, pyruvate:quinone oxidoreductase (pyruvate “oxidase”), D-lactate dehydrogenase, L-lactate dehydrogenase, glycerol-3phosphate dehydrogenase, and proline dehydrogenase. As in other gram-positive bacteria, menaquinone was identified as the only isoprenoid quinone present in C. glutamicum, with dihydromenaquinone-8 (i.e., menaquinone containing eight isoprene units with one double bond hydrogenated) and menaquinone-9 as minor components and dihydromenaquinone-9 as major component [11,12,29]. Menaquinone biosynthesis starts from chorismate, an intermediate in the biosynthesis of the aromatic amino acids. Inspection of the genome sequence led to the identification of all genes required for menaquinone synthesis, i.e., menF (isochorismate synthase), menD (2-oxoglutarate decarboxylase and 2-succinyl-6-hydroxy-2,4-cyclohexadiene-1-carboxylate synthase), menC (o-succinylbenzoate synthase), menE (o-succinylbenzoyl-CoA synthetase), menB (1,4-dihydroxy-2-naphthoate synthase), menA (1,4-dihydroxy-2-naphthoate prenyltransferase), and menG (demethylmenaquinone methyltransferase) [5].

13.2.1 NADH DEHYDROGENASE Early inhibitor studies with amytal and rotenone by Sugiyama et al. [70] had indicated the absence of a mitochondrial complex I-type NADH dehydrogenase in C. glutamicum. This result was confirmed by the genome sequence [23,26], revealing the absence of the nuo operon encoding this multisubunit enzyme in bacteria. Instead,

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TABLE 13.1 Compilation of C. glutamicum Genes Encoding Respiratory Chain Components, F1F0-ATP Synthase, Menaquinone Biosynthesis, Heme Biosynthesis, and Cytochrome c Maturation Gene

ndh sdhC sdhA sdhB mqo pqo dld lldD glpD putA etfB etfA qcrC qcrA qcrB ctaD ctaC ctaE ctaF cydA cydB narG narH narJ narI

Proven or Anticipated Function of the Gene Product

Cgl No.

NCgl No.

Respiratory Chain Components NADH dehydrogenase, non-protonCgl1465 NCgl1409 pumping Succinate dehydrogenase, cytochrome b Cgl0370 NCgl0359 subunit Succinate dehydrogenase, flavoprotein Cgl0371 NCgl0360 subunit Succinate dehydrogenase, iron-sulfur Cgl0372 NCgl0361 protein subunit Malate:quinone oxidoreductase Cgl2001 NCgl1926 pyruvate:quinone oxidoreductase, Cgl2610 NCgl2521 (“pyruvate oxidase“) D-Lactate dehydrogenase, (using MQ as Cgl0901 NCgl0865 acceptor) L-Lactate dehydrogenase, (using MQ as Cgl2918 NCgl2817 acceptor) Glycerol-3-phosphate dehydrogenase Cgl1646 NCgl1584 L-Proline dehydrogenase Cgl0099 NCgl0098 Electron-transferring flavoprotein, Cgl1230 NCgl1182 subunit Electron-transferring flavoprotein, Cgl1231 NCgl1183 subunit α Cytochrome bc1 complex, cytochrome c1 Cgl2191 NCgl2111 subunit Cytochrome bc1 complex, Rieske ironCgl2190 NCgl2110 sulfur protein Cytochrome bc1 complex, cytochrome b Cgl2189 NCgl2109 subunit Cytochrome aa3 oxidase, subunit I Cgl2523 NCgl2437 Cytochrome aa3 oxidase, subunit II Cgl2195 NCgl2115 Cytochrome aa3 oxidase, subunit III Cgl2192 NCgl2112 Cytochrome aa3 oxidase, subunit IV Cgl2194 NCgl2114 Cytochrome bd oxidase, subunit I Cgl1150 NCgl1104 Cytochrome bd oxidase, subunit II Cgl1149 NCgl1103 Nitrate reductase, α-subunit, Mo Cgl1189 NCgl1142 cofactor-containing Nitrate reductase, β-subunit, iron-sulfur Cgl1188 NCgl1141 protein Nitrate reductase, δ-subunit, assembly Cgl1187 NCgl1140 factor Nitrate reductase, γ-subunit, cytochrome b Cgl1186 NCgl1139

Cg No.

Reference

cg1656

[42,46]

cg0445 cg0446 cg0447 cg2192 cg2891

[45]

cg1027

[5]

cg3227 cg1853 cg0129 cg1386 cg1387 cg2405

[48,49,68]

cg2404

[48,49,68]

cg2403

[48,49,68]

cg2780 cg2409 cg2406 cg2408 cg1301 cg1300 cg1344

[48,49,58] [48,49,58] [48,49,58] [49] [34] [34]

cg1343 cg1342 cg1341

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TABLE 13.1 (continued) Compilation of C. glutamicum Genes Encoding Respiratory Chain Components, F1F0-ATP Synthase, Menaquinone Biosynthesis, Heme Biosynthesis, and Cytochrome c Maturation Gene

atpI atpB atpE atpF atpH atpA atpG atpD atpC

menF menD

menC menE menB menA menG

hemA hemC hemN hemH hemD hemB hemE hemG hemL ccsX ccdA ccsB ccsA ctaB

Proven or Anticipated Function of the Gene Product

Function unknown a-Subunit of F0 part c-Subunit of F0 part b-Subunit of F0 part δ-Subunit of F1 part α-Subunit of F1 part γ-Subunit of F1 part β-Subunit of F1 part ε-Subunit of F1 part

Cgl No.

NCgl No.

Cg No.

NCgl1158 NCgl1159 NCgl1160 NCgl1161 NCgl1162 NCgl1163 NCgl1164 NCgl1165 NCgl1166

cg1361 cg1362 cg1363 cg1364 cg1365 cg1366 cg1367 cg1368 cg1369

Biosynthesis Cgl1292 NCgl1243 Cgl0467 NCgl0450

cg1462 cg0552

F1F0-ATP Synthase [64] Cgl1205 Cgl1206 Cgl1207 Cgl1208 Cgl1209 Cgl1210 Cgl1211 Cgl1212 Cgl1213

Menaquinone Isochorismate synthase 2-Oxoglutarate decarboxylase and 2succinyl-6-hydroxy-2,4cyclohexadiene-1-carboxylate synthase o-Succinylbenzoate synthase o-Succinylbenzoyl-CoA synthase 1,4-Dihydroxy-2-naphthoate synthase 1,4-Dihydroxy-2-naphthoate prenyltransferase Demethylmenaquinone methyltransferase

Cgl0466 Cgl0450 Cgl0463 Cgl0448

NCgl0449 NCgl0435 NCgl0446 NCgl0433

cg0551 cg0533 cg0548 cg0531

Cgl0471

NCgl0454

cg0556

Heme Biosynthesis and Cytochrome c Maturation Glutamyl-tRNA reductase Cgl0417 NCgl0402 Porphobilinogen deaminase Cgl0418 NCgl0403 Coproporphyrinogen III, dehydrogenase Cgl2292 NCgl2212 Ferrochelatase Cgl1537 NCgl1479 Uroporphyrinogen III synthase Cgl0429 NCgl0414 δ-Aminolevulinic acid dehydratase Cgl0431 NCgl0416 Uroporphyrinogen decarboxylase Cgl0435 NCgl0420 Protoporphyrinogen oxidase Cgl0436 NCgl0421 Glutamate-1-semialdehyde-2,1Cgl0437 NCgl0422 aminomutase Periplasmic thioredoxin Cgl0439 NCgl0424 Disulfide interchange protein Cgl0440 NCgl0425 Cytochrome c biogenesis protein B Cgl0441 NCgl0426 Cytochrome c biogenesis protein A Cgl0442 NCgl0427 Heme o synthase (protoheme IX Cgl1573 NCgl1511 farnesyltransferase)

cg0497 cg0498 cg2517 cg1734 cg0510 cg0512 cg0516 cg0517 cg0518 cg0520 cg0522 cg0523 cg0524 cg1773

Reference

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TABLE 13.1 (continued) Compilation of C. glutamicum Genes Encoding Respiratory Chain Components, F1F0-ATP Synthase, Menaquinone Biosynthesis, Heme Biosynthesis, and Cytochrome c Maturation Proven or Anticipated Function of the Gene Product

Gene ctaA ccdA2

glbO

Cgl No.

NCgl No.

Cg No.

Cgl1569

NCgl1508

cg1769

Cgl0018 Cgl0019

NCgl0017 NCgl0018

cg0025 cg0026

Others Cgl2448

NCgl2362

cg2689

Heme a synthase (heme o monooxygenase) Cytochrome c biogenesis, periplasmic protein-disulfide isomerase

Hemoglobin-like protein

NADH

Reference

NADH-DH II (ndh)

Succinate

Succinate DH (sdhABC)

L-Malate

Malate:MK OR (mqo)

Pyruvate

Pyruvate:MK OR (pqo)

D-Lactate

D-Lactate DH (dld )

L-Lactate

L-Lactate DH (lldD)

Glycerol-3phosphate

Glyc-3-P DH (glpD)

Proline

L-Proline DH (putA)

Cytochrome bc1-aa3 supercomplex (qcrCAB, ctaCDEF) Menaquinone (MQ-8, MQ-9)

O2

Cytochrome bd oxidase (cydAB)

O2

Nitrate reductase (narGHI )

NO3

? Acyl-CoA

ETF (etfAB)

FIGURE 13.1 Overview on the components constituting the respiratory chain of C. glutamicum. (Reprinted from [5] with permission.)

transfer of electrons from NADH to MQ in C. glutamicum is catalyzed by a type-II NADH dehydrogenase (NDH-II) encoded by the ndh gene [46]. This enzyme is not an integral membrane protein but associated with the inner side of the cytoplasmic membrane and is not coupled to ion translocation (EC 1.6.99.3). The ndh gene product consists of 467 amino acid residues (51.0 kDa) and shows 27% and 32% sequence identity to NDH-II of E. coli and B. subtilis, respectively. A C. glutamicum

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out +

2 H+

2e

2e−

2e−

Malate Oxaloacetate Pyruvate Acetate D-Lactate Pyruvate L-Lactate Pyruvate + 2H+ + CO2 + 2H+ + 2H+

2H+

bH bL

MK 2H+

[4Fe-4S] [2Fe-2S] FAD 2e−

b558

2

H2O

b595 2H

+

CydA

d

4H+ 2 MKH2 MK MKH2

Fumarate + 2H

bL bH 2H

+

−

NO2− + H2O

NO3 + 2H+

2H+ [2Fe-2S]c1

c

CuA

CuB

a QcrC

2O

CydB

MKH2 MK Membrane

1/

2e− +

CtaF CtaC

Succinate

2H+

Molybdopterin dinucleotide

a3

1/

2 O2

CtaE

2e−

Glycerol- Dihydroxy L-Proline Pyrroline-53-P aceton-P carboxylate + 2H+ + 2H+

out +

[3Fe-4S] 3[4Fe-4S]

NarH

PutA

2e−

bL

[3Fe-4S]

FAD

GlpD

FAD

bH

SdhB

2H+

MKH2 MKH 2 MK MK

SdhC

MKH2

SdhA

− in

MKH2 MK

QcrB QcrA

Membrane

2H+ NarI

out +

− in

2e−

NarG

NAD + H+

TPP

−

2H+ FMN

FAD

CtaD

+

2H+

FAD

Mqo

Ndh 2e

−

NADH

2H+

FAD

LldD

2 H+ FAD

MKH2 MK

MKH2 MK

Dld

− in

MKH2 MK

MKH2 MK

MKH2 MK

Pqo

Membrane

2H+ 2H+ H O 2

FIGURE 13.2 Schematic representation of subunit composition, cofactors, topology, electron flow, and involvement in the generation of a proton-motive force for the components of the C. glutamicum respiratory chain. Heme groups are indicated by black squares, copper ions by black circles; bH high potential heme b; bL, low potential heme b; TPP, thiamine pyrophosphate. Reprinted from [5] with permission.

mutant with a disrupted ndh gene (ndh::pEMndhint) has completely lost membranebound NADH dehydrogenase activity, indicating that NDH-II is solely responsible for this activity under the conditions tested. The viability of the ndh mutant is presumably due to the consecutive action of malate dehydrogenase (encoded by the mdh gene) and malate:quinone oxidoreductase (encoded by the mqo gene, see Section 13.2.3), which together catalyze electron transfer from NADH to MQ in the absence of NDH-II. Malate dehydrogenase reduces oxaloacetate with NADH to malate, which is the thermodynamically strongly favored direction, and malate is

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then reoxidized by malate:quinone oxidoreductase using MQ as electron acceptor [46]. This explanation is supported by the finding that a mqo ndh double mutant failed to grow under conditions that allowed growth of the ndh mutant. In addition, NADH dehydrogenase activity was reconstituted in vitro with membranes from the ndh mutant supplemented with malate and purified malate dehydrogenase [46]. Besides NADH, NADPH was slowly oxidized by membranes of the C. glutamicum wild-type, but not of the Δndhint mutant strain. This clearly indicates that NDH-II rather than a specific NADPH dehydrogenase [41] is responsible for this activity [46]. NADH dehydrogenase was purified 16-fold from C. glutamicum membranes to apparent homogeneity, resulting in a single polypeptide with an apparent mass of 55 kDa and the N-terminal sequence SVNPTRPEGGR, which is identical to the protein sequence deduced from ndh except for the lack of the N-terminal methionine residue [42]. Purified NADH dehydrogenase could be activated by FAD, but not by FMN, confirming that FAD is the cofactor of this enzyme. It had a specific NADH:UQ1 reductase activity of 46.5 U/mg and a specific NADPH oxidase activity of 4.5 U/mg. Whereas the NADH:UQ1 reductase activity was maximal at pH 6.5, the NADPH:UQ1 reductase activity was highest at pH 4.5 and negligible at pH 6.5, indicating that NADPH oxidation by NDH-II is virtually absent at physiological pH.

13.2.2 SUCCINATE DEHYDROGENASE Succinate dehydrogenase (SDH, EC 1.3.5.1) is part both of the citric acid cycle and of the aerobic respiratory chain (for reviews, see [19,35]). C. glutamicum SDH has not yet been characterized, but many properties can be deduced by a comparison of the genome-derived protein sequences with that of extensively characterized SDH enzymes, in particular those for which a crystal structure has been determined, i.e., E. coli fumarate reductase [24] and Wolinella succinogenes fumarate reductase [36]. C. glutamicum SDH consists of three subunits, SdhA, SdhB, and SdhC, which are encoded by the sdhCAB operon [80]. According to the functional classification [19], C. glutamicum SDH belongs to subclass 3, which includes those enzymes that catalyze the oxidation of succinate and the reduction of a low-potential quinone, in this case MQ. Succinate:MQ reductase from B. subtilis [20] or MQH2:fumarate reductase from W. succinogenes [33] are prominent representatives of this subclass. The flavoprotein subunit SdhA of C. glutamicum (625 residues) shows 32 to 34% sequence identity to SdhA of B. subtilis, FrdA of W. succinogenes, and FrdA of E. coli. The FAD prosthetic group is presumably linked to the histidine residue at position 50 of C. glutamicum SdhA and the active site histidine residue is located at position 248. The iron-sulfur-protein subunit SdhB of C. glutamicum (249 residues) has 29 to 30% sequence identity to SdhB of B. subtilis, FrdB of W. succinogenes, and FrdB of E. coli. C. glutamicum SdhB contains all 11 cysteine residues that are required to form the [2Fe-2S] cluster (Cys58, Cys63, Cys66, Cys84), the [4Fe-4S] cluster (Cys159, Cys162, Cys165, Cys223), and the [3Fe-4S] cluster (Cys169, Cys219, and Cys213). The membrane-anchored subunit SdhC (257 residues) shows only 25% sequence identity to SdhC of B. subtilis and FrdC of W. succinogenes [5], but shares important properties with the two homologs, i.e., the presence of five transmembrane helices and of four conserved histidine residues,

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Membrane

out +

C

His65

His173

His124

His221

− in N

FIGURE 13.3 Proposed transmembrane topology of the cytochrome b subunit (SdhC) of succinate:menaquinone oxidoreductase of C. glutamicum. The potential axial ligands of the low-potential heme iron (His65 and His173) and of the high-potential heme iron (His124 and His221) are indicated. (Reprinted from [5] with permission.)

which presumably serve as axial ligands of a high-potential heme group (heme bH) and a low-potential heme group (heme bL). In Figure 13.3, a model of the transmembrane topology of the C. glutamicum SdhC subunit is shown. Succinate oxidase activity of bacterial cells that use MQ (E0’ –73 mV) as the respiratory quinone, including C. glutamicum, is inhibited by uncouplers, whereas succinate oxidase activity from bacteria using UQ (E0’ +110 mV) as acceptor is not influenced [61]. It was therefore proposed that electron transport from succinate to MQ by succinate:MQ oxidoreductase involves a reversed electron transfer across the cytoplasmic membrane from the inner to the outer side via two heme b groups, which is driven by the proton or electrical potential. In support of this proposal, an electrochemical proton potential was formed when SDH of B. subtilis functioned as fumarate reductase [62]. In the model of electron flow through SDH of C. glutamicum (Figure 13.2), which was adapted from that of B. subtilis [20,35], it is evident that two protons are released in the cytoplasm during succinate oxidation and two protons are consumed at the outside for MQ reduction, corresponding to a net influx of two protons.

13.2.3 MALATE:QUINONE OXIDOREDUCTASE C. glutamicum possesses not only a cytoplasmic, NAD+-dependent malate dehydrogenase (MDH; EC 1.1.1.37), but also a membrane-associated malate:quinone oxidoreductase (MQO; EC 1.1.99.16), which oxidizes malate to oxaloacetate and transfers the reducing equivalents to MQ [45]. Owing to the different redox potentials of NAD+ and MQ, oxidation of malate by MDH is highly endergonic with a ΔG0’ of +28.6 kJ/mol, whereas oxidation by MQO is highly exergonic with a ΔG0’ of –18.9 kJ/mol. The mqo gene was isolated by complementing a C. glutamicum mutant obtained by treatment with 1-methyl-3-nitro-1-nitrosoguanidine that grew more slowly than the wild-type on all substrates and lacked detectable MQO activity [45]. The identity

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of the gene product (500 residues, 54.8 kDa) with MQO was confirmed by showing that the purified His-tagged protein possesses high 2,6-dichlorindophenol reductase activity. Whereas a mutant lacking the gene for the NAD+-dependent malate dehydrogenase (Δmdh) has no obvious growth phenotype, a Δmqo mutant is unable to grow in minimal medium [46]. This indicates that MQO, but not MDH, is essential for malate oxidation to oxaloacetate in C. glutamicum. Growth of the Δmqo mutant, but not of a Δmqo Δmdh double mutant, on minimal medium could be restored by the addition of nicotinamide (1 mg/L), which led to an increase of the intracellular NAD+ concentration from 0.22 to 1.53 mM and of the NADH concentration from 0.05 to 0.54 mM. However, the growth rate was reduced by 10% on glucose, by 40% on pyruvate, and by 75% on acetate. These data indicate that in the presence of elevated NAD+ levels MDH could partially complement the absence of MQO. Biochemical characterization of MQO [45] indicated that it is a peripheral membrane protein that can be easily detached. If cells are fractionated in the presence of Mg2+ and Ca2+, ~18% of the total MQO activity was associated with the membrane, whereas this value dropped to 6% if the chelators EDTA and EGTA were present [45]. The 2,6-dichlorindophenol reductase activity of solubilized MQO was strongly activated by the addition of certain quinones, e.g., 2-methyl-1,4-naphthoquinone (menadione, vitamin K3), UQ0, and UQ1, the latter being most efficient. Surprisingly, MQ4 (vitamin K2) did not stimulate the activity. Lipids and FAD stimulated the activity only after inactivation of MQO with the chaotropic salt KSCN, indicating that both the FAD prosthetic group and lipid are tightly bound to the enzyme in the solubilized state and only released upon addition of the thiocyanate ion. Partially purified MQO reduced UQ1 also in the absence of 2,6-dichlorindophenol, confirming that UQ1 is a substrate rather than a cofactor or otherwise stimulating factor.

13.2.4 PYRUVATE:QUINONE OXIDOREDUCTASE (PYRUVATE OXIDASE) Inspection of the genome sequence revealed that C. glutamicum contains a gene (pqo), whose deduced protein (579 amino acids, 62.0 kDa) shows 47% sequence identity to PoxB of E. coli. The E. coli enzyme catalyzes the oxidative decarboxylation of pyruvate to acetate and CO2 using a quinone as electron acceptor. It thus functions as pyruvate:quinone oxidoreductase (EC 1.2.2.2). The protein PoxB from E. coli is a peripheral membrane protein that contains FAD, thiamine pyrophosphate, and Mg2+. It is strongly activated by a variety of phospholipids that in E. coli increase the maximal velocity about 20-fold and lower the concentration of pyruvate needed for enzyme saturation about 10-fold [17]. The physiological function of Pqo in C. glutamicum is unknown.

13.2.5 LACTATE DEHYDROGENASES C. glutamicum can grow on D-lactate and L-lactate as the sole carbon and energy sources. An inspection of the genome sequence revealed two genes that most likely encode D-lactate dehydrogenase (dld) and L-lactate dehydrogenase (lldD). Both are

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presumably peripheral membrane proteins that use MQ as electron acceptor, in contrast to the soluble NAD+-dependent lactate dehydrogenase (EC 1.1.1.27; encoded by the ldhA gene), which is also present in C. glutamicum. Under standard conditions, lactate oxidation with NAD+ as acceptor is highly endergonic, whereas lactate oxidation with MQ is highly exergonic. The NAD+-dependent lactate dehydrogenase is presumably used for reoxidation of NADH under conditions where oxidation by the respiratory chain is limiting. Under these conditions, lactate is formed as a fermentation product. In contrast, the quinone-dependent lactate dehydrogenases Dld and LldD most likely serve to oxidize lactate when it is used as a carbon and energy source. The protein deduced from dld (571 residues, 63.7 kDa) shows 46% sequence identity to D-lactate dehydrogenase (EC 1.1.1.28) from E. coli and other bacteria. The crystal structure of the E. coli enzyme, which serves as a model for studying protein–lipid interactions, revealed that it is composed of three domains, an FADbinding domain, a cap domain, and a membrane-binding domain [15]. The residues involved in the noncovalent FAD binding are conserved in C. glutamicum Dld as well as the proposed active site residues (Ile-142 and Ser-144). The structure of the E. coli Dld protein indicated that association of the enzyme with the membrane is mediated by electrostatic interactions between an electropositive surface composed of several arginine and lysine residues in the membrane-binding domain and the electronegative phospholipid head groups of the membrane. Several of these basic residues are also present in C. glutamicum D-lactate dehydrogenase, suggesting a similar association mechanism. D-Lactate dehydrogenase was solubilized from membranes of the C. glutamicum strain ATCC14310-DL4 with N,N-dimethyldodecylamine N-oxide (LDAO) and then purified 775-fold to a specific activity of 310 U/mg protein. After SDS-PAGE, one dominant protein with an apparent mass of 66 kDa was observed, which corresponds to the predicted mass of the dld gene product (63.7 kDa). Loss of enzyme activity during purification could be reversed by addition of FAD, but not FMN. In addition, the purified enzyme was shown to contain FAD, which thus was unequivocally identified as the prosthetic group of D-lactate dehydrogenase from C. glutamicum [5]. The protein deduced from lldD is composed of 420 residues and shows 38% sequence identity to LldD of E. coli and 32% identity to L-lactate ferricytochrome c reductase (cytochrome b2) from Saccharomyces cerevisiae, which in addition to the bacterial enzymes contains an N-terminal protoheme IX binding domain. Biochemical data on bacterial L-lactate dehydrogenases (EC 1.1.2.3) and the crystal structure of the yeast enzyme [82] have identified FMN as prosthetic group.

13.2.6 GLYCEROL-3-PHOSPHATE DEHYDROGENASE Glycerol-3-phosphate dehydrogenase catalyzes the oxidation of glycerol-3-phosphate to dihydroxyacetone phosphate and reduces quinone in the cytoplasmic membrane. It functions to salvage glycerol and glycerol phosphates generated from the breakdown of phospholipids and triacylglycerol. C. glutamicum possesses a gene (glpD) encoding a protein of 574 residues that shows 34% sequence identity to the aerobic glycerol-3-phosphate dehydrogenase (EC 1.1.99.5) of E. coli (glpD gene

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315

product). The E. coli GlpD protein (501 residues, 57 kDa) was purified and shown to be a homodimer with noncovalently bound FAD as a cofactor [63]. The primary sequences of E. coli GlpD and C. glutamicum GlpD do not contain putative transmembrane helices and therefore these proteins are presumably peripheral membrane proteins.

13.2.7 PROLINE DEHYDROGENASE C. glutamicum can grow on L-proline as sole carbon and nitrogen source, albeit slowly (our own unpublished results). The catabolism of proline presumably involves the protein encoded by the putA gene (1,152 amino acids), which shows 30% sequence identity to the E. coli proline dehydrogenase PutA (EC 1.5.99.8 and EC 1.5.1.12). This is a bifunctional enzyme that catalyzes the oxidation of L-proline to Δ1-pyrroline-5-carboxylate with a membrane-bound quinone as electron acceptor and the further oxidation of Δ1-pyrroline-5-carboxylate to L-glutamate with NAD+ as electron acceptor [40]. The E. coli proline dehydrogenase (P09546; 1,320 amino acids) enabling the use of proline as carbon, nitrogen, and electron source, is a homodimer (2 x 144 kDa) and contains noncovalently bound FAD as a cofactor [6]. When the proline concentration is high, PutA is enzymatically active as a dehydrogenase. In the absence of excess proline, this protein accumulates in the cytoplasm and binds to the put operon control region, thereby repressing put operon expression. The binding of proline to PutA results in increased hydrophobicity of the protein and an increased affinity for membrane-binding sites [14]. In contrast to the E. coli proline dehydrogenase, PutA of C. glutamicum probably does not function as transcriptional repressor, because a sequence alignment (not shown) reveals that domain II of E. coli PutA, which has been implicated in DNA-binding [38], is largely absent in the C. glutamicum protein.

13.2.8 ELECTRON-TRANSFERRING FLAVOPROTEIN The C. glutamicum genome contains two genes whose protein products show clear homology to the α- and β-subunits of the electron-transferring flavoprotein ETF (etfA and etfB, respectively). It has been shown for other bacteria that the reducing equivalents formed during the oxidation of fatty acyl-CoA to trans-Δ2-enoyl-CoA by acyl-CoA dehydrogenases are transferred to the FAD prosthetic group of the α-subunit of ETF. The reduced FADH2 is subsequently reoxidized by ETF:quinone oxidoreductase (EC 1.5.5.1), which transfers the electrons to the membrane-bound quinone pool. This enzyme contains a [4Fe-4S] cluster and FAD as prosthetic groups [3,55,66]. Remarkably, there is no obvious homolog of ETF:ubiquinone oxidoreductase in C. glutamicum. The same situation appears to be true for all other Grampositive bacteria of known genome sequence, indicating that the oxidation of reduced ETF must be catalyzed by a different enzyme in the Gram-positives. This might be related to the much lower redox potential of MQ compared with UQ. The standard redox potential of FAD varies between 0 mV in the ETF of human [59] and pig [21] and +200 mV in the ETF from Methylophilus methylotrophicus [8]. The redox potentials of FAD and the iron-sulfur cluster in mammalian ETF:UQ oxidoreductase

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are around 0 mV and +50 mV, respectively [51]. Thus, electron transfer from ETF to MQ would be energetically unfavorable under standard conditions and eventually require the electrochemical proton potential as driving force, similar to the case of succinate dehydrogenase described above.

13.3 ELECTRON TRANSFER FROM MENAQUINOL TO OXYGEN Reduced-minus-oxidized difference spectra obtained with intact cells, cell extracts, and membranes of C. glutamicum revealed peaks at 600 nm, 560 nm, and 552 nm (Figure 13.4), indicating the presence of a-, b- and c-type cytochromes, respectively [48,69,70]. In the CO-reduced-minus-reduced difference spectrum, a peak at 427 nm and a trough at 443 nm were found, typical for a cytochrome aa3-type terminal oxidase [74]. Isolated membranes exhibited oxidase activity with the artificial electron donor tetramethyl-p-phenylendiamine (TMPD), which was almost completely inhibited by 0.1 mM cyanide [41]. In contrast, oxidation of other substrates, such as NADH, succinate, lactate, and NADPH, was less sensitive to cyanide. With the TMPD oxidase system, a single Km value for oxygen of 56 μM was determined, whereas with the NADH oxidase system two Km values of 18 μM and 48 μM were obtained. The H+/O ratios with endogenous substrates was 5.2 for cells from the mid-log phase and decreased to 3.0 after inhibition with 2 mM cyanide. In addition, for cells harvested in the stationary phase, a decreased H+/O ratio of 3.9 was determined [41]. When C. glutamicum (formerly Brevibacterium thiogenitalis) was grown in copper-free medium, cytochromes a and c disappeared and instead cytochrome d was detected by its absorption at 630 nm in redox difference spectra. NADH oxidase activity of the cell-free extracts from copper-deficient cells was

Absorbance

ΔA = 0.02

Wt

ΔctaD Δqcr

500

550

600

650

Wavelength (nm)

FIGURE 13.4 Redox difference spectra (dithionite-reduced minus ferricyanide-oxidized) of membranes (30 mg protein/mL) isolated from C. glutamicum wild-type (Wt), the ctaD deletion mutant (ΔctaD), and the qcr deletion mutant (Δqcr) grown aerobically in BHI medium with 2% (w/v) glucose. (Reprinted from [48] with permission.)

Respiratory Energy Metabolism

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inhibited only 20% by 1 mM cyanide in contrast to 63% inhibition with coppersufficient cells [70]. From these data, the presence of at least two terminal oxidases could be inferred: (i) a cytochrome aa3 terminal oxidase, which receives electrons from a bc1 complex and which is inhibited by micromolar concentrations of cyanide, and (ii) an alternative menaquinol oxidase of the bd-type that is more resistant to cyanide and that has a higher affinity for oxygen but a lower energetic efficiency with respect to proton translocation.

13.3.1 CYTOCHROME bc1 COMPLEX The C. glutamicum cytochrome bc1 complex is encoded by the qcrCAB genes (Figure 13.5) for cytochrome c1 (30 kDa), Rieske iron-sulfur protein (45 kDa), and cytochrome b (60 kDa), respectively [48,68]. The deduced protein sequences show a number of differences from classical representatives of the bc1 complex, such as an extension of about 120 amino acids at the C-terminus of cytochrome b (QcrB) and the presence of three putative transmembrane helices in the N-terminal region of the Rieske iron-sulfur protein (QcrA) rather than only one. Most remarkably, cytochrome c1 (QcrC) was found to have two CXXCH motifs for covalent heme attachment, indicating that it is a diheme c-type cytochrome. A qcrCAB deletion mutant of C. glutamicum shows a severe growth defect (Figure 13.6), indicating that the bc1 branch of the respiratory chain is of primary importance and cannot be easily substituted by alternative menaquinol oxidation pathways [48]. Staining of proteins separated by SDS-PAGE for covalently bound heme indicated that there is only a single c-type cytochrome with an apparent mass of 31 kDa present in C. glutamicum wild-type [48,68]. This protein was missing in the ΔqcrCAB mutant, confirming that it represents cytochrome c1. Reduced-minusoxidized difference spectra of the mutant (Figure 13.4) also revealed the absence of the 552-nm cytochrome c-peak, whereas a- and b-type cytochromes were still present [48]. Cytochrome c1 was purified from wild-type membranes by a three-step chromatographic protocol. The final preparation had a heme content of 62 nmol/mg protein, confirming that QcrC is a diheme cytochrome c1 [68]. Interestingly, neither

ctaD SU I 1 kb

ctaC

ctaF

SU II SU IV

Cytochrome aa3 oxidase

ctaE

qcrC

SU III Cyt. c1

qcrA FeS

qcrB Cyt. b

Cytochrome bc1 complex

FIGURE 13.5 Physical map of the C. glutamicum genome region comprising ctaC, ctaF, and ctaE (encoding subunit II, IV, and III of cytochrome aa3 oxidase, respectively), and qcrCAB (encoding cytochrome c1, Rieske iron-sulfur protein, and cytochrome b of the cytochrome bc1 complex, respectively). The ctaD gene encoding subunit I of cytochrome aa3 oxidase is located separately 345 kbp upstream of ctaC.

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FIGURE 13.6 Growth of the wild-type strain C. glutamicum ATCC13032 and of mutants lacking either the ctaD gene for subunit I of cytochrome aa3 oxidase, or the ctaF gene for subunit IV of cytochrome aa3 oxidase, or the cydAB genes for cytochrome bd oxidase, or the qcrCAB genes for the cytochrome bc1 complex. The bacteria were grown for 2 d at 30°C on brain-heart infusion agar supplemented with 0.5 M sorbitol.

QcrB nor QcrA were copurified with QcrC, indicating weak interactions between these proteins. The same conclusion was obtained from the purification of streptagged QcrB from a C. glutamicum mutant lacking cytochrome aa3 oxidase, which led to the isolation of a complex consisting mainly of QcrB and QcrA, whereas QcrC was lacking (Figure 13.7) [49]. Site-directed mutagenesis of QcrC revealed that covalent attachment of both heme groups to apo-cytochrome c1 is essential for assembly and/or stability of the whole bc1 complex. If either the N-terminal or the C-terminal CXXCH heme binding motif was changed to SXXSH, a similar growth defect as in the ΔqcrCAB strain was observed. In membranes of these mutant strains, no c-type cytochrome was detectable and the QcrB level was clearly reduced [49].

13.3.2 CYTOCHROME aa3 OXIDASE C. glutamicum cytochrome aa3 oxidase is composed of four subunits encoded by the genes ctaD (subunit I), ctaC (subunit II), ctaE (subunit III), and ctaF (subunit IV). The ctaD gene is located separately from ctaC, ctaE, and ctaF and encodes a protein of 65 kDa whose sequence shows all features typical for subunit I of hemecopper terminal oxidases [48]. The other three genes are located immediately upstream of qcrCAB (Figure 13.5). The CtaC protein of C. glutamicum (40 kDa) differs from classical subunit II representatives by an insertion of about 30 amino acids in the substrate binding domain, which is proposed to play a role in the interaction with cytochrome c1 [58]. Subunit III (CtaE, 22 kDa) of C. glutamicum

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FIGURE 13.7 Purification of cytochrome bc1 and cytochrome aa3 single complexes and supercomplexes from C. glutamicum by affinity chromatography on StrepTactin sepharose. The proteins were denatured, separated on an 8–16% Tris⋅HCl gradient SDS gel (Biorad) and stained with Coomassie blue. Lane 1, cytochrome aa3 oxidase (7.2 μg protein) purified from a ΔqcrCAB strain transformed with a plasmid encoding subunit I of cytochrome aa3 oxidase with a C-terminal StrepTag II. Lane 2, cytochrome bc1-aa3 supercomplex (4.7 μg) purified from a ΔctaD strain transformed with a plasmid encoding subunit I of cytochrome aa3 oxidase modified with a C-terminal StrepTag II. Lane 3, cytochrome bc1-aa3 supercomplex (5.7 μg) purified from the ΔqcrCAB strain transformed with a plasmid encoding subunit III of cytochrome aa3 oxidase and the three subunits of the bc1-complex; cytochrome b contained a C-terminal StrepTag II. Lane 4, cytochrome ‚bc1’complex (4.3 μg) purified from the ΔctaD strain. (Adapted from [49] with permission.)

lacks the N-terminal part of the homologous protein from P. denitrificans comprising two transmembrane helices [48]. Subunit IV (CtaF, 16 kDa) was identified only recently [49]. It is unique to the actinomycetales (Figure 13.8) and does not show significant homology to subunit IV representatives of other heme-copper oxidases, such as CtaF [60] (caa3-type cytochrome oxidase) and QoxD [76] (aa3-type quinol oxidase) from B. subtilis or CtaH from P. denitrificans [81]. C. glutamicum mutants lacking either ctaD or ctaF show severe growth defects (Figure 13.6) [48,49]. Reduced-minus-oxidized difference spectra revealed that both mutants lacked the 600-nm cytochrome aa3 peak and that the 552-nm cytochrome c1 peak was much smaller than in the wild-type. By contrast, the 600-nm peak was

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FIGURE 13.8 Sequence alignment of CtaF proteins from different species of actinomycetes. Putative transmembrane helices are indicated by lines and numbered. Amino acids identical in at least five sequences are shaded in black, conservative exchanges in gray. The bacteria have been abbreviated as follows: Cgl, C. glutamicum (NCgl2114); Cdi, C. diphtheriae (NC_002935); Mtu, Mycobacterium tuberculosis (Rv2199c); Mbo, M. bovis (NC_002945); Mle, M. leprae (ML0876); Sco, Streptomcyces coelicolor (NP_626410); Tfu, Thermobifida fusca (Tfus_p_278). (Reprinted from [49] with permission.)

not changed in the ΔqcrCAB strain compared with the wild-type (Figure 13.4). This indicates that cytochrome aa3 oxidase is important for the stability of cytochrome c1 but not vice versa. Purification of cytochrome aa3 oxidase from membranes of C. glutamicum wildtype by conventional chromatography resulted in a three-subunit complex comprising CtaD, CtaC, and CtaE [58]. The heme of subunit I is presumably heme aS, which contains a geranylgeranyl side chain (C20H33-) instead of the farnesyl group (C15H25–) of heme a. In agreement with the presence of a lipoprotein signal sequence in subunit II (CtaC), Cys-29 was identified as the N-terminal residue of the mature protein and its mass was 3 kDa lower than predicted from the complete open reading frame. The purified cytochrome aa3 complex catalyzed the oxidation of TMPD, horse-heart cytochrome c, and yeast cytochrome c with turnover numbers of 0.61, 0.23, and 2.54 s–1 per cytochrome aa3, respectively [58]. When cytochrome aa3 oxidase of C. glutamicum was purified by StrepTactin affinity chromatography, a four-subunit complex was isolated containing CtaF in addition to CtaD, CtaC, and CtaE (Figure 13.7) [49]. For this purpose, the ΔqcrCAB strain transformed with a plasmid encoding a strep-tagged CtaD variant was used. The turnover numbers of the isolated four-subunit complex with TMPD, bovine heart cytochrome c, and yeast cytochrome c were 1.14, 1.18, and 0.93 s–1 per cytochrome aa3, respectively. These values are in the same range as those reported for the three-subunit complex, indicating that the fourth subunit (CtaF) is most likely not required for catalytic activity. However, the strong growth defect of the ΔctaF mutant, which is similar to that of the ΔctaD mutant, clearly showed that CtaF is essential for the formation of an active cytochrome aa3 oxidase and therefore presumably required for its assembly and/or stability [49].

Respiratory Energy Metabolism

13.3.3 IDENTIFICATION

OF A

321

CYTOCHROME bc1-aa3 SUPERCOMPLEX

Since cytochrome c1 is most likely the only c-type cytochrome in C. glutamicum, electron transfer from the bc1 complex to cytochrome aa3 oxidase either involves a different, yet unknown electron carrier or a direct transfer from cytochrome c1 to the CuA center in subunit II of cytochrome aa3 oxidase. The presence of two c-type hemes in cytochrome c1 suggested that the second heme group takes over the function of a separate cytochrome c and supported a direct electron transfer. Such a mechanism would require intimate contact between the bc1 complex and cytochrome aa3 oxidase and therefore the formation of a bc1-aa3 supercomplex was suggested [48]. While purification of cytochrome c1 [68] and of cytochrome aa3 oxidase [58] by classical chromatographic techniques gave no hints on the existence of such a complex, the use of StrepTactin affinity chromatography allowed the isolation of the supercomplex [49]. Purification of strep-tagged cytochrome b (QcrB) led to a preparation containing the three subunits of the bc1 complex and the four subunits of cytochrome aa3 oxidase. Vice versa, purification of strep-tagged subunit I (CtaD) resulted in a preparation containing not only the four subunits of cytochrome aa3 oxidase, but also the three subunits of the bc1 complex (Figure 13.7) [49]. The isolated bc1-aa3-supercomplexes had quinol oxidase activity of 1.5 to 1.7 U/mg protein with 2,3-dimethylnaphthoquinol (DMNH2) as substrate, indicating functional electron transfer between cytochrome c1 and the CuA center of subunit II (CtaC). Four additional proteins were copurified with the known subunits of the bc1 complex and cytochrome aa3 oxidase (Figure 13.7), three of which could be identified by peptide mass fingerprinting. Since deletion of the corresponding genes did not alter the growth behavior of the mutant strains compared with the wild-type, the copurified proteins are neither required for the assembly of the bc1-aa3 supercomplex nor for its catalytic activity [49].

13.3.4 CYTOCHROME bd MENAQUINOL OXIDASE The early studies by Sugiyama et al. [69,70] had shown that under copper deficiency a d-type cytochrome appears in membranes of C. glutamicum. This d cytochrome is part of the menaquinol oxidase cytochrome bd (Figure 13.2), which is encoded by the genes cydA and cydB [34]. Subunit I of C. glutamicum (= CydA) and other high-G+C Gram–positive bacteria lack a significant part of the Q loop present in the E. coli CydA protein. This is typical for the cytochrome bd subfamily found in low-G+C Gram-positive bacteria, cyanobacteria, chlamydia, archaea, and some proteobacteria. The C. glutamicum cydAB genes form a cluster with the cydCD genes encoding an ABC transporter that might function as a cysteine exporter [53]. Cytochrome bd oxidase was isolated from membranes of stationary phase cells of C. glutamicum ATCC14067 (formerly Brevibacterium flavum) cultivated aerobically in a copper-deficient medium [34]. The purified enzyme consists of two subunits of 56.4 and 41.5 kDa, similar to the homologous enzymes of other bacteria [25]. The redox difference spectrum revealed two major α-band peaks at 627 and 560 nm and a minor peak at 595 nm, corresponding to cytochrome d, low-spin heme

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b and high-spin heme b, respectively. The enzyme catalyzed the oxidation of naphthoquinols, with activities increasing in the order menadiol, MQ3-H2, MQ2-H2, DMNH2, and MQ1-H2. Preincubation of the enzyme with menaquinones stimulated the quinol oxidase activity up to fivefold. The efficiency of the various quinones in activating the quinol oxidase was clearly different from the substrate specificity and increased in the order MQ1, MQ3, and MQ2. Almost no activation was observed with DMN and menadione. Therefore, the presence of an additional quinone-binding site besides the active site for substrate oxidation was suggested. The quinol oxidase activity with DMNH2 was inhibited by cyanide with a Ki value of 5.3 mM. In many bacteria, e.g., Mycobacterium smegmatis [27], cytochrome bd oxidase is induced under microaerobic conditions because the high oxygen affinity of this enzyme allows respiration even under very low oxygen tensions. A similar type of regulation might also occur in C. glutamicum but has not yet been demonstrated. Under aerobic conditions, a C. glutamicum ΔcydAB mutant has no obvious growth defects (Figure 13.6), confirming that under these conditions the bc1-aa3 branch of the respiratory chain is of prime importance.

13.3.5 ALTERNATIVE OXIDASE ACTIVITIES Trutko et al. [74] and Matsushita et al. [41] reported on cyanide-resistant respiration by C. glutamicum cells that did not contain cytochrome d in redox difference spectra. Also Kusumoto et al. [34] mentioned a third oxidase activity besides cytochrome aa3 and cytochrome bd that was insensitive to cyanide up to 30 mM. Inspection of the genome sequence, however, indicates that C. glutamicum does not contain an additional terminal oxidase besides cytochrome aa3 and cytochrome bd. Although the presence of a new type of terminal oxidase cannot be excluded per se, it seems more likely that the cyanide-resistant oxidase activity mentioned above is caused by reactions in which molecular oxygen chemically oxidizes redox enzymes, forming superoxide [65]. Several types of such reactions have been described, i.e., the autoxidation of reduced flavoenzymes, such as succinate dehydrogenase [43] and the autoxidation of semiquinone radicals, i.e., the semiquinone bound at the Qp site of the cytochrome bc1 complex [1,22,75].

13.4 ELECTRON TRANSFER FROM MENAQUINOL TO NITRATE C. glutamicum is able to reduce nitrate to nitrite and this property is routinely used in taxonomical tests [39]. The nitrate reductase responsible for this reduction is encoded by the narKGHJI gene cluster, which has not yet been analyzed experimentally, but has high similarity to the narKGHJI operon of E. coli. In E. coli, NarK presumably functions as a nitrate/nitrite transporter [10] and NarG (α-subunit), NarH (β-subunit), and NarI (γ-subunit) constitute the respiratory nitrate reductase A (for review, see [4,54]). Subunit NarG is a molybdenum cofactor-containing subunit catalyzing nitrate reduction to nitrite, NarH is an electron transfer subunit that binds four iron-sulfur clusters, and NarI is an membrane-integral quinol dehydrogenase subunit with two heme b groups (Figure 13.2). NarJ (δ-subunit) is not part of the

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nitrate reductase holo-enzyme but is required for the insertion of the molybdenum cofactor into NarG in E. coli. Subunits NarG and NarH of C. glutamicum show 46 and 53% sequence identity to the corresponding E. coli proteins, respectively. Three cysteine residues in the N-terminal part of NarG, which are essential for activity as well as nine cysteine residues in NarH that coordinate three [4Fe-4S] clusters and one [3Fe-4S] cluster, are conserved in C. glutamicum. Subunit NarI of C. glutamicum has only 28% sequence identity to E. coli NarI, but the functionally important residues His56/His205 and His66/His187 of E. coli NarI, which ligate heme bH and heme bL, respectively, are present. In analogy to its E. coli counterpart, nitrate reduction by the NarGHI complex of C. glutamicum might generate an electrochemical proton gradient because quinol oxidation occurs at the outer side of the cytoplasmic membrane and nitrate reduction in the cytoplasm (see Figure 13.2). However, in contrast to E. coli, there are no reports on anaerobic growth of C. glutamicum by nitrate respiration, which might be due to the inability to reduce the toxic nitrite. In general, nitrite can either be reduced via nitric oxide (NO) and nitrous oxide (N2O) to dinitrogen (N2) in the process of denitrification or converted directly to ammonium (NH4+) via a sixelectron reduction. Neither genes required for denitrification nor genes encoding dissimilatory nitrite reductases were detected in C. glutamicum. One gene of C. glutamicum (Cgl2817) was annotated to encode a cytoplasmic ferredoxin-dependent nitrite reductase (EC 1.7.7.1), but it is clustered with those for phosphoadenosine phosphosulfate reductase (EC 1.8.99.4) and both subunits of sulfate adenylyltransferase (EC 2.7.7.4) and therefore probably encodes a sulfite reductase (EC 1.8.7.1). Sulfite reductase and nitrite reductase belong to the same family of sirohemecontaining proteins and both can catalyze the reduction of sulfite and nitrite, but with largely different specificity [13]. The physiological role of the dissimilatory nitrate reductase in C. glutamicum is unclear. In mycobacteria, nitrate reductase was induced in the transition to the nonreplicating state, and a mutant of M. bovis BCG lacking nitrate reductase showed a decreased persistence in some tissues of immunocompetent mice [16,77,79]. Since hypoxia is considered to be a major factor for inducing nonreplicating persistence [78], these results support an involvement of nitrate reductase in the adaptation to anoxic conditions in M. bovis.

13.5 HEME BIOSYNTHESIS AND CYTOCHROME C MATURATION Although no experimental data are available for C. glutamicum concerning heme biosynthesis and cytochrome c maturation, the corresponding pathways [50] can be deduced by genome analysis, at least partially [5]. δ-Aminolevulinic acid, the precursor of porphyrins, is synthesized via the C5 pathway from glutamate by glutamyl-tRNA synthetase (gltX), glutamyl-tRNA reductase (hemA), and glutamate-1-semialdehyde aminomutase (hemL). The conversion of protoheme IX (heme b) into heme a involves protoheme IX farnesyl transferase (ctaB) [56,57] and heme o monooxygenase (ctaA) [7]. Formation of holo-cytochrome c1 requires the covalent attachment

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1 hemB 2 3

4

hemE hemG

hemL

5 ccsX ccdA ccsB

ccsA 1 kb

FIGURE 13.9 Physical map of the C. glutamicum genome region comprising heme biosynthesis genes and cytochrome c maturation genes. The genes code for following proteins: hemD, uroporphyrinogen III synthase (EC 4.2.1.75, Cgl0429); hemB, δ-aminolevulinic acid dehydratase (EC 4.2.1.24, Cgl0431); hemE, uroporphyrinogen decarboxylase (EC 4.1.1.37, Cgl0435); hemG, protoporphyrinogen oxidase (EC 1.3.3.4, Cgl0436); hemL, glutamate-1semialdehyde aminotransferase (EC 5.4.3.8, Cgl0437); ccsX, periplasmic thioredoxin (Cgl0439), ccdA, disulfide interchange protein (Cgl0440); ccsB, cytochrome c biogenesis protein B (Cgl0441); ccsA, cytochrome c biogenesis protein A (Cgl0442); 1–3, hypothetical proteins (Cgl0430, Cgl0432, Cgl0433); 4, putative cation transporting P-type ATPase (Cgl0434); 5, protein with homology to phosphoglyceratmutase/fructose-2,6-bisphosphatase.

of two protoheme IX molecules to the two CXXCH motifs of the apo-protein [32,72]. This reaction presumably takes place on the outer side of the cytoplasmic membrane and involves transport of apo-cytochrome c1 via the Sec protein secretion system and export of protoheme IX via a still unknown transport system. After reduction of the cysteine residues of the CXXCH motifs by a thioreductase, the thioether bonds with protoheme IX are formed. In C. glutamicum, four putative genes for cytochrome c biogenesis, i.e., ccsX, ccdA, ccsB, and ccsA (names adapted from [2]), are clustered with genes involved in protoheme IX biosynthesis (Figure 13.9). Protein CcsX (24% sequence identity to ResA of B. subtilis) possesses a CXXC motif typical for members of the thioredoxin family and contains a lipoprotein signal sequence, indicating a periplasmic location of the mature protein. Protein CcdA (33% sequence identity to CcdA of B. subtilis) is an integral membrane protein with homology to the thiol:disulfide exchange protein DipZ from E. coli whose function is to transfer electrons across the membrane that are used to reduce disulfide bonds. Together, CcsX and CcdA might be responsible for the thiol-mediated reduction of the cysteines in apo-cytochrome c1. The CcsA and CcsB proteins (25% sequence identity to ResC and ResB of B. subtilis, respectively [37,71]) most likely function in heme delivery/ligation to apo-cytochrome c [5]. Besides the four genes described above, the C. glutamicum genome contains a gene (Cgl0018) whose protein product shows 24% sequence identity to the CcdA protein. Interestingly, this gene is located upstream of a gene encoding a putative periplasmic protein-disulfide isomerase (Cgl0019). These two genes might also be involved in cytochrome c maturation.

13.6 IMPACT OF F1F0-ATP SYNTHASE ON METABOLISM The electrochemical proton potential generated by the respiratory chain is used to drive the phosphorylation of ADP by F1F0-ATP synthase. The C. glutamicum genes

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encoding the eight subunits of this enzyme are organized as in most other bacteria in the atpIBEFHAGDC operon, which includes the atpI gene of unknown function. The promoter-proximal genes atpB, atpE, and atpF encode the three components of the membrane-integral F0 part, i.e., the a-, c-, and b-subunits, respectively. The promoter-distal genes atpH, atpA, atpG, atpD, and atpC encode the five components of the peripheral F1 part, i.e., the δ-, α-, γ-, β-, and ε-subunits, respectively. The importance of F1F0-ATP synthase for the metabolism of C. glutamicum was recently demonstrated by the analysis of the mutant strain F172-8, whose ATPase activity was decreased to 25% of the wild-type level owing to a point mutation in the γ-subunit (Ser-273-Pro) [64]. This strain was almost unable to produce glutamate, the glucose consumption per cell was increased by 70%, the respiration rate was doubled, and significant amounts of the pyruvate-derived metabolites lactate, L-alanine and L-valine, were formed. A spontaneous revertant of strain 172-8, named R2-1, in which Pro273 of the γ-subunit was changed to leucine, had regained 70% of the ATPase activity of the wild-type. The properties of this revertant were very similar to that of the wild-type, indicating that the phenotype of strain F172-8 was in fact due to the defect in F1F0-ATP synthase activity. This defect might cause an ATP shortage, which in turn could trigger an increased glucose catabolism. As a consequence, the rate of NADH formation would increase, leading to an enhanced respiration rate. By an unknown mechanism, the catabolism of pyruvate via the pyruvate dehydrogenase complex and the citric acid cycle seems to be inhibited, leading to the absence of glutamate synthesis and to the formation of lactate, alanine, and valine. Obviously, regular metabolism of C. glutamicum requires an intact F1F0-ATP synthase, and additional studies will be necessary to elucidate the mechanism by which defects in this enzyme affect metabolism.

13.7 INFLUENCE OF RESPIRATORY CHAIN COMPOSITION ON THE ATP YIELD Present knowledge indicates that the aerobic respiratory chain of C. glutamicum involves three enyzmes that couple electron transfer to the generation of an electrochemical proton gradient across the cytoplasmic membrane, i.e., the cytochrome bc1 complex, cytochrome aa3 oxidase, and cytochrome bd oxidase. In analogy to wellstudied systems, the number of protons formally transported across the membrane per electron (H+/e–) is three for the cytochrome bc1-aa3 supercomplex and one for cytochrome bd oxidase [47]. Synthesis of one ATP molecule by F1F0-ATP synthase requires transport of three to four protons through F0 from the outside to the cytoplasm. Using these values, transfer of two electrons from NADH to O2 via the bc1-aa3 branch would allow the formation of 1.5 to 2 ATP, whereas the bd branch (or nitrate reductase and nitrate as terminal electron acceptor) would yield only 0.5 to 0.7 ATP. The same values are obtained if the electrons are transferred to MQ by malate:quinone oxidoreductase, D-lactate dehydrogenase, L-lactate dehydrogenase, glycerol-3-phosphate dehydrogenase, or L-proline dehydrogenase rather than by NADH dehydrogenase, since all these enzymes are peripheral membrane proteins

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attached to the inner surface of the cytoplasmic membrane and consequently, the protons for the reduction of MQ stem from the cytoplasm (Figure 13.2). The situation is different for succinate dehydrogenase, since electron transfer by this enzyme involves a membrane-integral subunit (SdhC) with two heme b groups and the reduction of MQ probably occurs at the outer surface of the cytoplasmic membrane, with protons coming from the outside rather than from the inside (Figure 13.2). Consequently, two-electron transfer from succinate to O2 via the bc1-aa3 branch would result in the net extrusion of four protons, but no net proton extrusion would occur if the bd branch is used. Taking the values mentioned above and a stoichiometry of 3 H+/ATP for F1F0ATP synthase, the complete aerobic oxidation of glucose via glycolysis, the citric acid cycle, and the bc1-aa3 branch of the respiratory chain would result in the formation of 4 ATP by substrate level phosphorylation and of 22.7 ATP by oxidative phosphorylation. Use of the cytochrome bd branch would result in the synthesis of only 6.7 ATP by oxidative phosphorylation. The complete aerobic oxidation of acetate by the citric acid cycle (after activation by the acetate kinase/phosphotransacetylase pathway [18]) would yield 7.3 ATP if the bc1-aa3 branch is used and only 2 ATP if the bd branch is used. These examples illustrate the impact of respiratory chain composition on the efficiency of oxidative phosphorylation and hence on the entire energetics of the cell. The rough calculations made above for C. glutamicum are supported by studies with E. coli [9] and B. subtilis [67].

13.8 BIOTECHNOLOGICAL ASPECTS As shown by the effects of copper [28,70], cyanide [74], or a defective F1F0-ATPsynthase [64], interferences with the system of oxidative phosphorylation can have a strong influence on metabolism in general and on amino acid production in particular. Further support for this finding comes from a number of patent applications reporting that disruption or overexpression of genes relevant to respiration can influence amino acid production. For example, a mutant with a disrupted sdhA gene (flavoprotein of succinate dehydrogenase) was reported to show increased glutamate formation [52]. Another example concerns a gene not yet mentioned in this review, i.e., the glbO gene encoding a hemoglobin-like protein (131 amino acids, Cgl2448), which presumably is able to bind oxygen. Overexpression of this gene was found to have a positive influence on L-lysine formation [44]. The molecular basis for this effect is unknown at present, however, since the discovery of Khosla and Bailey [31] that heterologous expression of a bacterial hemoglobin improves the growth properties of E. coli, numerous studies were performed on this topic. Another example for the influence of the respiratory chain on L-lysine formation is provided by Toma et al. [73]. They showed that the best lysine yield obtained with a C. glutamicum ATCC 1407 derivative (Brevibacterium flavum LD22) grown in chemostat culture at varying growth rates and at varying oxygen tensions was always observed when cyanide-resistant respiration was minimal, i.e., when the efficiency of oxidative phosphorylation was maximal.

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13.9 CONCLUDING REMARKS The recent experimental studies in several laboratories and the information obtained by the genome sequence have established a good picture of the aerobic respiratory chain of C. glutamicum. It involves several dehydrogenases, menaquinone, a cytochrome bc1-aa3 supercomplex, and cytochrome bd menaquinol oxidase. Cytochrome c1 represents the only c-type cytochrome of C. glutamicum and is unusual in that it contains two covalently bound heme groups, both of which are essential for assembly, stability, and activity of the bc1 complex. The bc1-aa3 branch is of major importance for aerobic growth in minimal medium. Menaquinone reduction by succinate catalyzed by succinate dehydrogenase presumably is an energy-driven process, which must be taken into account in bioenergetic calculations. There are several hints that the composition of the respiratory chain can profoundly influence growth, metabolism, and amino acid production, and future studies with defined C. glutamicum mutants should throw more light on this topic. Eventually, it will be possible to improve product formation by engineering the system of oxidative phosphorylation.

ACKNOWLEDGMENTS Support of the studies on Corynebacterium glutamicum in the laboratory of M.B. by the EU project “VALPAN,” the BMBF “Genomik” program (Genome research on bacteria relevant for agriculture, environment, and biotechnology; cluster IV: Corynebacteria), the BMBF “Proteomics” program (New methods for proteome analysis: Application and combination with metabolome analysis using Corynebacterium glutamicum as an example), the Degussa AG, and the DFG-Graduiertenkolleg “Molekulare Physiologie: Stoff- und Energieumwandlung” is gratefully acknowledged. The authors thank Volker Wendisch for stimulating discussions and Hermann Sahm for continuous and generous support.

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43. Messner KR and Imlay JA. (2002) Mechanism of superoxide and hydrogen peroxide formation by fumarate reductase, succinate dehydrogenase, and aspartate oxidase. J. Biol. Chem. 277:42563–42571. 44. Möckel B, Marx A, and Pfefferle W. (2002) Novel nucleotide sequences coding for the glbO gene. Patent EP1287143. 45. Molenaar D, van der Rest ME, and Petrovic S. (1998) Biochemical and genetic characterization of the membrane-associated malate dehydrogenase (acceptor) from Corynebacterium glutamicum. Eur. J. Biochem. 254:395–403. 46. Molenaar D, van der Rest ME, Drysch A, and Yucel R. (2000) Functions of the membrane-associated and cytoplasmic malate dehydrogenases in the citric acid cycle of Corynebacterium glutamicum. J. Bacteriol. 182:6884–6891. 47. Nicholls DG and Ferguson SJ. (2002) Bioenergetics 3. Academic Press, London. 48. Niebisch A and Bott M. (2001) Molecular analysis of the cytochrome bc1-aa3 branch of the Corynebacterium glutamicum respiratory chain containing an unusual diheme cytochrome c1. Arch. Microbiol. 175:282–294. 49. Niebisch A and Bott M. (2003) Purification of a cytochrome bc1-aa3 supercomplex with quinol oxidase activity from Corynebacterium glutamicum. Identification of a fourth subunit of cytochrome aa3 oxidase and mutational analysis of diheme cytochrome c1. J. Biol. Chem. 278:4339–4346. 50. O’Brian MR and Thöny-Meyer L. (2002) Biochemistry, regulation and genomics of haem biosynthesis in prokaryotes Adv. Microb. Physiol. 46:257–318. 51. Paulsen KE, Orville AM, Frerman FE, Lipscomb JD, and Stankovich MT. (1992) Redox properties of electron-transfer flavoprotein ubiquinone oxidoreductase as determined by EPR-spectroelectrochemistry. Biochemistry 31:11755–11761. 52. Pfefferle W, Marx A, and Möckel B. (2001) Polynucleotide sequences from Corynebacterium glutamicum coding for succinate dehydrogenase (sdhA, sdhB, sdhC). Patent EP1106684. 53. Pittman MS, Corker H, Wu G, Binet MB, Moir AJ, and Poole RK. (2002) Cysteine is exported from the Escherichia coli cytoplasm by CydDC, an ATP-binding cassettetype transporter required for cytochrome assembly. J. Biol. Chem. 277:49841–49849. 54. Richardson DJ, Berks BC, Russell DA, Spiro S, and Taylor CJ. (2001) Functional, biochemical and genetic diversity of prokaryotic nitrate reductases. Cell. Mol. Life Sci. 58:165–178. 55. Ruzicka FJ and Beinert H. (1977) A new iron-sulfur flavoprotein of the respiratory chain. A component of the fatty acid beta oxidation pathway. J. Biol. Chem. 252:8440–8445. 56. Saiki K, Mogi T, and Anraku Y. (1992) Heme O biosynthesis in Escherichia coli: the cyoE gene in the cytochrome bo operon encodes a protoheme IX farnesyltransferase. Biochem. Biophys. Res. Commun. 189:1491–1497. 57. Saiki K, Mogi T, Hori H, Tsubaki M, and Anraku Y. (1993) Identification of the functional domains in heme O synthase. Site-directed mutagenesis studies on the cyoE gene of the cytochrome bo operon in Escherichia coli. J. Biol. Chem. 268:26927–26934. 58. Sakamoto J, Shibata T, Mine T, Miyahara R, Torigoe T, Noguchi S, Matsushita K, and Sone N. (2001) Cytochrome c oxidase contains an extra charged amino acid cluster in a new type of respiratory chain in the amino-acid-producing Gram-positive bacterium Corynebacterium glutamicum. Microbiology 147:2865–2871. 59. Salazar D, Zhang L, deGala GD, and Frerman FE. (1997) Expression and characterization of two pathogenic mutations in human electron transfer flavoprotein. J. Biol. Chem. 272:26425–26433.

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60. Saraste M, Metso T, Nakari T, Jalli T, Lauraeus M, and van der Oost J. (1991) The Bacillus subtilis cytochrome c oxidase. Variations on a conserved protein theme. Eur. J. Biochem. 195:517–525. 61. Schirawski J and Unden G. (1998) Menaquinone-dependent succinate dehydrogenase of bacteria catalyzes reversed electron transport driven by the proton potential. Eur. J. Biochem. 257:210–215. 62. Schnorpfeil M, Janausch IG, Biel S, Kröger A, and Unden G. (2001) Generation of a proton potential by succinate dehydrogenase of Bacillus subtilis functioning as a fumarate reductase. Eur. J. Biochem. 268:3069–3074. 63. Schryvers A, Lohmeier E, and Weiner JH. (1978) Chemical and functional properties of the native and reconstituted forms of the membrane-bound, aerobic glycerol-3phosphate dehydrogenase of Escherichia coli. J. Biol. Chem. 253:783–788. 64. Sekine H, Shimada T, Hayashi C, Ishiguro A, Tomita F, and Yokota A. (2001) H+ATPase defect in Corynebacterium glutamicum abolishes glutamic acid production with enhancement of glucose consumption rate. Appl. Microbiol. Biotechnol. 57:534–540. 65. Shvinka JE, Toma MK, Galinina NI, Skards IV, and Viesturs UE. (1979) Production of superoxide radicals during bacterial respiration. J. Gen. Microbiol. 113:377–382. 66. Simkovic M, Degala GD, Eaton SS, and Frerman FE. (2002) Expression of human electron transfer flavoprotein-ubiquinone oxidoreductase from a baculovirus vector: kinetic and spectral characterization of the human protein. Biochem. J. 364:659–667. 67. Sone N, Tsukita S, and Sakamoto J. (1999) Direct correlationship between proton translocation and growth yield: An analysis of the respiratory chain of Bacillus stearothermophilus. J. Biosci. Bioeng. 87:495–499. 68. Sone N, Nagata K, Kojima H, Tajima J, Kodera Y, Kanamaru T, Noguchi S, and Sakamoto J. (2001) A novel hydrophobic diheme c-type cytochrome. Purification from Corynebacterium glutamicum and analysis of the qcrCBA operon encoding three subunit proteins of a putative cytochrome reductase complex. Biochim. Biophys. Acta 1503:279–290. 69. Sugiyama Y, Kitano K, and Kanzaki T. (1973) Purification of cytochrome a of an Lglutamate-producing microorganism, Brevibacterium thiogenitalis. Agric. Biol. Chem. 37:1607–1612. 70. Sugiyama Y, Kitano K, and Kanzaki T. (1973) Role of copper ions in regulation of L-glutamate biosynthesis. Agric. Biol. Chem. 37:1837–1847. 71. Sun G, Sharkova E, Chesnut R, Birkey S, Duggan MF, Sorokin A, Pujic P, Ehrlich SD, and Hulett FM. (1996) Regulators of aerobic and anaerobic respiration in Bacillus subtilis. J. Bacteriol. 178:1374–1385. 72. Thöny-Meyer L. (1997) Biogenesis of respiratory cytochromes in bacteria. Microbiol. Mol. Biol. Rev. 61:337–376. 73. Toma MK, Shvinka YE, Ruklisha MP, Sakse AK, and Baburin LA. (1984) Cyanideresistant oxygen uptake in lysine-synthesizing bacteria Brevibacterium flavum 22 LD. Prikl. Biokhim. Mikrobiol. 20:95–100. 74. Trutko SM, Kuznetsova NN, Balitskaya RM, and Akimenko VK. (1982) Effect of supersynthesis of glutamic acid on development of cyanide-resistant respiration in the bacterium Corynebacterium glutamicum. Biochemistry Moscow 47:1356–1364. 75. Turrens JF. (1997) Superoxide production by the mitochondrial respiratory chain. Biosci. Rep. 17:3–8. 76. Villani G, Tattoli M, Capitanio N, Glaser P, Papa S, and Danchin A. (1995) Functional analysis of subunits III and IV of Bacillus subtilis aa3-600 quinol oxidase by in vitro mutagenesis and gene replacement. Biochim. Biophys. Acta 1232:67–74.

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77. Wayne LG and Hayes LG. (1998) Nitrate reduction as a marker for hypoxic shiftdown of Mycobacterium tuberculosis. Tuber. Lung. Dis. 79:127–132. 78. Wayne LG and Sohaskey CD. (2001) Nonreplicating persistence of Mycobacterium tuberculosis. Annu. Rev. Microbiol. 55:139–163. 79. Weber I, Fritz C, Ruttkowski S, Kreft A, and Bange FC. (2000) Anaerobic nitrate reductase (narGHJI) activity of Mycobacterium bovis BCG in vitro and its contribution to virulence in immunodeficient mice. Mol. Microbiol. 35:1017–1025. 80. Wendisch VF. (2003) Genome-wide expression analysis in Corynebacterium glutamicum using DNA microarrays. J. Biotechnol. 104:273–285. 81. Witt H and Ludwig B. (1997) Isolation, analysis, and deletion of the gene coding for subunit IV of cytochrome c oxidase in Paracoccus denitrificans. J. Biol. Chem. 272:5514–5517. 82. Xia ZX and Mathews FS. (1990) Molecular structure of flavocytochrome b2 at 2.4 Å resolution. J. Mol. Biol. 212:837–863.

14

Nitrogen Metabolism and Its Regulation A. Burkovski

CONTENTS 14.1 Introduction ..................................................................................................333 14.2 Uptake of Nitrogen Sources ........................................................................334 14.2.1 Ammonium Uptake Systems ...........................................................335 14.2.2 Uptake of Urea and Urease Activity ...............................................335 14.2.3 Transport of Other Nitrogen Sources ..............................................336 14.3 Assimilation of Ammonium.........................................................................337 14.3.1 Glutamate Dehydrogenase ...............................................................339 14.3.2 The Glutamine Synthetase/Glutamate Synthase Pathway...............339 14.4 Signal Transduction......................................................................................340 14.4.1 Regulation of GS Activity ...............................................................340 14.4.2 The GlnK/UTase Pathway ...............................................................341 14.5 Regulation of Transcription .........................................................................342 14.5.1 The Global Regulator Protein AmtR ...............................................342 14.5.2 Influence of Two-Component Signal Transduction Systems ..........344 14.5.3 Nitrogen Control and Sigma Factors...............................................345 14.6 Nitrogen Control in Corynebacteria ............................................................345 14.7 Open Questions ............................................................................................345 References..............................................................................................................346

14.1 INTRODUCTION Almost all of the macromolecules in a bacterial cell, e.g., proteins, nucleic acids, and cell wall components, contain nitrogen. Thus, most prokaryotes have developed elaborate mechanisms to provide an optimal nitrogen supply for metabolism and to overcome and survive situations of nitrogen starvation. The uptake and assimilation of nitrogen sources in Corynebacterium glutamicum, as well as connected regulatory mechanisms, will be described in the following sections (for review, see also [5,6]).

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14.2 UPTAKE OF NITROGEN SOURCES The first step in the utilization of a nitrogen source is its uptake into the cell. This can be mediated either simply by passive diffusion or by active transport processes. In C. glutamicum a considerable number of transport systems for the uptake of nitrogen sources have been characterized biochemically and on the genetic level (Figure 14.1). Transport(er) (Gene)

Substrate

Driving force

NH3

concentration gradient

Amt (amt )

NH4+

membrane potential

AmtB (amtB)

NH4+ or NH3

?

NH4+ or NH3

?

GltS (gltS ) GluABCD (gluABCD)

L-glutamine/Na+

membrane potential

L-glutamate/Na+

membrane potential

L-glutamate

urea/H+ urea

ATP-hydrolysis

membrane potential concentration gradient

FIGURE 14.1 Uptake of nitrogen sources in C. glutamicum. Some nitrogen sources, e.g., ammonia and urea, can enter the cell simply by diffusion; for others, specific uptake systems have to be synthesized. In C. glutamicum, transporters for the uptake of ammonium [22,42], urea [43], L-glutamine [41], and L-glutamate [7,21,53] have been studied. For different uptake systems, the corresponding genes have been identified (shown in parentheses). Biochemically characterized transport proteins for which the corresponding genes are unknown until now are shaded in grey.

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14.2.1 AMMONIUM UPTAKE SYSTEMS Ammonium is an excellent nitrogen source for C. glutamicum. In the presence of high concentrations of this solute in the environment, diffusion of uncharged ammonia (NH3), which is in equilibrium with the protonated ammonium (NH4+), across the cytoplasmic membrane into the cytoplasm is sufficient for growth. When diffusion becomes limiting for metabolism, special transporters are activated to cope with this situation of nitrogen shortage (Figure 14.1). The characterization of the C. glutamicum amt gene, and of the corresponding transport protein [42], was the first study of a bacterial ammonium uptake system on the molecular level. Amt mediates the transport of methylammonium — an ammonium analog that does not serve as a nitrogen source in C. glutamicum — into the cell, with a high affinity of 44 ± 7 μM and a maximal velocity of 25 ± 5 nmol min–1 (mg dry weight)–1. Uptake is strictly dependent on the membrane potential and stops immediately after the addition of protonophores such as carbonyl cyanide m-chlorophenylhydrazone (CCCP) [42]. In the presence of 50 μM methylammonium, 10 μM ammonium is sufficient for a half-maximal inhibition of methylammonium uptake, indicating an even higher affinity of Amt to this solute [22]. As shown by the determination of the apparent Km at different pH values, ammonium and methylammonium, but not ammonia and methylamine, are substrates of the Amt permease [22]. In addition to amt, another gene encoding an ammonium permease has been isolated in C. glutamicum, originally named amtP, for amt paralog [17]. Based on a great number of homologs cloned from different bacteria, sequence similarity analyses, and a typical chromosomal arrangement together with the glnK gene (see Section 14.4), which is conserved in many Eubacteria and Archaea [51], this gene was later renamed and is designated as amtB now [18,22]. The deduced amino acid sequences of Amt and AmtB show an amino acid sequence identity of 38%. In contrast to Amt, the AmtB carrier exclusively accepts ammonium as transported substrate. This property makes an exact kinetic characterization of transport difficult, since rather than following the uptake of radioactive label into the cell, the decrease of ammonium in the medium has to be measured, either by the use of an ammoniumselective electrode or by colorimetric tests. Nevertheless, the determination of ammonium consumption rates in the wild-type and in different mutant strains carrying single and double mutations of the amt and amtB gene indicates that (i) AmtB transports ammonium into the cell and (ii) even a third ammonium uptake system might be present in C. glutamicum, since a considerable ammonium consumption rate has been observed in the absence of Amt and AmtB, even when diffusion was negligible due to a very low ammonium concentration of 200 μM [22].

14.2.2 UPTAKE

OF

UREA

AND

UREASE ACTIVITY

Urea is excreted by a variety of organisms into the environment and is therefore a readily available nitrogen source. In bacteria that are able to utilize urea, the molecule is effectively hydrolyzed by urease to ammonium and CO2 (for review, see [24]). Beginning in the 1960s urease activity has been used to classify L-glutamic acid–producing bacteria [48], and C. glutamicum has been described as urease-positive, which

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is in line with the fact that urea can serve as sole nitrogen source for this organism. Although urea is small and uncharged and can therefore easily pass the bacterial cytoplasmic membrane, C. glutamicum, like many prokaryotes, synthesizes energydependent transport systems for its uptake. When present in high concentrations, urea crosses the C. glutamicum cytoplasmic membrane by passive diffusion, a process for which a permeability coefficient of 9 × 10–7 cm s–1 has been determined [43]. Only under conditions of nitrogen starvation is an energy-dependent urea uptake system synthesized. Carrier-mediated urea transport occurs by a secondary transport system, most likely in symport with protons. With a Km for urea of 8 μM, the affinity of this uptake system is much higher than the affinity of urease toward its substrate (Km approximately 55 mM) [43]. The maximal urea uptake rate depends on the level of expression and is, at only 2.0 to 3.5 nmol min–1 (mg dry weight)–1, relatively low [43]. While the gene encoding the urea permease is unknown, those coding for urease were recently isolated and sequenced [25,32]. The structural urease subunits α, β, and γ are encoded by the ureABC genes, which are transcribed together and form a common mRNA [25], and the ureEFGD genes, which encode accessory urease subunits. A regulation of the ureABCEFGD gene cluster on the level of transcription could not be observed by RNA hybridization experiments or RT-PCR [25]; however, experiments using DNA microarrays showed a clear up-regulation of transcription in response to nitrogen starvation (G. Beckers, A.K. Bendt, A. Burkovski, and M. Farwick, unpublished data). The latter data are in accordance with urease activity measurements. These show a relatively low activity of 0.9 μmol min–1 (mg protein)–1 in extracts of cells grown in complex medium and the highest activity of 7.8 μmol min–1 (mg protein)–1 in extracts from nitrogen-starved cells [25]. Depending on the growth conditions, urease activity varies between 0.9 and 2.2 μmol min–1 (mg protein)–1 for cells grown in ammonium-rich minimal medium [25,32] between 1.0 and 1.6 μmol min–1 (mg protein)–1, when glutamine is used as the nitrogen source [32] and between 2.0 and 6.1 μmol min–1 (mg protein)–1 when different concentrations of urea were added to the medium [25,32].

14.2.3 TRANSPORT

OF

OTHER NITROGEN SOURCES

C. glutamicum can utilize L-glutamine as a nitrogen source, which can be easily converted to glutamate by glutaminase or glutamate synthase. L-glutamine uptake is mediated by a secondary active permease with a Km of 36 μM and a Vmax of 12.5 nmol min–1 (mg dry weight)–1. Uptake is driven by the membrane potential and at least one sodium ion is transported together with each L-glutamine molecule. Consequently, the addition of Na+-ionophores or uncouplers impairs or totally abolishes L-glutamine transport across the membrane [41]. The Km for the co-transported Na+ is, at 1.4 mM, much higher than that for L-glutamine [41]. This has consequences for the utilization of this amino acid as a nitrogen source; L-glutamine is an excellent nitrogen source, comparable with ammonium, only when a sufficient sodium concentration is present in the medium. In contrast, low Na+ concentrations impair L-glutamine uptake, leading to a situation of nitrogen limitation even in the presence of high L-glutamine concentrations.

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Although less effective than glutamine, glutamate serves as a nitrogen source for C. glutamicum as well (e.g., [40,52]). Two different L-glutamate uptake systems have been characterized: a binding protein–dependent ABC transporter [20] and a secondary, sodium-coupled carrier [7]. The primary glutamate uptake system is encoded by the gluABCD cluster. It comprises an ATP binding protein (encoded by gluA), an L-glutamate binding protein (the gluB gene product), and two integral membrane proteins (encoded by gluC and gluD). The uptake of L-glutamate occurs with high affinity (Km 0.5 to 1.3 μM) and a Vmax of 15 nmol min–1 (mg dry weight)–1. It is under control of glucose catabolite repression and therefore down-regulated in media using glucose or sucrose as the carbon source [21]. Recently performed DNA microarray experiments indicate that expression of the gluABCD genes is also under nitrogen control [6], i.e., transcription is repressed under high nitrogen supply. This is in accordance with the observation that high ammonium concentrations inhibit growth of C. glutamicum when L-glutamate is used as the carbon source [40]. The secondary L-glutamate uptake system, which is predominantly active in complex media, is characterized by a moderate Km of 0.6 mM for L-glutamate and a Vmax of about 15 nmol min–1 (mg dry weight)–1. Transport is driven by the membrane potential and is dependent on sodium ions; uncouplers or Na+ ionophores inhibit transport [7]. For the co-transported sodium ions, a relatively low affinity (Km of 3.3 mM) was determined. Recently, the corresponding gene, designated gltS, was identified. It encodes a highly hydrophobic protein with 11 to 13 transmembrane helices depending on the assumed start codon [53]. A number of other amino acids can also serve as nitrogen sources for C. glutamicum, e.g., L-alanine, L-asparagine, L-serine, and L-threonine. However, systematic studies on the utilization of different amino acids have not been published. Furthermore, for some amino acids, contradictory observations about their utilization were made, e.g., for the utilization of L-proline. This may be the result of strainspecific differences or of a low affinity of amino acid uptake systems for their coupling ion Na+, as described for glutamine and glutamate. Additionally, substrate mixtures were used in some studies, leading to confusion about the utilization of single amino acids, e.g., L-aspartate [32], as sole nitrogen source. A special pitfall with respect to false-positive results is the presence of high internal L-glutamate and L-glutamine pools in C. glutamicum. Up to 200 mM L-glutamate [20] and 50 mM L-glutamine [50] have been determined in cells grown in minimal medium. These allow — depending on the preculture conditions — approximately two duplications of the cells (with a growth rate comparable to that of cells grown with high nitrogen supply) even in the complete absence of any nitrogen source [22] and may mimic growth with a certain nitrogen source if an appropriate control culture without a nitrogen source is lacking.

14.3 ASSIMILATION OF AMMONIUM C. glutamicum is able to assimilate ammonium via two main pathways, the glutamate dehydrogenase (GDH) and the glutamine synthetase/glutamate synthase (also desig-

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GSinactive (glnA) ?

AMP

ATase (glnE)

GS activity

nitrogen supply

GSactive (glnA)

glutamate

glutamine

GOGAT (gltBD)

oxoglutarate

oxoglutarate

glutamate

GDH (gdh)

glutamate

FIGURE 14.2 Ammonium assimilation and activity regulation of glutamine synthetase. Ammonium, directly taken up [22,42] or generated inside the cell from the cleavage of urea by urease [25,43], is assimilated via the glutamate dehydrogenase (GDH) or glutamine synthetase/glutamate synthase (GS/GOGAT) pathway. Since GDH has only a low affinity to its substrates, at ammonium concentrations below 5 mM [49] the GS/GOGAT pathway takes over, although at an extra cost of one mole of ATP per mole of ammonium fixed. To adapt GS activity exactly to the cellular L-glutamine demand in order to avoid a loss of energy, GS is regulated at the level of activity by adenylyltransferase (ATase, glnE gene product) [17,26]. Grey arrows indicate the direction of signal relay; the question mark shows an unknown sensory input.

nated glutamine synthetase/glutamate-oxoglutarate transferase, GS/GOGAT) pathways (Figure 14.2). Other mechanisms to assimilate ammonium, e.g., fixation by alanine dehydrogenase, were not observed, when in vivo flux measurements of ammonium assimilation were carried out [50]. When present in high concentrations, ammonium is primarily fixed by glutamate dehydrogenase (GDH), thereby oxidizing one mole of NADPH per mole of ammonium assimilated. At ammonium concentrations below approximately 5 mM [49], the glutamine synthetase/glutamate synthase (GS/GOGAT) system takes over, although at the cost of an extra mole of ATP per mole of ammonium fixed. Thus, it is advantageous to down-regulate the GS/GOGAT pathway when ammonium is present at high concentrations to prevent a waste of energy. However, even under nitrogen surplus, GS must remain at least partially active to satisfy the cellular demand for glutamine. In fact, the GS/GOGAT pathway is significantly up-regulated in response to nitrogen deprivation, while GS is active during nitrogen surplus as

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well. In vivo flux analyses revealed that without nitrogen limitation 28% of the ammonium is assimilated via the GS reaction in C. glutamicum [50]. This is an unusually high fraction compared, for example, to only 13% in E. coli [33], and it was speculated that this situation reflects the higher glutamine demand of C. glutamicum for cell wall synthesis [50].

14.3.1 GLUTAMATE DEHYDROGENASE Due to its glutamate synthesis function, the GDH enzyme was investigated with respect to its biochemical properties and influence on L-glutamate production starting in the early 1960s [19,30,38] in Brevibacterium flavum, now known as C. glutamicum ssp. flavum. With Km values of 3.08 mM for ammonium and 5.72 mM for oxoglutarate, the GDH enzyme has only a low affinity for its substrates. The affinity for its product L-glutamate is even lower, with a Km of 100 mM [38]. The C. glutamicum gdh gene has been cloned and characterized [2] and a defined gdh mutant strain was constructed. Surprisingly, analysis of the gdh mutant showed that a functional GDH is not required for growth and glutamate production [3,49], although GDH activity in the wild-type is orders of magnitude higher than that of glutamate synthase [3,44,52]. For example, GDH activities between 1.8 and 2.5 μmol min–1 (mg protein)–1 were determined for cells grown in different minimal media with glucose as carbon and energy source [2,3,50] while GOGAT activities of only 0.003 and 0.05 μmol min–1 (mg protein)–1 were reported [1,3,49,50]. It was proposed that an unknown regulatory mechanism recruits the GS/GOGAT pathway in the absence of GDH to take over glutamate synthesis [3], and in fact, in a gdh mutant strain a high GOGAT activity was observed even when cells were grown under nitrogen surplus [49].

14.3.2 THE GLUTAMINE SYNTHETASE/GLUTAMATE SYNTHASE PATHWAY The biochemical investigation of C. glutamicum glutamine synthetase and glutamate synthase enzymes started in 1978 [39,44,45,52]. From analytical ultracentrifugation and gel-filtration experiments, an octameric structure built from identical subunits was proposed for glutamine synthetase [31,46], which would be in contrast to GS-I enzymes from other bacteria, which are composed of 12 identical subunits (for review, see [23]). The first reported Km values for this enzyme were astonishingly high: 7.9 mM for glutamate, 5 mM for ammonium, and 1.2 mM for ATP [46]. However, later studies showed that these values change dramatically depending on the concentration of divalent cations in the enzyme assay buffer [47,54]. For example, the maximal GS activity at an optimal concentration of Mn2+ was about three times higher than that at the optimal Mg2+concentration and the apparent Km for Mn2+ATP was much lower than that for Mg2+-ATP, about 40 μM versus 600 μM [54]. Therefore, the physiological relevance of the published in vitro measurements remains unclear. Indeed, the increase of GS activity in response to nitrogen limitation hints toward a significantly higher ammonium affinity of this enzyme in vivo. Isolation and analysis of the glutamine synthetase–encoding glnA gene revealed that C. glutamicum possesses a GS-Iβ-subtype enzyme [16]. This GS type is characterized by a regulation via post-translational modification, namely adenylylation.

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GS-Iα-subtype proteins, which are mainly distributed in low-G+C Gram-positive bacteria, are not post-translationally modified, and also the heat-labile GS-II enzymes are (with few exceptions) not modified [4,23]. Analyses of a glnA deletion mutant showed that glnA is essential for glutamine synthesis, since cells lacking this gene are glutamine-auxotrophic. Recently, a gene proposed to encode a GS-Iα-subtype glutamine synthetase was identified and designated glnA2. It forms an operon together with glnE, which encodes an adenylyltransferase. Transcription of glnA2 seems to be constitutive and neither its deletion nor its overexpression exhibits any effect on glutamine synthetase activity, even in an glnA deletion background [26]. Therefore, the physiological function of the glnA2 gene product remains unclear. When cells were grown under nitrogen surplus, no GOGAT activity could be detected by in vitro enzyme assays [1,52] or in vivo flux measurements [50]. This is in accordance with the results of reporter gene assays and RNA hybridization experiments that revealed that transcription of the GOGAT-encoding gltBD operon strictly depends on nitrogen starvation [1,37], and only in this situation is the GOGAT enzyme synthesized. The expression of gltBD is under control of the global repressor protein AmtR (see Section 14.5.1) as demonstrated by gel-shift experiments and analysis of gltB transcription in an amtR deletion strain [1]. In contrast to its pivotal function in other bacteria such as Bacillus subtilis [8], Escherichia coli [10,13], and Klebsiella aerogenes [10], glutamate synthase exhibits no crucial role in C. glutamicum. Inactivation of the gltB or gltD gene, which results in a total loss of GOGAT activity, has no detrimental effect on growth when cells are cultured in standard minimal medium, and only a minor increase in the doubling time of the corresponding mutant strains has been observed in the presence of limiting amounts of nitrogen sources such as ammonium or urea. Moreover, these mutants show a normal L-glutamate production phenotype, indicating that glutamate synthase has no central function in L-glutamate production by C. glutamicum [1]. Moreover, while glutamate synthase is discussed as a central enzyme interconnecting nitrogen and carbon metabolism in B. subtilis [8], its activity in C. glutamicum is not influenced by changes in carbon availability [1,37].

14.4 SIGNAL TRANSDUCTION 14.4.1 REGULATION

OF

GS ACTIVITY

Amino acid sequence analyses of the C. glutamicum glutamine synthetase revealed a typical adenylylation site [16], a tyrosine residue at amino acid position 405. Two lines of evidence showed that GS activity is controlled by adenylylation/deadenylylation via adenylyltransferase (ATase). First, a glnA allele encoding a phenylalanine instead of a tyrosine residue at position 405 was constructed by site-directed mutagenesis. In the corresponding mutant strain, GS activity is, in contrast to the wild-type, not down-regulated upon addition of ammonium to a nitrogen-starved culture [17]. Second, the gene encoding adenylyltransferase, glnE, which was identified during the systematic sequencing of the C. glutamicum genome, was inactivated. As expected, a glnE deletion mutant revealed no GS activity regulation [26]. Further GS activity measurements using cell extracts from a glnK deletion strain

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showed that C. glutamicum adenylyltransferase works independently from the GlnK nitrogen signal transduction protein (see Section 14.4.2). While GlnK/AmtR-mediated transcription control is lost in this mutant (see Section 14.5.1), the regulation of GS activity is not impaired [6].

14.4.2 THE GLNK/UTASE PATHWAY GlnK and uridylyltransferase are key components for nitrogen signaling in C. glutamicum [17,27]. Although these polypeptides are highly similar to their E. coli counterparts, it has been shown that the nitrogen regulation cascade in C. glutamicum functions by a new mechanism (Figure 14.3). The genes encoding uridylyltransferase and GlnK, glnD and glnK, are organized in an operon together with amtB, which codes for an ammonium permease (see Section 14.2.1). It has been shown that GlnK is essential for nitrogen control and that signal transduction is transmitted by modification of this protein at a specific amino acid residue, tyrosine 51 [27]. As a consequence, also a glnD deletion strain lacking uridylyltransferase is impaired in its response to nitrogen shortage and reveals a decreased growth rate in the presence of limiting amounts of ammonium ?

UTase (glnD)

GlnK (glnK)

GlnK (glnK)

AMP

AmtR (amtR)

AmtR (amtR)

repression of transcription expression of AmtR-controlled genes

LOW

NITROGEN SUPPLY

HIGH

FIGURE 14.3 Nitrogen-starvation–dependent transcription control. Expression of nitrogencontrolled genes is governed by the master regulator AmtR [18,27], which itself is controlled by a signal cascade of uridylyltransferase (UTase, glnD gene product) and GlnK. When ammonium is present at high concentrations, AmtR blocks transcription of various genes (see Table 14.1). In response to nitrogen starvation, the UTase enzyme adenylylates the GlnK signal transduction protein, which is assumed to directly interact with AmtR in this modified form, leading to the release of repression. Grey arrows indicate the direction of signal relay; the question mark shows an unknown sensory input.

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or urea [27]. Proteome analyses by [35S]-methionine in vivo labeling and twodimensional gel electrophoresis followed by autoradiography showed that the glnD mutant is impaired in its response to nitrogen starvation on the protein level [27] and provided first proteome data for the nitrogen control of the split D,L-diaminopimelate synthesis pathway. In the wild-type tetrahydrodipicolinate succinylase, thioredoxin, and a protein with unknown function were newly synthesized in response to nitrogen starvation while synthesis of 50S RNA subunit L12 ceased [27]. Tetrahydrodipicolinate succinylase is part of one of the two pathways for the synthesis of m-diaminopimelate present in C. glutamicum, the succinylase and dehydrogenase pathways [36,55]. While diaminopimelate dehydrogenase is active at high concentrations of ammonium, the succinylase pathway is ammonium-independent and is obviously activated in response to nitrogen starvation. These changes in the protein profile were not observed in the glnD mutant strain, demonstrating that synthesis of these proteins is under control of the uridylyltransferase/GlnK pathway [27]. In contrast to E. coli, in which glnD is constitutively expressed, in C. glutamicum transcription of glnD is up-regulated upon nitrogen limitation. This indicates that the glnD gene product is likely not the primary sensor of the nitrogen status in C. glutamicum, as has been shown for enterobacteria. In accordance with this idea, the overexpression of glnD leads to a transcription of nitrogen-starvation–controlled genes even in the presence of high ammonium concentrations, which is difficult to explain with a sensor function of UTase. Furthermore, quantification of cytoplasmic amino acid pools excluded the possibility that a drop in the glutamine concentration is perceived as the signal for nitrogen starvation by C. glutamicum, as is found in enterobacteria. Up to 200 mM internal glutamate [20] and 50 mM internal glutamine [50] have been determined in C. glutamicum cells grown in minimal medium. While the expression of nitrogen-starvation–controlled genes occurs within a few minutes after the onset of nitrogen limitation, the internal L-glutamate and L-glutamine pools are only gradually consumed during several hours of starvation. Obviously, these internally accumulated amino acids serve as buffers for the nitrogen and/or carbon supply of the C. glutamicum cell, rather than as indicators of the nitrogen status. This is in contrast to the situation in Salmonella typhimurium, B. subtilis, and Klebsiella pneumoniae, in which nitrogen limitation is perceived as a drop in the internal glutamine pool [14,15,35]. In C. glutamicum, determination of intracellular ammonium concentrations in cells grown under nitrogen surplus or in nitrogenstarved cells showed a correlation of the decrease in the intracellular ammonium level in response to nitrogen shortage and the response on the level of transcription. This has led to the idea that the intracellular ammonium pool might reflect the cellular nitrogen status in C. glutamicum [27].

14.5 REGULATION OF TRANSCRIPTION 14.5.1 THE GLOBAL REGULATOR PROTEIN AMTR The expression of many nitrogen-starvation–induced genes in C. glutamicum is controlled by the AmtR protein, a TetR-type repressor protein [18]. This master

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343

TABLE 14.1 AmtR-Binding Motifs in C. glutamicum

Gene or Operon amtB-glnK-glnD

Protein(s)

Position AmtR-Binding (Relative to the Sequence Start Codon)

Reference

Ammonium transporterGlnK-UTase Ammonium/methylammonium permease-putative ornithine decarboxylaseputative sarcosine oxidase glutamine synthetase (GS-Iβ-subtype) ABC transporter for the uptake of L-glutamate

ATCTATAGN1-ATAG ATCTATAGN4-ATAG

–140 to –128

[18]

–61 to –46 and –103 to –87

[18]

ATCTAT

–142 to –137

[26]

ATCTAT-N6ATAG

–190 to –175

gltBD hkm

Glutamate synthase Putative regulator of glutamate synthase

ATCTATAG ATCTATAG

–112 to –105 –99 to –92

ureABCEFGD

Urease

ATCTATA

–130 to –124

G. Beckers, A. Burkovski, J. Kalinowski; unpublished results [1] G. Beckers, A. Burkovski, J. Kalinowski; unpublished results G. Beckers, A. Burkovski, J. Kalinowski; unpublished results

amt-ocd-soxA

glnA gluABCD

Note: The binding of AmtR was shown by deletion analyses and gel-shift experiments for amtB-glnKglnD, amt-ocd-soxA, glnA, and gltBD; other motifs were identified by database searches using HiddenMarkov models and AmtR control was verified by RNA hybridization experiments (gluABCD, hkm, and ureABCEFGD). The AmtR-binding DNA regions are located either on the coding DNA strand (e.g., upstream of amt and amtB) or on the noncoding DNA strand (e.g., upstream of the gltB gene).

regulator represses transcription of more than 20 genes during nitrogen surplus (Table 14.1). Consequently, deletion of the amtR gene unblocks expression control and leads to a deregulated transcription of AmtR-regulated genes [18]. In the wildtype, AmtR is controlled by the UTase/GlnK pathway (Figure 14.3, see also Section 14.4.2), most likely by a direct interaction with the modified GlnK protein. Deletion of the glnD or glnK gene or replacement of the crucial tyrosine residue 51 by phenylalanine in GlnK interrupts the transfer of the nitrogen starvation signal to AmtR and leads to a permanent repression of AmtR-controlled genes [27].

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Gel-retardation experiments with extracts from amtR-expressing E. coli cells and with purified AmtR protein indicated that AmtR binds to a particular region upstream of amt, with the sequence ATCTATAGN4ATAG. This was also shown in vivo using a yeast-one-hybrid system [18]. Subsequent studies showed that AmtR not only regulates transcription of the amt gene, but also expression of the ocd and soxA genes, which are located downstream of amt and form an operon together with this gene. The soxA gene encodes a putative sarcosine oxidase, and ocd codes for a putative ornithine cyclodeaminase. Based on the genetic organization of the corresponding genes, a role of these proteins in nitrogen metabolism might be expected. However, functional data are as of yet not available. Besides amt-ocd-soxA transcription, AmtR controls transcription of the amtBglnK-glnD operon encoding an ammonium carrier, GlnK, and UTase [18]; the gltBD operon encoding glutamate synthase [1]; and the glnA gene coding for glutamine synthetase [26]. Interestingly, the stringency of AmtR repression differs for the particular genes, most likely due to variations in the AmtR binding motif. Upstream of the strictly regulated amt gene, two ATCTATAGN4ATAG sites have been identified [18], whereas upstream of the weakly repressed glnA gene, only a truncated AmtR motif with the sequence ATCTAT is found [26]. Known AmtR targets and binding motifs identified up to now are summarized in Table 14.1.

14.5.2 INFLUENCE OF TWO-COMPONENT SIGNAL TRANSDUCTION SYSTEMS In many bacteria, two-component signal transduction systems, which comprise a sensor kinase and a response regulator protein, regulate the cellular adaptation to changing environmental conditions. The NtrB/NtrC system, for example, activates the transcription of σ54-dependent promoters in response to nitrogen limitation in E. coli and other enterobacteria. A two-component system with a similar function is most likely absent in C. glutamicum, since the investigation of mutant strains of the thirteen two-component signal transduction systems encoded in the C. glutamicum genome revealed no hint of a participation of these proteins in nitrogen regulation (M. Bott, A. Burkovski, A. Faust, and S. Schaffer, unpublished data). A gene identified upstream of the gltBD operon was designated hkm due to the similarity of the corresponding protein to a histidine kinase of a two-component signal transduction system from Mesorhizobium loti [37]. However, the homologous protein region does not span the typical phosphorylation domain or ATP-binding site of histidine kinases but is confined to a so-called PAS domain, which is widely distributed in pro- and eukaryotes and found in a variety of proteins besides histidine kinases including chemo- and photoreceptors, circadian clock proteins, cyclic nucleotide phosphodiesterases, threonine/serine kinases, and voltage-activated channels. Based on gene disruption analyses, growth tests and enzyme activity measurements, a function of the hkm gene product in the regulation of GOGAT activity was suggested [37]. Reporter gene studies revealed that transcription of the hkm gene is moderately up-regulated in response to nitrogen limitation [37]. A detailed characterization of the function of this gene is still missing.

Nitrogen Metabolism and Its Regulation

14.5.3 NITROGEN CONTROL

AND

345

SIGMA FACTORS

Alternative sigma factors of RNA polymerase control the expression of genes that encode proteins necessary to survive conditions of starvation and stress in many bacteria. Although the presence of such sigma factors was reported for C. glutamicum ssp. lactofermentum [29] and ssp. flavum [11,12], no indication has yet been found that these alternative sigma factors are involved in the regulation of nitrogen metabolism. On the contrary, analyses of transcripts from amt [18], glnA [26], and gltB [1,37] showed identical transcriptional start sites for the corresponding mRNAs isolated from cells grown under nitrogen surplus or under nitrogen limitation. These data make a regulatory function of alternative sigma factors for the transcription of these genes rather unlikely.

14.6 NITROGEN CONTROL IN CORYNEBACTERIA The schematic presentation of nitrogen metabolism and its regulation as shown in Figures 14.2 and 14.3 seems to be valid for corynebacteria other than C. glutamicum. The genes encoding components of nitrogen metabolism and nitrogen control present in the Corynebacterium diphtheriae genome and their organization showed striking similarities with their C. glutamicum counterparts. In the sequence published by the Sanger Centre, genes encoding AmtR, AmtB, GlnK, uridylyltransferase, adenylyltransferase, glutamate dehydrogenase, GS-Iα, and GS-Iβ were identified [28]. Furthermore, the proposed functions for AmtR and uridylyltransferase were verified by disruption of the corresponding genes and characterization of the corresponding mutant strains by RNA hybridization experiments [28]. After release of the completely annotated sequence, further work will be necessary to validate that both organisms control their nitrogen metabolism in an identical way, or to show distinct differences, which might reflect the pathogenic lifestyle of C. diphtheriae. Corynebacterium efficiens is a new L-glutamic-acid–producing Corynebacterium species that was isolated from soil samples and onion bulbs [9]. The genome sequence of this organism was determined and database searches revealed that homologs of C. glutamicum proteins discussed in this review are present in C. efficiens strain YS-314 as well: an amt-ocd-soxA cluster, an amtB-glnK-glnD operon, amtR, gdh, glnA, glnE-glnA2, gltBD, a gluABCD cluster, and a putative ureABCEFGD operon were annotated, and for the corresponding amino acid sequences, in general, a high percentage of identical amino acids, between 60 and 80% compared to C. glutamicum proteins, was observed. Thus, it can be proposed that nitrogen metabolism and its regulation is similar in both species.

14.7 OPEN QUESTIONS Although a detailed picture for ammonium assimilation and its regulation in C. glutamicum has been established in recent years, a number of open questions are still to be answered. The most urgent ones to address are the identification of the nitrogen sensor(s) of C. glutamicum and the identity of the molecule(s) whose concentration reflect(s) the cellular nitrogen status.

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More information on nitrogen regulation will be derived in the future from global analysis techniques such as transcriptome analyses via DNA microarrays and proteome investigations using two-dimensional SDS-polyacrylamide gel electrophoresis (2-D SDS-PAGE), which are reviewed in other chapters of this book. The use of 2-D SDS-PAGE has already led to first indications of a connection of the global regulatory networks of nitrogen control and the regulation of carbon and energy metabolism in C. glutamicum [34], and provided proteome data for the nitrogen control of the split D,L-diaminopimelate synthesis pathway [27]. In addition, preliminary DNA microarray data hint at a crosstalk of nitrogen control and respiratory chain regulation (A. Burkovski, A. Hüser, J. Kalinowski, and M. Silberbach, unpublished data), opening new interesting fields of research.

REFERENCES 1. Beckers G, Nolden L, and Burkovski A. (2001) Glutamate synthase of Corynebacterium glutamicum is not essential for glutamate synthesis and is regulated by the nitrogen status. Microbiology 147:2961–2970. 2. Börmann ER, Eikmanns BJ, and Sahm H. (1992) Molecular analysis of the Corynebacterium glutamicum gdh gene encoding glutamate dehydrogenase. Mol. Microbiol. 6:317–326. 3. Börmann-El Kholy ER, Eikmanns BJ, Gutmann M, and Sahm H. (1993) Glutamate dehydrogenase is not essential for glutamate formation in Corynebacterium glutamicum. Appl. Environ. Microbiol. 59:2329–2331. 4. Brown JR, Masuchi Y, Robb FT, and Doolittle WF. (1994) Evolutionary relationships of bacterial and archaeal glutamine synthetases. J. Mol. Evol. 38:566–576. 5. Burkovski A. (2003) I do it my way: Regulation of ammonium uptake and ammonium assimilation in Corynebacterium glutamicum. Arch. Microbiol. 179:83–88. 6. Burkovski A. (2003) Ammonium assimilation and nitrogen control in Corynebacterium glutamicum and its relatives: an example for new regulatory mechanisms in actinomycetes. FEMS Microbiol. Rev. 27:617–628. 7. Burkovski A, Weil B, and Krämer R. (1996) Characterization of a secondary uptake system for L-glutamate in Corynebacterium glutamicum. FEMS Microbiol. Lett. 136:169–173. 8. Faires N, Tobisch S, Bachem S, Martin-Verstraete I, Hecker M, and Stülke J. (1999) The catabolite control protein CcpA controls ammonium assimilation in Bacillus subtilis. J. Mol. Microbiol. Biotechnol. 1:141–148. 9. Fudou R, Jojima Y, Seto A, Yamada K, Rimura E, Nakamatsu T, Hirashi A, and Yamanaka S. (2002) Corynebacterium efficiens sp. Nov., a glutamic-acid-producing species from soil and plant material. Int. J. Syst. Evol. Microbiol. 52:1127–1131. 10. Goss TJ, Perez-Matos A, and Bender RA. (2001) Roles of glutamate synthase, gltBD, and gltF in nitrogen metabolism of Escherichia coli and Klebsiella aerogenes. J. Bacteriol. 183:6607–6619. 11. Halgasova N, Bukovska G, Timko J, and Kormanec J. (2001) Cloning and transcriptional characterization of two sigma factor genes, sigA and sigB, from Brevibacterium flavum. Curr. Microbiol. 43:249–254. 12. Halgasova N, Bukovska G, Ugorcakova J, Timko J, and Kormanec J. (2002) The Brevibacterium flavum sigma factor SigB has a role in the environmental stress response. FEMS Microbiol. Lett. 216:77–84.

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13. Helling RB. (1994) Why does Escherichia coli have two primary pathways for synthesis of glutamate? J. Bacteriol. 176:4664–4668. 14. Hu P, Leighton T, Ishkhanova G, and Kustu S. (1999) Sensing of nitrogen limitation by Bacillus subtilis: comparison to enteric bacteria. J. Bacteriol. 181:5042–5050. 15. Ikeda TP, Shauger AE, and Kustu S. (1996) Salmonella typhimurium apparently perceives external nitrogen limitation as internal glutamine limitation. J. Mol. Biol. 259:589–607. 16. Jakoby M, Tesch M, Sahm H, Krämer R, and Burkovski A. (1997) Isolation of the Corynebacterium glutamicum glnA gene encoding glutamine synthetase I. FEMS Microbiol. Lett. 154:81–88. 17. Jakoby M, Krämer R, and Burkovski A. (1999) Nitrogen regulation in Corynebacterium glutamicum: Isolation of genes involved and biochemical characterization of corresponding proteins. FEMS Microbiol. Lett. 173:303–310. 18. Jakoby M, Nolden L, Meier-Wagner J, Krämer R, and Burkovski A. (2000) AmtR, a global repressor in the nitrogen regulation system of Corynebacterium glutamicum. Mol. Microbiol. 37:964–977. 19. Kimura K. (1962) The significance of glutamic dehydrogenase in glutamic acid fermentation. J. Gen. Appl. Microbiol. 8:253–260. 20. Krämer R and Lambert C. (1990) Uptake of glutamate in Corynebacterium glutamicum. 2. Evidence for a primary active transport system. Eur. J. Biochem. 194:937–944. 21. Kronemeyer W, Peekhaus N, Krämer R, Sahm H, and Eggeling L. (1995) Structure of the gluABCD cluster encoding the glutamate uptake system of Corynebacterium glutamicum. J. Bacteriol. 177:1152–1158. 22. Meier-Wagner J, Nolden L, Jakoby M, Siewe R, Krämer R, and Burkovski A. (2001) Multiplicity of ammonium uptake systems in Corynebacterium glutamicum: Role of Amt and AmtB. Microbiology 147:135–143. 23. Merrick MJ and Edwards RA. (1995) Nitrogen control in bacteria. Microbiol. Rev. 59:604–622. 24. Mobley HL, Island MD, and Hausinger RP. (1995) Molecular biology of microbial ureases. Microbiol. Rev. 59:451–480. 25. Nolden L, Beckers G, Möckel B, Pfefferle W, Nampoothiri KM, Krämer R, and Burkovski A. (2000) Urease of Corynebacterium glutamicum: organisation of corresponding genes and investigation of activity. FEMS Microbiol. Lett. 189:305–310. 26. Nolden L, Farwick M, Krämer R, and Burkovski A. (2001) Glutamine synthetases in Corynebacterium glutamicum: transcriptional control and regulation of activity. FEMS Microbiol. Lett. 201:91–98. 27. Nolden L, Ngouoto-Nkili C-E, Bendt AK, Krämer R, and Burkovski A. (2001) Sensing nitrogen limitation in Corynebacterium glutamicum: The role of glnK and glnD. Mol. Microbiol. 42:1281–1295. 28. Nolden L, Beckers G, and Burkovski A. (2002) Nitrogen assimilation in Corynebacterium diphtheriae: pathways and regulatory cascades. FEMS Microbiol. Lett. 208:287–293. 29. Oguiza JA, Marcos AT, Malumbres M, and Martin JF. (1996) Multiple sigma factor genes in Brevibacterium lactofermentum: characterization of sigA and sigB. J. Bacteriol. 178:550–553. 30. Oshima K, Tanaka K, and Kinoshita S. (1964) Studies on glutamic acid fermentation. XI. Purification and properties of L-glutamic acid dehydrogenase from Micrococcus glutamicus. Agric. Biol. Chem. 28:714–722. 31. Park M-E, Choi S-Y, Kim S-J, and Min K-H. (1989) Properties of glutamine synthetase from Corynebacterium glutamicum. Korean Biochem. J. 22:128–132.

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32. Puskás LG, Inui M, and Yukawa H. (2000) Structure of the urease operon of Corynebacterium glutamicum. DNA Seq. 11:383–394. 33. Reitzer LJ and Magasanik B. (1987) Ammonium assimilation and the biosynthesis of glutamine, glutamate, aspartate, asparagine, L-alanine, and D-alanine. In Neidhardt EC, Ingraham JL, Magasanik B, Schaechter M, and Umbarger HE (Eds.), Escherichia coli and Salmonella. American Society of Microbiology, Washington, D.C, pp. 302–320. 34. Schmid R, Uhlemann E-M, Nolden L, Wersch G, Hecker R, Hermann T, Marx A, and Burkovski A. (2000) Response to nitrogen starvation in Corynebacterium glutamicum. FEMS Microbiol. Lett. 187:83–88. 35. Schmitz RA. (2000) Internal glutamine and glutamate pools in Klebsiella pneumoniae grown under different conditions of nitrogen availability. Current Microbiol. 41:357–362. 36. Schrumpf B, Schwarzer A, Kalinowski J, Pühler A, Eggeling L, and Sahm H. (1991) A functionally split pathway for lysine synthesis in Corynebacterium glutamicum. J. Bacteriol. 173:4510–4516. 37. Schulz AA, Collett HJ, and Reid SJ. (2002) Regulation of glutamine synthetase and glutamate synthase in Corynebacterium glutamicum ATCC 13032. FEMS Microbiol. Lett. 205:361–367. 38. Shiio I and Ozaki H. (1970) Regulation of nicotinamide adenine dinucleotide phosphate-specific glutamate dehydrogenase from Brevibacterium flavum, a glutamateproducing bacterium. J. Biochem. 68:633–647. 39. Shiio I and Ujigawa K. (1978) Enzymes of the glutamate and aspartate synthetic pathways in glutamate-producing Brevibacterium flavum. J. Biochem. 84:647–657. 40. Shiio I, Ozaki H, and Mori M. (1982) Glutamate metabolism in a glutamate-producing bacterium, Brevibacterium flavum. Agric. Biol. Chem. 46:493–500. 41. Siewe RM, Weil B, and Krämer R. (1995) Glutamine uptake by a sodium-dependent secondary transport system in Corynebacterium glutamicum. Arch. Microbiol. 164:98–103. 42. Siewe RM, Weil B, Burkovski A, Eikmanns BJ, Eikmanns M, and Krämer R. (1996) Functional and genetic characterization of the (methyl)ammonium uptake carrier of Corynebacterium glutamicum. J. Biol. Chem. 271:5398–5403. 43. Siewe RM, Weil B, Burkovski A, Eggeling L, Krämer R, and Jahns T. (1998) Urea uptake and urease activity in Corynebacterium glutamicum. Arch. Microbiol. 169:411–416. 44. Sung HA, Tachiki T, Kumagai H, and Tochikura T. (1984) Production and preparation of glutamate synthase from Brevibacterium flavum. J. Ferment. Technol. 62:569–575. 45. Sung HA, Tamaki H, Tachiki T, Kumagai H, and Tochikura T. (1985) Ammonia assimilation by glutamine synthase/glutamate synthase system in Brevibacterium flavum. J. Ferment. Technol. 63:5–10. 46. Tachiki T, Wakisaka S, Kumagai H, and Tochikura T. (1981) Crystallization of Micrococcus glutamicus glutamine synthetase brought about by divalent cations. Agric. Biol. Chem. 47:287–292. 47. Tachiki T, Wakisaka S, Suzuki H, Kumagai H, and Tochikura T. (1981) Variation of properties of glutamine synthetase from Micrococcus glutamicus: effect of nitrogen sources in culture medium on enzyme formation and some properties of crystalline enzyme. Agric. Biol. Chem. 45:1487–1492. 48. Takayama K, Abe S, and Kinoshita S. (1965) Taxonomical studies on glutamic acid producing bacteria part 1: On the taxonomical characters. [In Japanese] Nippon Nogeikagaku Kaishi 39:328–342.

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49. Tesch, M, Eikmanns BJ, de Graaf AA, and Sahm H. (1998) Ammonia assimilation in Corynebacterium glutamicum and a glutamate dehydrogenase-deficient mutant. Biotechnol. Lett. 20:953–957. 50. Tesch M, de Graaf AA, and Sahm H. (1999) In vivo fluxes in the ammoniumassimilatory pathways in Corynebacterium glutamicum studied by 15N nuclear magnetic resonance. Appl. Environ. Microbiol. 65:1099–1109. 51. Thomas G, Coutts G, and Merrick M. (2000) The glnKamtB operon: a conserved gene pair in prokaryotes. Trends Genet.16:11–14. 52. Tochikura T, Sung H-C, Tachiki T, and Kumagai H. (1984) Occurrence of glutamate synthase in Brevibacterium flavum. Agric. Biol. Chem. 48:2149–2150. 53. Trötschel C, Kandirali S, Diaz-Achirica P, Meinhardt A, Morbach S, Krämer R, and Burkovski A. (2003) GltS, the sodium-coupled L-glutamate uptake system of Corynebacterium glutamicum: Identification of the corresponding gene and impact on L-glutamate production. Appl. Microbiol. Biotechnol. 60:738–742. 54. Wakisaka S, Tachiki T, and Tochikura T. (1990) Properties of Brevibacterium flavum glutamine synthetase in an “in vivo-like” system. J. Ferm. Bioeng. 70:182–184. 55. Wehrmann A, Phillipp B, Sahm H, and Eggeling L. (1998) Different modes of diaminopimelate synthesis and their role in cell wall integrity: a study with Corynebacterium glutamicum. J. Bacteriol. 180:3159–3165.

15

Sulfur Metabolism and Its Regulation H.-S. Lee

CONTENTS 15.1 15.2 15.3 15.4

Introduction ..................................................................................................351 Assimilation of Sulfur..................................................................................352 Transport of Sulfate .....................................................................................357 Biosynthesis and Degradation of Sulfur-Containing Amino Acids ............358 15.4.1 Cysteine ............................................................................................358 15.4.2 Methionine .......................................................................................360 15.5 Regulatory Mechanisms...............................................................................366 15.5.1 Regulation of Cysteine Biosynthesis...............................................366 15.5.2 Regulation of Methionine Biosynthesis ..........................................366 15.5.3 Significance of the Parallel Pathways..............................................369 15.6 Constructing Methionine-Producing Strains ...............................................369 15.7 Conclusions and Perspectives ......................................................................370 References..............................................................................................................371

15.1 INTRODUCTION Sulfur is essential for microbial growth because it is a constituent of cysteine, methionine, iron-sulfur proteins, coenzyme A, lipoic acid, mycothiol, biotin, thiamine, and tRNA. Cysteine and methionine are the predominant sulfur-containing molecules within the cell. Usually, only oxidized sulfur compounds like sulfate are available for the cell, which have to be reduced to the level of sulfide before incorporation into the organic sulfur compounds (assimilatory reduction). Some organisms use inorganic sulfur compounds for energy metabolism. However, energygenerating metabolic processes involving sulfur, such as dissimilatory reduction and oxidation, have not been reported for Corynebacterium species, and there is also no indication from the genome sequence that such processes exist in C. glutamicum and C. efficiens. In this chapter, the present knowledge on the biosynthesis pathways of the sulfur-containing amino acids cysteine and methionine in C. glutamicum will be summarized, including the sulfur assimilation pathway as derived from genome sequence analysis. In addition, the regulatory mechanisms governing the expression of genes involved in cysteine and methionine biosynthesis will be presented. 351

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15.2 ASSIMILATION OF SULFUR Sulfur is abundant and widely distributed in nature. Its natural reservoirs include fossil fuels, elemental sulfur deposits, rocks, and minerals. Inorganic sulfur occurs naturally in three different oxidation states: 0 (S0, elemental sulfur), –2 (S2–, inorganic sulfide), and +6 (SO42–, sulfate). In the lithosphere, inorganic sulfur is largely present as sulfate and sulfide, such as CaSO4 and FeS. Less abundant forms of inorganic sulfur include thiosulfates (S2O32–), thionates (SnO62–), and elemental sulfur. In the aerobic biosphere, sulfate (SO42–) and its derivatives, such as thiosulfate, are the main forms of inorganic sulfur assimilated by bacteria, and, for this reason, sulfate is the most common source of inorganic sulfur in laboratory media. The sulfur content of C. glutamicum is 0.40 ± 0.01% (g/g dry weight) when grown in CGXII minimal medium [37] with glucose as a carbon source (see also Table 2.1). After uptake of sulfate into the cytoplasm by primary or secondary transport systems (see below), it must be reduced to sulfide. Since the redox potential of the sulfate/sulfite pair is very low (E′ο = –454 mV), sulfate has to be activated to adenosine5′-phosphosulfate (APS), which lowers the redox potential and allows reduction to sulfite by NADPH (E′ο = –320 mV) or its equivalent. C. glutamicum is likely to utilize ATP sulfurylase (EC 2.7.7.4) for the activation of sulfate to APS as the genes cysD and cysN encoding the two subunits of ATP sulfurylase are clearly identified in the genome sequence (Table 15.1 and Figure 15.1); CysD and CysN form a heterodimer. Whereas CysD catalyzes the formation of APS and PPi from ATP and sulfate, CysN functions as a GTPase catalyzing the hydrolysis of GTP to GDP and Pi. The hydrolysis of GTP is energetically coupled to ATP sulfurylation and is required to shift the equilibrium of this reaction in favor of the product APS [44,86]. Thus, the overall reaction catalyzed by ATP sulfurylase is as follows: ATP4– + SO42– + GTP4– + H2O → APS2– + PPi4– + GDP3– + Pi2– + H+. The next step, the formation of sulfite from APS, is likely to be catalyzed by APS reductase encoded by cysH. Unlike C. glutamicum, E. coli forms sulfite from APS via 3′-phosphoadenosine 5′phosphosulfate (PAPS) by a two-step process catalyzed by APS kinase (EC 2.7.1.25), encoded by cysC, and by PAPS reductase, encoded by cysH. The APS and PAPS reductases share high sequence identities and were thus given the identical gene name cysH. However, APS reductases contain the signature sequences CCXXRKXXPL and SXGCXXCT, both containing potentially mechanistically relevant cysteine residues, whereas PAPS reductases lack these motifs [90]. Since CysH of C. glutamicum exhibits these motifs, APS is expected to be reduced in one step to sulfite (Figure 15.1). In support of this assumption, no gene is present in the C. glutamicum genome whose product shows significant homology to APS kinase (CysC) of E. coli. However, the bypass of the requirement for the formation of PAPS in C. glutamicum has not been demonstrated experimentally. Originally identified in plants, APS reductase is present in several bacteria, including Sinorhizobium meliloti [1], Pseudomonas aeruginosa [6], Bacillus subtilis [90], and Mycobacterium species [90]. These reductases are thioredoxin-dependent [6] and the same presumably holds true for C. glutamicum APS reductase. Mycobacterium species possess both APS reductase and APS kinase. PAPS formed by APS kinase is used as substrate for sulfotransferases, which transfer

Cgl0870* Cgl1051* Cgl1473*

Cgl0216*

Cgl0214*

Cgl0213*

Cgl2816 (cysH)* Cgl2817 (cysI)* Cgl1998 (cysG)* Cgl0701* Cgl1220 (ssuD)*

ABC-type transporter, permease component ABC-type trasporter, periplasmic component ABC-type transporter, ATPase component Sulfate permease Sulfate permease Sulfate permease

Sulfate adenyltransferase subunit 1 Sulfate adenyltransferase subunit 2 APS reductase Sulfite reductase Siroheme synthase Thiosulfate sulfurtransferase Alkanesulfonate monooxygenase

Cgl2814 (cysN)*

Cgl2815 (cysD)*

Protein Function

ORF (Gene)2

518 579 462

304

268

279

231 561 250 301 381

304

433

Length (aa) Protein Function

APS reductase Nitrite/sulfite reductase Siroheme synthase Probable thiosulfate sulfurtransferase Sufonate monooxygenase

ATP sulfurylase subunit 2

C. perfringens M. tuberculosis Pseudomonas sp.

E. coli

CysA, ATP-binding component of sulfate permease A protein Probable sulfate permease Sulfate transporter Sulfate permease

Sulfate (thiosulfate) Transport E. coli CysU, sulfate/thiosulfate transport system permease T protein E. coli Molybdate-binding periplasmic protein

S. coelicolor S. coelicolor E. coli M. tuberculosis P. putida

E. coli

Sulfur Assimilation E. coli ATP sulfurylase subunit 1

Organism

Homologs3

TABLE 15.1 Corynebacterium glutamicum Genes and Candidate ORFs Involved in Sulfur Metabolism1

551 560 495

365

257

277

236 565 457 297 381

302

475

Length (aa)

35 51 51

37

31

30

45 54 38 60 50

46

47

Identity (%)

data data data data

Genome data Genome data Genome data

Genome data

Genome data

Genome data

Genome Genome Genome Genome 61

Genome data

Genome data

Source

Sulfur Metabolism and Its Regulation 353

Serine acetyltransferase Cysteine synthase Cysteine synthase Putative transcriptional regulator

Homoserine O-acetyltransferase Cystathionine γ-synthase Cystathionine β-lyase O-acetylhomoserine sulfhydrylase Methionine synthase I, B12-dependent Methionine synthase II, B12-independent

Cgl2563 (cysE)* Cgl2562 (cysK)* Cgl2136 (cysM)* Cgl2928*

Cgl0652 Cgl2446 Cgl2309 Cgl0653 Cgl1507

Cgl1139 (metE)

(metX) (metB) (aecD) (metY) (metH)

Protein Function

ORF (Gene)2

745

377 386 368 437 1,221

182 318 317 308

Length (aa)

coli coli coli coli

—

— — — — —

E. E. E. E.

Protein Function

—

Methionine — — — — —

Cysteine Serine acetyltransferase O-Acetylserine sulfhydrylase A O-Acetylserine sulfhydrylase B CysB, transcriptional regulator

Organism

Homologs3

TABLE 15.1 (continued) Corynebacterium glutamicum Genes and Candidate ORFs Involved in Sulfur Metabolism1

—

— — — — —

273 323 303 324

Length (aa)

—

— — — — —

40 53 31 25

Identity (%)

65

57 28 64 29 65

Genome Genome Genome Genome

data data data data

Source

354 Handbook of Corynebacterium glutamicum

Glycosyl transferase — — Acetyltransferase

Spermidine synthase

Cgl0401 (mshA)* Cgl1100 (mshB)* Cgl1514 (mshC)* Cgl2576 (mshD)*

Cgl2702 (speE)*

3

2

1

513

418 290 420 292

382

407 213 246 356 478

326

523

480 303 414 315

Mycothiol tuberculosis MshA glycosyl transferase (Rv0486) tuberculosis MshB deacetylase (Rv1170) tuberculosis MshC ligase (Rv2130) tuberculosis MshD acetyltransferase (Rv0819)

Polyamine M. tuberculosis Spermidine synthase

M. M. M. M.

— — — — 495

296

379

— — — — S-Adenosylhomocysteine hydrolase

5,10-Methylenetetrahydrofolate reductase

Cystathionine γ-lyase

B. subtilis

— — — — M. tuberculosis

E. coli

56

51 38 56 35

33

— — — — 74

33

data data data data

Genome data

Genome Genome Genome Genome

Genome data

22 61 Genome data 68 Genome data

65

Genes with known function (or ORFs with identifiable function) are listed in the table. The protein function and gene designation of the ORFs from the genome data are putative. Asterisks (*) indicate that the gene function is not yet proven. The listed homologs do not necessarily mean the one giving the highest similarity scores.

Cgl2786*

Cgl1603 (metK) Cgl2941 (mcbR) Cgl2445* Cgl1776 (cglIM) Cgl0752 (ahcY)*

5,10-Methylenetetra-hydrofolate reductase Methionine adenosyl-transferase McbR, transcriptional repressor Putative transcriptional regulator 5′-Cytosine methyltransferase S-Adenosylhomocysteine hydrolase Cystathionine γ-lyase

Cgl2171 (metF)*

Sulfur Metabolism and Its Regulation 355

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Sulfate (SO42-) Outside ATP Inside ADP

Sulfate ATP

ATP sulfurylase (cysD, cysN)

P-P ATP

APS

APS kinase (cysC)

ADP

Alkanesulfonate

Alkanesulfonate monooxygenase (ssuD )

APS reductase (cysH)

TRXRED

AMP

TRXOX

PAPS TRXRED TRXOX

Sulfite (SO32-)

PAPS reductase (cysH)

3NADPH + 3H+ Sulfite reductase (cysI)

3NADP+

Sulfide (H2S)

Cysteine, methionine biosynthesis

FIGURE 15.1 Proposed pathway of sulfate assimilation in C. glutamicum. The pathways have been constructed by utilizing the genome data. Reactions shown in gray are likely to be absent from C. glutamicum but present in E. coli. Instead of APS reductase, E. coli utilizes APS kinase and PAPS reductase. Abbreviations: APS, adenosine 5′-phosphosulfate; PAPS, 3′-phosphoadenosine 5′-phosphosulfate; TRX, thioredoxin.

sulfuryl groups from PAPS to other molecules, such as glycolipids [49]. The sulfated glycolipids are known as mediators of cell–cell contacts, such as host–pathogen interactions. In addition, in Mycobacterium species, CysN (GTPase) and CysC (APS kinase) form a single polypeptide encoded by a single gene cysNC [90]. The CysC part is located in the C-terminus of the CysNC protein. Although such configuration

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357

is not uncommon as in S. meliloti [71], P. aeruginosa [27], and Xanthomonas oryzae [74], such a feature is not found in C. glutamicum. The reduction of sulfite to sulfide is presumably catalyzed by sulfite reductase (EC 1.8.1.2; Figure 15.1). In the genome of C. glutamicum, a gene (cysI) is located upstream of cysH (Figure 15.7), which encodes a protein with clear similarity to the α-subunit of sulfite reductases. However, its involvement in sulfite reduction has not yet been proven. In E. coli, sulfite reductase is composed of two subunits encoded by cysJ (subunit α, a flavoprotein) and cysI (subunit β, a hemoprotein), which are part of the cysJIH operon [72]. The subunit structure is presumably α8β8 [94]. The α8 flavoprotein contains four FAD and four FMN, whereas each of the β-subunits contains an Fe4S4 cluster and a siroheme prosthetic group. Electron flow presumably proceeds as follows: NADPH → FAD → FMN → Fe4S4 → siroheme → sulfite. Remarkably, no gene encoding a CysJ homolog is found in the C. glutamicum genome, raising the question on the in vivo electron donor for sulfite reduction. In E. coli, the cysG gene product is involved in the synthesis of siroheme, which is also found in nitrite reductase [77]. A cysG homolog (Cgl1998) is found in the genome of C. glutamicum. The encoded CysG protein composed of 250 amino acids is significantly smaller than the E. coli counterpart composed of 457 amino acids. The sequence identity of the overlapping region with the C-terminal half of the E. coli protein is 38%. Utilization of other sulfur sources, such as thiosulfate and organic sulfur sources, is less well known even in E. coli. A gene (Cgl0701) that may encode thiosulfate sulfurtransferase (EC 2.8.1.1), catalyzing the breakage of the S–S bond present in thiosulfate into sulfur and sulfite, is found in the genome of C. glutamicum. However, as the physiological roles of sulfurtransferases found in many kinds of organisms are still in question, the involvement of the corynebacterial sulfurtransferase in sulfur metabolism is not clear. Alternatively, thiosulfate can be assimilated into cysteine through a different route (see following section for details). Since most of the sulfur in aerobic soils is present in organic form, such as sulfate esters, sulfonates, and sulfur-containing amino acids, there might be ways to utilize these compounds also by C. glutamicum. In other organisms, organic sulfur sources are generally converted to sulfate or sulfite by the action of appropriate enzymes, such as sulfatases and oxygenases [38,82]. Although no in-depth studies have been performed on C. glutamicum, some genes that might be involved in the metabolism of alkane sulfonates and organic sulfur compounds are found in the genome sequence. Rey et al. [61] reported on one of the genes that is a putative alkanesulfonate monooxygenase encoded by ssuD (Figure 15.1). Interestingly, the expression of ssuD is affected by the regulatory gene McbR, which governs the expression of several genes involved in the biosynthesis of methionine (see following section for details).

15.3 TRANSPORT OF SULFATE Scrutiny of the genome sequence of C. glutamicum reveals two types of possible sulfate transport systems: ABC (ATP-binding cassette) type of sulfate-thiosulfate transporters and sulfate permeases. In E. coli, the membrane components of the ABC carrier are encoded by cysU and cysW, whereas cysA encodes the ATPase component

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of the permease. This single transport system utilizes two periplasmic binding proteins encoded by cysP and sbp, which are specific for thiosulfate and sulfate, repectively [42,76]. Among the candidate ORFs of C. glutamicum, ORF Cgl0213, Cgl0214, and Cgl0216 show a maximum of 37% amino acid sequence identity with E. coli permease component, periplasmic component, and ATPase component, respectively (Table 15.1). However, since the ORFs are located near genes involved in molybdenum cofactor metabolism, the involvement of the encoded proteins in molybdenum and molybdate transport cannot be ruled out. In addition to the ATP-dependent transporter, genes that may encode secondary active transporters are also found. These include Cgl0870, Cgl1051, and Cgl1473 encoding proteins composed of 518, 579, and 462 amino acids, respectively (Table 15.1). Although the presence of such genes in the C. glutamicum genome looks interesting with respect that such genes are rarely found in the genome of E. coli and M. tuberculosis, the functional and physiological significance of the genes is unknown.

15.4 BIOSYNTHESIS AND DEGRADATION OF SULFUR-CONTAINING AMINO ACIDS 15.4.1 CYSTEINE The synthesis of cysteine and its subsequent use is the predominant route by which reduced sulfur is incorporated into cellular compounds. Cysteine is used for protein synthesis and it also donates its reduced sulfur to methionine, lipoic acid, thiamine, mycothiol, coenzyme A, or to other organic molecules. The cysteine biosynthetic pathway appears to be identical in many different microorganisms [42,80] and occurs via a two-step process under conditions of sulfide availability (Figure 15.2). The first step is the acetylation of serine to O-acetylserine catayzed by serine acetyltransferase (EC 2.3.1.30) encoded by the cysE gene. The C. glutamicum CysE protein shows 40% sequence identity to CysE from E. coli (Table 15.1). The next step is catalyzed by O-acetylserine sulfhydrylase A encoded by the cysK gene, which is located immediately upstream of cysE, or by O-acetylserine sulfhydrylase B encoded by the cysM gene, which is located elsewhere on the chromosome (Figure 15.7). The CysK and CysM proteins from C. glutamicum show 53% and 31% sequence identity to the corresponding proteins from E. coli. The protein CysM from E. coli can also use thiosulfate instead of sulfide as sulfur donor leading to the synthesis of S-sulfocysteine, which may subsequently be converted to cysteine by sulfonatases [72]. In C. glutamicum, neither the kinetic parameters of CysK and CysM nor their specific functions have been determined. However, both contain binding motifs for pyridoxalphosphate (PLP) and therefore are most likely PLP-dependent enzymes like their E. coli counterparts. The protein CysK was clearly identified in proteome studies as a dominant protein of C. glutamicum [25,69] and recently also shown to be a phosphoprotein [5], in accordance with the presence of PLP in this enzyme. Degradation of cysteine into pyruvate, H2S, and NH3 is possible in C. glutamicum by cysteine desulfhydrase (Figure 15.2) encoded by the aecD gene. The AecD gene product was characterized as a C-S lyase with α,β-elimination activity [64]. As

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FIGURE 15.2 Biosynthesis and degradation of cysteine in C. glutamicum.

expected, disruption of aecD stimulates cysteine production by C. glutamicum [84]. The AecD protein is also involved in methionine biosynthesis owing to its cystathionine β-lyase activity, and this appears to be the natural function of the protein, and therefore it is also termed MetC [39,65] (see following section for details). Cysteine can be utilized for the biosynthesis of a low-molecular-mass thiol, mycothiol, which in actinomycetes is the functional replacement of glutathione. Like glutathione of Gram-negative bacteria, mycothiol is a mediator of the cellular redox homoeostasis and plays an important role in protecting cells from oxidative stress [54]. In addition, mycothiol is associated with the protection of mycobacteria from harmful thiol-reactive substances, such as formaldehyde and antibiotics. Mycothiol consists of N-acetyl-L-cysteine linked to a pseudodisaccharide, D-glucosamine, and myo-inositol [52]. Although the biosynthesis of mycothiol has not been studied in C. glutamicum, the biosynthetic route appears to be identical to that of Mycobacterium species, as highly homologous genes are identified in C. glutamicum (Table 15.1). In Mycobacterium species, the biosynthesis of mycothiol proceeds through a four-step pathway [3,10,54], and involves at least four enzymes MshA, MshB, MshC, and MshD. First, the formation of 1-D-myo-inosityl-2-acetamido-2deoxy-α-D-glucopyranoside from as yet unidentified precursors is catalyzed by MshA glycosyltransferase [55], followed by deacetylation by MshB deacetylase [53]. The resulting 1-D-myo-inosityl-2-amino-2-deoxy-α-D-glucopyranoside is ligated with

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a cysteine and the reaction is catalyzed by MshC ligase [67]. Finally, MshD acetylase catalyzes the acetylation of 1-D-myo-inosityl-2-(L-cysteinyl)amido-2-deoxy-α-D-glucopyranoside to produce mycothiol (1-d-myo-inosityl-2-(N-acetyl-L-cysteinyl)amido2-deoxy-α-D-glucopyranoside) [40]. The encoded protein products of ORFs Cgl0401, Cgl1100, Cgl1514, and Cgl2576 from C. glutamicum show protein sequence identities of 51%, 38%, 56%, and 35% to MshA, MshB, MshC, and MshD from M. tuberculosis, respectively (Table 15.1).

15.4.2 METHIONINE Methionine plays many important biological roles. Methionine is not only required for protein synthesis, but also involved in methylation reactions, in polyamine biosynthesis and in quorum sensing, for instance [72]. Methionine belongs to the aspartate family of amino acids, which are derived from oxaloacetate, a TCA cycle intermediate. In C. glutamicum, aspartate is converted to aspartylphosphate, aspartate semialdehyde, and homoserine, by consecutive reactions catalyzed by aspartate kinase, aspartate semialdehyde dehydrogenase, and homoserine dehydrogenase, respectively. Homoserine then serves as the common precursor of isoleucine, threonine, and methionine synthesis. In C. glutamicum, the first specific step of methionine biosynthesis, the acylation of homoserine to O-acetylhomoserine, is catalyzed by homoserine acetyltransferase (EC 2.3.1.31), which is encoded by metX (formerly metA) (Figure 15.3). The metX gene of C. glutamicum, which was islolated by the complementation of an E. coli metA mutant, codes for a polypeptide composed of 379 amino acids with molecular weight of 41,380 [57]. Its deduced protein sequence shows high amino acid sequence identity with other homoserine acetyltransferases encoded by metX (e.g., 48% identity with M. tuberculosis metX). The purified enzyme uses acetyl-CoA as the acyl donor, which cannot be replaced by succinyl-CoA [46]. The Km values for homoserine and acetyl CoA are 2.8 and 0.05 mM, respectively [46]. The kinetic parameters derived from crude extract determinations are comparable [33]. The activity of the enzyme regulates the flux of homoserine into multiple biosynthetic pathways and, therefore, represents a critical control point for cell growth and viability (see below). Unlike C. glutamicum, many microorganisms including E. coli utilize O-succinylhomoserine, and not O-acetylhomoserine as the acylated homoserine intermediate for methionine biosynthesis. In this case, homoserine succinyltransferase (EC 2.3.1.4), encoded by metA, is responsible for the synthesis of O-succinylhomoserine. The low homology between C. glutamicum MetX and E. coli MetA (27% identity) may reflect such difference. Subsequent formation of homocysteine from O-acetylhomoserine can be accomplished through two different routes in C. glutamicum: a transsulfuration pathway via cystathionine utilizing cysteine as the sulfur donor and a direct sulfhydrylation pathway utilizing sulfide as the sulfur donor (Figure 15.3). In the transsulfuration pathway, O-acetylhomoserine is converted to cystathionine by cystathionine γ-synthase (EC 2.5.1.48), the product of metB. Unlike cystathionine γ-synthases from E. coli and S. typhimurium, which prefer the use of succinylated substrates, the

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361

FIGURE 15.3 Methionine biosynthetic pathways in C. glutamicum. Abbreviation: THF, tetrahydrofolate. Symbols: *, transsulfuration pathway; #, direct sulfhydrylation pathway.

corresponding enzyme from C. glutamicum prefers acetylated substrates [28,29,57]. The corynebacterial metB gene, isolated by the complementation of an E. coli metB mutant strain, encodes a protein composed of 386 amino acids with a molecular weight of 41,655 [28], and the putative protein product shows in part very high amino acid sequence identity to its counterparts in other organisms, e.g., 63% identity with M. tuberculosis MetB. However, as with the metX gene product, the identity between the MetB proteins of C. glutamicum and E. coli is relatively low (41%). In all cases investigated, including E. coli [26], cystathionine γ-synthase (MetB) was found to be a tetramer of identical or closely related 40- to 50-kDa subunits with one PLP cofactor bound per monomer. Although the native molecular weight of the corynebacterial MetB is unknown, conserved motifs for PLP binding are found in the amino acid sequence of MetB (Figure 15.4). In the transsulfuration pathway, cystathionine is further hydrolyzed to homocysteine, pyruvate, and ammonia by cystathionine β-lyase (EC 4.4.1.8, Figure 15.3) encoded by aecD (also known as metC). The aecD gene was previously isolated by its ability to confer resistance to S-(2-aminoethyl)-L-cysteine (AEC), a toxic lysine analog [64] that is structurally analogous to cystathionine. Later, a C. glutamicum

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Consensus: MetB:

195

AecD::

307

MetY:

207

[DQ]-[LIVMF] -x-x-x-[STAGC] -[STAGCI] -T-K-[FYWQ]-[LIVMF] -x-G-[HQ]-[SGNH]

H A VLH S T M W LDF A G D V VVA S L

TK Y I GGH S TK I E EAP S TK F Y TGN G ∗

176 46 215

FIGURE 15.4 Binding motifs of pyridoxal 5′-phosphate (PLP) in three enzymes of methionine biosynthesis of C. glutamicum. PLP binds to the lysine (K) residue, which is marked with an asterisk.

gene, isolated by complementation of an E. coli metC mutant, turned out to be identical to the previously isolated aecD, suggesting that the natural function of aecD, encoding a C-S lyase, is probably mediating methionine biosynthesis [39]. In support of this, deletion of aecD in the metY background (see below) results in methionine auxotrophy in C. glutamicum [65]. Wada et al. [84] purified the enzyme and confirmed the enzymatic activities as a C-S lyase. The aecD gene encodes a protein composed of 368 amino acids with a molecular weight of 40,735. It shows only low sequence identity to MetC proteins of other organisms, e.g., 26% with E. coli MetC. The cystathionine β-lyase from E. coli is composed of six identical subunits of 43 kDa [17]. Each monomer contains a firmly bound PLP prosthetic group as a cofactor. The enzyme of C. glutamicum requires PLP for maximal activity [34] and exhibits structural features of PLP-binding enzymes (Figure 15.4). In accordance with this, the enzyme requires PLP for maximal activity [34]. Corynebacterial cystathionine β-lyase (AecD) uses cystathionine, AEC, cysteine, and cystine as substrates [64,84]. Among the β C-S bond containing substrates, cysteine is a poorer substrate than cystathionine or cystine. The Km value of cystathionine β-lyase for L-cystathionine is 5 mM [34]. As mentioned in the previous section, it also possesses cysteine desulfhydrase activity, which is important for degrading cysteine to pyruvate, H2S, and NH3 [84]. C. glutamicum can synthesize homocysteine from O-acetylhomoserine also by direct sulfhydrylation with sulfide [29]. This reaction is catalyzed by O-acetylhomoserine sulfhydrylase (EC 2.5.1.49) encoded by the metY gene (Figure 15.3). Because of the presence of parallel pathways, a single deletion of metB, metC, or metY does not result in methionine auxotrophy [29,39,64,65,84], whereas a double deletion of metB and metY (or aecD and metY) causes methionine auxotrophy [29,65]. In C. glutamicum, both methionine biosynthetic pathways may function equally well because the growth pattern of the metY mutant strain is identical to that of the metB mutant strain [29]. The metY gene of C. glutamicum is located immediately upstream of metX (Figure 15.7) and encodes a protein of 437 amino acids with a molecular weight of 46,751. The metX and metY genes are separated by 143 bp, and are expressed independently; a putative rho-independent transcriptional stop signal is found at the downstream region of the metY gene and deleting the metY gene does not affect the metA expression [29]. The molecular weight of native O-acetylhomoserine sulfhydrylase (MetY) from C. glutamicum has been reported

Sulfur Metabolism and Its Regulation

363

to be 360,000 [56], whereas in other organisms the enzyme is usually composed of four subunits and has a molecular weight of approx. 200,000 [93]. C. glutamicum MetY requires PLP as cofactor [29,56] and does not use O-succinylhomoserine in place of O-acetylhomoserine as substrate [56]. In terms of utilizing sulfide as a substrate, the enzyme is functionally homologous to O-acetylserine sulfhydrylase. However, its activity with O-acetylserine as substrate is almost negligible [56]. The Km values of O-acetylhomoserine sulfhydrylase for H2S and O-acetylhomoserine, which were determined with partially purified protein, are 0.08 mM and 2.0 mM, respectively. Recently, Bendt et al. [5] reported that MetY is a phosphoprotein, reflecting the binding of PLP to the enzyme (Figure 15.4). The final step, the methylation of homocysteine to methionine, is catalyzed by methionine synthase (Figure 15.3). In C. glutamicum, two forms of methionine synthase are present, a B12-independent enzyme encoded by metE and a B12-dependent enzyme encoded by metH [65]. The methyl group donors of C. glutamicum MetE and MetH have not yet been determined. In E. coli, MetH utilizes N5-methyltetrahydrofolate or its polyglutamyl derivative as substrate, whereas MetE utilizes N5-methyl-tetrahydropteroyl-triglutamate. A C. glutamicum metE deletion mutant was methionine auxotrophic on solid media, but not in liquid culture or on solid media supplemented with vitamin B12. Rückert et al. [65] speculated that this phenotype might be caused by the fact that the amount of vitamin B12 synthesized under aerobic conditions by C. glutamicum is not sufficient to compensate for the loss of cob(I)alamin by oxidation. These authors therefore suggested that MetH could be used during microaerobic growth and MetE under aerobic growth. In a proteome analysis of C. glutamicum grown aerobically in glucose minimal medium, the MetE protein was clearly identified [69]. Moreover, a recent phosphoproteome analysis of C. glutamicum indicated that MetE is a phosphoprotein [5]. The physiological significance of a phosphorylation has not yet been clarified, however. The methyl tetrahydrofolate substrate is formed by the consecutive action of serine hydroxymethyltransferase (glyA gene product) and N5,N10-methylene tetrahydrofolate reductase (metF gene product) (Figure 15.3). An additional step, the conversion of methionine to S-adenosylmethionine (SAM), is catalyzed by SAM synthase (Figure 15.3) encoded by the metK gene. The metK gene of C. glutamicum has a size of 1,224 bp and encodes a protein with molecular weight of 44.2 kDa [22]. Protein MetK shows 57% sequence identity to the MetK protein of E. coli. The metK transcript was only detected in cells of the logarithmic growth phase, but not in cells of the stationary phase, indicating a strict transcriptional regulation of metK [22]. SAM is an essential molecule and plays important roles in the regulation of methionine biosynthesis, in methylation reactions, and in polyamine biosynthesis, for instance [21,66,72,80]. Among the many methyltransferases found in the genome of C. glutamicum, Schaffer et al. [68] identified the cglIM gene to be responsible for the methylation of cytosine residues in DNA. The encoded 5′-cytosine methyltransferase protein contains highly conserved motifs, found in other 5′-cytosine methyltransferases, one of which presumably constitutes the SAM-binding site. Among the genes involved in polyamine biosynthesis, such as speA (arginine decarboxylase), speB (agmatine ureohydrolase), speC (ornithine decarboxylase), speD (SAM decarboxylase), and speE (spermidine

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FIGURE 15.5 Degradation of methionine in C. glutamicum. Transaminase and methionine γ-lyase activities have been reported, but not the identification of the corresponding genes. The formation of methanesulfonate is speculative.

synthase) of E. coli, only the speE (Cgl2702) homolog is identified in the C. glutamicum genome (Table 15.1). Regarding the degradation of methionine, evidence was provided recently that C. glutamicum can convert L-methionine to α-ketobutyrate, methanethiol (CH3SH), and ammonia by the activity of L-methionine γ-lyase (Figure 15.5). Alternatively, a two-step pathway may be used involving a transaminase converting α-ketoglutarate and L-methionine to L-glutamate and α-keto-γ-methyl-thiobutyrate and an unknown enzyme converting α-keto-γ-methyl-thiobutyrate to methanethiol and α-ketobutyrate [9]. The fate of methanethiol is unknown and so is the question of how C. glutamicum utilizes methionine as a sole sulfur source. In P. putida, methanethiol is converted via an unknown pathway to methanesulfonate, which is then desulfonated to yield sulfite [83]. Presence of the ssuD gene encoding methanesulfonate monooxygenase in the C. glutamicum genome [61] may suggest the similarity to P. putida in the degradation of methanethiol to sulfite, as shown in Figure. 15.5. Some bacteria possess the so-called reverse transsulfuration pathway to enable the use of methionine for cysteine synthesis. In P. aeruginosa, methionine that is provided as the sole sulfur source is converted to cysteine via demethylation to homocysteine and subsequent reverse transsulfuration via cystathionine (by cystathionine

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β-synthase and cystathionine γ-lyase) [23,83]. In Saccharomyces cerevisiae and Streptomyces venezuelae, cystathionine is generated by cystathionine β-synthase, which catalyzes the condensation of homocysteine and serine [13,15]. Although cysteine is mainly synthesized by the thiolation of O-acetylserine in S. venezuelae, the interconversion of preexisting sulfur-containing amino acids by reverse transsulfuration appears to have a supplementary role. Although no experimental evidence is available, Rückert et al. [65], basing on the genome data, suggested a possibility for the presence of the reverse transsulfuration pathway in C. glutamicum. Figure 15.6 shows the putative reverse transsulfuration reactions of C. glutamicum.

Methionine S-Adenosylmethionine synthase (metK)

S-Adenosyl methionine Various methyltransferases

S-Adenosylhomocysteine S-Adenosylhomocysteine hydrolase (ahcY)

Homocysteine Cystathionine β-synthase

Cystathionine Cystathionine γ-lyase

Cysteine FIGURE 15.6 The proposed pathway of reverse transsulfuration in C. glutamicum. The pathways have been constructed by utilizing genome data.

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15.5 REGULATORY MECHANISMS 15.5.1 REGULATION

OF

CYSTEINE BIOSYNTHESIS

In microorganisms, the process of sulfur assimilation and cysteine biosynthesis is tightly regulated by the end-product cysteine and the intermediate O-acetylserine [12,42,51,80]. The biosynthesis of cysteine in E. coli ceases almost entirely when the cells are grown on cysteine or cystine as a result of the feedback inhibition of CysE by cysteine. The work by Wada et al. [84] suggests similar pattern of regulation in C. glutamicum. Indeed, feedback inhibition–insensitive CysE is necessary for the overproduction of cysteine in C. glutamicum. In E. coli, N-acetylserine, an isomerization product of O-acetylserine, together with the transcriptional activator CysB, which is a LysR-type regulater, induce the genes involved in sulfate reduction. A CysB homolog (Cgl2928) is found in the genome of C. glutamicum. Although the number of amino acids in the corynebacterial protein (308) is comparable to that of E. coli (324), the identity of the amino acid sequence is low (25%), requiring further investigations for clarification of a functional relationship. Recently, a corynebacterial repressor protein McbR was identified and found to regulate several genes involved in the biosynthesis of methionine and cysteine, such as cysK [61]. The presence of McbR and regulatory mechanism by the protein may suggest that the regulatory mechanism of cysteine biosynthesis in C. glutamicum may be different from that of E. coli.

15.5.2 REGULATION

OF

METHIONINE BIOSYNTHESIS

In general, methionine biosynthesis is tightly regulated at the gene expression and enzyme activity levels, and the regulatory mechanisms are also closely associated with sulfur assimilation. Because methionine belongs to the aspartate family of amino acids, its biosynthesis is partly controlled by other members of the family, such as lysine and threonine. Aspartate kinase plays an important role in the control of the biosynthesis of amino acids of the aspartate family. The asparte kinase of C. glutamicum is almost completely feedback inhibited by excess amounts of lysine and threonine (Table 15.2). Regulation at the aspartate semialdehyde branch is also important in the biosynthesis of methionine. Homoserine dehydrogenase, catalyzing the conversion of aspartate semialdehyde to homoserine and leading to the biosynthesis of threonine and methionine, is inhibited by threonine. Therefore, the overall biosynthesis of the aspartate family of amino acids can be controlled at the enzyme activity level by aspartate kinase and homoserine dehydrogenase. Additionally, the genes hom (homoserine dehydrogenase) and thrB (homoserine kinase), which constitute an operon and are involved in threonine biosynthesis, are repressed by methionine (Table 15.2). This may suggest that methionine is preferentially synthesized to threonine. In support of this, the repression of homoserine kinase, the first enzyme for threonine biosynthesis, by methionine, is not so strong as that of homoserine acetyltransferase, the first enzyme for methionine biosynthesis (see below) [46]. In C. glutamicum, methionine biosynthesis is highly regulated by inhibition and repression (Table 15.2). Regarding the control of methionine biosynthesis at the

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TABLE 15.2 Regulation of Genes and Enzymes Involved in Methionine Biosynthesis in C. glutamicum Enzyme Aspartate kinase Aspartate semialdehyde dehydrogenase Homoserine dehydrogenase Homoserine acetyltransferase Cystathionine γ-synthase Cystathionine β-lyase O-acetylhomoserine sulfhydrylase Methionine adenosyltransferase

Mw (Da) Gene

Operon

Inhibition

44,300 18,584 36,275

lysCα lysCαβasd Lysine plus threonine lysCβ asd lysCαβasd

46,136

hom

41,380

metX

41,655

metB

40,735

aecD

46,751

metY

44,200

metK

homthrB

Threonine

Repression

Literature

—

32, 78 31

Methionine

Methionine, SAM, Methionine, O-acetylhomoserine SAM, cysteine SAM Methionine Methionine, cysteine, Methionine glycine Methionine, Methionine O-acetylsereine Methionine

16, 19, 59, 60, 33, 35, 57, 75 28, 34 34, 39, 56, 64, 84 29, 75 35

level of enzyme activity, there is clear evidence that the MetX and MetY proteins are feedback inhibited by the end product methionine. In particular, the inhibition of MetX by methionine or SAM is labile and dependent on the presence of sulfhydryl compounds, such as dithiothreitol or cysteine, in the assay mixture. The concentrations of methionine and SAM required to give 50% inhibition of the enzyme activity are 4.8 and 0.26 mM, respectively. Unlike the inhibition by SAM, the inhibition of MetA by methionine may not be physiologically meaningful, as assessed by the concentration of methionine required for inhibition. Unlike E. coli, no synergistic action between methionine and SAM has been observed in C. glutamicum. O-Acetylhomoserine, which is a reaction product of MetX, also inhibits the activity by 50% at 10 mM. MetB of C. glutamicum is inhibited by SAM (50% inhibition by 3 mM SAM), but not by methionine. However, AecD is not inhibited by methionine or SAM, but is inhibited by cysteine (83% inhibition by 10 mM cysteine) and glycine (45% inhibition by 5 mM glycine). The physiological meaning of the inhibition by glycine is not clear. MetY of C. glutamicum is not inhibited by SAM or cystathionine, but by methionine (50% inhibition by 10 mM methionine) and by O-acetylserine to some extent (30% inhibition by 10 mM methionine). Under the situation in which MetX is highly inhibited by SAM, the inhibition of MetY by methionine may not be physiologically meaningful. In addition, a temperature sensitivity of corynebacterial MetX has been sugested [29,43]. In organisms like E. coli, Aerobacter aerogenes, Klebsiella pneumoniae, Serratia marcescens, S. typhimurium, and B. polymyxa, the MetA proteins (equivalent to MetX of C. glutamicum) are inactivated at

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higher temperatures [24,62,63,91]. Due to this, their growth rates at elevated temperatures are controlled by the limiting availability of endogenous methionine. As the MetA protein of E. coli has been suggested to play a role in triggering the heat shock response [7,8], MetX of C. glutamicum may perform a similar function. In the control of methionine biosynthesis at the level of gene expression (Table 15.2), the expression of metX, metB, aecD, and metY genes are repressed by methionine. The synthesis of MetY is more strongly repressed by methionine than the synthesis of MetB. Addition of 0.1 mM methionine to the growth medium reduces the MetY activity by 80%, whereas that of MetB is reduced by 50%. In addition, MetY is also highly expressed in the absence of methionine, although the physiological meaning of such a phenomenon is unclear. In addition, like the O-succinylhomoserine sulfhydrylase of P. aeruginosa [18], no methionine or cysteine residues are found in the coding region of C. glutamicum metY, suggesting that amino acids or compounds containing sulfur may regulate metY expression. The repression of aecD by methionine is relatively weak compared with the repression of metX, metB, and metY [34,56]. Recently, a transcriptional regulator for genes involved in sulfur-containing amino acids has been isolated from C. glutamicum [61]. The putative transcriptional repressor McbR, which was identified among the proteins binding to the promoter region of metY, is composed of 213 amino acids and has a molecular weight of 23,743. The N-terminal region (from amino acid 21 to 60) of the deduced polypeptide shows a multihelical DNA-binding domain known to be characteristic for the TetR family of transcriptional regulators. Apparently, it functions as a transcriptional repressor by acting on several genes involved in the biosynthesis of amino acids methionine and cysteine. In an mcbR mutant strain, genes for methionine biosynthesis, such as metY and metK, and genes for sulfur assimilation and cysteine biosynthesis, such as cysK and cysI, are derepressed as identified by two-dimensional PAGE analysis. In addition, proteins encoded by hom (homoserine dehydrogenase) and ssuD (alkanesulfonate monooxygenase) are more abundant in the mcbR deletion mutant strain. However, the substrate that modulates the McbR activity is unknown, although a substance that appears in the cell when methionine is limiting may play such a role [61]. Since only a few met genes are repressed by the McbR protein, there must exist additional regulators for other met genes of C. glutamicum. In E. coli, MetJ, an aporepressor that binds SAM for activation, is responsible for the repression of all met genes, except for metH [21,87,89]. In E. coli, the metJ and metB genes are adjacent to each other and transcribed divergently. In C. glutamicum, an ORF (Cgl2445) encoding a Tet-R type of regulator is present adjacent to metB, although the establishment of a functional relationship with metJ of E. coli requires further investigation. A positive regulator MetR is involved in the expression of metE and metH of E. coli [81]. MetR also activates the metA gene, in which homocysteine plays a negative role [45]. Homologs of MetJ and MetR cannot be identified in the genome of C. glutamicum by homology-based search. With the exception of metX and metY, the genes involved in methionine biosynthesis are evenly scattered over the entire chromosome of C. glutamicum (30; Figure 15.7).

Sulfur Metabolism and Its Regulation

369

Cgl2928

metX metY

metH

metE

metK

ahcY

0

2.0 Mb

ssuD

Cgl0701

Cgl0213 Cgl0214 Cgl0216

Cgl0870 Cgl1051

mcbR

3.0 Mb

mshC

mshB

Cgl2786

speE

cglIM

1.0 Mb

mshA

metF aecD Cgl2445 metB

mshD

cysG cysM

cysK cysE

cysN cysD cysH cysI

Cgl1473

FIGURE 15.7 Physical map of C. glutamicum genes and candidate ORFs involved in sulfur metabolism. Gene designations for some ORFs are putative (see Table 15.1). The involvement of some ORFs in sulfur metabolism is not obvious.

15.5.3 SIGNIFICANCE

OF THE

PARALLEL PATHWAYS

Genetic and biochemical data, such as growth analysis with mutant cells and enzymatic activities under different growth conditions, suggest that C. glutamicum utilizes both transsulfuration and direct sulfhydrylation pathways with almost equal efficiency [29]. In addition, the metY gene of C. glutamicum is efficiently expressed [29]. The stronger repression of MetY by methionine than that of MetB suggests a distinctive role for the direct sulfhydrylation pathway. By analogy with the biosynthetic pathways for lysine in C. glutamicum, where the succinylase or dehydrogenase pathways are utilized depending on the availability of ammonium ions [70], one could speculate about a similar role for the methionine biosynthetic pathways in terms of sulfur availability. In organisms utilizing both transsulfuration and direct sulfhydrylation pathways, such as Schizosaccharomyces pombe, N. crassa, Aspergillus nidulans, and higher plants, the direct sulfhydrylation pathway (Figure 15.3) plays a minor role in methionine biosynthesis [11,20,58,88,93]. As evidenced by radioisotopic studies, the majority of homocysteine sulfur atoms are derived from cysteine via transsulfuration in these organisms. However, in P. aeruginosa and P. putida, the direct sulfhydrylation pathway appears to be favored over transsulfuration [83] because, under most growth conditions, cysteine is not converted to methionine via cystathionine and homocysteine. In addition, the enzymes catalyzing the transsufuration pathway, such as cystathionine γ-synthase and cystathionine β-lyase, are only expressed when cysteine is supplied as a sole sulfur source.

15.6 CONSTRUCTING METHIONINE-PRODUCING STRAINS Several methods have been described to produce L-methionine, namely, the enzymatic production of methionine from hydantoin [85], the chemical synthesis of D,Lmethionine from acrolein and methyl mercaptan via 3′-methylthiopropionaldehyde,

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its continuous production in immobilized cell or enzyme reactors [73], and its production by fermentation using analog-resistant mutant strains [35,36,47]. Because of the complexities of its biosynthetic pathways and metabolic regulation, the fermentative production of methionine using analog-resistant mutant strains has been only marginally successful, up to now. Using such a technique, Kase and Nakayama [36] isolated a C. glutamicum mutant strain that produced 250 mg/l of methionine. By successive mutagenesis, a mutant resistant to ethionine, selenomethionine, and methionine hydroxamate was developed that produced 2 g l–1 of methionine [35]. In this strain, the methionine biosynthetic genes are derepressed. Ethionine and selenoethionine are known to inhibit MetX (homoserine acetyltransferase) [75], but a direct inhibition of MetX activity is not confirmed. Mondal and Chatterjee [47] reported on an ethionine-resistant strain of Brevibacterium heali that produced 13 g l–1 of methionine. The mutation method was also applied to obligate and facultative methylotrophs, and resulted in mutant strains producing up to 800 mg l–1 methionine [48,92]. In an obligate methylotrophic mutant strain, MetA (homoserine succinyltransferase) is no longer feedback inhibited by SAM. A mutant strain of E. coli resistant to a threonine analog and ethionine was found to produce 2 g/l of methionine and threonine [14]. More recently, Nakamori et al. [50] isolated an analog-resistant E. coli strain producing 910 mg l–1 of methionine, which was attributed to a point mutation in the MetJ protein, causing derepression of the methionine biosynthetic genes.

15.7 CONCLUSIONS AND PERSPECTIVES Sulfur is an essential component of cells. Many genes are directly and indirectly associated with sulfur metabolism, and the understanding of sulfur metabolism is crucial for the production of the sulfur-containing amino acids methionine and cysteine. However, despite their importance, the metabolic pathways involving sulfur in C. glutamicum are largely unknown. Although the main features of sulfur metabolism are known from experimental and genome data, a number of aspects, such a sulfite reduction, recycling of sulfur compounds, and regulatory mechanisms, deserve further study. The limited success achieved at isolating methionine-producing C. glutamicum strains probably reflects the complexities of the regulatory networks that govern sulfur-containing molecules. However, the recent establishment of a competitive cysteine production process based on an engineered E. coli strain illustrates that in principle such regulations can be overcome. C. glutamicum is distinguished from other microorganisms because it utilizes parallel biosynthetic pathways for methionine. The two pathways involved are mediated by independent genes and enzymes, and tightly regulated by methionine. Although the physiological significance of the two pathways is unknown, the presence of such duplication provides some insight into the metabolic flexibility and potential efficiency of this organism, and proffers C. glutamicum as a candidate for the construction of methionine-producing strains.

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371

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Sareen D, Steffek M, Newton GL, and Fahey RC. (2002) ATP-dependent L-cysteine: 1D-myo-inosityl 2-amino-2-deoxy-α-D-glucopyranoside ligase, mycothiol biosynthesis enzyme, MshC, is related to Class I cysteinyl-tRNA synthetases. Biochemistry 41:6885–6890. Schäffer A, Tauch A, Droste N, Pühler A, and Kalinowski J. (1997) The Corynebacterium glutamicum cglIM gene encoding a 5′-cytosine methyltransferase enzyme confers a specific DNA methylation pattern in an McrBC-deficinet Escherichia coli strain. Gene 203:95–101. Schaffer S, Weil B, Nguyen VD, Dongmann G, Günther K, Nickolaus M, Hermann T, and Bott M. (2001) A high-resolution reference map for cytoplasmic and membraneassociated proteins of Corynebacterium glutamicum. Electrophoresis 22:4404–4422. Schrumpf B, Schwarzer A, Kalinowski J, Pühler A, Eggeling L, and Sahm H. (1991) A functionally split pathway for lysine synthesis in Corynebacterium glutamicum. J. Bacteriol. 173:4510–4516. Schwedock JS, Changxian L, Leyh TS, and Long SR. (1994) Rhizobium meliloti NodP and NodQ form a multifunctional sulfate-activating comples requiring GTP for activity. J. Bacteriol. 176:7055–7064. Sekowska A, Kung H-F, and Danchin A. (2000) Sulfur metabolism in Escherichia coli and related bacteria: facts and fiction. J. Mol. Microbiol. Biotechnol. 2:145–177. Sharma S and Gomes J. (2001) Effect of dissolved oxygen on continuous production of methionine. Eng. Life Sci. 1:69–73. Shen Y, Sarma P, da Silva FG, and Ronald P. (2002) The Xanthomonas oryzae pv. oryzae raxP and raxQ genes encode and ATP sulfurylase and adenosine-5′-phosphosulphate kinase that are required for AvrXaq21 avirulence activity. Mol. Microbiol. 44:37–48. Shiio I and Ozaki H. (1981) Feedback inhibition by methionine and S-adenosylmethionine, and desensitization of homoserine O-acetyltransferase in Brevibacterium flavum. J. Biochem. 89:1493–1500. Sirko A, Zatyka M, Sadowy E, and Hulanicka D. (1995) Sulfate and thiosulfate transport in Escherichia coli K-12: evidence for a functional overlapping of sulfateand thiosulfate-binding proteins. J. Bacteriol. 177:4134–4136. Stroupe ME, Leech HK, Daniels DS, Warren MJ, and Getzoff ED. (2003) CysG structure reveals tetrapyrrole-binding features and novel regulation of siroheme biosynthesis. Nat. Struct. Biol. 10:1064–1073. Thierbach G, Kalinowski J, Bachmann B, and Pühler A. (1990) Cloning of a DNA fragment from Corynebacterium glutamicum conferring amino ethyl cysteine resistance and feedback resistance to aspartokinase. Appl. Microbiol. Biotechnol. 32:443–448. Thomas D and Surdin-Kerjan Y. (1997) Metabolism of sulfur amino acids in Saccharomyces cerevisiae. Microbiol. Mol. Biol. Rev. 61:503–532. Umbarger HE. (1978) Amino acid biosynthesis and its regulation. Annu. Rev. Biochem. 47:533–606. Urbanowski ML, Stauffer LT, Plamann LS, and Stauffer GV. (1987) A new methionine locus, metR, that encodes a trans-acting protein required for activation of metE and metH in Escherichia coli and Salmonella typhimurium. J. Bacteriol. 169:1391–1397. van der Ploeg JR, Eichhorn E, and Leisinger T. (2001) Sulfonate-sulfur metabolism and its regulation in Escherichia coli. Arch. Microbiol. 176:1–8. Vermeij P and Kertesz MA. (1999) Pathways of assimilative sulfur metabolism in Pseudomonas putida. J. Bacteriol. 181:5833–5837.

376 [84]

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Handbook of Corynebacterium glutamicum Wada M, Awano N, Haisa K, Takagi H, and Nakamori S. (2002) Purification, characterization and identification of cysteine desulfhydrase of Corynebacterium glutamicum, and its relationship to cysteine production. FEMS Microbiol. Lett. 217:103–107. Wagner T, Hantke B, and Wagner F. (1996) Production of L-methionine from D,L(2-methylthioethyl) hydantoin by resting cells of a new mutant strain of Arthrobacter species DSM 7330. J. Biotechnol. 46:63–68. Wei J, Liu C, and Leyh TS. (2000) The role of enzyme isomerization in the native catalytic cycle of the ATP sulfurylase-GTPase system. Biochemistry 39:4704–4710. Weissbach H and Brot N. (1991) Regulation of methionine synthesis in Escherichia coli. Mol. Microbiol. 5:1593–1597. Wiebers JL and Garner HR. (1967) Homocysteine and cysteine synthetases of Neurospora crassa. Purification, properties, and feedback control of activity. J. Biol. Chem. 242:12–23. Wild CM, McNally T, Phillips SEV, and Stockley PG. (1996) Effects of systematic variation of the minimal Escherichia coli met consensus operator site: in vivo and in vitro met repressor binding. Mol. Microbiol. 21:1125–1135. Williams SJ, Senaratne RH, Mougous JD, Riley LW, and Bertozzi CR. (2002) 5′Adenosinephosphosulfate lies at a metabolic branch point in Mycobacteria. J. Biol. Chem. 277:32606–32615. Wyman A, Shelton E, and Paulus H. (1975) Regulation of homoserine transacetylase in whole cells of Bacillus polymyxa. J. Biol. Chem. 250:3904–3908. Yamada H, Morinaga Y, and Tani Y. (1982) L-Methionine overproduction by ethionine-resistant mutants of obligate methylotroph strain OM33. Agric. Biol. Chem. 46:47–55. Yamagata S. (1989) Roles of O-acetyl-L-homoserine sulfhydrylases in microorganisms. Biochimie 71:1125–1143. Zeghouf M, Fontecave M, and Covès J. (2000) A simplified functional version of the Escherichia coli sulfite reductase. J. Biol. Chem. 275:37651–37656.

16

Phosphorus Metabolism V.F. Wendisch and M. Bott

CONTENTS 16.1 Introduction ..................................................................................................377 16.2 Genomic Survey of Genes/Proteins Involved in Phosphorus Metabolism...................................................................................................378 16.2.1 Phosphorus Uptake ..........................................................................378 16.2.2 Extracytoplasmic Phosphorus Mobilization ....................................384 16.2.3 Alternative Phosphorus Sources ......................................................385 16.2.4 Phosphorus Assimilation and Polyphosphate Metabolism..............386 16.3 The Phosphate Starvation Response............................................................388 16.4 Regulation of the Phosphate Starvation Response......................................392 16.5 Comparison of Phosphorus Metabolism and Its Regulation in C. glutamicum, E. coli, B. subtilis, and M. tuberculosis.............................392 References..............................................................................................................394

16.1 INTRODUCTION Phosphorus (P) is one of the major constituents of the cell, making up about 1.5 to 2.1% of the cell dry weight. Phosphorus occurs in inorganic form as orthophosphate (Pi), pyrophosphate (PPi) or polyphosphate (polyPi) and in numerous organophosphates, such as nucleotides, sugar phosphates, and phospholipids. Major P-containing components of the cells are the polymers RNA, DNA, and polyphosphate. Phosphorus plays a central role in energy metabolism, since the free energy obtained by the oxidation of substrates is used to generate ATP from ADP and Pi. Moreover, many cellular regulatory mechanisms depend on Pi, e.g., in the phosphotransfer reactions of two-component signal transduction systems or of serine/threonine protein kinases. Studies on the P metabolism of Corynebacterium glutamicum were initiated only recently and have focused on polyphosphate metabolism [22], the characterization of pyrophosphatase [32], the definition of the phosphate starvation stimulon [18], and the elucidation of the phosphoproteome [4]. In this chapter, the information on P metabolism deducible from the genome sequence as well as the available experimental data will be summarized.

377

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16.2 GENOMIC SURVEY OF GENES/PROTEINS INVOLVED IN PHOSPHORUS METABOLISM The availability of the complete genome sequence of C. glutamicum ATCC13032 [17,19] allows us to search for genes encoding proteins, which presumably participate in P uptake and metabolism owing to their high sequence similarity with proteins of other bacteria, which have been shown experimentally to be involved in P uptake or metabolism. In the following paragraphs, an overview of such genes and proteins in C. glutamicum will be given. Their genomic organization is shown in Figure 16.1, and a number of basic properties of the proteins are summarized in Table 16.1. It has to be pointed out that with very few exceptions none of these proteins from C. glutamicum has been experimentally analyzed hitherto.

16.2.1 PHOSPHORUS UPTAKE Three putative uptake systems for inorganic phosphate (PitA, NptA, Pst) are encoded in the genome of C. glutamicum. First, NCgl0445 (pitA) presumably encodes a lowaffinity, high-velocity phosphate uptake system because the derived protein shows 28% sequence identity to PitA of Escherichia coli, which appears to be formed constitutively and is the major Pi uptake system when Pi is in excess [33,34]. Divalent cations such as Mg2+ or Ca2+ are essential for PitA activity and PitA presumably transports a soluble neutral metal phosphate complex (MeHPO4) in symport with a proton [40]. The involvement of PitA in the transport of divalent cations is supported by an increased resistance of a pitA mutant to toxic external zinc concentrations, and it was also suggested that PitA plays a role in zinc efflux [3]. The reported apparent Km values of PitA for Pi vary between 25 μM [33] and 38 μM [43] in whole cells and 11.9 μM in vesicles [40], and Vmax was determined to be 55 nmol Pi⋅min–1⋅(mg protein)–1 [43]. The absence of PitA has no effect on growth with Pi or on Pho regulon control in E. coli [42]. Preliminary evidence indicates that C. glutamicum pitA is not expressed constitutively but repressed under Pi limitation (our unpublished results). Second, C. glutamicum NCgl2648 (nptA) codes for a putative Na+-dependent Pi transporter. This type of transporter was originally found in animals, but recently also in Vibrio cholerae [23]. Evidence was provided that the V. cholerae NptA protein, which is significantly smaller (383 amino acid residues) than its eukaryotic homologs (~640 amino acid residues), is a functional Na+-phosphate symporter with apparent Km values of 300 μM for Pi and 75 mM for Na+ and a Vmax value with respect to Pi of 9 pmol⋅min–1⋅(mg protein)–1 [23]. These values were obtained with whole cells of E. coli expressing V. cholerae nptA. The C. glutamicum NptA protein shows 29% sequence identity to the V. cholerae homolog and expression of the C. glutamicum nptA gene was not induced under Pi starvation conditions [18]. Third, the C. glutamicum pstSCAB operon (NCgl2483-2486; see Figure 16.1) most likely codes for an ABC-type transporter catalyzing ATP-driven high-affinity phosphate uptake. PstS functions as an extracytoplasmic phosphate-binding protein because it contains a lipoprotein signal sequence (Ala-Leu-Val-Ala-Cys*-Ser) typical

Phosphorus Metabolism

pnuX

phoH1

citM

0063 0066 Riboflavin transporter

0444 0446

psiB

pctD pctC pctB

Lipoate- PhnB- Benzoate Protein like transport ligase protein 1 protein

1328 1334

pstB pstA

ugpA ugpE

ppa

glpQ1

ugpC

sigA

Glycero phosphoryldiester phosphodiesterase 1

phoD 2185 tRNA- Alkaline tRNA- tRNAMet- phosphatase Asn- AsnCAT GTT GTT

psiC

phoB

2253 2255

pstC

2371 Alkaline tRNAphoB phosphatase HisGTG

nucH

pstS 2502 2504 Glutathione peroxidase

speE

Nuclease

ppk2B

2606 2608

Ferrochelatase

groEL

26192621

D-alanylInorganic Spermidine D-alanine pyrophosphatase synthase carboxypeptidase

NAD-dependent aldehyde dehydrogenase

nptA 26472649 MFS transporter Na +/phosphate Epimerase symporter

Exopoly- tRNAphosphatase Leu2 TAA

Glycerol-3-phosphate transporter

Phosphate Phosphate ABC transporter transport reuglatory protein

dacB

ugpB

1834 1836 Inositol Polyphosphate- RNA polymerase mono- glucokinase sigma factor A phosphatase

Molecular chanperone

phoU

ppx2

Enolase

dnaJ

PhoH2 protein

GroES chaperone

eno

ppgK

pctA

ABC transporter

phoH2

2481 2487

Sugar kinase

0935 0938

lplA phnB1 1029 1031

PhnA- MarR-type like regulator protein

1401 1406 RpiRtype regulator

2207 2210

Naphthoate synthase

Polyphosphate kinase 2A

phnA

psiA

0570 0572

0879 0881 MFS-type transport system

1017 1019

pitA Low-affinity inorganic phosphate transporter

ppk2A

0814 0816

UDP-sugar hydrolase

Glycosyl transferase

proC Pyrroline-5carboxylate reductase

Exopolyphosphatase 1

ushA

0321 0323

Citrate-proton symporter

PhoH1 protein

ppx1 0395 0398

379

phnB2 27212726 PhnB-like protein 2

tctA

tctB

glpQ2

tctC

tricarboxylate transporter

Chaperonin GroEL

gntP

28062808 Trans- Glycerophosphoryl Gluconate locase diester phospho- permease diesterase 2

proP 28192823

29582961 ICC-family phosphohydrolase

Inositol dehydrogenase

Phosphoesterase

Proline/ectoine uptake system

FIGURE 16.1 Organization of C. glutamicum genes shown or suggested to be involved in phosphorus metabolism. The order of the gene loci shown follows that found in the genome sequence. Numbers refer to NCgl synonyms of the genes as in NCBI NC_003450.

of Gram-positive bacteria [37]. C. glutamicum PstS shows 30% sequence identity to E. coli PstS, for which a Kd value for Pi of ~1 μM was determined [20]. The proteins PstA and PstC are integral membrane proteins, and PstB contains the features of an ATP-binding protein. The C. glutamicum PstA, PstB, and PstC proteins show up to 56% sequence identity to the corresponding counterparts in E. coli. The

0571 140 0815 200 0880 279 0938 321 1018 114 1030 137

0064 458 0322 694 0396 281 0445 461

aa

aa

aa

aa

aa

aa

aa

aa

aa

aa

NCgl No.

0596 143 0849 200 1479 279 0977 321 1063 114 1075 137

0065 374 0328 694 0408 309 0460 425

aa

aa

aa

aa

aa

aa

aa

aa

aa

aa

Cgl No.

0689 143 0971 200 917 279 1115 321 1209 114 1224 137

0085 425 0397 694 0488 309 0545 425

aa

aa

aa

aa

aa

aa

aa

aa

aa

aa

Cg No.

phnB1

phnA

ppx2

ppk2A

psiB

psiA

pitA

ppx1

ushA

phoH1

Gene

PhnB-like protein 1

PhnA-like protein

Exopolyphosphatase 2

Polyphosphate kinase 2A

Hypothetical protein

Low-affinity Pi transporter, presum. uptake of a metal phosphate in symport with H+ Hypothetical protein

ATPase related to Pi starvation inducible protein UDP-sugar hydrolase/5′nucleotidase Exopolyphosphatase 1

Putative Product or Function

14,826

12,020

34,763

37,614

21,968

15,065/15,409

48,092/43,962

72,288 69,485 30,189/33,038

50,199/40,772/46,739

Mass (Da)

0

0

0

0

4

0

10 10

0

1

0

TMH

No

No

No

No

No?

No

No

No

1–28

No

Signal Peptide

TABLE 16.1 Overview of Genes and Proteins Involved in Phosphorus Metabolism in C. glutamicum

No

No

No

No

No

No

No

PAFA ↓ DE

No

Cleavage Site

C

C

C

C

CM

CM

C

CM

C

Location (C, CM, EC)

No

No

No

No

Yes

Yes

No

No

Yes

Yes

Pi Starvation Induced1

380 Handbook of Corynebacterium glutamicum

1402 282 1403 268 1404 268 1405 350 1835 250 2185 516 2208 350 2254 534

1329 301 1330 278 1331 438 1332 408 1333 235

aa

aa

aa

aa

aa

aa

aa

aa

aa

aa

aa

aa

aa

1458 282 1459 268 1460 268 1461 350 1910 276 2265 520 2288 350 2336 536

1383 301 1384 278 1385 438 1386 408 1387 239

aa

aa

aa

aa

aa

aa

aa

aa

aa

aa

aa

aa

1649 282 1650 268 1651 268 1652 350 2091 250 2485 516 2513 339 2566 531

1568 310 1569 278 1570 438 1571 408 1572 235

aa

aa

aa

aa

aa

aa

aa

aa

aa

aa

aa

aa

aa

psiC

phoH2

phoD

ppgK

pctA

pctB

pctC

pctD

glpQ1

ugpC

ugpB

ugpE

ugpA

Phosphodiesterase/alkaline phosphatase D (EC 3.2.3.1) ATPase related to Pi starvation inducible protein Hypothetical protein

ABC transporter, ATP binding protein ABC transporter, alkyl phosphonate binding protein Polyphosphate glucokinase

ABC transporter, permease

G3P ABC transporter, G3P binding protein G3P ABC transporter, ATP binding protein Glycerophosphoryl diester phosphodiesterase (EC 3.1.4.46) ABC transporter, permease

G3P ABC transporter, permease

G3P ABC transporter, permease

56,716/56,943/56,389

56,888/57,376 p 53,844/53844 m 38,766/37,640

36,687 p 33,869 m 26,687/29,645

28,953

28,656

30,495

25,405/25,942

47,098 p 44,760 m 44,137

31,216

34,072/35,028

1

0

0

0

0

4

5

0

0

0

6

6

Sec (1-29)

Tat (1No

Sec (1-27) No

No

No

No

No

Tat/Sec? (1-24) No

No

No

AANA ↓ VE

No

PARA ↓ EE

No

ALVG ↓ CS

No

No

No

No

No

TLAA ↓ CA

No

No

CM

C

EC

C

EC

C

CM

CM

C

C

EC

CM

CM

Yes

No

No

No

No

Yes

Yes

No

Yes

Yes

Yes

Yes

Yes

Phosphorus Metabolism 381

2371 473 2482 244 2483 257 2484 307 2485 338 2486 375 2503 916 2607 158

aa

aa

aa

aa

aa

aa

aa

aa

NCgl No.

2457 473 2571 244 2572 299 2573 307 2574 338 2575 375 2592 916 2700 158

aa

aa

aa

aa

aa

aa

aa

aa

Cgl No.

2700 473 2842 244 2843 257 2844 307 2845 355 2846 375 2868 916 2988 158 aa

aa

aa

aa

aa

aa

aa

aa

Cg No.

ppa

nucH

pstS

pstC

pstA

pstB

phoU

phoB

Gene

Inorganic pyrophosphatase

Pi ABC transporter, Pi binding protein Nuclease

Pi ABC transporter, permease

Pi ABC transporter, ATP binding protein Pi ABC transporter, permease

Pi uptake regulator

Alkaline phosphatase

Putative Product or Function

38,788 p 36,402 m 96,444 p 93,736 m 17,905

35,707/37,593

32,738

28,213/32,932

50,797 p 47,918 m 28,130

Mass (Da)

0

1

0

6

5

0

0

0

TMH

No

Sec (1-24) Yes

No

No

No

Sec (1-29) No

Signal Peptide

TABLE 16.1 (continued) Overview of Genes and Proteins Involved in Phosphorus Metabolism in C. glutamicum

No

AATA ↓ NP

ALVA ↓ CS

No

No

No

No

TAAA ↓ QS

Cleavage Site

C

CM

EC

CM

CM

C

C

EC

Location (C, CM, EC)

No

Yes

Yes

Yes

Yes

Yes

No

No

Pi Starvation Induced1

382 Handbook of Corynebacterium glutamicum

2714 306 aa 2744 388 aa 2821 138 aa 2907 273 aa 2922 539 aa 3064 1344 a

3007 299 aa 3039 436 aa 3122 138 aa 3215 253 aa 3232 539 aa 3393* 1461 a 40,948/46,145

Na+-dependent Pi transporter PhnB-like protein 2 Glycerophosphoryl diester phosphodiesterase (EC 3.1.4.46) Secreted phosphohydrolase, ICC family Putative secreted phosphoesterase (*signal peptide only predicted for cg3393)

nptA phnB2 glpQ2 0

59,544 p 56,589 m 157,168 p 154,968 m 1

0

0

10/10

0

31,432/28,922

15,355

36,044/35,305

Polyphosphate kinase 2B

ppk2B

Sec/Tat? (1-28) Sec (1-22)

No

No

No

No

ALA ↓ LG

AIA ↓ DE

No

No

No

No

CM

EC

C

CM

CM

C

Yes

No

No

No

No

No

Abbreviations: p, preprotein; m, mature protein; TMH, transmembrane helix; C, cytoplasmic; CM, cytoplasmic membrane; EC, extracytoplasmic. In the case of predicted lipoprotein, the N-terminal cysteine residue of the mature protein is indicated in bold and underlined. 1 As determined by Ishige et al. [18]. The numbers below the gene numbers give the respective length of the derived polypeptides in aminoacyl residues (aa). Note that upon prolonged Pi starvation additional genes may change expression.

2620 306 aa 2648 388 aa 2722 138 aa 2807 273 aa 2821 539 aa 2959 1497 a

Phosphorus Metabolism 383

384

Handbook of Corynebacterium glutamicum

apparent Km value for Pi of the E. coli Pst system was found to be around 0.2 μM [33,43]. As is true for many bacteria, the pstSCAB operon of C. glutamicum is strongly induced by phosphate starvation [18]. Besides the uptake systems for Pi described above, at least one organophosphate uptake system is present in C. glutamicum. The ugpAEBC operon (NCgl1329-1332; see Figure 16.1) presumably encodes an ABC transport system for the uptake of glycerol-3-phosphate. Protein UgpB functions as an extracytoplasmic glycerol-3phosphate-binding protein and contains a characteristic lipoprotein signal sequence (Thr-Leu-Ala-Ala-Cys*-Ala); UgpA and UgpE are integral membrane proteins; and UgpC contains features of an ATP-binding protein. The corresponding proteins show up to 49% (UgpC) sequence identity with the corresponding E. coli counterparts. The E. coli Ugp system has a Km for glycerol-3-phosphate of ~1.5 μM and a Vmax of ~3 nmol⋅min–1⋅(mg protein)–1 [35]. Remarkably, the E. coli Ugp system cannot transport glycerol-3-phosphate for use as a carbon source, presumably because high intracellular Pi inhibits uptake [7]. As in many other bacteria, the C. glutamicum ugpAEBC operon and presumably also the adjacent, but divergently transcribed glpQ1 gene encoding a glycerophosphoryldiester phosphodiesterase, are induced when C. glutamicum is starved for phosphate [18]. In other bacteria, several further uptake systems for organophosphates are known, e.g., GlpT for glycerol-3-phosphate; UhpT for hexose-6-phosphates; and PgtP for phosphoenolpyruvate, 2-phosphoglycerate, and 3-phosphoglycerate [42]. The C. glutamicum genome lacks homologs of the corresponding genes. However, the pctABCD operon (NCgl1402-1405, see Figure 16.1) encoding an ABC transport system was induced after Pi starvation [18]. Sequence comparisons revealed that the corresponding proteins have low similarity to phosphonate ABC transporters. The periplasmic binding protein PctA contains a lipoprotein signal sequence (Ala-LeuVal-Gly-Cys*-Ser). The PctABCD ABC transporter might be involved in the uptake of an as-yet-unidentified P compound.

16.2.2 EXTRACYTOPLASMIC PHOSPHORUS MOBILIZATION Under phosphate limitation, many bacteria form periplasmic or extracellular phosphatases that hydrolyze organophosphates and, subsequently, the liberated free phosphate is taken up by the Pi transport systems. Such enzymes are presumably present in C. glutamicum, as well. First, NCgl0322 (ushA) encodes a protein active as a UDP sugar hydrolase (EC 3.6.1.45), which catalyzes the hydrolytic cleavage of UDP sugars to UMP and sugar-1-phosphate and/or as a 5′-nucleotidase (EC 3.1.3.5), which hydrolyzes 5′-(desoxy)ribonucleotides to (desoxy)ribonucleosides and Pi (our unpublished results). The protein shows 29% identical amino acids to UshA from E. coli and it contains a signal peptide and presumably a C-terminal transmembrane helix. Thus, C. glutamicum UshA is a transmembrane protein with most of the protein facing outside, or, alternatively, it is processed to yield a cell wall-associated or secreted protein. Under Pi starvation conditions, the C. glutamicum ushA gene is strongly induced [18] whereas in E. coli ushA is not phosphate-starvation inducible. In E. coli, dephosphorylation of 5′-AMP by the nucleotidase activity of UshA is required for

Phosphorus Metabolism

385

growth of E. coli on 5′-AMP as sole carbon source [27,44]. In C. glutamicum, ushA is essential to utilize AMP or UDP-glucose as the sole P source (our unpublished results). This is commensurate with the view that in C. glutamicum UshA primarily functions in phosphorus metabolism, whereas in E. coli UshA is important for nucleotide salvage. Second, NCgl2503 (nucH) encodes a protein that could function as nuclease. The predicted signal peptide and a single C-terminal transmembrane helix suggest that NucH is a membrane protein. However, processing of the protein to yield a cell wall–associated or secreted protein cannot be ruled out. The nucH gene is strongly induced after Pi limitation [18] and may serve a role in liberating Pi from extracellular nucleic acids. Third, NCgl2371 (phoB) presumably encodes an alkaline phosphatase because it shows 36% sequence identity to the alkaline phosphatase PhoB of B. subtilis [16]. It contains a signal peptide of 29 residues and presumably is exported. In contrast to B. subtilis, the C. glutamicum phoB gene was only slightly induced in some of the phosphate starvation experiments (our unpublished results). Fourth, NCgl2185 (phoD) encodes a second protein that may function as an alkaline phosphatase, since it shows 38% sequence identity to the alkaline phosphatase PhoD of B. subtilis [11] and 36% identity to the PhoD ortholog from Streptomyces griseus [25]. At the N-terminus of C. glutamicum PhoD, a signal peptide of 29 residues is present that contains the Tat consensus motif SRRxFL [5]. Since C. glutamicum possesses both TatA (NCgl1434, NCgl2498) and TatC (NCgl1433) orthologs, it is possible that PhoD is exported by the Tat system, as was shown for B. subtilis [28]. In contrast to B. subtilis, the C. glutamicum phoD gene was not induced after Pi limitation (our unpublished results). Therefore, the function of PhoB and PhoD in C. glutamicum is unclear at present. Fifth, NCgl2959 encodes a protein containing a putative phosphoesterase domain in the carboxyterminal region. Depending on the proposed start codon, the protein may contain a signal peptide (cg3393, 1461 aa) or may not (NCgl2959, 1497 aa; Cgl3064, 1344 aa). At the carboxyterminal end, a putative transmembrane helix is present, in front of which a putative sortase motif (APGTG) was detected. Therefore, the NCgl2959 protein may function as a cell wall–associated phosphatase. Orthologous proteins are found in C. efficiens (CE2911, 72% sequence identity) and in B. subtilis (YvnB, 33% sequence identity), but like the C. glutamicum protein they have not been functionally characterized. The NCgl2959 gene showed a 12-fold increase in mRNA level 120 min after a shift from Pi excess to Pi starvation conditions, indicating that the corresponding protein could be directly involved in the adaptation process [18].

16.2.3 ALTERNATIVE PHOSPHORUS SOURCES Besides Pi, C. glutamicum can also utilize pyrophosphate, polyphosphate, glycerol3-phosphate, 3-phosphoglycerate, phosphoenolpyruvate, glucose-6-phosphate, glucose1-phosphate, AMP, UDP-glucose, and DNA as P sources for growth, but neither aminophosphonate nor ethylphosphonate (our unpublished results). C. glutamicum is able to degrade the organophosphorous pesticide demeton-S-methyl during cometabolism

386

Handbook of Corynebacterium glutamicum

with glucose, fructose, or acetate [12]. During this biotransformation, the S–C bond of demeton-S-methyl is reductively cleaved to yield dimethyl thiophosphate, which is not utilized further but accumulates in the culture medium.

16.2.4 PHOSPHORUS ASSIMILATION

AND

POLYPHOSPHATE METABOLISM

Phosphorus is assimilated in the form of phosphate. Table 16.2 lists enzymatic reactions contributing to phosphate assimilation in C. glutamicum. Many reactions of the central carbon and energy metabolism contribute to the assimilation of phosphate into ATP and other central metabolites. However, with ATP as precursor of nucleotide metabolism most of the assimilated phosphate will end up in polymers, particularly in RNA. C. glutamicum possesses a number of genes putatively coding for enzymes of the pyrophosphate or polyphosphate metabolism. Two genes for class II polyphosphate kinases (NCgl0880 and NCgl2620), two genes encoding exopolyphosphatases (NCgl0396 and NCgl0938), and genes for a polyphosphate-dependent glucokinase (NCgl1835) and a pyrophosphatase (NCgl2607; ppa) can be found in its genome. Although none of the proteins encoded by these genes, except pyrophosphatase, have been characterized biochemically, it is clear that C. glutamicum can synthesize polyphosphate as revealed by 31P-NMR (Figure 16.2) [22]. Furthermore, electrondense particles within C. glutamicum cells have been interpreted as polyphosphatecontaining granules [9]. The pyrophosphatase gene ppa of C. glutamicum (Brevibacterium lactofermentum) ATCC13869 was shown to complement an E. coli ppa mutant and to encode a soluble pyrophosphatase (EC 3.6.1.1) [32]. This protein was identified because it copurified with the cell division protein FtsZ, and, interestingly, a ppa-gfp gene fusion revealed that the gene product mainly accumulated at the cell poles in both E. coli and B. lactofermentum [32]. It remains unclear whether the ppa-encoded protein serves functions in addition to its enzymatic role in hydrolysis of inorganic pyrophophate. The proteins encoded by NCgl0396 and NCgl0938 show about 25% sequence identity to each other and to the E. coli exopolyphosphatase Ppx (EC 3.6.1.11), which contains 200 additional amino acids at its C-terminus that are absent in the C. glutamicum proteins. C. glutamicum possesses two genes (NCgl0880 and NCgl2620) for putative class II polyphosphate kinases. The corresponding proteins, PPK2A (encoded by NCgl0880) and PPK2B (encoded by NCgl2620), show 57% and 64% sequence identity with Pseudomonas aeruginosa PPK2 [45], respectively, and 51% sequence identity with each other. Genes encoding class I polyphosphate kinases [45], such as E. coli ppk, are absent from the genome of C. glutamicum or other corynebacteria, but are present in M. tuberculosis and other mycobacteria [45]. The roles and biochemical properties of the putative C. glutamicum exopolyphosphatases and polyphosphate kinases remain to be studied. A polyphosphate-dependent glucokinase similar to the biochemically characterized enzymes from other actinomycetes is likely encoded by C. glutamicum NCgl1835. It shares 45% identical amino acids with polyphosphate/ATP glucomannokinase of Arthrobacter sp. strain KM [26] and 50% with polyphosphate glucokinase of

TCA cycle Acetate excretion

Oxidative phosphorylation Glycolysis

Pathway

→

Glyceraldehyde-3-phosphate + Pi + NAD+ 1,3-bisphosphoglycerate + ADP Succinyl-CoA + ADP + Pi Acetyl-CoA + Pi Acetyl-phosphate + ADP → → → →

→

ADP + Pi

Reaction

Proton-pumping F1F0-ATPase Glyceraldehyde-3-phosphate dehydrogenase Phosphoglycerate kinase Succinyl-CoA synthetase Phosphotrans acetylase Acetate kinase

1,3-bisphosphoglycerate + NADH + H+ 3-phosphoglycerate + ATP Succinate + ATP + CoA Acetyl-phosphate + CoA Acetate + ATP

Enzymes ATP

TABLE 16.2 Pathways Contributing to Phosphate Assimilation in C. glutamicum

atpIBEFHAGDC (NCgl1158-1166) gap (NCgl1526) gap2 (NCgl900) pgk (NCgl1525) sucCD (NCgl2476-2477) pta (NCgl2657) ack (NCgl 2656)

Genes

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FIGURE 16.2 In-vivo 31P-NMR spectrum of a C. glutamicum cell suspension. Signals of (1) phosphomonoesters, such as NMP, NADP(H), hexose-, pentose- and triose-phosphates; (2) intracellular inorganic phosphate; (3) extracelluar inorganic phosphate; (4) PEP and phosphodiesters, such as tRNA, low-molecular-weight RNA and DNA; (5) NTPγ and NDPβ; (6) NDPα and NTPα; (7) NAD(P)(H) and NDP-Glucose; (8) NDP-Glucose; (9) NTPβ; and (10) core phosphate units of poly P (PP4) are marked. (Courtesy of S. M. Schoberth, Institute for Biotechnology, Forschungszentrum Jülich.)

M. tuberculosis H37Rv [14]. Glucokinases of this class accept both ATP and polyphosphate as phosphoryl donors, and they are distinct from exclusively ATP-dependent glucokinases, which occur throughout the domains of life, as well as from strictly polyphosphate-dependent glucokinases, which to date have been identified only in the polyphosphate-accumulating bacterium Microlunatus phosphovorus [38].

16.3 THE PHOSPHATE STARVATION RESPONSE In the standard minimal medium CGXII with 40 g of glucose per liter as the C-source and 13 mM potassium phosphate as the sole P source, C. glutamicum ATCC 13032 reaches a final OD600 of approximately 60. When the cells are washed and used to inoculate CGXII-glucose medium lacking a P source, they still grow and reach a final OD600 of about 12, indicating the existence of internal P sources, such as polyphosphate [18,22]. C. glutamicum cells precultured under Pi-limiting (0.13 mM) conditions show almost no growth in the absence of a P source but exhibit a proportional increase in the cell yield as well as in the growth rate with increasing Pi concentrations (Figure 16.3). The Monod constant, i.e., the Pi concentration supporting growth with a half-maximal growth rate, is about 0.1 mM [18]. The Pi starvation stimulon and the kinetics of the global gene expression changes during a shift from Pi sufficient to limiting conditions were characterized recently [18]. Hierarchical cluster analysis of the observed gene expression changes (Figure 16.4) revealed five groups or subclusters of genes that showed similar expression profiles. Subcluster 1 comprised 25 genes whose mRNA level increased early after the Pi downshift and remained high until 180 min after the onset of Pi limitation. Except for NCgl1123, a gene encoding a hypothetical protein similar to yceI in E. coli, this

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FIGURE 16.3 Growth rate as a function of the Pi concentration (0.13–13 mM) with starved or unstarved inoculum, deducing KS, and importance of intracellular P storage compounds. Reprinted from Ishige et al. [18] with permission.

subcluster contained all of the genes that showed increased expression in cells grown for 7.5 h under Pi-limiting as compared with Pi-saturating conditions [18]: pstSCAB, glpQ1, ugpAEBC, nucH, and the hypothetical genes NCgl0571 and NCgl2254. In addition, subcluster 1 comprises genes presumably involved in P metabolism, including ushA, the phosphate-starvation-inducible gene phoH, two genes of the pctABCD operon, and a gene encoding a phosphoesterase (NCgl2959). Furthermore, a number of genes not obviously related to P metabolism showed increased RNA levels after the shift to Pi limitation, including those encoding methylcitrate lyase (prpB1), methylcitrate synthase (prpC1), ferrochelatase (NCgl2505), two hypothetical proteins (NCgl2254 and ORF 3362), a permease of unknown function (NCgl2052), and glucose-1-dehydrogenase (NCgl2053). It is not yet clear whether these proteins play a role, direct or indirect, in the adaptation of C. glutamicum to Pi starvation. With regards to the kinetics of expression changes, the genes of subcluster 1 can be ordered. Expression of the high-affinity Pi uptake system genes pstSCAB increased first after the shift to phosphate starvation conditions followed by increases in expression of the sn-glycerol 3-phosphate uptake genes ugpAEBC and glpQ1 and of nucH, ushA, NCgl0571, and NCgl2554. All other genes in subcluster 1 exhibited smaller and slower increases in expression. Expression of a transcriptional fusion of the pstS promoter region to the promoterless chloramphenicol acetyltransferase reporter gene on the promoter-probe vector pET2 [41] increased immediately after the Pi downshift and reached maximal levels about 60 min after the shift to Pi limitation. The temporal expression profile of pstS-cat is in good agreement with

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the profile obtained in the DNA microarray experiments (Figure 16.5). Similarly, expression of a transcriptional fusion of the ugpA promoter region increased after the Pi downshift, but with a delay as compared with pstS-cat (Figure 16.5). Thus, transcription of the pstSCAB and ugpAEBC operons is induced in response to reduced Pi availability. Subcluster 2 (Figure 16.4) included nine genes that showed transiently increased mRNA levels after the Pi downshift. Five of these genes are part of a chromosomal locus encoding a copper-exporting ATPase, an associated protein, a two-component regulatory system, a hypothetical protein, a multicopper oxidase, and a thiol-disulfide interchange protein (NCgl2860 to NCgl2866). Additionally, genes encoding a response regulator (NCgl2518), a nonheme ferritin (NCgl2439), a putative cation efflux protein, and two hypothetical proteins (NCgl2877 and NCgl1232) belong to subcluster 2. Subcluster 3 (Figure 16.4) contained nine genes whose mRNA levels increased rapidly after the Pi downshift but decreased to pre-downshift levels already after 30 min (Figure 16.4). Five of these genes (NCgl 2306, 2309, 2310, 2314, and NCgl1032) encode enzymes or subunits of enzymes predicted to be involved in the degradation of protocatechuate, which is present in CGXII medium to facilitate iron uptake, and similar compounds to yield acetyl-CoA and succinyl-CoA. Additionally, the genes encoding a putative transporter (NCgl2816), the arginine repressor and ornithine carbamoyltransferase (NCgl1344 and 1345), and a sensor kinase (NCgl2517) showed short-term expression increases after the onset of Pi limitation. Interestingly, expression of the response regulator gene NCgl2518 located next to the sensor kinase gene NCgl2517 transiently increased in a similar manner. Subcluster 4 (Figure 16.4) comprised 29 genes that showed decreased RNA levels 60 min after the onset of Pi limitation and continued to exhibit low levels at later times. Most of these genes code for proteins involved in translation or DNA replication; 15 genes code for ribosomal proteins; 2 genes code for translation initiation factors (NCgl0536 and NCgl1324); and 1 gene codes for the singlestranded DNA-binding protein (NCgl2880). Reduced expression of these genes, as well as of several other genes, was correlated with reduced growth after the shift to Pi-limiting conditions. Subcluster 5 (Figure 16.4) included 18 genes that exhibited transiently reduced expression after the Pi downshift. The RNA levels were lower 10, 30, 60, and 90 min after the onset of Pi limitation, but after 120 and 180 min the RNA levels were the same as those before the shift. Fifteen of these genes are presumably involved in iron metabolism and putatively encode a heme transport system and associated proteins, two ferric siderophore transport systems and an associated protein, two ferric dicitrate-binding proteins, and a siderophore utilization protein. Besides these putative iron metabolism genes, two genes encoding hypothetical proteins and a gene encoding a transcriptional regulator belong to subcluster 5.

FIGURE 16.4 (opposite page) (Color insert follows page 208.) Hierarchical cluster analysis of gene expression changes 10, 30, 60, 90, 120 and 180 min after C. glutamicum cells were shifted to phosphate-starvation conditions. Reprinted from Ishige et al. [18] with permission.

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FIGURE 16.5 Kinetics of expression changes of the pstSCAB and ugpAEBC operons after C. glutamicum cells were shifted to phosphate-starvation conditions as monitored by transcriptional fusion analyses. Reprinted from Ishige et al. [18] with permission.

16.4 REGULATION OF THE PHOSPHATE STARVATION RESPONSE In many bacteria, the adaptation to Pi starvation conditions is controlled by twocomponent regulatory systems, e.g., the PhoRB system in E. coli or the PhoPR and ResDE systems in B. subtilis. In the genome of C. glutamicum, genes encoding 13 two-component systems have been identified. Screening of C. glutamicum strains lacking the genes for one of the two-component systems revealed one mutant that was severely affected in the response to Pi starvation (our unpublished results). Preliminary studies indicate that the two-component system lacking in this mutant is of major importance for eliciting the phosphate starvation response in C. glutamicum. Most, but not all, of the genes known to be involved in the adaptation to Pi starvation were no longer induced in the mutant (our unpublished results). However, there was also evidence for the participation of a second, yet unknown regulatory system. Studies are in progress to further clarify the phosphate starvation response in C. glutamicum.

16.5 COMPARISON OF PHOSPHORUS METABOLISM AND ITS REGULATION IN C. GLUTAMICUM, E. COLI, B. SUBTILIS, AND M. TUBERCULOSIS The characterization of the C. glutamicum Pi starvation stimulon revealed similarities as well as differences with the Pi starvation stimulons of E. coli and B. subtilis. In all three bacteria, Pi starvation conditions result in increased expression of the pst

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genes encoding ABC transporters for high-affinity Pi uptake [16,42]. Similarly, the induction of the ugp genes for sn-glycerol 3-phosphate ABC uptake systems (except in B. subtilis) and glycerophosphoryl diester phosphodiesterases provides these bacteria with the capacity to utilize sn-glycerol 3-phosphate-yielding organophosphates [16,42]. In E. coli as well as in B. subtilis, induction of genes for alkaline phosphatases allows access to Pi from nontransportable organophosphates [16,42]. In contrast, expression of C. glutamicum phoB increases only slightly and that of phoD remains unchanged under the Pi starvation conditions employed by Ishige et al. [18]. The regulation of the phosphate starvation response in E. coli and B. subtilis involves two-component regulatory systems [16,42]. In E. coli, the sensor kinase PhoR and the response regulator PhoB represent the central Pi regulatory system [42]. In B. subtilis, Pi starvation regulation requires the specific phosphate regulatory system PhoPR, as well as Spo0A for termination of the Pi starvation response and subsequent initiation of sporulation and ResDE for full induction of the Pho regulon genes [6,16,36]. As outlined above, a two-component regulatory system as well as a second regulatory system are involved in eliciting the Pi starvation response of C. glutamicum. E. coli and Salmonella enterica serovar Typhimurium are able to utilize phosphonates as P sources when Pi is scarce [42]. The majority of the phosphatestarvation-inducible genes in these bacteria, i.e., 21 of 38 genes, play a role in the uptake and the degradation of phosphonates, e.g., the phnCDEFGHIJKLMNOP operon, and their expression is regulated by PhoR [42]. In contrast, the C. glutamicum genome [17,19] lacks homologs of genes for phosphonate degradation as well as the capability to utilize aminophosphonate or ethylphosphonate as sole P sources for growth (our unpublished results). On the other hand, the C. glutamicum ushA gene encoding a putative 5′-nucleotidase or related esterase showed increased expression upon Pi starvation, but the E. coli ushA homolog is not a known member of the Pi starvation stimulon [42]. As outlined above, ushA is required for utilization of AMP or UDP-glucose as sole P source by C. glutamicum (our unpublished results). In contrast to C. glutamicum and E. coli, the response of B. subtilis to Pi starvation also involves increased expression the tuaABCDEFGH operon coding for teichuronic acid biosynthesis and reduced expression was observed for the tagAB and tagDEF teichoic acid biosynthesis operons [2,16,21,29,30]. As a consequence, upon Pi starvation B. subtilis replaces teichoic acids in the cell wall with the non-phosphatecontaining teichuronic acids. Neither teichoic nor teichuronic acids are present in C. glutamicum cell walls and homologs of the B. subtilis tua or tag operons were not identified in the C. glutamicum genome. In B. subtilis, but not in C. glutamicum [18], Pi starvation leads to the induction of genes of the general stress response [2,21,29,30], which is mediated by σB [13,31]. Thus, the induction of the general stress response as well as the replacement of teichoic acids by teichuronic acids distinguishes the Pi starvation response of B. subtilis from that of C. glutamicum. In M. tuberculosis, it was recognized that the most immunogenic antigen, Pab, is present at higher levels under Pi starvation conditions [1]. This protein is a phosphate-binding protein similar to E. coli PstS [8]. It was then realized that the genome of M. tuberculosis harbors three different phosphate-binding protein genes in three operons (pstBS1C1A2, pstS2, and pstS3C2A1) at one locus [10,24] and that

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the phosphate-binding proteins are surface attached [24]. By using translational fusions, two Pi starvation-responsive promoters were identified for the pstBS1C1A2 operon [39]. In the related M. smegmatis, alkaline phosphatase activity was shown to increase under Pi starvation conditions [39]. In view of the close phylogenetic relationship of mycobacteria and corynebacteria, it can be expected that many features of the Pi starvation response of C. glutamicum will also be true for mycobacteria.

REFERENCES 1. Andersen AB, Ljungqvist L, and Olsen M. (1990) Evidence that protein antigen B of Mycobacterium tuberculosis is involved in phosphate metabolism. J. Gen. Microbiol. 136:477–480. 2. Antelmann H, Scharf C, and Hecker M. (2000) Phosphate starvation-inducible proteins of Bacillus subtilis: Proteomics and transcriptional analysis. J. Bacteriol. 182:4478–4490. 3. Beard SJ, Hashim R, Wu G, Binet MR, Hughes MN, and Poole RK. (2000) Evidence for the transport of zinc(II) ions via the pit inorganic phosphate transport system in Escherichia coli. FEMS Microbiol. Lett. 184:231–235. 4. Bendt AK, Burkovski A, Schaffer S, Bott M, Farwick M, and Hermann T. (2003) Towards a phosphoproteome map of Corynebacterium glutamicum. Proteomics 3:1637–1646. 5. Berks BC, Sargent F, and Palmer T. (2000) The Tat protein export pathway. Mol. Microbiol. 35:260–274. 6. Birkey SM, Liu W, Zhang XH, Duggan MF, and Hulett FM. (1998) Pho signal transduction network reveals direct transcriptional regulation of one two-component system by another two-component regulator:Bacillus subtilis PhoP directly regulates production of ResD. Mol. Microbiol. 30:943–953. 7. Brzoska P, Rimmele M, Brzostek K, and Boos W. (1994) The Pho regulon dependent Ugp uptake system for glycerol-3-phosphate in Escherichia coli is trans-inhibited by Pi. J. Bacteriol. 176:15–20. 8. Chang ZY, Choudhary A, Lathigra R, and Quiocho FA. (1994) The immunodominant 38-kDa lipoprotein antigen of Mycobacterium tuberculosis is a phosphate-binding protein. J. Biol. Chem. 269:1956–1958. 9. Coello N, Pan JG, and Lebeault JM. (1992) Corynebacterium glutamicum: Morphological and ultrastructural changes of L-lysine producing cells in continuous culture. Appl. Microbiol. Biotechnol. 38:34–38. 10. Cole ST, Brosch R, Parkhill J, Garnier T, Churcher C, Harris D, Gordon SV, Eiglmeier K, Gas S, Barry CE III, Tekaia F, Badcock K, Basham D, Barrell BG, et al. (1998) Deciphering the biology of Mycobacterium tuberculosis from the complete genome sequence. Nature 393:537–544. 11. Eder S, Shi L, Jensen K, Yamane K, and Hulett FM. (1996) A Bacillus subtilis secreted phosphodiesterase/alkaline phosphatase is the product of a Pho regulon gene, phoD. Microbiology 142:2041–2047. 12. Girbal L, Hilaire D, Leduc S, Delery L, Rols JL, and Lindley ND. (2000) Reductive cleavage of demeton-S-methyl by Corynebacterium glutamicum in cometabolism on more readily metabolizable substrates. Appl. Environ. Microbiol. 66:1202–1204. 13. Hecker M and Völker U. (1998) Non-specific, general and multiple stress resistance of growth-restricted Bacillus subtilis cells by the expression of the σB regulon. Mol. Microbiol. 29:1129–1136.

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14. Hsieh PC, Shenoy BC, Samols D, and Phillips NFB. (1996) Cloning, expression, and characterization of polyphosphate glucokinase from Mycobacterium tuberculosis. J. Biol. Chem. 271:4909–4915. 15. Hulett FM. (2002) The Pho regulon. In: Sonenshein JA and Losick RM (eds) Bacillus subtilis and its closest relatives:from genes to cells. ASM Press, Washington, D.C., pp 193–201. 16. Hulett FM, Kim EE, Bookstein C, Kapp NV, Edwards CW, and Wyckoff HW. (1991) Bacillus subtilis alkaline phosphatases III and IV: Cloning, sequencing, and comparisons of deduced amino acid sequence with Escherichia coli alkaline phosphatase three-dimensional structure. J. Biol. Chem. 266:1077–1084. 17. Ikeda M and Nakagawa S. (2003) The Corynebacterium glutamicum genome: features and impacts on biotechnological processes. Appl. Microbiol. Biotechnol. 62:99–109. 18. Ishige T, Krause M, Bott M, Wendisch VF, and Sahm H. (2003) The phosphate starvation stimulon of Corynebacterium glutamicum determined by DNA microarray analyses. J. Bacteriol. 185:4519–4529. 19. Kalinowski J, Bathe B, Bartels D, Bischoff N, Bott M, Burkovski A, Dusch N, Eggeling L, Eikmanns BJ, Gaigalat L, Goesmann A, Hartmann M, Huthmacher K, Krämer R, Linke B, McHardy AC, Meyer F, Möckel B, Pfefferle W, Pühler A, Rey DA, Rückert C, Rupp O, Sahm H, Wendisch VF, Wiegrabe I, and Tauch A. (2003) The complete Corynebacterium glutamicum ATCC 13032 genome sequence and its impact on the production of L-aspartate-derived amino acids and vitamins. J. Biotechnol. 104:5–25. 20. Kubena BD, Luecke H, Rosenberg H, and Quiocho FA. (1986) Crystallization and X-ray diffraction studies of a phosphate-binding protein involved in active transport in Escherichia coli. J. Biol. Chem. 261:7995–7996. 21. Lahooti M, Pragai Z, and Harwood CR. (2000) Phosphate regulation. In Schumann W, Ehrlich SD, and Ogasawara N (Eds.), Functional Analysis of Bacterial Genes: A Practical Manual. Wiley, Chichester, pp. 234–244. 22. Lambert C, Weuster-Botz D, Weichenhain R, Kreutz EW, de Graaf AA, and Schoberth SM. (2002) Monitoring of inorganic polyphosphate dynamics in Corynebacterium glutamicum using a novel oxygen sparger for real time P-31 in vivo NMR. Acta Biotechnol. 22:245–260. 23. Lebens M, Lundquist P, Soderlund L, Todorovic M, and Carlin NIA. (2002) The nptA gene of Vibrio cholerae encodes a functional sodium-dependent phosphate cotransporter homologous to the type II cotransporters of eukaryotes. J. Bacteriol. 184:4466–4474. 24. Lefevre P, Braibant M, DeWit L, Kalai M, Roeper D, Grotzinger J, Delville JP, Peirs P, Ooms J, Huygen K, and Content J. (1997) Three different putative phosphate transport receptors are encoded by the Mycobacterium tuberculosis genome and are present at the surface of Mycobacterium bovis BCG. J. Bacteriol. 179:2900–2906. 25. Moura RS, Martin JF, Martin A, and Liras P. (2001) Substrate analysis and molecular cloning of the extracellular alkaline phosphatase of Streptomyces griseus. Microbiology 147:1525–1533. 26. Mukai T, Kawai S, Matsukawa H, Matuo Y, and Murata K. (2003) Characterization and molecular cloning of a novel enzyme, inorganic polyphosphate/ATP-glucomannokinase, of Arthrobacter sp. strain KM. Appl. Environ. Microbiol. 69:3849–3857. 27. Neuhard J and Kelln RA. (1996) Biosynthesis and conversion of pyrimidines. In Neidhardt FC, Curtiss III R, Ingraham JL, Lin ECC, Low KB, Magasanik B, Reznikoff WS, Riley M, Schaechter M and Umbarger HE (Eds.). Escherichia coli and Salmonella: Cellular and Molecular Biology. ASM Press, Washiington, D.C., Vol. 1, pp. 580–599.

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28. Pop O, Martin U, Abel C, and Müller JP. (2002) The twin-arginine signal peptide of PhoD and the TatAd/Cd proteins of Bacillus subtilis form an autonomous Tat translocation system. J. Biol. Chem. 277:3268–3273. 29. Pragai Z and Harwood CR. (2002) Regulatory interactions between the Pho and σBdependent general stress regulons of Bacillus subtilis. Microbiology 148:1593–1602. 30. Pragai Z, Allenby NEE, O’Connor N, Dubrac S, Rapoport G, Msadek T, and Harwood CR. (2004) Transcriptional regulation of the phoPR operon in Bacillus subtilis. J. Bacteriol. 186:1182–1190. 31. Price CW, Fawcett P, Ceremonie H, Su N, Murphy CK, and Youngman P. (2001) Genome-wide analysis of the general stress response in Bacillus subtilis. Mol. Microbiol. 41:757–774. 32. Ramos A, Adham SAI, and Gil JA. (2003) Cloning and expression of the inorganic pyrophosphatase gene from the amino acid producer Brevibacterium lactofermentum ATCC 13869. FEMS Microbiol. Lett. 225:85–92. 33. Rosenberg H, Gerdes RG, and Chegwidden K. (1977) Two systems for uptake of phosphate in Escherichia coli. J. Bacteriol. 131:505–511. 34. Rosenberg H, Gerdes RG, and Harold FM. (1979) Energy coupling to the transport of inorganic phosphate in Escherichia coli K12. Biochem. J. 178:133–137. 35. Schweizer H, Argast M, and Boos W. (1982) Characteristics of a binding proteindependent transport system for sn-glycerol-3-phosphate in Escherichia coli that is part of the Pho regulon. J. Bacteriol. 150:1154–1163. 36. Sun GF, Birkey SM, and Hulett FM. (1996) Three two-component signal-transduction systems interact for Pho regulation in Bacillus subtilis. Mol. Microbiol. 19:941–948. 37. Sutcliffe IC and Harrington DJ. (2002) Pattern searches for the identification of putative lipoprotein genes in Gram-positive bacterial genomes. Microbiology 148:2065–2077. 38. Tanaka S, Lee SO, Hamaoka K, Kato J, Takiguchi N, Nakamura K, Ohtake H, and Kuroda A. (2003) Strictly polyphosphate-dependent glucokinase in a polyphosphateaccumulating bacterium, Microlunatus phosphovorus. J. Bacteriol. 185:5654–5656. 39. Torres A, Juarez MD, Cervantes R, and Espitia C. (2001) Molecular analysis of Mycobacterium tuberculosis phosphate specific transport system in Mycobacterium smegmatis. Characterization of recombinant 38 kDa (PstS-1). Microb. Pathog. 30:289–297. 40. van Veen HW, Abee T, Kortstee GJJ, Konings WN, and Zehnder AJB. (1994) Translocation of metal phosphate via the phosphate inorganic transport system of Escherichia coli. Biochemistry 33:1766–1770. 41. Vasicova P, Abrhamova Z, Nesvera J, Patek M, Sahm H, and Eikmanns B. (1998) Integrative and autonomously replicating vectors for analysis of promoters in Corynebacterium glutamicum. Biotechnol. Tech. 12:743–746. 42. Wanner BL. (1996) Phosphorus assimilation and control of the phosphate regulon. In Neidhardt FC, Curtiss III R, Ingraham JL, Lin ECC, Low KB, Magasanik B, Reznikoff WS, Riley M, Schaechter M and Umbarger HE (Eds.). Escherichia coli and Salmonella: Cellular and Molecular Biology. ASM Press, Washington, D.C., Vol. 1, pp. 561–579. 43. Willsky GR and Malamy MH. (1980) Characterization of two genetically separable inorganic phosphate transport systems in Escherichia coli. J. Bacteriol. 144:356–365. 44. Zalkin H and Nygaard P. (1996) Biosynthesis of purine nucleotides. In Neidhardt FC, Curtiss III R, Ingraham JL, Lin ECC, Low KB, Magasanik B, Reznikoff WS, Riley M, Schaechter M and Umbarger HE (Eds.). Escherichia coli and Salmonella: Cellular and Molecular Biology. ASM Press, Washington, D.C., Vol. 1, pp. 561–579. 45. Zhang HY, Ishige K, and Kornberg A. (2002) A polyphosphate kinase (PPK2) widely conserved in bacteria. Proc. Natl. Acad. Sci. USA 99:16678–16683.

17

Vitamin Synthesis: Carotenoids, Biotin, and Pantothenate G. Sandmann and H. Yukawa

CONTENTS 17.1 Carotenoids in Corynebacteria.....................................................................397 17.1.1 Carotenoid Synthesis in C. glutamicum ..........................................399 17.1.2 Carotenogenic Genes of Corynebacteria .........................................402 17.1.3 Conclusion........................................................................................405 17.2 Biotin and C. glutamicum............................................................................405 17.2.1 Synthesis of Biotin...........................................................................405 17.2.2 Functional Identification of bio Genes ............................................407 17.2.3 The bio Loci in Corynebacterium ...................................................407 17.3 Synthesis of Pantothenate ............................................................................408 17.3.1 Pantothenate Synthesis in C. glutamicum .......................................408 17.3.2 Pantothenate Production ..................................................................411 Acknowledgments..................................................................................................411 References..............................................................................................................412

17.1 CAROTENOIDS IN CORYNEBACTERIA Carotenoids are widespread in bacteria. In some cases, the presence of special derivatives of these pigments is characteristic for an individual taxon. Among the bacterial carotenoids, acyclic C40 compounds derived from neurosporene and lycopene as well as cyclic carotenoids like β-carotene derivatives or isorenieratene with aromatic end groups are found. Specific for bacteria are C30 carotenoids synthesized in parallel to the C40 pathway from shorter prenyl pyrophosphates and C50 carotenoids. As lipophilic molecules, carotenoids are integrated into membranes. There, they may have a protective function as antioxidants or act as a membrane stabilizer. Within the corynebacteria, unpigmented species and those pigmented by carotenoids exist. These carotenoids are predominantly of the C50 type. The Halobacterium species occupy the only other taxon in which C50 carotenoids are found [16]. The carotenoid end products in the biosynthetic pathways of corynebacteria are given in Figure 17.1. They include acyclic as well as cyclic compounds with β-ionone 397

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Carotenoids

Species CH2 OH

HOH2 C

Corynebacterium glutamicum MJ233C Decaprenoxanthin C6 H11O5 CH2O

OH2C C6H11O 5

Arthrobacter spec. M3 Agromyces mediolanus (formerly Flavobacterium dehydrogenans)

Decaprenoxanthin diglucosid

OH

Arthrobacter glacialis HO

OH

Bacterioruberin

Curtobacterium flaccumfaciens (formerly Corynebacterium poinsettiae)

HO

HO

OH

Curtobacterium flaccumfaciens (formerly Corynebacterium poinsettiae)

Bisanhydrobacterioruberin

CH2 OH HOH2C

Cp 450

FIGURE 17.1 C50 carotenoids occurring in different corynebacteria.

and ε-ionone rings. Decaprenoxanthin and its glucoside with two ε-ionone groups are the dominant carotenoids in Corynebacterium glutamicum [28], Agromyces mediolanus, and Arthrobacter spec. M3 [3,22,54]. Another species from the Arthrobacter genus, Arthrobacter glacialis, forms the acyclic bacterioruberin [4]. This carotenoid together with its hydration product bisanhydrobacterioruberin is found in Curtobacterium flaccumfaciens [33]. In addition, this bacterium synthesizes carotenoid Cp 450, which structurally resembles decaprenoxanthin and differs only by two β-ionone instead of ε-ionone groups. Interestingly, among the C. glutamicum isolates, those forming decaprenoxanthin derivatives and those unable to synthesize these carotenoids exist, or those synthesizing them in very weak quantities. An example of this is C. glutamicum ATCC13032, which possesses the carotenogenic gene cluster for the complete pathway of decaprenoxanthin synthesis, but only small amounts of precursor carotenes are present. Obviously, carotenoids are not essential for the growth of C. glutamicum. A similar situation is known for Streptomyces species, with some isolates being carotenogenic and some not. In the case of uncarotenogenic S. griseus, all genes for the synthesis of isorenieratene exist but are not expressed [50]. Another possibility of an incomplete pathway is a mutation in a gene encoding an early enzyme in

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carotenoid biosynthesis. An example is a mutant of C. glutamicum ssp. flavum MJ233C with an inactivated phytoene synthase, which is devoid of carotenoids [24]. Other mutants of this strain accumulate either lycopene or acyclic C45 and C50 precursors of decaprenoxanthin [28]. It should be pointed out that not all carotenogenic corynebacteria are able to synthesize C50 carotenoids. For example, the carotenoids of Brevibacterium linens are the aromatic C40 isorenieratene together with its 3-HO and 3,3’-(HO)2 derivatives [23].

17.1.1 CAROTENOID SYNTHESIS

IN

C.

GLUTAMICUM

The biosynthesis of C50 carotenoids initially proceeds via the typical C40 pathway along to the branch point lycopene (Figure 17.2). The synthesis of lycopene involves a phytoene synthase for the condensation of two molecules of geranylgeranyl pyrophosphate (GGPP) to phytoene and a single desaturase capable of introducing four double bonds into phytoene [47]; GGPP is also the precursor for undecaprenol phosphate (C55) in C. glutamicum [15], required as a lipophilic carrier in cell wall synthesis. In most bacteria, the prenyl pyrophosphate precursors like GGPP are synthesized via the deoxyxylulose 5-phosphate (DXP) pathway rather than via the mevalonate pathway [30]. The absence of a gene for hydroxymethylglutaryl-coenzyme A reductase from the latter pathway and the presence of the genes encoding deoxyxylulose 5-phosphate synthase and deoxyxylulose 5-phosphate reductoisomerase in the genome indicate that in C. glutamicum the DXP pathway supplies the carotenoid precursors. This conclusion is supported by the accumulation of 2-Cmethyl-D-erythritol-2,4-cyclopyrophosphate, an intermediate of the DXP pathway in C. ammoniagenes [39]. The detailed reaction sequence to decaprenoxanthin diglucoside fatty acid esters has been elucidated in C. glutamicum MJ233C by the identification of several intermediates in carotenogenic mutants [28] and by stepwise establishment of the final part of the pathway in Escherichia coli by introduction of carotenogenic genes [24]. Lycopene is the C40-carotenoid substrate for extension of the carbon chain to yield mature carotenoids (Figure 17.2). The intermediates nonaprene, sarcinene, and decaprenoxanthin were identified in an overproducing intensively yellow mutant and the wild-type of C. glutamicum MJ233C. Thus, the following reaction sequence can be proposed. Lycopene is converted into the C45-carotenoid nonaprene by addition of a C5 unit and subsequently cyclized. At the other side of the molecule, a second addition of another C5 unit and cyclization results in the formation of the C50carotenoid sarcinene. In the next step, hydroxylation leads via dehydroxydecaprenoxanthin to decaprenoxanthin followed by glucosylations of the hydroxy groups to decaprenoxanthin monoglucoside and decaprenoxanthin diglucoside (DDG), the main carotenoids of C. glutamicum. A final esterification at the glucosides yields DDG monoester and DDG diester, which were found in small amounts in C. glutamicum MJ233C. The esterified fatty acids are C16:0 and C18:1. The analysis of a pink mutant yielded interesting information of the first steps of decaprenoxanthin synthesis [28]. In the mutant, the acyclic carotenoids nonaflavuxanthin with 45 carbon atoms and flavuxanthin with 50 carbon atoms were identified (Figure 17.3A). Their occurrence is a strong indication against a simultaneous

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FIGURE 17.2 Biosynthesis of decaprenoxanthin in wild-type C. glutamicum MJ233C (adapted from Krubasik et al. [28] with permission). The genes of the pathway have been identified except those encoding the enzymes for C-4″/C-4 hydroxylation, glucosylation and glucoside ester formation.

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A +C5

Lycopene

hydroxylation

+C5

Nonaflavuxanthin

hydroxylation

HOH2C

CH2 OH

Flavuxanthin

HOH2C4′′

B 1 2

+

R

R 2

+

- PP - H+

+ OH

DMAPP 16

1

R

to Decaprenoxanthin

HO

R

to Bacteriorubin

FIGURE 17.3 Carotenoid biosynthesis in the carotenogenic mutant MV70 of C. glutamicum MJ233C (A) and putative mechanism of the elongation step (B) (adapted from Krubasik et al. [28] with permission). DMAPP: dimethylallyl pyrophosphate, PP: the released pyrophosphate, and R: the remainder of the lycopene backbone.

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elongation/cyclization reaction. Thus, the initial step toward decaprenoxanthin is the elongation of the lycopene molecule by C5 units at each end. As indicated in Figure 17.3B, this reaction starts by an electrophilic attack of an isoprenoid cation at the C-1,2 double bond of lycopene. The resulting postulated carbocation with an attached C5-side chain at C-2 is stabilized by proton abstraction, yielding first C45 nonaflavuxanthin with one C-1,16 double bond and then the symmetrical elongated C50 flavuxanthin when the same enzymatic reaction modifies the other side of the molecule. The mechanism of this elongation reaction resembles in principle the mechanism for elongation in the synthesis of bacterioruberin (see structure in Figure 17.1). In the latter case, formation of the prenylated carbocation from lycopene should be initially the same but its stabilization proceeds via the addition of OH– at C-1 and C-1′, respectively, instead of proton abstraction (Figure 17.3B). Bacterioruberin, which differs from flavuxanthin only by the absence of the C-1,16 and C-1′,16′ double bonds and the presence of hydroxyl groups at C-1 and C-1′ (Figure 17.1), was identified as end product of the carotenoid pathway in Curtobacterium flaccumfaciens [33] and A. glacialis [4]. The mechanism of subsequent formation of an ε-end group from flavuxanthin in the pathway to decaprenoxanthin may be similar to the lycopene cyclization reaction. Protonation of the C-1,16 double bond should be the initial reaction during ring formation. In the wild-type, hydroxylations at C-4″ and C-4 (Figure 17.2) follows the formation of the ε-end groups, whereas the accumulating acyclic precursors are also hydroxylated in the pink mutant.

17.1.2 CAROTENOGENIC GENES

OF

CORYNEBACTERIA

Before the complete genome sequence of C. glutamicum ATCC13032 was established, a carotenogenic gene cluster was cloned from strain MJ233C carotenoid transposon mutants and all crt genes were therein functionally identified [24]. The genes for the three steps in the carotenoid pathway from GGPP to lycopene — crtE encoding a GGPP synthase, crtB encoding a phytoene synthase, and crtI encoding a phytoene desaturase — were identified by sequence similarities to homologous genes from other organisms (Figure 17.4) and their function assigned by the involvement of their products in lycopene synthesis [27]. Heterologous expression of a plasmid with these three genes demonstrated that their products are responsible for lycopene synthesis. It also showed that one open-reading frame within the gene cluster is not necessary for carotenoid biosynthesis in C. glutamicum MJ233C (Figure 17.4). This gene shares some similarities with lipid transport proteins from mycobacteria. One could speculate that its product may be involved in carotenoid transportation. The products of another three genes are involved in the subsequent biosynthetic steps from lycopene to decaprenoxanthin. Their individual functions were elucidated in a lycopene-producing E. coli strain (Figure 17.5). With crtEb alone, a lycopeneproducing E. coli transformant converts lycopene via the C45 intermediate nonaflavuxanthin to the acyclic C50 carotenoid flavuxanthin by addition of two C5 isoprenoid groups at position C-2 and C-2′. Expression of gene crtYe together with crtYf has no effect on lycopene conversion. However, these genes mediate the formation of

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403

Corynebacterium glutamicum crtE

crtYe crtYf

crtI

crtB

crtEb

Corynebacterium diphteriae crtB

crtI

Agromyces mediolanus idi

IPP isomerase

crtE

crtB

crtYe crtYf

crtI

Phytoene desaturase

GGPP synthase Phytoene synthase

crtEb

Lycopene elongase

C 45/50 Carotenoid ε-cyclase

FIGURE 17.4 Organization of carotenogenic genes in the genomes of corynebacteria. Information originates from data bases, Jessen et al. [22], and Kimura et al. [24].

Carotenoids (μg/g dw)

Genes expressed

crtYf crtYe

L

N

F

crtEb

18.9

2.2

87.6

crtEb

15.7

1.7

50.8

crtEb

9.7

0.8

42.7

crtYe crtYf crtYe crtYf

D

DMG DDG

24.1

crtEb

35.1

56.6

40.8

FIGURE 17.5 Carotenoids formed by a lycopene-accumulating E. coli strain transformed with plasmids carrying combinations of the genes crtEb, crtYe, and crtYf (adapted from Kimura et al. [24] with permission). L: lycopene, N: nonaflavuxanthin, F: flavuxanthin, D: decaprenoxanthin, DMG: decaprenoxanthin monoglucoside, DDG: decaprenoxanthin diglucoside.

decaprenoxanthin derivatives in the presence of crtEb (necessary for the synthesis of flavuxanthin). The combination of crtYe with crtYf is decisive. One of these gene products alone is not sufficient to catalyze the cyclization to decaprenoxanthin. Therefore, both gene products encode the carotenoid C45/50 ε-cyclase.

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The arrangement of the carotenogenic genes as shown in Figure 17.4 for C. glutamicum ATCC 13032 is identical in the C. glutamicum strain MJ233C and C. efficiens. In addition, in the corynebacteria analyzed, the crt genes of the operon share the same direction of transcription and the arrangement and order of the genes is the same. Another phytoene synthase-like gene (NCgl2347) is present in the genome of C. glutamicum ATCC13032. Since phytoene and squalene synthases share significant similarities, it may encode a squalene synthase. In Agromyces mediolanus, crtE and crtB are not separated by an open-reading frame unrelated to carotenogenesis [22]. In addition, the idi gene encoding isopentenyl pyrophosphate isomerase — the initial enzyme for the formation of prenyl pyrophosphates as precursors of carotenoids and other terpenoids — is part of the operon. Common to all species are the adjacent genes crtYe and crtYf, together with the elongase gene crtEb at the downstream end of the operon. In C. diphteriae, only a small carotenogenic operon, including a putative phytoene synthase and desaturase, is present. The formation of carotenoids has not been reported for this species. However, if these genes were expressed, they could mediate carotenoid formation up to lycopene only. The lycopene elongase CrtEb is a C5 prenyl unit transferring enzyme as concluded from the products made in vivo in the heterologous host E. coli (Figure 17.5). It belongs to the UbiA prenyl transferases and thus shares structural motifs with other prenyl transferases utilizing allylic prenyl pyrophosphates. These include not only prenyl transferases from Methanobacterium thermoautotrophicum and Synechocystis sp. PCC6803, but also a bacteriochlorophyll synthase from Heliobacillus mobilis, an octaprenyl transferase from Bacillus subtilis, a farnesyl pyrophosphate synthase from rat, and the geranylgeranyl pyrophosphate synthases from Rhodobacter capsulatus and Erwinia uredovora. The C45/50 carotenoid ε-cylcase is a heterodimeric protein consisting of CrtYe and CrtYf. These proteins exhibit sequence similarities with CrtYc and CrtYd of Brevibacterium linens [27] and Mycobacterium aurum [56], where they were characterized for the first time as a special type of cyclase. In contrast to the cyclase in C. glutamicum, this cyclase is specific for C40 carotenoids and forms a β-ionone end group. These differences in substrate specificity and ring formation are reflected at the sequence level. Whereas the CrtYe and CrtYc polypeptides of the two cyclases exhibit high identities among each other, this is not the case for CrtYf and CrtYd. Therefore, these latter polypeptides could well be responsible for one of the specific features of the two types of cyclases. Interestingly, Sulfolobus solfataricus possesses a lycopene β-cyclase in which the two polypeptides are fused but separated in corynebacteria although the arrangement of the domains is retained. Surprisingly, hydroxylated C50 carotenoids were formed in the complementation experiments with the lycopene-producing E. coli strain expressing crtEb of C. glutamicum, and C50 carotenoid glucosides were obtained in combination with crtYe and crtYf (Figure 17.5). These results and the absence of genes with sequence similarities to known hydroxylase and glucosylase genes from the crt gene cluster of C. glutamicum may suggest that these modifications are catalyzed by unspecific enzymes that are present in C. glutamicum and E. coli.

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17.1.3 CONCLUSION The synthesis of C50 carotenoids is restricted to a small number of bacteria. The most abundant C50 carotenoid, decaprenoxanthin including its glucosides and glucoside fatty acid esters, is characteristic for corynebacteria like C. glutamicum ssp. flavum MJ233C. Other C50 carotenoids are acyclic or contain β-ionone instead of ε-ionone groups. Important enzymes for the final steps of the pathway are an elongase that initiates the formation of decaprenoxanthin by the extension of the C40 intermediate lycopene and a new type of C45/50 carotenoid ε-cylcase consisting of two polypeptides that together catalyze the cyclization reaction. The organization of the carotenogenic genes in the genome of the corynebacteria analyzed is rather uniform. Although the steps from lycopene to decaprenoxanthin are now well understood, comparatively little is known about the C-4″/C-4 hydroxylation and the decaprenoxanthin modification reactions as well as the genes involved.

17.2 BIOTIN AND C. GLUTAMICUM Biotin is a colorless water-soluble member of the B-complex vitamins. Its skeleton consists of a bi-heterocyclic core (a ureido ring fused to a sulfur-containing ring) to which a carboxybutyl side chain is attached [10] (Figure 17.6). Biotin is of particular interest for C. glutamicum since biotin auxotrophy resulted in its discovery as a glutamate producer. Understanding how this mechanism leads to L-glutamate overproduction is still a major challenge (see Chapter 19). In addition, recent work attempts to use this biotin auxotrophy to uncouple the growth of C. glutamicum from its biosynthetic capacities. The reason is that C. glutamicum is rather unsusceptible to autolysis, thus enabling the reuse of C. glutamicum in production processes. For instance, C. glutamicum strain R can be used up to 30 times for the conversion of precursors to L-isoleucine [54], L-valine [55] or L-aspartate [58]. In C. glutamicum, biotin is present as a covalently linked prosthetic group in just two polypeptides. This is AccBC, the α-subunit of acetyl-CoA carboxylase [21] and the pyruvate carboxylase [41].

17.2.1 SYNTHESIS

OF

BIOTIN

Biotin biosynthesis has mainly been studied in Escherichia coli but also in other organisms. The pathway from pimeloyl-CoA is apparently identical in all (Figure 17.6). However, the steps leading to pimeloyl-CoA, as well as the final step of biotin formation, are not yet fully characterized. The known synthesis of biotin beginning from pimeloyl-CoA requires 7-keto-8-aminopelargonic acid synthase activity (bioF) to condense the activated pimelate with L-alanine [42]. This enzyme, whose three-dimensional structure is known, as is that of the two subsequent enzymes [49], shares similarities with other acyl-CoA synthases. The next step is the conversion to 7,8-diaminopelargonic acid by the 7,8-diaminopelargonic acid synthase (bioA). This enzyme is a unique amino transferase since it utilizes S-adenosyl-L-methionine as an amino group donor. The structure of the protein is very similar

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NH3+

O −O

COO−

SCoA +

Pimeloyl CoA + L-Alanin

bioF

KAPA synthase

7-Keto-8-aminopelargonic acid (KAPA) DAPA synthase SAM

bioA

7,8-Diaminopelargonic acid (DAPA) DTB synthetase ATP, CO2

bioD

Dethiobiotin (DTB) Biotin synthase SAM, NADPH, S

bioB O NH H H

HN H S

O O−

Biotin

FIGURE 17.6 The biotin biosynthesis pathway. The genes given in boldface are present and functionally active in C. glutamicum.

to the structure of the 7-keto-8-aminopelargonic acid synthase, suggesting an evolutionary relationship between the two proteins. In the subsequent reaction, the dethiobiotin synthetase (bioD) introduces the ureido ring into the molecule [37]. The final step is the conversion of dethiobiotin to biotin by biotin synthase. This reaction is poorly understood. In addition to biotin synthase, other proteins or components might be involved since a sulfur atom is introduced and an additional reducing system is required [5,20,48]. The bioB encoded biotin synthase belongs to the family of AdoMet-dependent enzymes that reductively cleave AdoMet into a deoxyadenosyl radical and that is also responsible for the homolytic cleavage of C-H bonds [32]. As mentioned, not only the mechanism of this last step in biotin synthesis, but also the initial step providing pimeloyl-CoA as a substrate is unclear. It could be synthesized by a modified fatty acid pathway starting with a malonyl thioester [32]. Several genes have been identified to act in the biotin synthesis pathway upstream of pimeloyl-CoA biosynthesis, such as the bioC and bioH genes of E. coli and S. typhimurium [1] or bioW and bioI of B. subtilis, the latter encoding a cytochrome P450 [17] that complements mutations in the bioC or bioH genes of

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407

E. coli [6]. Pimelate can be converted to pimeloyl-CoA by the enzyme 6-carboxyhexanoate-CoA ligase [42]. Biotin is covalently linked via a specific lysine residue to proteins like AccBC and pyruvate carboxylase by the action of BirA. In C. glutamicum, BirA is encoded by NCgl0679 and that of C. efficiens by CE0740. Interestingly, in both organisms birA is directly adjacent, but divergently transcribed, to dtsR1 and dtsR2, which appear to encode β-subunits interacting with the biotin-containing AccBC, supposedly the α-subunit of acyl-CoA carboxylases [24]. In addition to the enzymatic function, the BirA proteins of B. subtilis and E. coli regulate the transcription of the biotin operon [40]. The domain responsible for regulation contains an HTH-motif of GntR-like bacterial transcription factors. However, the corynebacterial BirA is at variance in the amino-terminus of the polypeptide although the other part of BirA shares identities of more than 29% with the corresponding E. coli part. Not only the capability to synthesize biotin (see below), but also the property to regulate bio genes, might be lost in C. glutamicum.

17.2.2 FUNCTIONAL IDENTIFICATION

OF BIO

GENES

Although C. glutamicum requires biotin for growth, early experiments indicated that supplementation with aminopelargonic acid derivatives to C. glutamicum can compensate for biotin [38]. An understanding of this phenomenon was obtained by detailed cross-feeding experiments between C. glutamicum and defined E. coli bio mutants [19]. The three different C. glutamicum strains used were ssp. flavum MJ233, ssp. lactofermentum ATCC13869, and C. glutamicum ATCC31831, which behaved identically. The experiments verified that C. glutamicum is still able to perform the individual reactions of the pathway starting from 7-keto-8-aminopelargonic acid. This might suggest that genome arrangements occurred during evolution resulting in the loss of some bio genes and the inability to synthesize biotin de novo. The genes identified from C. glutamicum by complementation of E. coli are bioA, bioD, and bioB, whereas neither bioF, bioH, nor bioC could be identified [18]. Since the latter two genes are involved in the synthesis of pimeloyl-CoA, it appears that the entire first part of the synthesis pathway is absent. The BioB protein is strongly conserved in C. glutamicum, C. efficiens, and M. tuberculosis. The proteins exhibit exceptional high identities of more than 71% among each other, whereas the B. subtilis or E. coli BioB share, at best, 37% identities with the protein of C. glutamicum.

17.2.3 THE

BIO

LOCI

IN

CORYNEBACTERIUM

In the genomes of C. glutamicum ATCC13032, C. efficiens, and C. diphteriae, bioB is always separated from the clustered bioA and bioD. This is also the case in C. glutamicum R [29], whose genome sequence consists of 3.314 Mbp and therefore is of a size similar to that of strain ATCC 13032, with 3.309 Mbp. In C. efficiens, the genes bioA and bioD overlap by four nucleotides, which is strong indication that these two genes form a transcriptional unit. This organization is also present in the other Corynebacterium strains. However, in M. tuberculosis, bioA and bioD are

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separated by bioF. These three genes together might form a transcriptional unit, as they do possibly also in M. marinum and Brevibacterium linens. As mentioned earlier, the bioB gene is separately transcribed. However, in all Corynebacterianeae, including Mycobacterium tuberculosis, M. bovis, M. marinum, and M. leprae, a small orf encoding a peptide that is 71 amino acyl residues in size in C. glutamicum is present. Since this arrangement is retained irrespective of the genomic context and since bioB overlaps with the small orf in some Corynebacterianeae, a functional connection of the orf with BioB is suggested. Because C. glutamicum is auxotroph for biotin, an interesting question is how biotin enters the cell. In Sinorhizobium meliloti, the gene bioM appears to be connected to biotin transport [14] and a similar gene is present in C. glutamicum (NCgl1884) encoding the ATP-binding protein of an ABC carrier. Downstream of bioM, a second gene encoding the permease of an ABC transporter is present. Even more intriguing is the fact that upstream of the bioM homolog bioY is located. The BioY protein is involved in bioconversion of pimelate into dethiobiotin although the exact function of the protein is unknown. The operon, likely to be formed by these three genes, is conserved in the Corynebacterium species glutamicum, efficiens, and diphteriae, which is strong evidence that the genes are related to biotin metabolism.

17.3 SYNTHESIS OF PANTOTHENATE D-Pantothenate (vitamin B5) is made from L-aspartate and 2-ketoisovalerate, the latter being also an intermediate of L-valine and L-leucine synthesis (Figure 17.7). Since 2-ketoisovalerate itself is made by the same enzymes synthesizing the L-isoleucine precursor 2-ketomethylvalerate, D-pantothenate synthesis is in fact linked to the synthesis of all three branched-chain amino acids. The pathway as given in Figure 17.7 including the genes of the respective enzymes has been established for E. coli and genome analysis revealed that in diverse eubacteria these genes are present. However, this is not true for archaea, in which only panB and panE genes have been found, and for plants and fungi, in which only panB and panC are present [31]. Thus, the final part in the pathway starting from ketopantoate and L-aspartate is at variance in different organisms. Indeed, at least three mechanisms of β-alanine formation are thought to be present in microorganisms [52]. In E. coli, the first reaction catalyzed by the ketopantoatehydroxymethyl transferase uses tetrahydrofolate and 2-ketoisovalerate to generate ketopantoate, which is reduced to D-pantoic acid. An aspartate-α-decarboxylase activity generates β-alanine, which is ligated with pantoic acid to yield D-pantothenate.

17.3.1 PANTOTHENATE SYNTHESIS

IN

C.

GLUTAMICUM

The ketopantoatehydroxymethyl transferase of C. glutamicum is encoded by panB. Its gene product of 28.5 kDa exhibits highest identities of 52% with that of M. tuberculosis, and enzyme activity determinations resulting in 12-fold-increased transferase activity with overexpressed panB verified its function [46]. The enzyme

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409

FIGURE 17.7 The biosynthesis of D-pantothenate. The genes given in boldface are present and functionally active in C. glutamicum. A functional ketopantoic acid reductase as typical for E. coli is not present in C. glutamicum.

of C. glutamicum appears not to be inhibited by D-pantothenate or D-pantoate as is the case in E. coli. As mentioned earlier, the further conversion of ketopantoate is at variance among organisms. Indeed, a very special situation is present in C. glutamicum. No specific enzyme exists, but the ilvC-encoded isomeroreductase catalyzing the reduction of either acetohydroxybutyrate or acetolactate (Figure 17.7) catalyzes also the reduction

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of ketopantoate. Consequently, when ilvC is inactivated, the reductase activity with ketopantoate as substrate is reduced from 12.6 nmol⋅min–1⋅mg–1 protein to less than 0.2 nmol⋅min–1⋅mg–1 protein [33]. Furthermore, growth of the ilvC mutant (supplied with L-isoleucine and 2-ketoisovalerate) is dependent on D-pantothenate, thus demonstrating that IlvC in vivo serves the generation of D-pantothenate. This is in accordance with the observation that also in E. coli IlvC exhibits ketopantoate reductase activity [13,43] and that no phenotype of a panE mutant exists. The basis for this overlapping specificity is on the one hand the striking structural resemblance of α-ketopantoate to the isomerized intermediates of the L-isoleucine and L-valine synthesis. On the other hand, IlvC shares structural similarities with PanE proteins of a number of bacteria, in particular to the N-terminal domain of PanE [31]. The rate of reduction of α-ketopantoate of the S. typhimurium isomeroreductase is 1/20 as compared with that with α-acetolactate as substrate [43]. The distinct physiological role of IlvC in C. glutamicum consisting in the utilization of three different substrates required for vitamin and amino acid synthesis might explain the transcriptional control of the ilvC gene. This gene forms together with ilvB and ilvN an operon [9]. Although ilvC occupies the third position in this operon, it is present on the three different transcripts formed and its transcript level is the most abundant [23]. This may ensure adequate IlvC formation and high activity of this enzyme even when a surplus of branched-chain amino acids is present leading to an attenuated expression of the operon [34]. The pantothenate ligase encoded by panC catalyses the ATP-dependent condensation of pantoic acid with β-alanine. The corresponding gene of C. glutamicum encodes a polypeptide of 35.8 kDa, and its ligase function has been demonstrated in enzyme assays leading to a 13-fold increased activity with plasmid encoded panC. [46]. Interestingly, ligase activity was obtained only with fragments containing panC together with the 5′-located panB gene. This, and the fact that both genes are overlapping by one nucleotide is strong evidence that both genes form an operon. This organization is also present in C. efficiens and Brevibacterium linens. In several bacteria, such as Bacillus and Salmonella species, panD is located as a third gene directly downstream of panBC. The β-alanine required for the final step of D-pantothenate synthesis is generated by L-aspartate-α-decarboxylase. The corresponding panD gene of C. glutamicum was cloned by heterologous complementation [11]. Its inactivation resulted in β-alanine auxotrophy, and its overexpression resulted in a more than 200-fold increase in decarboxylase activity. However, the PanD polypeptide itself does not possess enzyme activity. Instead, and as is known from E. coli [45], the inactive proenzyme (the π-protein, PanD), is at least in part autocatalytically cleaved and processed. This yields a β-subunit with XOH at its C-terminus and an α-subunit with a pyruvoyl group at its N-terminus, derived from serine at the cleavage site. The cleavage site between Gly-24 and Ser-25 is strictly conserved in the 14.1-kDa PanD polypeptide of C. glutamicum, and gel analyses proved the formation of the larger α-subunit of 11.5 kDa from the C. glutamicum π-protein. The active enzyme is a tetramer consisting of α-subunits with the catalytic pyruvoyl groups necessary for decarboxylation at three active sites and an ester resulting from the autocatalytic self-processing at the fourth [2].

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411

17.3.2 PANTOTHENATE PRODUCTION D-Pantothenate

is produced with E. coli strains derived by undirected mutagenesis. They contain plasmid-encoded panBC and have a deregulated 2-ketoisovalerate supply. Upon addition of the cheap β-alanine, they excrete D-pantothenate into the medium in concentrations exceeding 60 g l–1 [35]. Already with E. coli K12, panE overexpression results in an approximately threefold increased formation of D-pantothenate up to 4.3 mg (g dry weight)–1 [13]. Similarily, with panD overexpressed in an E. coli wild-type, an increased accumulation of D-pantothenate can be obtained [11]. Notable is the profitable use of the heterologous gene of C. glutamicum as compared with that of E. coli, which could be due to an increased processing of the π-protein and therefore a higher specific activity of the C. glutamicum enzyme [11]. Brevibacterium ammoniagenes, has already been reported to accumulate coenzyme A, which is synthesized from D-pantothenate [51,52]. To study the capability of C. glutamicum for D-pantothenate accumulation, a strain with increased flux toward 2-ketoisovalerate was made by overexpressing ilvBNCD in an ilvA background [46]. Such a strain accumulates 0.87 μM D-pantothenate, whereas the wildtype accumulates about 0.04 μM. Upon addition of β-alanine, the same strain reaches 0.53 mM, illustrating the strict control and limiting flux via aspartate-α-decarboxylase activity. In fact, panD overexpressed alone in the wild-type results already in an approximately 60-fold increase in D-pantothenate accumulation [53]. Furthermore, D-pantothenae accumulation increases from 0.53 mM to 4.2 mM when the panBC genes and the ilvBNCD genes are concomitantly overexpressed. This illustrates the feasibility of the engineering approach to increase flux. Since 2-ketoisovalerate is the ultimate precursor for L-valine, the strain overexpressing ilvBNCD in the ilvA background was also used to direct the flux to L-valine by panBC deletion, which yielded L-valine concentrations of up to 91 mM [44]. Therefore, an even higher formation of D-pantothenate should be achieved if it would be possible to direct the entire 2-ketoisovalerate flux toward D-pantothenate. This conclusion is supported by flux analysis [8]. In a fed-batch cultivation, the strain accumulating in batch 0.92 g l–1 D-pantothenate accumulated 1.53 g l–1 of the vitamin, but at the same time unidentified compounds were found [7]. Therefore, still-unknown limitations might exist that prevent full conversion of ketoisovalerate to D-pantothenate by C. glutamicum. One option would be limiting the export; indeed, deletion of the branched-chain amino acid aminotransferase ilvE and overexpression of ilvBNCD and panBC did not result in concentrations exceeding 5.8 mM [12], although no 2ketoisovalerate can be converted to L-valine. This is evidence for limiting export, but further analyses are required to confirm this suggestion. The in-depth analysis of the D-pantothenate accumulation of C. glutamicum shows the enormous synthesis capability of this organism but also demonstrates the advantage and possibly the necessity to use undirected methods to achieve high-level production strains.

ACKNOWLEDGMENTS H. Y. thanks A. A. Vertès and M. Inui for help during preparation of the manuscript.

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REFERENCES 1. Akatsuka H, Kawai E, Sakurai N, and Omori K. (2003) The Serratia marcenscens bioH gene encodes an esterase. Gene 302:1–2. 2. Albert A, Dhanaraj V, Genschel U, Khan GL, Ramjee MK, Pulido R, Sibanda BL, von Delft F, Witty M, Blundell TL, Smith AG, and Abell C. (1998) Crystal structure of aspartate decarboxylase at 2.2 angstrom resolution provides evidence for an ester in protein self-processing. Nature Struct. Biol. 5:289–293. 3. Arpin N, Fiasson J, Norgard S, Borch G, and Liaaen-Jensen S. (1972) Bacterial carotenoids, XLVI. C50-carotenoids 14. C50-Carotenoids from Arthrobacter glacialis, Acta Chem. Scand. B 29:921–929. 4. Arpin N, Liaaen-Jensen S, and Trouilloud M. (1972) Bacterial carotenoids. XXXVIII C50-carotenoids. 9. Isolation of decaprenoxanthin mono- and diglucoside from an Arthrobacter sp., Acta Chem. Scand. 26:2524–2526. 5. Birch OM, Fuhrmann M, and Shaw NM. (1995) Biotin synthase from Escherichia coli, an investigation of the low molecular weight and protein components required for activity in vitro. J. Biol. Chem. 270:19158–19165. 6. Bower S, Perkins J, Yocum RR, Serror P, Sorokin A, Rahaim P, Howitt CL, Prasad N, Ehrlich SD, and Pero J. (1995) Cloning and characterization of the Bacillus subtilis birA gene encoding a repressor of the biotin operon. J. Bacteriol. 177:2572–2575. 7. Chassagnole C, Diano A, Létisse F, and Lindley ND. (2003) Metabolic network analysis during fed-batch cultivation of Corynebacterium glutamicum for pantothenic acid production: first quantitative data and analysis of by-product formation. J. Biotechnol. 104:261–272. 8. Chassagnole C, Létisse F, Diano A, and Lindley ND. (2002) Carbon flux analysis

in a pantothenate overproducing Corynebacterium glutamicum strain. Mol. Biol. Rep. 29:129–134. 9. Cordes C, Möckel B, Eggeling L, and Sahm H. (1992) Cloning, organization and functional analysis of ilvA, ilvB and ilvC genes from Corynebacterium glutamicum. Gene 112:113–116. 10. De Clercq PJ. (1997) Biotin: A timeless challenge for total synthesis. Chem. Rev. 97:1755–1792. 11. Dusch N, Pühler A, and Kalinowski J. (1999) Expression of the Corynebacterium glutamicum panD gene encoding L-aspartate-alpha-decarboxylase leads to pantothenate overproduction in Escherichia coli. Appl. Environ. Microbiol. 65:1530–1539. 12. Eggeling L and Sahm H. (2002) Method for the production of D-pantothenic acid by fermentation. Patent No. WO 02/055711 A2. 13. Elischewski F, Pühler A, and Kalinowski J. (1999) Pantothenate production in Escherichia coli K12 by enhanced expression of the panE gene encoding ketopantoate reductase. J. Biotechnol. 75:135–146. 14. Entcheva P, Phillips DA, and Streit WR. (2002) Functional analysis of Sinorhizobium meliloti genes involved in biotin synthesis and transport. Appl. Environ. Microbiol. 68:2843–2848. 15. Gibson KJ, Eggeling L, Maughan WN, Krumbach K, Gurcha SS, Nigou J, Puzo G, Sahm H, and Besra GS. (2003) Disruption of Cg-Ppm1, a polyprenol monophosphomannose synthase and the generation of lipoglycan-less mutants in Corynebacterium glutamicum. J. Biol. Chem. 278:40842–40850. 16. Goodwin TW. (1980) The Biochemistry of Carotenoids, Vol. 1, Chapman and Hall, London.

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17. Green AJ, Rivers SL, Cheeseman M, Reid GA, Quaroni LG, Macdonald ID, Chapman SK, and Munro AW. (2001) Expression, purification and characterization of cytochrome P450 Biol: a novel P450 involved in biotin synthesis in Bacillus subtilis. J. Biol. Inorg. Chem. 6:523–533. 18. Hatakeyama K, Hohama K, Vertes AA, Kobayashi M, Kurusu Y, and Yukawa H. (1993) Genomic organization of the biotin biosynthetic genes of coryneform bacteria: cloning and sequencing of the bioA-bioD genes from Brevibacterium flavum. DNA Seq. 4:177–184. 19. Hatakeyama K, Kohama K, Vertes AA, Kobayashi M, Kurusu Y, and Yukawa H. (1993) Analysis of the biotin biosynthesis pathway in coryneform bacteria: cloning and sequencing of the bioB gene from Brevibacterium flavum. DNA Seq. 4:87–93. 20. Ifuku O, Kishimoto J, Haze S, Yanagi M, and Fukushima S. (1992) Conversion of dethiobiotin to biotin in cell-free extracts of Escherichia coli. Biosci. Biotechnol. Biochem. 56:1780–1785. 21. Jäger W, Peters-Wendisch PG, Kalinowski J, and Pühler A. (1996) A Corynebacterium glutamicum gene encoding a two-domain protein similar to biotin carboxylases and biotin-carboxyl-carrier proteins. Arch. Microbiol. 166:76–82. 22. Jessen HJ, deSouza ML, Amore F, Gokarn RR, and Schroeder WA. (2002) Cloning, characterization and expression of a gene cluster involved in synthesis of a C50 carotenoid from Agromyces mediolanus ATCC 13930, Proc. 13th Intern. Carotenoid Symposium, Hawaii, p. 152. 23. Keilhauer C, Eggeling L, and Sahm H. (1993) Isoleucine synthesis in Corynebacterium glutamicum: molecular analysis of the ilvB–ilvN–ilvC operon. J. Bacteriol. 175:5595–5603. 24. Kimura E, Abe C, Kawahara Y, Nakamatsu T, and Tokuda H. (1997) A dtsR genedisrupted mutant of Brevibacterium lactofermentum requires fatty acids for growth and efficiently produces L-glutamate in the presence of an excess of biotin. Biochem. Biophys. Res. Commun. 234:157–161. 25. Kohl W, Achenbach H, and Reichenbach H. (1983) The pigments of Brevibacterium linens: aromatic carotenoids. Phytochemistry 22:207–210. 26. Krubasik P, Kobayashi M, and Sandmann G. (2001) Expression and functional analysis of a gene cluster involved in the synthesis of decaprenoxanthin reveals the mechanisms for C50 carotenoid formation. Eur. J. Biochem. 268:3702–3708. 27. Krubasik P and Sandmann G. (2000) A carotenogenic gene cluster from Brevibacterium linens with novel lycopene cyclase genes involved in the synthesis of aromatic carotenoids. Mol. Gen. Genet. 263:423–432. 28. Krubasik P, Takaichi S, Maoka T, Kobayashi M, and Sandmann G. (2001) Detailed biosynthetic pathway to decaprenoxanthin diglucoside in Brevibacterium flavum and identification of novel intermediates. Arch. Microbiol. 176:217–223. 29. Kurusu Y, Kainuma M, Inui M, Satoh Y, and Yukawa H. (1990) Electroporationtransformation system for coryneform bacteria by auxotrophic complementation. Agric. Biol. Chem. 54:443–447. 30. Lange BM, Rujan T, Martin W, and Croteau R. (2001) Isoprenoid biosynthesis: the evolution of two ancient and distinct pathways across genomes. Proc. Natl. Acad. Sci. USA 97:13172–13177. 31. Lobley CM, Schmitzberger F, Kilkenny ML, Whitney H, Ottenhof HH, Chakauya E, Webb ME, Birch LM, Tuck KL, Abell C, Smith AG, and Blundell TL. (2003) Structural insights into the evolution of the pantothenate-biosynthesis pathway. Biochem. Soc. Trans. 31:563–571.

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32. Marquet A, Bui BT, and Florentin D. (2001) Biosynthesis of biotin and lipoic acid. Vitam. Horm. 61:51–101. 33. Merkamm M, Chassagnole C, Lindley ND, and Guyonvarch A. (2003) Ketopantoate reductase activity is only encoded by ilvC in Corynebacterium glutamicum. J. Biotechnol. 104:253–260. 34. Morbach S, Junger C, Sahm H, and Eggeling L. (2000) Attenuation control of ilvBNC in Corynebacterium glutamicum: evidence of leader peptide formation without the presence of a ribosome binding site. J. Biosci. Bioeng. 90:501–507. 35. Moriya T, Hikichi Y, Moriya Y, and Yamaguchi T. (1996) Process for producing D-pantoic acid and D-pantothenoic acid or salts thereof. PCT Patent, WO 97/10340. 36. Norgard S, Aasen AJ, and Liaaen-Jensen S. (1970) Bacterial carotenoids. XXXII. C50-Carotenoids. 6. Carotenoids from Corynebacterium poinsettiae including four new C50-diols, Acta Chem. Scand. 24:2183. 37. Ogata K, Izumi K, Aoike K, and Tani Y. (1973) Further characterization of ureido ring synthetase from Pseudomonas graveolens. Agric. Biol. Chem. 37:1093–1099. 38. Okumura S, Tsugawa R, Tsunoda T, and Morisaki S. (1962) Studies on L-glutamic acid fermentation. Part II. Activities of various pelargonic acid compounds to promote fermentation. J. Agric. Chem. Soc. 36:204–211. 39. Ostrovsky D, Diomina G, Lysak E, Matveeva E, Ogre O, and Trutko S. (1998) Effect of oxidation stress on the biosynthesis of 2-C-methyl-D-erythritol-2,4-cyclopyrophosphate and isoprenoids by several bacterial strains. Arch. Microbiol. 171:69–72. 40. Perkins JB, Bower S, Howitt CL, Yocum RR, and Pero J. (1996) Identification and characterization of transcripts from the biotin biosynthetic operon of Bacillus subtilis. J. Bacteriol. 178:6361–6365. 41. Peters-Wendisch PG, Kreutzer C, Kalinowski J, Patek M, Sahm H, and Eikmanns BJ. (1998) Pyruvate carboxylase from Corynebacterium glutamicum: characterization, expression and inactivation of the pyc gene. Microbiology 144:915–927. 42. Ploux O, Soularue P, Marquet A, Gloeckler R, and Lemoine Y. (1992) Investigation of the first step of biotin biosynthesis in Bacillus sphaericus. Purification and characterization of the pimeloyl-CoA synthase, and uptake of pimelate. Biochem. J. 287:685–690. 43. Primerano DA and Burns RO. (1983) Role of acetohydroxy acid isomeroreductase in biosynthesis of pantothenic acid in Salmonella typhimurium. J. Bacteriol. 153:259–269. 44. Radmacher E, Vaitsikova A, Burger U, Krumbach K, Sahm H, and Eggeling L. (2002) Linking central metabolism with increased pathway flux: L-valine accumulation by Corynebacterium glutamicum. Appl. Environ. Microbiol. 68:2246–2250. 45. Ramjee MK, Genschel U, Abell C, and Smith AG. (1997) Escherichia coli L-aspartatealpha-decarboxylase: preprotein processing and observation of reaction intermediates by electrospray mass spectrometry. Biochem. J. 323:661–669. 46. Sahm H and Eggeling L. (1999) D-Pantothenate synthesis in Corynebacterium glutamicum and use of panBC and genes encoding L-valine synthesis for D-pantothenate overproduction. Appl. Environ. Microbiol. 65:1973–1979. 47. Sandmann G. (2001) Carotenoid biosynthesis and biotechnological application. Arch. Biochem. Biophys. 385:4–12. 48. Sanyal I, Gibson KJ, and Flint DH. (1996) Escherichia coli biotin synthase: an investigation into the factors required for its activity and its sulfur donor. Arch. Biochem. Biophys. 326:48–56. 49. Schneider G and Lindqvist Y. (2001) Structural enzymology of biotin biosynthesis. FEBS Lett. 495:7–11.

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50. Schuhmann G, Nürnberger H, Sandmann G, and Krügel H. (1996) Activation and analysis of cryptic crt genes for carotenoid biosynthesis from Streptomyces griseus. Mol. Gen. Genet. 252:658–666. 51. Shimizu S, Esumi A, Komaki R, and Yamada H. (1984) Production of coenzyme A by a mutant of Brevibacterium ammoniagenes resistant to oxypantetheine. Appl. Environ. Microbiol. 48:1118–1122. 52. Shimizu S and Yamada H. (1992) Enzymatic synthesis of chiral intermediates for D-pantothenate synthesis. In Heinemann R and Wolnak B (Eds.), Opportunities with Industrial Enzymes. Bernard and Associates, Chicago, pp. 227–241. 53. Tauch A, Götker S, Pühler A, Kalinowski J, and Thierbach G. (2002) The alanine racemase gen alr is an alternative to antibiotic resistance genes in cloning systems for industrial Corynebacterium glutamicum strains. J. Biotechnol. 99:79–91. 54. Terasawa M, Inui M, Goto M, Kurusu Y, and Yukawa H. (1991) Depression of byproduct formation during L-isoleucine production by a living-cell reaction process. Appl. Microbiol. Biotechnol. 35:348–351. 55. Terasawa M, Inui M, Goto M, Shikata K, Imanari M, and Yukawa H. (1990) Living cell reaction process for L-isoleucine and L-valine production. J. Ind. Microbiol. 5:289–294. 56. Viveiros M, Krubasik P, Sandmann G, and Houssaini-Iraqui M. (2000) Structural and functional analysis of the gene cluster encoding carotenoid biosynthesis in Mycobacterium aurum A+. FEMS Microbiol. Lett. 187:95–101. 57. Weeks OB and Garner RJ. (1967) Biosynthesis of carotenoids in Flavobacterium dehydrogenans Arnaudi. Arch. Biochem. Biophys. 121:35–49. 58. Yamagata H, Terasawa M, and Yukawa H. (1994) A novel industrial process for L-aspartic acid production using an ultrafiltration membrane. Catalysis Today 22:621–627.

18

Osmoregulation S. Morbach and R. Krämer

CONTENTS 18.1 The Impact of Osmotic Stress on Bacterial Physiology .............................417 18.2 Response of C. glutamicum to Hypoosmotic Stress ...................................418 18.3 Response of C. glutamicum to Hyperosmotic Stress..................................420 18.3.1 Potassium and Glutamate Response ................................................420 18.3.2 Accumulation of Compatible Solutes by Biosynthesis...................421 18.3.3 Accumulation of Compatible Solutes under N-Limitation .............421 18.3.4 Accumulation of Compatible Solutes from the Environment.........423 18.4 Biosynthesis of Compatible Solutes ............................................................424 18.4.1 Regulation of Proline Biosynthesis .................................................424 18.4.2 Regulation of Trehalose Biosynthesis .............................................425 18.5 Uptake of Compatible Solutes.....................................................................426 18.5.1 The Betaine Uptake System BetP ...................................................426 18.5.2 The Proline/Ectoine Uptake System ProP.......................................430 18.5.3 The Ectoine/Betaine/Proline Uptake System EctP..........................430 18.5.4 The Betaine/Ectoine Permease LcoP...............................................430 18.6 Relevance of Osmostress for Fermentation Processes................................431 References..............................................................................................................432

18.1 THE IMPACT OF OSMOTIC STRESS ON BACTERIAL PHYSIOLOGY The cytoplasmic membrane is freely permeable to water but forms an effective barrier for solutes of the environment or the cytoplasm. In general, the total concentration of osmotically active solutes within a cell is higher than in the environment, causing water to flow down its chemical potential into the cell. As a result, a hydrostatic pressure, the so-called turgor, is exerted by the cytoplasmic membrane toward the cell wall. Consequently, turgor balances the difference in osmotic pressure between the cell interior and its surroundings. Turgor is maintained throughout the growth cycle as the cell elongates, and it is considered necessary for enlargement of the cell envelope and thus for growth and division [27]. Although turgor is very difficult to quantify in bacteria [60], values of 3 to 5 bar for Gram-negative bacteria

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and approximately 20 bar for Gram-positive bacteria have been estimated [7,19,56]. The much higher value in Gram-positive bacteria is thought to reflect the large cytoplasmic solvent pool needed for expansion of the multilayer peptidoglycan. Because the environment of a bacterial cell is frequently subjected to fluctuations in osmolality, bacteria were forced to develop efficient adaptation mechanisms to cope with both, a decrease or an increase in the external osmolality, or, in other words, with a hypoosmotic shift (osmotic downshift) or a hyperosmotic shift (osmotic upshift). The two main adaptation strategies found in Corynebacterium glutamicum are the release of solutes out of the cytoplasm after an osmotic downshift and the accumulation of so-called compatible solutes after an osmotic upshift [5,31]. Betaine, proline, glutamine, ectoine, and trehalose were found to be effective in C. glutamicum (Figure 18.1) [13–16,22,40,49,59]. Compatible solutes combine two fundamental properties. In the cellular context, they cause rehydration of the cytoplasm by elevating the internal osmolality because they can be accumulated up to molar concentrations without disturbing cellular functions [7]. At the molecular level, compatible solutes are thought to stabilize and protect enzymes because they are excluded from the protein surface, thus leading to a preferential hydration of the protein (Figure 18.1) [3,61].

18.2 RESPONSE OF C. GLUTAMICUM TO HYPOOSMOTIC STRESS An osmotic downshift leads immediately to an excessive water influx and consequently to a dramatically increased turgor pressure. To avoid the risk of cell lysis, C. glutamicum, as with other bacteria, activates within milliseconds emergency release valves, the mechanosensitive (MS) channels (Figure 18.2). As a result, cytoplasmic solutes are effectively released into the environment and the driving force for water entry is reduced. C. glutamicum releases solutes like betaine or proline with a very high velocity of 6000 μmol⋅min–1⋅(mg dry weight)–1 [44]. Betaine and proline are preferred to other amino acids of similar size, and furthermore ATP is not excreted after an osmotic downshift [44]. Therefore, the corynebacterial MS channels seem to confer higher substrate specificity than the stretch-activated channels of E. coli, which are regarded as unspecific pores leading even to the excretion of small proteins like thioredoxin [1]. In C. glutamicum, two channel activities have been detected by means of patch clamp analyses [43]. The conductance obtained varied between 600 to 700 pS and 1,200 to 1,400 pS, respectively. These values are similar to those of MscS and MscL (MS channel of small and large conductance) of E. coli [4]. In the genome of C. glutamicum, two open-reading frames were identified [33] whose derived amino acid sequences show similarities with the MscL and the MscS (YggB) family of MS channels [28,51]. The MscL homolog (135 amino acid residues) shares 57% identity with its mycobacterial counterpart and 38% with MscL of E. coli. In contrast to the highly conserved MscL family, the sequences and lengths of MscS homologs are less conserved. The corynebacterial MscS (533 amino acid residues) has a higher identity to the corresponding homologs

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A Compatible solutes COOHO

HN -OOC

N+ H2

Proline

HOCH2 O

HO

N+ N+ H

COO-

Ectoine

HO

Betaine

OH

OH

O HO

O

H2 COH

Trehalose

B Action of compatiple solutes

Water NaCl Compatible solute

Low osmolality

Osmotic stress

Preferential exclusion

FIGURE 18.1 Compatible solutes and their proposed mode of action through preferential exclusion from protein surfaces [3,61]. (A) Compatible solutes active in C. glutamicum. (B) A model of the action of compatible solutes on proteins. At low osmolality, proteins are completely hydrated (left), whereas hyperosmotic stress, or more precisely, a decrease in water activity (middle), leads to a decrease in the hydration state of proteins and thus causes protein denaturation as indicated by the change in the shape of the protein. If compatible solutes are present (right), they are excluded from the immediate hydration shell of the protein, resulting in a preferential hydration of the protein surface. The disruption of the ordered water structure around proteins by local unfolding is thus energetically not favored, and the native protein conformation is stabilized.

FIGURE 18.2 Activation of mechanosensitive channels after an osmotic downshift. The black arrows indicate the outwardly directed turgor, which increases dramatically after an osmotic downshift owing to the instant influx of water into the cell (gray arrows). To prevent cell lysis, MS channels like MscL and MscS, which are localized in the cytoplasmic membrane, are immediately activated, which leads to a release of solutes, a decrease in the internal osmolality, and concomitantly to a reduction of the driving force for water entry. Cytoplasmic membrane (solid black line); cell wall (thick dashed gray line).

of different mycolic acids containing actinomycetes (up to 36% identity) than to MscS of E. coli (289 amino acid residues), where the identity of 29% is restricted to the three transmembrane segments present in both proteins [33]. Analyses of a

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double knock-out strain of C. glutamicum revealed that, besides an MscL and an MscS channel, at least one additional type of channel is present. The latter seems to have transport velocity and capacity similar to those of MscL and MscS because when a severe osmotic downshift was applied almost no differences between the mutant and the wild-type were detected in biochemical and cell survival assays [33]. Interestingly, a severe downshock was lethal in the corresponding E. coli double deletion mutant [28], indicating that C. glutamicum probably has a type of mechanosensitive channel with high permeability that is as yet unidentified. In terms of physiology, the redundancy of the systems guarantees a high degree of safety against hypoosmotic conditions for C. glutamicum.

18.3 RESPONSE OF C. GLUTAMICUM TO HYPEROSMOTIC STRESS If bacteria are faced to a sudden osmotic upshift, instant water efflux occurs and the cells dehydrate. As a consequence, growth is slowed down or even stopped. Bacteria respond to hyperosmotic stress in three overlapping phases [60] as illustrated for C. glutamicum in Figure 18.3A. In the first phase (within seconds and up to a minute), an instant but passive dehydration of the cytoplasm occurs. In the second phase, the cytoplasm rehydrates by adjustment of the cytoplasmic solvent composition owing to an accumulation of ions or compatible solutes (up to an hour). This phase overlaps with the third phase, the remodeling of the cell, which is characterized by changes of gene expression profiles and by exchange of ionic osmolytes against compatible solutes (up to one or more hours). As a result, growth is resumed.

18.3.1 POTASSIUM

AND

GLUTAMATE RESPONSE

Many bacteria respond to a hyperosmotic upshock instantaneously by a fast but transient uptake of potassium ions [12,56–57]. In C. glutamicum, the situation concerning this immediate response is not as clear. Guillouet and Engasser [15] reported that not K+ but Na+ accumulates eightfold, up to 40 mg Na+ (g dry weight)–1 [corresponding to an increase from approx. 200 to 1800 μmol (g dry weight)–1], within 30 min after shifting the osmolality from 0.4 to 1.5 Osm. In contrast, it was recently shown [59] that the internal K+ concentration increased within 5 min from 450 to 800 μmol (g dry weight)–1 after the osmolality was shifted from 0.9 to 2.4 Osm. The search for known potassium uptake systems in the genome of C. glutamicum revealed that only a homolog of the E. coli low-affinity and lowcapacity Kup system is present (NCBI annotation NCgl0682), whereas the highly efficient transporters KdpFABC and Trk of E. coli [46] or the Ktr system of Vibrio alginolyticus and B. subtilis [18,32] are missing. Even if the uptake of potassium is not part of the osmotic response, a K+ import system is essential for the formation of turgor pressure. Therefore, C. glutamicum should possess a high-efficient K+ uptake system. Unfortunately, no experimental data concerning the kinetics of potassium uptake or the systems involved are available. Future work must reveal how

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potassium uptake is mediated in C. glutamicum and whether hyperosmotic upshifts induce Na+ or K+ accumulation. In many bacteria, glutamate synthesis is increased concomitantly with K+ uptake to counterbalance the positive charges of the accumulated potassium ions [12]. In C. glutamicum, elevated glutamate concentrations were observed in the first hour after an osmotic upshift [15,59]. Although the synthesis of glutamate occurs directly after K+ accumulation, it seems questionable whether glutamate plays an important role in the osmostress response of C. glutamicum because the observed increase from 240 to 350 μmol (g dry weight)–1 was not very pronounced [59] (Figure 18.4).

18.3.2 ACCUMULATION OF COMPATIBLE SOLUTES BY BIOSYNTHESIS For a non-halophilic bacterium like C. glutamicum, the accumulation of K+, Na+, or glutamate after an osmotic upshock inevitably leads to problems owing to high internal ion (charge) concentrations, which may interfere with cell metabolism. Therefore, in the second phase of the osmotic response, ions are exchanged against compatible solutes either by uptake or synthesis [60] (Figure 18.3B). If compatible solutes are not present in the environment, C. glutamicum is able to synthesize glutamine, proline, and trehalose after an osmotic upshift (Figures 18.4 and 18.5) [14–16,22,40,49,59]. The most pronounced response is the accumulation of proline. Depending on the initial osmolality (0.4 or 0.9 Osm), an increase from 10 or 100 μmol up to a level of 900 μmol (g dry weight)–1 at an external osmolality of 2.4 was detected [58].

18.3.3 ACCUMULATION OF COMPATIBLE SOLUTES UNDER N-LIMITATION Several independent investigations suggested proline and not trehalose to be the major de novo-synthesized compatible solute of C. glutamicum [15–16,22,40,49]. Trehalose differs from all other osmoprotectants used by C. glutamicum by the absence of nitrogen in its chemical structure. Accumulation and physiological significance of those compatible solutes might thus depend on the availability of nitrogen. Indeed, it was shown that under nitrogen starvation the synthesis of compatible solutes was shifted at the expense of proline and glutamine toward trehalose [58,59]. The trehalose concentration was about sixfold increased, up to 600 μmol (g dry weight)–1. Therefore, trehalose becomes the predominant compatible solute of C. glutamicum under these conditions (Figures 18.3A and 18.5). Apart from the availability of nitrogen, the kind of carbon source influences the osmolyte pool. Under osmostress conditions, maltose-grown cells possess a three-times-higher trehalose content, of approx. 300 μmol (g dry weight)–1 as sucrose-grown cells, with a concomitant decrease in the cytoplasmic proline concentration [59]. Thus, osmoadaptation by compatible solute synthesis does not follow a rigid scheme but is flexibly adjusted to a variety of environmental parameters sensed by the cell in addition to osmolality, which are transduced to the osmoregulation network.

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B

Uptake of solutes: +

K

BetP

EctP

Efflux of solutes: LcoP

ProP

?

out

CM in

MscL

MscS

?

Biosynthesis of compatible solutes: α-ketoglutarate + NH4

Gdh

ProB

glutamate

glutamate

glutamine

maltodextrins

GS

glutamate

ProA

ProC

proline

TreY

TreZ

trehalose

FIGURE 18.3 Responses of C. glutamicum to osmotic stress. (A) Chronology of reactions of C. glutamicum after a hyperosmotic shift. First phase: a sudden osmotic upshift causes an immediate water efflux (indicated by gray arrows) and concomitantly a reduction or loss of a (positive) turgor pressure (indicated by black arrows), cell shrinkage, and dehydration of the cytoplasm. Second and third phases: to reverse the water fluxes, the cell increases the internal osmolality by accumulation of compatible solutes by different means depending on the environmental conditions. If compatible solutes are not present in the environment they are synthesized de novo. Under nitrogen starvation, only trehalose (I) is accumulated whereas under nitrogen surplus (II) the initially synthesized glutamate, glutamine, and trehalose are

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FIGURE 18.4 Typical changes in the internal concentration of compatible solutes in C. glutamicum ATCC 13032 induced by a severe osmotic upshift from 0.9 to 2.4 Osm by the addition of 750 mM NaCl under N-surplus conditions. Proline (gray dashed line); glutamate (black solid line); glutamine (black dotted line), and trehalose (gray solid line) [59].

18.3.4 ACCUMULATION OF COMPATIBLE SOLUTES FROM THE ENVIRONMENT Owing to lower energy costs, C. glutamicum prefers the uptake of external compatible solutes to their de novo synthesis [31]. In media of high osmolality, C. glutamicum shows improved growth in the presence of betaine (and its precursor dimethyl glycine), ectoine, and proline [6,13,23]. Betaine, which cannot be synthesized or degraded in C. glutamicum [13,23], is the most efficient compatible solute in terms of growth enhancement [13]. The accumulation of betaine in the cytoplasm (Figure 18.3A) changed both the concentration and the pattern of the protecting solutes synthesized [23,40]. The most pronounced effect is observed for proline, the concentration of which is reduced by at least about 60%, indicating that the biosynthesis of osmoprotectants is a well-coordinated process in C. glutamicum to prevent the waste of energy. FIGURE 18.3 (continued) replaced against proline, which is the predominant solute in the third phase of the adaptation process (II). If present in the environment, betaine is imported via BetP, EctP, and LcoP. As a consequence, in the third phase of the stress response the concentration of proline is reduced to less than 50% of the level observed in the absence of external betaine. At the end of these processes (I– III), the cell is hydrated again and therefore adapted to the elevated osmolality. (B) Systems involved in the osmostress response of C. glutamicum. The adaptation to an osmotic downshift is mediated by the mechanosensitive channels MscL and MscS, whereas in response to an osmotic upshift either uptake carriers are activated to import compatible solutes or the biosynthesis of osmolytes is increased. CM, cytoplasmic membrane, Gdh, glutamate dehydrogenase, GS, glutamine synthetase

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FIGURE 18.5 Consequences of different nitrogen availabilities on the intracellular trehalose and proline content after a severe osmotic up-shock, from 0.9 to 2.4 Osm by KCl addition. The left panel shows the accumulation of intracellular trehalose and the right panel the internal proline content. The gray dashed lines are for the solutes at nitrogen surplus, the black dashed lines for nitrogen limitation, and the black solid lines for nitrogen starvation conditions [59].

18.4 BIOSYNTHESIS OF COMPATIBLE SOLUTES 18.4.1 REGULATION

OF

PROLINE BIOSYNTHESIS

Under surplus nitrogen conditions, proline is the major endogenously synthesized osmoprotectant in C. glutamicum. The corynebacterial genes encoding the enzymes of the proline biosynthesis pathway were identified [2,47] and named proB (encoding the γ-glutamyl kinase), proA (encoding γ-glutamyl phosphate reductase), and proC (encoding the Δ1-pyrroline-5-carboxylate reductase, or P5C) (Figure 18.3B). The genes proB and proA are separated by a gene of unkown function, unk, whereas proC is located elsewhere in the chromosome. The C. glutamicum genome sequence revealed that no additional open-reading frames with similarities to proB, proA, or proC are present. Since C. glutamicum possesses only one set of enzymes for anabolism and osmostress regulation, a coordinated regulation of proline biosynthesis either at the level of enzyme activity or gene expression is necessary. It must be ensured that in the absence of osmotic stress a comparatively low proline pool of approx. 10 μmol (g dry weight)–1 is maintained, which must be elevated to 900 μmol (g dry weight)–1 under severe osmotic stress [29,58]. No data on the activity regulation of the proline biosynthesis enzymes are available. However, concerning expression control of proline biosynthesis, Kawahara et al. [22] observed that the enzymatic activity of ProC, but not of ProB, was elevated three times, from 67.3 to 220 nmol⋅min–1⋅(mg of protein)–1 in cells grown in the presence of 1 M sorbitol. Meanwhile, expression analyses showed that proB, proA, and proC are induced in an osmostress-dependent manner [29].

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glucose-6-P UDP-glucose

( α-(1,4) maltodextrins)n

maltose

OtsA

TreY ( α−(1,1) trehalosylmaltodextrins)n

TreS

trehalose-6-P

OtsB

TreZ

HOCH2 HO

O

HO

OH

OH

( α-(1,4) maltodextrins)n-2

O HO

HO

O

H2COH

trehalose FIGURE 18.6 Trehalose metabolism in C. glutamicum. Three different pathways involved in trehalose biosynthesis (OtsAB and TreYZ) or degradation (TreS) have been identified [53,59]. For details see text.

18.4.2 REGULATION

OF

TREHALOSE BIOSYNTHESIS

Genome searches revealed that C. glutamicum possesses five genes encoding the enzymes of three different trehalose biosynthesis pathways. The OtsAB pathway, the most common route known to be involved in the stress response of E. coli and yeast, proceeds from UDP-glucose and glucose-6-phosphate to form trehalose-6phosphate, which is subsequently dephosphorylated to yield free trehalose (Figure 18.6). The reactions are catalyzed by trehalose-6-phosphate synthase and trehalose-6-phosphate phosphatase, respectively. Less-prominent routes for trehalose synthesis are the TreYZ [11,26,30] and the TreS pathways [11,52] (Figure 18.6). The substrates of the TreYZ route are oligomaltodextrins or glycogen [11,30,53]. In the first reaction step, TreY (maltooligosyl trehalose synthase) transglycosylates a terminal maltosyl residue into a trehalosyl residue before trehalose is liberated through the activity of TreZ (maltooligosyl trehalose hydrolase). Finally, it was described that TreS (trehalose synthase) transforms maltose in a single transglycosylation reaction into trehalose [11,52]. Since many bacteria harbor only a single trehalose biosynthesis pathway, the question arose why C. glutamicum possesses three routes. As a matter of fact, trehalose is not absolutely essential for C. glutamicum because it was possible to construct trehalose-deficient mutants by deleting the trehalose biosynthesis genes. However, these strains were severely impaired in growth [53,59]. Recently, the functions of the trehalose pathways were partially assigned [59]. As mentioned

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above, trehalose is important for the osmostress response under certain environmental conditions, such as N starvation or if maltose as a carbon source is present. In C. glutamicum, trehalose synthesis in response to osmotic stress does not depend on the OtsAB pathway but is almost entirely TreYZ-mediated, which is in contrast to many other bacteria [59]. Employing TreYZ for stress adaptation seems reasonable because the typical bacterial storage carbohydrate, glycogen, represents a rapidly accessible substrate that does not require uptake mechanisms, which might be hampered after a hyperosmotic shock [60]. Concerning the physiological function of TreS, several findings led to the conclusion that TreS acts in vivo not as a trehalose synthase, which is the described function of the enzyme in the literature, but rather as a trehalose-degrading enzyme [59]. In contrast to the TreYZ and the TreS pathway, it has not so far been possible to assign a particular metabolic function to the OtsAB pathway in C. glutamicum. It might be related to the synthesis of mycolic acids, which are major constituents of cell wall lipids in corynebacteria, mycobacteria, and nocardiae [39]. The reason is that trehalosyl monomycolate (mycolates esterified with a trehalosyl residue) is thought to be the precursor of all other types of mycolic acids, namely, cell wall–bound corynemycolate and trehalosyl dimycolate and free corynemycolate [48]. Since the OtsAB pathway is the sole source of trehalose-6-phosphate, one might suggest that the OtsA activity is essential for mycolate biosynthesis, but recent investigations showed that OtsA-deficiency alone has no effect on mycolate biosynthesis. Only strains deficient in both, the OtsAB and the TreYZ pathways, are unable to synthesize trehalose and are concomitantly devoid of mycolic acids [53,58–59]. This indicates that both pathways provide trehalose for mycolic acid formation in C. glutamicum.

18.5 UPTAKE OF COMPATIBLE SOLUTES The uptake of compatible solutes is mediated by five secondary transport systems, whereas in other bacteria, for example, E. coli [60] and B. subtilis [25], primary transport systems also participate in the stress response. One of the five carriers of C. glutamicum, PutP, is not osmoregulated in its activity and is responsible for proline uptake for anabolic purposes [35]. The other transporters, BetP, ProP, EctP, and LcoP, are regulated in their activity in dependence of the external osmolality and are therefore relevant for the osmostress response [13,34,37,50]. They have overlapping substrate spectra and different affinities with their substrates, allowing C. glutamicum to use a broad variety of compatible solutes found at different concentrations in the environment (Table 18.1).

18.5.1 THE BETAINE UPTAKE SYSTEM BETP BetP (595 amino acids), a typical secondary transport system of the betaine/carnitine/choline transporter (BCCT) family (Table 18.1) [13,34], is predicted to possess 12 transmembrane segments as well as two highly charged domains. These are located at its N- and C-terminal ends and consist of either 62 or 55 amino acids, respectively (Figure 18.7) [42]. The biochemical characterization showed that BetP

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TABLE 18.1 Substrate Specificity, Kinetic Parameters, and Activity Regulation of Uptake Systems for Compatible Solutes in C. glutamicum Transporter (Transporter Family) BetP (BCCT1) EctP (BCCT)

LcoP (BCCT) ProP (MHS2) PutP (SSS4) 1

Substrates

Co-substrate

Betaine Ectoine Betaine Proline Betaine Ectoine Proline Ectoine Proline

Na+ Na+

Na+ H+ Na+

Km μM) (μ 8.9 63 333 1,200 154 539 48 132 7.6

± ± ± ± ± ±

0.4 4.9 45 180 15 41

Vmax (nmol/[min mg dry weight])

Activity Optimum (Osm)

110 27 34 34 n.d. n.d. 713 1293 20

1.3 1.3

1.6 1.3 Not osmoregulated

Betaine/carnitine/choline transporter family (Transport Protein Database of http://tcdb.ucsd.edu/tcdb/tcclass.php). 2 Metabolite H+ symporter family. 3 Values were derived after induction of plasmid-encoded proP in E. coli MKH13/pHP5. 4 Solute: sodium symporter family. n.d. = not determined.

M.

Saier:

is a Na+-coupled, highly specific betaine uptake system (Km of 8.9 μM) [6,34] that is able to build up extremely high betaine gradients of 4 × 106 (inside/outside ratio) [13]. The carrier is osmoregulated at the level of activity. In vivo, almost no transport was detected at low external osmolalities between 0.2 and 0.4 Osm. Higher osmolalities resulted in a stress-dependent activation of the carrier, with a maximal activity at approx. 1.3 Osm [13,34]. Interestingly, after heterologous expression of betP in E. coli, the carrier was still osmoregulated but the activity optimum was found at a lower osmolality, approx. 0.6 Osm. This was the first indication that BetP itself possesses three functions [34], the catalytic function of betaine transport, the ability to detect hyperosmotic stress (osmosensing), and the capability to regulate the transport activity in dependence of the extent of osmotic stress (osmoregulation). To prove this hypothesis, it was necessary to reduce the complexity of the cellular system by reconstitution of purified BetP in liposomes made from E. coli phospholipids. In this in vitro system, BetP showed kinetic properties similar to those of intact cells and retained its osmodependent activity regulation [41]. Therefore, BetP meets the demands of an osmosensor and osmoregulator. The underlying mechanisms — how the osmotic stress is sensed — are not well understood and are a matter of current research for several osmoregulated carriers [31,38]. By using the proteoliposomal system, it was possible to identify the physicochemical stimulus that is sensed by BetP as a measure of osmotic stress. It could be excluded that turgor pressure, liposomal shrinkage (membrane strain), external conditions, or

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A IV

V

VI

VII

VIII

IX

X

XI

XII

Cytoplasmic membrane

III

II

I

Cytoplasm

-

Modulator

Sensor

+ + + K+ sensitive region

N

C

FIGURE 18.7 BetP and its osmostress-dependent activation. (A) Model of the BetP topology and domains involved in osmosensing and regulation. A stretch of 13 residues within the last 25 amino acids of the positively charged C-terminal extension of BetP is directly involved in sensing K+ [45], whereas the negatively charged N-terminal domain seems to modulate only the activity. (B) In proteoliposomes, the sole trigger for carrier activation is the concentration increase in luminal K+ (or of related ions like Rb+), whereas other cations showed no effect [42]. (C) Influence of the lipid composition of proteoliposomes on the activation threshold of BetP. The ratio between negatively charged phosphatidyl glycerol (PG), the main phospholipid of C. glutamicum, and E. coli phospholipid, comprising mainly neutral phosphatidyl ethanolamine (PE), defines the activation threshold of BetP. With increasing PG contents, higher osmolalities are necessary to activate BetP in proteoliposomes [41].

changes of the internal ionic strength or osmolality are triggering the activity of BetP in proteoliposomes [42,45]. Exclusively, the increase of the cytoplasmic K+ concentration was found to be responsible for BetP activation in proteoliposomes (Figure 18.7B) [42,45]. Meanwhile, investigations were carried out to clarify which part of BetP is responsible for the osmodependent activity regulation. Variants of BetP were generated carrying either truncations at the N- or the C-terminal domain [36]. The in vivo

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429

FIGURE 18.7 (continued).

analysis revealed that mutants with deletions in the N-terminal extension were still osmoregulated, but the activity optimum was shifted to a higher osmolality, 2.6 Osm, indicating that this domain has only a modulating influence on BetP activity (Figure 18.7A). The effects of C-terminal deletions were more pronounced. Stepwise truncations of 12, 25, or 45 amino acids resulted either in a partly deregulated (BetPΔ12) or a completely deregulated activity (BetPΔ25 or BetPΔ45), meaning that the carrier was fully active, independent of the external osmolality [36,45]. This indicates that the C-terminal domain is involved in the K+ sensing process. The experimental verification of this hypothesis was carried out in the proteoliposomal system. C-Terminally truncated BetP proteins were purified and reconstituted in liposomes. The analysis of the K+ dependence of BetP activation revealed that K+ sensitivity is still detectable if only the terminal 12 amino acid residues were deleted but is completely lost if the truncation comprises the last 25 residues [45]. Therefore, al least a part of the K+ sensor seems to be located within this part of BetP. The sensitivity of BetP to hyperosmotic stress is influenced by the kind of phospholipids used for preparing the proteoliposomes [41]. The higher the share of negatively charged phosphatidyl glycerol in the proteoliposomes, the higher the external osmolalities needed to activate BetP (Figure 18.7C). The opposite was found for the fraction of E. coli lipids in the liposomes. This observation may explain why BetP showed in vivo a different sensitivity toward osmotic stress in C. glutamicum and E. coli cells: phosphatidyl glycerol is the major phospholipid of the C. glutamicum plasma membrane, whereas the E. coli plasma membrane consists mainly of phosphatidyl ethanolamine [17]. In recent years, the triggers for activation of five different osmosensitive systems were studied, namely of the secondary systems BetP [42] and ProP [10], of the ABC transporter OpuA [54] as well as of two sensor kinases of two-component systems in E. coli, namely KdpD and EnvZ [20–21]. Interestingly, the stimulus indicating hyperosmotic stress was either the internal ionic strength (ProP, KdpD, OpuA) or the internal potassium concentration (EnvZ, BetP), although these systems are not functionally or structurally related at all. It must thus be concluded that osmoregulated systems — irrespective of a completely different evolutionary origin and irrespective of the fact that a large number of stimuli are in principle available (see above) — seem to use closely similar stimuli for signal perception.

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18.5.2 THE PROLINE/ECTOINE UPTAKE SYSTEM PROP The C. glutamicum proP gene was isolated by complementation of an E. coli strain deficient for the uptake of betaine and proline [37]. The ProP transporter (504 amino acids) belongs to a subgroup of the major facilitator superfamily (MFS), the metabolite H+ symporter family, and shows a high identity of 38% to ProP of E. coli [8]. In contrast to the E. coli protein, ProP of C. glutamicum is predicted to possess only 11 and not 12 transmembrane helices. The topology prediction furthermore indicates that ProP possesses an N- and a C-terminal extension of 69 and 57 amino acids, respectively [37]. The biochemical characterization showed that ProP prefers proline as substrate (Km of 48 μM) to ectoine (Km 132 of μM). Furthermore, the carrier is regulated at the level of activity independent of the external osmolality with a regulation pattern very similar to BetP [37]. No further investigations were carried out to prove whether the C-terminal extension of ProP is involved in the osmoregulation of the carrier. Interestingly, the coiled-coil motif located in the C-terminal extension of the E. coli homolog, which seems to be necessary for the osmostressdependent activation [9], is not found in the C. glutamicum counterpart.

18.5.3 THE ECTOINE/BETAINE/PROLINE UPTAKE SYSTEM ECTP EctP was isolated by complementation of a C. glutamicum strain, deficient for the uptake of compatible solutes [37]. Sequence comparisons showed that EctP belongs to the same transporter family as BetP, i.e., the BCCT family. In contrast to the highly specific BetP, EctP has a broader substrate specificity and accepts ectoine, betaine, and proline with different affinities (Table 18.1). The gene codes for a 615residue protein, which is predicted to contain 12 transmembrane segments and extensions at its N-and C-terminal ends, with a length of 23 and 108 residues, respectively. Interestingly, there are sequence similarities between EctP and BetP in the central part of the carriers only, where the highly conserved motif of the BCCT family is located (at the cytoplasmic side of helix 8 and the downstream loop). However, the hydrophilic extensions are not conserved with respect to primary sequence, length, or charge distribution. Since transport by EctP is also regulated at the level of activity with an optimum at an external osmolality of 1.3 Osm [37], the question was addressed whether these extensions are involved in the osmoregulation of EctP, too. For this approach, EctP variants were generated truncated either at the N- or C-terminal domain. Indeed, an EctP mutant missing the complete N-terminal domain became activated at higher osmolalities, whereas the C-terminally truncated mutant was active independent of the external osmolality [50]. It seems that the hydrophilic domains located at the N- and C-terminal parts of carriers of the BCCT family have similar functions in regulating the carrier activity.

18.5.4 THE BETAINE/ECTOINE PERMEASE LCOP Another osmoregulated transport system for compatible solutes, designated LcoP, was identified in the C. glutamicum genome as a result of sequence similarities shared with the BCCT family [50]. This Na+-dependent carrier accepts betaine and

Osmoregulation

431

ectoine as substrates with Km values of 155 μM and 539 μM, respectively but showed a low Vmax of 1.3 nmol⋅min–1⋅(mg dry weight)–1 (Table 18.1). In view of its low transport capacity, the physiological function of this system remains to be elucidated.

18.6 RELEVANCE OF OSMOSTRESS FOR FERMENTATION PROCESSES Only few investigations directly address the question how the amino acid production process is influenced by high osmolality or by the availability of compatible solutes, although hyperosmotic conditions routinely occur during fermentation processes owing to high substrate or product concentrations. Kawahara et al. [24] showed with an L-lysine production strain that the addition of betaine to the fermentation broth has beneficial effects on growth as well as on sugar consumption and the L-lysine production rate. Recently, the effects of betaine on L-lysine production of C. glutamicum strain MH20-22B during growth at different constant osmolalities of 1.0, 1.5, and 2.5 Osm were investigated. In this study, growth rates and cytoplasmic volumes decreased linearly with increasing osmotic stress. Interestingly, under all conditions tested the presence of 10 mM betaine led to an increase of the cell volume but did not influence the maximal growth rates [40]. Severe osmotic stress led to a betaine accumulation of up to 250 mM, with a concomitant reduction of the cytoplasmic proline content from 750 to 300 mM [40]. Betaine was found to be more efficient in counteracting osmotic stress because the overall concentration of compatible solutes in the cell was lower in the presence of betaine. This again illustrates why betaine uptake is preferred to the endogenous synthesis of compatible solutes. Although betaine has a positive effect on the cell’s vitality, its impact on L-lysine production is rather low, since the production of L-lysine is stimulated only under severe osmotic stress. At 2.5 Osm, the addition of betaine shifted L-lysine production in MH20-22B to earlier fermentation times and increased both product concentration and yield in these phases, but did not improve the final L-lysine yield. Because under severe osmotic stress total L-lysine yield was reduced by about 70% from 11 to 4 g l–1, it seems preferable to avoid high osmolalities during the fermentation process rather than to add compatible solutes [40]. Using a completely different setup, the negative effect of osmotic stress on L-lysine productivity was shown by Varela et al. [55], too. In this approach, the adaptation of an L-lysine production strain to a linear osmotic gradient from 0.2 to 1.8 Osm was analyzed in continuous culture. A decrease of the L-lysine yield was detected after a threshold level of 0.85 Osm was exceeded. Osmotic stress seems to change the energy demands of C. glutamicum. In a glucose-limited continuous culture of an L-lysine production strain, it was shown that the specific glucose uptake rate increased concomitantly with the osmolality whereas the biomass yield was reduced [16]. Recently, these observations were expanded by metabolic flux analysis done by Varela et al. [55]. They showed that the specific fluxes, via both the glucose PTS system and the TCA cycle, increased with increasing osmolalities. As a result, the ATP production rate was stimulated by about 50% [55]. This indicates that the share of energy for maintenance requirements (maintenance energy) is elevated under osmotic stress, which might therefore be the reason for the reduced L-lysine yield observed.

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34. Peter H, Burkovski A, and Krämer R. (1996) Isolation, characterization, and expression of the Corynebacterium glutamicum betP gene, encoding the transport system for compatible solute glycine betaine. J. Bacteriol. 178:5229–5234. 35. Peter H, Bader A, Burkovski A, Lambert C, and Krämer R. (1997) Isolation of the putP gene of Corynebacterium glutamicum and characterization of a low affinity uptake system for compatible solutes. Arch. Microbiol. 168:143–151. 36. Peter H, Burkovski A, and Krämer R. (1998) Osmo-sensing by N- and C-terminal extensions of the glycine betaine uptake system BetP of Corynebacterium glutamicum. J. Biol. Chem. 273:2567–2574. 37. Peter H, Weil B, Burkovski A, Krämer R, and Morbach S. (1998) Corynebacterium glutamicum is equipped with four secondary carriers for compatible solutes: identification, sequencing, and characterization of the proline/ectoine uptake system, ProP, and the ectoine/proline/glycine betaine carrier, EctP. J. Bacteriol. 180:6005–6012. 38. Poolman B, Blount P, Folgering JH, Friesen RH, Moe PC, and van der Heide T. (2002) How do membrane proteins sense water stress? Mol. Microbiol. 44:889–902. 39. Puech V, Bayan N, Salim K, Leblon G, and Daffé M. (2000) Characterization of the in vivo acceptors of the mycoloyl residues transferred by the corynebacterial PS1 and the related mycobacterial antigens 85. Mol. Microbiol. 35:1026–1041. 40. Rönsch H, Krämer R, and Morbach S. (2003) Impact of osmotic stress on volume regulation, cytoplasmic solute composition and lysine production in Corynebacterium glutamicum MH20-22B. J. Biotechnol. 104:87–97. 41. Rübenhagen R, Rönsch H, Jung H, Krämer R, and Morbach S. (2000) Osmosensor and osmoregulator properties of the betaine carrier BetP from Corynebacterium glutamicum in proteoliposomes. J. Biol. Chem. 275:735–741. 42. Rübenhagen R, Morbach S, and Krämer R. (2001) The osmoreactive betaine carrier BetP from Corynebacterium glutamicum is a sensor for cytoplasmic K+. EMBO J. 20:5412–5420. 43. Ruffert S, Berrier C, Krämer R, and Ghazi A. (1999) Identification of mechanosensitive channels in the cytoplasmic membrane of Corynebacterium glutamicum. J. Bacteriol. 181:1673–1676. 44. Ruffert S, Lambert C, Peter H, Wendisch VF, and Krämer R. (1997) Efflux of combatible solutes in Corynebacterium glutamicum mediated by osmoregulated channel activity. Eur. J. Biochem. 247:572–580. 45. Schiller D, Rübenhagen R, Krämer R, and Morbach S. (2004) The C-terminal domain of the betaine carrier BetP of Corynebacterium glutamicum is directly involved in sensing K+ as an osmorelevant stimulus. Biochemistry 43:5583–5591. 46. Schlösser A, Meldorf M, Stumpe S, Bakker EP, and Epstein W. (1995) TrkH and its homologue, TrkG, determine the specificity and kinetics of cation transport by the Trk system of Escherichia coli. J. Bacteriol. 177:1908–1910. 47. Serebrijski I, Wojcik F, Reyes O, and Leblon G. (1995) Multicopy suppression by asd gene and osmotic stress-dependent complementation by heterologous proA in proA mutants. J. Bacteriol. 177:7255–7260. 48. Shimakata T and Minatogawa Y. (2000) Essential role of trehalose in the synthesis and subsequent metabolism of corynomycolic acid in Corynebacterium matruchotii. Arch. Biochem. Biophys. 380:331–338. 49. Skjerdal OT, Sletta H, Flenstad SG, Josefsen KD, Levine DW, and Ellingsen TE. (1996) Changes of the intracellular composition in response to hyperosmotic stress of NaCl, sucrose or glutamic acid in Brevibacterium lactofermentum and Corynebacterium glutamicum. Appl. Microbiol. Biotechnol. 44:635–642.

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50. Steger R. (2002) Vergleichende Studien zur Aktivitätsregulation osmosensitiver Transporter aus Corynebacterium glutamicum. PhD thesis, Universität zu Köln. 51. Sukharev SI, Blount P, Martinac B, Blattner FR, and Kung C. (1994) A largeconductance mechanosensitive channel in E. coli encoded by mscL alone. Nature 368:265–268. 52. Tsusaki K, Nishimoto T, Nakada T, Kubota M, Chaen H, Sugimoto T, and Kurimoto M. (1996) Cloning and sequencing of trehalose synthase gene from Pimelobacter sp. R48. Biochim. Biophys. Acta 21:1–3. 53. Tzvetkov M, Klopprogge C, Zelder O, and Liebl W. (2003) Genetic dissection of trehalose biosynthesis in Corynebacterium glutamicum: inactivation of trehalose production leads to impaired growth and an altered cell wall. Microbiology 49:1659–1673. 54. van der Heide T, Stuart MC, and Poolman B. (2001) On the osmotic signal and osmosensing mechanism of an ABC transport system for glycine betaine. EMBO J. 20:7022–7032. 55. Varela C, Agosin E, Baez M, Klapa M, and Stephanopoulos G. (2003) Metabolic flux redistribution in Corynebacterium glutamicum in response to osmotic stress. Appl. Microbiol. Biotechnol. 60:547–555. 56. Whatmore AM and Reed RH. (1990) Determination of turgor pressure in Bacillus subtilis: a possible role for K+ in turgor regulation. J. Gen. Microbiol. 136:2521–2526. 57. Whatmore AM, Chudek JA, and Reed RH. (1990) The effects of osmotic upshock on the intracellular solute pools of Bacillus subtilis. J. Gen. Microbiol. 136:2527–2535. 58. Wolf A. (2002) Trehalosesysnthese in Corynebacterium glutamicum. Ph.D. Thesis, Universität zu Köln. 59. Wolf A, Krämer R, and Morbach S. (2003) Three pathways for trehalose metabolism in Corynebacterium glutamicum ATCC13032 and their significance in response to osmotic stress. Mol. Microbiol. 49:1119–1134. 60. Wood JM. (1999) Osmosensing by bacteria: signals and membrane-based sensors. Microbiol. Mol. Biol. Rev. 63:230–262. 61. Youxing Q, Bolen CL, and Bolen W. (1998) Osmolyte-driven contraction of a random coil protein. Proc. Natl. Acad. Sci. USA 95:9268–9273.

Part VI Synthesis and Production

19

L-Glutamate

Production

E. Kimura CONTENTS 19.1 Introduction ..................................................................................................439 19.2 Glutamate Technology .................................................................................440 19.2.1 Carbon Source..................................................................................440 19.2.2 Fermentation Process .......................................................................441 19.2.3 Crystallization ..................................................................................442 19.2.4 Waste Reduction...............................................................................443 19.3 Induction of L-Glutamate Overproduction...................................................444 19.3.1 From Leak Model to Metabolic Flux Change Model.....................445 19.4 Cellular Characteristics of Glutamate Overproduction ...............................445 19.4.1 Reactions Leading to 2-Oxoglutarate ..............................................445 19.4.2 Ammonia Incorporation...................................................................447 19.4.3 The Significance of ODHC Activity ...............................................448 19.4.4 Cloning and Analysis of the odhA Gene.........................................448 19.4.5 Cloning of dtsR Genes.....................................................................451 19.4.6 Characterization of dtsR Mutants ....................................................452 19.4.7 Regulation of dtsR1 Expression ......................................................453 19.4.8 Metabolic Flux Analysis of Glutamate Overproduction .................454 19.5 Next Generation Glutamate Producer: Corynebacterium efficiens .............454 19.5.1 Comparative Analysis of C. efficiens...............................................456 19.6 Future Prospects...........................................................................................457 Acknowledgments..................................................................................................457 References..............................................................................................................457

19.1 INTRODUCTION In 1908, Prof. K. Ikeda of Tokyo University discovered that the major ingredient of a seaweed known as kon-bu was monosodium glutamate (MSG). MSG has a unique flavor, which is called umami in Japanese [78]. One year later the company Ajinomoto put Col., Inc., MSG into the market. The sources for MSG at that time were wheat, soybean, and other plant protein material, from which MSG was extracted after hydrolysis by hydrochloric acid. However, in 1956 an L-glutamate–producing bacterium, Corynebacterium glutamicum, was isolated from nature, revolutionizing MSG production [50,104].

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All of the successive studies on C. glutamicum have started from this revolutionary finding. Soon a number of further isolates were found to be able to excrete L-glutamate; these were named Brevibacterium lactofermentum or B. flavum [109], for instance. However, all these bacteria were later determined to be Corynebacterium species [58] (see also Chapter 2), and they are referred to in this chapter as coryneform bacteria. The mechanism of L-glutamate [49] overproduction by these bacteria is exciting, and recent discoveries add very much to the understanding of the ability of these bacteria to overproduce L-glutamate. One particular feature of C. glutamicum is that L-glutamate excretion occurs only under special treatments, or growth conditions, such as growth under biotin limitation. In this review aspects of L-glutamate production and of the properties of C. glutamicum will be described, together with a putative model leading to production. The current major suppliers of MSG and L-glutamate, which are used as a flavor enhancer and also as a drug and as a precursor of drugs, cosmetics, and further pharmaceutical compounds, are Ajinomoto, Miwon, and Cheil-Jedang. Whereas the annual production of MSG in 1957 was less than 100,000 t, and in 1985 it was about 300,000 t, in 1996 900,000 t were produced. At the end of the last millennium about 1,000,000 t of MSG were made, with the volume still increasing. Glutamate is exclusively produced by the fermentative method using coryneform bacteria, with a majority of the production plants located in South Asia.

19.2 GLUTAMATE TECHNOLOGY 19.2.1 CARBON SOURCE The costs of substrates are among the variable costs in fermentations, and it is imperative to reduce these costs as much as possible. Consequently, a number of different substrates, e.g., sugars, acetate, n-paraffins, and methanol, have been studied for cost-effective utilization in L-glutamate production with C. glutamicum. However, use of sugars such as cane molasses, beet molasses, or hydrolysates from corn or cassava became standard, with the type of sugar depending on the geographical location of the production plant. Whereas starch hydrolysate from corn is the most important carbon source in North America, molasses is common in Europe and South America, and starch hydrolysate from cassava is preferentially used in South Asia [28]. At the beginning of L-glutamate fermentation, pure sugars were favorable as compared to molasses, because some types of molasses contain biotin concentrations inhibitive for L-glutamate production with C. glutamicum at that time. However, this is of no concern with the current sophisticated mutants and procedures developed in which any biotin present does not interfere with the excellent production properties of the strains. Therefore, molasses, which in addition to sugar contain specific nutrients, trace elements, and osmoprotectants whose presence is advantageous for the process, are cost-effectively used. However, depending on the type of molasses or hydrolysates further medium components may have to be adjusted, and specific strains may be used.

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Production

441

Fed-batch Culture (FC)

Continuous Culture (CC)

Cell Recycled Continuous Culture (CRCC)

Feed

Feed

Feed

Sugar, Nutrients

Sugar, Nutrients

Sugar, Nutrients

Pull out

Pull out Broth

Cells Broth

Glutamate

FIGURE 19.1 Different types of fermenter systems suitable for L-glutamate production.

19.2.2 FERMENTATION PROCESS The fermentative productivity of L-glutamate is represented simply by the following equation: Pglu = ρglu × X × V where Pglu = L-glutamate productivity by fermentation, L-glutamate (g)/time (h) ρglu = L-glutamate productivity by unit cells, L-glutamate (g)/cell (g)/time (h) X = cell density, cell (g)/volume (l) V = volume, l Obviously, fermenter volume and cell density affect the productivity of the fermentation process. In addition, the cellular specific productivity, ρglu, is decisive for the productivity of the process. Although ρglu depends largely on the capability of the mutant strain, it is also strongly affected by the process conditions, such as for instance cultivation temperature, pH, medium composition, and aeration. Basically, industrial amino acid fermentation is performed using batch or fedbatch culture. The fed-batch culture process has several advantages: the sugar concentration can be kept low to reduce by-product formation, including lactate or acetate formation; and at the same time a high oxygen demand, which might occur during exponential growth and which would exceed the capacity of the fermenter thereby also reducing product yield [28], is prevented. Other fermentation processes have been studied for the further improvement of the productivity (Figure 19.1). In the case of continuous culture, the glutamate yield obtained was 55% with a

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productivity of up to 8.3 g l–1 h–1 [111], which is twice the productivity of a typical batch culture. Since the cell density (X) is directly proportional to Pglu, cell retention is of course a consequent development [51]. In such a process, called cell-recycled continuous culture (CRCC) (Figure 19.1), basically culture supernatant without cells is removed from the fermentation tank. This is done, for instance, by separating cells by centrifugation and recycling them into the fermenter. Using this method, fermentation yield and productivity are maintained at a high level for a relatively long time. Another approach is the use of immobilized cells [38]. However, high cost of the carrier to immobilize the bacteria and refinement of the carrier exchange technique were difficulties encountered.

19.2.3 CRYSTALLIZATION Most of the L-glutamic acid in the fermentation broth can be crystallized by pH adjustment with acid to its isoelectric point (pH 3.2) because of the low solubility of L-glutamic acid (Figure 19.2). There are two polymorph crystal forms of L-glutamic acid at its isoelectric point [74,75], and the α form is the preferred one for harvesting

α-form

β-form L-Glutamate broth pH adjust (low) α-form crystallization α to β transition Dissolving (NaOH) Decoloration (Active carbon) Concentration L-glutamate-Na Dry

FIGURE 19.2 Two crystal forms of L-glutamate (A), and the downstream processing of monosodium glutamate production (B).

L-Glutamate

Production

443

because it has a granular shape and is therefore easier to separate from the fermentation broth than the plate-like form when using the separation machine (super decanter). Fortunately, the α form crystallizes faster than the form. Following the first separation step, solvent-mediated conversion of the metastable α form to the β form occurs, taking advantage of the low solubility of the form. By this recrystallization most of the impurities are easily removed [77]. In this manner, the different crystallization speeds, solubilities, and crystal sizes are advantageously used to obtain a high-quality product, which after dissolution in water, neutralization with NaOH, and further processing eventually yields monosodium L-glutamic acid monohydrate crystals (MSG) suitable for use as a flavor enhancer in food additives.

19.2.4 WASTE REDUCTION An attractive option to reduce the tremendous material fluxes and waste loads during glutamate fermentation is the simultaneous production of glutamate together with lysine. In lysine fermentation, sulfate usually accumulates in the fermentation medium as a counterion of the positively charged lysine, whereas during glutamate fermentation, ammonium accumulates as a counterion of the negatively charged glutamate. Both ions might become by-products in the respective process, representing severe cost factors and environmental issues. Consequently, we explored the use of a C. glutamicum strain suitable for the simultaneous production of both amino acids to eliminate the counterion by-products. For this purpose, we derived from the lysine-overproducing strain C. glutamicum ssp. lactofermentum AJ12937 by nitrosoguanidine treatment strain EK117 as a Tween-40 temperature-sensitive mutant. As expected, EK117 overproduced glutamate when the cultivation temperature is shifted to 37˚C even without Tween-40 addition. A simplified flux analysis of EK117 before and after temperature shift is shown in Figure 19.3. At the lower temperature almost no glutamate is formed, whereas at Sugar

Lys

0.23

0.23

Sugar

1.00 Lys

0.02

Biomass

Glu

EK117 30°C

0.48 0.24

1.52 0.32

Biomass

Glu

EK117 37°C

FIGURE 19.3 Simple flux analysis of strain EK117 producing either L-lysine or L-lysine plus L-glutamate as induced by a temperature shift. Relative fluxes are given, with the sugar uptake flux at 30˚C set to one.

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the higher temperature the relative flux to glutamate is 0.32%, compared to the sugar uptake flux at the lower temperature [40,93]. Most interestingly, due to glutamate production the specific sugar consumption rate is increased by 50%, which is a wellknown feature of glutamate-producing conditions of C. glutamicum, in which the specific sugar consumption rate may be increased twofold as compared to nonproducing conditions. A possible reason could be the reduced oxoglutarate dehydrogenase activity under producing conditions and a reduced energy status of the cell interacting with the regulation of sugar uptake. Due to the higher sugar uptake and precursor supply the flux toward lysine is also increased. This example shows the profitable simultaneous production of glutamate and lysine, which has interesting consequences on the metabolism of C. glutamicum and a high potential as an environmental issue for waste reduction.

19.3 INDUCTION OF L-GLUTAMATE OVERPRODUCTION As already mentioned, C. glutamicum was isolated in a screening system by the fortunate coincidence that the medium used was devoid of biotin [50,104]. Systematic studies revealed that L-glutamate overproduction is induced when wild-type strains are cultured at biotin-limiting concentrations of 0.3 ng ml–1 [88]. Also, in the presence of excess biotin glutamate overproduction can be induced by the addition of 3 mg ml–1 of specific detergents such as polyoxyethylene sorbitane monopalmitate (Tween 40) or polyoxyethylene sorbitane monostearate (Tween 60) [13,97]. However, monolaurate or monooleate esters, (Tween 20 and Tween 80, respectively) are not effective. Meanwhile a number of techniques are known to induce glutamate overproduction (Table 19.1). For instance, addition of some beta-lactam antibiotics,

TABLE 19.1 Leak Model and Metabolic Flux Change (MFC) Model Leak Model

MFC Model

Trigger factor

Biotin limitation Surfactant addition Penicillin addition Oleate auxotrophy Glycerol auxotrophy

Biotin limitation Surfactant addition Penicillin addition Oleate auxotrophy

Targets

Cell surface Cytoplasmic membrane Fatty acid biosynthesis Cell wall

Biotin-containing complex DtsR1 AccBC dtsR1-regulator protein (DRP)

Hypothetical effect

Increase of membrane permeability

Decrease of DtsR1 containing complex activity Metabolic flux change

Glutamate leakage through membrane

L-Glutamate

Production

445

in particular penicillin, induces L-glutamate production [67]. Furthermore strains that are auxotrophic for glycerol or fatty acids accumulate L-glutamate in the culture medium [35,64,70]. Different from other amino acid fermentative productions, Lglutamate production by the wild-type of C. glutamicum was originally supposed to be linked to growth inhibition. However, in the case of the dtsR1 mutant (see Section 19.4.6) growth inhibition appears not to be a necessary prerequisite for Lglutamate production.

19.3.1 FROM LEAK MODEL

TO

METABOLIC FLUX CHANGE MODEL

It is challenging to cast the diverse observations leading to glutamate overproduction into a coherent model. Historically, the leak model was formulated to explain glutamate formation with C. glutamicum. In this model, the focus is on fatty acid synthesis and membrane alteration: Biotin is the cofactor of acetyl-CoA carboxylase, which is the first enzyme for fatty acid biosynthesis. Due to a decreased fatty acid and phospholipid content under biotin limitation [71,88], the cell membrane permeability for glutamate is enhanced. It has also been considered that detergents and penicillin directly attack the cell membrane and cell wall, respectively, acting in the end in a similar manner [11,23,64]. However, in this model L-glutamate secretion is a passive process [4,11,23,64,71,80–82,98], and recent evidence disagrees with this model. It has been demonstrated that membrane permeability is not changed, since other amino acids, as well as acetate and ions such as H+, K+, and Cl– do not leak from the cell during glutamate overproduction [23,27,52]. Moreover, it has been suggested that a glutamate exporter is present in C. glutamicum. Furthermore, it was confirmed that the membrane fluidity does not change under biotin-limited conditions [65]. Due to these inconsistencies and a number of recent observations pointing to changes within the cytoplasm — and not merely to changes to the cell wall — the metabolic flux change model was introduced. This model is supported by the central role of the 2-oxoglutarate dehydrogenase (ODHC) activity [92]. ODHC competes with oxoglutarate dehydrogenase for the common substrate oxoglutarate. At a reduced ODHC activity, the metabolic flux is thought to be shifted from energy production via the tricarboxylic acid cycle toward L-glutamate production [36]. The details of the metabolic flux change model accommodating the various observations and dynamic flux changes are already described [44,45,47,48] and will be further outlined in the following sections.

19.4 CELLULAR CHARACTERISTICS OF GLUTAMATE OVERPRODUCTION 19.4.1 REACTIONS LEADING

TO

2-OXOGLUTARATE

As a prerequisite to the study of intracellular fluxes in C. glutamicum, a number of tools, including cloning techniques [31], the genome sequence [32], and labeling techniques, are now well-established. Using such procedures, most of the genes associated with L-glutamate biosynthesis (sugar metabolism [59,103], glycolysis and central metabolism [14,15,33,61,68,72,108], the TCA cycle [16,17,62,73,105],

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CO2 PEPC

Phosphoenolpyruvate PC

Ppyruvate Oxaloacetate

Acetyl-CoA CS

Fatty acids

Citrate NADP Isocitrate ICDH NADPH Oxoglutarate NADP GDH NADPH Glutamate FIGURE 19.4 Major reactions of relevance for L-glutamate synthesis within the central metabolism.

nitrate assimilation [3,30,34], transport [54,101], and energetics [79,94]) have been cloned and characterized. The flux over the Embden-Meyerhof pathway (EMP) and the hexose monophosphate pathway (HMP) was analyzed with 13C-labeled substrate because the latter supplies the reducing power for biosynthesis, including production of glutamate. It was shown that the flux into the HMP decreases during glutamate overproduction [29]. The EMP/HMP ratio was estimated to be 80/20 during glutamate production, in sharp contrast to lysine production where this ratio is 30/70 to 40/60. From these results, it is supposed that regulation of the EMP/HMP ratio has an important role in maintaining the balance of the metabolic network under L-glutamate overproduction. The anaplerotic reactions present at the junction between glycolysis and the TCA cycle are of particular importance for glutamate synthesis, since net carboxylation must occur (Figure 19.4). Individual enzymes of these reactions have been studied recently with respect to glutamate and lysine formation [7,76]. Interestingly, during glutamate overproduction, as triggered by a temperature increase, phophoenolpyruvate carboxylase (PEPC) activity carries up to 70% of the glutamate flux, whereas pyruvate carboxylase (PC) is responsible for the remaining 30% [10]. This is in agreement with the fact that almost no PC protein was detectable under glutamate-producing conditions (unpublished data), and agrees furthermore with the fact that PC is a biotin-binding enzyme [88], but nevertheless biotin-limitation can be used for efficient glutamate production. It is interesting that L-glutamate production by wild-type C. glutamicum in response to detergent or penicillin is more or less comparable to biotin-limiting conditions, even though PC might be active in the former case where an adequate biotin level is present. This shows the robustness of the anaplerotic reactions.

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Production

447

Citrate synthase (CS), encoded by gltA, catalyzes the initial reaction of the TCA cycle and is supposed to be the rate-controlling enzyme for the entry of substrates into the cycle. However, the specific activity of CS in C. glutamicum was found to be independent of the level of substrate and of the phase of growth. The enzyme was not affected by NADH or 2-oxoglutarate, and was only weakly inhibited by ATP [89]. Amplification of gltA did not result in increased glutamate production, and its inactivation resulted in glutamate auxotrophy indicating that only one single CS is present in C. glutamicum [17]. The isocitrate dehydrogenase (ICDH) of C. glutamicum is constitutively expressed because its specific activity is independent of the substrate and the growth phase [2]. The enzyme is a monomer, in contrast to the dimeric enzyme of Escherichia coli. In addition to the different tertiary structure, the C. glutamicum enzyme exhibits a 10-fold increased activity as compared to the E. coli enzyme and also a striking increased specificity toward NADP and isocitrate. The enzyme is weakly inhibited by oxaloacetate, 2-oxoglutarate, and citrate, and is strongly inhibited by oxaloacetate plus glyoxylate [16]. An ICDH-deficient mutant (icd) was a glutamate auxotroph, and a strain overexpressing icd showed no detectable alteration of L-glutamate production, even when the glutamate dehydrogenase gene gdh was simultaneously overexpressed. These results indicate that some factor other than ICDH is rate-limiting for L-glutamate production [16]. Since both ICDH and glutamate dehydrogenase (GDH) of coryneform bacteria are dependent on NADP(H), a direct coupling, at least when considering the cellular NADP(H) balance, seems to be important for effective L-glutamate overproduction (Figure 19.4). Moreover, since much NADPH is certainly required for fatty acid synthesis, an NADPH-type GDH might be an important feature of glutamate overproduction as a physiological link to fatty acid synthesis.

19.4.2 AMMONIA INCORPORATION Oxoglutarate is reductively aminated to afford L-glutamate synthesis, and there are two principal mechanisms to achieve this. As in many bacteria (and described in detail in Chapter 14) either glutamate dehydrogenase (GDH) [18,19,20,56,60,63,84, 95,96,99,100,102] or the glutamine synthetase (GS) [30]/glutamate synthase (also known as glutamine:2-oxoglutarate aminotransferase, GOGAT) system is operative [34,101]. GDH was genetically and enzymatically analyzed in B. flavum [84], C. glutamicum, and C. callunae [19]. These studies indicated that the formation of glutamate in coryneform bacteria is mainly dependent on GDH, since (i) GDHdefective mutants of B. flavum showed L-glutamate auxotrophy [84] and lower L-glutamate production in the presence of high ammonia concentration [96], (ii) the GS/GOGAT system of coryneform bacteria was repressed at high ammonia concentrations, and (iii) GDH activity was far higher than GOGAT activity [19,95]. However, as shown with the cloned gdh gene available [18], GDH is in principle not essential for L-glutamate synthesis and excretion with the wild-type [56] albeit the situation for high-level production might be different. In gdh mutants the GS/GOGAT system substitutes the absent dehydrogenase activity [19,20]. Interestingly, upon gdh

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overexpression the intracellular glutamate pool is increased without resulting in increased excretion [56], which indicates a limiting export system. Although GDH and GS/GOGAT are often depicted as alternative mechanisms, both systems are operating in parallel, as for instance derived by an in vivo flux analysis [100]. One important aspect is that GS activity has to be regarded as a reaction removing glutamate. Indeed, glutamine synthesized by GS from glutamate is an undesirable by-product in L-glutamate production. Careful pH control of the fermentation is necessary, since at a lower pH considerable formation of glutamine occurs [63]. Recently, components of the respiratory chain and the ATPase of C. glutamicum have been studied [37,54,55,60,77,102]. Interestingly, with the mutated AtpG subunit of the H+-ATPase [79] glutamate production was abolished, although the mutant accumulated pyruvate-derived metabolites in large concentrations, as well as a considerable concentration of oxoglutarate, suggesting major cellular changes due to the altered energy situation of the mutant. Since glutamate excretion is strictly energy-dependent [5,26,27,69], the energy situation in the mutant might be unfavorable to allow export of glutamate.

19.4.3 THE SIGNIFICANCE

OF

ODHC ACTIVITY

Originally, in coryneform bacteria, ODHC activity was hardly verifiable [85], leading to the idea that the cellular concentration of oxoglutarate is rather high, therefore favoring flux towards L-glutamate. According to the leak model, when efflux of glutamate occurs, the intracellular glutamate concentration is reduced, thus abolishing any inhibition of the entire glutamate biosynthetic pathway and therefore sustaining the further diffusive efflux [83]. However, several facts clearly disagree with this view. For instance, significant ODHC activity was detected in C. glutamicum [9,86], and flux quantifications revealed a functional TCA cycle even during glutamate-overproduction [53,87,106]. Moreover, the cytoplasmic membrane may in effect be altered, but it is certainly not generally permeable [23,26,52,65]. Therefore, the metabolic flux changes related to L-glutamate overproduction were reconsidered [23,26,52,65]. Although ODHC is active in the modified model, this enzyme occupies a key position within the chain of events leading eventually to glutamate production. E. coli mutants in which the TCA is inactivated at the level of ODCH and/or succinyl-CoA synthetase overproduce L-glutamate [57,87,106]. Whereas this might be expected, it is important that in coryneform bacteria like Brevibacterium sp. a 40 to 60% decrease in ODHC activity is observed under biotin-limiting conditions as compared to biotin-rich conditions [92]. It is even more important that under different conditions resulting in glutamate overproduction the ODHC activity is reduced [36]. This is illustrated in Figure 19.5. This finding clearly relates glutamate formation of coryneform bacteria with a reduced ODHC activity.

19.4.4 CLONING

AND

ANALYSIS

OF THE ODHA

GENE

The ODHC enzyme complex has three individual enzyme activities, which are 2-oxoglutarate dehydrogenase (E1o), dihydrolipoyl transacylase (E2o), and dihydrolipoyl dehydrogenase (E3). In E. coli [8] and Bacillus subtilis [6], the genes encoding

449

1.5

300

OD 200

Glu

100

0

1.0

0.5

0

10

20

Growth (OD)

Glutamate (mM)

A

Production

B ODHC (sp A)

L-Glutamate

0.06

Biotin rich

0.04 0.02 0

0 40

30

0.08

Biotin lim. 0

OD 200

1.0

100

0.5

0

Glu 0

10

20

30

D ODHC (sp A)

1.5

300

Growth (OD)

Glutamate (mM)

C

0.04 0.03 0.02

Tween 40

0.01 0

100

0.5

0

10

20

30

Culture time (hr)

0 40

F ODHC (sp A)

1.0

Growth (OD)

Glutamate (mM)

200

0

40

10

20

30

40

Culture time (hr)

OD

Glu

30

0

0 40

1.5

300

20

0.05

Culture time (hr)

E

10

Culture time (hr)

Culture time (hr)

0.05 0.04 0.03

Penicillin

0.02 0.01 0 0

10

20

30

40

Culture time (hr)

FIGURE 19.5 Initiation of glutamate overproduction and correlation with oxoglutarate dehydrogenase activity by different means. Shown is biotin limitation (A, B), Tween-40 addition (C, D), and penicillin addition (E, F), where the arrow marks the time of the additions. On the left are shown growth and L-glutamate accumulation, whereas on the right the specific activity of the oxoglutarate dehydrogenase in the cultures at the various time points is plotted.

E1 and E2 are adjacent and form an operon, and the E3 polypeptide of ODHC and that of pyruvate dehydrogenase are identical. Complementation of an E. coli mutant devoid of E1o [105] yielded an ODHC gene (odhA) of C. glutamicum ssp. lactofermentum with interesting structure. The polypeptide of 12,457 amino acid residues consists of two distinct domains (Figure 19.6). Whereas the C-terminal part, occupying about two-thirds of the protein exhibits very high identity (44%) with the 2-oxoglutarate dehydrogenase component E1o of E. coli, the first one-third of the protein has a sequence identity of about 28% with the dihydrolipoyl transsuccinylase component of B. subtilis. This suggests that the C. glutamicum odhA gene encodes a novel bifunctional protein that combines both E1o and E2o activities. As evident

450

Handbook of Corynebacterium glutamicum

E2o

1257aa E1o

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

C. glutamicum E1o: 941aa

E2o:417 aa .......... .......... .......... .......... ..........

E1o: 933aa

B. subtilis

E2o:405 aa .......... .......... .......... .......... ..........

E. coli

Dehydrogenase .......... .......... .......... .......... ..........

Succinyltransferase E3-binding

Lipoyl

Flexible segment FIGURE 19.6 Structure of oxoglutarate dehydrogenase complex proteins. On top is shown the large OdhA polypeptide of C. glutamicum consisting of clearly identifiable E2o and E1o domains. Below are shown the separate E1o and E2o polypeptides of B. subtilis and E. coli, respectively.

from the genome sequences, this peptide fusion of ODHC is also present in other high-G+C Gram-positive bacteria, such as Mycobacterium tuberculosis, M. leprae, Rhodococcus, and Streptomyces coelicolor. The gene adjacent to pyruvate carboxylase in C. glutamicum might represent the third ODHC component (E3). In order to further strengthen the relationship between the decrease in ODHC activity and L-glutamate production by coryneform bacteria, an odhA-disrupted mutant was constructed from wild-type C. glutamicum ssp. lactofermentum (ATCC13869). This odhA-disrupted mutant accumulates L-glutamate in the presence of excess biotin at a concentration of 265 mM at a final cellular dry weight of 1.3 g l–1, whereas the wild-type accumulates merely 1.3 mM L-glutamate under these conditions at a cell concentration of 3.0 g l–1 (Y. Asakura et al., manuscript in preparation). This high concentration is similar to that obtained with the wild-type by biotin limitation. The fatty acid composition of the odhA-disrupted mutant was almost the same as that of the wild-type [1]. These results confirm that the mechanism of L-glutamate production by coryneform bacteria is not primarily related to the membrane structure and that L-glutamate production is caused by a change in metabolic flux toward L-glutamate synthesis. Indeed, using metabolic flux analysis, the decrease in ODHC activity has recently been shown to have the strongest impact on flux distribution at the metabolic branchpoint of 2-oxoglutarate [91].

L-Glutamate

Production

451

A tgtga-n6 -tcaca

T

P

T P

359

dtsR1

1987

2331

dtsR2

3941

B Propionyl-CoA carboxylase (M. leprae, human) Acetyl-CoA carboxylase (E. coli)

PccA AccB

AccC

AccBC

Acyl-CoA carboxylase (C. glutamicum)

Biotin binding site Biotin carboxy- BCCP lase domain domain

PccB AccD

AccA

DtsR1 DtsR2

Carboxyltransferase domain

FIGURE 19.7 Shown is the close proximity of dtsR1 and dtsR2 (A), and the structural similarities of the encoded proteins with domains and polypeptides of acyl-CoA carboxylases.

19.4.5 CLONING

OF DTSR

GENES

Since reduced ODHC activity is decisive for L-glutamate excretion, and ODHC redu