Handbook of Corynebacterium glutamicum

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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

Visit the Taylor & Francis Web site at http://www.taylorandfrancis.com Taylor & Francis Group is the Academic Division of T&F Informa plc.

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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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

12


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.


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


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


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


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


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


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 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


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


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.

68


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


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

70


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].

74


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


75

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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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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Handbook of Corynebacterium glutamicum 76. Yoshihama M, Higashiro K, Rao EA, Akedo M, Shanabruch WG, Follettie MT, Walker GC, and Sinskey AJ. (1985) Cloning vector system for Corynebacterium glutamicum. J. Bacteriol. 162:591–597. 77. Zhang Y, Praszkier J, Hodgson A, and Pittard AJ. (1994) Molecular analysis and characterization of a broad-host-range plasmid, pEP2. J. Bacteriol. 176:5718–5728. 78. Zheng Z-X, Ma C-P, Yan W-Y, He P-F, Mao Y-X, Sun W, Lei Z-Z, Zhu P, and Wu J-F. (1987) Restriction map of plasmid pXZ10145 of Corynebacterium glutamicum and construction of an integrated plasmid. Chin. J. Biotechnol. 3:183–188.

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

Regulation of Gene Expression

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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


85

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


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


91

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


93

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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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.

98 [69]

[70] [71]

[72]

[73]

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


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


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


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



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


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



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

Proteomics

107


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

108



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.

110


pl 3

7

10

Molecular mass (KDa)

180 130 100 70 55 45 35 25 15

10 B


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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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


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


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


133

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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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

150


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

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

151

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


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


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



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


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


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


463

373 422

448

406 187

424

378

459

486

393 315

447

480

407 445

433

413

468



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


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.


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


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



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


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


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


446 417

% Identity

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



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


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


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


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



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


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


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


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



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


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


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


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



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


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


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


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



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


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


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#


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



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


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


Na+


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


67% 54/80

85% 231/270

19% 23/116

36% 54/149

% Identity


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


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


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


3

2

2

d

Evidence

3.A.11.X.X

3.A.5.1.1

Protein TC#

DNA

Proteins


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


87% 386/440

66% 75/112

% Identity


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


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


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


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


89% 477/532

81% 212/259

65% 173/264

77% 463/597

% Identity


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


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


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


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


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




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.

180


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.


181

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].

182


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


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.

184


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.


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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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

188


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].

Export of Amino Acids and Other Solutes

189

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

190


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


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

192


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


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

194


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


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

196


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


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

198


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


199

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

200


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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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

215

216


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

218


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


219

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.



221

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

222


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

224


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


225

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


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

Central Metabolism: Sugar Uptake and Conversion 227

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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


229

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.

230


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


231

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

232


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


233

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.

234


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


235

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].

236


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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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

242


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.

Central Metabolism: Tricarboxylic Acid Cycle and Anaplerotic Reactions

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.

244


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


245

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


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

Central Metabolism: Tricarboxylic Acid Cycle and Anaplerotic Reactions 247

248


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


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

250


[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,


251

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,

252


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


253

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].

254

11.2.2 CARBON FLUX

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


255

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

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