-Myxobacteria Multicellularity and Differentiation
EDITED BY
DAVID E. WHITWORTH
Department of Biological Sciences, University of Warwick, Coventry, United Kingdom
ASM PRESS Washington, DC
Copyright 0 2008
ASM Press American Society for Microbiology 1752 N Street, N.W. Washington, DC 20036-2904
Library of Congress Cataloging-in-Publication Data Myxobacteria :multicellularity and differentiation / edited by David E. Whitworth. p. ;cm. Includes index. ISBN 978-1-5558 1-420-5 1. Myxobacterales. 2. Cell differentiation. I. Whitworth, David E. 11. American Society for Microbiology. [DNLM: 1. Myxococcales. 2. Cell Differentiation. QW 150 M999 20081 QR82.M95M98 2008 579.3’2-dc22 2007038056
All Rights Reserved Printed in the United States of America 1 0 9 8 7 6 5 4 3 2 1 Address editorial correspondence to: ASM Press, 1752 N St., N.W., Washington, DC 20036-2904, U.S.A. Send orders to: ASM Press, P.O. Box 605, Herndon, VA 20172, U.S.A. Phone: 800-546-2416; 703-661-1593 Fax: 703-66 1- 1501 Email:
[email protected] Online: estore.asm.org
Contents
Contributors Preface xu
...
vaaa
I Myxobacterial Biology
1
1 From Glycerol to the Genome 3 DALEKAISERAND MARTINDWORKIN 2 Why Cooperate? The Ecology and Evolution of Myxobacteria 17 GREGORY J. VELICER AND KRISTINAL. HILLESLAND
II Development and Motility
41
43 3 Initiation and Early Developmental Events MICHELLE E. DIODATI,RONALD E. GILL,LYNDAPLAMANN, AND MITCHELL SINGER 4
Contact-Dependent Signaling in Myxococcus xanthus: the Function 77 of the C-Signal in Fruiting Body Morphogenesis LOTTES0GAARD-ANDERSEN
5
Reversing Myxococcus xanthus Polarity DALEKAISER
6
103 Gliding Motility of Myxococcus xanthus PATRICIA HARTZELL, WENYUAN SHI, AND PHILIPYOUDERIAN
93
7 The Frz Chemosensory System of Myxococcus xanthus DAVIDR. ZUSMAN, YUKIF. INCLAN, AND TAMMIGNOT
123 V
CONTENTS
vi
III Regulatory Mechanisms
133
8 Chemosensory Signal Transduction Systems in Myxococcus xantbus 135 JOHNR. KIRBY,JAMESE. BERLEMAN,SUSANNEMULLER, DI LI, JODIEC. SCOTT,AND JANETM. WILSON 9 Transcriptional Regulatory Mechanisms during Myxococcus xantbus Development 149 LEEKROOSAND SUMIKOINOUYE 10 Two-Component Signal Transduction Systems of the Myxobacteria 169 AND PETER J. A. COCK DAVIDE. WHITWORTH
11 Protein Ser/Thr Kinases and Phosphatases in Myxococcus xanthus 191 SUMIKOINOUYE, HIROFUMI NARIYA, AND JOSEMUROZ-DORADO 12 Carotenogenesis in Myxococcus xantbus: a Complex Regulatory Network 211 MONTSERRAT EL~AS-ARNANZ, MARTAFONTES,AND S. PADMANABHAN
N
Structure and Metabolism
227
13 Composition, Structure, and Function of the Myxococcus xanthus Cell Envelope 229 ZHAOMIN YANG,XUE-YAN DUAN,MEHDIESMAEILIYAN, AND HEIDI B. KAPLAN 14 Metabolic Pathways Relevant to Predation, Signaling, and Development 241 J. SHIMKETS PATRICK D. CURTISAND LAWRENCE 15 Secondary Metabolism in Myxobacteria HELGE B. BODE AND ROLFMULLER
259
V Myxobacterial Genomics and Postgenomics
283
16 The Genomes of Myxococcus xantbus and Stigmatella aurantiaca 285 AND WILLIAM C. NIERMAN CATHERINE M. RONNING 299 17 A Postgenomic Overview of the Myxobacteria SUEN,BARRYS. GOLDMAN, AND ROYD. WELCH GARRET
VI Stigmatella and Sorangium
313
18 The Challenge of Structural Complexity: Stigmatella aurantiaca as an Alternative Myxobacterial Model 315 WULFPLAGA 329 19 Sorangium cellulosum KLAUSGERTH,OLENAPERLOVA, AND ROLFMULLER
CONTENTS
VII Analogous Systems
vii
349
20 Bdellovibrio: Lone Hunter “Cousin” of the “Pack Hunting” 351 Myxobacteria K. J. EVANS,L. HOBLEY, C. LAMBERT, AND R. E. SOCKETT 21 Bacillus subtilis Sporulation and Other Multicellular Behaviors 363 P. MORAN, JR. LEEKROOS,PATRICK J. PIGGOT,AND CHARLES 22 Developmental Control in Caulobacter crescentus: Strategies for Survival in Oligotrophic Environments 3 85 DEANNE L. PIERCEAND YVESV. BRUN 23 Developmental Biology of Heterocysts, 2006 JINDONG ZHAOAND C. PETERWOLK
397
24 Multicellular Development in Streptomyces 4 19 MARIEA. ELLIOT,MARK J. BUTTNER,AND JUSTINR. NODWELL 25 A Eukaryotic Neighbor: Dictyostelium discoideum DERRICK BRAZILLAND RICHARDH. GOMER
439
26 Multispecies Interactions and Biofilm Community Development 453 S. JAKUBOVICS, AND PAULE. KOLENBRANDER, NICHOLAS NATALIA I. CHALMERS
VIII Myxobacterial Methods
463
27 Myxococcus xanthus: Cultivation, Motility, and Development 465 PENELOPE I. HIGGS AND JOHNP. MERLIE, JR. 28 Myxococcus xanthus: Expression Analysis 479 FRANK-DIETRICH MULLER AND JIMMY SCHOUV JAKOBSEN 29 Genetic Tools for Studying Myxococcus xanthus Biology A. MURPHY AND ANTHONY G. GARZA KIMBERLY
503 30 Sorangium cellulosum Methods ANKETREUNER-LANGE, SABRINA Dog, AND TINAKNAUBER Index
513
491
Contributors
E. BERLEMAN Dept. of Microbiology, University of Iowa, 51 Newton Rd., Iowa City, IA 52242
JAMES
HELGE B. BODE Institut fur Pharmazeutische Biotechnologie, Universitat des Saarlandes, Postfach 15 11 50, 66041 Saarbrucken, Germany DERRICK BRAZILL Dept. of Biology, Hunter College, 695 Park Ave., New York, NY 10021
YVESV. BRUN Dept. of Biology, Indiana University, Bloomington, IN 47405
MARK J. BUTTNER Dept. of Molecular Microbiology, John Innes Centre, Colney Lane, Norwich, NR4 7UH, United Kingdom
NATALIA I. CHALMERS Oral Biofilm Communication Unit, Oral Infection and Immunity Branch, National Institute of Dental and Craniofacial Research, National Institutes of Health, Bethesda, MD 20892, and University of Maryland School of Dentistry, Baltimore, MD
PETERJ. A. COCK MOAC Doctoral Training Centre, University of Warwick, Coventry CV4 7AL, United Kingdom
PATRICK D. CURTIS Dept. of Microbiology, University of Georgia, Athens, GA 30602
...
VZZZ
CONTRIBUTORS MICHELLE E. DIODATI Section of Microbiology, University of California-Davis, Davis, CA 95616 SABRINA DoB Dept. of Microbiology and Molecular Biology, Justus-Liebig-University Giessen, Heinrich-Buff-Ring 26-32, D-35392 Giessen, Germany XUE-YANDUAN Dept. of Molecular and Cellular Biology, Baylor College of Medicine, Houston, TX 77030
MARTIN DWORKIN Dept. of Microbiology, University of Minnesota, Minneapolis, M N 55455-0312 MONTSERRAT EL~AS-ARNANZ Departamento de Genetica y Microbiologia (Unidad Asociada a1 IQFR-CSIC), Facultad de Biologia, Universidad de Murcia, 30100 Murcia, Spain
MARIE A. ELLIOT Dept. of Biology, McMaster University, 1280 Main St. West, Hamilton, ON, Canada L8S 4K1 MEHDIESMAEILIYAN Dept. of Natural Sciences, University of HoustodDowntown, Houston, TX 77002 K. J. EVANS Institute of Genetics, School of Biology, University of Nottingham Medical School, Nottingham NG7 2UH, United Kingdom MARTAFONTES Departamento de Genetica y Microbiologia (Unidad Asociada a1 IQFRCSIC), Facultad de Biologia, Universidad de Murcia, 30100 Murcia, Spain
ANTHONYG. GARZA Dept. of Biology, Syracuse University, Syracuse, NY 13244 KLAUSGERTH Helmholtz-Zentrum fur Infektionsforschung GmbH, InhoffenstraBe 7, 3 8 124 Braunschweig, Germany
RONALDE. GILL Dept. of Microbiology, University of Colorado Health Sciences Center, Denver, CO 80262 BARRYS. GOLDMAN Monsanto Company, St. Louis, M O 63167 RICHARDH. COMER Howard Hughes Medical Institute and Dept. of Biochemistry and Cell Biology, MS-140, Rice University, 6100 S. Main St., Houston, TX 77005-1892
ix
CONTRIBUTORS
X
PATRICIA HARTZELL Dept. of Microbiology, Molecular Biology and Biochemistry, University of Idaho, Moscow, ID 83844
PENELOPE I. HIGGS Dept. of Ecophysiology, Max Planck Institute for Terrestrial Microbiology, Marburg, Germany 35043
KRISTINAL. HILLESLAND Dept. of Civil and Environmental Engineering, University of Washington, Seattle, WA 98195-2700 L. HOBLEY Institute of Genetics, School of Biology, University of Nottingham Medical School, Nottingham NG7 2UH, United Kingdom YUKIF. INCLAN Dept. of Molecular and Cell Biology, University of California, Berkeley, CA 94720-3204 SUMIKOINOUYE Dept. of Biochemistry, Robert Wood Johnson Medical School, Piscataway, NJ 08854 SCHOUVJAKOBSEN 194 Chemin du Siege, Residence le Mirabaou, F-06140 Vence, France
JIMMY
NICHOLAS S. JAKUBOVICS Oral Biofilm Communication Unit, Oral Infection and Immunity Branch, National Institute of Dental and Craniofacial Research, National Institutes of Health, Bethesda, MD 20892
DALEKAISER Dept. of Biochemistry and Dept. of Developmental Biology, Stanford University Medical School, Stanford, CA 94305
HEIDIB. KAPLAN Dept. of Microbiology and Molecular Genetics, University of Texas Medical School, Houston, TX 77030 R. KIRBY Dept. of Microbiology, University of Iowa, 51 Newton Rd., Iowa City, IA 52242
JOHN
TINAKNAUBER Dept. of Microbiology and Molecular Biology, Justus-Liebig-University Giessen, Heinrich-Buff-Ring 26-32, D-35392 Giessen, Germany
PAULE. KOLENBRANDER Oral Biofilm Communication Unit, Oral Infection and Immunity Branch, National Institute of Dental and Craniofacial Research, National Institutes of Health, Bethesda, MD 20892 LEEKROOS Dept. of Biochemistry and Molecular Biology, Michigan State University, East Lansing, MI 48 824
CONTRIBUTORS C. LAMBERT Institute of Genetics, School of Biology, University of Nottingham Medical School, Nottingham NG7 2UH, United Kingdom
DI LI Dept. of Microbiology, University of Iowa, 51 Newton Rd., Iowa City, IA 52242 JR. JOHNP. MERLIE, Dept. of Molecular and Cell Biology, University of California-Berkeley, Berkeley, CA 94720
TAMMIGNOT Laboratoire de Chimie Bacttrienne, 31, Chemin Joseph Aiguier, 13009 Marseille, France
P. MORAN, JR. CHARLES Dept. of Microbiology and Immunology, Emory University School of Medicine, Atlanta, GA 30322 MULLER FRANK-DIETRICH Dept. for Ecophysiology, Max Planck Institute for Terrestrial Microbiology, Karl-von-Frisch-Strage, D-35043 Marburg, Germany ROLFMULLER Institut fur Pharmazeutische Biotechnologie, Universitat des Saarlandes, Postfach 15 11 50,66041 Saarbriicken, Germany SUSANNE MULLER Dept. of Microbiology, University of Iowa, 51 Newton Rd., Iowa City, IA 52242
JosB MUAOZ-DORADO Departamento de Microbiologia, Facultad de Ciencias, Universidad de Granada, E-18071 Granada, Spain KIMBERLY A. MURPHY Dept. of Biology, Syracuse University, Syracuse, NY 13244
HIROFUMI NARIYA Dept. of Biochemistry, Robert Wood Johnson Medical School, Piscataway, NJ 08854 WILLIAMC. NIERMAN J. Craig Venter Institute, 9712 Medical Center Dr., Rockville, MD 20850 R. NODWELL Dept. of Biochemistry and Biomedical Sciences, Health Sciences Centre, McMaster University, Hamilton, ON, Canada L8N 325
JUSTIN
S. PADMANABHAN Instituto de Quimica-Fisica “Rocasolano,” CSIC, 28006 Madrid, Spain OLENAPERLOVA Institut fur Pharmazeutische Biotechnologie, Universitat des Saarlandes, Postfach 15 11 50,66041 Saarbrucken, Germany
xi
CoNTRIB UTORS
xii
DEANNE L. PIERCE Dept. of Biology, Indiana University, Bloomington, IN 47405
J. PIGGOT PATRICK Dept. of Microbiology and Immunology, Temple University School of Medicine, Philadelphia, PA 19140
WULFPLAGA Zentrum fur Molekulare Biologie der Universitat Heidelberg (ZMBH), University of Heidelberg, 69120 Heidelberg, Germany LYNDAPLAMANN School of Biological Sciences, Cell Biology and Biophysics, University of Missouri-Kansas City, Kansas City, M O 64110
CATHERINE M. RONNING J. Craig Venter Institute, 9712 Medical Center Dr., Rockville, MD 20850
C. SCOTT Dept. of Microbiology, University of Iowa, 51 Newton Rd., Iowa City, IA 52242
JODIE
WENYUAN SHI Dept. of Oral Biology, School of Dentistry, and Dept. of Microbiology, Immunology and Molecular Genetics, School of Medicine, University of California, Los Angeles, Los Angeles, CA 90095 LAWRENCE J. SHIMKETS Dept. of Microbiology, University of Georgia, Athens, GA 30602
MITCHELL SINGER Section of Microbiology, University of California-Davis, Davis, CA 95616 R. E. SOCKETT Institute of Genetics, School of Biology, University of Nottingham Medical School, Nottingham NG7 2UH, United Kingdom LOTTES0GAARD-ANDERSEN Max Planck Institute for Terrestrial Microbiology, Karl-von-Frisch Str., 35043 Marburg, Germany
GARRET SUEN Dept. of Biology, Syracuse University, Syracuse, NY 13244 ANKETREUNER-LANGE Dept. of Microbiology and Molecular Biology, Justus-Liebig-University Giessen, Heinrich-Buff-Ring 26-32, D-35392 Giessen, Germany GREGORY J. VELICER Dept. of Biology, Indiana University, Bloomington, IN 47405, and Max Planck Institute for Developmental Biology, 72076 Tiibingen, Germany ROY D. WELCH Dept. of Biology, Syracuse University, Syracuse, NY 13244
CONTRIBUTORS DAVIDE. WHITWORTH Dept. of Biological Sciences, University of Warwick, Coventry CV4 7AL, United Kingdom M.WILSON Dept. of Microbiology, University of Iowa, 51 Newton Rd., Iowa City, IA 52242
JANET
C. PETERWOLK MSU-DOE Plant Research Laboratory and Dept. of Plant Biology, Michigan State University, E. Lansing, MI 48824
ZHAOMIN YANG Dept. of Biology, Virginia Polytechnic Institute and State University, Blacksburg, VA 24060
PHILIPYOUDERIAN Dept. of Biology, Texas A & M University, College Station, TX 83843 JINDONG
ZHAO
State Key Laboratory of Protein and Plant Genetic Engineering, College of Life Sciences, Peking University, Beijing 100871, China DAVIDR. ZUSMAN Dept. of Molecular and Cell Biology, University of California, Berkeley, CA 94720-3204
...
xzzz
Preface
Since their discovery, the myxobacteria have proven to be enduring sources of wonder and inspiration for microbiologists. Myxobacteria exhibit several behaviors that are rare within the bacterial world but commonplace in eukaryotes, including multicellular development and cellular differentiation. They have consequently been used as amenable model organisms for studies into the general principles of such behaviors-studies that have addressed the evolutionary, ecological, and molecular mechanisms involved. This book focuses on myxobacterial multicellularity and differentiation, but attempts to paint a broader canvas by also describing analogous behaviors seen in a wide range of microbiological systems. In recent decades biology has been revolutionized by the emergence of techniques capable of probing the molecular basis of cellular behavior. The ability to manipulate an organism’s genes, to characterize their protein products, and to extract information from molecular sequences has led to an ever-increasing understanding of cellular processes. In 1984, the first book on myxobacteria was published (edited by E. Rosenberg), and this was followed in 1993 by a second volume (edited by M. Dworkin and D. Kaiser; American Society for Microbiology). In these works it is apparent that the appropriate genetic tools had been developed to enable research into fundamental features of myxobacterial behavior, with particular attention being paid to the regulation of motility and multicellular development. With the advent of a new millennium, further technological advances have continued to revolutionize biology. The ability to routinely determine the entire genome sequence of an organism, to simultaneously assess expression of every gene within that genome, and to identify changes in that organism’s global pool of proteins has resulted in a flood of molecular biological data. These large sets of data are now being actively generated and exploited by researchers of myxobacterial biology. The complete genome sequence for the model myxobacterium Myxococcus xanthus has recently become available, along with those of three xv
xvi
PREFACE less well-characterized myxobacteria (Sorangium cellulosum, Stigmatella aurantzaca, and Anaeromyxobacter dehalogenans). The historical voyage of discovery from glycerol induction of sporulation to the advent of the Myxococcus xanthus genome sequence provides a plot for the introductory chapter of this volume (chapter l),kindly provided by the editors of the last collected volume on myxobacterial biology. The picture that emerges is that the myxobacteria are extremely intricate organisms, with complexity rivaling that of many eukaryotes. Dworkin and Kaiser’s preface to the 1993 book stated that “we are thus hopeful and cautiously optimistic that the next edition of this book will see the emergence of insights about fruiting body morphogenesis, the mechanisms of multicellular communication and coordination, the mechanism and function of rippling, and the mechanism of gliding motility, to name only a few of the fascinating aspects of myxobacterial biology.” This optimism was not misplaced. Fourteen years later, chapters in this book review major progress in our understanding of many features of myxobacterial biology. Indeed, some topics that were single chapters in the 1993 book are now entire sections in this volume. For example, descriptions of the mechanisms of motility span four chapters in this book (chapters 5 through 8). Similarly, our understanding of multicellular development has progressed significantly (chapters 3 and 4), as has knowledge of regulatory mechanisms (chapters 9, 10, 11, and 15). Myxobacteria continue to attract significant pharmaceutical interest through their production of bioactive secondary metabolites, and myxobacterial metabolism forms the topic of two chapters (chapters 13 and 14). The genome sequence of M . xanthus ushered myxobacterial research into the post-genomic era (chapter 17). Since then, comparative genomic analyses have continued to provide insights into the molecular biology of other myxobacteria, particularly Sorangium cellulosum (chapter 19) and Stigmatella aurantiaca (chapters 16 and 18). The first chapters in the book underpin the others by providing historical and ecological/evolutionary contexts for contemporary myxobacterial research (chapters 1 and 2). In their preface to the 1993 book, Dworkin and Kaiser correctly warned about the dangers of focusing on a single model organism. In addition to descriptions of M . xanthus, chapters in this volume present the biology of two other myxobacteria: Sorangium cellulosum and Stigmatella aurantiaca (chapters 18 and 19). Behaviors exhibited by the myxobacteria can also be found in other (often very different) organisms. I am therefore delighted to include seven diverse examples of microbial multicellularity and differentiation (chapters 20 through 26), enabling myxobacterial biology to be set in a much broader context. I am especially grateful to the authors for chapters on proteobacterial predation and differentiation, cyanobacterial differentiation, development and sporulation in gram-positive bacteria, eukaryotic multicellularity and differentiation, and multispecies biofilm development. The myxobacterial research community currently numbers around 40 laboratories across the globe, and further expansion is to be encouraged. To aid researchers who are unfamiliar with the myxobacteria but wish to start working with these organisms, chapters have also been included that describe the most commonly used methods for cultivating, manipulating, and characterizing myxobacteria (chapters 27 to 30). I hope that these chapters will also act as a compendium of techniques for existing myxobacteria researchers. At the start of a new millennium it is tempting to think that we are finally beginning to comprehend the complex behavior of the myxobacteria. While huge leaps have indeed been made in our knowledge of molecular mechanisms (particularly those governing motility and fruiting body formation), the significance of
PREFACE
xvii
those mechanisms for the physiology of the myxobacteria in their natural environment is still largely unappreciated. Molecular ecology and evolutionary analyses are starting to address such gaps in our understanding, but there is still much to learn. I hope this volume stimulates seasoned myobacteriologists and interested amateurs alike, and it is to be hoped that the next book on the myxobacteria will be able to claim a true understanding of this most complex of prokaryotes. Particular thanks must go to Heidi Kaplan (University of Texas, Houston), who helped greatly during the conception and early stages of this project. I would also like to thank Carolyn Love (also at the University of Texas, Houston) for clerical assistance, and Greg Payne and Ellie Tupper at ASM Press for their invaluable and humane support. All chapter authors deserve special thanks for timely submission of manuscripts. Every chapter in this volume has been rigorously peer reviewed, and I would like to thank all the reviewers for assisting with this process. Finally, I would very much like to thank all the members of the myxobacteria research community, and my family, for their encouragement and support for this project and for the stimulating environment that they provide.
David E. Whitworth W a r w i c k , 2007
References Rosenberg, E. (ed). 1984. Myxobacteria. Development and Cell Interactions. SpringerVerlag, New York, NY. Dworkin, M., and D. Kaiser (ed.). 1993. Myxobacteria 11. American Society for Microbiology, Washington, DC.
Mvxobacteria 1 siblogy
1
Myxobacteria: Multiceiiularity and Differentiation Edited by David E. Whitworth 0 2008 ASM Press, Washington, D.C.
Dale Kaiser Martin Dworkin
From Glycerol to the Genome
GLYCEROL INDUCTION OF MYXOSPORES Dorothy Powelson’s laboratory at Purdue University had been studying the nature of the cell surface of Myxococcus xanthus and had been attempting to compare the chemical composition of the cell walls of vegetative cells and myxospores. Seeking a more convenient method of collecting myxospores than harvesting fruiting bodies, her group had explored techniques for converting vegetative cells into myxospores in liquid culture. The method they settled on involved exposing the cells to high concentrations of sucrose, which induced the conversion of vegetative cells to round, optically refractile, somewhat resistant cells, which they concluded were spores (Adye and Powelson, 1961).Dworkin’s laboratory at the University of Minnesota was interested in the processes that were involved in the cellular morphogenesis of the rod-shaped vegetative cells to the round myxospores. However, when they examined the sucrose induction method more closely they became convinced, based on the sequence of morphological events during the conversion, that the cells were converting to osmotically resistant spheroplasts rather than to myxospores. Nevertheless, the conversion of a rod-shaped cell to a sphere, albeit not a bona fide myxospore, was sufficiently interesting
1
that they decided to try to understand the mechanism of the conversion to spheroplasts. They looked at the effect of a variety of other polyhydroxy compounds and found that glycerol at an optimal concentration of 0.5 M was able to induce the conversion of vegetative cells to myxospores rapidly (within 120 min), synchronously, and quantitatively (Dworkin and Gibson, 1964).A more detailed description of the process followed, which demonstrated that glycerol-induced myxospores were able to germinate, were resistant to elevated temperature, W irradiation, and sonication, and mimicked the sequence of morphological stages during the formation of fruiting body myxospores (Dworkin and Sadler, 1966; Ramsey and Dworkin, 1968; Sadler and Dworkin, 1966; Sudo and Dworkin, 1969). While David Zusman (Zusman, 1984) pointed out some important differences between glycerol-induced and fruiting body myxospores, glycerol induction quickly became a favorite vehicle for comparing the properties and processes of vegetative cells and myxospores, albeit with careful qualifications. Following the pioneering work of Powelson, White et al. (White et al., 1968) characterized the peptidoglycan of M. xanthus and showed that unlike the murein of Escherichia coli and other gram-negative bacteria, which
Dale Kaiser, Departments of Biochemistry and Developmental Biology, Stanford University Medical School, Stanford, CA 94305. Martin Dworkin, Department of Microbiology, University of Minnesota, Minneapolis, MN 55455-03 12.
3
4 existed as a continuous bag-shaped macromolecule (Weidel and Pelzer, 1965), the sacculus of M. xanthus existed as discrete patches of peptidoglycan held together by trypsin-sensitive material. Moreover, the peptidoglycan of vegetative cells contained substantial amounts of covalently bound glucose. Bacon et al. (Bacon et al., 1975) showed that during glycerol-induced myxospore formation there was a substantial shift in carbon flow to polysaccharide synthesis, which was accompanied by an increased amount of cross-linking via diaminopimelic acid and a substantial decrease in the amount of peptidoglycan-linked glucose in the myxospore cell wall. White (White, 1984) suggested that the patchy quality of the peptidoglycan and the above changes during glycerol induction were causally related to the shape change during myxospore formation. Despite structural differences between fruiting body and glycerol-induced myxospores, the TnSlac insertion mutation, Q7536, described below, simultaneously blocked the development of glycerol-induced spores as well as fruiting body spores as they changed their shape from rod to sphere (Licking et al., 2000). The discovery of a common step encouraged the study of glycerolinduced conversion of the cylindrical vegetative rods to the spherical myxospores as a simple model of cellular morphogenesis.
MYXO FILMS A series of momentous events in the history of myxobacterial research took place between 1965 and 1974. During this time Hans Reichenbach (Fig. 1)was working on his Ph.D. dissertation research in the laboratory of Hans Kiihlwein (Fig. 2) at the University of Gottingen. Reichenbach, with the collaboration of the Institut fur den Wissenschaftlichen Film in Gottingen, created a series of masterful time-lapse photomicrographic films demonstrating the behavior of a variety of different myxobacteria. These films illustrated their cellular morphology, cell division, gliding motility, swarming behavior, aggregation, fruiting body formation, myxospore and sporangiole formation and germination, feeding on prey bacteria, and pervasive cell-cell interactions (Kuhlwein and Reichenbach, 1968; Reichenbach, 1965, 1966, 1968, 1974). One of the films illustrated a unique and mysterious rippling wave behavior that Reichenbach referred to as rhythmic oscillations (Reichenbach, 1965). In 1974, excerpts from these films were edited into a 13-min-long mosaic by Dworkin, which has since served to introduce the myxobacteria to large numbers of fascinated microbiologists, some of whom were influenced to study these organisms in their laboratories. (This film can be
MYXOBACTERIAL BIOLOGY
Figure 1 Hans Reichenbach in 1980, collecting samples of soil in the Loire Valley during the Myxo meeting in Poitiers, France.
viewed as part of the chapter “The Myxobacteria” in the online edition of The Prokaryotes). The “myxo movies,” as they have come to be known, document the unique myxobacterial grade of multicellularity, one of Nature’s many explorations of that state. They challenged myxobacteriologists to explain it, and some of the challenges have been taken up.
DENSITY-DEPENDENT GROWTH ON CASEIN By 1976 it had already become clear that a defining feature of myxobacterial behavior was the pervasive tendency of cells to maintain a high cell density. An examination of Reichenbach’s films revealed that even though individual cells could momentarily leave the swarm, the tendency was for the swarm to remain intact throughout feeding and development (A- and S-motility were yet to be distinguished). The demonstration that experimental induction of fruiting body formation required a high cell density inoculum (Wireman and Dworkin, 1975) was consistent with the broader notion that “. . . the life cycle of the myxobacteria is directed at all times to preserving the existence or the potential for the swarm” (Dworkin, 1972). Furthermore, the ability of the myxobacteria to hydrolyze a wide variety of
TO THE GENOME 1. FROMGLYCEROL
macromolecules as a nutrient source had led to the proposal that “. . . generation of optimal concentrations of lower molecular weight subunits of these polymers will depend on a certain optimal density of cells excreting the hydrolases. In other words, a wolf-pack effect” (Dworkin, 1973). Experimental support for this idea was absent until Eugene Rosenberg began thinking about the problem. Rosenberg had begun work on the myxobacteria while a Professor of Microbiology at UCLA and continued that work after his emigration to Tel Aviv University in Israel. On his sabbatical in Minneapolis in 1976 he formulated an experimental strategy for addressing the problem. We called on our colleague Ken Keller, a chemical engineer, to help guide us through the mathematical analyses; from that collaboration there emerged clear and definitive proof that while a single cell of M. xanthus could grow perfectly well on an enzymatic hydrolyzate of casein, alternatively, when that cell was presented with the intact casein molecule as a substrate, growth was clearly cell density-dependent (Rosenberg et al., 1977). This was an obvious reflection of the fact that the cells, when growing on a macromolecular substrate, were at the mercy of diffusion of their hydrolytic enzymes away from the cell and diffusion of the low-molecular-weight products of the hydrolysis toward the cell. Slowly, the modus vivendi of the myxobacteria began to make sense-their predilection for insoluble macromolecular substrates, their ability to move by gliding on a solid surface, the density dependence of their growth and fruiting body formation, and the collection of myxospores in fruiting bodies in what were probably optimally sized packages of resistant resting cells poised to form a swarm upon germination-which helps put the entire episode of density-dependent growth into a larger biological perspective.
CAROTENOIDS Myxobacteria are well known for their carotenoid production, and the exciting story of its regulation is told in chapter 12. Experiments on carotenogenesis began in 1964 in Dworkin’s group. Robert Burchard, then a graduate student, was trying to isolate a bacteriophage for M. xanthus. At that time, no phages for any of the myxobacteria had been isolated. Lawns of M. xanthus were exposed to soil extracts and, after incubation, examined for characteristic plaques. After a series of unsuccessful attempts, one series of plates was left on a windowsill after having been incubated for about 1 week. When preparing to discard the plates, Burchard noticed that the bacterial lawn on the uppermost plate in the
5
Figure 2 Martin Dworkin (left) and Hans Kiihlwein (right), during the Myxo meeting in Poitiers, France. pile had pockmarks suggestive of phage plaques. However, repeated attempts to transfer the plaque material to fresh lawns produced no new plaques. Eventually, we realized that the top plate in the pile had been exposed to the sunlight shining through the window. Subsequent experiments also revealed that the plates that had been incubated in the dark did not produce the characteristic myxobacterial carotenoids, which we later showed served a photoprotective purpose and were photoinduced. Furthermore, the photosensitizing pigment was identified as protoporphyrin IX, a cytochrome precursor, which accumulated in the cells only as they entered the stationary phase (Burchard and Dworkin, 1966).
FRUITING BODY DEVELOPMENT The most famous segments of Reichenbach’s movies deal with aggregation and fruiting body formation of several different genera of myxobacteria. How does the order evident in the shape of a species-specific fruiting body arise from the apparent disorder of a swarm? Although each species built a different structure, all were initiated by starvation. How does starvation induce fruiting body development? To investigate starvation, it was first necessary to find which nutrients were required. Dworkin had shown
6 that although M . xanthus grew well in Casitone and preferred peptides, it could be grown in a chemically defined medium that contained 17 amino acids with a generation time of 8 to 10 h (Dworkin, 1962). Despite improvements that shortened the generation time to 6.5 h (Witkin and Rosenberg, 1970), it was not clear which amino acids in the medium were essential and which were serving as sources of carbon. In 1978, Anthony Bretscher identified leucine, isoleucine, and valine as essential amino acids and found that vitamin B,, was essential for the synthesis of methionine (Bretscher and Kaiser, 1978). Those requirements were confirmed in 2005 by the absence in the M. xanthus genome of genes for the biosynthesis of the essential branched-chain amino acids (Goldman et al., 2006). Dworkin (Dworkin, 1962) observed that phenylalanine addition stimulated growth, and Bretscher verified its low rate of synthesis that severely limited growth. Using Bretscher’s minimal synthetic medium, Colin Manoil found that limitation for any amino acid, whether essential or nonessential, induced fruiting body development (Manoil and Kaiser, 1980). He found that starvation for carbon, energy, or phosphorus also induced development, while Kimsey showed that neither purine nor pyrimidine starvation would induce development (Kimsey and Kaiser, 1991). These observations suggested how starvation might be recognized. Since a complete set of aminoacyl tRNAs is essential for protein synthesis, the absence of one or more aminoacyl tRNAs could readily be perceived by a halt in protein synthesis. In M. xanthus, as in many other bacteria, the absence or shortage of any one of the charged tRNAs causes a ribosome to synthesize guanosine tetraphosphate (and pentaphosphate), (p)ppGpp, by transferring P-P from ATP to GTP, to trigger a stringent response. Mitchell Singer showed that M. xanthus has a relA gene (ppGpp synthetase) and that (p)ppGpp was necessary and sufficient to initiate fruiting body development (Singer and Kaiser, 1995). It soon was recognized that this was a multicellular response.
MOLECULAR GENETICS It was clear in 1972 that genetic experiments would be necessary to complement the ongoing biochemical and cellular studies, if the program of fruiting body development were to be found. Mutations that blocked a single step in development would need to be analyzed, and this would require a means for gene transfer. In a search for transduction, we found that E. coli phage PlCM both adsorbed to and injected DNA into M. xanthus (Kaiser and Dworkin, 1975). Subsequently, transposon Tn5,
MYXOBACTERIAL BIOLOGY which conferred kanamycin resistance, was found to tag interesting mutations (Kuner et al., 1981; Sodergren and Kaiser, 1983).Meanwhile, several generalized transducing phages, Mx4, Mx8, and Mx9, were found for Myxococcus (Campos et al., 1978; Martin et al., 1978). Lee Kroos constructed TnSlac, which carried a promoterless trp-lac fusion fragment inserted near the end of the transposable element. Transposition of Tn5lac into M. xanthus transcriptionally fused lacZ to an adjacent promoter, creating a reporter for that promoter (Kroos and Kaiser, 1984). Kroos and Adam Kuspa created a library of TnSlac insertions that were expressed at different times in development (Kroos et al., 1986).Leon Avery developed transposon replacement in situ, enabling construction of doubly marked mutant strains with transposons at two different sites in the same strain that brought resistance to two different antibiotics (Avery and Kaiser, 1983). The transposon library made it possible to analyze signaling mutants (Kuspa et al., 1986). Electroporation is effective for introducing M. xanthus DNA into M. xanthus (Ramaswamy et al., 1997) and has been systematically improved (Youderian et al., 2003).
CELLS SIGNAL EACH OTHER Two groups (Hagen et al., 1978, and LaRossa et al., 1983) analyzed the same set of M. xanthus mutants that were conditionally defective in the formation of sporefilled fruiting bodies. Interestingly, these mutants were unable to develop on their own, but when mixed with wild-type cells or with certain other mutants, they were able to develop. Pairwise testing of 57 mutants divided them into four groups (A to D), and a fifth group was discovered by John Downard (Downard, 1993). Complementation did not result from cross-feeding of essential intermediary metabolites that were diffusible because the mutants, with the exception of group E, could grow on a minimal defined medium (LaRossa et al., 1983). It now appears that groups A and C define extracellular signals that are exchanged between cells, while the ByD, and E groups are more likely consequences of cell contact-dependent exchanges of materials between cells, like stimulation (Kaiser, 2004; Nudleman et al., 2005).
A-Signal and Responses to A-Signal Medium conditioned by Myxococcus development was found to contain a heat-stable and a heat-labile form of A-signal activity. Lynda Plamann found that heat-labile A-signal was a mixture of proteases and proteins that were sensitive to those proteases (Plamann et al., 1992). Then, Adam Kuspa showed that heat-stable A-signal is a set of amino acids and small peptides containing those
1. FROMGLYCEROL TO THE GENOME amino acids (Kuspa et al., 1992). Most likely amino acids are the primary A-signal molecules, while the extracellular release of proteases and proteins generates first peptides and then A-signal amino acids. Because M. xanthus can develop fruiting bodies in a medium devoid of external amino acids, developmental proteins are synthesized at the expense of cellular reserves. Mitchell Singer found that A-signal helps M . xanthus assess the nutrient available to it for development so it can complete the program (Singer and Kaiser, 1995). As Myxococcus faces starvation, it must choose between initiating fruiting body development with differentiation of spores and slow growth at a rate compatible with whatever level of nutrient happens to be available. Spore counts indicate that fewer than 1% of cells undertaking fruiting body development eventually become spores. If nutrient is on its way to eventual exhaustion, then slowing growth to match the level of residual nutrient will lead to slower and slower growth, until death ensues. Since either option kills the majority of cells, the better choice from the cell’s perspective depends on its projection of nutrient availability in its near future. M . xanthus appears to use a stringent response and the cell-density-dependent A-signal to predict the nutrient available to it. Heidi Kaplan, analyzing suppressors of asg mutants, called sas mutants (Kaplan et al., 1991), found that a sensor histidine kinase, Sass, and a response regulator, SasR, constituted a two-component system that senses the extracellular level of A-signal. If the level is adequate, sasR protein triggers the expression of certain A-signaldependent genes, such as a 4 5 2 1 (Kaplan and Plamann, 1996; Keseler and Kaiser, 1995).
C-Signal Protein Active C-signal was purified from detergent-extracted cell membranes. A bioassay of restoring aggregation and sporulation, in vitro, to a mutant lacking a csgA gene was employed to monitor purification (Kim and Kaiser, 1990a).Because csgA mutants arrest fruiting body development having only formed traffic jams (Kaiser, 2003) and very few viable spores, the sporulation assay was quite sensitive. A 17-kDa protein that could restore development was purified from starved wild-type cells (Kim and Kaiser, 1990a). No C-signal activity was recovered from extracts of cells that were not starved, or from csgA mutant cells (Kim and Kaiser, 1990a). Nevertheless, the genomic sequence of the csgA gene predicted a 25-kDa protein homologous to the short-chain alcohol dehydrogenase family of enzymes (Lee et al., 1995). The discrepancy in size was resolved by Sune Lobedanz, then a graduate student working with Lotte Sargaard-Andersen.
7 Using antibodies to fragments of p25, Lobedanz showed that p17 corresponded to a C-terminal fragment of p25, in agreement with Kim’s amino acid sequence data (Kim and Kaiser, 1990b). Lobedanz also detected serine protease activity in an M. xanthus cell surface fraction that was capable of cleaving p25, removing an N-terminal peptide with the NAD+ binding site, leaving p17 adhering to the surface (Lobedanz and Sargaard-Andersen, 2003). Evidently, p17 is the signal, and processing by a cell-surface protease ensured that the signal is transmitted to another cell; a cell never signals itself.
C-Signal Transmission An unexpected observation that nonmotile mutants of M. xanthus arrested fruiting body development at the same morphological stage as a csgA mutant (Kroos et al., 1988) suggested that motility might be required for C-signaling because it took place between cells in end-to-end contact. Seung Kim tested this hypothesis by mechanically forcing nonmotile cells into end-to-end alignment (Kim and Kaiser, 1 9 9 0 ~ )He . used the asymmetry of the long rod-shaped M. xanthus cells to orient them lengthwise as they fell into the narrow grooves produced by scoring agar with a fine-grained aluminum oxide abrasive paper. Phase-contrast microscopy revealed that cells, which had settled into the grooves, were indeed oriented with their long axes parallel to the axis of the groove (Kim and Kaiser, 1 9 9 0 ~ )Then, . a second independent line of experiments on traveling waves also led to the conclusion that C-signal transmission was specific for end-to-end contact, as opposed to contact with the side of a cell. Traveling wave crests, colliding at any angle, were never seen to interfere with each other (Sager and Kaiser, 1994). Instead, colliding wave crests reflected from one another, providing additional evidence that the C-signal is transmitted through the ends of two cells in contact. Moreover, signaling restricted to cell ends was in quantitative agreement with the mathematical analysis of traveling waves (Igoshin et al., 2001; Welch and Kaiser, 2001). Thus, C-signal carries information about the local cell density as well as the orientation with respect to neighboring cells.
C-Signal Transduction Work from several laboratories has shown that C-signal is a morphogen that, in a dose-dependent manner, manages cell movement, initiates the expression of many developmentally regulated genes, and triggers sporulation (Kim and Kaiser, 1991; Kruse et al., 2001; Li et al., 1992; Sargaard-Andersen et al., 1996). Thomas Gronewold discovered a positive-feedback loop in the C-signal response circuit that is controlled by the act operon of
MYXO BACTERIAL BIOLOGY
8
five cotranscribed genes (Gronewold and Kaiser, 2001, 2002). This feedback is responsible for raising the number of C-signal molecules per cell from a few at 3 h poststarvation to several hundred by 18 h (Gronewold and Kaiser, 2001; Kim and Kaiser, 1991). The rise serves to time C-signal-dependent gene expression and to restrict that expression to the fruiting body (Julien et al., 2000; Kroos and Kaiser, 1987). Small aggregates enlarge when the number of Csignal molecules per cell rises above a moderate threshold. Above that threshold, a responding cell decreases its reversal frequency and increases its speed (Jelsbak and Ssgaard-Andersen, 1999, 2002; Ssgaard-Andersen et al., 2003). Responding cells tend to form a chain or stream whose cells are in frequent end-to-end contact with each other. Upon contact, they signal each other (Jelsbak and Ssgaard-Andersen, 2000). Streaming was observed by Brian Sager, when he tracked individual cells inside nascent fruiting bodies. He observed one-half of the cells to circulate clockwise and one-half counterclockwise (Sager and Kaiser, 1993).None of the tracked cells reversed during the experiment. As cells stream, they have more opportunities to C-signal each other and there is still more positive feedback. Finally, when the number of C-signal molecules per cell rises to the higher threshold for sporulation, the dev operon is expressed (Ellehauge et al., 1998; Kroos et al., 1986). dev Operon The last three genes of the dev operon, devT, devR, and devS, have been characterized (Boysen et al., 2002; Kroos et al., 1990; Thony-Meyer and Kaiser, 1993). Linda Thony-Meyer showed that devR devS double mutants are able to aggregate, but fail to sporulate (Thony-Meyer and Kaiser, 1993). Bryan Julien showed that dev expression was spatially restricted to the fruiting body (Julien et al., 2000). Ellen Licking and Lisa Gorski showed that since the Tn51ac::R7536 mutant aggregates normally and fails to sporulate, this reporter gene could be placed in a sporulation pathway downstream of dev (Licking et al., 2000). A consequence of dev action is that the differentiation of myxospores occurs only after aggregation is complete.
MOTILITY Many years ago myxobacterial motility was described by Jahn (Jahn, 1924), and by Schmidt-Lorenz and Iciihlwein (Schmidt-Lorenz and Kuhlwein, 1968) in terms of the microscopic structure of cells and their appendages. Henrichsen distinguished gliding by myxobacteria from the flagellar swimming motility observed in many
other bacteria as movement on a surface usually in the direction of a cell’s long axis (Henrichsen, 1972).A large stride forward was taken by Hans Reichenbach, working with Hans Kiihlwein, who made movies that showed cell movement during cell division, swarming, and the building of fruiting bodies by several different species (Kuhlwein and Reichenbach, 1968; Reichenbach, 1966, 1968,1974). Experimental genetic investigations of myxobacterial motility began with the isolation and examination of mutants by Burchard (1970), and by MacRae and McCurdy (1976). Then, Jonathan Hodgkin investigated many motility mutants all derived from the same genetically characterized strain. Comparisons between mutants revealed two different swarm patterns, indicative of two different gliding engines, referred to as engine A and engine S (Hodgkin and Kaiser, 1979a, 1979b). Hodgkin found roughly equal numbers of mutants that lacked either engine A or engine S, which he distinguished by their different swarm patterns. Normally the two engines cooperate with each other, but when there is only engine A (A+,?-),the cell clusters in swarms are long and strung out; when there is only engine S (A-S’), the clusters are short and stubby. In both types of swarms, cells move singly and in groups. A-S- colonies have sharp edges, much like those of E. coli, and the cells are unable to swarm or to form fruiting bodies.
S-Motility The S-engines are composed of type IV pili (TFP), long thin retractile hairs that extend from the front end of a cell and have the ability to retract, pulling the cell forward. M. xanthus TFP share at least 15 proteins that are involved in extending and retracting pili with motile Neisseria and Pseudomonas (whose motility is traditionally called twitching despite its relation to gliding), and with Synechocystis (Wall and Kaiser, 1999). The role played by a number of the Pi1 proteins has been worked out (Nudleman and Kaiser, 2004). For instance, PilT, an inner membrane protein, is an AAA motor protein necessary for pilus retraction. PilT mutants have pili, but because those pili are unable to retract, the cells lack S-motility (Li et al., 2003; Wu et al., 1997).Like the PilT mutants, another group of S- mutants that was discovered by David Morandi-the dsp (dispersed growth) mutants-have pili but nevertheless lack S-motility. The Dsp mutants lack fibrils.
Fibrils Stimulated by a report on “filaments” given at the 1978 Myxo meeting at Spring Hill, MN, M. Dworkin
TO THE GENOME 1. FROMGLYCEROL
saw their similarity to “myxonemata” described 15 years earlier by Walter Fluegel, who had viewed them by staining living cells with India ink. Dworkin recalls that both findings were received with mild interest and then quietly ignored until 1988, when Arnold and Shimkets showed that these filaments, which they termed “fibrils,” were responsible for cell-cell cohesion of M. xanthus (Arnold and Shimkets, 1988a, 198813). The late Rich Behmlander morphologically characterized the fibrils by means of low-voltage scanning electron microscopy. He showed that fibril formation required cells to be at a high cell density on a solid surface and that they consisted of a polysaccharide matrix with associated proteins (Behmlander and Dworkin, 1991, 1994a, 199413).He demonstrated that the so-called fibrils were not preparational artifacts by showing that when fibrils were decorated with carbon particles, examination of the cells by phase-contrast microscopy revealed the presence of fibrils which had not been subjected to fixation or dehydration (Behmlander and Dworkin, 1994a, 1994b). Arnold and Shimkets (Arnold and Shimkets, 1988a, 1988b) provided unambiguous evidence that the fibrils were required for the effective social and developmental behavior of M . xanthus. This was supported by the experiments of Chang and Dworkin (Chang and Dworkin, 1994) showing that isolated fibrils, added to a fibril-minus dsp mutant, were able to rescue cohesion and development of a dsp mutant. Subsequently, Yang et al. (Yang et al., 2000) showed that isolated fibrils could also partially rescue cohesion and development in dif mutants, one class of dsp mutants. Li et al. (Li et al., 2003) provided evidence that the TFP at a leading end of a cell of M. xanthus attached to amine-containing polysaccharide from fibrillar material deposited on the agar and triggered pilus retraction. This result was consistent with an earlier observation that glucosamine, one of the components of fibril polysaccharide, blocked cellcell cohesion (Behmlander and Dworkin, 1994b).In sum, S-motility results when a pilus from one cell attaches to a network of fibrils that encloses a group of cells ahead. As the attached pilus retracts, the piliated cell pulls up to the leading cells.
A-Motility Gliding M . xanthus leaves a phase-bright trail of “slime” behind on the agar surface, which has the same width as the cell. Reichenbach’s movies showed examples, and Lars Jelsbak photographed three isolated cells at higher magnification laying trails as they moved (online movie available in Kaiser, 2003). Wolgemuth et al. (2002) observed the extrusion of ribbons of slime-like material uniquely from one end of the cell and proposed that
9 polar slime secretion from the more than 100 pores visible at the back end of a cell pushes the cell forward. This proposal found support in the work of Rosa Yu and Kaiser (2007),who showed that A-motility is perfectly correlated with unipolar slime secretion. Bipolar slime secretion in the mglA mutants results in loss of motility (Kaiser and Yu, 2005).
Elasticotaxis It must be a source of great satisfaction for an author to find that a paper written over 50 years ago continues to provoke interest and to generate new experiments. Roger Stanier was one of the most outstanding microbiologists of our era, who had made major contributions to our understanding of the phototrophic bacteria, the pseudomonads, the cyanobacteria, and Caulobacter, but whose sole published contribution to the myxobacteria was one little note. As a young master’s student at UCLA, Stanier characterized the gliding motility, nutritional physiology, and taxonomy of the marine cytophagas and concluded casually, following an earlier suggestion by Krzemieniewska, that they be classified as nonfruiting myxobacteria in the order Myxobacteriales (sic) (Stanier, 1940). Stanier then joined Kees van Niel’s laboratory at the Hopkins Marine Station in Pacific Grove, CA, where his Ph.D. dissertation resulted in a classic monograph on the CytophagalSporocytophaga group (Stanier, 1942b). He characterized the group thoroughly, provided a much more detailed rationale for classifying them among the myxobacteria, and mentioned extensive, unpublished experience with the so-called “higher” myxobacteria while in van Niel’s laboratory. It was during this period that he noted that cultures of members of the Myxococcaceae when streaked on agar slants oriented themselves and their subsequent fruiting bodies in a consistently ordered fashion relative to the streak lines on the agar. He demonstrated by means of a simple and elegant experiment that this behavior was reproducible and that the gliding cells of Chondrococcus (Myxococcus) exiguus (now classified by Reichenbach as Corallococcus exiguus) oriented themselves parallel to lines of stress in the agar substrate and subsequently formed fruiting bodies at right angles to their direction of movement. He called the phenomenon “elasticotaxis” (Stanier, 1942a).It is interesting that the later Cytophagu monograph contained numerous photographs showing a similar tactic behavior as the cellulose-decomposing cytophagas oriented themselves to the cellulose fibers. A segment in one of Reichenbach’s films showing myxobacterial lysis of prey bacteria, in which the myxobacteria seemed to head directly for the clusters of Sarcina lutea prey (Reichenbach, 1968) particularly
MYXOBACTERIAL BIOLOGY
10 intrigued Dworkin. It seemed to him a good system for demonstrating myxobacterial chemotaxis, and he set about intending to do so. As a first control, to eliminate the possibility that the myxobacteria were responding to some nonspecific physical presence of the cells, Dworkin substituted 10-pm-diameter polystyrene latex beads for the clumps of prey bacteria. To his astonishment some of the myxobacterial swarms seemed to head directly to the plastic beads. To eliminate the possibility that the beads contained a diffusible chemical perceived by the cells, Dworkin substituted washed, incinerated glass beads and watched while the cells repeated their directed movement. After many hours of watching in disbelief as the swarms moved toward the beads, and after subjecting the process to a statistical analysis, Dworkin concluded that the cells were indeed detecting the physical presence of the beads on the agar (Dworkin, 1983). A careful rereading of Stanier’s elasticotaxis paper revealed that in one of his experiments the cells had also responded in an elasticotactic fashion to sterile glass beads which he had scattered over the agar surface. This made Dworkin comfortable in suggesting that the directed movement was an elasticotactic response. More recently, Marta Fontes and Kaiser (Fontes and Kaiser, 1999) quantified the assay for elasticotaxis and, drawing on a collection of A- and S- mutants, showed that the elasticotactic response in M. xanthus requires A-motility but not S-motility. Indeed, the tactic response was enhanced by the absence of S-motility. This suggested to them that S-motility, which requires that the cells move toward other cells ahead of them, competes with elasticotaxis for setting the direction of cell movement. Since in most cases the stress direction would be different from the pilus direction, competition would result. They also observed the progressive reorientation of cells that happened to start parallel to lines of stress in agar. Almost all those cells became perpendicular within 15 min, the time required for gliding a couple of cell lengths. Kaiser (Kaiser, 2003) suggested that elasticotaxis was like following a slime trail and that oriented agarose chains in compressed agar or slime polymer chains in a trail would be expected to have similar orientating effects on an A-motile cell. It seems reasonable to suggest that in nature, the ability of myxobacteria to perceive the presence of a colony of potential prey bacteria resting on their own deformable pad of slime would be useful for predation. It would allow the myxobacteria to move directly to the prey by following a stress line, just as Dworkin’s photographs record (Dworkin, 1983). Dworkin also observed that movement toward a bead depended on A-motility but not on S-motility. Since so little is known about the
specific niches occupied by myxobacteria in the soil, this suggestion is tentative.
Stimulation Early in his investigation of the genetics of gliding motility, Hodgkin made a startling discovery concerning contacts between 111. xanthus cells. He observed that several nonmotile (A-S-) mutants could be stimulated to move transiently by contact with wild-type cells or with cells of a different mutant type. Most of his motility mutants were not rescuable, but tgl, cglB, cglC, cglD, cglE, and cglF mutants could be stimulated (Hodgkin and Kaiser, 1977). Because stimulation did not occur when contact between cells was prevented, he also learned that stimulation depends on close apposition of interacting cells (Hodgkin and Kaiser, 1977). Subsequent progress in understanding stimulation required knowing the molecular function and the cellular location of stimulatable proteins. Jorge Rodriguez-Soto showed that Tgl protein was essential for S-motility, not for A-motility, and that it is a 27-kDa lipoprotein to be found in the outer membrane of M. xanthus (Rodriguez-Soto and Kaiser, 1997a, 1997b). Dan Wall, Sam Wu, and Kaiser (1998) showed that tgl mutants make ample quantities of PilA pilin, but fail to assemble it into pili. Nudleman, Wall, and Kaiser (Nudleman et al., 2006) demonstrated that Tgl is required for the assembly of PilQ monomers into a multimeric secretin channel in the outer membrane. The pilus elongates through the assembled PilQ channel as it extends outside the cell (Nudleman et al., 2006). These authors also showed that Tgl for PilQ assembly could be provided by stimulation from another cell. Finally, by separating the stimulated recipient cells from donor cells, Nudleman et al. (2005) demonstrated that Tgl protein was transferred from donor to recipient cell. They also demonstrated the transfer from donor to recipient of CglB protein, which is essential for A-motility (Nudleman et al., 2005). CglB had been found to be a lipoprotein (Rodriguez and Spormann, 1999) that also localized to the outer membrane of M. xanthus (Simunovic et al., 2003). The concentration of Tgl and CglB proteins in stimulated cells was found to be similar to the concentration in donor cells, as if the donor and recipient cells shared their mobile outer membrane proteins equally, which thereby created a primitive tissue (Nudleman et al., 2005).
Reversing the Engines Reichenbach’s movies of myxobacterial swarms show individual cells moving alternately along both directions of their long pole-to-pole axis (Kuhlwein and Reichenbach, 1968).Jelsbak’s movie, noted above, also
TO THE GENOME 1. FROMGLYCEROL
illustrates frequent alternation of gliding direction. No cell has either a permanent head or a permanent tail. Studies of the frizzy mutants of M. xanthus have shown that cells have a well-defined average frequency of gliding reversal that is inherited and controlled by the frz genes (Blackhart and Zusman, 1985). Tracking cells in the traveling waves of M. xanthus has shown very regular reversals at 8-min intervals (Welch and Kaiser, 2001). M. xanthus cells clearly do not reverse randomly in time; instead, each cell appears to have its own reversal clock (Igoshin et al., 2004). Finally, reversal is not coupled to the cell cycle because M. xanthus cells can reverse their gliding direction 20 or more times in traveling waves in a single cell cycle. Wild-type M. xanthus colonies on the surface of agar are flat disk-shaped swarms that taper down to a monolayer of cells at their edge. That edge is observed to spread outward symmetrically at a constant rate for many days (Burchard, 1974; Kaiser and Crosby, 1983).The colony of an A-S- strain, which lacks motility, measures the colony expansion that is due to growth. Since an A-Scolony expands at about one-eighth the rate at which the motile (A+S+)swarm expands, the outward spreading of the swarm can be attributed largely to motility, although growth is essential (Kaiser and Crosby, 1983). Since the swarm rate is observed to increase with the cell density of the inoculum, individual cells appear to help each other move outwards (Kaiser and Crosby, 1983). The A-engines cooperate through slime trail following; the S-engines employ pili that bind fibrils on another cell. The advantage of swarming is evident for colonies on the surface of nutrient agar: by spreading the cells out, swarming decreases competition between feeding cells for nutrient that comes from below the agar surface. The selective advantage of swarming appears to be the enhancement of nutrient absorption by the swarm. As described above, both engines are polar. There is evidence that the two engines are located at opposite poles of the cell, and while the S-engine pulls at one end, the A-engine pushes at the other. In addition, the maximum swarming rate of an A+S+strain is 1.6 pm min-' while the sum of the maximum rates of two strains, each having one of the two engines, is 1.0 pm min-l (Kaiser and Crosby, 1983). This synergism implies that the Sand A-engines occupy opposite poles. It also indicates that for each cell, reversal of one engine is highly correlated with reversal of the other engine. Finally, there is evidence that reversal is necessary for swarming. A mutant with Tn5 inserted in the frzE gene reverses only once every 2 h (Shi and Zusman, 1995) compared to the wild type with a reversal every 7 min, and the frzE mutant fails to swarm (Shi et al., 1993). Qualitatively, outward
11 swarming might be explained by periodic reversals that need not be coordinated between cells. A quantitative theory of swarming is needed to relate the reversal frequency, the speed, and the direction of individual cells to the overall rate of swarm expansion. A quantitative theory could be used to test whether periodic reversals that are independent from cell to cell are sufficient to explain swarming, or whether reversals of different cells in the same area must be coordinated by a gradient of nutrient availability. Although the frequency of reversals depends on the genotype and whether cells are growing or are starving (Jelsbak and Sargaard-Andersen, 2000,2002), directional bias in response to an attractant has yet to be demonstrated (Ward and Zusman, 1997). Nevertheless, it has been proposed that 16:l phosphatidylethanolamine, a lipid found in the outer membrane of M. xanthus, may serve as an attractant during fruiting body aggregation because it appears to change the reversal frequency (Kearns et al., 2001).
THE GENOME M . xanthus exhibits multicellular behavior: it feeds as a coordinated group of cells, and when its food supply nears exhaustion, many thousands of cells cooperate to build a fruiting body and to differentiate spores within it. The recently released genome sequence of M. xanthus by Monsanto and The Institute for Genomic Research sheds new light on the origin and regulation of myxobacterial multicellularity (GenBank assession no. CPOOOll3). The sequence revealed a large genome of 9.14 Mb, 7,388 predicted coding sequences, and a gene density comparable to that of E. coli. The sequence confirms the membership of M. xanthus in the delta subgroup of proteobacteria, but the six other sequenced deltaproteobacteria range from 3.6 to 3.9 Mb in size. This raises the question-how did the M. xanthus genome grow to 9.1 Mb from the size of 3.9 Mb or below that is found in all other deltaproteobacteria? A myxobacterium-specific genome expansion is suggested by the similarity in size of M. xanthus and two species of Stigmatella (a rough draft of one Stigmatella sequence is available at The Institute for Genomic Research). A substantial part of the expansion can be traced to the duplication of individual genes followed by differentiation of function between duplicates. About one-half (48%)of the 7,388 predicted coding sequences in M. xanthus are members of families of closely related sequences. Moreover, 16% of the M. xanthus genome constitutes families of paralogous proteins. Paralogs are more closely related to one another than they are to any protein from any other organism whose genome has been sequenced. Since it appears that many of these paralogs
MYXOBACTERIAL BIOLOGY
12
arose by gene duplication in the ancestors of M. xanthus, they most likely represent myxobacterial-lineage-specific duplications. The lineage-specific duplications found in 111.xanthus are far from a random sample of the genome; rather, they involve particular functions, and because the duplications survived, those functions must have been important to M. xanthus. Among the duplications are genes for sensing and signaling, proteolysis, predation, and development. For sensing, the lineage-specific duplications include Ser/Thr protein kinases (a total of 99 proteins; at least 20 are lineage specific), sigma 54 enhancer-binding proteins (about 50), 137 sensor and hybrid histidine protein kinases and a roughly equal number of response regulators, and about 40 extracytoplasmic sigma factors that respond to extracellular signals (Helmann, 2002). In combination these proteins may create complex sensory circuits for regulating transcription in M. xanthus. There is experimental evidence for a Ser/Thr protein kinase, sigma 54 enhancer-binding protein combination (Jelsbak et al., 2005). Such multistep regulators would resemble those that control embryonic development in multicellular eukaryotes in having sensory input at multiple steps of the circuit. Apparently the multistep regulators were gained by M. xanthus at the expense of one-component regulators that are common in bacteria. The total number of transcriptional regulatory proteins in M. xanthus lies in the range expected for a genome of 9.1 Mb, but the fraction devoted to multistep regulators is increased compared to other soil organisms with large genomes, like Streptomyces coelicolor. For predation and scavenging, the lack of the ZlvC and ilvD genes, which are necessary for branched-chain amino acid biosynthesis, results in the observed requirement for leucine, isoleucine, and valine in the laboratory and suggests that their natural diet consists mainly of protein. In addition, the genome includes many genes whose products may be used for predation. Among the duplicated genes are copies of chaperones and related proteases-DnaK (15copies), DnaJ, GrpE, ClpX, ClpAB, ClpP, HslU, HslV, HtpX, GroEL (2 copies), and Lon (2 copies). Almost 9 % of the M. xanthus genome encodes enzymes that produce secondary metabolites. This amounts to twice the capacity of S. coelicolor or Streptomyces avermitilis, organisms that are well-recognized producers of secondary metabolites. Considering the large variety of duplicated and differentiated genes, the proposed increase in genome size from an ancestral deltaproteobacterium is likely to have occurred over a long stretch of time. It seems likely that predation involving cooperative cell interactions evolved first, allowing M. xanthus to hunt like a pack of wolves
(Dworkin, 1973). Complex sensory modules, such as those now found in M. xanthus, may have enabled scavenging and predation of whole surface colonies of bacteria found in nature. Given the capacity to hunt, fruiting body development could have evolved to enhance survival of the colony when food is exhausted. Many fruiting body cysts are elevated above the surface, sticky, and attached by a thin stalk-properties that would permit a package of spores to be broken off and then to stick to a small animal that is hunting for food. Thousands of spores would thereby be transported together, and it is likely that fruiting bodies are optimally sized and positioned packages of spores. Insects can move long distances in soil and have sophisticated senses for finding food. When the animal encounters nutrients and feeds, the package of myxospores may be deposited in the organic matter. The spores would germinate together and instantly create a feeding swarm of myxobacteria. This scenario, obviously speculative, calls out for experimental tests and refinements. Those experiments would constitute no more than one step in the overall task of understanding the whole organism that is revealed to us in the M. xanthus genome.
References Adye, J. C., and D. M. Powelson. 1961. Microcyst of Myxococcus xanthus: chemical composition of the wall. J. Bacterial. 81:780-785. Arnold, J. W., and L. Shimkets. 1988a. Inhibition of cell-cell interactions in Myxococcus xanthus by Congo red. J. Bacterial. 1705765-5770. Arnold, J. W., and L. J. Shimkets. 1988b. Cell surface properties correlated with cohesion in Myxococcus xanthus. J. Bacteriol. 1705771-5777. Avery, L., and D. Kaiser. 1983. In situ transposon replacement and isolation of a spontaneous tandem genetic duplication. Mol. Gen. Genet. 191:99-109. Bacon, K. D., R. H. Clutter, M. Kottel, M. Orlowski, and D. White. 1975. Carbohydrate accumulation during myxospore formation in Myxococcus xanthus. J. Bacteriol. 124:16351636. Behmlander, R. M., and M. Dworkin. 1991. Extracellular fibrils and contact-mediated cell interactions in Myxococcus xanthus. J. Bacteriol. 173:7810-7821. Behmlander, R. M., and M. Dworkin. 1994a. Integral proteins of the extracellular matrix fibrils of Myxococcus xantbus. J. Bacteriol. 176:6304-6311. Behmlander, R. M., and M. Dworkin. 1994b. Biochemical and structural analyses of the extracellular matrix fibrils of Myxococcus xanthus. J. Bacteriol. 176:6295-6303. Blackhart, B. D., and D. Zusman. 1985. Frizzy genes of Myxococcus xanthus are involved in control of frequency of reversal of gliding motility. Proc. Natl. Acad. Sci. USA 82~8767-8770.
TO THE GENOME 1. FROMGLYCEROL
Boysen, A., E. Ellehauge, B. Julien, and L. Ssgaard-Andersen. 2002. The DevTprotein stimulates synthesis of FruA, a signal transduction protein required for fruiting body morphogenesis in Myxococcus xanthus. J. Bacteriol. 184:1540-1546. Bretscher, A. P., and D. Kaiser. 1978. Nutrition of Myxococcus xanthus, a fruiting myxobacterium. J. Bacteriol. 133:763768. Burchard, R. P. 1970. Gliding motility mutants of Myxococcus xanthus. J. Bacteriol. 104:940-947. Burchard, R. P. 1974. Growth of surface colonies of the gliding bacterium Myxococcus xanthus. Arch. Microbiol. 96:247254. Burchard, R. P., and M. Dworkin. 1996. Light-induced lysis and carotenogenesis in Myxococcus xanthus. J. Bacteriol. 180535-545. Campos, J., J. Geisselsoder, and D. Zusman. 1978. Isolation of bacteriophage MX4, a generalized transducing phage for Myxococcus xanthus. 1.Mol. Biol. 119:167-178. Chang, B. Y., and M. Dworkin. 1994. Isolated fibrils rescue cohesion and development in the Dsp mutant of Myxococcus xanthus. J. Bacteriol. 176:7190-7196. Downard, J. 1993. Identification of esg, a genetic locus involved in cell-cell signaling during Myxococcus xanthus development. ]. Bacteriol. 175:7762-7770. Dworkin, M. 1962. Nutritional requirements for vegetative growth of Myxococcus xanthus. J. Bacteriol. 84:250-257. Dworkin, M., and S. Gibson. 1964. A system for studying microbial morphogenesis: rapid formation of microcysts in Myxococcus xanthus. Science 146:243-244. Dworkin, M., and W. Sadler. 1966. Induction of cellular morphogenesis in Myxococcus xanthus. I. General description. J. Bacteriol. 91:15 16-151 9. Dworkin, M. 1972. Myxobacteria: new directions in studies of prokaryotic development. Crit. Rev. Microbiol. 1:435-452. Dworkin, M. 1973. Cell-cell interactions in the Myxobacteria. Symp. Soc. Gen. Microbiol. 23:125-147. Dworkin, M. 1983. Tactic behavior of Myxococcus xanthus. J. Bacteriol. 154:452-459. Ellehauge, E., M. Norregaard-Madsen, and L. Ssgaard-Andersen. 1998. The FruA signal transduction protein provides a checkpoint for the temporal coordination of intercellular signals in M . xanthus development. Mol. Microbiol. 30:807-813. Fontes, M., and D. Kaiser. 1999. Myxococcus cells respond to elastic forces in their substrate. Proc. Natl. Acad. Sci. USA 96~8052-8057. Goldman, B. S., W. C. Nierman, D. Kaiser, S. C. Slater, A. S. Durkin, J. A. Eisen, C. M. Ronning, W. B. Barbazuk, M. Blanchard, C. Field, C. Halling, G. Hinkle, 0. Iartchuk, H. S. Kim, C. Mackenzie, R. Madupu, N. Miller, A. Shvartsbeyn, S. A. Sullivan, M. Vaudin, R. Wiegand, and H. B. Kaplan. 2006. Evolution of sensory complexity recorded in a myxobacterial genome. Proc. Natl. Acad. Sci. USA 103: 15200-15205. Gronewold, T. M. A., and D. Kaiser. 2001. The act operon controls the level and time of C-signal production for M. xanthus development. Mol. Microbiol. 40:744-756. Gronewold, T. M. A., and D. Kaiser. 2002. act operon control of developmental gene expression in Myxococcus xanthus. J. Bacteriol. 184:1172-1179.
13 Hagen, D. C., A. P. Bretscher, and D. Kaiser. 1978. Synergism between morphogenetic mutants of Myxococcus xanthus. Dev. Biol. 64:284-296. Helmann, J. D. 2002. The extracytoplasmic function (ECF) sigma factors. Adv. Microb. Physiol. 46:47-110. Henrichsen, J. 1972. Bacterial surface translocation: a survey and a classification. Bacteriol. Rev. 36:478-503. Hodgkin, J., and D. Kaiser. 1977. Cell-to-cell stimulation of movement in nonmotile mutants of Myxococcus. Proc. Natl. Acad. Sci. USA 74:2938-2942. Hodgkin, J., and D. Kaiser. 1979a. Genetics of gliding motility in M . xanthus (Myxobacterales): genes controlling movement of single cells. Mol. Gen. Genet. 171:167-176. Hodgkin, J., and D. Kaiser. 197913. Genetics of gliding motility in M . xanthus (Myxobacterales):two gene systems control movement. Mol. Gen. Genet. 171:177-191. Igoshin, O., A. Mogilner, R. Welch, D. Kaiser, and G. Oster. 2001. Pattern formation and traveling waves in myxobacteria: theory and modeling. Proc. Natl. Acad. Sci. USA 98~14913-14918. Igoshin, O., A. Goldbetter, D. Kaiser, and G. Oster. 2004. A biochemical oscillator explains the developmental progression of myxobacteria. Proc. Natl. Acad. Sci. USA 101:15760-15765. Jahn, E. 1924. Beitrage zur botanischen Protistologie. I. Die Polyangiden. Gebruder Borntraeger, Leipzig, Germany. Jelsbak, L., and L. Ssgaard-Andersen. 1999. The cell-surface associated C-signal induces behavioral changes in individual M . xanthus cells during fruiting body morphogenesis. Proc. Natl. Acad. Sci. USA 965031-5036. Jelsbak, L., and L. Ssgaard-Andersen. 2000. Pattern formation: fruiting body morphogenesis in Myxococcus xanthus. Curr. Opin. Microbiol. 3:637-642. Jelsbak, L., and L. Ssgaard-Andersen. 2002. Pattern formation by a cell-surface associated morphogen in M . xanthus. Proc. Natl. Acad. Sci. USA 99:2032-2037. Jelsbak, L., M. Givskov, and D. Kaiser. 2005. Enhancer-binding proteins with a forkhead-associated domain and the sigma54 regulon in Myxococcus xanthus fruiting body development. Proc. Natl. Acad. Sci. USA 102:3010-3015. Julien, B., A. D. Kaiser, and A. Garza. 2000. Spatial control of cell differentiation in Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 97:9098-9103. Kaiser, A. D., and C. Crosby. 1983. Cell movement and its coordination in swarms of Myxococcus xanthus. Cell Motil. 3:227-245. Kaiser, D., and M. Dworkin. 1975. Gene transfer to myxobacterium by Escherichia coli phage P1. Science 187:653-654. Kaiser, D. 2003. Coupling cell movement to multicellular development in myxobacteria. Nut. Rev. Microbiol. 1:45-54. Kaiser, D. 2004. Signaling in myxobacteria. Annu. Rev. Microbiol. 58:75-98. Kaiser, D., and R. Yu. 2005. Reversing cell polarity: evidence and hypothesis. Curr. Opin. Microbiol. 8:216-221. Kaplan, H. B., A. Kuspa, and D. Kaiser. 1991. Suppressors that permit A signal-independent developmental gene expression in Myxococcus xanthus. J. Bacteriol. 173:1460-1470. Kaplan, H. B., and L. Plamann. 1996. A Myxococcus xanthus cell density-sensing system required for multicellular development. FEMS Microbiol. Lett. 139539-95.
14
MYXOBACTERIAL BIOLOGY
LaRossa, R., J. Kuner, D. Hagen, C. Manoil, and D. Kaiser. Kearns, D. B., A. Venot, J. T. Bonner, B. Stevens, G.-J. Boons, 1983. Developmental cell interactions in Myxococcus: analand L. J. Shimkets. 2001. Identification of a developmental ysis of mutants. J. Bacteriol. 153:1394-1404. chemoattractant in Myxococcus xanthus through metabolic engineering. Proc. Natl. Acad. Sci. USA 98:13990-13994. Lee, B.-U., K. Lee, J. Mendez, and L. J. Shimkets. 1995. A tactile sensory system of Myxococcus xanthus involves an Keseler, I. M., and D. Kaiser. 1995. An early A-signalextracellular NAD(P)+-containingprotein. Genes Dev. 9: dependent gene in Myxococcus xanthus has a sigma-54-like promoter. J. Bacteriol. 177:4638-4644. 2964-2973. Li, S., B. U. Lee, and L. Shimkets. 1992. csgA expression Kim, S. K., and D. Kaiser. 1990a. Purification and properties of Myxococcus xanthus C-factor, an intercellular signaling entrains Myxococcus xanthus development. Genes Dev. 6:401-410. protein. Proc. Natl. Acad. Sci. USA 873635-3639. Li, Y., H. Sun, X. Ma, A. Lu, R. Lux, D. Zusman, and W. Shi. Kim, S. K., and D. Kaiser. 1990b. C-factor: a cell-cell signal2003. Extracellular polysaccharides mediate pilus retraction ling protein required for fruiting body morphogenesis of during social motility of Myxococcus xanthus. Proc. Natl. M. xanthus. Cell 61:19-26. Acad. Sci. USA 1005443-5448. Kim, S. K., and D. Kaiser. 1990c. Cell alignment required in difLicking, E., L. Gorski, and D. Kaiser. 2000. A common step ferentiation of Myxococcus xanthus. Science 249:926-928. for changing the cell shape in fruiting body and starvationKim, S. K., and D. Kaiser. 1991. C-factor has distinct aggregaindependent sporulation of Myxococcus xanthus. J. Bactetion and sporulation thresholds during Myxococcus develrial. 182:3553-3558. opment. J. Bacteriol. 173:1722-1728. Lobedanz, S., and L. Ssgaard-Andersen. 2003. Identification of Kimsey, H. H., and D. Kaiser. 1991. Targeted disruption of the the C-signal, a contact-dependent morphogen coordinating Myxococcus xanthus orotidine 5’-monophosphate decarmultiple developmental responses in Myxococcus xanthus. boxylase gene: effects on growth and fruiting-body developGenes Dev. 17:2151-2161. ment. /. Bacteriol. 173:6790-6797. T. H., and H. D. McCurdy. 1976. Gliding motilMacRae, Kroos, L., and D. Kaiser. 1984. Construction of TnSlac, a ity mutants of Myxococcus xanthus. Can. J. Microbiol. transposon that fuses lacz expression to exogenous promot22: 1282-1 292. ers, and its introduction into Myxococcus xanthus. Proc. Manoil, C., and D. Kaiser. 1980. Accumulation of guanosine Natl. Acad. Sci. USA 81:5816-5820. tetraphosphate and guanosine pentaphosphate in MyxococKroos, L., A. Kuspa, and D. Kaiser. 1986. A global analysis cus xanthus during starvation and myxospore formation. of developmentally regulated genes in Myxococcus xanthus. J. Bacteriol. 141:297-304. Dev. Biol. 117:252-266. Martin, S., E. Sodergren, T. Masuda, and D. Kaiser. 1978. Kroos, L., and D. Kaiser. 1987. Expression of many develSystematic isolation of transducing phages for Myxococcus opmentally regulated genes in Myxococcus depends on a xanthus. Virology 88:44-53. sequence of cell interactions. Genes Dev. 15340-854. Nudleman, E., and D. Kaiser. 2004. Pulling together with type Kroos, L., P. Hartzell, K. Stephens, and D. Kaiser. 1988. A IV pili. J. Mol. Microbiol. Biotechnol. 752-62. link between cell movement and gene expression argues that Nudleman, E., D. Wall, and D. Kaiser. 2005. Cell-to-cell motility is required for cell-cell signalling during fruiting transfer of bacterial outer-membrane lipoproteins. Science body development. Genes Dev. 2:1677-1685. 309~125-127. Kroos, L., A. Kuspa, and D. Kaiser. 1990. Defects in fruiting Nudleman, E., D. Wall, and D. Kaiser. 2006. Polar assembly body development caused by Tn5lac insertions in M. xanof the type IV pilus secretin in Myxococcus xanthus. Mol. thus. J. Bacteriol. 172:484-487. Microbiol. 60:16-29. Kruse, T., S. Lobendanz, N. M. S. Bertheleson, and L. SsgaardPlamann, L., A. Kuspa, and D. Kaiser. 1992. Proteins that Andersen. 2001. C-signal: a cell surface-associated morphorescue A-signal-defective mutants of Myxococcus xanthus. gen that induces and coordinates multicellular fruiting body J. Bacteriol. 174:3311-3318. morphogenesis and sporulation in M. xanthus. Mol. Microbiol. 40: 156-1 68. Ramaswamy, S., M. Dworkin, and J. Downard. 1997. Identification and characterization of Myxococcus xanthus mutants Kuhlwein, H., and H. Reichenbach. 1968. Swarming and Morphogenesis in Myxobacteria, Archangium, M~XOCOCCUS, deficient in calcofluor white binding.]. Bacteriol. 179:28632871. Chondrococcus, Chondromyces. Film C893/1965. Institut fur den Wissenschaftlichen. Film, Gottingen, Germany. Ramsey, W. S., and M. Dworkin. 1968. Microcyst germination in Myxococcus xanthus. J. Bacteriol. 95:2249-2257. Kuner, J., L. Avery, D. E. Berg, and D. Kaiser. 1981. Uses of transposon Tn5 in the genetic analysis of Myxococcus xanReichenbach, H. 1965. Rhythmic motion in swarms of Myxothus, p. 128-132. In D. Schlessinger (ed.), Microbiologybacteria. Ber. Dtsch. Bot. Ges. 78:102-105. 2 981. American Society for Microbiology, Washington, DC. Reichenbach, H. 1966. Myxococcus spp. (Myxobacterales) Kuspa, A., L. Kroos, and D. Kaiser. 1986. Intercellular signalSchwarmentwicklung und Bildung uon Protocysten. Institut ing is required for developmental gene expression in Myxofur den Wissenschaftlichen Film, Gottingen, Germany. coccus xanthus. Dev. Biol. 117:267-276. Reichenbach, H. 1968. Archangium violaceum (Myxobacteriales) Schwarmentwicklung und Bildung von Protocysten. Kuspa, A., L. Plamann, and D. Kaiser. 1992. Identification of heat-stable A-factor from Myxococcus xanthus. J . Bacteriol. Film E 777/1965. Institut fur den Wissenschaftlichen Film, 1 7 4 ~ 319-3326. 3 Gottingen, Germany.
1. FROMGLYCEROL TO THE GENOME Reichenbach, H. 1974. Chondromyces apiculatus (Myxobacteriales) Schwarmentwicklung und Morphogenese. Film E 779/1965. Institut fur den Wissenschaftlichen Film, Gottingen, Germany. Rodriguez, A. M., and A. M. Spormann. 1999. Genetic and molecular analysis of cglB, a gene essential for single-cell gliding in Myxococcus xanthus. J. Bacteriol. 181:438 1-4390. Rodriguez-Soto, J. P., and D. Kaiser. 1997a. The tgl gene: social motility and stimulation in Myxococcus xanthus. J. Bacteriol. 179:4361-4371. Rodriguez-Soto, J. P., and D. Kaiser. 1997b. Identification and localization of the tgl protein, which is required for Myxococcus xanthus social motility. J. Bacteriol. 179:4372-4381. Rosenberg, E., K. H. Keller, and M. Dworkin. 1977. Cell density-dependent growth of Myxococcus xanthus on casein. J. Bacteriol. 129:770-777. Sadler, W., and M. Dworkin. 1966. Induction of cellular morphogenesis in Myxococcus xanthus. 11. Macromolecular synthesis and mechanism of inducer action. J. Bacteriol. 91:1520-1525. Sager, B., and D. Kaiser. 1993. Two cell-density domains within the Myxococcus xanthus fruiting body. Proc. Natl. Acad. Sci. USA 90:3690-3694. Sager, B., and D. Kaiser. 1994. Intercellular C-signaling and the traveling waves of Myxococcus. Genes Dev. 8:2793-2804. Schmidt-Lorenz, W., and H. Kuhlwein. 1968. Intracellulare Bewegungsorganellen der Myxobakterien. Arch. Mikrobiol. 60:95-98. Shi, W., T. Kohler, and D. R. Zusman. 1993. Chemotaxis plays a role in the social behaviour of Myxococcus xanthus. Mol. Microbiol. 9:601-611. Shi, W., and D. R. Zusman. 1995. The frz signal transduction system controls multicellular behavior in Myxococcus xanthus, p. 419-430. In J. A. Hoch and T. J. Silhavy (ed.), TwoComponent Signal Transduction. ASM Press, Washington, DC. Simunovic, V., F. C . Gherardini, and L. J. Shimkets. 2003. Membrane localization of motility, signaling, and polyketide synthase proteins in Myxococcus xanthus. J. Bacteriol. 185:5066-5075. Singer, M., and D. Kaiser. 1995. Ectopic production of guanosine penta- and tetra-phosphate can initiate early developmental gene expression in Myxococcus xanthus. Genes Dev. 9:1633-1644. Sodergren, E., and D. Kaiser. 1983. Insertions of Tn5 near genes that govern stimulatable cell motility in Myxococcus. J. Mol. Biol. 167:295-310. Ssgaard-Andersen, L., F. Slack, H. Kimsey, and D. Kaiser. 1996. Intercellular C-signaling in Myxococcus xanthus involves a branched signal transduction pathway. Genes Dev. 10:740-754. Ssgaard-Andersen, L., M. Overgaard, S. Lobedanz, E. Ellehauge, L. Jelsbak, and A. A. Rasmussen. 2003. Coupling gene expression and multicellular morphogenesis during fruiting body formation in Myxococcus xanthus. Mol. Microbiol. 48:l-8. Stanier, R. Y. 1940. Studies on the cytophagas. J . Bacteriol. 40:619-635.
15 Stanier, R. Y. 1942a. Elasticotaxis in myxobacteria. J . Bacterial. 44:405-412. Stanier, R. Y. 194213. The cytophaga group: contributions to the biology of the myxobacteria. Bacteriol. Rev. 6:143196. Sudo, S. Z., and M. Dworkin. 1969. Resistance of vegetative cells and microcysts of Myxococcus xanthus. J. Bacteriol. 98:8 83-887. Thony-Meyer, L., and D. Kaiser. 1993. devRS, an autoregulated and essential genetic locus for fruiting body development in Myxococcusxanthus. J. Bacteriol. 175:74507462. Wall, D., and D. Kaiser. 1999. Type IV pili and cell motility (MicroReview). Mol. Microbiol. 32:l-10. Wall, D., S. S. Wu, and D. Kaiser. 1998. Contact stimulations of Tgl and type IV pili in Myxococcus xanthus. J. Bacteriol. 180:759-761. Ward, M. J., and D. R. Zusman. 1997. Regulation of directed motility in Myxococcus xanthus. Mol. Microbiol. 245385893. Weidel, W., and H. Pelzer. 1964. Bagshaped macromoleculesa new outlook on bacterial cell walls. Adv. Enzymol. 26: 193-232. Welch, R., and D. Kaiser. 2001. Cell behavior in traveling wave patterns of myxobacteria. Proc. Natl. Acad. Sci. USA 98:14907-14912. White, D., M. Dworkin, and D. J. Tipper. 1968. Peptidoglycan of Myxococcus xanthus: structure and relation to morphogenesis. J. Bacteriol. 95:2186-2197. White, D. 1984. Structure and function of myxobacteria cells and fruiting bodies, p. 51-67. In E. Rosenberg (ed.), Myxobacteria, Development and Cell Interactions. SpringerVerlag, New York, NY. Wireman, J., and M. Dworkin. 1975. Morphogenesis and developmental interactions in the Myxobacteria. Science 189~516-523. Witkin, S., and E. Rosenberg. 1970. Induction of morphogenesis by methionine starvation in Myxococcus xanthus: polyamine control. J. Bacteriol. 103:641-649. Wolgemuth, C., E. Hoiczyk, D. Kaiser, and G. Oster. 2002. How myxobacteria glide. Curr. Biol. 12:369-377. Wu, S. S., J. Wu, and D. Kaiser. 1997. The Myxococcus xanthus pilT locus is required for social gliding motility although pili are still produced. Mol. Microbiol. 23:109-121. Yang, Z., X. Ma, L. Tong, H. B. Kaplan, L. J. Shimkets, and W. Shi. 2000. The Myxococcus xanthus dif genes are required for the biogenesis of cell surface fibrils essential for social gliding motility. J. Bacteriol. 1825793-5798. Youderian, P., N. Burke, D. J. White, and P. L. Hartzell. 2003. Identification of genes required for adventurous gliding motility in Myxococcus xanthus with the transposable element mariner. Mol. Microbiol. 49555-570. Yu, R., and D. Kaiser. 2007. Gliding motility and polarized slime secretion. Mol. Microbiol. 63:454-467. Zusman, D. 1984. Developmental program of Myxococcus xanthus, p. 185-213. In E. Rosenberg (ed.), Myxobacteria. Springer, New York, NY.
Myxobacteria: Multicellularity and Differentiation Edited by David E. Whitworth 0 2008 ASM Press, Washington, D.C.
Gregory J. Velicer Kristina L. Hillesland
Why Cooperate? The Ecology and Evolution of Myxobacteria
The myxobacteria fascinate through their ability to swarm and hunt as packs and to cooperatively crawl together into complex fruiting bodies. But when starvation strikes, why should tens of thousands of individuals aggregate to sporulate in social structures rather than staying put and sporulating as individuals? What are the selective forces that have led myxobacteria to move, hunt, and develop as social units rather than as dispersed individuals? What were the first evolutionary steps in myxobacterial sociality upon which later innovations were built and integrated? What other microbes do myxobacteria eat, and what can eat the myxobacteria? Within the myxobacteria, who meets whom and who cooperates with whom? Molecular biologists have made great strides in understanding the genetics and molecular mechanisms that underlie social behavior in Myxococcus xanthus and a few other species. Much territory remains to be explored on the front of M. xanthus social genetics, with new genes that affect social traits and their combinatorial interactions being identified and analyzed at an increasing rate. However, as M. xanthus and other species become increasingly well characterized at the genetic and biochemical levels, the challenges of understanding
3
the natural ecology, evolutionary histories, and diversity of myxobacteria also loom large on the research horizon. Many fundamental ecological and evolutionary questions remain to be satisfactorily answered. Ecological and evolutionary research seeks to understand the many forces that shape the diversity and distribution of organisms, a complex endeavor that remains in its exciting infancy for the myxobacteria. In this chapter, we first describe research on general ecological and evolutionary issues with the myxobacteria such as their diversity and distribution, population structure, and issues relating to their predatory behavior. Then, we discuss what is known about the sociobiology of these organisms.
ECOLOGY AND EVOLUTION OF THE MYXOBACTERIA Investigating the Abundance, Distribution, and Diversity of Species Where do myxobacteria live and why? Why are they so abundant in some places but not others? What is their relationship to the communities in which they live? These questions form the basis of myxobacterial ecology. More generally, the science of ecology is concerned ~
~~~
Gregory J. Velicer, Department of Biology, Indiana University, Bloomington, IN 47405. Kristina L. Hillesland, University of Washington, Department of Civil and Environmental Engineering, Seattle, WA 98195-2700.
17
18 with identifying the factors that define the distribution and abundance of species in nature and the processes responsible. These factors can be abiotic, such as temperature or humidity, or biotic, such as the abundance of species that are competitors or predators. Such ecological variables may mediate the action of natural selection across genetic variants within populations and thereby differentially affect their relative frequencies. Our understanding of ecology therefore also informs core issues in evolutionary biology. Evolutionary biology seeks not only to illuminate the historical relationships of species, lineages, genomes, and genes, but also to understand the forces and processes that generated the full diversity of organismic traits (from genomes to social behaviors) in the past and those that continue to shape them in the present. Investigation of the myxobacteria from an evolutionary perspective therefore involves answering challenging questions such as the following. How do myxobacteria benefit from forming fruiting bodies, maintaining multiple motility systems, or hunting in groups? How did the tendency to produce various complex fruiting bodies arise, and under what conditions might this ability be lost? Are some myxobacterial traits more likely to be found among populations living in soil than those living in sand or water? There are two complementary research strategies to address such a wide range of ecological and evolutionary questions. The first approach is to observe, describe, and analyze features of natural populations and communities. This approach has greatly inspired and informed our current understanding of ecology and evolution. For example, the most common approach to evolutionary studies is the comparative method, in which phylogenetic relationships or past evolutionary forces are inferred from patterns of trait variation among extant organisms. Evolutionary biologists might also follow changes in genotype frequencies and other traits over time in natural populations to better understand evolutionary forces at play in the present (Grant and Grant, 2002; Hanslti and Saccheri, 2006). An ecologist may measure the abundance of multiple populations across space and time and relate that to physiological properties of the organism and other measured properties of the environment or community. Most studies of natural myxobacteria populations have involved isolating species from diverse environments across the globe and looking for relationships between particular environmental features and the ability to isolate particular species. While these approaches with natural populations are of unquestionable value and are ultimately necessary for understanding natural communities of myxobacteria,
MYXOBACTERIAL BIOLOGY they also have inherent limitations. First, as with all microorganisms, it is difficult to observe and manipulate them directly in their natural environment. Modern molecular approaches employed in environmental microbiology such as sequencing DNA and RNA from environmental samples and fluorescence in situ hybridization improve our ability to link the presence or absence of species and genes to particular communities and environments (Amann et al., 1995; Olsen et al., 1986), but it is still difficult to measure phenotypes such as the rate of swarming across a surface or the rate of prey killing in natural conditions. Second, because natural environments are so complex, it is difficult to discern which features of the environment are responsible for changes in population abundance or mediate the action of natural selection (Endler, 1986).This issue is especially relevant in evolutionary studies employing the comparative method for which there may be poor knowledge of the selective environments and other forces that shaped lineages of interest in the past. Third, rigorous tests of some hypotheses may require combinations of ecological variables that do not occur naturally. Thus, complete reliance on studies of natural populations to inform our understanding of myxobacterial ecology and evolution would limit the range of questions that can be addressed. These limitations can be overcome by using model species such as 211. xanthus to test ecological and evolutionary hypotheses under controlled laboratory conditions. Ecological questions can be addressed by constructing alternative laboratory environments with only one or a few variables that differ between them. This approach allows the researcher to rigorously test the role of spatial structure or prey abundance on the performance of a species, or stability of a simple community (Bohannan and Lenski, 2000; Elena and Lenski, 2003; Jessup et al., 2004; Kassen and Rainey, 2004). Another powerful technique is experimental evolution, in which an organism is propagated in one or more simple, controlled environments for many generations. Phenotypic characteristics of the evolved populations, particularly competitive fitness, can then be measured and compared to those of the ancestral genotype to test a hypothesis of interest (Lenski et al., 1991). Experimental evolution has been used to address evolution of aging (Rose, 1984), host-parasite coevolution (Bohannan and Lenski, 2000; Elena, 2002), the role of chance and history in shaping evolutionary trajectories (Travisano et al., 1995), the evolution of social behavior (Rainey and Rainey, 2003; Turner and Chao, 1999; Velicer and Yu, 2003),and a variety of other topics that have been reviewed by Elena and Lenski (2003) and Feldgarden et al. (2003).
AND EVOLUTION OF MYXOBACTERIA 2. ECOLOGY
As described in this chapter, such laboratory-based approaches to testing ecological and evolutionary hypotheses have been applied to the myxobacteria in recent years. These approaches not only are enhancing our understanding of the myxobacteria per se, but also have the potential to inform our understanding of ecology and evolution more generally. Microorganisms such as the myxobacteria have several unique features which increase the range and power of experimental evolution for addressing hypotheses about adaptation relative to what is possible with many eukaryotic model organisms (Elena and Lenski, 2003). First, they can be stored frozen for long periods and then resuscitated. This property allows for direct comparisons of ancestors and evolved clones if both are stored in the freezer as they are generated. Second, many microbes have very short generation times and thus allow evolution experiments thousands of generations long to be conducted within the course of a Ph.D. degree. Third, asexual reproduction allows independent populations derived from a single ancestral organism to be evolved under the same conditions. Such repetition of evolutionary trials allows the researcher to distinguish between changes caused by natural selection and those that were likely due to random forces. Since Roland Thaxter first formally described the myxobacteria late in the nineteenth century (Thaxter, 1892), most ecological and evolutionary research has focused on the isolation, description, and classification of species from a variety of environments and the definition of their phylogenetic relationships (Shimkets et al., 2005). In this section we briefly summarize this research as well as newer cultivation and molecular studies that have advanced our understanding of the diversity of the myxobacteria, their habitats, and the structure of their natural populations. We then proceed to describe some laboratory studies addressing the effects of abiotic variables on ecologically relevant phenotypes and the interactions of myxobacteria with prey species and how such interactions affect predator evolution.
Natural Diversity and Distributions
Who Are the Myxobacteria? The myxobacteria form a monophyletic group in the order Myxococcales in the delta subgroup of the proteobacteria. They are most closely related to sulfate-reducing bacteria and Bdellovibrio species, which are also predators of bacteria (Shimkets et al., 2005). For a century, taxonomic classification of myxobacterial species was based almost exclusively on morphological traits such as fruiting body shape, size, and color, swarm patterns,
19
and cell shape. However, morphological similarity does not necessarily represent genetic similarity, such that morphology-based phylogenies may fail to accurately model ancestral relationships among species and phenotypes. The advent of DNA-sequence-based classification has opened the possibility of fully understanding not only the patterns of ancestral relatedness among myxobacterial species but also the degree to which social phenotypes reflect those patterns. Sproer et al. (1999) first classified 54 myxobacterial strains representing 10 previously named genera by traditional morphological criteria such as fruiting body phenotype. Subsequently, near-complete 16s rRNA gene sequences obtained for each strain were used to construct a molecular phylogeny. Strains assigned to the same genus by morphological classification tended to cluster tightly in the 16s rRNA phylogeny, providing strong evidence that myxobacterial phenotypes reflect overall genetic relatedness at this level, at least for this one essential gene. The order Myxococcales has traditionally been subdivided into the two suborders Cystobacterineae and Sorangiineae, and the 16s rRNA gene analysis of Sproer et al. (1999) confirms a deep phylogenetic bifurcation between these suborders (Fig. 1).Within the Sorangiineae, the family Nannocystaceae (genus Nannocystis) is deeply divergent from that of Polyangiaceae (genera Chondromyces, Polyangium, and Sorangium). Some have classified the Nannocystaceae as a distinct suborder (Shimkets et al., 2005). The characteristics that define these taxonomic groups are thoroughly described by Shimkets et al. (2005) and Reichenbach (1993), but we briefly summarize the information here. The first suborder, Cystobacterineae, includes the two most thoroughly studied species, Myxococcus xanthus and Stigmatella aurantiaca. The suborder Cystobacterineae contains the families Myxococcaceae (genera M ~ X O C O C CArchangium, US, and Corallococcus) and Cystobacteriaceae (genera Cystobacter, Hyalangium, Melittangium, and Stigmatella). Vegetative cells in this suborder tend to be long rods with tapered ends. Myxospores are much shorter and rounder and tend to have capsules. Swarms of vegetative cells usually remain on the surface of the agar and tend to make striking patterns. In contrast, swarms of the suborder Sorangiineae are often embedded in the agar, sometimes forming pits in the agar surface, and they may not produce pronounced swarming patterns on surfaces. The morphology of vegetative cells of the Sorangiineae differ from those in suborder Cystobacteriaceae in that they are stout with bluntly shaped ends. Myxospores of this suborder tend to be similar in shape to vegetative cells and do not have visible capsules but are always grouped
A
\
Cystobacterineae
AND EVOLUTION OF MYXOBACTERIA 2. ECOLOGY
into sporangioles, which are thick-walled casings for groups of myxospores. All species in the Cystobacteriacede are bacteriolytic, but some species of Sorangiineae are cellulose decomposers. The suborder Sorangiineae includes the family Polyangium, and the genera Sorangium (all cellulose decomposers), Haploangium, Chondromyces (which contain some of the most elaborate fruiting bodies), Byssophaga, and Jahnia. The Nunnocystineae are closely related to the Sorangiineae. They differ morphologically from species in the other two suborders of the Myxobacteria in that species do not produce fruiting bodies, although they may produce sporangioles or spores (Shimkets et al., 2005). Images of fruiting body morphologies representative of the various taxonomic groups can be found in Reichenbach (1993) and Shimkets et al. (2005). Below the genus level, the Sproer et al. (1999)phylogeny does not clearly follow the morphologically based species definitions. This lack of resolution highlights the perennial problem of choosing species definition criteria across bacteria more generally (Gevers et al., 2005) and the question of what constitutes a myxobacterial species in particular. Recent advances in sequencing technology (Margulies et al., 2005; Velicer et al., 2006) should soon allow the definition of entire genome sequences for multiple isolates of each classified species and thus a nearoptimal data set for mapping evolutionary relationships among strains and comprehensive sequence-based criteria for species level classification.
Where Do Myxobacteria Live? Myxobacteria have been isolated all over the globe from a variety of substrates, but the most common environment for most species is soil in tropical to temperate regions (Reichenbach, 1993, 1999; Shimkets et al., 2005). Nutrient-rich soils tend to harbor more myxobacterial
21
species, but they can also be found on rocky surfaces and in pure sand (Reichenbach, 1999). Myxobacteria have also been isolated from animal dung, decaying plant material, animal bark, marine and freshwater environments, and uranium-contaminated U.S. Department of Energy sites (Petrie et al., 2003; Reichenbach, 1999). Most cultured species prefer mild temperatures (20 to 30"C), neutral pH, and high concentrations of organic matter but low ionic concentrations (Reichenbach, 1993, 1999; Shimkets et al., 2005). Thus, there is a higher abundance of myxobacteria and greater density of species in tropical and temperate soils than in locations with more extreme climatic conditions such as Antarctica or highly acidic or alkaline soils. Although myxobacteria have been cultured from these more extreme environments, it has been unclear in many cases whether the cultured organisms were actually capable of growth there (Reichenbach, 1993,1999; Shimkets et al., 2005). In recent years, the range of environmental conditions that may support active growth of myxobacteria has been shown to be greater than expected. Dawid et al. (1988) documented Polyangium and Nunnocysti5 species isolated from Antarctica that grow at 4°C but are unable to grow at moderate temperatures, while Gerth and Miiller (2005) readily isolated strains from warm arid climates with optimum growth temperatures much higher than those of previous myxobacterial isolates (42 to 44°C). Zhang et al. (2005) reported that a marine isolate of Huliangium ochraceum fails to grow in the absence of sodium chloride. Two additional halotolerant strains in the Nannocystineae suborder have also been isolated from marine environments (Iizuka et al., 2003, 1998; Zhang et al., 2005). In their vegetative state, most myxobacterial species require significant aeration, yet a new strain has been isolated from anaerobic soil enrichments. Initial phylogenetic analyses
Figure 1 (A) Neighbor-joining tree of 16s rRNAs showing the phylogenetic position of the type strains of different genera of the order Myxococcales and isolates that were assigned to myxobacterial species on the basis of morphological characteristics (e.g., fruiting bodies, myxospores, and color). The sequences of gram-negative, sulfate-reducing bacteria were used to root the dendrogram. Numbers within the dendrogram indicate the percentages of occurrence of the branching order in 100 bootstrapped trees. The bar represents 10 nucleotide substitutions per 100 nucleotides. Reprinted with permission from Sproer et al. (1999). (B) Images of fruiting bodies of various species of Myxobacteria. In the Sorangiineae, from left to right: (i) Chondromyces spp. isolated during the Microbial Diversity course at Woods Hole (copyright 1995, D. E. Graham); (ii) Chondromyces crocatus fruiting bodies (photo courtesy of Hans Reichenbach);(iii) Polyungium fumosum fruiting bodies (used with permission from Shimkets et al., 2005). In the Nannocystaceae: (i)Koflerza flava swarm on agar (not fruiting bodies). Used with permission from Shimkets et al., 2005. In the Cystobacterineae: (i) Cystobacter badius fruiting bodies on agar (used with permission from Shimkets et al., 2005); (ii) M. xalzthus fruiting bodies in soil and a fruiting body of M. xanthus on agar (photos courtesy of M. Vos and S. Kadam, respectively).
22 indicate that this strain, called “Anaeromyxobacter,” branches deeply within the Myxococcales and is capable of reducing metals and other pollutants during anaerobic growth on electron donors such as acetate (He and Sanford, 2003; Sanford et al., 2002). That myxobacteria with such diverse optimal growth conditions have been cultivated in these and other studies (Neil et al., 2005; Watve et al., 1999) suggests that our knowledge of the distribution of these organisms is limited by our methods of cultivation and scale of sampling. It has been estimated by Watve et al. (1999) that finer-scale biogeographical sampling will reveal far more species of myxobacteria than have previously been classified, perhaps severalfold more numerous. Varying the temperature, pH, and other parameters in incubation of enrichments and greater utilization of molecular methods may reveal an even greater diversity and broader geographic range of myxobacteria than has been previously known. Modern molecular approaches promise to revolutionize our ability to define myxobacterial communities and populations and are beginning to be used and developed (Vos and Velicer, 2006; Wu et al., 2005; Jiang et al., 2007). Given that the preferred growth conditions of myxobacterial isolates vary substantially, it is reasonable to expect that the distribution of species and strains across environments is nonrandom, i.e., that some species may be more prosperous in certain environments than in others. This issue has been addressed in part by perhaps the most comprehensive study of species distribution patterns to date. Dawid (2000) examined 1,398 soil samples from 64 nations or states on all continents by using standard protocols (including three different isolation methods) throughout the long-term study. Strikingly, one or more myxobacterial species could be isolated from the vast majority (91%) of soil samples worldwide, demonstrating the ubiquitous success of this order in occupying a wide spectrum of terrestrial soils. Acidic soils and soils from low-temperature zones yielded fewer species than samples with more moderate parameters, and the highest average species diversity was found in samples from tropical climate zones. Four species (Corallococcuscoralloides, Archangium gephyra, Myxococcus fulvus, and a Polyangium species) were found in more than 40% of samples, 6 species in 10 to 35% of samples, 8 species in 5 to 10% of samples, and 21 species appeared in fewer than 5 % of samples. The species distribution reported by Dawid differed from that of previous studies, highlighting the limitations of comparisons across independently designed isolation studies and the need to develop sophisticated molecular tools for rapid quantification of myxobacterial species types and distributions within soil samples.
MYXOBACTERIAL BIOLOGY Spatial Structure in the Global M. xanthus Population Genetic diversity in natural populations of higher organisms is highly structured due t o both limited migration and adaptation to local conditions, such that spatially distinct populations are often genetically differentiated at numerous genetic loci. However, the degree to which genetic variation in microbial species is nonrandomly distributed is a long-standing problem in microbial ecology (Baas Becking, 1934; Fenchel, 2003; Martiny et al., 2006). Geographically distant populations of thermophiles (Whitaker et al., 2003) and pathogens that occupy specialized, noncontiguous habitats (Linz et al., 2007) are known to show distinct patterns of genetic variation. However, a recent study has demonstrated that genotypes of the soil bacterium M. xanthus, which has a relatively contiguous distribution across terrestrial soil ecosystems, are also nonrandomly distributed across large spatial scales (Vos, 2006). In a survey of M. xanthus populations sampled at the meter scale within each of 1 0 globally distributed sites, it was found that most populations (38 of 45 pairwise comparisons) show significantly distinct patterns of genetic variation across several highly conserved genes (Fig. 2). Moreover, the degree of genetic divergence between populations was found to correlate significantly with distance between the populations. Several European populations separated by only hundreds of kilometers showed no significant divergence, but all populations separated by >1,700 km were found to be distinct. The distribution of variation below the 100-km scale in Germany was found to be largely homogeneous, indicating thorough dispersal of genotypes at this scale. M. xanthus populations tend to be genetically distinct when separated by large distances, but what evolutionary mechanisms drive this nonrandom distribution? One possibility is that populations are adapting to a wide range of ecological habitats across broad spatial scales and that genetic differences between populations reflect such adaptation. Alternatively, populations might be limited by dispersal such that they diverge by randomly accumulating different sets of selectively neutral nucleotide changes (a process termed “genetic drift”). Baas Becking (1934) famously speculated that dispersal does not limit the distribution of microbes due to their immense numbers and small size, but rather local adaptation to heterogeneous environments determines what species and genotypes are found across different habitats. Although Whitaker et al. (2003) presented evidence that limited dispersal can indeed lead to genetic divergence between hot-spring thermophile populations, soil bacteria
2. ECOLOGY AND EVOLUTION OF MYXOBACTERIA
A
O0..?8 0.7
23
&
1
o = 5
A
rn I
5.5
6.5
6
7
7.5
a
distance (log km) Figure 2 F,, values between 10 different meter scale populations from around the globe plotted against distance between populations ( y 2 = 0.38, P > 0.05, n = 45). F,, values provide a measure of the degree of genetic diversity between populations relative to within populations, in this case based on the sequences of several highly conserved genes (Vos, 2006). Significantly differentiated population pairs are indicated by triangles, and nonsignificantly differentiated population pairs are indicated by squares.
that occupy large swaths of terrestrial soils should disperse more freely. Formation of stress-resistant spores should facilitate survival during dispersal events. Therefore, M. xanthus provides a conservative test of whether such broadly distributed soil bacteria can be limited by dispersal. As was found for several housekeeping genes (Fig. 2), genetic differentiation at the highly variable pilA gene among the 10 global populations of M. xanthus described above was found to increase significantly with distance. However, the average degree of differentiation between populations at pilA was found to be substantially lower than at the housekeeping genes (M. Vos, unpublished data). This result indicates that some piZA genes spread across populations via recombination and are maintained by selection in multiple locations. Nonetheless, both diversity and population differentiation at piZA still increase with distance, thus indicating that dispersal is limited. Furthermore, the increase of differentiation at pilA with spatial scale is caused as much by differences at synonymous sites as at nonsynonymous sites (Vos, unpublished data), as is expected if dispersal is limiting and populations are differentiating by genetic drift. The degree to which genetic differentiation of myxobacterial populations is caused by limited dispersal and subsequent genetic drift versus local adaptation (or some combination of these mechanisms) is a fundamental problem for future research seeking to understand the forces that drive myxobacterial biogeography.
Defining a Local Population The isolation studies described above provide a general picture of how species are distributed across diverse environments, but what does any particular local population of a myxobacterial species look like? How genetically similar are isolates that are close together in the same patch of soil? Which genes underlying social traits are under selection? These questions were addressed in a recent isolation study in which substantial genetic diversity was found among 78 isolates of M. xanthus within a 16- by 16-cm patch of soil in Tubingen, Germany, an area within which encounters among resident genotypes due to local migration are likely (Vos and Velicer, 2006). One hundred soil samples separated by 1.6 cm in a grid design yielded 78 successful clone isolations (one clone per sample for successful isolations), with isolates classified as M. xanthus based on morphology and 16s rRNA gene sequences. Fragments of the csgA, fibA, and pilA genes were sequenced for each, and concatemers of the three fragments revealed 21 distinct genotypes among the 78 clones, generating a prediction that a total of -26 genotypes would have been found in the plot with large increases in sampling effort. Thus, the genetic variation sampled largely reflected the total variation actually present at this spatial scale that was accessible from the utilized isolation protocol. Importantly, there was no statistically significant clustering of identical genotypes within the plot, indicating that the average clonal patch size in this location was smaller than 1.6 cm.
MYXOBACTERIAL BIOLOGY
24
A H o . 0010
/"
A4
99.2
\ 95'2
\
DK1622 '
97.4
A1 7 A5,h12
A75
\ 8e\o
A1
B -0.01
A12, A17 A53,A75
C
A66, A75
-0.1
A2,A3 A9, A53
57.i
A12
A25 A0 A98
The 21 detected genotypes formed six distinct phylogenetic branches, but the structure of the concatemer phylogeny was largely determined by variation in the pilA fragment due to its far greater diversity among isolates than csgA or fibA. Multiple tests provided strong evidence that diversifying natural selection is maintaining greater diversity in a highly variable region of the pilA gene than would be predicted by chance. In contrast, purifying natural selection appears to be purging more variation in the csgA gene than is expected by chance, indicating that differences in the CsgA signal are not responsible for developmental or motility incompatibilities observed among these isolates (see below). There was no evidence that any form of selection was acting on the sequenced portion of fibA. The degree of genetic exchange among individuals of the same species is a crucial determinant of how genomes evolve, and the evolutionary effects of sexual versus asexual modes of reproduction have been extensively modeled, investigated, and debated (Xu, 2004). Among microbes, the degree of genetic exchange (horizontal gene transfer [HGT])might range from levels approaching linkage equilibrium (when there is no statistical correlation between the presence of particular alleles across genes) to the complete absence of recombination among genomes. There have been no reports of spontaneous genetic exchange between M. xanthus cells under laboratory conditions, and tests for linkage disequilibrium among genes suggest that M. xanthus genome evolution is predominantly clonal (Vos and Velicer, 2006). Nonetheless, standard tests for recombination within genes provided evidence that recombination is likely to have occurred within six of nine genes examined among the lineages of 20 randomly chosen isolates. Moreover, comparison of the structure of the csgA, fibA, and pilA gene phylogenies provides additional evidence that HGT does occur in M. xanthus. The relative positions of several strains on the three gene trees were found to be radically incongruent (Fig. 3 ) , indicating transfer of alleles across evolving genome lineages rather than purely clonal Figure 3 Neighbor-joining trees of the csgA (A), fibA (B), and pilA (C) gene fragments. One or more clones were selected as representatives of each major clade in a csgA-fibApilA concatemer phylogeny generated for 78 local isolates from Tubingen, Germany. The laboratory strain DK1622 is included for comparison. Note the highly incongruent positions of strains A12, A17, and A75 across the three gene trees. The bootstrap value (1,000 replicates) is given at each node. Trees are not drawn to the same scale, and values in the upper left corner are genetic distances calculated with the Kimura two-parameter distance model. Reprinted with permission from Vos and Velicer (2006).
2. ECOLOGY AND EVOLUTION OF MYXOBACTERIA diversification, which would have resulted in congruent phylogenetic structures. Natural HGT in M. xanthus might occur by uptake of environmental DNA, ingestion of DNA from prey cells (which might include cannibalized cells of other myxobacteria), phage transduction, or via an undiscovered mode of conjugative transfer. Evidence suggesting HGT events from eukaryotes to myxobacteria has been reported (Porta and Rocha-Sosa, 2001; Quillet et al., 1995) but the frequency of such gene transfer across major taxonomic divisions or among strains of the same species remains unclear.
Ecological Determinants of Genotypic Performance Which myxobacterial phenotypes do we expect to find in particular environments? Answers to this question depend on how genotypes and environments interact to produce particular phenotypes and the relationship between those phenotypes and fitness. Although there have been significant advances in understanding the genetic mechanisms responsible for swarming and fruiting body morphogenesis, little systematic study of phenotypic variability in these traits across genotypes and environments has been conducted. Recently researchers have begun to compare phenotypic variability in swarming (Hillesland and Velicer, 2005; Shi and Zusman, 1993), fruiting body development at different population densities (Kadam and Velicer, 2006), and predatory ability of diverse genotypes (Bull et al., 2002; Pham et al., 2005) in the laboratory. The reproductive rate and survival of myxobacteria genotypes depend on their ability to survive in the absence of resources and on how quickly they can gain access to new ones. For myxobacteria living on surfaces, the latter parameter is tied to their rate of movement in search of food relative to other genotypes. In M. xanthus, swarming toward new food sources is accomplished by two genetically and physiologically distinct motility systems (A-motility and S-motility; see chapter 6 for details [Hodgkin and Kaiser, 1979a, 1979bl). Shi and Zusman (1993) showed that genotypes harboring either A-motility, S-motility, or both vary significantly in their swarming rate on different surfaces. Dually motile genotypes were able to produce large swarms on a range of surfaces, but on soft (0.3%) agar solely A-motile genotypes swarmed quite slowly while solely S-motile genotypes swarmed as fast as dually motile genotypes. On hard (1.5%) agar, solely A-motile genotypes swarmed much faster than solely S-motile genotypes. Further research by Hillesland and Velicer (2005) showed that the differential performance of A- and S-motility on hard versus soft surfaces is qualitatively
25
maintained across a wide range of nutrient concentrations. However, each motility system was found to respond differently to nutrient concentration, and this affected the relative performance of genotypes on the two surface types. The swarming rates of a dually motile genotype, a solely A-motile genotype, and a solely Smotile genotype were measured on hard and soft agar at varying nutrient concentrations. Genotypes with Amotility swarmed proficiently on hard agar across most nutrient concentrations, but S-motility swarming was almost undetectable at most low nutrient concentrations. Above 0.32% Casitone concentrations, the rate of swarming by S-motility on soft agar increased dramatically and was faster than A-motility swarming on hard agar. Figure 4 shows the relationships between the swarming rates of these genotypes across nutrient concentrations. At almost all nutrient concentrations, swarming by the solely A-motile genotype was faster than swarming by the solely S-motile genotype on hard agar (Fig. 4a). This ranking was reversed on soft agar, but the dominance of S-motility swarming was not significant except at high nutrient concentrations. At high nutrient concentrations, the dominance of S-motility swarming over A-motility swarming was very high (Fig. 4b). Finally, the genotype with both motility systems was superior to the solely A-motile genotype across nutrient concentrations on hard agar, indicating that S-motility enhances swarming by A-motility (Fig. 4c). However, A-motility does not seem to enhance S-motility swarming on soft agar (Fig. 4d). These results suggest that in a population containing multiple motility genotypes, solely A-motile genotypes would be at a disadvantage relative to genotypes harboring S-motility provided that nutrients were abundant and the swarming surface resembled soft agar. On hard agar, or at low nutrient concentrations, or both, solely S-motile genotypes should be at a significant disadvantage relative to genotypes that have A-motility. Another feature of 211. xanthus that may be important to fitness in nature and also exhibit variation across genotypes is the relationship between sporulation efficiency and population density. Recently, Kadam and Velicer (2006) documented considerable variation between nine genetically distinct natural isolates in their ability to form fruiting bodies and sporulate efficiently at low population densities. Several strains failed to form fruiting bodies or sporulate effectively at low densities, whereas other strains performed dramatically better at the same low densities (Fig. 5). All strains showed a threshold density below which sporulation efficiency rapidly decreased, but this threshold for one unusual strain was severalfoldlower than that of the strain with the second-lowest threshold. Why
MYXOBACTERIAL BIOLOGY
26
b
4 h
(u c
CJ 3 2 a, c
P
CO C -
-2
0
0.001 0.01
0.1
1
10
% casitone
0
0.001 0.01
0.1
1
10
% casitone
Figure 4 Relative A-motility, S-motility, and dual-motility swarming rates on hard and soft agar across a range of nutrient concentrations. Shown are the natural logs of the ratios of absolute swarming rates for solely A-motile versus solely S-motile genotypes on hard agar (a), solely S-motile versus solely A-motile genotypes on soft agar (b), dually motile versus solely A-motile genotypes on hard agar (c), and dually motile versus solely S-motile genotypes on soft agar (d). Shaded boxes indicate the half of the graph where data points should fall if the indicated genotype swarms comparatively faster than the alternative strain. Closed symbols indicate ratios that were significantly different from zero in a one-sample, one-tailed t test after sequential Bonferroni's correction for multiple comparisons ( P < 0.05 for eight comparisons in the same surface type; some comparisons not shown here). Error bars indicate bounds of the 95% confidence interval about the mean. Used with permission from Hillesland and Velicer (2005)
would there be phenotypic diversity in the ability to form fruiting bodies and spores at low population densities? It is possible that this variability is simply a by-product of variation across natural habitats on another, genetically linked phenotypic trait. However, it may be advantageous in some (but not all) habitats to form fruiting bodies and spores at low population densities. For example, populations evolving in typically nutrient-poor soils may tend to have lower population densities than populations living in habitats that fluctuate between high and low nutrient conditions. Thus, it may be advantageous to form fruiting bodies at low population densities in consistently poor nutrient environments.
Predation and Its Evolution Predators can affect the structure of entire food webs and the diversity of communities depending on whether they prey on competitively dominant or inferior species (Bohannan and Lenski, 2000; Lubchenco, 1978; Spiller and Schoener, 1998).Myxobacteria prey upon a diversity
of prokaryotic species, including cyanobacteria (Daft et al., 1985), various gram-positive organisms, Escherichiu coli, Rhizobiurn, and other gram-negative species (Beebe, 1941; Rosenberg and Varon, 1984). Many species of myxobacteria that have been tested are capable of utilizing multiple species as prey, although distinct species and isolates vary in the range of prey species they are capable of killing (Beebe, 1941; Bull et al., 2002; Rosenberg and Varon, 1984). A given population of myxobacteria may therefore significantly affect the density of a variety of organisms within a community and hence may be important players in the soil food web. To the extent that they prey upon organisms tied to animal or plant health, myxobacteria may also influence communities of macroorganisms. Bull and colleagues (2002) isolated several clones of Myxococcus spp. from organic and conventionally tilled strawberry fields. Assays of predatory capabilities of these myxobacteria indicated that they are capable of enhancing strawberry plant health by inhibiting the growth of fungal plant pathogens.
AND EVOLUTION OF MYXOBACTERIA 2. ECOLOGY
8.0
O
Log (initial cell density) 9.0 9.5
8.5
10.0
t
-1 h
$ -2
.-a, 0 E
;
-3 0 .c m 2 -4 0
Q v)
v
0 3 -5
9
J
-6 -7
I
Figure 5 Log-transformed sporulation efficiencies (spores produced/initial population size) of nine isolates of M. xanthus at five cell densities. Solid lines are used for isolates that sporulate efficiently at -3 x lo8cells/ml, whereas dashed lines indicate isolates that sporulate poorly at this density. Data points show the grand mean of all three replicate estimates, and error bars indicate 95% confidence intervals about the mean. Reprinted with permission from Kadam and Velicer (2006).
However, the Myxococcus spp. isolates were also capable of preying on fungi that may be used to control plant pathogens, so they could have negative effects on plant health as well. Pseudomonads, another potential biocontrol agent, were also susceptible to attack by Myxococcus spp. but could protect themselves by producing antibiotics. This study demonstrates the complexity of possible predator-prey interactions between myxobacteria and the diversity of soil microorganisms. It also shows that natural isolates of myxobacteria vary in their predatory ability. Different Myxococcus isolates exhibited different extents of predation on some species of fungi. Why would there be variation in predatory ability between different isolates from the same genus? To attack prey, myxobacteria must first search them out, a process that requires at least one functional motility system and might be enhanced by elasticotaxis (Fontes and Kaiser, 1999) and one or more chemotaxis systems (Kearns and Shimkets, 2001). Once prey have been found, myxobacteria use bacteriolytic enzymes, proteases, and possibly antibiotics (Reichenbach and Hofle, 1993; Rosenberg and Varon, 1984) to lyse them open and move across
27
the patch of prey and may further employ chemotaxis systems to ensure that they remain within the vicinity of prey populations. The genome contains multiple copies of a variety of proteases and chaperones that may be used in “digesting” prey (Goldman et al., 2006). Other genes, such as those involved in motility and development, may also affect predation. Pham et al. (2005) surveyed the effects of several mutations on the rate of lysis of lawns of prey on nitrocellulose membranes and streaks of prey on agar surfaces. They found that when Serratia marcescens lawns on nitrocellulose membranes were the only available resource for M . xanthus, functional A-motility was necessary for predation, but mutations affecting the S-motility system did not typically affect the rate of lysis. Mutations in the frz chemotaxis system that regulates cell reversal frequencies also negatively affected predation. The frz chemotactic system has also been implicated in predation by studies of single cells devouring microcolonies of E. coli (McBride and Zusman, 1996). In addition to these genes involved in swarm movement, prey lysis was reduced relative to the wild type when each of three genes (the sdeK, asgA, and csgA genes) necessary for early stages of fruiting body formation was mutated. The effects of these mutations on lysis varied significantly depending on the type of prey (Pham et al., 2005). These results show that predation involves many loci, and mutations in these loci may result in diverse predatory phenotypes on different prey species. Thus, significant genetic variation in predatory ability within species is likely, yet largely unexplored. Predatory rates by myxobacteria may also be significantly affected by features of the environment in which predation is taking place. This possibility has been examined for three variables: surface type, prey type, and the density of prey patches (Hillesland et al., 2007). In these experiments, patches were deposited onto a buffered agar surface in a grid configuration, M. xanthus was added to the center patch of the plate, and the plate was incubated for 2 weeks at 32°C (Fig. 6). During this incubation, M. xanthus swarms expanded radially outward across the agar surface in search of patches and then consumed them, as shown in Fig. 6c and d. All three variables influenced the percentage of total patches that was encountered by the swarm in this incubation period. A higher proportion of patches was encountered at high patch density than at low patch density, on hard agar than on soft agar, and on E. coli than on Micrococcus luteus (unless E. coli and M . luteus were on soft agar, in which case the ranking was reversed). Most of the prey within a patch were killed within a matter of a few hours if M. xanthus was equally distributed across the patch regardless of the surface type or which prey was used.
Next Page
28
Figure 6 Growth of M. xanthus on plates covered in patches of prey. Each plate consisted of buffered agar which was overlaid with thick patches of E. coli. A clone of M. xanthus was added to a central patch of E. coli, and photos were taken after 1 day of swarming at high (a) and low (b) patch density and again after 14 days of swarming at high (c) and low (d) patch density.
However, prey were more likely to be recovered from the patch after 24 h of incubation if the patches were distributed on soft agar, indicating that some environments offer greater protection to prey cells than others. Given the possibility of variation in predatory phenotypes due to both genetic and ecological causes, what predatory phenotypes might be expected to evolve in different environments? This question can be addressed through experimental evolution. The density of prey or prey patches affects many predators in the same manner that it affects M. xanthus (Holling, 1959). As shown experimentally above, more prey patches are consumed per unit time when they are densely distributed. At lower densities, prey patches are farther apart and the predator therefore has to spend relatively more time searching for prey at low densities than high densities. Thus, a reasonable hypothesis is that genotypes with faster search rates will have a competitive advantage at low patch densities. At high patch densities, the effect of searching rate on predation will be diminished and the competitive advantage of genotypes with higher searching rates will be less significant. This hypothesis was tested by allowing several replicate populations to evolve at high or low patch density for 2 1 0 0 generations
MYXOBACTERIAL BIOLOGY (Fig. 6). The searching rates of these evolved populations and the ancestor were then estimated by measuring the rate of swarm expansion on buffered agar in the absence of prey. All 16 populations evolved faster swarming on this surface, but as expected the increase in searching ability was much greater for the 8 populations that evolved at low density (-7-fold improvement) than it was for the 8 populations that evolved at high density (-2-fold improvement) (Hillesland, 2005). In another experiment, the fate of predatory ability after evolution in a prey-free environment was tested. Biologists have documented losses in traits that were unnecessary in a variety of species, including E . coli (Cooper et al., 2001; Velicer et al., 1998). Given the complex combination of genes potentially involved in predation and the likely metabolic cost of producing lytic enzymes, we tested whether predatory ability would be easily lost if it was not necessary for competitive success. The predatory abilities of eight populations that had evolved in batch culture in the absence of prey for 1,000 generations and eight populations that had evolved in the absence of prey on hard agar plates were measured in multiple environments along with their common ancestor. The populations that had evolved in the batch culture environment exhibited significant declines in their ability to encounter patches of prey in the grid environment shown in Fig. 6. They were also worse than the ancestor at killing prey in assay environments where searching for patches was not required (Hillesland, 2005; Velicer and Stredwick, 2002). This result suggests that predatory ability is readily lost during evolution in an environment where resources are abundant but prey are unavailable. However, most of the eight populations that had evolved on hard agar plates under selection for improved motility did not exhibit significant changes in predatory ability (Hillesland, 2005; Velicer and Stredwick, 2002).
SOCIOBIOLOGY OF THE MYXOBACTERIA Sophisticated cooperation is the hallmark of myxobacterial behavior, yet it can be argued that the precise evolutionary benefits of cooperation have not been clearly demonstrated for any myxobacterial trait, at least in natural habitats or relevant approximations thereof. While plausible speculation about such benefits has been appropriately abundant, the lack of relevant empirical data highlights the enduring difficultly of rigorous in situ study of microbial behavioral ecology. Beyond defining the precise manner in which cooperation benefits those organisms that engage in it, the evolutionary stability
Previous Page
2. ECOLOGYAND EVOLUTIONOF MYXOBACTERIA of cooperation remains one of the biggest problems in evolutionary biology. This is because cooperative behavior, by definition, benefits the evolutionary fitness of recipient individuals other than a cooperative actor and therefore can potentially be exploited by selfish cheaters. How can cooperation be stable if some individuals increase the representation of their genes in future generations by noncooperation (or “defection”) at the expense of cooperators? The combined challenges of (i) identifying traits that confer fitness benefits to organisms other than a cooperative actor; (ii) defining precisely what those benefits are and who receives them and in what degree; and (iii) understanding the evolutionary, behavioral, and molecular mechanisms that allow cooperative traits to be stably maintained are core themes in sociobiology. Although these challenges were first defined and explored in the context of metazoan cooperation, many microbiologists and evolutionary biologists have recognized that they apply with equal force to a wide variety of behavioral traits expressed by microbes, including the myxobacteria (Axelrod and Hamilton, 1981; Crespi, 2001; Keller and Surette, 2006; Parsek and Greenberg, 2005; Sachs et al., 2004; Smith and Szathmary, 1995; Velicer, 2003; West et al., 2006; Zahavi and Ralt, 1984).
Why Live in Groups or Cooperate in the First Place? Lions live in prides, migratory geese fly together, and newborn mice sleep in piles rather than alone. Ducklings swim in group formation, wolves, African wild dogs, and humans hunt in packs, and Hymenopteran insects construct complex breeding societies in which labor tasks are divided. The pervasiveness of group living across the biological spectrum indicates that organisms often enjoy greater reproductive success as members of social groups than they would with relatively individualistic life histories. However, the precise nature and quantitative degree of evolutionary benefits derived from group living remain unclear for many cooperative organisms, perhaps especially for microbes (Keller and Surette, 2006; Redfield, 2002; Travisano and Velicer, 2004; West et al., 2006). When nonmotile bacteria divide by fission in viscous habitats, physical proximity to colony mates in highdensity groups is unavoidable. Under such forced social proximity, natural selection will favor behavioral traits that succeed best when group life is the only option, and such traits are likely to involve cooperation between cells. However, cells with the capacity to “opt out” of sociality by actively migrating away from neighbors will maintain grouping behavior only if it is advantageous relative to a dispersal strategy (Shimkets, 1999). The
29
A-motility system of M. xanthus readily allows the movement of single cells (Hodgkin and Kaiser, 1979a, 1979b), yet the vast majority of M. xanthus natural isolates produce adhesions and slime tracks that chemically and physically hinder cells from migrating alone (Reichenbach, 1999).Why are such constraints on individualistic dispersal maintained? Here we revisit basic questions about why myxobacterial group behaviors (particularly as represented by M. xanthus) might be evolutionarily advantageous.
Communal Digestion Is there a benefit to myxobacterial grouping specific to food acquisition that requires extracellular catabolism? It has been proposed that individuals in high-density groups of myxobacteria enjoy a higher effective local concentration of extracellular hydrolytic enzymes during predation than single cells or low-density groups and therefore reproduce at a faster rate (Rosenberg et al., 1977). Rosenberg et al. (1977)demonstrated such a benefit of high density in the spatially extreme habitat of thoroughly shaken liquid medium. Cell density had no effect on growth rate when M. xanthus was provided with free amino acids and short peptides that can be transported directly into the cell. In contrast, populations fed with complex, nonhydrolyzed peptides that required extracellular catabolism prior to ingestion grew faster at high density than at low density, presumably due to a higher concentration of catabolic enzymes in the liquid. However, shaken liquid is a radical departure from the dining forum that terrestrial myxobacteria normally experience in the soil. In spatially structured soil habitats, enzyme diffusion should be very limited relative to shaken liquid and therefore density may have a much less pronounced effect on growth rate than in the Rosenberg et al. (1977) experiments. Although the Rosenberg et al. (1977) results are strongly suggestive, it has yet to be demonstrated that cooperative feeding provides a selective advantage in contexts more relevant to most natural myxobacterial habitats. Numerous myxobacteria are cellulolytic, and it will also be of interest to test whether population growth rate is density dependent when extracellular breakdown of cellulose is necessary for energy acquisition in spatially structured habitats. It has been intriguingly proposed that density-enhanced growth during predation may have been foundational to other forms of myxobacterial cooperation (Shimkets, 1990). Stress Support Groups? It has been reported that individual cells of some species are unable to grow in isolation, even in luxurious complex medium (Reichenbach, 1999). However, it is
30 unclear whether this phenomenon is due to actual synergistic interactions among cells or rather to high rates of cell death or inviability upon exposure to laboratory media. If the latter hypothesis is correct, inoculation populations of a minimum size would be necessary to statistically ensure obtaining one or more cells that proceed to grow. Even if synergism between cells is involved, the inability of single cells to grow is not likely to be an inherent trait of these genotypes across all environments. Rather, laboratory media may have parameters that are physiologically stressful for many strains relative to the low-nutrient soil habitats that myxobacteria normally inhabit, much the same way that moderate temperatures are stressful to genotypes adapted to cold habitats (Dawid et al., 1988).Group life may buffer against environmental stress even during growth. Addition of spent medium has been found to sometimes improve the growth of low-density populations, leading to speculation that a quorum-sensing mechanism may be in place to “prevent the futile growth of individual cells” (Reichenbach, 1999).However, it is difficult to imagine the ability of single cells to grow as being futile in an evolutionary sense, because successful population growth (from any starting density) is central to the definition of evolutionary success. Even if the competitive success of cells is greater at high densities during growth, growth from a single cell inherently moves a population toward the advantageous state of high cell density and therefore the retention of this ability should be favored by natural selection. It is therefore expected that the ability to grow as single cells is the norm in environments to which particular genotypes are well adapted. In contrast, some forms of physiological stress might be mitigated by cell-cell interactions within groups and therefore cause growth rate to positively correlate with population density when stress is high. Initial research indicates that pH stress causes the growth rate of M. xanthus strain DK1622 to be highly density dependent on complex, prehydrolyzed medium to a degree that is not observed in the absence of pH stress (H. Peitz and G. Velicer, unpublished results). Stress mitigation during vegetative growth may be a foundational benefit of group living in the myxobacteria. Group Motility Pilus-mediated S-motility in M. xanthus is costly to maintain when it is not important for fitness (Velicer et al., 1998; Velicer et al., 2002), so its maintenance in natural isolates indicates that it confers a significant overall fitness benefit. S-motility (in addition to A-motility) may confer several benefits that favor its evolutionary maintenance. First, studies of swarming by motility mutants on hard
MYXOBACTERIAL BIOLOGY agar surfaces suggest that the A- and S-motility systems are likely to be synergistic over a wide range of natural surfaces (Hillesland and Velicer, 2005; Shi and Zusman, 1993). Second, S-motility may facilitate the action of kin selection in M. xanthus (Hillesland and Velicer, 2005; West et al., 2006). The type IV pili that drive S-motility promote group cohesion and a kin-clustered population structure, thus ensuring that recipients of other forms of cooperative behavior (e.g., developmental signaling) are likely to be close relatives. Third, S-motility is necessary for tight packing of spores within fruiting bodies (Wu et al., 1998), which may itself provide several benefits to M. xanthus, discussed below. Finally, at least in the M. xanthus lab strain DK1622, the A-motility and S-motility systems appear to exhibit distinct swarming “specializations” that are specific to different surface types (hard agar for A-motility and soft agar for S-motility) (Hillesland and Velicer, 2005; Shi and Zusman, 1993). However, the faster swarming of S-motility on soft laboratory surfaces is observed only under abundant resource conditions, suggesting that such environmental specificity of performance may be only a minor component of the total fitness advantage conferred by the maintenance of S-motility. Some natural habitats may harbor ecologically successful strains of M. xanthus (or other species) that lack pilus-driven S-motility. Finally, individual prey and prey microcolonies may more frequently be “bumped into” by a group of several cells moving as a raft than they would by an isolated cell, which would cover a much smaller area. Sporulating Together Why do many myxobacteria sporulate in fruiting bodies rather than alone? A compelling answer to this question is still lacking. Laboratory mutants that sporulate effectively without normal fruiting body formation exist (Velicer et al., 1998), and some species of Nannocystineae do not make fruiting bodies at all (Shimkets et al., 2005). This raises the possibility that numerous myxobacteria in the wild may sporulate without fruiting body construction rather than bothering to pack themselves into a very tight spot with thousands of neighbors and a high probability of not differentiating into spores. Many methods of myxobacterial isolation are biased toward obtaining fruiting genotypes, and there may be many natural nonfruiting strains of species known for their fruiting ability in the laboratory (Velicer et al., 2002; Jiang et al., 2007). An enhanced probability of dispersal to a new, foodrich habitat is a commonly proposed benefit of group sporulation (Kaiser, 2001). Although plausible for some species that make tall fruiting bodies or exhibit
AND EVOLUTION OF MYXOBACTERIA 2. ECOLOGY
light-induced development (White et al., 1980),the status of this hypothesis remains uncertain for any myxobacterial species. The lack of prominent fruiting body stalks or chemotaxis toward warmth and light in some species gives pause when considering the dispersal hypothesis. There are several additional (although nonexclusive) potential benefits to group fruiting as well. Fruiting body sporulation facilitates germination and growth in highdensity groups, which may begin sooner and occur at a faster rate, respectively, than for isolated cells or lowdensity groups (Kaiser, 2001; Rosenberg et al., 1977). There may be some aspect of differentiation within fruiting bodies (e.g., autolytic release of compounds beneficial to spores [Wireman and Dworkin, 19771) that generates individual spores of higher quality than can be produced by asocial or low-density sporulation, thus resulting in greater longevity under stress. Further, the extracellular matrix surrounding spores in simple fruiting bodies (such as those of M. xanthus) and the spore-bearing sporangioles of the suborder Sorangiineae may provide a degree of protection against environmental stresses that would not be available to relatively isolated spores. Such stresses include starvation duration, temperature and pH extremes, caustic biotic (e.g., antibiotics produced by competitors) and abiotic molecules, and ingestion or digestion by would-be predators. The ultimate evolutionary causes and benefits of myxobacterial cooperation remain open to much exploration.
The Physiological Cost of Cooperation All behaviors, whether cooperative or not, are produced at a short-term physiological cost and can be maintained by natural selection only when they yield a net gain in evolutionary fitness across generational time scales. Some cooperative microbial behaviors, such as intercellular signal production, may generate a fitness benefit that exceeds the cost of production only in a group context (Travisano and Velicer, 2004), such that natural selection will favor nonexpression or loss during evolution under conditions where social interaction is limited. Rapid loss of fruiting behavior in myxobacteria and complex multicellular phenotypes in other bacteria during laboratory cultivation is a widespread observation among microbiologists (Branda et al., 2001; Dawid, 2000; Sproer et al., 1999; Velicer et al., 1998).Such losses during domestication may often cause large fitness gains under luxurious laboratory conditions due to reallocation of metabolic resources away from unnecessary functions. The evolutionary fate of unnecessary social functions was examined systematically by allowing 12 independent lineages of M. xanthus all derived from a
31
single clone of strain DIC1622 to evolve for 1,000 generations in nutrient-rich liquid medium (Velicer et al., 1998).This selective regime did not favor proficiency at motility, predation, or development. The evolved populations showed significant adaptation to this “asocial” regime by achieving an average growth rate increase of -28 %. However, this adaptation was associated with major losses in social capacities, with most populations showing complete loss of S-motility and/or fruiting body development. The rapidity (many losses occurred between 200 and 500 generations [our unpublished data]) and convergence of losses across populations indicated that the mutations responsible for inactivation of social capacities rose to high frequency via natural selection rather than chance. In a subsequent study, mutations that caused the loss of S-motility in several lines were localized to the pi1 genes responsible for the generation and function of type IV pili necessary for S-motility (Velicer et al., 2002). It was shown that genetic inactivation of essential pi1 genes in the ancestral strain conferred a fitness advantage to the mutants during growth in the liquid evolution regime. Reciprocally, partial restoration of S-motility in deficient evolved genotypes was detrimental to fitness in liquid medium. These results demonstrated the advantage of eliminating social capacities that are costly to maintain when they do not enhance fitness.
Genetic Conflict over Public Goods Microbial public goods, such as intercellular developmental signals in the myxobacteria, are made accessible to other cells at an immediate cost to the producer. To be maintained in populations over evolutionary time, therefore, the cost of public-good production must be outweighed by direct or indirect fitness benefits, or both, for the producing genotype. Enter cheaters, the perennial bane of social harmony and productivity. Public-good contributions by cooperative actors need not be reciprocated by all recipients. Because such contributions are costly, failure by an individual to proportionately contribute to a public good (“defection”)may be advantageous for fitness in the short run, In this case, natural selection favors selfish cheaters rather than cooperative contributors. If mutational pathways (e.g., in social microbes) or behavioral plasticity (e.g., in social metazoans) readily allows the appearance of cheating behavioral strategies in the presence of cooperators, evolutionary game theory predicts that such cheaters should be a persistent feature in the long-term evolution of cooperative biological systems. When cheating can appear, it will spread unless there exist mechanisms to stop it (Travisano and Velicer, 2004; West et al., 2006).
MYXOBACTERIAL BIOLOGY
32 Various forms of cheating are common in social animals and insects. For example, entire species of social ants and wasps are obligate social parasites in which the queens do not generate their own worker offspring but rather usurp colonies of their social hosts and utilize host workers to raise their own reproductive brood (Bourke and Franks, 1995; Lorenzi et al., 2004). In the myxobacteria, social cheating might occur during social swarming, group predation, or multicellular development, and in fact developmental cheaters can be readily obtained. Velicer et al. (Velicer et al., 2000; Velicer and Stredwick, 2002) examined several genotypes derived from DK1622 that are defective at spore production to varying degrees in pure culture. Some of these defectors were defined mutants defective at A- or C-signal production, whereas others carried undefined defect mutations that had accumulated during evolution in liquid culture (Velicer et al., 1998). Strikingly, more than one-half of the defective genotypes sporulated more efficiently than the fully proficient strain DK1622 when mixed as a small minority (1%) with DK1622, despite their defects in pure culture (Fig. 7). These cheaters included two defined mutants that presumably differ from DK1622 by only their mutations in either asgB or csgA. This result demonstrates the ease of crossing the thin evolutionary line between cooperation and cheating by simple mutational pathways. The full range of M. xanthus genes in which single mutations can cause cheating has yet to be determined. Theory predicts the short-term evolutionary success of cheaters whenever there exists a cooperatively produced public good and accessible mutational or neurological pathways to cheating behavior. Consistent with this expectation, cheating has been documented in several microbial public-good scenarios other than that of M. xanthus development (Greig and Travisano, 2004; Griffin et al., 2004; MacLean and Gudelj, 2006; Rainey and Rainey, 2003; Turner and Chao, 1999; Vulic and Kolter, 2001). Most published examples of microbial social exploitation share the common theme of defectors gaining an advantage by failing to contribute to a group-generated public good in a genetically determined manner, a mode of exploitation that has been termed “obligate cheating.” Obligate cheaters are inherently poor performers under clonal conditions in which the exploited public good is important for fitness. Numerous other microbial public-good scenarios may also be subject to cheating, including quorum sensing and biofilm formation in many species and group predation and social motility in the myxobacteria. Obligate cheating constitutes a potentially enormous problem for microbial cooperation, and mechanisms
-3
-2
-1 Log,, (initial mixing ratio)
0
1
Figure 7 Spore production of an evolved cheater genotype when mixed with its wild-type progenitor (DK1622)at nine initial ratios (a) and the corresponding relative sporulation efficiencies (b). The cheater produces no detectable spores during development in pure culture (data not shown). (a) Squares, triangles, and circles indicate total, DK1622, and cheater spore production, respectively. The expected production of the evolved clones under the hypothesis that DK1622 does not improve the defective strain’s sporulation efficiency ( H l ) is represented by the solid line. The expected production of evolved clones under the hypothesis that the defective strain is rescued to the same efficiency as DK1622 (H2)is represented by the dotted line. The spore production of DK1622 in independent pure cultures is represented by the dashed line. Error bars indicate 95% confidence intervals. (b) Sporulation efficiency of the cheater relative to that of DK1622 for these same initial mixing ratios. The dashed line indicates a relative efficiency of 1. Reprinted with permission from Velicer et al. (2006).Copyright (2006)National Academy of Sciences, United States.
therefore must exist to limit its appearance or effects (Travisano and Velicer, 2004; West et al., 2006). Such mechanisms may include negative pleiotropic effects of defector mutations, targeting of benefits to cooperators, targeted punishment of cheaters, and group-level selection in which individuals in groups without cheaters outperform individuals in distinct groups burdened by heavy cheating loads.
2. ECOLOGY AND EVOLUTION OF MYXOBACTERIA An alternative mode of microbial exploitation does not involve an inherent social defect in the exploiter. In this case, a genotype that is fully proficient by itself at the exploited social task is able to perform that task more efficiently in the presence of a particular competitor (and to that competitor’s expense) than in clonal isolation. Such facultative exploitation has been documented in natural isolates of the eukaryotic slime mold Dictyostelium discoideum and of M. xanthus during painvise development competition mixes with other natural isolates of the same respective species (Fiegna and Velicer, 2005; Strassmann et al., 2000). That obligate cheating can be so readily generated in the lab and that facultative exploitation is so easily detected among natural isolates suggest that both forms of exploitation may be common in natural microbial populations.
The Group Perils of Cheating Obligate cheaters that make no spores in isolation can invade cooperative populations when cheats are rare, but they inherently diminish group productivity when they reach high frequencies. When the exploited cooperative trait is important for survival, obligate cheaters may drive populations to outright extinction. To examine the competitive fates and population-level effects of obligate developmental cheaters, three distinct cheater genotypes were allowed to compete against marked variants of DK1622 through several successive rounds of starvation and growth (Fiegna and Velicer, 2003). Although all three cheaters fail to make any spores in isolation, they had very different effects on population dynamics. One cheater with a defect in CsgA production was maintained at high levels in the population without causing major population crashes. A second cheater was also maintained but caused large decreases in spore production upon reaching high frequencies. A third and particularly virulent cheater caused outright extinction events due to the short-term success of its own “selfish” behavior during development. In some replicate competitions with the latter cheater the entire population went extinct due to cheater-induced sporulation crashes, whereas in other replicates only the cheater (or neither competitor) was driven to extinction. Such cheater-induced extinctions demonstrate the concept of evolutionary suicide, in which selfish behaviors that improve the fitness of some genotypes in the short run are disastrous for populations or species in the long run (Rankin and Lopez-Sepulcre, 2005). The observed variation in cheater effects at the population level is presumably due to differences in cheating intensity and the frequency thresholds at which the cheaters lose their advantage and cause large drops in spore production. Cheaters with high short-term fitness
33
in nature are expected to hinder the success of others in the groups that they exploit.
Evolutionary Escape from Social Defects Re-Evolution of Development Myxobacteria can rapidly lose their social functions during evolution under favorable growth conditions, suggesting that socially defective strains may be common in nature. This raises the question of whether such social degeneration is an evolutionary dead end or whether accessible mutational pathways exist that can allow an evolutionary escape from a defect under conditions in which restored cooperation would be advantageous. Two recent studies demonstrate that M. xanthus has a remarkable capacity to reevolve social functions from a socially defective genomic background and does so through novel genetic pathways relative to the original genomic state prior to mutational deterioration of social functions. First, the extinction-inducing cheater described above (hereafter abbreviated OC for “obligate cheater”) did not always cause total self-destruction. In one replicate population, OC did drive the population to the brink of extinction, but a spontaneous mutant of OC not only survived the population crash but did so by reevolving developmental proficiency (Fiegna et al., 2006). The newly evolved genotype (PX for “Phoenix”) sporulated even more efficiently than a marked variant of the proficient ancestor of OC (GJV2) in pure culture and showed higher developmental fitness than both GJV2 and OC in mixed populations over a wide range of mixing frequencies. The appearance of PX represents not only an evolutionary escape from a state of obligate social cheating but also the emergence of a competitively superior cooperative strategy and the spontaneous evolution of resistance to cheating. In this transition, obligate cheating served as a stepping-stone to a novel and superior form of cooperation rather than being an evolutionarily terminal fate. Using two sequencing technologies, one new (“sequencing-by-synthesis”) and one traditional ( “Sanger” sequencing), the entire PX genome was sequenced in search of the mutational basis of the evolutionary transitions to OC and PX (Velicer et al., 2006). The resulting data are likely to have revealed all mutational differences between PX, OC, and GJVl. Such comprehensive mutation identification is a crucial beginning to the complex task of understanding the genetic basis of evolutionary change and adaptation in laboratory evolutionary experiments. While 15 mutational differences between PX and GJVl were discovered, only one was unique to PX. The others were all present in OC and
MYXOBACTERIAL BIOLOGY
34 distinguish it from its cooperative ancestor (Fig. 8). Strikingly, PX was shown to have arisen from just this single regulatory mutation. This mutation occurred upstream of an uncharacterized GNAT-acetyltransferase enzyme. It increases the production of this enzyme early in development and causes many other changes in PX gene expression that appear to drive a novel mechanistic route to social development in 111. xanthus (Fiegna et al., 2006; Kadam, 2006). That a social trait can be restored by evolutionary conversion of a cheater into a cooperator expands our view of how social systems can evolve. That a new genetic basis for social cooperation can arise with “builtin” resistance to cheating increases our understanding of how the effects of cheaters on microbial social systems might be limited over evolutionary time scales. That such a dramatic behavioral restoration can be accomplished
D K I622
5 mutations
I
Tn5
? gen.
by a single spontaneous mutation highlights how much we have yet to learn about existing and potential genetic networks that underlie social traits in M. xanthus and other social organisms.
Reevolution of Social Swarming Another study shows that the ability of 211. xanthus to reevolve lost social functions extends to group motility as well as development (Velicer and Yu, 2003). In this case, S-motility was intentionally eliminated by deletion of a large portion of the pilA gene that encodes the pilin protein used to construct type IV pili necessary for S-motility. Multiple populations derived from the ApilA ancestors were allowed to undergo multiple 2-week cycles of growth on a soft agar surface on which S-motility is required to drive effective swarming in the lab strain DK1622. Thus, the ApilA populations initially
\ “‘+ioOs
PX
14 mutations
GJVl
b
GVB207.3
1000 generations
b
growth in rich liquid medium
b
four cycles of alternating starvation and growth
Figure 8 Mutational history of the PX mutant. The previously sequenced strain DK1622 and its derivative clone GJVl are separated by five mutations and an unknown number of generations of lab stock cultivation. The lineage from GJVl to GVB207.3 incurred 1 4 mutations over 1,000 generations of growth in liquid medium (Velicer et al., 1998), one or more of which eliminated the ability to undergo multicellular development. OC was generated by integration of a Tn5 transposon (which confers resistance to kanamycin) into the GVB207.3 genome. OC evolved into PX by regaining the ability to sporulate via social development during an extended developmental competition against a marked variant of GJV1. Only one mutation was found to distinguish PX from OC, and this mutation was subsequently shown to cause the restoration of developmental proficiency in PX. Reprinted with permission from Fiegna et al. (2006).
2. ECOLOGYAND EVOLUTION OF MYXOBACTERIA expanded very little during the early transfer cycles. At the end of each cycle, however, cells from the point on the population perimeter furthest from the initial inoculation point were transferred to a fresh plate, thus favoring any mutants capable of swarming outward more effectively than the ancestral defective population. After extended evolution, six populations showed significant but relatively small increases in soft-agar swarming rates relative to their ApiZA ancestor, whereas two other populations gained much larger increases (Fig. 9). The evolved mutants showing the largest gains in swarming performance did not regain soft-agar swarming by restoring their ancestral capacity for pilin-mediated S-motility. Rather, they compensated by evolving a new mechanism to drive swarming on soft agar that does not involve pili (Velicerand Yu, 2003). The evolved swarmers were shown to lack pilin just like their ancestors, but they had reevolved the ability to produce cohesive extracellular fibril material that had been eliminated by the ApilA deletion. This fibril material and the intact A-motility system were both necessary for the reevolved swarming phenotypes. Thus, soft-agar swarming was restored by
35
the evolution of a novel (and as yet undefined) relationship between the M. xanthus extracellular matrix and the A-motility system. Moreover, it was shown that social interactions between cells that are mediated by fibrils, and not mere fibril production per se, were involved in generating the evolved swarming phenotypes. Therefore, M . xanthus reevolved a socially mediated behavior (swarming on soft agar) but found a novel molecular mechanism for how to drive this behavior.
Social Divergence in Natural Populations A fundamental question in myxobacterial sociobiology is the following: Who cooperates with whom? Selfpropelled motility and external migration vectors should frequently lead to encounters between distinct species and strains of myxobacteria. In such cases of external encounter of different colony types, how distinct can two genotypes be at various genetic loci and still swarm, aggregate, or sporulate together? Suboptimal cooperators (or even outright antagonists) might also arise within groups, since every bacterial colony above a certain population size contains some degree of genetic
Figure 9 Swarming phenotypes of ancestral ApilA genotypes (third and fifth positions clockwise from top) and their evolved descendants (second and fourth positions clockwise from top, respectively) relative to DK1622 (top position) on soft agar. The DK1622 swarm was 3 days old, and the ApilA, and evolved strain swarms were 7 days old. Reprinted with permission from Velicer and Yu (2003).
MYXOBACTERIAL BIOLOGY
36 diversity. For example, the normal spontaneous mutation rate of M. xanthus may be similar to that of E. coZi (-1 to 5 x 10-lo per base pair per generation [Lenski et al., 2003; Velicer et al., 2006]), such that one in every several hundred cell divisions may result in a new genetic variant within colonies of M. xanthus. Because single spontaneous mutations can dramatically affect cell surface molecules, intercellular signal production, or cell behavior (Fiegna et al., 2006; Velicer et al., 2006), the appearance of genetically distinct neighbors by simple mutational steps that reduce compatibility to the majority genotype may occur frequently. In the first study to examine developmental compatibility across myxobacterial genotypes, Smith and Dworkin (1994) mixed two strains classified as Myxococcus virescens and M. xanthus at the onset of starvation. These two genotypes appear to have fully segregated into species-specific fruiting bodies, and the presence of the M. virescens strain strongly inhibited spore production by the M . xanthus isolate. Subsequently, Fiegna and Velicer (2005) quantified developmental compatibility across all possible pairwise combinations of nine M. xanthus strains isolated from distant global locations. Most pairings resulted in greatly reduced sporulation for at least one competitor and reduced total spore production (relative to expectations from pure-culture controls). Strain fitness relationships were predominately hierarchical rather than circular (i.e., nontransitive), with three strains dominating over six inferior strains, which in turn exhibited a linear rank hierarchy. Such competitive asymmetries are likely to be due to differences in one or more components of cell surface or diffusible “secretomes” across competitors. In several instances, superior competitors actually sporulated more efficiently in the presence of their inferior partner than they did as clonal cultures, thus demonstrating the possibility of social exploitation among socially proficient myxobacteria in the wild (Fig. 10). That isolates from across the globe have inadvertently evolved social incompatibility in isolation from one another is perhaps not surprising. But what happens when closely related neighbors encounter one another? In a study parallel in design to that of Fiegna and Velicer (2005), pervasive developmental antagonism has been observed among nine isolates from the 16- by 16-cm Tubingen sample plot (see above), including between isolates that have identical csgA, fibA, and pilA alleles (Vos, 2006). Moreover, a swarm compatibility test was conducted for all possible pairs of distinct isolates from the plot that share an identical csgA-fi6A-piZA concatemer sequence. In most cases, the paired swarms (inoculated near one another on an agar plate) failed
D
”j -7
I
I
G
1
Figure 10. Facultative and antagonistic exploitation by two natural isolates of M . xanthus (D against I and I against G) during mixed development. The log-scale effect of mixing strains i and j on the sporulation efficiency of strain i is termed Ci(j). Open bars show the effect of mixing on sporulation efficiency for the dominant, exploitative competitor in each pair (D and I, respectively) in response to its inferior competitor. Shaded bars indicate the effect of mixing on the inferior strain (I and G, respectively). Error bars indicate 95 % confidence intervals. Reprinted with permission from Fiegna and Velicer (2005).
to merge, whereas control pairs of two swarms of the same isolate always merged (Vos, 2006). These results indicate that divergence into incompatible social genotypes occurs at a very fine scale of genomic divergence that is not reflected by variation in the highly variable piZA gene. Extrapolating these results across whatever portion of the -150 million km2 of terrestrial earth surface that M. xanthus has colonized (which is certainly a substantial fraction), it appears that M. xanthus has diverged into many trillions of distinct social genotypes that are in most cases socially incompatible with one another to some degree. Distinct species of myxobacteria can often be isolated from the same soil sample (Dawid, 2000), prompting the question of how such coexisting species might interact. N. Dahanukar performed a study of swarm interactions between seven distinct myxobacterial isolates that appeared to represent both classified species ( M . xanthus, M . fulvus, and StigmateZla aurantiaca) and previously unclassified species. The results suggested that some species strongly dominate over others in pairwise swarm encounters on agar plates, even to the point of invading competitor territory and decimating competitor populations (Dahanukar, 2003). Intriguingly, the average degree of dominance over competitors during swarm mixes appeared to positively correlate with
2.
ECOLOGY AND EVOLUTION OF MYXOBACTERIA
the stalk length that each isolate exhibited during development. Further research is needed to determine whether there are general patterns of dominance across species, or whether intraspecific social diversification is so great that competitive dominance between any two random isolates cannot be predicted by species classification.
CONCLUSION Laboratory-based approaches to the ecology and evolution of myxobacteria have begun to open up a new range of largely unexplored questions in this field. Moreover, we stand on the verge of great increases in our power to ask fundamental questions about the distribution, ecology, and evolution of myxobacteria in their native habitats by using current and future molecular tools. These research trends promise to tell us much more not only about the myxobacteria themselves but also about broader biological issues such as the evolution of group living, cooperation, conflict, and multicellularity.
References Amann, R. I., W. Ludwig, and K. H. Schleifer. 1995. Phylogenetic identification and in-situ detection of individual microbial cells without cultivation. Microbiol. Rev. 59:143-169. Axelrod, R., and W. D. Hamilton. 1981. The evolution of cooperation. Science 211:1390-1396. Baas Becking, L. G. M. 1934. Geobiologie: of inleiding tot de milieukunde. W. P. Van Stockum & Zoon N. V., The Hague, The Netherlands. Beebe, J. M. 1941. Studies on the Myxobacteria. Iowa St. Coll. J. Sci. 15:307-337. Bohannan, B. J. M., and R. E. Lenski. 2000. Linking genetic change to community evolution: insights from studies of bacteria and bacteriophage. Ecol. Lett. 3:362-377. Bourke, A. F. G., and N. R. Franks. 1995. Social Evolution in Ants. Princeton University Press, Princeton, NJ. Branda, S. S., J. E. Gonzalez-Pastor, S. Ben-Yehuda, R. Losick, and R. Kolter. 2001. Fruiting body formation by Bacillus subtilis. Proc. Natl. Acad. Sci. USA 98:11621-11626. Bull, C. T., K. G. Shetty, and K. V. Subbarao. 2002. Interactions between Myxobacteria, plant pathogenic fungi, and biocontrol agents. Plant Dis. 869894396. Cooper, V. S., D. Schneider, M. Blot, and R. E. Lenski. 2001. Mechanisms causing rapid and parallel losses of ribose catabolism in evolving populations of Escherichia coli. B. J. Bucteriol. 183:2834-2841. Crespi, B. J. 2001. The evolution of social behavior in microorganisms. Trends Ecol. Evol. 16:178-183. Daft, M. J., J. C. Burnham, and Y. Yamamoto. 1985. Lysis of Phormidium luridum by Myxococcus fulvus in continuous flow cultures. J. Appl. Bacteriol. 59:73-80. Dahanukar, N. 2003. Species, morphological and behavioral diversity in myxobacteria. Abasaheb Garware College, Pune, India.
37
Dawid, W. 2000. Biology and global distribution of Myxobacteria in soils. FEMS Microbiol. Rev. 24:403-427. Dawid, W., C. A. Gallikowski, and P. Hirsch. 1988. Psychrophilic myxobacteria from antarctic soils. Polarforschung 58~217-278. Elena, S. F. 2002. Restrictions to RNA virus adaptation: an experimental approach. Antonie Leeuwenhoek 81:135-142. Elena, S. F., and R. E. Lenski. 2003. Evolution experiments with microorganisms: the dynamics and genetic bases of adaptation. Nut. Rev. Genet. 4:457-469. Endler, J. A. 1986. Natural Selection in the Wild. Princeton University Press, Princeton, NJ. Feldgarden, M., D. M. Stoebel, D. Brisson, and D. E. Dykhuizen. 2003. Size doesn’t matter: microbial selection experiments address ecological phenomena. Ecology 84:1679-1687. Fenchel, T. 2003. Biogeography for bacteria. Science 301:925926. Fiegna, F., and G. J. Velicer. 2003. Competitive fates of bacterial social parasites: persistence and self-induced extinction of Myxococcus xanthus cheaters. Proc. R. SOC. London B 270~1527-1534. Fiegna, F., and G. J. Velicer. 2005. Exploitative and hierarchical antagonism in a cooperative bacterium. PLoS Biol. 3 :1980-1 987. Fiegna, F., Y. T. N. Yu, S. V. Kadam, and G. J. Velicer. 2006. Evolution of an obligate social cheater to a superior cooperator. Nature 441:310-314. Fontes, M., and D. Kaiser. 1999. Myxococcus cells respond to elastic forces in their substrate. Proc. Nut. Acad. Sci. USA 96:8052-8057. Gerth, K., and R. Muller. 2005. Moderately thermophilic Myxobacteria: novel potential for the production of natural products isolation and characterization. Environ. Microbiol. 72374-8 80. Gevers, D., F. M. Cohan, J. G. Lawrence, B. G. Spratt, T. Coenye, E. J. Fed, E. Stackebrandt, Y. Van de Peer, P. Vandamme, F. L. Thompson, and J. Swings. 2005. Re-evaluating prokaryotic species. Nut. Rev. Microbiol. 3:733-739. Goldman, B., W. Nierman, D. Kaiser, S. Slater, A. Durkin, J. Eisen, C. Ronning, W. Barbazuk, M. Blanchard, C. Field, C. Halling, G. Hinkle, 0. Lartchuk, H. Kim, C. Mackenzie, R. Madupu, N. Miller, A. Shvartsbeyn, S. Sullivan, M. Vaudin, R. Wiegand, and H. Kaplan. 2006. Evolution of sensory complexity recorded in a myxobacterial genome. Proc. Nutl. Acad. Sci. USA 103:15200-15205. Grant, P. R., and B. R. Grant. 2002. Unpredictable evolution in a 30-year study of Darwin’s finches. Science 296:707-711. Greig, D., and M. Travisano. 2004. The Prisoner’s Dilemma and polymorphism in yeast SUC genes. Proc. R. Soc. London 271:S25-S26. Griffin, A. S., S. A. West, and A. Buckling. 2004. Cooperation and competition in pathogenic bacteria. Nature 430:10241027. Hanski, I., and I. Saccheri. 25 April 2006. Molecular-level variation affects population growth in a butterfly metapopulation. PLoS Biol. 4:e129. [Epub ahead of print.] He, Q., and R. A. Sanford. 2003. Characterization of Fe(II1) reduction by chlororespiring Anaeromxyobacter dehalogenuns. Appl. Environ. Microbiol. 69:2712-2718.
38 Hillesland, K. L. 2005. Evolutionary ecology of predation by the soil bacterium, Myxococcus xanthus. Ph.D. thesis. Michigan State University, East Lansing. Hillesland, K. L., R. E. Lenski, and G. J. Velicer. 2007. Ecological variables affecting predatory success in Myxococcus xanthus. Microb. Ecol. 53571-578. Hillesland, K. L., and G. J. Velicer. 2005. Resource level affects relative performance of the two motility systems of Myxococcus xanthus. Microb. Ecol. 49558-566. Hodgkin, J., and D. Kaiser. 1979a. Genetics of gliding motility in Myxococcus xanthus (Myxobacterales): two gene systems control movement. Mol. Gen. Genet. 171:177-191. Hodgkin, J., and D. Kaiser. 1979b. Genetics of gliding motility in Myxococcus xanthus (Myxobacterales):genes controlling movement of single cells. Mol. Gen. Genet. 171:167-176. Holling, C. S. 1959. Some characteristics of simple types of predation and parasitism. Can. Entomol. 91:385-398. Iizuka, T., Y. Jojima, R. Fudou, M. Tokura, A. Hiraishi, and S. Yamanaka. 2003. Enhygromyxa salina gen. nov., sp nov., a slightly halophilic myxobacterium isolated from the coastal areas of Japan. Syst. Appl. Microbiol. 26:189-196. Iizuka, T., Y. Jojima, R. Fudou, and S. Yamanaka. 1998. Isolation of myxobacteria from the marine environment. FEMS Microbiol. Lett. 169:317-322. Jessup, C. M., R. Kassen, S. E. Forde, B. Kerr, A. Buckling, P. B. Rainey, and B. J. M. Bohannan. 2004. Big questions, small worlds: microbial model systems in ecology. Trends Ecol. Evol. 19~189-197. Jiang, D.-M., Z.-H. Wu, J.-Y. Zhao, and Y.-Z. Li. 2007. Fruiting and non-fruiting myxobacteria: a phylogenetic perspective of cultured and uncultured members of this group. Mol. Phylogenet. Evol. 44545-552. Kadam, S. 2006. Ecology and evolution of multicellular development in the social bacterium Myxococcus xanthus. Doctoral thesis, Universitat Tubingen, Tubingen, Germany. Kadam, S. V., and G. J. Velicer. 2006. Variable patterns of density-dependent survival in social bacteria. Behav. Ecol. 17:833-838. Kaiser, D. 2001. Building a multicellular organism. Annu. Rev. Genet. 35:103-123. Kassen, R., and P. B. Rainey. 2004. The ecology and genetics of microbial diversity. Annu. Rev. Microbiol. 58:207-231. Kearns, D.B., and L. J. Shimkets. 2001. Lipid chemotaxis and signal transduction in Myxococcus xanthus. Trends Microbiol. 9:126-129. Keller, L., and M. G. Surette. 2006. Communication in bacteria: an ecological and evolutionary perspective. Nut. Rev. Microbiol. 4:249-258. Lenski, R. E., M. R. Rose, S. C. Simpson, and S. C. Tadler. 1991. Long-term experimental evolution in Escherichia coli. I. Adaptation and divergence during 2,000 generations. Am. Nut. 138~1315-1341. Lenski, R. E., C. L. Winkworth, and M. A. Riley. 2003. Rates of DNA sequence evolution in experimental populations of Escherichia coli during 20,000 generations. J. Mol. Evol. 56~498-508. Linz, B., F. Balloux, Y. Moodley, A. Manica, H. Liu, P. Roumagnac, D. Falush, C. Stamer, F. Prugnolle, S. W. van der Menve, Y. Yamaoka, D. Y. Graham, E. Perez-Trallero, T.
MYXOBACTERIAL BIOLOGY Wadstrom, S. Suerbaum, and M. Achtman. 2007. An African origin for the intimate association between humans and Helicobacter pylori. Nature 445:915-918. Lorenzi, M. C., R. Cervo, F. Zacchi, S. Turillazzi, and A. G. Bagneres. 2004. Dynamics of chemical mimicry in the social parasite wasp Polistes semenowi (Hymenoptera: Vespidae). Parasitology 129:643-651. Lubchenco, J. 1978. Plant species diversity in a marine intertidal community: importance of herbivore food preference and algal competitive abilities. Am. Nut. 112:23-39. MacLean, R. C.,and I. Gudelj. 2006. Resource competition in bacteria: an ecological and evolutionary perspective. Nut. Rev. Microbiol. 4:249-258. Margulies, M., M. Egholm, W. E. Altman, S. Attiya, J. S. Bader, L. A. Bemben, J. Berka, M. S. Braverman, Y. J. Chen, Z. Chen, S. B. Dewell, L. Du, J. M. Fierro, X. V. Gornes, B. C. Godwin, W. He, S. Helgesen, C. H. Ho, G. P. Irzyk, S. C. Jando, M. L. Alenquer, T. P. Jarvie, K. B. Jirage, J. B. Kim, J. R. Knight, J. R. Lanza, J. H. Leamon, S. M. Lefkowitz, M. Lei, J. Li, K. L. Lohman, H. Lu, V. B. Makhijani, K. E. McDade, M. P. McKenna, E. W. Myers, E. Nickerson, J. R. Nobile, R. Plant, B. P. PUC,M. T. Ronan, G. T. Roth, G. J. Sarkis, J. F. Simons, J. W. Simpson, M. Srinivasan, K. R. Tartaro, A. Tomasz, K. A. Vogt, G. A. Volkmer, S. H. Wang, Y. Wang, M. P. Weiner, P. Yu, R. F. Begley, and J. M. Rothberg. 2005. Genome sequencing in microfabricated high-density picolitre reactors. Nature 437:376-380. Martiny, J. B., B. J. Bohannan, J. H. Brown, R. K. Colwell, J. A. Fuhrman, J. L. Green, M. C. Homer-Devine, M. Kane, J. A. Krumins, C. R. Kuske, P. J. Morin, S. Naeem, L. Ovreas, A. L. Reysenbach, V. H. Smith, and J. T. Staley. 2006. Microbial biogeography: putting microorganisms on the map. Nut. Rev. Microbiol. 4:102-112. McBride, M. J., and D. R. Zusman. 1996. Behavioral analysis of single cells of Myxococcus xanthus in response to prey cells of Escherichia coli. FEMS Microbiol. Lett. 137:227231. Neil, R. B., D. Hite, M. I. Kelrick, M. L. Lockhart, and K. Lee. 2005. Myxobacterial biodiversity in an established oak-hickory forest and a savanna restoration site. Curr. Microbiol. 50:88-95. Olsen, G. J., D. J. Lane, S. J. Giovannoni, N. R. Pace, and D. A. Stahl. 1986. Microbial ecology and evolution: a ribosomal RNA approach. Annu. Rev. Microbiol. 40:337-365. Parsek, M. R., and E. P. Greenberg. 2005. Sociomicrobiology: the connections between quorum sensing and biofilms. Trends Microbiol. 13:27-33. Petrie, L., N.N. North, S. L. Dollhopf, D. L. Balkwill, and J. E. Kostka. 2003. Enumeration and characterization of iron(II1)reducing microbial communities from acidic subsurface sediments contaminated with uranium(V1). Appl. Environ. Microbiol. 69:7467-7479. Pham, V. D., C. W. Shebelut, M. E. Diodati, C. T. Bull, and M. Singer. 2005. Mutations affecting predation ability of the soil bacterium Myxococcus xanthus. Microbiology 1 51:1865-1 874. Porta, H., and M. Rocha-Sosa. 2001. Lipoxygenase in bacteria: a horizontal transfer event? Microbiology 147:3199-3200. Quillet, L., S. Barray, B. Labedan, F. Petit, and J. GuespinMichel. 1995. The gene encoding the beta-1,4-endoglucanase
AND EVOLUTION OF MYXOBACTERIA 2. ECOLOGY
(CelA) from Myxococcus xanthus: evidence for independent acquisition by horizontal transfer of binding and catalytic domains from actinomycetes. Gene 158:23-29. Rainey, P. B., and K. Rainey. 2003. Evolution of cooperation and conflict in experimental bacterial populations. Nature 425 ~72-74. Rankin, D. J., and A. Lopez-Sepulcre. 2005. Can adaptation lead to extinction? Oikos 111:616-619. Redfield, R. J. 2002. Is quorum sensing a side effect of diffusion sensing? Trends Microbiol. 10:365-370. Reichenbach, H. 1993. Biology of the Myxobacteria: ecology and taxonomy, p. 13-62. In M. Dworkin and D. Kaiser (ed.), Myxobacteria II. American Society for Microbiology, Washington, DC. Reichenbach, H. 1999. The ecology of the Myxobacteria. Environ. Microbiol. 1:15-21. Reichenbach, H., and G. Hofle. 1993. Biologically active secondary metabolites from Myxobacteria. Biotechnol. Adv. 11~219-277. Rose, M. R. 1984. Laboratory evolution of postponed senescence in Drosophila melanogaster. Evolution 38:10041010. Rosenberg, E., K. H. Keller, and M. Dworkin. 1977. Cell density-dependent growth of Myxococcus xanthus on casein. J. Bacteriol. 129:770-777. Rosenberg, E., and M. Varon. 1984. Antibiotics and lytic enzymes, p. 109-125. In E. Rosenberg (ed.),Myxobacteria: Development and Cell Interactions. Springer-Verlag, New York, NY. Sachs, J. L., U. G. Mueller, T. P. Wilcox, and J. J. Bull. 2004. The evolution of cooperation. Q. Rev. Biol. 79:135-160. Sanford, R. A., J. R. Cole, and J. M. Tiedje. 2002. Characterization and description of Anaeromyxobacter dehalogenans gen. nov., sp nov., an aryl-halorespiring facultative anaerobic myxobacterium. Appl. Environ. Microbiol. 689393900. Shi, W., and D. R. Zusman. 1993. The two motility systems of Myxococcus xanthus show different selective advantages on various surfaces. Proc. Natl. Acad. Sci. USA 90:33783382. Shimkets, L. J. 1999. Intercellular signaling during fruiting-body development of Myxococcus xanthus. Annu. Rev. Microbiol. 53:525-549. Shimkets, L. J. 1990. Social and developmental biology of the myxobacteria. Microbiol. Rev. 54:473-501. Shimkets, L. J., H. Reichenbach, and M. Dworkin. 2005. The Myxobacteria. In M. Dworkin (ed.), The Prokaryotes, 3rd ed., vol. 7, p. 31-115. Springer, New York, NY. Smith, D. R., and M. Dworkin. 1994. Territorial interactions between two Myxococcus species. J. Bacteriol. 176:12011205. Smith, J. M., and E. Szathmary. 1995. The Major Transitions in Evolution. W. H. Freeman Spektrum, New York, NY. Spiller, D. A,, and T. W. Schoener. 1998. Lizards reduce spider species richness by excluding rare species. Ecology 79503516. Sproer, C., H. Reichenbach, and E. Stackebrandt. 1999. The correlation between morphological and phylogenetic
39
classification of myxobacteria. Int. J. Syst. Bacteriol. 49~1255-1262. Strassmann, J. E., Y. Zhu, and D. C. Queller. 2000. Altruism and social cheating in the social amoeba Dictyostelium discoideum. Nature 408:965-967. Thaxter, R. 1892. On the Myxobacteriaceae, a new order of Schizomycetes. Bot. Gaz. 17:389-406. Travisano, M., J. A. Mongold, A. F. Bennett, and R. E. Lenski. 1995. Experimental tests of the roles of adaptation, chance, and history in evolution. Science 26787-90. Travisano, M., and G. J. Velicer. 2004. Strategies of microbial cheater control. Trends Microbiol. 12:72-78. Turner, P. E., and L. Chao. 1999. Prisoner’s dilemma in an RNA virus. Nature 398:441-443. Velicer, G. J. 2003. Social strife in the microbial world. Trends Microbiol. 11:330-337. Velicer, G. J., L. Kroos, and R. E. Lenski. 2000. Developmental cheating in the social bacterium Myxococcus xanthus. Nature 404598-601. Velicer, G. J., L. Kroos, and R. E. Lenski. 1998. Loss of social behaviors by Myxococcus xanthus during evolution in an unstructured habitat. Proc. Natl. Acad. Sci. USA 95:1237612380. Velicer, G. J., R. E. Lenski, and L. Kroos. 2002. Rescue of social motility lost during evolution of Myxococcus xanthus in an asocial environment. J. Bacteriol. 184:2719-2727. Velicer, G. J., G. Raddatz, H. Keller, S. Deiss, C. Lanz, I. Dinkelacker, and s. C. Schuster. 2006. Comprehensive mutation identificationin an evolved bacterial cooperator and its cheating ancestor. Proc. Natl. Acad. Sci. USA 103:8107-8112. Velicer, G. J., and K. L. Stredwick. 2002. Experimental social evolution with Myxococcus xanthus. Antonie Leeuwenhoek 81:155-164. Velicer, G. J., and Y. N. Yu. 2003. Evolution of novel cooperative swarming in the bacterium Myxococcus xanthus. Nature 42575-78. Vos, M. 2006. Natural variation in the social bacterium Myxococcus xanthus. Doctoral thesis. Universitat Tiibingen, Tiibingen, Germany. Vos, M., and G. J. Velicer. 2006. Genetic population structure of the soil bacterium Myxococcus xanthus at the centimeter scale. Appl. Environ. Microbiol. 72:3615-3625. Vulic, M., and R. Kolter. 2001. Evolutionary cheating in Escherichia coli stationary phase cultures. Genetics 158:519526. Watve, M. G., A. M. Shete, N. Jadhav, S. A. Wagh, S. P. Shelar, S. S. Chakraborti, A. P. Botre, and A. A. Kulkarni. 1999. Myxobacterial diversity in Indian soils: how many species do we have? Curr. Sci. 77:1089-1095. West, S. A., A. S. Griffin, A. Gardner, and S. P. Diggle. 2006. Social evolution theory for microbes. Nat. Rev. Microbiol. 4597-607. Whitaker, R. J., D. W. Grogan, J. W. Taylor. 2003. Geographic barriers isolate endemic populations of hyperthermophilic Archaea. Science 301:976-978. White, D., W. Shropshire, and K. Stephens. 1980. Photocontrol of development by Stigmatella aurantiaca. J. Bacteriol. 142:1023-1024.
40 Wireman, J. W., and M. Dworkin. 1977. Developmentally induced autolysis during fruiting body formation by Myxococcus xanthus. J. Bacteriol. 129:796-802. Wu, S. S., J. Wu, Y. L. Cheng, and D. Kaiser. 1998. The pilH gene encodes an ABC transporter homologue required for type IV pilus biogenesis and social gliding motility in Myxococcus xanthus. Mol. Microbiol. 29:1249-1261. Wu, Z. H., D. M. Jiang, P. Li, and Y. Z. Li. 2005. Exploring the diversity of myxobacteria in a soil niche by myxobacteriaspecific primers and probes. Environ. Microbiol. 71602-1610.
MYXOBACTERIAL BIOLOGY Xu, J. P. 2004. The prevalence and evolution of sex in microorganisms. Genome 47:775-780. Zahavi, A., and D. Ralt. 1984. Social adaptations in myxobacteria, p. 215-220. In E. Rosenberg (ed.), Myxobacteria: Development and Cell Interactions. Springer-Verlag, New York, NY. Zhang, Y. Q., Y. Z. Li, B. Wang, Z . H. Wu, C. Y. Zhang, X. Gong, Z. J. Qiu, and Y. Zhang. 2005. Characteristics and living patterns of marine myxobacterial isolates. Appl. Enviyon. Micro biol. 71:33 3 1-3 336.
DeueloDment
Myxobacteria: Multicellulurity and Differentiation Edited by David E. Whitworth 02008 ASM Press, Washington, D.C.
Michelle E. Diodati, Ronald E. Gill, Lynda Plamann, Mitchell Singer
Initiation and Early Developmental Events
Myxococcus xanthus is a rod-shaped, gram-negative soil bacterium that, when subjected to nutrient deprivation, undergoes a developmental process culminating in the formation of a multicellular fruiting body filled with spores. These predatory bacteria utilize proteins, peptides, and amino acids as their primary source of carbon, nitrogen, and energy and acquire these essential nutrients by cooperatively pooling extracellular hydrolytic enzymes to degrade other bacteria in soil (Bretscher and Kaiser, 1978; Dworkin, 1962). The colonial association of M . xanthus provides a competitive advantage over single, dispersed cells, and upon nutrient depletion, the formation of fruiting bodies cluster spores together, ensuring that new microcolonies arise after germination of myxospores under favorable environmental conditions. Elucidating the relationship between nutrient limitation and the mechanisms that cells employ to recognize and respond to it is fundamental for understanding how these organisms adapt to their environment. In order for M. xanthus cells to initiate development, three criteria must be met: (i) cells must be on a solid surface to allow gliding motility to occur (Kroos et al.,
3
1988; Wireman and Dworkin, 1975); (ii) cells must be at an appropriate density (Shimkets and Dworkin, 1981; Wireman and Dworkin, 1975);and finally, (iii)cells must be able to perceive a nutrient downshift, such that some energy capacity for protein synthesis remains (Dworkin, 1963; Hemphill and Zahler, 1968). The process of M . xanthus development requires approximately la5cells, so it is necessary that individual cells monitor not only their own nutritional status at the cellular level but also the nutritional status at the population level. Once the developmental program is initiated by starvation, it proceeds in a predictable manner with aggregation, mound formation, fruiting body formation, and sporulation within the fruiting bodies in a 24- to 48-h timeline (depicted in Fig. 1).For the purposes of this chapter, early development can be defined as events occurring from initiation to the start of aggregation at approximately the first 6 h poststarvation. In 1995, Singer and Kaiser proposed a dual model of starvation recognition for developing 111. xanthus cells (Singer and Kaiser, 1995). First, individual cells need to recognize starvation at the cellular level, and second, cells
~
Michelle E. Diodati and Mitchell Singer, Section of Microbiology, University of California-Davis, Davis, CA 95616. Ronald E. Gill, Department of Microbiology, University of Colorado Health Sciences Center, Denver, CO 80262. Lynda Plamann, School of Biological Sciences, Cell Biology and Biophysics, University of Missouri-Kansas City, Kansas City, MO 641 10.
43
DEVELOPMENT AND MOTILITY
44 Starvation
I
0 hours
Aggregation
1
6 hour$
FB and spore maturation
Mounds
t
I
i
"
12 hours
24 hours
TPM
Figure 1 Pictorial and photographic representations of the developmental process in M. xunthus DK1622. The diagram shows approximate times for each step in the process: starvation (0 h), aggregation (6 to 8 h), mound formation (12 h), fruiting body formation and sporulation (24 to 48 h). The first row represents development in an MC7 submerged culture system (ICuner and Kaiser, 1982), and the second row represents development on TPM starvation agar plates at a magnification of X40. This figure is adapted from Tzeng et al., 2006.
need to know that the population as a whole is starving. These two pieces of information, cellular starvation and population starvation, are perceived and integrated by these individual cells to activate the developmental program. In retrospect, this model as first described is relatively simplistic, yet the three basic tenets remain the same. Over the last 10 years, new experimental evidence pertaining to early development and sequence information obtained from the M. xanthus genome has increased our understanding of how M. xanthus cells recognize and respond to nutrient limitation. In this chapter, the following tenets are addressed: how individual M. xanthus cells recognize starvation; how these cells perceive population starvation; and how individual cells integrate this information to ultimately initiate fruiting body formation and cellular differentiation.
INITIATION OF DEVELOPMENT: CELLULAR STARVATION RECOGNITION M . xanthus is a representative of the proteolytic myxobacteria, relying primarily on proteins, peptides, fatty acids, and tricarboxylic acid cycle intermediates for their carbon, nitrogen, and energy needs (Bretscher and Kaiser, 1978; Dworkin, 1962). Like most developing microbes, the trigger for the developmental process in M. xanthus is nutrient deprivation, in conjunction with high cell density and a solid surface, as described above. Understanding how cells recognize a nutritional downshift
at the molecular level is critical for discerning the transition from vegetative growth to development.
Conditions That Induce Development and Other Starvation Conditions There are various types of starvation, not all of which induce fruiting body formation or differentiation in M. xanthus. These are listed in Table 1.Starvation for amino acids, carbon (such as pyruvate), or phosphate induces the developmental response (Dworkin, 1996; Manoil
Table 1 Conditions known to initiate development and
induce a stringent response" Condition Carbon starvation Amino acid starvation Essential amino acids Nonessential amino acids (auxotrophs) Amino acid analogues Phosphate starvation Purine starvation Decoyinine
Aggregation Sporulation (p)ppGpp
+
+
t
+ + + +
+ +
t 1'
+ +
t t
-
-
NC
Mycophenolic acid
-
-
NC
Pyrimidine starvation
-
-
ND
aAbbreviations and symbols: +, induces the condition; -, does not induce the condition; t,increases the intracellular concentration of (p)ppGpp; NC, no change in (p)ppGpplevels; ND, not determined.
EVENTS 3. INITIATION AND EARLYDEVELOPMENTAL
45 initiate the sporulation process (Ochi et al., 1982).In M . xanthus, there is a small decrease in GTP levels during the stringent response, but this decrease alone is unable to initiate the developmental program based on studies with inhibitors of in situ GTP synthesis decoyinine and mycophenolic acid (Singer and Kaiser, 1995).These studies led to the hypothesis that M. xanthus uses (p)ppGpp as an internal starvation signal, and the level of this molecule either by itself or in concert with other factors leads to the initiation of fruiting body formation.
and Kaiser, 1980b; Shimkets, 1984, 1987). However, unlike what is observed for Bacillus subtilis, starvation for guanine nucleotides does not instigate the developmental process (Singer and Kaiser, 1995); and while the addition of excess purines can lead to development (Campos and Zusman, 1975), this has been shown to be an artifact that indirectly causes a nutritional imbalance (Manoil and Kaiser, 1 9 8 0 ~ )Furthermore, . Kimsey and Kaiser have shown that pyrimidine starvation hinders growth but does not trigger fruiting body development (Kimsey and Kaiser, 1991). A common feature of all conditions that induce development is that they also induce a stringent response in M . xanthus (Manoil and Kaiser, 1980b).It has been previously demonstrated that one of the earliest responses in M. xanthus development is the increase in the intracellular concentration of (p)ppGpp (Manoil and Kaiser, 1980a, 1980b; Singer and Kaiser, 1995; for reviews see Cashel et al., 1996, and Chatterji and Ojha, 2001). This alarmone accumulates during amino acid starvation and acts as an intracellular starvation signal that is both necessary and sufficient for the initiation of early developmental gene expression in M . xanthus (Singer and Kaiser, 1995).This is in contrast to B. subtilis sporulation, where a dramatic decrease in the GTP pool has been shown to
The Stringent Response: a General Model Coordinating Metabolism to Global Gene Expression The stringent response was first observed in Escherichia coli more than 50 years ago (Cashel and Gallant, 1969; Pardee and Prestidge, 1956; Sands and Roberts, 1952; Stent and Brenner, 1961). Since then, components of the system have been found in virtually every bacterium examined to date (Chatterji and Ojha, 2001; Ogata et al., 2001; Sun et al., 2001; M. E. Diodati and M. Singer, personal communication), and the E. coli stringent response remains the paradigm. A model of the E. coli stringent response is shown in Fig. 2. Interestingly, an analogous system with RelNSpoT homologues has also
p+ i T p p k
= ADP ATP
PPPG (GTP)
Ndk
1;:- I
PPG(GW
DksA+
1~ p ' " + ~ p i
~
SpoT PPPGPP
~
-
7
0
Positively controlled genes
PPGPP
PPi
Negatively controlled genes
Figure 2 Diagram of the E. coli stringent response. The enzymes involved in the (p)ppGpp metabolism are shown in bold. Ribosome-associated RelA or SpoT catalyzes the synthesis of pppGpp from ATP and GTP upon amino acid or carbon starvation, respectively. Gpp (or Ppx) dephosphorylates pppGpp to make ppGpp. (p)ppGpp accumulates in the cell and interacts with RNAP and DksA [which has been shown to play an important role in (p)ppGppdependent transcriptional regulation (Paul et al., 2004, 2005; Perederina et al., 2004)] to positively and negatively control transcription to respond to starvation. (p)ppGpp levels are modulated by SpoT, and when nutrient conditions change, SpoT hydrolyzes ppGpp to GDP (ppG). Ppk is involved in ATP synthesis, and Ndk forms GTP from ATP and GDP (ppG).For reviews, consult Cashel et al., 1996, and Chatterji and Ojha, 2001.
46
been found in plants (van der Biezen et al., 2000; Givens et al., 2004). Comparisons of the Stringent Response in Bacteria The stringent response links amino acid availability to the rate of protein synthesis through the signaling mol[(p)ppGpp]. ecules guanosine-Sr-(tri)di-3’-diphosphate The response directly inhibits stable RNA synthesis and protein elongation and activates the transcription of amino acid biosynthetic operons. There is a plethora of secondary or indirect effects which include the inhibition of ribosomal protein, phospholipid, and cell wall constituents synthesis, inhibition of DNA replication, and an increase in the production of stress proteins (for reviews see Cashel et al., 1996, and Chatterji and Ojha, 2001). Therefore, the stringent response allows cells to rapidly respond to nutrient limitation by modulation of metabolic pathways, rRNA synthesis, and stressadaptive genes, thus allowing cells to respond and adapt to starvation. In E. coli, two related proteins, RelA and SpoT, modulate the intracellular levels of (p)ppGpp either by the transfer of pyrophosphate from ATP to the 3’ hydroxyl of GTP to form pppGpp or by hydrolyzing ppGpp to pyrophosphate and GDP, respectively (Cashel and Gallant, 1969; Fiil et al., 1977; Laffler and Gallant, 1974). RelA is a ribosome-associated protein required for the synthesis of (p)ppGpp in response to stalled ribosomes due to a decrease in charged tRNAs binding to the acceptor site during amino acid starvation (Block and Haseltine, 1974; Cochran and Byrne, 1974; Haseltine and Block, 1973; Pedersen et al., 1973) (Fig. 2). Thus, (p)ppGpp acts as a monitor of amino acid availability for translation. In contrast, SpoT does not appear to be associated with the ribosomes, and it is a bifunctional enzyme that has both biosynthetic and degradative properties. The activity of SpoT, whether it is biosynthetic or hydrolytic, is influenced by a variety of environmental signals, although it is unclear how SpoT’s activity is regulated at a molecular level (Murray and Bremer, 1996). As mentioned above, E. coli has two distinct and separable enzymes for (p)ppGpp metabolism, RelA and SpoT (Cashel and Gallant, 1969; Laffler and Gallant, 1974). Although many enterics and other species have homologues of these two proteins, recent studies have demonstrated that there is divergence in the strict conservation of these homologues. Many bacteria, both gram negative and gram positive, have only a single bifunctional ribosome-associated protein that is both SpoT- and RelA-like, including B. subtilis (Wendrich and Marahiel, 1997), Sinorhizobium meliloti (Wells and
DEVELOPMENT AND MOTILITY Long, 2002), and Rhodobacter capsulatus (Masuda and Bauer, 2004). Sequence analysis of the M. xanthus RelA protein (MXAN3204) revealed that it contains both the hydrolytic and biosynthetic domains found in E. coli SpoT (Sun et al., 2001; Diodati and Singer, personal communication), implying that the M. xanthus RelA protein is a bifunctional enzyme similar to B. subtilis RelA (Wendrich and Marahiel, 1997),Streptomyces coelicolor RelA and RshA (Chakraburtty et al., 1996; Sun et al., 2001), Streptococcus equisimilis Re1 (Mechold and Malke, 1997), and RelA in the gram-negative bacterium Rhodobacter capsulatus (Masuda and Bauer, 2004). This is very intriguing because it has been suggested that the relA and spoT genes in gram-negative organisms evolved from a duplicated gram-positive re1 (rsh)-like gene (Mittenhuber, 2001) and the majority of relAlspoT hybrid (rsh)genes are found in gram-positive organisms (Jain et al., 2006). Interestingly, Harris et al. (Harris et al., 1998) found two bands with a 311-bp relA probe that mapped to two different regions of the M. xanthus chromosome. This suggested the possibility of two relAlspoT-like genes in 111. xanthus. This initial observation is supported by the identification of a second putative relAlspoT-like gene, in addition to relA, in the M. xanthus genome (Goldman et al., 2006). Stringent Response-Related Homologues in M. xanthus Sequence analysis revealed that M. xanthus encodes all the known components of the (p)ppGpp cycle. It has a single copy of ndk (nucleoside diphosphate kinase), gpp (guanosine pentaphosphatase), ppx (exopolyphosphatase), and ppk (polyphosphate kinase), with the last two being involved in polyphosphate metabolism, as well (for a review, see Cashel et al., 1996).
Two relA/spoT-like homologues in M. xanthus The M. xanthus genome (Goldman et al., 2006) has a second putative relAlspoT-like gene (MXAN1364).The product of MXAN1364 has strong sequence similarity to the N-terminal hydrolase domain of SpoT; therefore, it has been designated “shd” for SpoT hydrolase domain. shd has been previously described by Diodati et al. as mx-1.594 (Diodati et al., 2006). shd is a small 420-bp gene and, based on sequence homology, is predicted to have a partial SpoT motif HD domain (Aravind and Koonin, 1998). Shd has the conserved HD domain with the substrate binding pocket and the HD doublet motifs found in the superfamily of metal-dependent hydrolases (Aravind and Koonin,
EVENTS 3 . INITIATIONAND EARLYDEVELOPMENTAL 1998; Hogg et al., 2004). Notably, E. coli RelA, which is solely a synthetase, has substitutions in these regions of the protein. The downstream RelA/SpoT domains that are necessary for (p)ppGppsynthesis are missing in Shd. Therefore, Shd may have hydrolytic properties and may possibly be involved in regulating (p)ppGpplevels. Although Shd has not yet been examined for SpoT activity, of particular interest is that the shd gene was previously implicated in M. xanthus development by O’Connor and Zusman (O’Connor and Zusman, 1990; O’Connor and Zusman, personal communication) as a temperature-sensitive aggregation (Tag) mutant. However, the exact nature of the Tag mutation in shd is not known.
M. xanthus has four DksA homologues Annotation of the M . xanthus genome has identified several homologues to dksA, recently shown in E . coli to play a critical and synergistic role in (p)ppGpp-dependent transcriptional regulation (Paul et al., 2004, 2005; Perederina et al., 2004). DksA binds to RNA polymerase and enhances ppGpp’s direct negative effect on rRNA promoters by reducing the open complex lifetime of RNA polymerase (RNAP) and inhibiting rRNA promoter activity (Paul et al., 2004; Perederina et al., 2004). DksA also directly and indirectly affects activation of amino acid promoters in concert with ppGpp (Paul et al., 2005). Furthermore, DksA has been shown to be involved in cell division, quorum sensing, expression of virulence factors, and the suppression of temperaturesensitive growth in dnaK mutants (Branny et al., 2001; Ishii et al., 2000; Kang and Craig, 1990; Turner et al., 1998).Although AdksA mutants have pleiotropic effects on cells including misregulation of numerous genes, mild UV sensitivity (Clifton et al., 1994), and filamentation (Ishii et al., ZOOO), it has been suggested that these are indirect effects of alterations in rRNA transcription and subsequent RNAP titration in the mutant (Paul et al., 2004). In d k s A mutant cells, rRNA promoters are unresponsive to changes in amino acid availability, growth rate, or growth phase (Paul et al., 2004). Interestingly, M . xanthus has four DksA homologues in its genome (Goldman et al., 2006) designated DksA (MXAN3200), DksB (MXAN3006), DksC (MXAN5718), and DksD (MXAN7086) (Table 2), instead of the typical single copy that exists in other organisms. These homologues were found by performing a BLAST analysis of the M. xanthus genome with E. coli DksA. MXAN3200 has the highest homology to E. coli DksA and therefore was named “DksA.” The M. xanthus homologues are slightly shorter than E. coli or B. subtilis DksA, but have the full-length
47
C-terminal 4-cysteine zinc finger suggestive of a transcriptional regulator.
EshA, PgpH, and HvrA Recently, three additional proteins, EshA, PgpH, and HvrA, have been shown to modulate (p)ppGpplevels or (p)ppGpp-dependent gene expression during vegetative growth in a variety of bacterial species (Liu et al., 2006; Masuda and Bauer, 2004; Saito et al., 2006). EshA is a cyclic AMP (CAMP)-binding protein that when disrupted, results in lower levels of (p)ppGppaccumulation during early to late growth phase in S. coelicolor (Saito et al., 2006). Based on the Saito et al. work, EshA is proposed to fine-tune and maintain a specific ppGpp level during stationary phase for antibiotic production via its nucleotide-binding domain. M. xanthus has two homologues to EshA (Table 2). In Listeria monocytogenes, PgpH is a putative integral membrane protein with an HD domain at its C terminus (Liu et al., 2006). The HD domain suggests that it may act as a metal-dependent phosphohydrolase (Aravind and Koonin, 1998). A cold-sensitive L. monocytogenes mutant with a transposon insertion in pgpH accumulates higher levels of (p)ppGppthan the wild-type cells. Therefore, PgpH is hypothesized to play a role in the cold-induced stress response by directly or indirectly modulating ppGpp levels, which are increased during low-temperature growth, and restoring them to normal vegetative levels (Liu et al., 2006). M . xanthus has a PgpH homologue defined by MXAN4737 (Table 2). Lastly, a third stringent-response-related protein, HvrA, was recently identified. In R. capsulatus, an hvrA mutation suppresses the lethality of a SPOTmutant (Masuda and Bauer, 2004). HvrA is a nucleoid-like protein, and in E. coli similar proteins coregulate (p)ppGppdependent genes by influencing the supercoiled state of the promoters (Johansson et al., 2000). Masuda and Bauer (2004)suggest that HvrA may be acting in concert with (p)ppGpp to regulate the transcription of specific promoters during growth. Although M. xanthus has homologues of the eshA and pgpH genes, no homologues have been found for hvrA (Table 2).
The Role of (p)ppGppin M. xanthus Development The initial starvation response occurs at the level of the individual cell, with each cell required to evaluate its own nutritional status. Because M. xanthus is unable to utilize carbohydrates, cells primarily rely on amino acids and a-keto acids to serve as carbon and energy sources, as well as substrates for protein synthesis. Physiological studies support the hypothesis that M. xanthus initially senses starvation by monitoring its translational capacity
DEVELOPMENT AND MOTILITY
48 Table 2
List of stringent-response-related homologues in M. xanthus“
Homologue name (organism)
Gene name (derivation) in M. xantbus
M x no.
MXAN no.
RelA (E. coli)
relA
4330
3204
SpoT HD domain (E. coli)
Shd (SpoT H D domain )
1594
1364
DksA (E. coli)
dksA
2229
3200
DksA ( E . coli)
dksB (DksA homologue B)
6139
3006
DksA (E. coli)
dksC (DksA homologue C)
0673
5718
DksA ( E . coli)
dksD (DksA homologue D)
5736
7086
EshA ( S . coelicolor)
MXAN6248
6739
6248
EshA ( S . coelicolor)
MXAN6249
6738
6249
PgpH (L. monocytogertes)
MXAN4737
3629
4737
HvrA (R. sphaeroides)
NA
NA
NA
Role in original organism
Comments
Actually an rsh (relAlspoT) gene; annotated as “stringentrespon” in M. xanthus Expression is modified in (p)ppGpp degradation (Diodati and Singer, nla4 mutant; incorrectly personal communicaannotated as “relA” in tion) M. xanthus Originally identified as (p)ppGpp-regulated transcription DnaK suppressor ( E . coli) (Paul et al., 2004) Partial DksA homologue, (p)ppGpp-regulated transcription also known as DksA3006 (Paul et al., 2004) (p)ppGpp-regulated Partial DksA homologue, transcription also known as DksA5718 (Paul et al., 2004) (p)ppGpp-regulated Partial DksA homologue, transcription also known as DksA7086 (Paul et al., 2004) Sustains (p)ppGpp dur- CAMP binding protein; ing late growth phase finely tunes ppGpp (Saito et al., 2006) threshold for antibiotic production Sustains (p)ppGpp dur- Overlap or duplication? ing late growth phase (Saito et al., 2006) Adjusts (p)ppGpp levels Putative integral membrane protein; has H D domain during low temp growth (Liu et al., 2006) Coregulates (p)ppGpp- Mutations suppress spoT lethality (Masuda and dependent genes durBauer, 2004); no homoing growth (Masuda logue in M. xanthus and Bauer, 2004) (p)ppGpp synthesis (Cashel et al., 1996)
“Mx and MXAN numbers refer to gene names from the original M 1 genome (Jakobsen et al., 2004) and the completed TIGR (The Institute of Genomic Research)/ Monsanto versions of the M. xanthus genome (Goldman et al., 2006),respectively. N o homologues of HvrA were found in M. xanthus. The corresponding references for determining the roles of the above genes are included in parentheses. NA, not applicable.
via the intracellular levels of (p)ppGpp (Manoil and Kaiser, 1980a, 1980b; Singer and Kaiser, 1995). This model is very attractive, as it provides a molecular link between metabolism and development of M. xanthus, and remains a starting point to understand this complex sensory pathway. Previously, Singer and Kaiser (1995) demonstrated that the M. xanthus RelA protein also functions as a ribosome-dependent (p)ppGpp synthetase and is required for the earliest aspects of development (Harris
et al., 1998; Manoil and Kaiser, 1980a, 1980b; Singer and Kaiser, 1995).Where it has been specifically examined, most developmentally regulated genes require an increase in the intracellular levels of (p)ppGpp for their expression, includingproduction of the extracellular A- and C- signals. relA mutants do not aggregate or form spores, and other genes that are known to decrease ppGpp levels within the cells, such as the csgA (Crawford and Shimkets, 2000b), nlal8 (Diodati et al., 2006), and nla4 (Diodati and Singer, personal communication; F. Ossa,
3. INITIATION AND EARLYDEVELOPMENTAL EVENTS M. E. Diodati, N. B. Caberoy, M. Singer, and A. G. Garza, unpublished data) mutants, also fail to sporulate. Interestingly, todK (MXAN6955), a gene that affects the timing of development, requires starvation for its expression but not (p)ppGppaccumulation (Rasmussen and Sngaard-Andersen, 2003). This suggests that todK may represent a member of a new class of genes that are starvation dependent and RelA independent. This is supported by preliminary DNA microarray data comparing global patterns of expression from reZA mutant and wildtype cells. Jose and Singer (I. R. Jose and M. Singer, personal communication) have identified a subset of genes whose expression is starvation induced and (p)ppGpp independent. In addition to relA, other genes have been implicated in (p)ppGpp regulation in M . xanthus, including two genes involved in C-signal regulation, socE and csgA (MXAN1294) (Crawford and Shimkets, 2000; see chapter 4 for a discussion on C-signal), and the nutrient sensor, Nsd (Brenner et al., 2004). Nsd (MXAN7402) is described in more detail in the nutrient sensors portion of this chapter. More recently, we found that the inactivation of two genes that encode os4-transcriptional activators, nZal8 (MXAN3692) (Caberoy et al., 2003; Diodati et al., 2006) and nZu4 (MXAN2516) (Caberoy et al., 2003; Ossa et al., unpublished), result in ppGpp accumulation defects, suggesting that at least two d4 promoters are regulating the expression of unknown genes whose gene products affect ppGpp levels. The exact mechanism of action for each of these gene products is not known and may be through RelA activity or stability, interaction with the ribosome, influencing the levels of (p)ppGppprecursors, or (p)ppGppmetabolism. The identification of these additional components that affect (p)ppGpp levels suggest that the regulation of the stringent response is more complex in M . xanthus. A list of genes and their (p)ppGppeffects upon inactivation is given in Table 3 .
49 levels necessary to support growth. Once the M . xanthus developmental process is under way, starvation must be monitored and available nutrients must be diverted toward completion of development. Crawford and Shimkets (2000b) have shown that when SocE is depleted, cell growth is arrested, and cells start to accumulate (p)ppGpp and initiate a stringent response even in the presence of sufficient nutrients to support growth. In addition, DNA and stable RNA synthesis is inhibited [which correlates with increased (p)ppGpplevels], and sporulation is induced upon socE depletion. Their data are consistent with SocE acting as a repressor of development. The C-signaling protein, CsgA, appears to induce the stringent response. It has been shown, through amino acid substitution studies of the CsgA protein and mutant analysis, that CsgA maintains growth arrest during development, stimulates (p)ppGpp synthesis in the absence of SocE, and mediates sporulation. Moreover, upon starvation, the csgA mutant can initiate a stringent response but accumulates only one-half of the (p)ppGppof wild-type cells and fails to sustain these higher levels during development (Crawford and Shimkets, 2000b). Therefore, it appears that although CsgA is not essential for the stringent response when it is initiated by amino acid starvation, it is important for maintaining the response during A-signaling and beyond. Interestingly, not only is the M. xunthus stringent response regulated by the SocE and CsgA proteins, but also their transcription is dependent on the stringent response. The transcription of socE is inhibited by increased (p)ppGpp levels, as may be expected for
Table 3 List of genes affecting (p)ppGpp accumulation in M. xanthus Gene name
SocE and CsgA The stringent response, the recognition of starvation and the subsequent increase in (p)ppGpp levels that accompany the limitation of amino acids, mediates multiple physiological and metabolic changes in the cell (Cashel and Rudd, 1989). A unique aspect of the M . xanthus stringent response is that it is regulated, in part, by the SocE and CsgA proteins. Maintenance of the response is important because the stringent response is coupled to the formation of the multicellular fruiting body. Within 2 h of development, A-signal, which is composed mostly of amino acids, is produced and cells need to decipher the extracellular signal levels of amino acids from the
relA socE csgA
nsd nlal8 nla4
(PjPPGPP effect when inactivated"
No accumulation Increased 2.5- to 5-fold in 1% CYE Decreased 2-fold after starvation Increased 2-fold in 0.5% CTT Decreased 2-fold (veg), 5.5-fold (dev) Decreased 2-fold (veg), 2.6-fold (dev)
Reference(sj Harris et al., 2001 Crawford and Shimkets, 2000b Crawford and Shimkets, 2000b Brenner et al., 2004 Diodati et al., 2006 Ossa et al., unpublished; Diodati and Singer, personal communication
"dev, development; veg, vegetative growth.
50 a repressor of development. In contrast, increased (p)ppGpplevels stimulate csgA transcription (Crawford and Shimkets, 2000a). This RelA-dependent transcriptional regulation of both csgA and socE RNA levels creates an increase in the ratio of CsgA to SocE and results in cessation of growth and the redirection of resources towards development (Crawford and Shimkets, 2000a, 2000b). In summary, the balance of SocE and CsgA proteins in the cell is critical for sustaining the developmental program past initiation and is just one example of the unique aspects of the stringent response in this organism.
Nla18 and Nla4 When nZal8 is inactivated, it results in pleiotropic effects in vegetative and developmental gene expression (Caberoy et al., 2003; Diodati et al., 2006). These mutants grow two to three times slower than wild-type cells, exhibit a mild temperature-sensitive phenotype, and are very prone to lysis due to disruptions in membrane permeability and overall integrity (Diodati et al., 2006). These mutants accumulate 18 to 50% less ppGpp than the wild type upon nutrient downshift and have similar developmental defects to relA mutants, albeit not as severe. The phenotype of the nla28 mutant cannot simply be explained by its ppGpp defect; the membrane protein defect is not seen with relA mutants, and vegetative microarray data show that nlal8 mutants affect the regulation of a variety of translation-related genes and genes encoding transcriptional regulators (Diodati et al., 2006). Therefore, Nlal8’s role in the accumulation of ppGpp appears to be indirect, because Nlal8 is required for overall balanced growth. Nla4 is a second d4transcriptional activator that is required for normal vegetative growth and development in M . xanthus. nla4 mutant cells fail to aggregate and sporulate normally, and when codeveloped with wildtype cells, the sporulation deficiency fails to be rescued (Caberoy et al., 2003). Further analysis of this mutant reveals multiple defects in developmental gene expression and signal production (Ossa et al., unpublished) and ppGpp accumulation (Diodati and Singer, personal communication; Ossa et al., unpublished). RelA regulation Although recent work (Harris et al., 1998; Manoil and Kaiser, 1980a, 1980b; Singer and Kaiser, 1995)has demonstrated the importance of the stringent response in starvation recognition and development in M. xanthus, little is known about the regulation and control of the key regulator RelA. Pertinent questions such as how the RelA protein modulates its biosynthetic and degradative
DEVELOPMENT AND MOTILITY properties in response to environmental cues still remain. It has recently been shown that the bifunctional RelM SpoT homologue in Streptococcus dysgalactiae subsp. equisimilis catalyzes opposing synthetase and hydrolase reactions at distinct active sites (Hogg et al., 2004). The two different conformations of the enzyme, in which either the synthetase or the hydrolase activity is turned on while the reciprocal activity is repressed, appear to involve transmission of a ligand-induced signal between the two active sites (Hogg et al., 2004). This reciprocal regulation of the two catalytic activities is governed by the C-terminal half of the Re1 protein (Mechold et al., 2002). Similar regulation has been found for the RelN SpoT homologue in Mycobacterium tuberculosis, as well (Avarbock et al., 2000). The regulatory process controlling M. xanthus RelA enzyme activities may be similar to these other bifunctional proteins. In vitro biochemical and kinetic tests with purified RelA have yet to be performed. One approach to begin to answer the question of RelA regulation is to first identify and characterize the components of the M . xanthus stringent response and then elucidate the mechanism that controls the levels and activities of the secondary messenger, (p)ppGpp.A model summarizing the M . xanthus (p)ppGppresponse is shown in Fig. 3.
Other Early Developmental Effectors: Nutrient Sensors versus Developmental Timers Over the last 10 to 15years, data from several laboratories have identified two general classes of genes that alter the timing of development: developmental timers and nutrient sensors. A partial list of genes involved in these two processes is provided in Table 4. Developmental timers are genes defined by mutations that either speed up or slow down the developmental process but still require starvation for activation of development. Nutrient sensors, on the other hand, are genes defined by mutations that cause a premature initiation of development under inappropriate conditions, i.e., conditions that would normally not initiate the developmental pathway by wild-type cells. Mutations in genes encoding developmental timers, such as espAB (MXAN0931 and -0932) (Cho and Zusman, 1999a), espC (MXAN6855) (Lee et al., 2005), redCDEF (MXAN0459 through -0462) (Higgs et al., 2005), todK (MXAN6955) (Rasmussen and Sogaard-Andersen, 2003), and rodK (MXAN0733) (Rasmussen et al., ZOOS), have been shown to either accelerate or delay the developmental program. However, in all cases nutrient limitation was still required to initiate the developmental response in these mutants. Intriguingly, many of the developmental timers uncouple the spatial requirement of aggregation and
3. INITIATION AND EARLYDEVELOPMENTAL EVENTS
51
P+ i T P p k
j:zp 1:c.(= ADP ATP
PPPG (GTP)
Ndk
+ ppi
DksA + + Positively controlled
-r'
GPPf Ppx P P P G P P . 7
Nla18
CsgA
SocE Nsd
Development
PPG(GDP)
PPi
PPGPP
E d
genes
1
Negatively controlled genes
Figure 3 Diagram of the M. xanthus (p)ppGpp response and genes involved in its activation. The enzymes involved in (p)ppGpp metabolism are shown in bold. The asterisk (") represents uncharged tRNA in the acceptor site of the ribosome that triggers the associated RelA to catalyze the synthesis of (p)ppGpp from ATP and GTP. See Figure 2 legend for more details. In M . xanthus, (p)ppGpp levels are maintained by balancing the hydrolase activity of RelA with its biosynthetic activity. Diodati and Singer have postulated that Shd, a gene product with homology to the hydrolase domain of E. coli SPOT, may play a role in (p)ppGpp degradation. In addition, five proteins (SocE [Crawford and Shimkets, 200Ob1, Nsd [Brenner et al., 20041, Nla18 [Diodati et al., 20061, Nla4 [Diodati and Singer, personal communication; Ossa et al., unpublished], and CsgA [Crawford and Shimkets, 2000bJ) have been shown to either inhibit or stimulate ppGpp accumulation in M. xanthus through as yet unknown mechanisms. For more details, consult text. With elevated (p)ppGpp levels, RNA polymerase (Eo*) is predicted to interact with DksA, to modulate changes in RNA polymerase activity to alter gene expression. To date no secondary (p)ppGpp biosynthetic pathway has been identified.
sporulation. Inactivation of espA, espC, or rodK results in sporulation outside the fruiting body (Cho and Zusman, 1999a; Lee et al., 2005; Rasmussen et al., 2005). In addition, it has been shown that overexpression of CsgA results in this same phenotype (Kruse et al., 2001), implying that a lack of inhibition of the C-signaling pathway leads to induction of sporulation independent of the high cell density requirement to reach the appropriate C-signal threshold. While increased C-signal production results upon loss of RodK and EspC function (Lee et al., 2005; Rasmussen et al., 2005), csgA and espA mutant mixing experiments suggest that the espA mutant produces less C-signal than wild-type cells (Lee et al., 2005). These data imply that EspA may have a function other than inhibiting C-signal production causing a mutant in
espA to sporulate outside the fruiting body. Elucidating the mechanisms of spatial coupling of fruiting bodies and sporulation can lead to fascinating insights into the intricate regulation of this process. Also, there are many other developmental mutants that affect the timing of development. The overall defect is usually a delay, which may be due to numerous factors, including defects in motility, outer membrane components, or poor growth (i.e., poor nutrient stores).
Nutrient Sensors Nutrient sensing encompasses at least four important elements: (i) the detection of nutrients in the environment, (ii) the uptake of nutrients into the cell, (iii) metabolism
DEVELOPMENT AND MOTILITY
52
Table 4 Partial list of genes implicated in nutrient sensing and developmental timing Gene or locus
Sensor or timer
asgD
Sensor
bcsA
Sensor
Che3 operon
Sensor
MXAN2 9 02 (mx-332 0) nsd
Sensor Sensor
sigC
Sensor
socE
Sensor
spdR
Sensor
espA espB esp C redCDEF rodK todK
Timer Timer Timer Timer Timer Timer
Null phenotype Hypersensitive to nutrient levels Forms fruiting bodies on rich media Forms fruiting bodies on rich media Hypersensitive to nutrient levels Forms fruiting bodies on rich media Forms fruiting bodies on rich media Forms fruiting bodies on rich media Forms fruiting bodies on rich media Speeds up development Slows down development Speeds up development Speeds up development Speeds up development Speeds up development
and utilization of those nutrients by the cell, and (iv) the regulation of developmentally specific signal transduction pathways necessary to respond to the abundance or lack of nutrients in the cell’s environment. We have previously described how (p)ppGpp can act as an internal starvation signal, using the cell’s translational capacity as a monitor of the cell’s nutritional status. However, it has become evident that (p)ppGpp is not the only participant in starvation recognition. Work from several labs has identified genes that have been implicated in affecting nutrient sensing. These genes include the previously described socE (Crawford and Shimkets, 2000a, 2000b), nsd (Brenner et al., 2004), the genes of the che3 cluster (crdB, mcp3A, mcp3B, cheA3 [MXAN5147 through 51521) (Kirby and Zusman, 2003), a component of the A-signaling generation complex, asgD (MXAN6996) (Cho and Zusman, 1999a), and a developmentally regulated os4-transcriptional activator, MXAN2902 (previously described as Mx-3320) (Jakobsen et al., 2004). Nsd Nsd (nutrient-sensing/utilizing defective) (Brenner et al., 2004) was originally identified as the gene controlled by the developmentally regulated promoter known as
04469 (IR nomenclature represents TnSlac insertions) (Kroos et al., 1986). Nsd appears to be important for sensing nutrients in the environment and acts as an inhibitor of development in the presence of nutrients. Brenner et al. (2004)have shown that, under low nutrient conditions, growth is decreased 2- to 2.5-fold in nsd mutants compared to wild-type cells. Upon further characterization in 0.5% Casitone-Tris (CTT) broth, nsd mutants accumulate twofold more (p)ppGppthan wild-type cells. Moreover, nsd mutant cells initiate development on nutrient agar, but development is compromised; viable spore count is reduced (Brenner et al., 2004). Interestingly, on higher-nutrient plates (>0.5% CTT), nsd mutants can go through development but produce heat- and sonicationresistant phase-dark cells instead of spores (Brenner et al., 2004). These data demonstrate that the nsd mutants can initiate development and aggregate in the presence of nutrients, yet appear to require additional factor(s) for wild-type sporulation under these conditions. T h e Che3 operon Che3 is a chemosensory cluster (which includes crdB, mcp3A, mcp3B, cheA3, and others) that has been shown to expedite development when the genes are inactivated. crdB, mcp3A, mcpSB, and cheA3 mutants form fruiting bodies earlier than wild-type cells in a density-independent manner (Kirby and Zusman, 2003). This is corroborated by the observation that developmental reporter fusions 04403, IR4411, and 04521 are expressed earlier and at higher levels in mutant cells during development, as well as during vegetative growth in the Amcp3A mutant (Kirby and Zusman, 2003). In addition, these mutants form distorted fruiting bodies with spores on nutrient agar. These data suggest that the proteins encoded by the che3 cluster genes are nutrient sensors that act by blocking developmental gene expression during growth. There is a divergently transcribed os4-transcriptionalactivator gene, crdA, upstream of the gene cluster. It is proposed, based on mutational and yeast two-hybrid analyses, that the Che3 system modulates the activity of CrdA, which controls expression of specific developmental genes (Kirby and Zusman, 2003). It is important to note that the rapidly formed fruiting bodies of the che3 mutants on clone fruiting (CF) media look similar to wild-type fruiting bodies but the mutants are unable to sporulate normally. The vegetative expression of development-specific genes, the bypassing of the highdensity requirement for development, and the general premature entry into development phenotype of the che3 mutants suggest that these temporal and/or signaling checkpoints of fruiting body formation are critical for wild-type sporulation.
3 . INITIATIONAND EARLYDEVELOPMENTAL EVENTS As gD The inactivation of the asgD gene results in cells that are hypersensitive to nutrients; their development is inhibited by limited amounts of nutrients that are low enough to induce development in wild-type cells. On CF medium (10 mM MOPS [3-4-morpholine propanesulfonic acid], 0.015% Casitone [Difco], 8 mMMgSO,, 1 mMKH,PO,,0.2% sodiumcitrate,0.02% (NH,),SO,, 0.1% pyruvate, and 1.5% agar), which contains asmall amount of nutrients for cells to undergo a gradual starvation, AasgD mutants form loose aggregates and do not sporulate (Cho and Zusman, 1999a). In contrast, on the more-stringent starvation medium MMC (10 mM MOPS buffer, 4 mM MgSO,, 2 mM CaCl,, and 1.5% agar), AasgD cells are able to form slightly irregular fruiting bodies and have 35% of wild-type sporulation (Cho and Zusman, 1999a).In order to decipher the component(s) in the CF media that were causing the developmental inhibition, the authors removed components and added them back individually. Eliminating citrate, pyruvate, or Casitone from the media increased the asgD mutant’s spore count to 0.25, 0.96, and 44% of wild-type numbers, respectively (Cho and Zusman, 1999a). Taken together, these data suggest that AsgD may be involved in nutrient sensing. When asgD is inactivated, the cells do not perceive starvation until nutrient concentrations are minuscule. Once these nutrients are consumed, the mutant cells may not have enough energy to sustain and complete development. Therefore, as opposed t o having a function of inhibiting development, like Nsd and the proteins of the Che3 cluster, AsgD appears to be acting by promoting cells to develop when there are still sufficient nutrients in the environment to support the developmental process. Furthermore, the AasgD mutant can be extracellularly complemented and appears to be a member of the “asg mutant” group (Cho and Zusman, 1999a). Since mutations in asgD also affect A-signal production, AsgD may serve as a link between starvation sensing and activation of the extracellular A-signal generation pathway described in the latter sections of this chapter. MXAN2 902 MXAN2902 (previously described as Mx-3320) is a crs4-transcriptional activator protein that appears to be involved in sensing nitrogen-related nutrients in the environment at about 12 h of development (Jakobsen et al., 2004). When the MXAN2902 gene is disrupted, mutants are defective in mound formation (Jakobsen et al., 2004). The mounds of the mutant are flatter than the wild type, and they have projecting tails of cells at
53 one end. Spores are found within the mound but also at the periphery and are concentrated in the protruding tail of lower-density cells. This density-independent manner of sporulation is similar to what is seen with the che3 cluster mutants and many developmental timer mutants described above. Like the asgD mutant, these cells had defective development on CF agar but show wild-type development on more-stringent starvation agar. When particular components were removed from the CF medium or added to the TPM medium (10 mM Tris-HC1 [pH 8.01, 1 mM KH,PO,, 8 mM MgSO,, and 1.5% agar) to interpret the nutrient-sensing defect, the mutants were hypersensitive to the nitrogen sources in the media (Jakobsen et al., 2004). With these results, and the phenotypic similarities that the MXAN2902 mutant shares with the asgD mutant in regards to nutrient sensing, MXAN2902 may act as a positive effector of development as well. Nutrient sensors can work by the promotion or inhibition of development, as described above. The method by which these nutrient sensors elicit their actions to affect entry into development is not yet known. Possible direct or indirect mechanisms could include detection, transport, or metabolism of nutrients as well as the regulation of signal transduction pathways that may or may not control these above-mentioned functions. As demonstrated by the characterization of the nutrient sensor genes, the overall decision to initiate development requires a highly regulated system with the input and cooperation of many factors in order to proceed.
Global Gene Regulation during the Cellular Response to Starvation Once nutrient limitation is detected, cells respond by redirecting transcription to prepare for the new environmental condition. M. xanthus cells must meet density requirements as well as have solid support. It has been shown that overall protein synthesis patterns are dramatically different during the early stationary phase and initiation of development, even though under both conditions, cells are entering a nutrient-limiting environment (Ueki and Inouye, 1998). It is evident that there are additional factors that influence the cells to initiate development, which is an expensive, energy-demanding process. Therefore, cells need to distinguish between a gradual, an immediate, or an absolute starvation state with enough nutrients to carry out development but not enough to sustain vegetative growth, coupled with a strong community consensus. For example, wild-type cells grown on rich agar media, such as CTT (Hodgkin and Kaiser, 1979) or CTTYE (CTT supplemented with
54 0.5% yeast extract), enter stationary phase but do not undergo fruiting body development or differentiation. These data suggest that M. xanthus cells and populations evaluate multiple facets of their environment and then respond accordingly. Strict control of gene expression is needed for the appropriate survival response to be evoked. Sigma factors are important regulatory elements that bacteria utilize to increase the specificity of largescale transcriptional responses. Control of induction of development and/or early developmental gene expression appears to rely, in part, on three global regulators: RpoN, SigD (the M . xanthus RpoS homologue), and SigC.
RpoN and Associated Activator Proteins If developmental criteria are met, M . xanthus cells respond by initiating a complex pathway that causes changes in behavior and culminates in cellular differentiation. Many of the known early developmentally regulated genes in M . xanthus are transcribed from d4like promoters. These include the previously characterized genes spi (MXAN4276) (Keseler and Kaiser, 1995), mbhA (MXAN7061) (Romeo and Zusman, 1991), asgE (MXAN1010) (Garza et al., 2000a, 2000b), and sdeK (MXAN1014) (Garza et al., 1998; Pollack and Singer, 2001), all of which are expressed within the first few hours of development. rpoN (MXANlOGl), which encodes d4, is usually associated with specialized metabolic functions, such as nitrogen regulation in E . coli and Salmonella (Kustu et al., 1989; Ninfa et al., 1995) or motility in Pseudomonas (Hobbs et al., 1993), Caulobacter (Brun and Shapiro, 1992), and M. xanthus (Wu and Kaiser, 1997). There is a single copy of rpoN in M. xanthus (Goldman et al., 2006). Uniquely, rpoN is essential for growth (Keseler and Kaiser, 1997). as4-dependent promoters require a positive activator for transcription. The identification of as4-likepromoter elements (based on their -12 and -24 sequences) 5’ to many of the earliest developmental genes suggests a role for rpoN and these associated transcriptional activators in early development. Enhancer binding proteins (EBPs) are transcriptional activators for d4promoters and are named after the eukaryotic enhancer binding proteins they resemble. These proteins typically bind 100 to 200 bp upstream of d4promoter elements (Morett and Buck, 1988) and interact with RNAP through a DNA looping mechanism that allows the isomerization of the RNAP from the closed to the transcriptionally active open complex in an ATP-dependent manner. (For reviews, see Morett and Segovia, 1993; Studholme and Dixon, 2003; and Xu and Hoover, 2001.)
DEVELOPMENT AND MOTILITY The NtrC protein of E . coli is one of the best characterized of these activator proteins, and the term “NtrClike activators” has been used interchangeably with “enhancer binding proteins” to describe these activators in M. xanthus. NtrC-like activators are usually associated with a subset of activators containing an N-terminal response regulator domain. For the purposes of this general discussion and to prevent confusion, the term “enhancer binding protein” or “EBP” will be used to describe these os4-transcriptional activators. Fifty-two potential EBPs have been identified by sequence analysis in M . xanthus (Caberoy et al., 2003; Gorski and Kaiser, 1998; Jakobsen et al., 2004; Jelsbak et al., 2005) and are listed in Table 5. EBPs typically have a three-domain structure that consists of a regulatory domain(s)at the N terminus, a highly conserved central ATPase domain, and a helix-turn-helix DNA binding motif at the C terminus (Jelsbak et al., 2005; Studholme and Dixon, 2003). These activators were identified in the M . xanthus genome sequence, using the conserved central ATPase domain. Based on the essential nature of rpoN in M. xanthus and the fact that many of the earliest known developmental genes have predicted RpoN-dependent promoters, Keseler and Kaiser (1997) suggested that early development may be a function of a succession or cascade of EBPs that drives progression through the early stages of development; analogous to the sigma factor cascade that drives B . subtilis sporulation (Kroos et al., 1999). The collective efforts of several labs to identify EBPs and to characterize knockout and insertion mutants have revealed 18 that have vegetative and/or developmental phenotypes (Caberoy et al., 2003; Gorski and Kaiser, 1998; Gronewold and Kaiser, 2001, 2002; Guo et al., 2000; Hager et al., 2001; Jakobsen et al., 2004; Jelsbak et al., 2005; Kaplan, 2003; Tse and Gill, 2002). Interestingly, most of these activators with developmental phenotypes are important for early developmental gene expression. Therefore, RpoN and associated EBPs may play an important role in coordinating the starvation recognition process and the early steps of development. Eleven EBPs are described in detail below. The seven remaining EBPs with vegetative and/or developmental defects have been attributed to motility or late developmental defects (Table 5 ) and are beyond the scope of this chapter.
EBPs with vegetative growth defects As previously described, nlal8 and nla4 mutants have severe vegetative defects including slow growth and decreased ppGpp accumulation (Diodati et al., 2006; Diodati and Singer, personal communication; Ossa
Table 5 Names, MXAN and M x numbers, N-terminal regulatory domains, gene knockout constructions, mutant phenotypes, and related references for all EBPs in M . xanthusa Regulatory domain(s)b
KO?
Phenotype and comments
Name(s) of EBP
MXAN no.
M x no.
ActB, Mxa259
3214
4338
RR
Yes: I, D
Late devdef; adjacent to RR
CrdA, Nla26, Mxa227
5153
1467
RR
Yes: I, D
Early devdef, nutrient sensor (see text); HK nearby
FrgC, Nla25
1128
223 8
RR
Yes: I
WT; adjacent to HK
HsfA
5364
1035
RR
No
MrpB, Mxa189
5124
5602
RR
Yes: I, D
Mx-1288
4339
1288
FHA
Yes: I
Mx-3098, Mxa198
5041
3098
GAF
Yes: I
Interacts with sigma 70 RNAP and activates lonD (bsgA)in vitro; adjacent to HK Early devdef (see text); adjacent to HK WT; adjacent to STK (divergently transcribed) WT; up at 12 h of development
Mx-3320
2902
3320
Not identified
Yes: I
Mx-3 725
3333
3725
FHA, GAF
Yes: I
Mx-48 85
4899
4885
FHA
Yes: I, D
Mxal91 Mxa211
0353 0172
1757 3558
FHA RR
Yes: I No
Mxa213 Mxa221 Mxa249 Mxa264 Mxa296, Mxa2lO
4020 4196 5672 0116 0180
1598 3656 2469 5079 5565
FHA, GAF RR FHA FHA Not identified
Yes: I No No Yes: I Yes: I
Nla 1 Nla2 Nla3 Nla4
5853 3381 4785 2516
3336 3471 3814 0839,0840
RR Not identified RR RR
Yes: I Yes: I Yes: I Yes: I
WT WT; adjacent to HK
Nla5
2501
1502
FHA, FHA
Yes: I
WT
Devdef, nutrient sensor MXAN 2902 (see text) WT; lacks GAFTGA motif; STK nearby Late devdef; near STK-associated KapB Slow growth (see text) HK nearby (divergently transcribed) Early dev; STK nearby Adjacent to HK
WT; adjacent to STK Early devdef (see text); HK nearby
S-motility defect; adjacent to HK
Slow growth, devdef (see text)
Reference(s) Gorski et al., 2000; Gorski and Kaiser, 1998; Gronewold and Kaiser, 2001,2002 Caberoy et al., 2003; Gorski and Kaiser, 1998; Kirby and Zusman, 2003 Caberoy et al., 2003; Cho et al., 2000 Ueki and Inouye, 2002
Gorski and Kaiser, 1998; Sun and Shi, 2001a, 2001b Jelsbak et al., 2005 Gorski and Kaiser, 1998; Jakobsen et al., 2004 Jakobsen et al., 2004 Jelsbak et al., 2005 Jelsbak et al., 2005 Gorski and Kaiser, 1998 Gorski and Kaiser, 1998; Kaufman and Nixon, 1996 Gorski and Kaiser, 1998 Kaufman and Nixon, 1996 Kaufman and Nixon, 1996 Gorski and Kaiser, 1998 Gorski and Kaiser, 1998; Diodati and Singer, personal communication Caberoy et al., 2003 Caberoy et al., 2003 Caberoy et al., 2003 Caberoy et al., 2003; Ossa et al., unpublished Caberoy et al., 2003
(Continued)
Table 5 Names, MXAN and M x numbers, N-terminal regulatory domains, gene knockout constructions, mutant phenotypes, and related references for all EBPs in M . xanthusa(Continued) Name(s) of EBP
Regulatory domain(s)*
KO?
Phenotype and comments
Reference(s)
MXAN no.
Mx no.
NlaG
4042
2063
RR
Yes: I
Nla7 Nla8 Nla9, TaR3
0937 4580 3952
2140 2840 2942
RR RR GAF
Yes: I Yes: I Yes: I
NlalO
5048
4193
Not identified
Yes: I
Nlall
6426
4170
Not identified
Yes: I
Nla12 Nla13 Nla14
2159 3811 3095
4562 6755 4901
FHA, GAF RR FHA
Yes: I Yes: I Yes: I
NlalS, Nla16 Nla17 Nla18
5680 3418 3692
0033, 0233 0517 0888,0889
RR' RR FHA
Yes: I Yes: I Yes: I
Nla20 Nla21
4252 4983
2594 4341
RR RtcR
Yes: I Yes: I
Nla22, Mx-4756
4240
4756
RR
Yes: I
Nla23, PilR2
5777
1973
RR
Yes: I
Nla24
7440
2057
RR
Yes: I
Nla27 Nla2 8
1345 1167
2176 1617
FHA RR
Yes: I Yes: I
Nla34
1565
4965
GAF
No
A- and S-motility defect; adjacent to HK WT Early devdef (see text); adjacent to HK Down regulated in nlal8 mutant
PilR, Mxa15
5784
3013
RR
Yes: I, D
S-motility defect; adjacent to HK
Gorski and Kaiser, 1998; Kaufman and Nixon, 1996; Wu and Kaiser, 1995, 1997
SasR, Mxa287
1245
0124
RR
Yes: I, D
Early devdef (see text); adjacent to HK
Gorski and Kaiser, 1998; Guo et al., 1996; Yang and Kaplan, 1997
Early devdef (see text); adjacent to HK WT; adjacent to HK WT; adjacent to HK WT, regulates antibiotic TA synthesis WT; adjacent to STK (divergently transcribed) WT; adjacent to STK (divergently transcribed) WT; adjacent to STK WT; adjacent to HK WT; adjacent to STK (divergently transcribed) WT WT, adjacent to HK Slow growth, devdef (see text); adjacent to STK (divergently transcribed) WT, adjacent to HK WT; upstream RNA 3' terminal phosphate cyclase (divergently transcribed) WT; up at 12 h of development; HK nearby S-motility defect; adjacent to HK
Caberoy et al., 2003 Caberoy et al., 2003 Caberoy et al., 2003 Caberoy et al., 2003; Y. Paitan, E. Orr, E. Z. Ron, and E. Rosenberg, unpublished data Caberoy et al., 2003 Caberoy et al., 2003 Caberoy et al., 2003 Caberoy et al., 2003 Caberoy et al., 2003 Caberoy et al., 2003 Caberoy et al., 2003 Caberoy et al., 2003; Diodati et al., 2006 Caberoy et al., 2003 Caberoy et al., 2003
Caberoy et al., 2003; Jakobsen et al., 2004 Caberoy et al., 2003; Jelsbak and Kaiser, 2005 Caberoy et al., 2003; Lancero et al., 2004 Caberoy et al., 2003 Caberoy et al., 2003 Diodati et al., 2006
3. INITIATION AND EARLYDEVELOPMENTAL EVENTS
3 0
v 4-2
8V
3
c
e?
*
d p:
Z 2 2 2
00
m c n m 3 r . c n
I\ 0
CA 3
m
z
!xa
"J
v)
0
W
Y
W
r
d d
.
2
r
.
m
f
--
57 et al., unpublished). Nla18 (MXAN3692) is an atypical EBP that has a forkhead-associated (FHA) domain. as its N-terminal regulatory module, instead of the more-common response regulator domain that Nla4 (MXAN2516) contains (Diodati et al., 2006; Jelsbak et al., 2005). Also, nlul8 is adjacent to pskB5, a gene that encodes a serine threonine kinase that is important for timely development (S. Inouye, personal communication), which is down regulated in an nlu18 mutant during vegetative growth (Diodati et al., 2006). Proteins with FHA domains have been shown to interact with threonine-phosphorylated substrates of serinekhreonine protein kinases in M. tuberculosis(Alderwick et al., 2006; Molle et al., 2003). Such activators may act as regulatory links to various signal transduction pathways mediated by serinehhreonine kinases (Alderwick et al., 2006; Jelsbak et al., 2005; Kroos, 2005; Molle et al., 2003). Twelve EBPs with FHA domains have been found in M. xanthus (Jelsbak et al., 2005), and four of these activators, including Nlal8, have vegetative or developmental phenotypes when their genes are disrupted. Inactivation of mxa213 (MXAN4020) (Gorski and Kaiser, 1998) or mx-4885 (MXAN4899) (Jelsbak et al., 2005) results in cells with developmental defects and is discussed below. Disruption of mxul91 (MXAN0353) (Gorski and Kaiser, 1998) results in a mutant with a slow growth rate similar to nlul8 or nlu4 mutants but, interestingly, displays wild-type development.
EBPs with early developmental defects Eight of the 18 EBPs with vegetative or developmental phenotypes affect early developmental events. These activators are SpdR (MXAN1078),CrdA (MXAN5153), SasR (MXAN1245), MrpB (MXAN5124), Mxa296 (MXAN0180), Mxa213 (MXAN4020), Nla6 (MXAN4092), and Nla28 (MXAN1167) (Caberoy et al., 2003; Gorski and Kaiser, 1998; Guo et al., 2000; Hager et al., 2001; Kirby and Zusman, 2003; Sun and Shi, 2001a, 2001b; Tse and Gill, 2002). SpdR is a typical EBP with a response regulator domain that is in an operon with its putative cognate histidine kinase, SpdS. It was found as a bypass suppressor for bsgA mutants (Hager et al., 2001; Tse and Gill, 2002) and is described in more detail in the cell signaling portion of this chapter. SpdR is expressed during vegetative growth and acts as a nutrient sensor that inhibits development in the presence of nutrients. Mutations in spdR alter a cell's vegetative patterns of gene expression, and spdR mutants express developmental genes on rich media. Also, an early A-signal-dependent developmental gene with a oS4promoter, spi (R4521), requires SpdR for its expression (Hager et al., 2001). Based on these data, SpdR may
Next Page
58 act between the assessment of nutrient conditions and signal-dependent gene expression. CrdA is an EBP that is negatively regulated by the nutrient-sensing genes of the Che3 gene cluster described in the previous section. The crdA gene is divergently transcribed from the che3 promoter region, and CrdA and CheA3 (a histidine kinase) interact very strongly with each other in yeast two-hybrid studies (Kirby and Zusman, 2003). Cells with mutations in crdA are delayed 12 to 24 h in development and reduce and delay the expression of mbhA during development. Furthermore, CrdA may be directly or indirectly controlling a wide variety of developmental genes because mutations in che3 cluster genes, of which CrdA is epistatic, affect a large number of developmental genes (Kirby and Zusman, 2003). The CheA3 kinase appears to modulate the activity of its cognate response regulator, CrdA, by inhibiting activation of the EBP to prevent developmental gene expression during vegetative growth. SasR was first identified and characterized by Gorski and Kaiser (1998) as Mxa287. The gene corresponding to Mxa287 was found, along with 1 2 other putative d4 transcriptional activators, with degenerate PCR probes to the conserved central domain (Gorski and Kaiser, 1998; Kaufman and Nixon, 1989). Mutants in mxa287 have both vegetative and developmental defects. On nutrient agar, the colonies are smaller and less cohesive, and on starvation media the cells are blocked early in development, appear flat, and are unable to express 04521. When complementation studies were carried out, mxa287 mutants could produce both A- and C-signal but failed to respond to the extracellular signals provided by the wild-type cells (Gorski and Kaiser, 1998). This led Gorski and Kaiser to hypothesize that Mxa287 may be acting as part of a signal reception pathway. These observations and conclusions corroborated very nicely with the sasR characterization described by Kaplan and colleagues (Guo et al., 2000; Yang and Kaplan, 1997). SasR is part of a signal transduction pathway involved in A-signal sensing (Guo et al., 2000). The sasR gene encodes a d4response regulator that is in an operon with the histidine kinase, sass. SasR is predicted to function downstream of Sass, as a positive regulator of spi (a4521) expression (Yang and Kaplan, 1997).SasR is discussed in more detail in the A-signaling section of this chapter. The mrp locus consists of a histidine kinase homologue (mrpA)and an NtrC-like response regulator gene ( m r p B )in an operon adjacent to an independently transcribed CAMP receptor protein-like transcriptional regulator (mrpc) (Sun and Shi, 2001b). MrpB is induced upon starvation, up-regulated during development, and
DEVELOPMENT AND MOTILITY required for both aggregation and sporulation. MrpB is also known as Mxa189, which was unable to be characterized due to difficulties in obtaining insertion mutants with the targeted PCR probe method (Gorski and Kaiser, 1998). The mrpB deletion mutants are completely flat on starvation media. Based on site-directed mutagenesis studies of the conserved aspartate of MrpB, the phosphorylated form of MrpB is required for aggregation while dephosphorylation of MrpB is needed for sporulation (Sun and Shi, 2001b). Sun and Shi (2001a, 2001b) proposed that MrpB functions after (p)ppGpp and A-signaling, but before C-signaling. The expression of mrpB is reduced in both a relA and an asgA mutant background, but not in a csgA mutant. In addition, mrpB mutants produce A-signal to 80% of wild-type levels and very little C-signal. Expression of six TnSluc fusions, including A- and C-signal-dependent markers, were reduced in the mrpB mutant (Sun and Shi, 2001a, 2001b). Although these data do support the claim that mrpB is acting after starvation initiation, the predicted function of MrpB after A-signaling does not account for the 41% reduction in the expression of the A- and Csignal-independent a4408 (sdeK) in the mrpB mutant. The role of MrpB in sdeK regulation requires further characterization. MrpB function prior to C-signaling is supported by data showing that MrpC is essential for the expression of fruA, a key transcription factor required for C-signaling, and that MrpB regulates mrpC expression (Nariya and Inouye, 2006). Taken together, the role of MrpB is complex and is an essential part of the development process in M. xanthus. Two EBPs with unique domain structures that affect early development are Mxa296 and Mxa213. Mxa296 is also known as Mxa210 because DNA sequence analysis of the mxa210 pLAGl (Gorski and Kaiser, 1998) plasmid insert reveals that it is identical to the central region of the mxa296 gene (Diodati and Singer, personal communication). mxa296 mutant cells are able to develop normally on TPM agar but fail to develop in submerged culture in polystyrene microtiter plates (Gorski and Kaiser, 1998). Consistent with this phenotype is the observation that 04521 is expressed on TPM agar but not in submerged culture. Interestingly, the mutant is able to express 04521 in TPM or MC7 buffered suspension (Kuner and Kaiser, 1982), suggesting that the polystyrene is somehow inhibiting development at the preaggregation stage (Gorski and Kaiser, 1998). Further investigation is needed to decipher the subtleties of this fascinating phenotype. Based on sequence analysis, Mxa296 has an unusual EBP domain structure in that it lacks a readily identifiable N-terminal sensory domain (Jelsbak et al., 2005). Only 6 of 52 activators have this
Previous Page
3. INITIATIONAND EARLYDEVELOPMENTAL EVENTS characteristic (Jelsbak et al., 2005; Diodati and Singer, personal communication; Table 5 ) . In contrast, Mxa213 is an EBP with two regulatory domains; it has both an FHA and a GAF domain at its N terminus (Jelsbak et al., 2005). As described previously, FHA domains potentially interact with threonine-phosphorylated substrates of serinehhreonine kinases (STKs) and there is a gene encoding a putative STK close to mxa213 (Jelsbak et al., 2005). GAF domains are found in cyclic GMP-specific and stimulated phosphodiesterases, adenylate cyclases, and E. coli FhlA and NifA related proteins (Studholme and Dixon, 2003). It is predicted that GAF domains may regulate signaling events via the binding of ligands such as nucleotides and small molecules (Aravind and Ponting, 1997). Insertional disruption of mxa213 causes cells to be blocked in development. On TPM agar, these mutants form large aggregates that have irregular shapes and patterns of darkening. The mxa223 mutant cells fail to sporulate, and the sporulation defect is not corrected by mixing with wild-type cells, indicating a cell autonomous defect in sporulation. Interestingly, these mutants are defective in the expression of the C-signal-dependent a 4 4 1 4 (Gorski and Kaiser, 1998). A closer examination of C-signal production in this mutant is needed to determine if it is lower than wild-type levels. Nla6 and Nla28 are EBPs that are NtrC-like response regulators with similar mutant phenotypes. In addition, the nla28 gene is downstream of a histidine kinase gene. Inactivation of nla28 or nla6 results in a short delay in aggregation, reduced production of A- and C-signals, and 50- to 500-fold fewer spores than wild-type cells, respectively (Caberoy et al., 2003). Even though these mutants are defective in the production of the early extracellular A-signal and show a defect in aggregation, more-detailed developmental characterization is needed to temporally pinpoint where the mutations first manifest themselves. RpoN and the associated EBPs that activate transcription at u54promoters are fundamental to the overall physiology of M. xanthus. Elucidating the roles of these activators can uncover the secrets to the unique essentiality of RpoN in M . xanthus. With the data from detailed mutational analyses of these activators and prospective genetic and direct binding studies, we can begin to build models of ordered networks of these regulators in M. xanthus in the near future.
SigD, the M. xanthus RpoS Homologue In E. coli, RpoS functions as a master stress-related sigma factor and is induced under a variety of general stress conditions including starvation, transition into stationary phase, and osmotic shock (Brown et al., 2002; Hengge-Aronis, 2002). The RpoS homologue,
59 SigD (MXAN2957), appears to have roles in early and late developmental gene expression (Viswanathan et al., 2006; Yoder and Kroos, 2004a, 2004b), as well as in stationary phase (Ueki and Inouye, 1998). SigD is essential for M . xanthus survival during stationary phase and for the expression of a large number of proteins that are produced or degraded under conditions of nutrient deprivation. During vegetative growth, AsigD mutants cease to grow past late exponential phase and lose viability when the cultures are plated on nutrient media after entry into stationary phase (Ueki and Inouye, 1998). In addition, two-dimensional gel electrophoresis of wild-type and AsigD mutants reveals that protein patterns are significantly different during late exponential phase in the two strains (Ueki and Inouye, 1998). Furthermore, AsigD mutants lack or have limited resistance to a variety of stresses. Mutants fail to grow after heat shock, h.ave reduced growth upon cold shock, are more sensitive to H,O,-induced oxidative stress, and fail to accumulate osmoprotective trehalose in response to osmotic shock (Ueki and Inouye, 1998). These data demonstrate the importance of SigD to overall cell viability in M. xanthus. In addition to the multiple growth phenotypes, the sigD mutant is delayed in development and defective in sporulation (Ueki and Inouye, 1998). Recently, a more detailed developmental characterization of the sigD mutant was performed, and SigD appears to play a largely positive role in regulating aspects of the developmental program (Viswanathan et al., 2006). Regulation of RpoS in E. coli is very complex and multifaceted; regulation can occur at the level of transcription, mRNA turnover, translation initiation, and proteolysis (Brown et al., 2002). In M. xanthus, based on studies with lac2 transcriptional and translational fusions, there is differential regulation of SigD at the level of transcription and translation during exponential growth and transition into stationary phase (Ueki and Inouye, 1998). The transcription of the sigD gene increases upon entry into stationary phase and then decreases during stationary phase. Conversely, the activity of the SigD translational fusion is increased during stationary phase and remains constant. This pattern of regulation appears to depend on whether cells are exposed to more-gradual nutrient-limiting conditions with accumulation of waste products (stationary phase), gradual nutrient-limiting conditions to initiate development (CF plates), or abrupt starvation conditions such as a shift to TPM liquid media (Ueki and Inouye, 1998; Viswanathan et al., 2006). When cells were plated on CF agar to induce development, the activity of both the transcriptional and translational fusions increased until
60 12 h and then decreased (Ueki and Inouye, 1998). Upon stringent starvation of cells in TPM media, sigD transcript levels fall after 20 min and stay low after 40 min (Viswanathan et al., 2006). These dissimilar patterns of expression under the three starvation conditions exemplify the fundamental physiological differences between these nutrient-restricted environments. Interestingly, in relA mutants, sigD transcript levels are substantially lower during exponential growth than in wild-type cells, but the sigD mRNA levels rise above wild-type levels after 40 min poststarvation (Viswanathan et al., 2006). This suggests that (p)ppGpp levels in the cell directly or indirectly regulate sigD transcription. SigC, a Sigma Factor and Nutrient Sensor SigC (MXAN6209)is a third sigma factor that is important for the cell’s decision to undergo development. Deletion mutants of sigC have normal vegetative growth and form fruiting bodies on CF agar, albeit slightly deformed and elongated, with wild-type sporulation (Apelian and Inouye, 1993). The sequence of SigC has homology to the heat shock sigma factor, 032, but when sigC is inactivated, heat shock proteins are still induced to wild-type levels (Ueki and Inouye, 2001). Also, in contrast to sigD mutants, AsigC mutants can produce normal amounts of trehalose upon osmotic shock and during sporulation (Apelian and Inouye, 1993). Interestingly, AsigC mutants form fruiting bodies on 0.5% CTT after 15 h and sporulate. Under these same conditions, wild-type DZFl cells formed few (<3% of As&) fruiting bodies and were unable to sporulate (Apelian and Inouye, 1993). This suggests that SigC may act as a nutrient sensor similar to Nsd and members of the Che3 cluster and may have a role in the transcription of genes that negatively regulate initiation of development.
A Model of Overall Nutrient Sensing and the Starvation Response in M . xanthus Based on the current data, the simplest model for nutrient sensing still focuses on the cell’s ability to utilize its translational capacity as an overall measurement of starvation. Data from the studies on nla18 (Caberoy et al., 2003; Diodati et al., 2006) and nla4 (Caberoy et al., 2003; Ossa et al., unpublished) support the premise previously described by Manoil and Kaiser ( 1 9 8 0 ~ ) that a disruption in balanced growth has severe effects on development. Mutations in either gene cause a severe vegetative growth defect and alter the normal pattern of ppGpp production in response to nutrient deprivation. This suggests that the vegetative state of the cell is important for its ability to properly recognize a starvation situation.
DEVELOPMENT AND MOTILITY When cells are starved for carbon, nitrogen, or phosphate, they immediately induce a stringent response by synthesizing (p)ppGpp. The ability of vegetative cells to mount this response is absolutely dependent upon RelA and possibly other nutrient sensors as well, such as Nsd (Brenner et al., 2004). Once the response has been triggered, regulation of gene expression is redirected, through direct and indirect effects of (p)ppGpp and DksA on RNAP. General starvation response genes are activated, and a subset of these genes is required for development. These developmental genes include several EBPs in addition to SigD (RpoS) (Ueki and Inouye, 1998). The expression and/or activation of these regulators allows the cells to respond to nutritional stress and initiate the first steps of fruiting body formation, including the induction of the extracellular signaling system, A-signal (see following sections). The mechanism by which many of these nutrientsensory systems, EBPs, and sigma factors act to integrate environmental cues and regulate starvation and development-specific gene expression remains a mystery. However, in some cases it is known that some proteins like SigD and MrpB act downstream of (p)ppGpp accumulation. The next challenge is to determine the various targets of these (p)ppGpp-dependent regulators and to establish the interactions of these genetic circuits. Progress is being made in this area. Work by Sun and Shi (2001a, 2001b) has demonstrated that mrpB expression is partially dependent upon (p)ppGppand that mrpB controls, at least partially, sdeK expression, which is required for passage through the aggregation stage of development (Garza et al., 1998; Pollack and Singer, 2001; Sun and Shi, 2001b). With the release of the M. xantbus sequence and the availability of M. xantbus DNA microarrays to study global transcriptional regulation, identifying members of these various regulatory pathways and determining how they are integrated to control the initiation of development can now be readily accomplished.
CELL-CELL SIGNALING IN EARLY DEVELOPMENT: POPULATION STARVATION RECOGNITION Over the past 30 years many excellent reviews have been written focusing on the isolation, characterization, and description of the five currently known extracellular signaling systems in M. xanthus (for reviews, see Kaiser, 2004, and Kaiser and Kroos, 1993). In this section the goal is not to reiterate these past findings, but to integrate more recent data and to expand on the models governing how early development initiates and responds to two of the earliest signals, the B- and the A-signaling systems.
EVENTS 3. INITIATION AND EARLYDEVELOPMENTAL
A Historical Overview of Cell-Cell Signaling in Myxobacteria Myxobacteria are social organisms that feed, move, and develop in a cooperative manner. They consume organic material from the soil by the secretion of extracellular enzymes, including proteases, lipases, cellulases, and other digestive enzymes; the products of these digestive reactions are then taken up by the cells and used as carbon, nitrogen, and energy sources (Bretscher and Kaiser, 1978; Dworkin, 1962). Based on this feeding behavior, the Myxobacteria have been described as bacterial “wolf packs,” preying on susceptible soil microbes (Dworkin, 1973).The ability of myxobacterial cells to “pool” these extracellular enzymes to benefit the colony was quantified by Rosenberg et al. (1977). This cooperativity in feeding provides a strong selection for colonial growth and may have provided the impetus for the evolutionary development of the myxobacterial fruiting body as a mechanism to promote the survival of a large number of cells in close physical association. Myxobacteria move by gliding, a complex mechanism that translocates cells across a solid surface (for reviews, see Doetsch and Hageage, 1968, and Kuhwein and Reichenbach, 1968). Genetic, biochemical, and physiological studies have clearly demonstrated that cell-cell interactions, including the production of extracellular signals, are involved in controlling motility. As with feeding, cell density also has a strong effect on the expansion rate of the myxobacterial colony (Kaiser and Crosby, 1983).Two genetically distinct motility systems, A (adventurous) and S (social), collectively result in gliding motility. Of interest for this discussion is that both motility systems require specific extracellular signaling pathways. For example, early genetic studies identified the tgl and cgl loci, which encode proteins involved in motility signaling in a population (Hodgkin and Kaiser, 1977). Based on these examples of feeding and movement, where the viability of the individual cell is tied to the population, it is not surprising that M . xanthus has evolved complex signaling systems to drive the developmental program. In 1962, McVittie et al. (1962) published a paper describing a pair of developmental mutants that individually could not develop but when cocultured could form fruiting bodies. Unfortunately, because one of the mutants was a pyrimidine auxotroph, and cross feeding could not be ruled out, this observation was ignored for several years. It was not until 1978 that the first set of nonautonomous developmental mutants were published (Hagen et al., 1978). Hagen et al. (1978) isolated 115 mutants that could be placed into four different extracellular complementation groups based on
61 cell-mixing experiments. These groups were designated A, B, C, and D. A fifth class, designated E, was later isolated by Downard et al. (1993). Each complementation group, when cultured by itself or with members of its own class, could not develop. However, development could progress when cells were cocultured with either wild-type cells or members of a different class. This extracellular complementation test would become a powerful tool for the isolation and characterization of mutants defining the extracellular signaling systems in M . xanthus. A great deal of information has been collected since these extracellular signals were first identified almost 30 years ago. Since several excellent reviews have been published regarding the isolation and early characterization of these signaling mutants and systems, data obtained since the publication of Myxobacteria I1 (Dworkin and Kaiser, 1993) are focused on. For discussion on the initiation of development, the A and B groups are the most relevant and are described below. The extracellular signals are described in a temporal manner; therefore, the B group is discussed before the A group.
The B-Signal, an Initiating Signal for Development The B-signaling mutants isolated by Hagen et al. are a genetically and phenotypically heterogeneous group (Gill and Bournemann, 1988; Hagen et al., 1978; LaRossa et al., 1983). The locus responsible for the B-signaling mutant phenotype has been determined only for three of the mutants tested. For these three strains, the B-signaling mutant phenotype is attributable to the loss of function of a single gene, homologous to the ATP-dependent protease lonD (MXAN3993), designated bsgA. Mutations in the bsgA gene, in an otherwise wild-type genetic background, demonstrate the B-signaling phenotype (Gill and Cull, 1986; Gill et al., 1988; Tojo et al., 1993a, 1993b). It should be noted at the outset of this discussion that other members of the Group B signaling mutants remain unmapped and are yet to be extensively characterized. This section exclusively discusses the properties of the bsgA mutants. Morphologically, bsgA mutants appear to be blocked quite early in the developmental program. They fail to produce fruiting bodies, and the production of refractile, sonication-resistant spores is less than 10-4 of wild-type levels. The effect of bsgA mutations on developmental gene expression has been assessed indirectly using collections of developmentally induced lacZ fusions, and directly using the expression of certain development-specific proteins (Gill and Cull, 1986; Kroos and Kaiser, 1987; LaRossa et al., 1983). In each of these studies, wild-type bsgA was required for normal levels
62 of expression of each of the genes tested, including representatives that are normally expressed immediately upon starvation. For many developmental genes used in these studies, mutations in bsgA abolished expression. Curiously, however, lac2 fusions with expression times prior to 6 h and certain fusions with expression times from 6 to 14 h had only a partial dependency on bsgA for expression as indicated by a reduced level of P-galactosidase expression (Kroos and Kaiser, 1987). This relationship has not been fully explained. Nevertheless, a reasonable hypothesis is that expression of the early developmental genes are responsive, perhaps in an additive fashion, to regulation by multiple levels of regulatory input, only one of which is dependent upon the bsgA product. The gene expression data discussed above corroborate morphological observations suggesting that BsgA is required very early in the developmental program. However, the precise temporal and regulatory relationships between nutritional starvation, the relA-mediated stringent response, and bsgA are less well defined. For example, expression of starvation-induced, relA-independent genes (Rasmussen and Ssgaard-Andersen, 2003; Jose and Singer, personal communication) are yet to be studied in a bsgA mutant background. Furthermore, LaRossa et al. (1983) reported that representative B-signaling mutants accumulate guanosine tetraphosphate to a level at least 50% of that of wild-type cells (as determined by the ratio of guanosine tetraphosphate to guanosine triphosphate levels measured after 1 h of incubation in nutrient-free buffer). On this basis, the authors suggest that the Bsignaling mutants exhibit a normal stringent response to starvation. However, it is tenuous to extrapolate these data to the bsgA mutants, since it has not yet been established whether or not the mutations used in this study have been reported to map to this locus. In addition, considering the rather subtle, but likely critical, effect of mutations in csgA on the stringent response (Crawford and Shimkets, 2000b), it is worth revisiting both the magnitude and kinetics of the stringent response in bsgA and other B-signaling mutants.
On the Nature of Extracellular B-Signaling Strains containing the bsgA mutation are described as extracellular B-signaling mutants according to their ability to sporulate when developing in mixtures of wild-type cells, or cells of any of the other extracellular complementation groups (Gill and Cull, 1986; Hagen et al., 1978). However, the nature of the signaling event remains elusive, and for some investigators (Kroos and Kaiser, 1987), somewhat controversial. Previously, Gill and Cull (1986) have shown that strains with a genetically defined null mutation in bsgA
DEVELOPMENT AND MOTILITY (bsgA330) undergo extracellular complementation when developing in mixtures with wild-type cells, which results in the formation of heat- and sonication-resistant spores derived from the bsgA parent (Gill and Cull, 1986). These studies also demonstrated that, for at least a limited number of developmentally induced genes or ZacZ fusions, extracellular complementation by wild-type cells restored developmental gene expression in the bsgA mutant. However, in a larger study, using a more comprehensive collection of developmentally induced ZacZ fusions, Kroos and Kaiser were unable to demonstrate extracellular complementation of either sporulation or developmental gene expression using strains containing the same bsgA330 mutant allele (Kroos and Kaiser, 1987). The discrepancy between the experiences of the two groups of investigators remains unexplained. The critical parameter(s) that must be optimized in order to demonstrate extracellular complementation of bsgA mutants requires additional studies. One clue to understanding the disparate results for extracellular complementation may lie in the observation that the bsgA mutant cells exert a strong inhibitory effect on the development of even wild-type cells (Gill and Cull, 1986; Hagen et al., 1978; Kroos and Kaiser, 1986; R. E. Gill, personal communication). While this may seem contradictory, extracellular complementation can be observed in mixtures containing no more than 50% bsgA mutant cells, and total cell densities up to approximately 2 X l o 7 cells per cm2 (Gill, personal communication). Beyond these limits, the inhibitory effect of the bsgA mutant in the mixtures becomes progressively more dramatic, reducing both the level of developmental gene expression and the production of spores by wild-type cells (Gill, personal communication). Considerable effort has been dedicated to identifying an extracellular or cell-associated signaling molecule from wild-type cells capable of rescuing the development of bsgA mutants. However, neither medium conditioned by developing wild-type cells nor cell envelope preparations of developing wild-type cells rescued the development of bsgA mutants. Taken together with the inhibitory nature of the mutants themselves, it is tempting to question whether bsgA mutants are true signaling mutants. On the other hand, this behavior may be a peculiarity of the bsgA mutants, and not necessarily characteristic of other B-signaling mutants. Efficient extracellular complementation has been demonstrated for certain other members of the B-signaling group with little or no evidence of the dramatic dominant-negative phenotype characteristic of bsgA mutants (Hagen et al., 1978; LaRossa et al., 1983; Gill, personal communication).
AND EARLYDEVELOPMENTAL EVENTS 3. INITIATION
Efforts to discover putative signaling molecules that rescue the development of these strains have not been reported. T h e bsgA Gene Product The bsgA (lonD) gene encodes an intracellular protease that shares approximately 45% amino acid identity with the Lon protease of E. coli (Gill et al., 1993; Tojo et al., 1993b). The cloned bsgA gene was expressed in E. coli, and the purified BsgA protein, like its E. coli homologue, was shown to have ATP-dependent protease activity (Gill et al., 1993). Curiously, M . xanthus also encodes a second Lon-like protease which appears to be required for cell viability (designated lonV) (MXAN2017) (Tojo et al., 1993a). As such, any role lonV may play in development has not been determined. Significantly, however, the finding of two very similar proteases, but with presumed nonredundant functions, suggests that the two proteases have distinct substrate specificities. As substrates are identified, this situation affords an elegant opportunity to examine the determinants of specificity for both the protease and the substrate. The presence of the BsgA protease has been demonstrated in both exponentially growing and developing cells (Gill and Bournemann, 1988; Tojo et al., 199313). This is consistent with the finding that bsgA mutations have an observable effect on the behavior of cells during both the vegetative and developmental phases of the M . xanthus life cycle (Gill and Cull, 1986; Kroos and Kaiser, 1987). Furthermore, the relative level of BsgA protein remains relatively constant during exponential growth and for several hours following the initiation of development-at least up to the onset of aggregation, and well after BsgA has exerted its regulatory effects on developmental gene transcription. Although levels of BsgA protein are relatively constant, changes in the relative activity of the protease upon entry into the developmental program have not been addressed. T h e Search for BsgA Protease Substrates Studies with purified BsgA protease used casein as a suitable substrate to demonstrate in vitro enzymatic activity (Gill et al., 1993). However, using the Lon protease of E. coli as a paradigm (Goldberg, 1992), it is likely that the function of BsgA is to degrade many species of aberrant, improperly folded proteins, as well as certain normal regulatory proteins. It is likely that one or more of the substrate regulatory proteins, rendered unstable and short-lived by the protease, mediates the transcriptional regulation that becomes perturbed in bsgA mutants. For example, BsgA-mediated degradation of a regulatory
63 protein that acts to repress or inhibit transcription of early developmental genes may be required for efficient entry into the developmental program. Both proteomic and genetic approaches have been used to identify possible regulatory substrates of the BsgA protease. Comparative two-dimensional gel electrophoresis of wild-type and bsgA mutant cell extracts failed to reveal reproducible differences between the protein profiles (Gill, personal communication). However, this technique is capable of displaying only a portion of the most abundant part of the proteome expressed at any given time. A genetic approach to identifying BsgA protease substrates has also been reported (Cusick et al., 2002; Hager et al., 2001). The Lon-like proteases are typically characterized by the highly processive and complete degradation of substrates (Goldberg, 1992). The genetic strategy is based on the isolation of suppressor mutations that bypass the developmental requirement for the BsgA protease. Suppressor mutations of this sort may arise in a number of ways, including (i) mutational inactivation of the substrate itself, making the protease dispensable, and (ii) alterations in downstream components of the bsgA regulatory pathway, which are not themselves substrates of the protease. Using this strategy, two bsgA suppressors, spdR (Hager et al., 2001; Tse and Gill, 2002) and bcsA (Cusick and Gill, 2005; Cusick et al., 2002), have been isolated and characterized. While neither appears to be a BsgA substrate, both shed light on the organization of the downstream regulatory pathway.
SpdR The spdR locus encodes a two-component regulatory system, composed of a sensor kinase (SpdS, MXAN1077) and an NtrC-like response regulator or EBP (SpdR, MXAN1078). Null mutations in spdR resulted in the following phenotypes: (i) they bypass the developmental defect in a bsgA mutant (i.e., they restore expression of bsgA-dependent, developmentally induced lacZ gene fusions and restore fruiting body and spore formation); (ii) they result in inappropriate expression of at least one developmental gene, tps (MXAN5432),during vegetative growth; (iii) they develop markedly faster than otherwise isogenic wild-type cells; and (iv) they form fruiting bodies and spores on semirich medium at nutrient concentrations sufficient to inhibit the development of wild-type cells (Hager et al., 2001; Tse and Gill, 2002). These characteristics suggest that SpdR acts as a nutrient sensor and a negative regulator of development, the inhibition being overcome by the BsgA protease or inactivation of SpdR.
64
BcsA The bcsA locus (MXAN3122) encodes a protein homologous to FAD-dependent monooxygenases. Mutations in this gene also suppress the developmental defect of the BsgA protease-deficient mutant, allowing the formation of fruiting bodies and spores. bcsA mutations also restore expression of a subset of late developmental genes to the protease mutant, but apparently bypass some of the regulatory requirements for early development. In addition, these mutants develop significantly faster than wild-type cells when plated on starvation agar at low cell density, similar to the faster density-independent sporulation phenotype of the Che3 cluster mutants. Furthermore, the bcsA mutants were found to complete development on medium containing nutrient levels sufficient to support vegetative growth in wild-type cells (Cusick and Gill, 2005; Cusick et al., 2002). Thus, BcsA appears to also play a role in nutrient sensing and shares characteristics of other nutrient sensors, such as Nsd, members of the Che3 cluster, and SigC. These observations are taken as evidence that the bcsA gene normally acts to inhibit progress through the developmental program. Although mutations in spdR and bcsA both suppress the sporulation defect and restore transcription of subsets of developmentally induced genes in the bsgA mutant, they have distinct and unexpected effects on certain other signaling mutants. Mutations in spdR suppress the developmental defect of bsgA and A-signaling mutants, but not C-signaling mutants. Conversely, mutations in bcsA suppress the developmental defect of bsgA and C-signaling mutants, but not the A-signaling mutants. These data suggest that there are at least two parallel BsgA-dependent regulatory pathways in early development; one that includes spdR and A-signaling and another that includes bcsA and C-signaling. Exciting new data that more clearly define a possible role for the BsgA protease in early developmental gene regulation were recently reported by Nariya and Inouye (2006). These investigators discovered that the transcriptional activator MrpC (MXAN5125) is apparently present in two distinct forms: a relatively less active full-length form and a highly active, truncated form (MrpC2).MrpC2 appears in cell extracts relatively early in development. It is likely to be the form responsible for activating transcription of the downstream transcriptional regulator, FruA (MXAN3117), which is in turn required for continued progression through the developmental program. However, MrpC2 was not detected in a bsgA (ZonD)mutant, where fruA expression is also low. These authors suggest the possibility that the BsgA protease may play an important role in the processing and activation of MrpC (Nariya and Inouye, 2006). BsgA
DEVELOPMENT AND MOTILITY may play a direct role if MrpC is a substrate of the protease. However, such proteolytic cleavage and processing of a substrate, as opposed to complete degradation, is somewhat difficult to reconcile with the typical highly processive activity of the Lon-like proteases. Alternatively, BsgA may be required for expression of the MrpC processing protease or a regulator of its activity.
The A-Signal, a Complex Cell Density Signal The group A mutants, henceforth to be called asg for A-signal generating, all display a similar phenotypic block in the developmental program; these cells produce irregular, loose mounds devoid of myxospores (LaRossa et al., 1983).
The First asg Mutants The initial screens by Hagen et al. (1978) for the isolation of mutants defective in production of extracellular signals identified three distinct loci for the asg mutants designated asgA, asgB, and as&. Recently, two additional asg loci have been reported, asgD (Cho and Zusman, 1999a) and asgE (Garza et al., 2000b). These are discussed in detail in the subsection below. From a historical perspective, considerable work has been done and extensive reviews have been written regarding the identification and characterization of the original three loci (Kaiser and Kroos, 1993; Kaplan and Plamann, 1996). Below is a summary of what is currently known for each of these three genes. asgA (MXAN2670) encodes a hybrid signal transduction protein consisting of a putative N-terminal receiver domain and a C-terminal histidine kinase domain (Plamann et al., 1995). In vitro phosphorylation studies show that AsgA can be autophosphorylated and that the chemical stability pattern is consistent with a histidyl-phosphate. Expression of asgA, using a TnSlac transcriptional fusion, demonstrated that asgA is expressed vegetatively and increases in expression during development approximately twofold (Plamann et al., 1995). asgB (MXAN2913)encodes a putative DNA-binding protein essential for both vegetative growth and development (Plamann et al., 1994). Like asgA, transcriptional fusion studies demonstrate that asgB is expressed during vegetative growth and increases approximately twofold during development. asgC (MXAN5204) encodes the vegetative sigma factor and is an allele of sigA, the rpoD homologue in M . xanthus. Of particular interest is that this rpoDlike gene encodes a modification in a region of RpoD that, in E. coli, alters the interaction with the alarmone (p)ppGpp (Hernandez and Cashel, 1995). Of the three
AND EARLYDEVELOPMENTAL EVENTS 3 . INITIATION
original asg loci, asgC mutants show the weakest developmental phenotypes. Defining and Quantifying the A-Signal As previously discussed, the asg mutants phenotypically resembled mutants of the By C, and D classes (Hagen et al., 1978; LaRossa et al., 1983). To resolve the individual phenotypes, LaRossa et al. (1983) examined the mutants for specific effects on known developmental markers, including (p)ppGpp, protein S, and myxobacterial hemagglutinin (MbhA). These initial experiments provided an approach to temporally rank the various signaling mutants. This work was expanded upon by Kroos et al. (1986) by using a TnSlac reporter system developed in the Kaiser laboratory (Kroos and Kaiser, 1984). With the concurrent advances in the genetic manipulation of 211. xanthus (Kuner and Kaiser, 1981; O’Connor and Zusman, 1983; Shimkets et al., 1983), this TnSlac reporter system provided a systematic approach to determine the temporal organization of the distinct signaling mutants. In addition, it provided a framework to order known genes on the temporal pathway to determine the dependencies of gene expression on the extracellular signaling mutants. Finally, this work directly led to the development of the A-signal bioassay; a biochemical approach to quantify and decipher the molecular identification of the A-signal (Kuspa et al., 1986). The A-signal bioassay takes advantage of the ability of exogenously added A-signal to rescue the asg mutants, whether it is in the form of whole cells, cell extracts, or specific molecules. This provides the researcher a tool to specifically probe the nature of the A-signal. The assay takes advantage of the fact that asg mutants are unable to express the a 4 5 2 1 TnSlac insertion (Kroos and Kaiser, 1984), and expression in this strain is dependent upon exogenously added A-signal. In the assay, an asgB480 mutant (DK4324) carrying the a 4 5 2 1 TnSlac insertion is used as the reporter strain and can be incubated with various exogenous sources of A-signal. A-signal activity is then quantified based on P-galactosidase activity; one A-signal unit is defined as the amount of substance required to produce one unit of P-galactosidase (1nmol of o-nitrophenyl-P-D-galactopyranoside per min for an entire test well containing 1.25 x lo8 cells) above the background in the defined assay condition (Kuspa et al., 1986). With the bioassay in hand, the biochemical identification of the A-signal was under way. Early work supported the hypothesis that two forms of A-signal were present in conditioned media. Approximately 40 to 60% of the A-signal activity was lost when heat treated, implying a heat-labile form (Plamann et al., 1992). In addition,
65
the components of the heat-stable fraction were of low molecular weight, less than 3 kDa in size, while the heatlabile material was much larger. Further analysis led to the discovery that the heat-labile material had proteolytic activity and could be correlated to at least two M. xanthus proteins, a 27- and a 10-kDa protein. Biochemical characterization of these proteases demonstrated different specificities. In addition, Plamann et al. (1992) demonstrated that exogenous proteases such as pronase, proteinase K, or trypsin also have A-signal activity, leading to the hypothesis that the heat-labile activity is due to the release of proteases by the cell during early development (Plamann et al., 1992). This model predicted that a product or products of these general proteases should also have A-signal activity. Simultaneous work by Kuspa et al. (1992a) demonstrated that the heatstable fraction of A-signal was in fact amino acids and that six amino acids, Tyr, Pro, Phe, Trp, Leu, and Ile, made up the majority of the heat-stable activity. It was also found that small peptides containing those amino acids possessed A-signal activity (Kuspa et al., 1992a). Consistent with the notion of amino acids acting as a cell-density monitor, Kuspa et al. (1992b) further demonstrated a proportional response between A-signal amino acids and the cell number, through the range of 5 X l o 8to 2.5 X 1O1O cells per ml. Therefore, Kuspa et al. (1992b) provided evidence of a “threshold” level of A-signal amino acids required to elicit the A-signal-dependent response. These data led to the hypothesis that during early development, cells release a cocktail of proteases that degrade peptides and proteins in the external milieu to amino acids and small peptides that in turn act as the A-signal.
Models for the Quorum-Sensing A-Signaling System This early work with the identification of three asg loci and the biochemical and physiological characterization of the nature of the A-signal provides overwhelming support for a model whereby the A-signal acts as cell density monitor, functionally equivalent to homoserine lactones used in many other gram-negative species (Engebrecht et al., 1983; Fuqua et al., 1996). In addition, researchers have been unable to demonstrate any role for homoserine lactones in M . xanthus development (H. B. Kaplan, personal communication; W. Shi, personal communication), nor have homoserine lactone production genes been identified in the newly released 211. xanthus genome sequence (Goldman et al., 2006). The quorum-sensing model proposes that once cells recognize starvation, they release extracellular proteases that begin to degrade proteins on their outer surface, liberating amino acids and
DEVELOPMENT AND MOTILITY
66 peptides into their environment. These amino acids and peptides would be enriched for the six most active amino acids. As more and more cells begin to release proteases, the concentration of A-signal amino acids would rise until a critical concentration is achieved, at which point the cells respond by activating gene expression. In this general quorum-sensing model, the activation and/ or release of the A-signal proteases is dependent upon individual cells recognizing and responding to starvation signals. Mutations that disrupt the recognition of cellular starvation or the production of the proteases would result in a block in A-signal production.
A-Signaling as a Linear Phosphorelay System Based on the molecular analysis of AsgA and AsgB, Kaplan and Plamann (1996) have proposed that AsgA and AsgB interact in a signal transduction phosphorelay, analogous to the B. subtilis phosphorelay (Burbulys et al., 1991; Hoch, 1993), where AsgA lies upstream of the signaling cascade to AsgB (Fig. 4). The pathway is activated by the cellular starvation system that leads to the phosphorylation of AsgA, which eventually transfers the phosphate to AsgB. AsgB is proposed to act as a repressor in its unphosphorylated form, and as either an activator or an inactive repressor in its phosphorylated form (AsgB-P). AsgB-P, in conjunction with SigA holoenzyme (as& is an allele of sigA, i.e., the M. xanthus rpoD homologue), activates the genes required for Asignal production. This linear phosphorelay model also predicts that there are still several missing components in the relay. The linear phosphorelay model (Fig. 4) is a very attractive model, and although it was formally proposed in 1996 by Kaplan and Plamann (1996), it has been the working hypothesis on the nature of the A-signal since its conception in the early 1990s. However, there are
several pieces of data that suggest that the nature of the A-signal and its regulation may be more complex. First, as Kaplan and Plamann (1996) point out, several predicted key components (such as the intermediary AsgX) are missing. This has led to the active pursuit of additional asg genes. Second, while proteases are able to restore nearly wild-type sporulation efficiencies to the asgB and as& mutants, the rescue of the sporulation defect of the asgA mutant was minimal (Kuspa et al., 1992a; Plamann et al., 1992). This suggests that while proteases and amino acids are critical for M. xanthus development and A-signaling, it cannot account for all of the A-signaling defects. Third, the asgA phenotype was reported to be much more severe than either the asgB or asgC phenotypes (Kuspa and Kaiser, 1989; Kuspa et al., 1992b).This is surprising based on a simple linear signal transduction pathway for these three genes. Finally, two additional asg genes have been identified based on extracellular complementation, response to the addition of A-signal components, and their A-signal bioassay phenotypes. These genes, asgD and asgE, do not display all of the same phenotypic characteristics initialIy used to define the asgA, asgB, and asgC mutants by Kuspa and colleagues (Kuspa and Kaiser, 1989) or intuitively fit into this simple phosphorelay scheme. This suggests that the A-signal and its regulation are much more complex than previously thought.
Two New asg Loci, asgD and asgE The asgD gene encodes a hybrid histidine protein kinase. AsgD is similar in domain organization to the AsgA protein, but with a large, 3 80-amino-acid long central region between the N-terminal receiver and C-terminal histidine kinase domains. AsgD is proposed to act as a nutrient sensor (as described previously) and as a component of the A-signaling complex. SigA (As&) ~
(PIPPGPP
--
AsgA
AsgX-P
AsgB
-b
AsgA-P
AsgX
AsgB-P
I I I
------
+
A-signal Proteases
Figure 4 Linear phosphorelay model of A-signaling in M. xanthus. Linear model of A-signal production by a phosphorelay (Kaplan and Plamann, 1996). In this model, AsgA recognizes a rise in (p)ppGpp levels, which leads to the phosphorylation of AsgA and initiates a phosphorelay that ultimately activates the production of the A-signal proteases, which in turn produce the A-signal amino acids, collectively known as A-signal. This model predicts the existence of a histidine phosphotransfer protein, designated AsgX as an intermediary between AsgA-P and AsgB. Phosphorylated AsgB in conjunction with SigA ( AsgC) activates expression of the A-signal proteases. Dashed lines represent predicted, yet mechanistically unknown interactions.
3. INITIATION AND EARLY DEVELOPMENTAL EVENTS The developmental defects of the asgD mutant cannot be complemented by the asg mutants: asgA, as@, or as&. Kuspa et al. (1992b) have shown that AsgB mutants can be rescued by the addition of A-signal amino acids. Addition of specific L-amino acids can rescue development of the asgD mutant, with leucine being the most effective, but some A-signal amino acids can actually inhibit its development (Cho and Zusman, 1999a). The recognition and response to the individual A-signal amino acids by the asgD mutant are varied. Based on these data, the A-signal amino acids can be separated into two groups based on the rescue of the asgD mutant. Moreover, asgD is developmentally expressed. In asgB mutants, asgD expression is lowered by 50%, indicating that asgD expression is partially dependent on AsgB (Cho and Zusman, 1999a). Interestingly, usgD mutants produce wild-type levels of a4521 (spi)(Cho and Zusman, 1999a). This gene has been shown to be absolutely dependent on A-signal for its expression, and this is one reason why it is used as a reporter for the A-signal bioassay. Based on the characterization of the asgD mutant, Cho and Zusman (1999a) suggest that inactivation of asgD may interfere with only a part of the A-signaling process and that AsgD may be acting downstream of a subgroup of A-signal amino acids to monitor nutritional conditions. The most recent addition to the A-signaling pathway is AsgE. Sequence analysis of asgE predicts that it encodes a protein with two membrane-spanning domains and has closest homology to a family of aminohydrolases. Strains carrying the asgE mutation are defective in the compaction of aggregates into mature fruiting bodies and in sporulation. These mutants can be rescued by codevelopment with wild-type cells, csgA cells, and esg cells,
I
I
SigA (AsgC)
AsgA
I
+
I
AsgE
I
Other
- - - - -I
b AsgD
67 but not asgA cells (Garza et al., 2000b). This suggests that the asgE mutants may be defective in production of A-signal, and this was supported by the A-signal bioassay data. Although not as severe as the asgA mutant, asgE cells produce only 30 to 50% of wild-type A-signal, based on restoration of spi (R4521)expression. The level of A-signal produced by asgE mutants is 1.5- to 2.5-fold higher than the level in usgA cells (Garza et al., 2000b). Further characterization of usgE cells revealed that the primary defect is in the production of heat-labile A-signal (Garza et al., 2000b); asgE cells produce 10-fold less of the wild-type levels. In contrast, usgE mutant cells are only down twofold for the production of heat-stable A-signal compared to wild-type cells. Most interestingly, asgE cells are not rescued by the addition of exogenous proteases, including pronase. This implies that while asgE cells are defective in a heat-labile component of A-signal, the defect of the asgE mutant is more intricate than a simple lack of an A-signal protease. A-Signaling as a Network of Several Signals With the identification and characterization of the asgD and asgE mutants and a reexamination of the earlier asgABC complementation data, a more complicated picture regarding the nature of A-signaling has emerged. Garza et al. (2000b) proposed that A-signaling may be a mixture of signals: specifically, the previously identified set of six amino acids and associated secreted proteases may represent one of several signals produced concurrently during early (0 to 6 h postinitiation) development. This alternative model (diagrammed in Fig. 5) proposes that AsgA sits on the top of a signaling hierarchy and acts as a master control protein regulating these early signaling systems in combination with signals from the
Proteases -b Amino Acids Peptides
Heat Labile Az-Signal
-
’
Collectively Quorum sensing A-signal
Figure 5 The network model of A-signaling in M. xanthus. An alternative model whereby AsgA recognizes a rise in (p)ppGpp levels and sits on top of a hierarchy of genes that are required to activate a variety of signals that collectively make up the quorum-sensing system of M. xanthus. In the simplest model, AsgA is required for all components of the A-signaling system. Alternatively, there may be requirements for other starvation signals [in addition to (p)ppGpp] or additional hierarchical regulators (like AsgD) that either directly or indirectly activate the system. These inputs are represented by dashed lines.
68
cellular starvation recognition system, such as (p)ppGpp and AsgD. AsgB and AsgE represent two parallel signaling systems, dependent upon input from AsgA but independent from each other, where AsgB controls the production or release of extracellular proteases and AsgE controls some other, not yet defined signal. This model could also explain the regulation of other proposed early signals such as an aggregation signal proposed by Kuspa (1989).In addition, H. B. Kaplan (personal communication) has recently reported the identification of another early developmental signal. There are several pieces of data that support a multiple signal hypothesis. First, asgA mutants are not rescued by proteases, suggesting there is a secondary block, although these cells are lacking heat-labile A-signal (Plamann et al., 1992) and they are poorly rescued by amino acids (Plamann et al., 1992). Although Kuspa et al. (1992a) suggest that this could be due to the cells being sensitive to the “quality” of the A-signal amino acids, whereby cells are responding to the concentration of the various activating and inhibitory amino acids generated, this does not sufficiently explain the asgD and asgE mutant phenotypes (Cho and Zusman, 1999a; Garza et al., 2000a, 2000b). Second, asgE mutants are defective in a heat-labile fraction of A-signal, yet they are not rescued by the addition of extracellular proteases (Garza et al., 2000b), suggesting they are missing a different heat labile component. Extracellular complementation tests also demonstrate that asgE can rescue asgB mutants and will rescue asgA mutants to the level of an asgE mutant (Garza et al., 2000b). However, the asgE mutant showed no increase in sporulation efficiency when codeveloped with the asgA mutant. Thus, AsgE appears to function downstream of AsgA in the A-signal production pathway (Garza et al., 2000a, 2000b). Taken together, these data suggest that asgE and asgB mutants are defective in different extracellular components of A-signal and that asgE mutants (as@ mutants have not been tested) can supply asgA mutants with all of the missing “A-signal components” except the asgE-dependent heatlabile component. Finally, predation studies with asg mutants on Serratia marcescens lawns show that asgA, a&, and asgE cells are defective in predation; each had zones of lysis ranging from 20.8 to 36.8% of wild-type levels (Pham et al., 2005a). In contrast, the asgB mutant cells had a wild-type predation phenotype on S. marcescens, suggesting a more complex A-signaling network than a linear phosphorelay system.
A-Signal Reception and Regulation Regardless of the nature of the A-signal, whether it is a complex mixture of signaling molecules or simply a
DEVELOPMENT AND MOTILITY collection of amino acids, cells respond by activating a specific set of genes, including spi (Kroos et al., 1986).As previously stated, the activation of spi (04521 TnSlac) by A-signal has allowed for the development of the Asignal bioassay and has been used to begin to dissect the mechanism of A-signal reception. Mutants able to express the a 4 5 2 1 TnSlac fusion in an asgB background have led to the identification of several genes postulated to be involved in A-signal reception. These mutations were designated Sas, for suppressors of asg. Sas mutants are able to express the 04521 fusion independently of A-signal. Two classes of mutants were identified: sasB mutants, which express 04521 both vegetatively and during development, and sasA mutants, which express 04521 independent of A-signal yet still require starvation for activation. Both classes were initially generated by W mutagenesis. Although the sas mutants were originally isolated in an asgB background, in all cases they render expression of a 4 5 2 1 independent of A-signal in asgA and asgC strains as well (Kaplan et al., 1991).This implies that the sas genes can bypass the A-signaling requirement regardless of the nature of A-signal itself. Recently several reviews have been written on the isolation and role of Sas in A-signaling (Kaplan and Plamann, 1996), and these past findings are summarized here. Additionally, work by S~gaard-Andersenand colleagues has identified a two-component system kinase, RodK, that has been implicated in A-signal regulation (Rasmussen et al., 2005).
SasB The sasB locus identified several putative regulatory proteins, including sass, sasR, and sasN (Kaplan et al., 1991). In the case of the sasB mutants, bypass of A-signal recognition is still dependent upon starvation. The six suppressor mutants that map to the sasB locus comprise three regulatory genes: sass (MXAN1249), sasR (MXAN1245),and sasN (MXAN1244) (Xu et al., 1998; Yang and Kaplan, 1997). The product of the sass gene has been shown to be essential in the reception of A-signal, and sequence analysis predicts it to be a transmembrane sensor histidine kinase typical of the type found in two-component signal transduction pathways (Yang and Kaplan, 1997).The N terminus consists of an input domain and two transmembrane domains similar to the N-terminal domains of methyl-accepting chemotactic proteins (Yang and Kaplan, 1997).The sasR gene is predicted to encode an NtrC-like activator protein or EBP (Kaufman and Nixon, 1996) and along with Sass acts as positive activator of 0 4 5 2 1 (Yang and Kaplan, 1997).This is supported by the phenotypes of null mutations in each of these genes. Yang and Kaplan (1997)
3 . INITIATION AND EARLYDEVELOPMENTAL EVENTS suggest that Sass and SasR represent a signal transduction pathway whereby Sass recognizes and responds to A-signal amino acids (peptides) and activates SasR by phosphorylation, which then activates gene expression. The third gene, sasN, lies just 5’to sasR and has no similarity to other proteins in the GenBank database (Xu et al., 1998).Mutations in sasN confer starvation and Asignal-independent expression of 04521, implying that SasN is a negative regulator with a role in the inhibition of 04521 expression (Xu et al., 1998). It has been suggested that SasN may “sense” the intracellular availability of A-signal amino acids and their levels dictate the activity of SasN (Xu et al., 1998). When the levels are low, SasN represses, and when levels are high, SasN is inactive, thus allowing for the SasSR system to function. Whether SasN acts directly on the 04521 promoter or indirectly through Sass and/or SasR is unclear. However, the role of these proteins is critical for the recognition of A-signal.
SasA Mutations in the sasA locus cause defective fruiting body formation, reduced sporulation, and the restoration of A-signal-dependent a 4 5 2 1 expression in the absence of A-signal. The wild-type sasA locus was sequenced, and three open reading frames were identified with sequence similarity to proteins involved in lipopolysaccharide (LPS) O-antigen biosynthesis. The first two genes, designated rfbA (MXAN4623)and rfbB (MXAN4622),show sequence similarity to the integral membrane domains and ATPase domains of the ATP-binding cassette (ABC) transporter required for the biosynthesis of LPS O-antigen in certain gram-negative bacteria (Guo et al., 1996). The third gene, rfbC (MXAN4621), encodes a predicted protein with similarity to the Yersinia enterocolitica 0 3 rfbH gene product, which is also required for O-antigen biosynthesis (Guo et al., 1996). As expected, sasA mutants are defective in LPS production (Kaplan et al., 1991). In addition, sasA mutants are defective in social motility, leading Kaplan and colleagues to suggest that along with pili and fibrils, LPS O-antigen represents a third cell surface component required for social motility (Guo et al., 1996). The identification of LPS O-antigen in A-signal reception was surprising, and the question of the role and mechanism by which LPS O-antigen is involved in Asignal reception remains today. Of particular interest is that while mutations in sasA bypass the requirement for A-signal, they still require starvation for activation and unlike mutations in s a d , sasA mutants are defective for development (Kaplan et al., 1991). One explanation is that the overall structure of the outer membrane
69 is important for A-signal reception and mutations in LPS O-antigen mimic the changes that occur during the A-signaling process, thereby bypassing the requirement for A-signal. This would imply that the role of LPS O-antigen is indirect. Future studies will be required to clarify the role of LPS O-antigen in S-motility and A-signal reception.
RodK RodK (MXAN0733) acts as a developmental timer that is important for the spatial coupling of the aggregation and sporulation processes of development (Rasmussen et al., 2005, 2006). A mutation in rodK results in increased A-signal production. When asgB cells are codeveloped with ArodK cells in submerged culture, the spi (fl4.521) P-galactosidase activity is 1.5- to 2-fold higher after 6 to 24 h than asgB cells codeveloped with wild-type cells (Rasmussen et al., 2005). Thus, intriguingly, RodK appears to be involved in the inhibition of A-signal production late in development. In addition, there is an A-signal-dependent decrease in RodK accumulation after 12 h of development in wild-type cells (Rasmussen et al., 2005). Since A-signal is produced at about 2 h into development and the decrease in the accumulation of RodK is not seen until much later, RodK accumulation may be regulated by a member of the A-signal regulatory pathway instead of the actual A-signal itself.
INTEGRATION OF THE CELLULAR AND POPULATION RECOGNITION PATHWAYS The initial developmentalchallenge for M . xanthus is to recognize the onset of nutrient limitation, such that cells will have a carbon and energy reserve sufficient for the completion of development. In addition, cells must monitor and respond to two different starvation levels, that of the individual cell and that of the population. This is accomplished by having two parallel pathways, one in response to cellular starvation and the other responding to population cues. The latter includes the quorum-sensingA-signaling system, as well as the contact-dependent C-signaling which occurs during the aggregation phase at 6 to 8 h postinitiation. The cellular starvation pathway includes the activation of the stringent response and the complex regulatory systems that both modulate (p)ppGpp levels and respond to them. The dual requirement for cellular and population starvation recognition by the developing M. xanthus cells has led to the creation of several models that focus on two parallel pathways converging at or around the aggregation phase of development (Kuspa, 1989; Singer and Kaiser, 1995). Figure 6 provides an updated model that encompasses new data over the past 10 years.
DEVELOPMENT AND MOTILITY
70 Population Starvation Response A-Signal
1
C-signal
7
I
4 A-signal reception
C-signal production
C-signal reception
FruA
Motility
I
Starvation I
I
Nsd
todK rodK
I
T
nsd
TodK RodK
A
4
I
Cellular Starvation Response
Figure 6 Model for dual starvation in M. xanthus. Schematic of major players, identified to date, that modulate entrance into the developmental process by monitoring and responding to nutrient levels. Proteins are identified in boldface type, and genes are in italics; direct interactions are represented by solid lines, presumed indirect interactions are shown as dashed lines, and proposed interactions are indicated by a dotted line and a question mark (?). Arrowheads indicate a positive interaction, and a blunt head indicates a negative interaction. Note: for simplicity, some important components of this process are not included in this model; thus, it is not all-inclusive. For more details, see text.
Briefly, vegetatively growing cells maintain homeostasis and balanced growth. Two integral regulatory pathways involved in maintaining balanced growth are the nZal8 and nlu4 regulatory systems. Mutations in either of these two genes disrupt balanced growth and prevent cells from launching a productive developmental response. Development is initiated when cells are starving for amino acids. 211. xunthus cells evaluate their nutritional status by monitoring their translational capacity-a mechanism that allows cells to couple their basic metabolism to development. As individual cells perceive nutrient limitation, (p)ppGpp levels rise due to activation of the stringent response. This response is modulated by a variety of other gene products including Nsd, SocE, and CsgA (Brenner et al., 2004; Crawford and Shimkets, 2000b). RelA activity and/or the
intracellular level of (p)ppGpp itself is positively regulated by Nla18, Nla4, and CsgA and negatively regulated by SocE and Nsd (Brenner et al., 2004; Crawford et al., 2000a, 2000b; Diodati et al., 2006; Ossa et al., unpublished). (p)ppGpp inhibits socE and stimulates csgA transcription (Crawford and Shimkets, 2000a). The intricate balance of SocE and CsgA is critical for maintaining the stringent response during development. Shd, based on its homology to the hydrolytic domain of E. coli SPOT,may modulate (p)ppGpp levels by affecting the nucleotide’s stability. Overall, increased levels of (p)ppGpp activate and repress a wide variety of genes involved in the general stress response and the developmental process. These alterations in gene expression are initially modulated by RNA polymerase, SigA ( AsgC), presumably in conjuction with DksA.
AND EARLYDEVELOPMENTAL EVENTS 3 . INITIATION
Once (p)ppGpp levels rise, two parallel pathways in the developmental process become activated: the cellular response and the population response. The population response includes the activation of the asg genes and the production of A-signal, as well as the expression of csgA and fruA which are required for the production and reception of C-signal (Harris et al., 1998).The SasRS system is involved in the reception of A-signal and regulates spi and fruA expression along with the inhibitor, SasN (Xu and Hoover, 2001; Yang and Kaplan, 1997). The products of the actABCDE operon are involved in a Csignal positive feedback loop that regulates the level and timing of C-signal (Gronewold and Kaiser, 200 1).Taken together, these processes lead to the propagation of the developmental response in concert with other cells. The second starvation response is at the level of the individual cell and includes the activation of genes that act independently of extracellular signals. The sigD gene, which encodes the M . xanthus RpoS homologue, is regulated by (p)ppGpp (Viswanathan et al., 2006). MrpABC acts downstream of (p)ppGpp and partially regulates the developmentally essential histidine kinase, SdeK (Sun and Shi, 2001a). SdeK is involved in the cellular response and acts in conjunction with BrgE (Pham et al., 2005b) to coordinate the individual cell with the starving population. In addition, both SdeK and BrgE inhibit nsd expression, thereby contributing to the maintenance of the higher (p)ppGpp levels needed for development to continue. The population and cellular starvation branches converge at the aggregation stage, approximately 6 to 8 h postinitiation. Downstream developmental genes that eventually lead to fruiting and sporulation, such as devTRS, 04400,04403,04406, and 04435, are dependent on extracellular signals, FruA and SdeK for their full expression (Kroos et al., 1986; Pollack and Singer, 2001). Finally, alternate yet undefined starvation signals activate TodK and RodK, which modulate both the spatial and temporal aspects of sporulation (Rasmussen et al., 2005; Rasmussen and Sargaard-Andersen, 2003). These two parallel pathways may serve as a checkpoint, to allow synchronous entry into the developmental program at the aggregation stage. This proposed synchrony could be at one or several levels, including cell cycle or physiological state. Cells may need to express a certain set of early genes prior to continuing through development, such as those required for responding to the Csignal. This ability to recognize and respond to both cellular and population starvation allows M . xanthus to coordinate complex behaviors and leads to the eventual construction of a multicellular fruiting body and the differentiation of vegetative rods into environmentally resistant and metabolically quiescent myxospores.
71
References Alderwick, L. J., V. Molle, L. Kremer, A. J. Cozzone, T. R. Daffron, G. S. Besra, and K. Futterer. 2006. Molecular structure of EmbR, a response element of Ser/Thr kinase signaling in Mycobacterium tuberculosis. Proc. Natl. Acad. Sci. USA 103:2558-2563. Apelian, D., and S. Inouye. 1993. A new putative sigma factor of Myxococcus xanthus. J. Bacteriol. 175:3335-3342. Aravind, L., and E. V. Koonin. 1998. The HD domain defines a new superfamily of metal-dependent phosphohydrolases. Trends Biochem. Sci. 23:469-472. Aravind, L., and C. P. Ponting. 1997. The GAF domain: an evolutionary link between diverse phototransducing proteins. Trends Biochem. Sci. 22:458-459. Avarbock, D., A. Avarbock, and H. Rubin. 2000. Differential regulation of opposing RelMtb activities by the aminoacylation state of a tRNA.ribosome.mRNA.RelMtbcomplex. Biochemistry 39:11640-11648. Block, R., and W. A. Haseltine. 1974. In vitro synthesis of ppGpp and pppGpp, p. 747-761. In M. Nomura, A. Tissieres, and P. Lengyel (ed.), Ribosomes. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. Branny, P., J. P. Pearson, E. C. Pesci, T. Kohler, B. H. Iglewski, and C. Van Delden. 2001. Inhibition of quorum sensing by a Pseudomonas aeruginosa dksA homologue. J. Bacteriol. 183:1531-15 39. Brenner, M., A. G. Garza, and M. Singer. 2004. nsd, a locus that affects the Myxococcus xanthus cellular response to nutrient concentration. J. Bacteriol. 186:3461-3471. Bretscher, A. P., and D. Kaiser. 1978. Nutrition of Myxococcus xanthus, a fruiting myxobacterium. J . Bacteriol. 133:763768. Brown, L., D. Gentry, T. Elliott, and M. Cashel. 2002. DksA affects ppGpp induction of RpoS at a translational level. J. Bacteriol. 184:4455-4465. Brun, Y. V., and L. Shapiro. 1992. A temporally controlled sigma factor is required for polar morphogenesis and normal cell division in Caulobacter. Genes Dev. 6:1395-1408. Burbulys, D., K. A. Trach, and J. A. Hoch. 1991. The initiation of sporulation in Bacillus subtilis is controlled by a multicomponent phosphorelay. Cell 64545-552. Caberoy, N. B., R. D. Welch, J. S. Jakobsen, S. C. Slater, and A. G. Garza. 2003. Global mutational analysis of NtrC-like activators in Myxococcus xanthus: identifying activator mutants defective for motility and fruiting body development. J. Bacteriol. 185:6083-6094. Campos, J., and D. Zusman. 1975. Regulation of development in Myxococcus xanthus: effect of CAMP, AMP, ADP, and nutrition. Proc. Natl. Acad. Sci. USA 72518-522. Cashel, M., and J. Gallant. 1969. Two compounds implicated in the function of the RC gene in Escherichia coli. Nature 221~838-841. Cashel, M., D. Gentry, J. Hernandez, and D. Vinella. 1996. The stringent response, p. 1458-1496. In F. C. Neidhardt (ed.), Escherichia coli and Salmonella: Cellular and Molecular Biology, 2nd ed., vol. 1. ASM Press, Washington, DC. Cashel, M., and K. E. Rudd. 1989. The stringent response, p. 1410-1438. In F. C. Neidhardt, K. B. Low, B. Magasanik,
72 M. Schaechter, and H. E. Umbarger (ed.), Escherichia coli and Salmonella: Cellular and Molecular Biology, vol. 2. ASM Press, Washington, DC. Chakraburtty, R., J. White, E. Takano, and M. Bibb. 1996. Cloning, characterization and disruption of a (p)ppGppsynthetase gene (relA) of Streptomyces coelicolor A3(2). Mol. Micro biol. 19:357-368. Chatterji, D., and A. K. Ojha. 2001. Revisiting the stringent response, ppGpp and starvation signaling. Curr. Opin. Microbiol. 4: 160-1 65. Cho, K., A. Treuner-Lange, K. A. O’Connor, and D. R. Zusman. 2000. Developmental aggregation of Myxococcus xanthus requires frgA, an frz-related gene. J. Bacteriol. 182~6614-6621. Cho, K., and D. R. Zusman. 1999a. AsgD, a new two-component regulator required for A-signalling and nutrient sensing during early development of Myxococcus xanthus. Mol. Microbiol. 34:268-28 1. Cho, K., and D. R. Zusman. 1999b. Sporulation timing in Myxococcus xanthus is controlled by the espAB locus. Mol. Microbiol. 34:714-725. Clifton, S. W., D. McCarthy, and B. A. Roe. 1994. Sequence of the rec-2 locus of Haemophilus influenzae: homologies to comE-ORF3 of Bacillus subtilis and msbA of Escherichia coli. Gene 146:95-100. Cochran, J. W., and R. W. Byrne. 1974. Isolation and properties of a ribosome-bound factor required for ppGpp and pppGpp synthesis in Escherichia coli. J. Biol. Chem. 249:353-360. Crawford, E. W., Jr., and L. J. Shimkets. 2000a. The Myxococcus xanthus socE and csgA genes are regulated by the stringent response. Mol. Microbiol. 37:788-799. Crawford, E. W., Jr., and L. J. Shimkets. 2000b. The stringent response in Myxococcus xanthus is regulated by SocE and the CsgA C-signaling protein. Genes Dev. 14:483-492. Cusick, J. K., and R. E. Gill. 2005. The bcsA gene influences multiple aspects of development in Myxococcus xanthus. Curr. Microbiol. 51:336-343. Cusick, J. K., E. Hager, and R. E. Gill. 2002. Characterization of bcsA mutations that bypass two distinct signaling requirements for Myxococcus xanthus development. ]. Bacteriol. 184:5141-5150. Diodati, M. E., F. Ossa, N. B. Caberoy, I. R. Jose, W. Hiraiwa, M. M. Igo, M. Singer, and A. G. Garza. 2006. Nla18, a key regulatory protein required for normal growth and development of Myxococcus xanthus. ]. Bacteriol. 188:1733-1743. Doetsch, R. N., and G. J. Hageage. 1968. Motility in prokaryotic organisms: problems, points of view and perspectives. Biol. Rev. Camb. Philos. SOC.43:317-362. Downard, J., S. V. Ramaswamy, and K. S. Kil. 1993. Identification of esg, a genetic locus involved in cell-cell signaling during Myxococcus xanthus development. ]. Bacteriol. 175:7762-7770. Dworkin, M. 1973. Myxobacterales, p. 191-202. In A. I. Laskin and H. A. Chevalier (ed.), Handbook of Microbiology. CRC Press, Boca Raton, FL. Dworkin, M. 1963. Nutritional regulation of morphogenesis in Myxococcus xanthus. J. Bacteriol. 86:67-72. Dworkin, M. 1962. Nutritional requirements for vegetative growth of Myxococcus xanthus. J. Bacteriol. 84:250-257.
DEVELOPMENT AND MOTILITY Dworkin, M. 1996. Recent advances in the social and developmental biology of the myxobacteria. Microbiol. Rev. 60~70-102. Dworkin, M., and D. Kaiser. 1993. Myxobacteria 11.American Society for Microbiology, Washington, DC. Engebrecht, J., K. Nealson, and M. Silverman. 1983. Bacterial bioluminescence: isolation and genetic analysis of functions from Vibrio fischeri. Cell 32:773-781. Fiil, N. P., B. M. Willumsen, J. D. Friesen, and K. von Meyenbur. 1977. Interaction of alleles of the relA, relC, and SPOT genes in Escherichia coli: analysis of the interconversion of GTP, ppGpp and pppGpp. Mol. Gen. Genet. 150:87-101. Fuqua, C., S. C. Winans, and E. l? Greenberg. 1996. Census and consensus in bacterial ecosystems: the LuxR-Lux1 family of quorum-sensing transcriptional regulators. Annu. Rev. Microbiol. 50:727-75 1. Garza, A. G., B. Z. Harris, B. M. Greenberg, and M. Singer. 2000a. Control of asgE expression during growth and development of Myxococcus xanthus. J. Bacteriol. 182:66226629. Garza, A. G., B. Z. Harris, J. S. Pollack, and M. Singer. 2000b. The asgE locus is required for cell-cell signalling during Myxococcus xanthus development. Mol. Microbiol. 35: 8 12-824. Garza, A. G., J. S. Pollack, B. Z. Harris, A. Lee, I. M. Keseler, E. F. Licking, and M. Singer. 1998. SdeK is required for early fruiting body development in Myxococcus xanthus. J. Bacteriol. 180:4628-4637. Gill, R. E., and M. G. Cull. 1986. Control of developmental gene expression by cell-to-cell interactions in Myxococcus xanthus. J. Bacteriol. 168:341-347. Gill, R. E., and C. Bournemann. 1988. Identification and characterization of the Myxococcus xanthus bsgA gene product. J. Bacteriol. 1705289-5297. Gill, R. E., M. G. Cull, and S. Fly. 1988. Genetic identification and cloning of a gene required for developmental cell interactions in Myxococcus xanthus. J. Bacteriol. 1705279-5288. Gill, R. E., M. Karlok, and D. Benton. 1993. Myxococcus xanthus encodes an ATP-dependent protease which is required for developmental gene transcription and intercellular signaling. J. Bacteriol. 175:4538-4544. Givens, R. M., M. H. Lin, D. J. Taylor, U. Mechold, J. 0.Berry, and V. J. Hernandez. 2004. Inducible expression, enzymatic activity, and origin of higher plant homologues of bacterial RelNSpoT stress proteins in Nzcotiana tabacum. J. Biol. Chem. 279:7495-7504. Goldberg, A. L. 1992. The mechanism and functions of ATPdependent proteases in bacterial and animal cells. Eur. J. Biochem. 203:9-23. Goldman, B. S., W. C. Nierman, D. Kaiser, S. C. Slater, A. S. Durkin, J. A. Eisen, C. M. Ronning, W. B. Barbazuk, M. Blanchard, C. Field, C. Halling, G. Hinkle, 0. Iartchuk, H. S. Kim, C. Mackenzie, R. Madupu, N. Miller, A. Shvartsbeyn, S. A. Sullivan, M. Vaudin, R. Wiegand, and H. B. Kaplan. 2006. Evolution of sensory complexity recorded in a myxobacterial genome. Proc. Natl. Acad. Sci. USA 103:15200-15205. Gorski, L., T. Gronewold, and D. Kaiser. 2000. A us4 activator protein necessary for spore differentiation within the fruiting body of Myxococcus xanthus. J. Bacteriol. 182:2438-2444.
AND EARLYDEVELOPMENTAL EVENTS 3. INITIATION
Gorski, L., and D. Kaiser. 1998. Targeted mutagenesis of aS4 activator proteins in Myxococcus xanthus. J. Bacteriol. 180~5896-5905. Gronewold, T. M., and D. Kaiser. 2001. The act operon controls the level and time of C-signal production for Myxococcus xanthus development. Mol. Microbiol. 40:744-756. Gronewold, T. M., and D. Kaiser. 2002. act operon control of developmental gene expression in Myxococcus xanthus. J. Bacteriol. 184:1172-1179. Guo, D., M. G. Bowden, R. Pershad, and H. B. Kaplan. 1996. The Myxococcus xanthus rfbABC operon encodes an ATP-binding cassette transporter homolog required for 0-antigen biosynthesis and multicellular development. J. Bacteriol. 178:1631-1639. Guo, D., Y. Wu, and H. B. Kaplan. 2000. Identification and characterization of genes required for early Myxococcus xanthus developmental gene expression. J. Bacteriol. 182~4564-4571. Hagen, D. C., A. P. Bretscher, and D. Kaiser. 1978. Synergism between morphogenetic mutants of Myxococcus xanthus. Dev. Biol. 64:284-296. Hager, E., H. Tse, and R. E. Gill. 2001. Identification and characterization of spdR mutations that bypass the BsgA protease-dependent regulation of developmental gene expression in Myxococcus xanthus. Mol. Microbiol. 39:765-780. Harris, B. Z., D. Kaiser, and M. Singer. 1998. The guanosine nucleotide (p)ppGpp initiates development and A-factor production in Myxococcus xanthus. Genes. Dev. 12:10221035. Haseltine, W. A., and R. Block. 1973. Synthesis of guanosine tetra- and pentaphosphate requires the presence of a codonspecific, uncharged transfer ribonucleic acid in the acceptor site of ribosomes. Proc. Natl. Acad. Sci. USA 70:15641568. Hemphill, H. E., and S. A. Zahler. 1968. Nutritional induction and suppression of fruiting in Myxococcus xanthus FB. J. Bacteriol. 9 5 1 0 18-1023. Hengge-Aronis, R. 2002. Signal transduction and regulatory mechanisms involved in control of the us(RpoS) subunit of RNA polymerase. Microbiol. Mol. Biol. Rev. 66:373-395. Hernandez, J., and M. Cashel. 1995. Changes in conserved region 3 of Escherichia coli a 7 0 mediate ppGpp-dependent function in vivo. J. Mol. Biol. 2 5 2 5 3 6 4 4 9 . Higgs, P. I., K. Cho, D. E. Whitworth, L. S. Evans, and D. R. Zusman. 2005. Four unusual two-component signal transduction homologs, RedC to RedF, are necessary for timely development in Myxococcus xanthus. J. Bacteriol. 187:8191-8195. Hobbs, M., E. S. Collie, P. D. Free, S. P. Livingston, and J. S. Mattick. 1993. PilS and PilR, a two-component transcriptional regulatory system controlling expression of type 4 fimbriae in Pseudomonas aeruginosa. Mol. Microbiol. 7~669-682. Hoch, J. A. 1993. Regulation of phosphorelay and the initiation of sporulation in Bacillus subtilis. Annu. Rev. Microbiol. 47:441-465. Hodgkin, J., and D. Kaiser. 1977. Cell-to-cell stimulation of movement in nonmotile mutants of Myxococcus. Proc. Natl. Acad. Sci. USA 74:2938-2942.
73 Hodgkin, J., and D. Kaiser. 1979. Genetics of gliding motility xanthus (Myxobacterales): two gene systems in M~XOCOCCUS control movement. Mol. Gen. Genet. 171:177-191. Hogg, T., U. Mechold, H. Malke, M. Cashel, and R. Hilgenfeld. 2004. Conformational antagonism between opposing active sites in a bifunctional RelNSpoT homolog modulates (p)ppGpp metabolism during the stringent response. Cell 117:5 7-6 8. Ishii, Y., H. Yamada, T. Yamashino, K. Ohashi, E. Katoh, H. Shindo, T. Yamazaki, and T. Mizuno. 2000. Deletion of the yhhP gene results in filamentous cell morphology in Escherichia coli. Biosci. Biotechnol. Biochem. 64:799807. Jain, V., M. Kumar, and D. Chatterji. 2006. ppGpp: stringent response and survival. J. Microbiol. 44:l-10. Jakobsen, J. S., L. Jelsbak, L. Jelsbak, R. D. Welch, C. Cummings, B. Goldman, E. Stark, S. Slater, and D. Kaiser. 2004. us4enhancer binding proteins and Myxococcus xanthus fruiting body development. J. Bacteriol. 186:43614368. Jelsbak, L., M. Givskov, and D. Kaiser. 2005. Enhancer-binding proteins with a forkhead-associated domain and the d4 regulon in Myxococcus xanthus fruiting body development. Proc. Natl. Acad. Sci. USA 102:3010-3015. Jelsbak, L., and D. Kaiser. 2005. Regulating pilin expression reveals a threshold for S-motility in Myxococcus xanthus. J. Bacteriol. 1872105-2112. Johansson, J., C. Balsalobre, S. Wang, J. Urbonaviciene, D. J. Jin, B. Sonden, and B. E. Uhlin. 2000. Nucleoid proteins stimulate stringently controlled bacterial promoters: a link between the CAMP-CRPand the (p)ppGppregulons in Escherichia coli. Cell 102:475-485. Kaiser, D. 2004. Signaling in myxobacteria. Annu. Rev. Microbiol. 58:75-98. Kaiser, D., and C. Crosby. 1983. Cell movement and its coordination in swarms of Myxococcus xanthus. Cell Motil. 3~227-245. Kaiser, D., and L. Kroos. 1993. Intercellular signaling, p. 257-284. In M. Dworkin and D. Kaiser (ed.), Myxobacteria 11.American Society for Microbiology, Washington, DC. Kang, P. J., and E. A. Craig. 1990. Identification and characterization of a new Escherichia coli gene that is a dosage-dependent suppressor of a dnaK deletion mutation. J. Bacteriol. 172:2055-2064. Kaplan, H. B. 2003. Multicellular development and gliding motility in Myxococcus xanthus. Curr. Opin. Microbiol. 6572-5 77. Kaplan, H. B., A. Kuspa, and D. Kaiser. 1991. Suppressors that permit A-signal independent developmental gene expression in Myxococcus xanthus. J. Bacteriol. 173:1460-1470. Kaplan, H. B., and L. Plamann. 1996. A Myxococcus xanthus cell density-sensing system required for multicellular development. FEMS Microbiol. Lett. 139:89-95. Kaufman, R. I., and B. T. Nixon. 1996. Use of PCR to isolate genes encoding as4-dependent activators from diverse bacteria. J. Bacteriol. 178:3967-3970. Keseler, I. M., and D. Kaiser. 1995. An early A-signal-dependent gene in Myxococcus xanthus has a as4-like promoter. J. Bacteriol. 1774638-4644.
74 Keseler, I. M., and D. Kaiser. 1997. us4,a vital protein for Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 94:1979-1984. Kimsey, H. H., and D. Kaiser. 1991. Targeted disruption of the Myxococcus xanthus orotidine 5'-monophosphate decarboxylase gene: effects on growth and fruiting-body development. J. Bacteriol. 173:6790-6797. Kirby, J. R., and D. R. Zusman. 2003. Chemosensory regulation of developmental gene expression in Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 100:2008-2013. Kroos, L. 2005. Eukaryotic-like signaling and gene regulation in a prokaryote that undergoes multicellular development. Proc. Natl. Acad. Sci. USA 102:2681-2682. Kroos, L., P. Hartzell, K. Stephens, and D. Kaiser. 1988. A link between cell movement and gene expression argues that motility is required for cell-cell signaling during fruiting body development. Genes Dev. 2:1677-1685. Kroos, L., and D. Kaiser. 1984. Construction of Tn5 lac, a transposon that fuses lacZ expression to exogenous promoters, and its introduction into Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 815816-5820. Kroos, L., and D. Kaiser. 1987. Expression of many developmentally regulated genes in Myxococcus xanthus depends on a sequence of cell interactions. Genes Dev. 1:840-854. Kroos, L., A. Kuspa, and D. Kaiser. 1986. A global analysis of developmentally regulated genes in Myxococcus xanthus. Dev. Biol. 117:252-266. Kroos, L., B. Zhang, H. Ichikawa, and Y. T. Yu. 1999. Control of sigma factor activity during Bacillus subtilis sporulation. Mol. Microbiol. 31:1285-1294. Kruse, T., S. Lobedanz, N. M. Berthelsen, and L. SsgaardAndersen. 2001. C-signal: a cell surface-associated morphogen that induces and co-ordinates multicellular fruiting body morphogenesis and sporulation in Myxococcus xanthus. Mol. Microbiol. 40:156-168. Kuhwein, H., and H. Reichenbach. 1968. Swarming and Morphogenesis in Myxobacteria (C893/1965). Institut fur den Wissenschaftlichen Film, Gottingen, Germany. Kuner, J. M., and D. Kaiser. 1982. Fruiting body morphogenesis in submerged cultures of Myxococcus xanthus. J. Bacterial. 151:458-461. Kuner, J. M., and D. Kaiser. 1981. Introduction of transposon Tn5 into Myxococcus xanthus for analysis of developmental and other nonselectable mutants. Proc. Natl. Acad. Sci. USA 7k425-429. Kuspa, A. 1989. Intercellular Signalling in the Regulation of Early Development in Myxococcus xanthus. Ph.D. dissertation. Stanford University, Stanford, CA. Kuspa, A., and D. Kaiser. 1989. Genes required for developmental signalling in Myxococcus xanthus: three asg loci. J. Bacteriol. 171:2762-2772. Kuspa, A., L. Kroos, and D. Kaiser. 1986. Intercellular signaling is required for developmental gene expression in Myxococcus xanthus. Dev. Biol. 117267-276. Kuspa, A., L. Plamann, and D. Kaiser. 1992a. Identification of heat-stable A-factor from Myxococcus xanthus. J. Bacteriol. 174~3319-3326. Kuspa, A., L. Plamann, and D. Kaiser. 1992b. A-signalling and the cell density requirement for Myxococcus xanthus development. J. Bacteriol. 174:7360-7369.
DEVELOPMENT AND MOTILITY Kustu, S., E. Santero, J. Keener, D. Popham, and D. Weiss. 1989. Expression of us4(ntrA)-dependentgenes is probably united by a common mechanism. Microbiol. Rev. 53:367-376. Laffler, T., and J. A. Gallant. 1974. Stringent control of protein synthesis in E. coli. Cell 3:47-49. Lancero, H., N. B. Caberoy, S. Castaneda, Y. Li, A. Lu, D. Dutton, X. Y. Duan, H. B. Kaplan, W. Shi, and A. G. Garza. 2004. Characterization of a Myxococcus xanthus mutant that is defective for adventurous motility and social motility. Microbiology 150:4085-4093. Lancero, H. L., S. Castaneda, N. B. Caberoy, X. Ma, A. G. Garza, and W. Shi. 2005. Analysing protein-protein interactions of the Myxococcus xanthus Dif signalling pathway using the yeast two-hybrid system. Microbiology 151:15351541. LaRossa, R., J. Kuner, D. Hager, C. Manoil, and D. Kaiser. 1983. Developmental cell interactions of Myxococcus xanthus: analysis of mutants.]. Bacteriol. 153:1394-1404. Lee, B., P. I. Higgs, D. R. Zusman, and K. Cho. 2005. EspC is involved in controlling the timing of development in Myxococcus xanthus. J. Bacteriol. 1875029-5031. Liu, S., D. 0.Bayles, T. M. Mason, and B. J. Wilkinson. 2006. A cold-sensitive Listeria monocytogenes mutant has a transposon insertion in a gene encoding a putative membrane protein and shows altered (p)ppGpp levels. Appl. Environ. Microbiol. 72:3955-3959. Manoil, C., and D. Kaiser. 1980a. Accumulation of guanosine tetraphosphate and guanosine pentaphosphate in Myxococcus xanthus during starvation and myxospore formation. J . Bacteriol. 141:297-304. Manoil, C., and D. Kaiser. 1980b. Guanosine pentaphosphate and guanosine tetraphosphate accumulation and induction of Myxococcus xanthus fruiting body development. J . Bacteriol. 141:305-315. Manoil, C., and D. Kaiser. 1980c. Purine-containing compounds, including cyclic adenosine 3',5'-monophosphate, induce fruiting of Myxococcus xanthus by nutritional imbalance. J. Bacteriol. 141:374-377. Masuda, S., and C. E. Bauer. 2004. Null mutation of HvrA compensates for loss of an essential relAlspoT-like gene in Rhodobacter capsulatus. J. Bacteriol. 186:235-239. McVittie, A., F. Messik, and S. A. Zahler. 1962. Developmental biology of Myxococcus. J. Bacteriol. 84546-551. Mechold, U., and H. Malke. 1997. Characterization of the stringent and relaxed responses of Streptococcus equisimilis. J. Bacteriol. 179:2658-2667. Mechold, U., H. Murphy, L. Brown, and M. Cashel. 2002. Intramolecular regulation of the opposing (p)ppGpp catalytic activities of Rel(Seq), the Rel/Spo enzyme from Streptococcus equisimilis. J. Bacteriol. 184:2878-28 88. Mittenhuber, G. 2001. Comparative genomics and evolution of genes encoding bacterial (p)ppGpp synthetases/hydrolases (The Rel, RelA and SpoT proteins). J. Mol. Microbiol. Biotechnol. 3585-600. Molle, V., L. Kremer, C. Girard-Blanc, G. S. Besra, A. J. Cozzone, and J. F. Prost. 2003. An FHA phosphoprotein recognition domain mediates protein EmbR phosphorylation by PknH, a Ser/Thr protein kinase from Mycobacterium tuberculosis. Biochemistry 42:15300-15309.
AND EARLYDEVELOPMENTAL EVENTS 3. INITIATION
Morett, E., and M. Buck. 1988. NifA-dependent in vivo protection demonstrates that the upstream activator sequence of nif promoters is a protein binding site. Proc. Natl. Acad. Sci. USA 85:9401-9405. Morett, E., and L. Segovia. 1993. The d4bacterial enhancerbinding protein family: mechanism of action and phylogenetic relationship of their functional domains. J. Bacteriol. 175:6067-6074. Murray, M. D., and H. Bremer. 1996. Control of the SpoTdependent ppGpp synthesis and degradation in Escherichia coli. J. Mol. Biol. 259:41-57. Nariya, H., and S. Inouye. 2006. A protein Ser/Thr kinase cascade negatively regulates the DNA-binding activity of MrpC, a smaller form of which may be necessary for the Myxococcus xanthus development. Mol. Microbiol. 60: 1205-1217. Ninfa, A. J., M. R. Atkinson, E. S. Kamberov, J. Feng, and E. G. Ninfa. 1995. Control of bacterial nitrogen assimilation by the NRI-NRII two component system of enteric bacteria, p. 64-88. In T. J. Silhavy and J. A. Hoch (ed.),Two-Component Systems of Bacteria. ASM Press, Washington, DC. Ochi, K., J. Kandala, and E. Freese. 1982. Evidence that Bacillus subtilis sporulation induced by the stringent response is caused by the decrease in GTP or GDP. J. Bacteriol. 151:1062-1 065. O’Connor, K. A., and D. R. Zusman. 1983. Coliphage P1mediated transduction of cloned DNA from Escherichia coli to Myxococcus xanthus: use for complementation and recombinational analyses. J. Bacteriol. 155:317-329. O’Connor, K. A., and D. R. Zusman. 1990. Genetic analysis of tag mutants of Myxococcus xanthus provides evidence for two developmental aggregation systems. J. Bacteriol. 172:3 868-3878. Ogata, H., S. Audic, P. Renesto-Audiffren, P. E. Fournier, V. Barbe, D. Samson, V. ROUX,P. Cossart, J. Weissenbach, J. M. Claverie, and D. Raoult. 2001. Mechanisms of evolution in Rickettsia conorii and R. prowazekii. Science 293~2093-2098. Pardee, A. B., and L. S. Prestidge. 1956. The dependence of nucleic acid syntheses on the presence of amino acids in Escherichia coli. J . Bacteriol. 71:677-683. Paul, B. J., M. M. Barker, W. Ross, D. A. Schneider, C. Webb, J. W. Foster, and R. L. Gourse. 2004. DksA: a critical component of the transcription initiation machinery that potentiates the regulation of rRNA promoters by ppGpp and the initiating NTP. Cell 118:311-322. Paul, B. J., M. B. Berkman, and R. L. Gourse. 2005. DksA potentiates direct activation of amino acid promoters by ppGpp. Proc. Natl. Acad. Sci. USA 102:78223-78228. Pedersen, F. S., E. Lund, and N. 0. Kjeldgaard. 1973. Codon specific, tRNA dependent in vitro synthesis of ppGpp and pppGpp. Nut. New. Biol. 243:13-15. Perederina, A., V. Svetlov, M. N. Vassylyeva, T. H. Tahirov, S. Yokoyama, I. Artsimovitch, and D. G. Vassylyev. 2004. Regulation through the secondary channel-structural framework for ppGpp-DksA synergism during transcription. Cell 188~297-309. Pham, V. D., C. W. Shebelut, M. E. Diodati, C. T. Bull, and M. Singer. 2005a. Mutations affecting predation ability
75 of the soil bacterium Myxococcus xanthus. Microbiology 15 1:1865-1 874. Pham, V. D., C. W. Shebelut, E. J. Zumstein, and M. Singer. 2005b. BrgE is a regulator of Myxococcus xanthus development. Mol. Microbiol. 57:762-773. Plamann, L., J. M. Davis, B. Cantwell, and J. Mayor. 1994. Evidence that asgB encodes a DNA-binding protein essential for growth and development of Myxococcus xanthus. J. Bacteriol. 176:2013-2020. Plamann, L., A. Kuspa, and D. Kaiser. 1992. Proteins that rescue A-signal-defective mutants of Myxococcus xanthus. J. Bacteriol. 174:3311-33 18. Plamann, L., Y. Li, B. Cantwell, and J. Mayor. 1995. The Myxococcus xanthus asgA gene encodes a novel signal transduction protein required for multicellular development. J. Bacteriol. 177:2014-2020. Pollack, J. S., and M. Singer. 2001. SdeK, a histidine kinase required for Myxococcus xanthus development. J. Bacteriol. 1 8 3 ~ 389-3596. 5 Rasmussen, A. A., S. L. Porter, J. P. Armitage, and L. SogaardAndersen. 2005. Coupling of multicellular morphogenesis and cellular differentiation by an unusual hybrid histidine protein kinase in Myxococcus xanthus. Mol. Microbiol. 56~1358-1372. Rasmussen, A. A., and L. Ssgaard-Andersen. 2003. TodK, a putative histidine protein kinase, regulates timing of fruiting body morphogenesis in Myxococcus xanthus. J. Bacteriol. 18.55452-5464. Rasmussen, A. A., S. Wegener-Feldbrugge, S. L. Porter, J. I? Armitage, and L. Ssgaard-Andersen. 2006. Four signalling domains in the hybrid histidine protein kinase RodK of Myxococcus xanthus are required for activity. Mol. Microbiol. 60525-534. Romeo, J. M., and D. R. Zusman. 1991. Transcription of the myxobacterial hemagglutinin gene is mediated by a as4-like promoter and a cis-acting upstream regulatory region of DNA. J. Bacteriol. 173:2969-2976. Rosenberg, E., K. H. Keller, and M. Dworkin. 1977. Cell density-dependent growth of Myxococcus xanthus on casein. J. Bacteriol. 129:770-777. Saito, N., J. Xu, T. Hosaka, S. Okamoto, H. Aoki, M. J. Bibb, and K. Ochi. 2006. EshA accentuates ppGpp accumulation and is conditionally required for antibiotic production in Streptomyces coelicolor A3(2).J. Bacteriol. 188:4952-4961. Sands, M. K., and R. B. Roberts. 1952. The effects of a tryptophan-histidine deficiency in a mutant of Escherichia coli. J. Bacteriol. 63505-511. Shimkets, L. 1984. Nutrition, metabolism, and the initiation of development, p. 92. In E. Rosenberg (ed.), Myxobacteria Development and Cell Interactions. Springer-Verlag, New York, NY. Shimkets, L. J. 1987. Control of morphogenesis in myxobacteria. Crit. Rev. Microbiol. 14:195-227. Shimkets, L. J., and M. Dworkin. 1981. Excreted adenosine is a cell density signal for the initiation of fruiting body formation in Myxococcus xanthus. Dev. Biol. 8451-60. Shimkets, L. J., R. E. Gill, and D. Kaiser. 1983. Developmental cell interactions in Myxococcus xanthus and the spoC locus. Proc. Natl. Acad. Sci. USA 80:1406-1410.
76 Singer, M., and D. Kaiser. 1995. Ectopic production of guanosine penta- and tetraphosphate can initiate early developmental gene expression in Myxococcus xanthus. Genes. Dev. 9~1633-1644. Stent, G. S., and S. Brenner. 1961. A genetic locus for the regulation of ribonucleic acid synthesis. Proc. Natl. Acad. Sci. USA 47~2005-2014. Studholme, D. J., and R. Dixon. 2003. Domain architectures of us4-dependent transcriptional activators. J. Bacteriol. 185:1757-1767. Sun, H., and W. Shi. 2001a. Analysis of mrp genes during Myxococcus xanthus development. J. Bacteriol. 183:6733-6739. Sun, H., and W. Shi. 2001b. Genetic studies of mrp, a locus essential for cellular aggregation and sporulation of Myxococcus xanthus. J. Bacteriol. 183:4786-4795. Sun, J., A. Hesketh, and M. Bibb. 2001. Functional analysis of relA and rshA, two relAlspoT homologues of Streptomyces coelicolor A3(2).J. Bacteriol. 183:3488-3498. Thony-Meyer, L., andD. Kaiser. 1993.devRS, an auto-regulated and essential genetic locus for fruiting body development in Myxococcus xanthus. J. Bacteriol. 175:7450-7462. Tojo, N., S. Inouye, and T. Komano. 1993a. Cloning and nucleotide sequence of the Myxococcus xanthus lon gene: indispensability of lon for vegetative growth. J. Bacteriol. 175~2271-2277. Tojo, N., S. Inouye, and T. Komano. 1993b. The lonD gene is homologous to the lon gene encoding an ATP-dependent protease and is essential for the development of Myxococcus xanthus. J. Bacteriol. 175:4545-4549. Tse, H., and R. E. Gill. 2002. Bypass of A- and B-signaling requirements for Myxococcus xanthus development by mutations in spdR.1. Bacteriol. 184:1455-1457. Turner, A. K., M. A. Lovell, S. D. Hulme, L. Zhang-Barber, and P. A. Barlow. 1998. Identification of Salmonella typhimurium genes required for colonization of the chicken alimentary tract and for virulence in newly hatched chicks. Infect. Immun. 66:2099-2106. Tzeng, L., T. N. Ellis, and M. Singer. 2006. DNA replication during aggregation phase is essential for Myxococcus xanthus development. J. Bacteriol. 188:2774-2779. Ueki, T., and S. Inouye. 1998. A new sigma factor, SigD, essential for stationary phase is also required for multicellular differentiation in Myxococcus xanthus. Genes Cells 3:371-385. Ueki, T., and S. Inouye. 2001. SigB, SigC, and SigE from Myxococcus xanthus homologous to u3’ are not required for heat
DEVELOPMENT AND MOTILITY shock response but for multicellular differentiation. J. Mol. Microbiol. Biotechnol. 3:287-293. Ueki, T., and S. Inouye. 2002. Transcriptional activation of a heat-shock gene, lonD, of Myxococcus xanthus by a two component histidine-aspartate phosphorelay system. J. Biol. Chem. 2776170-6177. van der Biezen, E. A., J. Sun, M. J. Coleman, M. Bibb, and J. D. G. Jones. 2000. Arabidopsis RelMSpoT homologs implicate (p)ppGpp in plant signaling. Proc. fiatl. Acad. Sci. USA 97:3747-3752. Viswanathan, P., M. Singer, and L. Kroos. 2006. Role of sigmaD in regulating genes and signals during Myxococcus xanthus development. J. Bacteriol. 188:3246-3256. Wells, D. H., and S. R. Long. 2002. The Sinorhizobium meliloti stringent response affects multiple aspects of symbiosis. Mol. Microbiol. 43:1115-1127. Wendrich, T. M., and M. A. Marahiel. 1997. Cloning and characterization of a relAlspoT homologue from Bacillus subtilis. Mol. Microbiol. 26:65-79. Wireman, J. W., and M. Dworkin. 1975. Morphogenesis and developmental interactions in myxobacteria. Science 1 8 9 516-523. Wu, S. S., and D. Kaiser. 1995. Genetic and functional evidence that Type IV pili are required for social gliding motility in Myxococcus xanthus. Mol. Microbiol. 18547-558. Wu, S. S., and D. Kaiser. 1997. Regulation of expression of the pilA gene in Myxococcus xanthus. J. Bacteriol. 179:77487758. Xu, D., C. Yang, and H. B. Kaplan. 1998. Myxococcus xanthus sasN encodes a regulator that prevents developmental gene expression during growth. J. Bacteriol. 180:62156223. Xu, H., and T. R. Hoover. 2001. Transcriptional regulation at a distance in bacteria. Curr. Opin. Microbiol. 4:138-144. Yang, C., and H. B. Kaplan. 1997. Myxococcus xanthus sass encodes a sensor histidine kinase required for early developmental gene expression.J. Bacteriol, 1797759-7767. Yoder, D. R., and L. Kroos. 2004a. Mutational analysis of the Myxococcus xanthus a 4 4 0 0 promoter region provides insight into developmental gene regulation by C-signaling. J. Bacteriol. 186:661-671. Yoder, D. R., and L. Kroos. 2004b. Mutational analysis of the Myxococcus xanthus R4499 promoter region reveals shared and unique properties in comparison with other C-signaldependent promoters. J. Bacteriol. 186:3766-3776.
Myxobacteria: Multicellularity and Differentiation Edited by David E. Whitworth 0 2008 ASM Press, Washington, D.C.
Lotte S~gaard-Andersen
Contact-Dependent Signaling in Myxococcus xanthus: the Function of the C-Signal in Fruiting Body Morphogenesis The past decade has witnessed a paradigm shift in microbiology. Previously, bacteria were largely regarded as autonomous individuals that did not interact with each other. The exceptions to the rule were the myxobacteria, which for more than 50 years had been suspected to interact during fruiting body morphogenesis, and Vibrio species, which were shown to emit bioluminescence in a cell-density-dependent manner in the 1960s. Today, the notion of bacteria as autonomous individuals that do not communicate has been replaced by one in which bacteria communicate extensively both within and between species by means of signaling molecules. Typically, these signals are small diffusible molecules such as acyiated homoserine lactones (AHLs) in gram-negative bacteria, modified oligopeptides in gram-positive bacteria, and autoinducer-2 (AI-2), which is found in gram-negative as well as in gram-positive bacteria. These signals are part of quorum-sensing systems that allow bacteria to regulate processes in response to changes in cell density. Analyses of the Myxococcus xanthus genome reveal that M . xanthus likely encodes neither Lux1 nor LuxS homologs, the two enzymes involved in synthesizing AHLs and AI-2, respectively. Consistently, the two biochemically and functionally characterized signaling molecules, i.e., the A-signal and the C-signal, in M. xanthus, the
model organism for analyzing intercellular communication in myxobacteria, are neither an AHL nor AI-2. Rather, the A-signal appears to be specific subsets of amino acids and small peptides, and the C-signal is a 17-kDa cell-surface-associated protein. The intercellular C-signal has a fundamental role in fruiting body morphogenesis in M. xanthus. It is used repeatedly during fruiting body formation and is the intercellular signal that induces and coordinates the three morphological processes involved in fruiting body formation, i.e., rippling, aggregation of cells into nascent fruiting bodies, and sporulation in those cells that have aggregated inside the nascent fruiting bodies. Also, the C-signal induces the expression of a large number of genes that are turned on after 6 h of starvation. In this chapter, our current understanding of how the C-signal acts at the molecular level to induce and coordinate events that are separated in time and space is discussed.
BIOFILM FORMATION IN M. XANTHUS Myxobacteria are found in the topsoil, where they feed on organic matter and prey on other microorganisms by secreting hydrolytic enzymes and antimicrobials
Lotte S~gaard-Andersen,Max Planck Institute for Terrestrial Microbiology, Karl-von-Frisch Str., 35043 Marburg, Germany.
77
DEVELOPMENT AND MOTILITY
78 (Reichenbach, 1999). If present on a solid surface at a high cell density, M. xanthus cells can form two morphologically distinct types of biofilms depending on the nutritional status of the cells. In the presence of nutrients, the motile, rod-shaped cells grow, divide, and form spreading, cooperatively feeding colonies. Cells at the edge of a colony spread coordinately over the surface, forming a thin, film-like structure. In the absence of nutrients, the spreading behavior is constrained and cells begin to aggregate. The aggregation process culminates in the formation of the multicellular, spore-filled fruiting bodies. Fruiting body formation proceeds in distinct morphological stages that are separated in time and space (Fig. 1).The first signs of fruiting body formation are evident after 4 to 6 h of starvation as cells begin to aggregate to form small aggregation centers. During aggregation, cells move into the aggregation centers organized in streams rather than entering the centers as single cells from all directions (O’Connor and Zusman, 1989).As more cells enter the aggregation centers, these centers increase in size and eventually become mound shaped. By 24 h, the aggregation process is complete and the nascent fruiting bodies each contain -lo5 densely packed cells. Inside the nascent fruiting bodies, the rod-shaped cells undergo morphological and physiological differentiation into spherical myxospores resulting in mature fruiting bodies. Spore maturation
Initial aggregation with formation of aggregation centers
is finished -72 h after the onset of starvation. Only 10 to 12% of cells undergo sporulation, and these cells are specifically those that have accumulated inside the fruiting bodies. Up to 30% of the cells remain outside the fruiting bodies. These cells remain rod shaped and differentiate to a cell type called peripheral rods. Even after extended periods of starvation, peripheral rods do not differentiate into spores (O’Connor and Zusman, 1991). Finally, the remaining cells undergo lysis (Rosenbluh et al., 1989). Aggregation and sporulation are the two invariable morphological processes in fruiting body formation. Under less stringent starvation conditions, fruiting body formation includes a third morphological process referred to as rippling. During rippling, cells accumulate in equispaced ridge-like structures separated by troughs of low cell density. The ridge-like structures move coordinately and synchronously as traveling waves over the surface (Reichenbach, 1965; Shimkets and Kaiser, 1982) (for a time-lapse movie of rippling cells, see Welch and Kaiser, 2001). Microscopic examination of cells during rippling has shown that individual cells essentially oscillate back and forth with no net movement, suggesting that colliding waves reflect each other (Sager and Kaiser, 1994; Welch and Kaiser, 2001). Rippling is typically initiated prior to aggregation. Later, during the aggregation process the wave structure disintegrates and cells
Mounds
Nascent fruiting bodies
Mature fruiting bodies
0 00 000 0000
00000 000000 0000000 00000000
ooooc3oooo
Morphological markers
0
0
~
_ I
Transcriptional markers 0 Hrs of starvation Intercellular signals
12
6
t
A
24
II-
72
C Min
Mar
Level of C-signaling
Figure 1 Schematic outline of fruiting body morphogenesis in M . xanthus. The different morphological stages are indicated. Triangles indicate genes that are induced at different time points during fruiting body formation. Filled and open triangles indicate C-signalindependent and -dependent genes, respectively. The times at which the intercellular A- and C-signals become important for development are indicated. The level of C-signaling that individual cells are exposed to is indicated by the level of gray. The grayscale below the timeline indicates C-signaling levels in individual cells.
0
0
~
4.
CONTACT-DEPENDENT SIGNALINGIN M. XANTHUS
aggregate into the nascent fruiting bodies. Fruiting body formation involves temporally coordinated changes in gene expression in which genes are turned on at specific time points during development (Inouye et al., 1979a; Kroos et al., 1986) (Fig. 1).Moreover, developmental gene expression is subject to spatial control in which genes that are turned on after 6 h are preferentially expressed in aggregating and sporulating cells whereas genes activated prior to 6 h are expressed in all cells including peripheral rods (Julien et al., 2000).
INTERCELLULAR SIGNALING DURING FRUITING BODY MORPHOGENESIS The first evidence for intercellular signals important for fruiting body formation came from the isolation of a collection of mutants that displayed nonautonomous developmental defects. Sporulation of the mutant cells was rescued by codevelopment with wild-type cells (Hagen et al., 1978). This rescue is referred to as extracellular complementation (Hagen et al., 1978) to emphasize that it does not involve transfer of genetic material from wild-type cells to mutant cells. Importantly, the nonautonomous mutants were prototrophs, suggesting that extracellular complementation did not involve crossfeeding (Hagen et al., 1978).Extracellular complementation experiments with pairs of nonautonomous mutants led to the classification of the nonautonomous mutants into five classes referred to as the asg (A-signal), bsg (B-signal), csg (C-signal), dsg (D-signal), and esg (E-signal) mutants, respectively (Downard et al., 1993; Hagen et al., 1978). In these experiments, mutants from one class rescue development of mutants from a different class, i.e., an asg mutant rescues sporulation of a bsg mutant and vice versa, whereas codevelopment of mutants belonging to the same class does not result in rescue of development. The nonautonomous developmental defects were hypothesized to depend on the inability of the mutant cells to produce an intercellular signal required for fruiting body formation. Along the same lines, extracellular complementation was suggested to depend on this missing signal being provided by the codeveloping strain (Hagen et al., 1978). The nonautonomous developmental defects inxsgmut a n s provided evidence that the xsg genes are required for fruiting body formation and that the defects can be rescued by extracellular complementation. Formally, however, these experiments do not provide evidence that the xsg genes are required for the synthesis of intercellular signals that induce specific responses. Among the five potential intercellular signals, referred to as the A- to E-signals, required for fruiting body formation,
79 only two-the A- and C-signals-have been characterized biochemically and functionally and shown to be true signaling molecules. It remains an open question whether the bsg, dsg, and esg mutants are deficient in the synthesis of an intercellular signal. The A-signal becomes important for development after 2 h of starvation (Kuspa et al., 1986) (Fig. 1)and consists of a subset of amino acids and peptides that are produced by the action of extracellular proteases, which are released in response to starvation (Kuspa et al., 1992b; Plamann et al., 1992). The A-signal functions as part of a system that measures the density of starving cells: each starving cell produces a constant amount of A-signal, and once a threshold concentration of A-signal is reached development proceeds (Kuspa et al., 1992a). Thus, the function of the A-signaling system seems to be to ensure that fruiting body morphogenesis does not proceed unless the density of starving cells is sufficiently high.
THE C-SIGNAL INDUCES FOUR RESPONSES THAT ARE SEPARATED IN TIME AND SPACE The C-signal comes into action after 6 h of starvation, coinciding with the first signs of morphogenesis (Kroos and Kaiser, 1987) (Fig. 1).Mutants that cannot synthesize the C-signal are unable to ripple, aggregate, and sporulate (Shimkets et al., 1983), and the expression of genes, which are normally turned on from 6 h, is reduced or abolished (Kroos and Kaiser, 1987). Synthesis of the C-signal depends on the csgA gene (Shimkets et al., 1983).Development of csgA mutant cells can be restored in two ways, by codevelopment with wild-type cells or with cells from either the asg, bsg, dsg, or esg classes of nonautonomous mutants or by purified, exogenously added C-signal (Kim and Kaiser, 1990d). Three lines of evidence suggest that the C-signal is the signal that induces rippling, aggregation, sporulation, and gene expression after 6 h. Kim and Kaiser (Kim and Kaiser, 1991) added increasing amounts of exogenous C-signal to starving csgA cells and observed that aggregation and expression of a C-signal dependent gene that is normally expressed at 6 h was induced by the addition of an intermediate amount of C-signal. Addition of a higher amount of purified C-signal molecule also induced sporulation as well as the expression of a late C-signal dependent gene. By manipulating expression of the csgA gene in vivo, Li et al. (Li et al., 1992) observed that a low level of csgA expression was sufficient to induce rippling, an intermediate level was sufficient to induce aggregation, and a high level of csgA expression was required to obtain sporulation. These experiments
80
showed that the C-signal molecule is required for aggregation, sporulation, and gene expression. To show that the C-signal is sufficient to induce these responses, Kruse et al. (Kruse et al., 2001) overexpressed the csgA gene in vivo, resulting in the accumulation of large amounts of the C-signal molecule early during development. Importantly, aggregation, sporulation, and C-signal-dependent gene expression were induced earlier than in wild-type cells, whereas the rippling stage was completely skipped. Thus, this experiment provided the crucial piece of evidence demonstrating that the C-signal is not only required but also sufficient to induce rippling, aggregation, sporulation, and full expression of developmental genes, which are turned on after 6 h.
THE MOLECULAR IDENTITY OF THE C-SIGNAL Originally, Kaiser and coworkers identified the C-signal as a 17-kDa protein encoded by the csgA gene (Kim and Kaiser, 1990c, 1990d). The C-signal was purified by detergent extraction and biochemical fractionation of starving M . xanthus cells on the basis of its ability to rescue development of csgA cells (Kim and Kaiser, 1990c, 1990d). The understanding that the 17-kDa protein was the C-signal was questioned by several observations, which suggested that the CsgA protein might act as a short-chain alcohol dehydrogenase (SCAD) to produce the C-signal. First, the csgA gene encodes a 25kDa CsgA protein (Kruse et al., 2001; Lee et al., 1995), which shows homology to SCADs (Baker, 1994; Lee et al., 1995). SCADs contain two conserved sequence motifs which are both present in the CsgA protein: an N-terminal motif corresponding to the NAD(P)-coenzyme binding pocket and a more C-terminal motif, which is part of the active site (Oppermann et al., 2003). Consistently, the full-length CsgA protein binds NAD+ in vitro (Lee et al., 1995). Moreover, csgA alleles encoding mutant versions of the full-length CsgA protein with substitutions in either the coenzyme binding pocket or in the active site did not complement a csgA mutant (Lee et al., 1995). Thirdly, exogenous full-length CsgA protein purified from Escherichia coli restores development of csgA cells, whereas exogenous full-length CsgA proteins carrying substitutions in either the coenzyme binding pocket or in the active site failed to restore development of csgA cells (Lee et al., 1995). Finally, overproduction of the SocA protein, which also shows homology to SCADs, restores development in a csgA mutant in vivo (Lee and Shimkets, 1994, 1996). Recently, these conflicting data were partially reconciled (Lobedanz and Ssgaard-Andersen, 2003). First it
DEVELOPMENT AND MOTILITY was shown that the CsgA protein exists in two forms, one with an approximate size of 25 kDa (designated p25), which corresponds to full-length CsgA protein, and one with an approximate size of 17 kDa (designated p17) (Kruse et al., 2001), which is similar in size to the C-signal protein purified by Kim and Kaiser (Kim and Kaiser, 1990c, 1990d). p17 and p25 are anchored in the outer membrane (Lobedanz and Ssgaard-Andersen, 2003). p25 is present in vegetative cells and accumulates during fruiting body formation, whereas p l 7 is detected only in starving cells (Kruse et al., 2001). Using the procedure of Kim and Kaiser for purification of the C-signal, Lobedanz and S~gaard-Andersenshowed that the C-signal copurifies with p17. p17 corresponds to the C-terminal17 kDa of p25; however, the precise N terminus of p17 has yet to be determined. Importantly, recombinant p17 proteins that correspond to the C-terminal 17 kDa of p25 and differ only in their N termini have C-signal activity. These recombinant p17 proteins lack the NAD+ coenzyme-binding pocket and are unable to bind NAD+ in vitro. Thus, these data strongly suggest that p17 does not depend on SCAD activity to engage in C-signaling. Rather, these data support the notion that p17 is the C-signal.
SYNTHESIS OF THE 17-kDa C-SIGNAL PROTEIN Having identified p17 as the C-signal, an important question became the mechanism by which p17 is produced. Shimkets and coworkers have shown that the start codon for p25 synthesis is essential for synthesis of the C-signal (Lee et al., 1995). To test whether p17 synthesis involved proteolytic processing of p25, Lobedanz and Ssgaard-Andersen used an in vitro protease assay in which total cell extracts were prepared from starving M . xanthus cells and then added to a recombinant p25 (Lobedanz and Ssgaard-Andersen, 2003). Using this assay, it was found that addition of M . xanthus cell extract to recombinant p25 resulted in the synthesis of p17. By adding protease inhibitors specific for different types of proteases, evidence was obtained that the protease involved in p25 processing is a serine protease. Consistent with the observation that p17 is detected only in developing cells (Kruse et al., 2001), it was observed that the activity of the protease is developmentally regulated. This protease is referred to as PopC for protease required for processing of the precursor of the C-signal. In cell fraction experiments, p25 and p17 are detected only in the outer membrane, arguing that the proteolytic processing of p25 occurs after the insertion in the outer membrane. This in turn argues that PopC is a secreted
4.
CONTACT-DEPENDENT SIGNALING I N M.XANTHUS
protease. To identify the popC gene, A. Rolbetzki and L. Sliigaard-Andersen (unpublished data) used a candidate approach in which genes in the M. xanthus genome likely to encode secreted serine proteases were identified. Among the 32 genes identified, 20 were likely to encode proteases with an inhibition profile similar to that of the serine protease involved in p25 cleavage as observed in the in vitro protease experiments by Lobedanz and Ssgaard-Andersen. Among these 20 protease genes, five were shown to be induced at the transcriptional level during development in genome-wide transcriptional profiling experiments using the M. xanthus DNA microarray. Inactivation of one of these genes, MXAN0206, which encodes a subtilisin-like serine protease, results in aggregation and sporulation defects. Most importantly, the MXAN0206 mutant is impaired in p17 synthesis whereas p25 synthesis is largely unaffected. Interestingly, MXAN0206 accumulates in vegetative cells; however, MXAN0206 appears to be secreted only in starvingcells. ThesedatasupporttheideathatMXAN0206 encodes PopC, and they suggest that p17 accumulation is restricted to starving cells by allowing MXAN0206 secretion only in starving cells. However, it still remains to be shown that the protease encoded by MXAN0206 directly cleaves p25. Overall, we now have a framework for understanding the molecular nature of the C-signal and the mechanism involved in its synthesis (Fig. 2A). However, the inability of mutant p25 proteins, which carry substitutions in the coenzyme binding pocket or in the active site, to rescue development of csgA cells in the C-signal bioassay (Lee et al., 1995) is intriguing. The substitutions in the coenzyme binding pocket may interfere with proteolytic processing of p25, and the substitutions in the active site may interfere with recognition by the C-signal receptor. A second unresolved issue is the potential enzymatic activity of p25. SCAD activity of full-length CsgA protein has not been demonstrated. Is p25 an enzymatic fossil, which functions only as a precursor for p17? Or does p25 still have SCAD activity? Clearly, this potential enzymatic activity is not required for fruiting body formation as development of csgA cells is rescued by exogenous p17.
CONTACT-DEPENDENT C-SIGNAL TRANSMISSION Early on, a striking correlation was observed between cell motility and C-signal transmission. First, nonmotile cells have an abnormal pattern of developmental gene expression that matches the pattern observed in csgA cells (Kroos et al., 1988). Secondly, in extracellular
81 complementation experiments donor cells of the C-signal as well as receiver cells of the C-signal need to be motile in order for C-signal transmission to occur (Kim and Kaiser, 1990b).Thirdly, the requirement for cell motility could be bypassed by artificially aligning cells in such a manner that extensive end-to-end contacts were established (Kim and Kaiser, 1990a). These experiments led to the suggestion that cell motility is required to establish specific cell-cell contacts between donor and receiver cells in order for C-signaling to occur and that these contacts are end-to-end contacts. Later, the observation that wild-type cells and csgA cells need to be in direct contact in order for extracellular complementation to occur further supported the contact-dependent C-signal transmission mechanism (Kim and Kaiser, 1990d). In addition, the observations that CsgA antibodies recognize epitopes that are located on the surface of developing cells (Shimkets and Rafiee, 1990)and that the 17-kDa C-signal protein is localized to the outer membrane (Lobedanz and Ssgaard-Andersen, 2003) showed that the C-signal is nondiffusible and, thus, further supported the idea that C-signal transmission involves a contact-dependent mechanism. Definitive evidence that C-signal transmission depends on specific end-to-end contacts is still lacking. However, analyses of cell behavior during rippling lend support to the notion that C-signal transmission involves end-to-end contacts (Sager and Kaiser, 1994).
THE C-SIGNAL TRANSDUCTION PATHWAY To understand how one signal may induce four responses, i.e., rippling, aggregation, sporulation, and developmental gene expression, which are separated temporally and spatially, it is crucial to clarify how the signal transduction pathway is structured. Random transposon mutagenesis followed by screening for mutants with deficiencies in C-signal-dependent responses, isolation of extragenic suppressors of a csgA insertion mutant, proteomics, and biochemical analyses have been instrumental in the identification of proteins in this pathway (Fig. 2A). The signaling event in the pathway is hypothesized to be the interaction between the 17-kDa C-signal protein located on one cell with a C-signal receptor located on a neighboring cell. This C-signal receptor has yet to be identified. A central protein in the pathway is the DNA binding response regulator FruA, which consists of an N-terminal receiver domain and a C-terminal DNA binding domain (Ellehauge et al., 1998; Ogawa et al., 1996). fmA mutants are unable to ripple, aggregate, and sporulate and are deficient in the expression of several C-signal-dependent genes (Ellehauge et al., 1998;
A.
I !I
I:
MrpC
n.=,ir
FruA
TodK
SdeK
c
4
\
c
I
t
Stariation
4 25 kDa CsgAprotein
-
T
I
J
I
Key:
RodK
17 kDa C-signal protein --(C-signai
receptor
---.---. outer membrane
inner membrane
B. FruA
t
fr"A
MrpC2 tLonD
I ~~
c14-P7
Pkn8
Pknl4
Pkn8-P
Signal? MrpA-P
mrpA
MrpB
mrpB
MrpC-P
+ Signal? +Starvation
mrpC
Starvation
Figure 2 The C-signal transduction pathway. (A) Model of the C-signal transduction pathway. Three different levels of phosphorylated FruA (FruA-P) are indicated, with the stippled circle indicating a low level and the heavy circle indicating a high level of phosphorylation. The encircled numbers indicate processes initiated at low, intermediate, and high levels of C-signaling and which may correspond to low, intermediate, and high levels of FruA phosphorylation, respectively. The shaded box indicates the site of convergence of the MXAN4899, SdeK, TodK, and RodK pathways with the C-signal transduction pathway. See the text for details. (B) Regulation of MrpC activity. See the text for details.
82
4.
CONTACT-DEPENDENT SIGNALING IN M .
XANTHUS
Horiuchi et al., 2002b; Ogawa et al., 1996; SsgaardAndersen and Kaiser, 1996). FruA activity is regulated at the transcriptional and posttranslational levels (Fig. 2A). fruA transcription is induced after 3 to 6 h of starvation. Several pathways converge to stimulate fruA transcription. The early acting A-signal by an unknown mechanism induces fruA transcription (Ellehauge et al., 1998; Ogawa et al., 1996). The DevT protein, which is encoded by the devTRS operon and which does not share similarity with other proteins in the databases, directly or indirectly stimulates transcription of fruA (Boysen et al., 2002). Finally, the MrpC protein, which is a homolog of the cyclic AMP receptor protein in E. coli (Sun and Shi, 2001b), binds directly to the fruA promoter and induces fruA transcription (Ueki and Inouye, 2003). Expression of mrpC, in turn, is activated by the MrpAB proteins, which constitute a two-component regulatory system with MrpA being a cytoplasmic histidine protein kinase and MrpB an enhancer binding protein of the NtrC type (Sun and Shi, 2001a, 2OOlb) (Fig. 2B). Expression of mrpAB is induced in response to starvation. In addition, MrpC directly stimulates transcription of mrpC (Nariya and Inouye, 2006; Sun and Shi, 2001a, 2001b). The activity of the MrpC protein is also regulated at the posttranslational level by phosphorylation and possibly also by proteolysis (Nariya and Inouye, 2005) (Fig. 2B). Specifically, the cytoplasmic Ser/Thr protein kinase Pknl4 phosphorylates MrpC. Pknl4, in turn, is phosphorylated by the integral membrane Ser/Thr kinase Pkn8. Phosphorylation of MrpC by Pknl4 appears to inhibit the activity of MrpC and results in decreased mrpC and fruA transcription. Moreover, MrpC seems to undergo proteolytic cleavage to MrpC2, which lacks the 25 N-terminal residues present in MrpC, in a manner that depends on LonD. MrpC2 binds with a higher affinity to the fruA and mrpC promoter regions. Importantly, Pknl4 does not phosphorylate MrpC2. Consistently, deletions of pkn8 and pkn14 result in increased accumulation of MrpC2 and FruA and faster progression through the developmental program. Accordingly, the Pkn8/Pknl4 kinase cascade negatively regulates the formation of MrpC2 by inducing phosphorylation of MrpC. In response to starvation, the Pknl8/Pkn14 kinase cascade would be inhibited and as a consequence MrpC2 would accumulate. The signal that regulates Pkn8, the upstream kinase in this cascade, remains to be identified, as does the signal that regulates MrpA activity. The regulatory mechanism that acts at the posttranslational level to activate FruA likely involves the phosphorylation of a conserved Asp residue in the N-terminal receiver domain (Ellehauge et al., 1998). Moreover, genetic evidence suggests that FruA
83 phosphorylation is induced by the C-signal (Ellehauge et al., 1998; Ssgaard-Andersen and Kaiser, 1996). The cognate FruA histidine protein kinase(s) has yet to be identified. Downstream from phosphorylated FruA the C-signal transduction pathway contains a branch point with one branch leading to rippling and aggregation and one leading to sporulation (Fig. 2A). The proteins in the cytoplasmic Frz chemosensory system are part of the motility branch (Ssgaard-Andersen and Kaiser, 1996; Ssgaard-Andersen et al., 1996). The Frz proteins share homology to proteins involved in chemotaxis responses in other bacteria (Ward and Zusman, 1999) and constitute a signal transduction system that controls several gliding motility parameters including the frequency of gliding reversals (Blackhart and Zusman, 1985; Jelsbak and Ssgaard-Andersen, 1999). In the motility branch, the C-signal induces methylation of the FrzCD protein (Ssgaard-Andersen and Kaiseq 1996), a methyl-accepting chemotaxis-protein (McBrideet al., 1989).C-signal-induced methylation of FrzCD depends on FruA and the FrzF methyltransferase (Ellehauge et al., 1998; SsgaardAndersen and Kaiser, 1996) (Fig. 2A). Genetic evidence suggests that during the rippling stage, the C-signal stimulates the Frz system and during the aggregation stage, the C-signal inhibits the Frz system (Sager and Kaiser, 1994; Ssgaard-Andersen and Kaiser, 1996) (Fig. 2A). The Frz system regulates the cellular reversal frequency by controlling and coordinating the frequency with which the two gliding engines in M. xanthus switch polarity (Mignot et al., 2005, 2007) (see chapter 6 for a description of gliding motility in M . xanthus). During the rippling stage, the C-signal-dependent Frz activation stimulates polarity switching of the two gliding engines and as a consequence, cells display a high reversal frequency (Sager and Kaiser, 1994; Welch and Kaiser, 2001). During the aggregation stage, the C-signal-dependent Frz inhibition results in inhibition of polarity switching of the two gliding engines and as a consequence, cells display a low reversal frequency (Jelsbak and SsgaardAndersen, 2002). The connection between the C-signal and the Frz system, its interaction with the gliding machinery, and the connection to cell behavior are discussed further below. The second branch downstream from phosphorylated FruA leads to sporulation (Ellehauge et al., 1998; Horiuchi et al., 2002b; Ssgaard-Andersen et al., 1996) (Fig. 2A). In this branch, FruA acts as a transcriptional regulator. Analyses of protein synthesized during development have shown that the C-signal and FruA jointly regulate the expression of at least 50 genes (Horiuchi et al., 2002b). Analyses of the expression of TnSlac promoter
84 fusions in csgA and fruA mutants led to the identification of the devTRS operon as a transcriptional target of FruA and the C-signal (Ellehauge et al., 1998). The DevTRS proteins are required for the expression of the sporulation gene tagged by the TnSluc 0 7 5 3 6 insertion (Licking et al., 2000). Moreover, DevT is required for full expression of the fruA gene (Boysen et al., 2002). In addition to regulating the expression of C-signal-dependent genes, FruA directly regulates the expression of at least eight genes including the dofA gene in a C-signal-independent manner (Horiuchi et al., 2002a, 2002b) (Fig. 2A). In the case of the dofA gene, which encodes a protein with an unknown function, the DNA binding domain of FruA directly binds to the promoter region (Ueki and Inouye, 2005b). Finally, the fdgA, s ~ Atps , (Ueki and Inouye, 2005a), and a 4 4 0 0 (Yoder-Himes and Kroos, 2006) genes have been shown to absolutely depend on FruA for full expression while only partially depending on the C-signal for full expression. The DNA binding domain of FruA has been documented to bind directly to the promoter region of the fdgA and 04400 genes, suggesting that FruA directly activates the transcription of these two genes. fdgA encodes an outer membrane lipoprotein involved in polysaccharide export (Ueki and Inouye, 2005a), sasA is important for biosynthesis of the lipopolysaccharide O-antigen (Guo et al., 1996), and tps encodes protein S , a major protein of the outer surface of myxospores (Inouye et al., 1979b). It is currently not known whether expression of FruA-dependent and Csignal-independent genes involves unphosphorylated FruA or whether FruA is phosphorylated in a C-signalindependent manner. A third branch in the C-signal transduction pathway is located upstream from FruA and by an unknown mechanisms leads to increased transcription of the csgA gene (Kim and Kaiser, 1991) (Fig. 2A). csgA is transcribed in vegetative cells, and transcription increases approximately fourfold during development (Li et al., 1992).The increase in csgA transcription in response to starvation involves RelA and the stringent response (Crawford and Shimkets, 2000) and the four proteins encoded by the act operon (Gronewold and Kaiser, 2001). ActA consists of an N-terminal receiver domain and a C-terminal GGDEF domain likely to be involved in cyclic-di-GMP synthesis (Jenal, 2004). ActB is similar to enhancer binding proteins of the NtrC family. ActC is a protein of unknown function containing an acetyltransferase domain and an epimerase/dehydratase domain. ActD does not contain conserved domains. ActA and ActB are both required for full transcription of csgA during development, whereas ActC and ActD are important for the correct timing of csgA transcription.
DEVELOPMENT AND MOTILITY The C-signal transduction pathway is activated in response to starvation. Several regulatory mechanisms help to restrict the activity of the pathway to starving cells. First, starvation induces the stringent response (Singer and Kaiser, 1995), which, in turn, induces csgA transcription (Crawford and Shimkets, 2000) and A-signal accumulation (Harris et al., 1998), which induces fruA transcription. Secondly, starvation induces mrpAB expression, and MrpAB induces mrpC transcription (Fig. 2A and B). Thirdly, by an unknown mechanism secretion of MXAN0206, the protease likely to cleave p25, is induced. For mrpAB it remains to be shown whether the increased transcription in response to starvation depends on the stringent response.
C-SIGNAL: A MORPHOGEN AND A TIMER OF DEVELOPMENT A hallmark in fruiting body formation is the temporal and spatial coordination of aggregation and sporulation: aggregation precedes sporulation, and only cells that have accumulated at a high cell density inside the fruiting bodies undergo sporulation. How is the C-signal transduction pathway structured to ensure the spatial and temporal coordination of aggregation and sporulation? The C-signal transduction pathway contains three amplification loops (Fig. 3). In the first loop, C-signaling induces aggregation and thus the accumulation of cells at a higher density. As C-signal transmission involves a contact-dependent mechanism, the prediction is that during the aggregation process the level of C-signaling that cells are exposed to increases. In the second loop, Csignaling results in increased csgA transcription, which results in p25 accumulation, which is processed to p17, which then engages in C-signaling with a neighboring cell. In the third loop, phosphorylated FruA induces transcription of the devTRS operon; the DevT protein, in turn, induces transcription of fruA. In combination, these three amplification loops may ensure that cells, which have engaged in C-signaling, are exposed to an ordered increase in the level of C-signaling. As outlined above, three lines of evidence suggest that the C-signal is the intercellular signal that induces rippling, aggregation, and sporulation (Kim and Kaiser, 1991; Kruse et al., 2001; Li et al., 1992).The same data suggest that the C-signal elicits distinct responses at different thresholds with rippling being induced at a low threshold, aggregation being induced at an intermediate threshold, and sporulation being induced at a high threshold (Fig. 1, 2A, and 3). Importantly, overexpression of the C-signal early during development not only
4.
CONTACT-DEPENDENT SIGNALING IN M . XANTHUS
85 Sporulation & late C-signal dependent gene expression
.-F (u
K
.-0) P
gregation & early C-signal pendent gene expression
% 0
0
f
Proteolytic
Starvation
-.+
csgA
-I
C-signaling
,(
expression fruA
Aggregation
)
phor:rylation
devT
Figure 3 A quantitative model for C-signal-induced responses. The three signal amplification loops in the C-signal transduction pathway are indicated. See the text for details.
results in premature aggregation and sporulation but also results in uncoupling of aggregation and sporulation with spores being formed outside fruiting bodies (Kruse et al., 2001). Together the C-signal thresholds in combination with the ordered increase in the level of C-signaling during development ensure that the C-signal induces first rippling, then aggregation and early gene expression, and finally, late gene expression and sporulation. According to this model, the C-signal is a timer of morphogenesis and a nondiffusible morphogen that induces distinct morphogenetic events at distinct thresholds. Currently, it is not known how different levels of C-signaling are transformed into different responses. However, one scenario would be that different levels of C-signaling are reflected in different levels of phosphorylated FruA and that these levels are subsequently transformed into the different responses (Fig. 2A). The spatial coordination of aggregation and sporulation is a direct consequence of the contact-dependent C-signal transmission mechanism (S~gaardAndersen et al., 2003). This signal transmission mechanism ensures that the level of C-signaling that a cell is exposed to reflects cell density and, thus, the position of a cell. The high level of C-signaling that induces late gene expression and sporulation is obtained only in cells that are closely packed inside the nascent fruiting bodies. As a consequence, only cells that have accumulated inside the fruiting bodies undergo sporulation. Thus, the mechanism of the C-signal allows cells to decode their position with respect to that of other cells and in that way
match gene expression and, ultimately, sporulation to their position. This model also provides an explanation for the spatial control of C-signal-dependent gene expression. C-signal-dependent genes are preferentially expressed in aggregating and sporulating cells (Julien et al., 2000). Peripheral rods are present at a low cell density outside the nascent fruiting bodies. Consequently, they only infrequently engage in contacts with other cells with C-signal transmission, Therefore, they experience a low level of C-signaling that allows neither C-signaldependent gene expression nor sporulation.
MULTIPLE SIGNAL TRANSDUCTION PATHWAYS CONTROL MORPHOGENESIS Genetic evidence suggests that several signal transduction pathways converge with the C-signal transduction pathway to regulate aggregation, sporulation, and gene expression after 6 h (Fig. 2A). Three of the pathways are defined by histidine protein kinases. For these three pathways, genetic evidence suggests that they converge with the C-signaling pathway downstream from FruA accumulation and upstream from the Frz system and devTRS expression. The SdeK histidine protein kinase is synthesized in a RelA-dependent manner immediately after the initiation of starvation, and the SdeK pathway by an unknown mechanism converges with the C-signal transduction pathway to stimulate aggregation, sporulation, and gene expression (Garza et al., 1998; Pollack
86 and Singer, 2001). The pathway defined by the TodK histidine protein kinase by an unknown mechanism inhibits aggregation, sporulation, and gene expression (Rasmussen and Ssgaard-Andersen, 2003). Synthesis of TodK is inhibited by starvation in a RelA-independent manner (Rasmussen and Ssgaard-Andersen, 2003). Genetic evidence suggests that the TodK-dependent inhibition of aggregation, sporulation, and gene expression is alleviated after 6 to 9 h of starvation (Rasmussen and SsgaardAndersen, 2003). SdeK and TodK are both predicted to be cytoplasmic proteins, and both contain PAS domains in their sensor part. PAS domains have been implicated in sensing changes in redox potential, oxygen, light, small ligands, and overall energy levels (Taylor and Zhulin, 1999). This led to the proposal that the kinase activities of TodK and SdeK are controlled by intracellular signals, which are indicative of the metabolic state of individual cells (Rasmussen and Ssgaard-Andersen, 2003). According to this model, starvation of cells results in the accumulation of these signals. This would subsequently trigger an alteration in the activity of the two kinases, resulting in the alleviation of the inhibitory effect of TodK and stimulating the activating effect of SdeK on the C-signal transduction pathway. RodK is the third kinase that defines a signal transduction pathway that converges with the C-signal transduction pathway (Rasmussen et al., 2005,2006). RodK is a cytoplasmic kinase and structurally highly complex with three C-terminal receiver domains in addition to the sensor and kinase domain. Genetic evidence suggests that RodK exerts an inhibitory effect on the C-signal transduction pathway. As in the case of TodK, the model for the action of RodK is that the signal that stimulates RodK activity vanishes during starvation, thus leading to the alleviation of the inhibition of the C-signal transduction pathway. The sensor domain in RodK is unique, and the signal recognized by RodK remains to be identified, as does the mechanism by which an output response is generated. A fourth pathway that converges with the C-signal transduction pathway is defined by the enhancer binding protein MXAN4899 (Jelsbak et al., 2005) (Fig. 2A). MXAN4899 is required for aggregation, sporulation, and C-signal-dependent gene expression, and genetic evidence suggests that the pathway defined by MXAN4899 converges with the C-signal transduction pathway downstream from FruA accumulation. MXAN4899 contains an N-terminal forkhead-associated domain. Forkheadassociated domains are involved in protein-protein interactions and preferentially bind to phospho-threonine residues in their targets (Li et al., 2000), suggesting that MXAN4899 activity is connected to, and modulated by, Ser/Thr kinase(s). The Ser/Thr kinase(s) potentially
DEVELOPMENT AND MOTILITY involved in regulating MXAN4899 remains to be identified, as does the signal(s) that may regulate the activity of this kinase(s). According to this model, the C-signal transduction pathway is an integration point at which the intercellular signals needed to coordinate multicellular efforts, the intracellular signals reflecting the energy status of individual cells, and possibly other signals are integrated. The advantage of this kind of integration would be that productive C-signaling and, thus, morphogenesis are strictly coordinated with the energy status of individual cells.
THE C-SIGNAL-DEPENDENT MOTILITY RESPONSE Fruiting body formation depends on changes in cell behavior from spreading to aggregation. Therefore, a complete understanding of the morphogenetic properties of the C-signal entails a description of how this signal molecule alters cell behavior. M. xanthus cells move by gliding and harbor two gliding systems (Hodgkin and Kaiser, 1979) (please refer to chapter 6 for a detailed description of the two motility systems). Briefly, in the social motility system motive force is generated by retraction of unipolarly localized type IV pili (Kaiser, 1979; Merz et al., 2000; Skerker and Berg, 2001; Sun et al., 2000; Wu and Kaiser, 1995). In the adventurous motility system, generation of motive force is less well understood. According to the current models, motive force is generated by the secretion of slime from nozzle-like structures located to the pole opposite to that containing type IV pili (Wolgemuth et al., 2002) and/ or by focal adhesion complexes (Mignot et al., 2007). During gliding, the speed is highly variable and, periodically, cells stop and then either resume gliding in the same direction or undergo a reversal in which the head becomes the tail and the tail becomes the head (Blackhart and Zusman, 1985; Jelsbak and Ssgaard-Andersen, 1999, 2002; Spormann and Kaiser, 1995). The molecular mechanism underlying a reversal has been proposed to involve the coordinated polarity switching of the two gliding engines (Kaiser, 2003; Ssgaard-Andersen, 2004; Mignot et al., 2005,2007). Fluorescent time-lapse videomicroscopy has been instrumental in analyzing how the C-signal modulates cell behavior. During the rippling stage cells display an increased reversal frequency (Sager and Kaiser, 1994; Welch and Kaiser, 2001), whereas aggregating cells display a decreased reversal frequency (Jelsbak and Ssgaard-Andersen, 2002). Thus, rippling cells display an oscillatory behavior whereas aggregating cells display a
SIGNALING IN M. XANTHUS 4. CONTACT-DEPENDENT unidirectional type of behavior. For a discussion of how the C-signal may induce opposite effects on the reversal frequency, please refer to chapter 5. Here, emphasis is on C-signal-induced aggregation. The C-signal-dependent decrease in the reversal frequency during the aggregation stage of fruiting body formation is in agreement with the genetic and biochemical evidence demonstrating that the C-signal is an input signal to the Frz chemosensory system (Ssgaard-Andersen and Kaiser, 1996) (Fig. 2A). Consistent with the observation that the C-signal induces a decrease in the reversal frequency, methylation of FrzCD correlates with a low reversal frequency (McBride et al., 1992). The C-signal-dependent decrease in the reversal frequency suggests that the C-signal inhibits polarity switching of the two gliding engines. The net result of this inhibition is that the two motility engines are locked in their polarity and, consequently, C-signaling cells are locked in a unidirectional mode of behavior.
C-SIGNAL-INDUCED AGGREGATIONA MODEL The ability of C-signaling cells to move a long distance is beneficial for aggregation to occur. However, in order for cells to aggregate they also need to move with a sense of direction. The identification of the motility parameters controlled by the C-signal during aggregation in combination with the contact-dependent C-signal transmission mechanism has allowed the generation of a model for C-signal-induced aggregation (Jelsbak and SsgaardAndersen, 2002). The model is composed of three discrete events (Fig. 4). The basic event is an end-to-end contact between two cells with C-signal transmission (Fig. 4A). Cells engaged in end-to-end contact with C-signal transmission gain the ability to move with a low reversal frequency and, thus, travel long net distances. In a field of starving cells, this event is predicted to occur at a high frequency and to result in the second discrete event, chain formation (Fig. 4B): the repeated end-to-end contacts with C-signal transmission are predicted to result in the sequential recruitment of cells into chains in which cells are moving with a low reversal frequency and in the same direction. The direction of movement is determined by the direction of movement of the cell at the leading end of the chain. In a chain, the information about the direction of movement is relayed from the leading to the lagging cell by the direct cellcell contacts. The arrangement and movement of cells in chains are predicted to result in the third discrete event, stream formation (Fig. 4C). Movement of one cell in a chain may create alignment of neighboring cells. This
87 will result in the formation of secondary chains that are associated with the initiating chain by lateral interactions. This could lead to the formation of the streams of cells that have been observed experimentally (Fig. 4D). Aggregation centers could form by collisions of streams or by a single stream turning on itself in a spiral movement. In this model, C-signal transmission is a local event between two cell ends, which occurs without reference to the global cellular pattern, and the result is a global organization of cells. Therefore, according to this model C-signal-induced aggregation is a self-organizing process.
SIGNAL INTEGRATION DURING FRUITING BODY FORMATION Formation of spore-filled fruiting bodies is an effective survival strategy in response to starvation. However, it is also costly, as only 10 to 20% of cells differentiate to spores. Therefore, the decision to initiate fruiting body formation is probably not taken lightly by M. xanthus cells. The stringent response and the intercellular A-signal system constitute two checkpoints, which operate to make sure that cells embark on fruiting body formation only when starvation is severe and anticipated to be long-lasting. The C-signal comes into action when the cells have passed the tests of the stringent response and A-signal. The primary functions of the C-signal are to induce and coordinate aggregation, sporulation, and full gene expression after 6 h. Thus, the C-signaling system cannot be regarded as a system whose primary function it is to monitor starvation. Regarded in this way, the stringent response and the A- and C-signaling systems essentially act positively to stimulate fruiting body formation. Predictably, fruiting body formation is also subject to intensive negative control (Cho and Zusman, 1999; Cusick et al., 2002; Hager et al., 2001; Kirby and Zusman, 2003; Munoz et al., 1991; Rasmussen and Ssgaard-Andersen, 2003; Rasmussen et al., 2005; Udo et al., 1996). The specific parameters monitored by these negative regulators are still unknown. However, the multitude of negative regulators of fruiting body formation indicates that cells are continuously monitoring the nutritional and possibly other conditions to decide whether or not fruiting body formation should continue. From a regulatory point of view, circuits that include the integration of positively as well as negatively acting pathways are highly robust and endow the system with the capacity to tailor the final decision, i.e., to continue or not to continue fruiting body formation, to the specific conditions that cells are exposed to.
DEVELOPMENT AND MOTILITY
88
A.
B.
C.
The basic event
Chain formation
Stream formation
1.
1
D.
1
2.
Change in cell-behavior: Reduction in reversal frequency
3.
Figure 4 Model for C-signal-induced aggregation. (A) The basic event with the end-to-end contact between two cells with C-signal transmission followed by a change in cell behavior. (B) Chain formation. This event is a consequence of end-to-end contacts with C-signal transmission between. The formation of a chain depends on the sequential recruitment of cells as shown in the four panels. Cells engaged in C-signal transmission are shown to move towards an aggregation center indicated in gray to the left with a low reversal frequency. Non-C-signaling cells move with a high reversal frequency as indicated by the double-headed arrow. (C)Stream formation. Movement of cells in a chain is predicted to create alignment of neighboring cells with the formation of secondary chains of cells (marked by dark color). Cells in secondary chains are associated with the primary chain by lateral cell-cell contacts and with other cells in the secondary chain by end-to-end contacts. Together, an initiating chain and its associated secondary chains will make up a stream. (D) Stream formation in vivo. Cell arrangements were visualized by fluorescence microscopy of GFP-labeled cells. GFP-labeled wild-type cells were codeveloped with nonfluorescent wild-type cells at a ratio of 1 to 40. Images were acquired after the indicated hours of starvation. Circles in the 6-h image indicate aggregation centers; arrows indicate streams. Scale bar, 50 ym.
CONCLUDING REMARKS We now have a general framework for understanding how the C-signal induces four responses that are separated in time and space. However, many questions are still unanswered. So far, the identification of components important for development has to a large extent depended on the often painstaking identification of relevant genes and proteins in a stepwise fashion. With the completed M. xanthus genome and functional genomics-based approaches such as proteome analysis (Horiuchi et al., 2002b) and DNA microarray-based analyses (Diodati et al., 2006; Overgaard et al., 2006) in place, the tools are
now available to pave the way for the systematic identification of additional important players in the regulatory pathways that regulate fruiting body formation. Moreover, the establishment of cell-biology-based methods with green fluorescent protein (GFP)fusion proteins and immunofluorescence are likely to result in the detailed understanding of the spatial organization of M. xanthus cells during starvation (Mignot et al., 2005, 2007; S. Leonardy and L. Sagaard-Andersen, unpublished data). I thank Penelope Higgs, Martin Overgaard, Anders Aa. Rasmussen, Sune Lobedanz, Eva Ellehauge, and Lars Jelsbak for many helpful discussions.
4.
CONTACT-DEPENDENT SIGNALINGIN M .
XANTHUS
References Baker, M. E. 1994. Myxococcus xantbus C-factor, a morphogenetic paracrine signal, is similar to Escherichia coli 3-oxoacyl-[acyl-carrier-protein] reductase and human 17betahydroxysteroid dehydrogenase. Biocbem. ]. 301:311-312. Blackhart, B. D., and D. R. Zusman. 1985. “Frizzy” genes of Myxococcus xanthus are involved in control of frequency of reversal of gliding motility. Proc. Natl. Acad. Sci. USA 82:8771-8774. Boysen, A., E. Ellehauge, B. Julien, and L. Ssgaard-Andersen. 2002. The DevT protein stimulates synthesis of FruA, a signal transduction protein required for fruiting body morphogenesis in Myxococcus xantbus. J. Bacteriol. 184:1540-1546. Cho, K., and D. R. Zusman. 1999. Sporulation timing in Myxococcus xanthus is controlled by the espAB locus. Mol. Microbiol. 34:714-725. Crawford, E. W., and L. J. Shimkets. 2000. The Myxococcus xanthus socE and csgA genes are regulated by the stringent response. Mol. Microbiol. 37:788-799. Cusick, J. K., E. Hager, and R. E. Gill. 2002. Characterization of 6csA mutations that bypass two distinct signaling requirements for Myxococcus xanthus development. ]. Bacteriol. 1845141-5150. Diodati, M. E., F. Ossa, N. B. Caberoy, I. R. Jose, W. Hiraiwa, M. M. Igo, M. Singer, and A. G. Garza. 2006. Nla18, a key regulatory protein required for normal growth and development of Myxococcus xantbus. J. Bacteriol. 188:1733-1743. Downard, J., S. V. Ramaswamy, and K. S. Kil. 1993. Identification of esg, a genetic locus involved in cell-cell signaling during Myxococcus xanthus development. J. Bacteriol. 175:7762-7770. Ellehauge, E., M. Nsrregaard-Madsen, and L. SsgaardAndersen. 1998. The FruA signal transduction protein provides a checkpoint for the temporal co-ordination of intercellular signals in M. xantbus development. Mol. Microbiol. 30:807-817. Garza, A. G., J. S. Pollack, B. Z. Harris, A. Lee, I. M. Keseler, E. F. Licking, and M. Singer. 1998. SdeIC is required for early fruiting body development in Myxococcus xanthus. J. Bacteriol. 180:4628-4637. Gronewold, T. M. A., and D. Kaiser. 2001. The act operon controls the level and time of C-signal production for Myxococcus xanthus development. Mol. Microbiol. 40:744-756. Guo, D., M. G. Bowden, R. Pershad, and H. B. Kaplan. 1996. The Myxococcus xanthus rfbABC operon encodes an ATP-binding cassette transporter homolog required for 0-antigen biosynthesis and multicellular development. J. Bacteriol. 178:163 1-1639. Hagen, D. C., A. P. Bretscher, and D. Kaiser. 1978. Synergism between morphogenetic mutants of Myxococcus xantbus. Dev. Biol. 64:284-296. Hager, E., H. Tse, and R. E. Gill. 2001. Identification and characterization of spdR mutations that bypass the BsgA proteasedependent regulation of developmental gene expression in Myxococcus xantbus. Mol. Microbiol. 39:765-780. Harris, B. Z., D. Kaiser, and M. Singer. 1998. The guanosine nucleotide (p)ppGpp initiates development and A-factor
89 production in Myxococcus xantbus. Genes Dev. 12:10221035. Hodgkin, J., and D. Kaiser. 1979. Genetics of gliding motility in Myxococcus xanthus (Myxobacterales): two gene systems control movement. Mol. Gen. Genet. 171:177-191. Horiuchi, T., T. Akiyama, S. Inouye, and T. Komano. 2002a. Analysis of dofA, a fruA-dependent developmental gene, and its homologue, dot%, in Myxococcus xanthus. J. Bacteriol. 184~6803-6810. Horiuchi, T., M. Taoka, T. Isobe, T. Komano, and S. Inouye. 2002b. Role of fruA and csgA genes in gene expression during development of Myxococcus xanthus. Analysis by two-dimensional gel electrophoresis. 1. Biol. Cbem. 27726753-26760. Inouye, M., S. Inouye, and D. R. Zusman. 1979a. Gene expression during development of Myxococcus xanthus: pattern of protein synthesis. Dev. Biol. 68579-591. Inouye, M., S. Inouye, and D. R. Zusman. 197910. Biosynthesis and self-assembly of protein S, a development specifc protein of Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 76:209-2 13. Jelsbak, L., and L. Ssgaard-Andersen. 1999. The cell surfaceassociated intercellular C-signal induces behavioral changes in individual Myxococcus xantbus cells during fruiting body morphogenesis. Proc. Natl. Acad. Sci. USA 96: 5031-5036. Jelsbak, L., and L. Ssgaard-Andersen. 2002. Pattern formation by a cell surface-associated morphogen in Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 99:2032-2037. Jelsbak, L., M. Givskov, and D. Kaiser. 2005. Enhancer-binding proteins with a forkhead-associated domain and the sigma54 regulon in Myxococcus xanthus fruiting body development. Proc. Natl. Acad. Sci. USA 102:3010-3015. Jenal, U. 2004. Cyclic di-guanosine-monophosphate comes of age: a novel secondary messenger involved in modulating cell surface structures in bacteria? Curr. Opin. Microbiol. 7185-1 91. Julien, B., A. D. Kaiser, and A. Garza. 2000. Spatial control of cell differentiation in Myxococcus xantbus. Proc. Natl. Acad. Sci. USA 97:9098-9103. Kaiser, D. 1979. Social gliding is correlated with the presence of pili in Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 7659.52-5956. Kaiser, D. 2003. Coupling cell movement to multicellular development in myxobacteria. Nat. Rev. Microbiol. 1:4554. Kim, S. K., and D. Kaiser. 1990a. Cell alignment required in differentiation of Myxococcus xanthus. Science 249: 926-928. Kim, S. K., and D. Kaiser. 1990b. Cell motility is required for the transmission of C-factor, an intercellular signal that coordinates fruiting body morphogenesis of Myxococcus xantbus. Genes Dev. 42396-904. Kim, S. K., and D. Kaiser. 1990c. Purification and properties of Myxococcus xantbus C-factor, an intercellular signaling protein. Proc. Natl. Acad. Sci. USA 873635-3639. Kim, S. K., and D. Kaiser. 1990d. C-factor: a cell-cell signaling protein required for fruiting body morphogenesis of M. xanthus. Cell 61:19-26.
90 Kim, S. K., and D. Kaiser. 1991. C-factor has distinct aggregation and sporulation thresholds during Myxococcus development. J. Bacteriol. 173:1722-1728. Kirby, J. R., and D. R. Zusman. 2003. Chemosensory regulation of developmental gene expression in Myxococcus xantbus. Proc. Natl. Acad. Sci. USA 100:2008-2013. Kroos, L., A. Kuspa, and D. Kaiser. 1986. A global analysis of developmentally regulated genes in Myxococcus xantbus. Dev. Biol. 117:252-266. Kroos, L., and D. Kaiser. 1987. Expression of many developmentally regulated genes in Myxococcus depends on a sequence of cell interactions. Genes Dev. 1:840-854. Kroos, L., P. Hartzell, K. Stephens, and D. Kaiser. 1988. A link between cell movement and gene expression argues that motility is required for cell-cell signaling during fruiting body development. Genes Dev. 2:1677-1685. Kruse, T., S. Lobedanz, N. M. S. Berthelsen, and L. SsgaardAndersen. 2001. C-signal: a cell surface-associated morphogen that induces and coordinates multicellular fruiting body morphogenesis and sporulation in M. xanthus. Mol. Microbiol. 40:156-168. Kuspa, A., L. Kroos, and D. Kaiser. 1986. Intercellular signaling is required for developmental gene expression in Myxococcus xanthus. Dev. Biol. 117:267-276. Kuspa, A., L. Plamann, and D. Kaiser. 1992a. A-signalling and the cell density requirement for Myxococcus xantbus development. J. Bacteriol. 174:7360-7369. Kuspa, A., L. Plamann, and D. Kaiser. 1992b. Identification of heat-stable A-factor from Myxococcus xantbus. J. Bacteriol. 174:33 19-3 326. Lee, B.-U., K. Lee, J. Mendez, and L. J. Shimkets. 1995. A tactile sensory system of Myxococcus xantbus involves an extracellular NAD(P)+-containing protein. Genes Dev. 9~2964-2973. Lee, K., and L. J. Shimkets. 1994. Cloning and characterization of the socA locus which restores development to Myxococcus xanthus C-signaling mutants. J. Bacteriol. 176: 2200-2209. Lee, K., and L. J. Shimkets. 1996. Suppression of a signaling defect during Myxococcus xanthus development. J. Bacteriol. 178:977-984. Li, J., G. I. Lee, S. R. Van Doren, and J. C. Walker. 2000. The FHA domain mediates phosphoprotein interactions. J. Cell Sci. 113:4143-4149. Li, S., B.-U. Lee, and L. J. Shimkets. 1992. csgA expression entrains Myxococcus xanthus development. Genes Dev. 6:4014 10. Licking, E., L. Gorski, and D. Kaiser. 2000. A common step for changing cell shape in fruiting body and starvation-independent sporulation in Myxococcus xanthus. J. Bacteriol. 182:3553-3558. Lobedanz, S., and L. Ssgaard-Andersen. 2003. Identification of the C-signal, a contact-dependent morphogen coordinating multiple developmental responses in Myxococcus xantbus. Genes Dev. 17:2151-2161. McBride, M. J., R. A. Weinberg, and D. R. Zusman. 1989. “Frizzy” aggregation genes of the gliding bacterium Myxococcus xantbus show sequence similarities to the
DEVELOPMENT AND MOTILITY chemotaxis genes of enteric bacteria. Proc. Natl. Acad. Sci. USA 86:424-428. McBride, M. J., T. Kohler, and D. R. Zusman. 1992. Methylation of FrzCD, a methyl-accepting taxis protein of Myxococcus xantbus, is correlated with factors affecting cell behavior. J. Bacteriol. 174:4246-4257. Merz, A. J., M. So, and M. P. Sheetz. 2000. Pilus retraction powers bacterial twitching motility. Nature 407:98-102. Mignot, T., J. P. Merlie, and D. R. Zusman. 2005. Regulated pole-to-pole oscillations of a bacterial gliding motility protein. Science 3105355-857. Mignot, T., J. W. Shaevitz, P. L. Hartzell, and D. R. Zusman. 2007. Evidence that focal adhesion complexes power bacterial gliding motility. Science 315:853-856. Munoz, D. J., S. Inouye, and M. Inouye. 1991. A gene encoding a protein serinekhreonine kinase is required for normal development of M. xanthus, a gram-negative bacterium. Cell 67~995-1006. Nariya, H., and S. Inouye. 2005. Identification of a protein Ser/Thr kinase cascade that regulates essential transcriptional activators in Myxococcus xanthus development. Mol. Microbiol. 58:367-379. Nariya, H., and S. Inouye. 2006. A protein Ser/Thr kinase cascade negatively regulates the DNA-binding activity of MrpC, a smaller form of which may be necessary for the Myxococcus xantbus development. Mol. Microbiol. 60:1205-1217. O’Connor, K. A., and D. R. Zusman. 1989. Patterns of cellular interactions during fruiting-body formation in Myxococcus xantbus. J. Bacteriol. 171:6013-6024. O’Connor, K. A., and D. R. Zusman. 1991. Development in Myxococcus xantbus involves differentiation into two cell types, peripheral rods and spores. J. Bacteriol. 173:33183333. Ogawa, M., S. Fujitani, X. Mao, S. Inouye, and T. Komano. 1996. FruA, a putative transcription factor essential for the development of Myxococus xanthus. Mol. Microbiol. 22~757-767. Oppermann, U., C. Filling, M. Hult, N. Shafqat, X. Wu, M. Lindh, J. Shafqat, E. Nordling, Y. Kallberg, B. Person, and H. Jornvall. 2003. Short-chain dehydrogenases/reductases (SDR):the 2002 update. Chem. Biol. Interact. 143-144:247253. Overgaard, M., S. Wegener-Feldbriigge, and L. Ssgaard-Andersen. 2006. The orphan response regulator DigR is required for synthesis of extracellular matrix fibrils in Myxococcus xanthus. J. Bacteriol. 188:4384-4394. Plamann, L., A. Kuspa, and D. Kaiser. 1992. Proteins that rescue A-signal-defective mutants of Myxococcus xanthus. J. Bacteriol. 174:3311-3318. Pollack, J. S., and M. Singer. 2001. SdeK, a histidine kinase required for Myxococcus xanthus development. J. Bacteriol. 183:3589-3596. Rasmussen, A. A., and L. Ssgaard-Andersen. 2003. TodK, a putative histidine protein kinase, regulates timing of fruiting body morphogenesis in Myxococcus xantbus. J. Bacteriol. 1855452-5464. Rasmussen, A. A., S. L. Porter, J. P. Armitage, and L. SsgaardAndersen. 2005. Coupling of multicellular morphogenesis
4. CONTACT-DEPENDENT SIGNALINGIN M. XANTHUS and cellular differentiation by an unusual hybrid histidine protein kinase during fruiting body morphogenesis in Myxococcus xanthus. Mol. Microbiol. 56:1358-1372. Rasmussen, A. A., S. Wegener-Feldbriigge, S. L. Porter, J. P. Armitage, and L. Ssgaard-Andersen. 2006. Four signalling domains in the hybrid histidine protein kinase RodK of Myxococcus xanthus are required for activity. Mol. Microbiol. 60525-534. Reichenbach, H. 1965. Rhythmische vorgange bei der Schwarmenfaltung von Myxobakterien. Bey. Dtsch. Bot. Ges. 78: 102-105. Reichenbach, H. 1999. The ecology of the myxobacteria. Enviyon. Microbiol. 1:15-21. Rosenbluh, A., R. Nir, E. Sahar, and E. Rosenberg. 1989. Cell-density-dependent lysis and sporulation of Myxococcus xanthus in agarose beads. J. Bacteriol. 171: 4923-4929. Sager, B., and D. Kaiser. 1994. Intercellular C-signaling and the traveling waves of Myxococcus. Genes Dev. 8: 2793-2804. Shimkets, L. J., and D. Kaiser. 1982. Induction of coordinated movement of Myxococcus xanthus cells. J. Bacteriol. 152~451-461. Shimkets, L. J., R. E. Gill, and D. Kaiser. 1983. Developmental cell interactions in Myxococcus xanthus and the spoC locus. Proc. Natl. Acad. Sci. USA 80:1406-1410. Shimkets, L. J., and H. Rafiee. 1990. CsgA, an extracellular protein essential for Myxococcus xanthus development. J . Bacteriol. 17252994306. Singer, M., and D. Kaiser. 1995. Ectopic production of guanosine penta- and tetraphosphate can initiate early developmental gene expression in Myxococcus xanthus. Genes Dev. 9:1633-1644. Skerker, J. M., and H. C. Berg. 2001. Direct observation of extension and retraction of type IV pili. Proc. Natl. Acad. Sci. USA 98:6901-6904. Ssgaard-Andersen, L., and D. Kaiser. 1996. C factor, a cellsurface-associated intercellular signaling protein, stimulates the cytoplasmic Frz signal transduction system in Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 93:2675-2679. Ssgaard-Andersen, L., F. J. Slack, H. Kimsey, and D. Kaiser. 1996. Intercellular C-signaling in Myxococcus xanthus involves a branched signal transduction pathway. Genes Dev. 10:740-754. Ssgaard-Andersen, L., M. Overgaard, S. Lobedanz, E. Ellehauge, L. Jelsbak, and A. A. Rasmussen. 2003. Coupling gene expression and multicellular morphogenesis during fruiting body formation in Myxococcus xanthus. Mol. Microbiol. 48:l-8.
91 Ssgaard-Andersen, L. 2004. Cell polarity, intercellular signalling and morphogenetic cell movements in Myxococcus xanthus. Curr. Opin.Microbiol. 7587-593. Spormann, A. M., and A. D. Kaiser. 1995. Gliding movements in Myxococcus xanthus. J. Bacteriol. 1775846-5852. Sun, H., D. R. Zusman, and W. Shi. 2000. Type IV pilus of Myxococcus xanthus is a motility apparatus controlled by the frz chemosensory system. Curr. Biol. 10:1143-1146. Sun, H., and W. Shi. 2001a. Genetic studies of mrp, a locus essential for cellular aggregation and sporulation of Myxococcus xanthus. J. Bacteriol. 183:4786-4795. Sun, H., and W. Shi. 2001b. Analyses of mrp genes during Myxococcus xanthus development. J. Bacteriol. 183:67336739. Taylor, B. L., and I. B. Zhulin. 1999. PAS domains: internal sensors of oxygen, redox potential, and light. Microbiol. Mol. Biol. Rev. 63:479-506. Udo, H., M. Inouye, and S. Inouye. 1996. Effects of overexpression of Pkn2, a transmembrane protein serinel threonine kinase, on development of Myxococcus xanthus. J. Bacteriol. 178:6647-6649. Ueki, T., and S. Inouye. 2003. Identification of an activator protein required for the induction of fruA, a gene essential for fruiting body development in Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 100:8782-8787. Ueki, T., and S. Inouye. 2005a. Identification of a gene involved in polysaccharide export as a transcription target of FruA, an essential factor for Myxococcus xanthus development. J. Biol. Chem. 280:32279-32284. Ueki, T., and S. Inouye. 2005b. Activation of a developmentspecific gene, dofA, by FruA, an essential transcription factor for development of Myxococcus xanthus. J. Bacteriol. 187:8504-8506. Ward, M. J., and D. R. Zusman. 1999. Motility in Myxococcus xanthus and its role in developmental aggregation. Cum Opin. Microbiol. 2:624-629. Welch, R., and D. Kaiser. 2001. Pattern formation and traveling waves in myxobacteria: experimental demonstration. Proc. Natl. Acad. Sci. USA 98:14907-14912. Wolgemuth, C., E. Hoiczyk, D. Kaiser, and G. Oster. 2002. How myxobacteria glide. Curr. Biol. 12:369-377. Wu, S. S., and D. Kaiser. 1995. Genetic and functional evidence that Type IV pili are required for social gliding motility in Myxococcus xanthus. 21101. Microbiol. 18: 547-558. Yoder-Himes, D. R., and L. Kroos. 2006. Regulation of the Myxococcus xanthus C-signal-dependent omega4400 promoter by the essential developmental protein FruA. J. Bacteriol. 188:5 167-5 176.
Myxobucteria: Multicellulurity and Differentiation Edited by David E. Whitworth 0 2008 ASM Press, Washington, D.C.
Dale Kaiser
Reversing Myxococcus xanthus Polarity
Colonies of M. xanthus on the surface of agar spread outward, and genetic studies of motility began with the isolation of mutants having abnormal spreading patterns (Burchard, 1970). Two independent gene systems that control gliding were revealed, called A and S (Hodgkin and Kaiser, 1979a, 197913). They were found to specify different gliding engines-the A-engine and the S-engine that are described below and detailed in chapter 6. As a rule, only double mutants having one mutation in system A and one mutation in system S failed to spread at all. Mgl mutants, which were common among Hodgkin’s early isolates, were an exception to the rule, and single mutations in mgZA blocked all spreading. MglA colonies have a sharp edge and a heaped center (Hodgkin and Kaiser, 1979a), whereas wild-type and other motility mutants are flat with an irregular edge. Hodgkin concluded that Mgl function was shared by system A and system S (Hodgkin and Kaiser, 1979a). In 30 years of subsequent investigation, MglA has not been found among the constituents of either the A or the S engine. Indeed, the A engine is working in an mglA mutant; it is likely that the S engine is working as well, although this has not been demonstrated directly. The function of Mgl is suggested by recent microscopic and genetic studies by Rosa Yu on A motility. Whereas all of her motile strains
5
produced slime only at one end of each cell at any point in time, mglA mutants, as well as other nonspreading mutants like cglB, secreted slime from both ends (Kaiser and Yu, 2005; Yu and Kaiser, 2007). Motile strains can produce slime from either end and simply switch the producing end when they reverse. Bipolar slime secretion suggested that by trying to move in both directions simultaneously, cells were unable to make progress in either. This new view of nonmotility is supported by the fact that Mgl cells oscillate rapidly back and forth at very low amplitude (Spormann and Kaiser, 1999).
TWO POLAR ENGINES Since chapter 6 offers a general description of both engines, the components important for engine reversal are emphasized here. In brief, the S engine depends on retraction of type IV pili at the leading pole of a cell (Hodgkin and Kaiser, 1979a, 1979b; Sun et al., 2000; Skerker and Berg, 2001; Kaiser, 2003). Pili attach to extracellular material, termed fibrils, on nearby cells (Behmlander and Dworkin, 1994; Dworkin, 1999; Li etal., 1992;NudlemanandKaiser,2004;Wuetal.,1997). No motion is produced by pilus extension because pili are thin (only 6 nm in diameter) and they bend rather than
Dale Kaiser, Departments of Biochemistry and Developmental Biology, Stanford University Medical School, Stanford, CA 94305.
93
94 penetrate a barrier (Merz and Forest, 2002). Many bacteria, including Pseudomonas, Neisseria, and Synechocystis share a core set of 10 similar type IV pilus genes (Nudleman and Kaiser, 2004). Generally type IV pili are polar structures; they are never seen to emerge from the sides of M. xanthus cells (MacRae and McCurdy, 1976a, 1976b; Kaiser, 1979).Virtually all M. xanthus cells have pili at only one of their poles (Kaiser, 1979),even though both poles are capable of bearing pili. Since pili pull, the piliated pole leads cell movement. The second engine, the A engine, is genetically independent of the S engine, which is to say that all mutants lacking S (S-) examined retain A motility. Pilus-independent gliding engines have been found in other bacteria, including Cytophaga and Flexibacter, but their mechanics are unknown (McBride, 2001). Gliding M. xunthus cells leave a visible trail of slime behind them when they glide on agar (Burchard, 1982; Rodriguez and Spormann, 1999). By electron microscopy E. Hoiczyk saw amorphous filaments, apparently of slime, that emerged from only one end of an M. xunthus cell. Light microscopy by Rosa Yu also revealed a single amorphous filament at one pole of wild-type cells that resembled the images of Hoiczyk (Fig. 1A). Hoiczyk also observed several hundred thick-walled ring structures, which he suggested were nozzles for slime secretion at both cell poles (Wolgemuth et al., 2002). Relatively few rings were found in nonpolar areas; 80% of the rings were polar. Thus, the microscopical evidence clearly shows that the A engine is unipolar, even though the nozzles (rings) are bipolar.
A AND S ENGINES COOPERATE When a circular patch of cells is deposited on agar, the cells spread outward at a roughly constant rate for many days (Burchard, 1974; Kaiser and Crosby, 1983).
DEVELOPMENT AND MOTILITY The increase in diameter of the patch reflects active cell movement, rather than growth, for the colony of an A-S- mutant, which lacks both engines, expands (due to cell division) at less than one-eighth the rate of an A’S+ colony (Kaiser and Crosby, 1983). Vibrio parahaemolyticus and other bacteria swarm over surfaces by using flagella (McCarter, 1995), but how the rotating flagella produce swarming is not understood. M. xanthus has no flagella and spreads over surfaces by using its A and S engines. Since their spreading gives rise to large, thin, and delicate structures, Reichenbach called those structures swarms (Reichenbach, 1984). His designation is biologically appropriate because it calls attention to the underlying interactions between swarming cells, as found in a “swarm” of bees. An mglA mutant, which is unable to reverse, cannot swarm; its colonies are mounded and have sharp edges (Stephens et al., 1989); therefore, swarming by M. xanthus requires reversal. Moreover, frzE mutants, which seldom reverse (Shi and Zusman, 1995), form smaller colonies than the wild type. These colonies have the abnormal frizzy structure, unlike normal swarms. Together the two engines promote swarming in A’S+ cells at three times the rate (1.6 p d m i n ) of cells that have one or the other engine alone (0.6 or 0.4 p d m i n [Fig. 21). Moreover, the data of Fig. 2 show that the maximum swarming rate of A+S+is 50% greater than the sum of the two single-engine rates (1.6 versus 0.6 + 0.4 p d m i n ) . This implies that the two engines are arranged so as to move a cell in the same direction. Since pili pull while the A engines are believed to push by secreting slime, they must be positioned at opposite ends of the cell: the pulling pili at the leading pole of the cell and the pushing A engines at the trailing pole. The swarm expansion rate data of Fig. 2 imply that the A engines and the S engines always help each other. Therefore, each cell is constructed so that its two engines are positioned at opposite poles.
A
DYNAMICS OF REVERSAL
Figure 1 Polarized slime secretion. (A) Wild type; (B) mglA mutant; ( C ) mglB mutant. Photographs from Yu and Kaiser, 2007.
Reichenbach’s movies of myxobacterial swarms show many individual cells moving alternately along both directions of their long pole-to-pole axis (Kuhlwein and Reichenbach, 1968). A movie of single cells reversing, made by Lars Jelsbak (online movie available in Kaiser, 2003), shows that cells simply stop momentarily before moving off in the opposite direction. Cell-tracking experiments make the same point (Jelsbak and SDgaard-Andersen, 2002; Spormann and Kaiser, 1999). Cells reverse at 8-min intervals on average. Not all stops lead to reversal, but when they do, head and tail switch within roughly 1 min and they appear to switch at the
5 . REVERSING M.XANTHUS POLARITY
95
1.8
1.4
1.6
1.0
-:
0.8 0.6 0.4'
0.2
Ol. 0
I
1
I
I
100
200
300
400
500
1000
DENSITY UNITS
0.9
I
I
I
I
0.8 0
I
a 0
"
0
w 0.f
0.3 0.2
0
I
I
1
I
100
200
300
400
500
1000
DENSITY UNITS
Figure 2 (A) Rate of swarm expansion versus initial cell density for the A'S' strain DK1622. Points are shown for six independent experiments. The best-fitting smooth curve has the following form: rate = 0.1 + 1.48 (1- e-densityi48). Reproduced from Kaiser and Crosby, 1983. (B) Rate of swarm expansion versus initial cell density for three A-S+ strains (closed symbols) and two A+S- strains (open symbols). Points are shown for six independent experiments. The smooth curve for the A-S+ has the following form: rate = 0.47 (1- e-densiv/190). The smooth curve for the A'S- has the following form: rate = 0.1 + 0.52 (1- e-density/20).
Singer, 2005). Since both engines are placed specifically to move cells in a particular direction, reversal of gliding direction involves moving both engines from one end to the other. When growing cells divide, both daughters are motile. Accordingly each new cell end gains an A engine or an S engine, whichever is opposite the old end retained by the daughter. Obviously, each cell has the capacity to synthesize both kinds of engines and to place them appropriately. Each frizzy mutant of M. xanthus, discussed in chapter 7, has a well-defined average frequency of gliding reversal that is inherited (Astling et al., 2006). The frz operon encodes a cytoplasmic chemosensory pathway that controls the frequency of reversal (Blackhart and Zusman, 1985a, 1985b). Frizzy proteins are related by amino acid sequence to the chemotaxis proteins of enteric and other bacteria (Bustamante et al., 2004). While the methyl-accepting chemosensory proteins (MCP)of chemotactic cells are membrane receptors, the MCP homologue in M. xanthus, FrzCD, is a cytoplasmic protein (McCleary et al., 1990; McCleary and Zusman, 1990a, 1990b). FrzCD lacks transmembrane and extracellular domains, and its amino-terminal domain bears no resemblance to the chemotaxis MCPs (Bustamante et al., 2004). FrzE, which receives a signal from FrzCD (Fig. 3 ) , has one domain homologous to a cheA histidine protein kinase, and it catalyzes its own phosphorylation (McCleary et al., 1990; McCleary and Zusman, 1990a, 1990b). FrzE also transfers that phosphate to its response regulator domain for output (Acuna et al., 1995). Although on average wild-type cells reverse at intervals of 8 min, a fyzE null mutant has a reversal period of 2 h. This is consistent with phosphorylated FrzE (FrzE-P) sending a reversal signal to change the direction of movement. A mutant that lacks the C-terminal end of FrzCD reverses at intervals of 2 min, as if methyl-FrzCD induced the phosphorylation of FrzE (Blackhart and Zusman, 1985a). The frizzy signaling circuit is summarized in Fig. 3 .
FrzCD
same time. Mignot and coworkers have observed pilus proteins switching periodically from one pole to the other (Mignot et al., 2005). Individual cells show no clear preference for either direction (Spormann and Kaiser, 1999; Tieman et al., 1996). Reversals are unrelated to the cell cycle because 20 or more reversals of gliding direction can be made during development (Welch and Kaiser, 2001), while DNA is replicating once (Tzeng and
FrzE-P
-
Reversal of polarity
Figure 3 The frizzy signaling pathway for vegetative cells. When FrzCD is methylated, it triggers the phosphorylation of FrzE. FrzE-P signals a reversal of polarity.
DEVELOPMENT AND MOTILITY
96
REVERSAL CLOCK C-signal, a 17-kDa cell surface protein, regulates the reversal frequency during development (Kim and Kaiser, 1990; Lobedanz and Sargaard-Andersen, 2003). How C-signaling gives rise to two different reversal patterns is explained by the signal transduction circuit, shown in Fig. 4. (For a detailed discussion of C-signaling, the reader is referred to chapter 4.) Briefly, the FruA response regulator is synthesized shortly after cells recognize starvation in response to the A-signal (Ellehauge et al., 1998; Ogawa et al., 1996). In response to reception of the C-signal, FruA is posttranslationally modified by phosphorylation (Ellehauge et al., 1998). FruA-P induces the methylation of FrzCD and then the phosphorylation of FrzE. At the beginning of fruiting body development, when the C-signal level is low and correspondingly small amounts of FruA-P are produced, cells respond by reversing their gliding direction. This response leads to developmental traveling waves (Igoshin et al., 2001; Welch and Kaiser, 2001). C-signal is required for traveling waves (Li et al., 1992; Sager and Kaiser, 1994). When two wave crests happen to collide within a system of traveling waves, cells in each crest signal cells in the other, and both sets of cells reverse their gliding direction. To analyze the role of C-signal in wave formation, the signal transduction circuit of Fig. 4 was simulated
mathematically. Unexpectedly, the simulation failed to produce traveling waves unless a negative-feedback loop from FrzE-P back onto the methylation of FrzCD was added. FrzE-P could inhibit the methylation of FrzCD, or it could enhance the methyltransferase. In either case, the negative-feedback loop, indicated in Fig. 4, causes the levels of the Frz intermediates to oscillate. Dubbed the “Frizilator,” the oscillator had the necessary refractory period (Igoshin et al., 2001) and the necessary time delay (Borner et al., 2002). Both the heaped-up traveling waves formed early in fruiting body development (Igoshin et al., 2001) and the accordion waves that form in a single layer of cells in culture (Sliusarenko et al., 2006) were found to depend upon the same negative feedback. Both sorts of waves involve synchronizing the pair of Frizilators in two cells that are C-signaling to each other, like those shown in Fig. 4. Cells moving in synchrony sharpen the crest of a wave and delineate it (Sliusarenko et al., 2006). Stevens and Sargaard-Andersen point out that the entire chain of molecular events that lead from C-signal reception to a gliding reversal, including the proposed negative-feedback loop, needs to be clarified biochemically (Stevens and S~gaard-Andersen,2005), which is correct, but they found no fault with the regulatory logic. They raise the possibility that the phenotypic effect of deleting the C-terminal of the FrzCD protein
A-signal
:
Reversal of gliding direction by inactivating old engines
csgA C-signal on cell surface
C-signal sensor (hypothetical)
Figure 4 C-signal transduction circuit. Early in development, reception of A-signal triggers FruA expression. When a cell receives C-signal by contact with another cell, FruA is phosphorylated. Double-headed open arrows are shorthand for a pair of reversed arrows as in Fig. 3 . Negative feedback, indicated by a minus sign (-) in the arrow from FrzE-P to the arrows between FrzCD and Me-FrzCD, causes the frizzy signaling circuit to oscillate. Phosphorylated FruA drives the oscillator, giving it a precise period. Also, when a cell receives C-signal, ActB is phosphorylated and the expression of csgA is increased. This raises the number of C-signal molecules on the cell surface. The arrow from FrzE-P to inactivation of old engines could have more than one step.
5. REVERSING 211. X A N T H U S POLARITY may be at odds with the Frizilator. However, until the structure of the C terminus of the FrzCD protein and its interactions with the FrzF methyltransferase, the FrzG methylesterase, and FrzE-P are clarified, there is no way to predict what the effect of such a deletion should be; the phenotype of the FrzCD mutant calls attention to the feedback but casts no doubt on the Frizilator itself. Despite the unknowns, the Frizilator explains two sorts of traveling waves semiquantitatively. Without further assumptions, the Frizilator also predicts a transition from waves to streaming that matches the experimental observations remarkably well (Stevens and Ssgaard-Andersen, 2005). The transition arises from a positive feedback in the C-signaling circuit that is generated by the act operon (Ellehauge et al., 1998; Gronewold and Kaiser, 2001; Ssgaard-Andersen and Kaiser, 1996; Ssgaard-Andersen et al., 1996).Whenever two cells make end-to-end contact and signal each other, expression of the csgA gene increases and more C-signal is produced on the surfaces of both cells (Gronewold and Kaiser, 2001,2002). act appears in Fig. 4. Increased expression of csgA requires ongoing starvation (Crawford and Shimkets, 2000). Each collision of wave crests adds more C-signal to the surface of the pairs of signaling cells. After many collisions the number of C-signal molecules on the cell surface rises to the threshold for streaming. Reversals are suppressed in streaming cells, and consequently their overall gliding speed increases (Jelsbak and Ssgaard-Andersen, 1999, 2000). At low levels of C-signaling, the rate-limiting step in the feedback loop of the Frizilator is the methylation of FrzCD and consequently there are waves, whereas at high levels of C-signaling the demethylation of FrzCD-CH, becomes rate limiting and there are streams (Igoshin et al., 2004). Since the demethylation rate determines the oscillation frequency, oscillations cease at high levels of C-signaling. As diagrammed in Fig. 5, at the streaming threshold, the large number of C-signal molecules per cell stops the Frizilator from oscillating. Moreover, it stops with FrzE in its unphosphorylated state (Igoshin et al., 2004). There being no FrzE-P, there is no signal to reverse, and consequently cells continue to move in their previous direction. This establishes the second developmental pattern of reversals-streaming. A stream is a chain of moving cells that make frequent end-to-end contact with the other cells just ahead or just behind them in their chain, due to their preference for gliding along their axis (Jelsbak and Sargaard-Andersen, 2002; Sager and I
97 Developmental time
4
FrzCD methylation rate
Figure 5 Oscillation frequency of the Frizilator as a function of the level of FruA-P, the signaling strength, measured as the level of FrzCD methylation. Oscillation ceases above a critical level of signaling as described in the text. Modified from Igoshin et al., 2004.
et al., 2001; Li et al., 1992). Once morphologically differentiated, myxospores are incapable of movement on their own. This program for development is based on experimental observations (Kaiser and Welch, 2004). The adequacy of this particular set of propositions to produce spherical fruiting bodies that contain spores has been demonstrated by two simulations (Sozinova et al., 2005,2006).
MglA IS SWITCHED BY THE FRIZILATOR Rosa Yu isolated and characterized many new nullmotility mutants and from the mutant phenotypes was able to draw two general conclusions about A-motility. First, all of her mutant strains that retained some A-motility, including mglB mutants, produced slime only at one end of each cell. The perfect correlation between A-motility and unipolar slime secretion is the basis for the conclusion that working A engines are to be found only at one pole at any instant in time. The second conclusion was that mglA and all other “nonmotile” mutants that she had detected were secreting slime from both ends (Kaiser and Yu, 2005; Yu and Kaiser, 2007). These mutants produce sharp-edged colonies, and a sample of Yu’s critical images are presented in Fig. 1. Bipolar slime secretion in the nonmotile mutants suggested that the cells were unable to make progress in either direction because they were secreting slime from both ends. MglA mutant cells had been observed to oscillate rapidly back and forth (Spormann and Kaiser, 1999). However, with
DEVELOPMENT AND MOTILITY
98 each oscillation they move less than 1.5 pm, which is less than one-fifth of a cell length, and thus can make no progress in either direction (Spormann and Kaiser, 1999).We suggest that the rapid reversals of mglA mutants are not due to signals from the reversal generator but are a statistical consequence of active slime secretion from both ends. Several hundred slime nozzles are visible at both cell ends, but to judge from the number of slime ribbons observed (Wolgemuth et al., 2002),fewer than 100 nozzles are working at any particular moment. The speed of AmglB cells (which have partial A-motility [Yu and Kaiser, 20071) fluctuates from 1 to >5.5 p d m i n (Spormann and Kaiser, 1999), and the variations in speed are probably due to variations in the number of nozzles that are working. Whether or not that is the reason for the observed fluctuations in A-engine speed, the speed distribution of AmglAB cells is predicted to be the difference between the speeds of two independent AmglB ends, if the statistical hypothesis is correct. Indeed, this expectation is borne out by Wolgemuth’s calculation shown in Fig. 6 (C. Wolgemuth, personal communication). The good agreement between observed and predicted speeds supports the suggestion that the rapid reversals are not due to switching engines but are the statistical consequence of simultaneous slime secretion from both ends.
50
The conclusion is that MglA mutants are not defective in either the A engine or the S engine but are unable to switch the polarity of their engines. Furthermore, since Amotile cells have slime at only one pole but mglA mutants have slime at both, it follows that the normal function of MglA protein is to inactivate engines. In terms of switching A-engine polarity, the function of MglA is to inactivate the old A engines. Because the A engines and the S engines are coordinately reversed, the old S engines are expected to be inactivated as well. A clear prediction of this scheme is that Mgl mutants will have pili at both ends, just as they have slime at both ends; however, this prediction has not yet been tested. If MglA inactivates the old engines, how are the new engines installed? Because growing and dividing cells routinely assemble new engines at both newly synthesized ends, engine assembly must be directed by the normal polarity inherent in myxobacterial cell growth, as illustrated in Fig. 7. A template in each cell causes new engines to be built at opposite poles without additional instruction as shown in Fig. 7. It is proposed that, as MglA specifically directs destruction of the old A engines and old S engines, normal growth processes are installing new engines at the poles opposite the old. A signal to inactivate old engines could come, directly or indirectly, from FrzE-P, as suggested
I
40 h
8 2 30 v
s
Lc
0 S
.-0 5 20 !$
LL
10
0 0
1
2
3
4
5
6
Speed (pm/min) Figure 6 Observed and predicted distribution of speeds of individual AmglAB cells. Black bars, observed data from Spormann and Kaiser, 1999. Gray bars, predicted as the difference in speed of two independent ends, each end distributed as observed for AmglB in Spormann and Kaiser, 1999.
5. REVERSING M . XANTHUS POLARITY
99
,
Incipient septum
old
new
new
Old
Figure 7 Growth produces two new polar engines that are always compatible with the polarity of the old ends.
in Fig. 4. MglA is a small Ras-like G-protein (Hartzell and Kaiser, 1991a, 1991b), and MglB is its putative guanine nucleotide release protein (GNRP),according to the GenBank record for protein AAA25388. By analogy to the activation of Ras by receptor tyrosine kinase Sev and GNRP Sos in Drosophila eye development (Simon et al., 1991) and to the activation of the flagellar switch complex in enteric bacteria by CheY-P (Stock et al., 1995), FrzE-P is proposed to activate (directly or indirectly) an MglAB switch for destruction of the old polar A and S engines. According to this view, FrzS would be localized as part of assembly of the new S engine, and this helps to explain why FrzS tracks the piliated pole without being required for reversal (Mignot et al., 2005). Both FrzE (McCleary et al., 1990; McCleary and Zusman, 1990a, 1990b) and MglA proteins (Hartzell and Kaiser, 1991a, 1991b)are found in the cytoplasm, where they would be able to interact with each other, and where MglA-GTP could access the cytoplasmic face of both poles for coordinately inactivating the A engine at one and the S engine at the other. MglA protein is found to be unstable in a AmglB mutant (Hartzell and Kaiser, 1991a, 1991b) as if MglA is prone to degradation, unless MglB is there to protect it, which fits an association between the two proteins. Spormann observed that the very high reversal frequency of AmglAB mutants is epistatic to the low reversal frequency of frzE mutants, consistent with an MglA location downstream of FrzE-P (Spormann and Kaiser, 1999) and with finding slime secretion from both poles of an mglA mutant, as illustrated in Fig. 1.
SOME ENGINE PROTEINS ARE UNIPOLAR AND OTHERS ARE BIPOLAR The S engines appear to be prepared for periodic inactivation and resynthesis. For example, pilA, which encodes the pilin monomer, is one of the most highly
transcribed genes in M. xanthus (Wu and Kaiser, 1997; Jelsbak and Kaiser, 2005). Most other proteins of the S engines remain intact and in position even as pili at the old poles disappear when the cell reverses. FrzS, which is associated with the S engine, is said to move from the old piliated pole to the new one (Mignot et al., 2005). However, synthesis of new FrzS for the new piliated pole has not been ruled out, and it is not clear that FrzS is required for pilus function. Like pili and FrzS, Tgl protein is found at only one cell pole, but Tgl is required for S motility. Moreover, Tgl is stimulatable whereas the other two proteins are not known to be (Nudleman et al., 2005). Stimulation implies that a newly piliated pole is able to use newly synthesized Tgl, because Tgl stimulation involves the transfer of Tgl protein from one cell to another with very high efficiency (Nudleman et al., 2005). Tgl, which has six tandem tetratrico peptide repeats for association with PilQ monomers, brings about assembly of the secretin into a gated channel that opens to allow the pilus to slide through (Nudleman et al., 2006). Assembled channels are very resistant to dissociation; they remain oligomeric in heated detergent solution, even though the monomers are not covalently linked (Nudleman et al., 2006). Since PilQ disassembles at the old piliated pole (Nudleman et al., 2006), the resistance of an assembled structure suggests that Tgl must be eliminated to disassemble the secretin. Immunochemical labeling experiments show that, in contrast to Tgl and FrzS, PilQ protein remains in place after reversal (Nudleman et al., 2005, 2006). Both poles exhibit fluorescent patches of PilQ that are superficially similar, and little PilQ fluorescence was evident along the sides of cells. Evidently PilQ monomers are not free to diffuse in the outer membrane. What holds PilQ in a patch at the nonpiliated pole? That patch of PilQ monomers dissociates into monomers in heated detergent because approximately one half of the PilQ in extracts of whole cells
DEVELOPMENT AND MOTILITY
2 00
migrates in sodium dodecyl sulfate gel electrophoresis as monomers and the other half migrates as assembled oligomers (Nudleman et al., 2006). Clearly the monomers are not covalently bonded to each other; they must be held together some other way. In fact, both polar patches of PilQ are associated with several other components of the pilus apparatus: PilM, PilN, PilO, Pill’, and PilG. These pi1 proteins are required for Tgl stimulation probably because they are needed to transport Tgl to the outer membrane, from which location they can be transferred by stimulation (Nudleman et al., 2006). Three other proteins, PilB, PilC, and PilT, of type IV pili that are highly conserved across bacterial species are located in the periplasm and inner membrane; they have been localized to both cell poles in Pseudomonus ueruginosu (Chiang et al., 2005). Most likely all these pi1 proteins are part of an assembly that extends outward from the inner membrane, passes through the periplasm, and attaches to the cell’s rigid peptidoglycan meshwork, providing the necessary mechanical rigidity for pilus retraction with forces in excess of 100 pN (Merz and Forest, 2002). Such an assembled multiprotein complex might hold PilQ monomers in a patch in the outer membrane. It is suggested that the complex (without pili or Tgl protein) survives S-engine inactivation, waiting to become part of a new pilus at the next reversal of polarity. The A engines also appear to be prepared for periodic inactivation and resynthesis. CglB stimulation of Amotility is due to a highly efficient transfer of CglB protein from donor to recipient cells (Nudleman et al., 2005). A-engine nozzles, viewed in cross sections as thick rings, are plentiful at both cell poles (Wolgemuth et al., 2002). However, according to the microscopic observations of Hoiczyk (Wolgemuth et al., 2002) and of Rosa Yu, shown in Fig. 1, at any given moment, slime is secreted from one pole only; evidently nozzles at the other end are not actively secreting. The thick walls of the rings and their uniform structure support the proposal that they are secretion nozzles (Wolgemuth et al., 2002). Nozzles in Phormidium pass through the peptidoglycan meshwork in the periplasm (Hoiczyk and Baumeister, 1998) and are likely to be anchored to that meshwork for the same mechanical reason suggested above for pili in the S engine. CglB, a stimulatable outer membrane lipoprotein essential for A-motility (Hodgkin and Kaiser, 1977; Nudleman et al., 2005; Rodriguez and Spormann, 1999; Simunovic et al., 2003), may be a pole-specific component of the slime nozzles. It is suggested that nozzles not secreting slime survive A-engine inactivation and wait to secrete slime until the next reversal. The two gliding engines seem to be controlled similarly by a common reversal generator that has the
capacity to distinguish old (used) engines from new engines that have not yet been completed. How old and new engines are distinguished is yet to be determined, but old S engines could, in principle, be distinguished by the processed pilin in their cytoplasmic membrane, remaining there from pilus retraction.
MECHANISM OF GLIDING REVERSAL The mechanism of gliding reversal that is proposed here explains how Mgl is shared by two engines that have no protein molecules in common. The frz reversal clock (the Frizilator) produces a pulse of frzE-P that signals an MglAB complex to bind GTP and to become active. Active MglA-GTP then recognizes the several A engines and S engines that are currently in use. GTP is hydrolyzed, and those engines are specifically inactivated. Meanwhile both sets of new engines are being completed at the opposite ends of the cell by the mechanisms associated with cellular growth. On completion of either or both new engines, the cell begins to glide in the opposite direction.
FUTURE CHALLENGES Many questions remain to be answered. Can the A engine nozzles be isolated from motile cells to test whether they secrete slime? How does frzE-P activate the mglAB switch? How are old and new engines distinguished from each other? Does the mglAB switch activate proteolysis? If there is proteolysis, what is the protease and what are its substrates?
References Acuna, G., W. Shi, K. Trudeau, and D. Zusman. 1995. The cheA and cheY domains of Myxococcus xanthus FrzE function independently in vitro as an autokinase and a phosphate acceptor, respectively. FEBS Lett. 358:31-33. Astling, D. P., J. Y. Lee, and D. R. Zusman. 2006. Differential effects of chemoreceptor methylation-domain mutations on swarming and development in the social bacterium MyxococGUS xanthus. Mol. Microbiol. 59:45-55. Behmlander, R. M., and M. Dworkin. 1994. Biochemical and structural analyses of the extracellular matrix fibrils of Myxococcus xanthus. J. Bacteriol. 176:6295-6303. Blackhart, B. D., and D. R. Zusman. 1985a. Cloning and complementation analysis of the frizzy genes of Myxococcus xanthus. Mol. Gen. Genet. 198:243-254. Blackhart, B. D., and D. Zusman. 1985b. Frizzy genes of Myxococcus xanthus are involved in control of frequency of reversal of gliding motility. Proc. Natl. Acad. Sci. USA 82:8767-8770. Borner, U., A. Deutsch, H. Reichenbach, and M. Bar. 2002. Rippling patterns in aggregates of myxobacteria arise from cell-cell collisions. Phys. Rev. Lett. 89:078 101.
5. REVERSING 111. XANTHUS POLARITY Burchard, R. P. 1970. Gliding motility mutants of Myxococcus xanthus. 1.Bacteriol. 104:940-947. Burchard, R. P. 1974. Growth of surface colonies of the gliding bacterium Myxococcus xanthus. Arch. Microbiol. 96:247254. Burchard, R. P. 1982. Trail following by gliding bacteria. J. Bacteriol. 152:495-501. Bustamante, V. H., I. Martinez-Flores, H. C. Vlamakis, and D. Zusman. 2004. Analysis of the Frz signal transduction system of Myxococcus xanthus shows the importance of the conserved C-terminal region of the cytoplasmic chemoreceptor FrzCD in sensing signals. Mol. Microbiol. 53:15011513. Chiang, P., M. Habash, and L. L. Burrows. 2005. Disparate su bcellular localization patterns of Pseudomonas aeruginosa type IV pilus ATPases involved in twitching motility. J. Bacteriol. 187:829-839. Crawford, E. W., Jr., and L. J. Shimkets. 2000. The Myxococcus xanthus socE and csgA genes are regulated by the stringent response. Mol. Microbiol. 37:788-799. Dworkin, M. 1999. Fibrils as extracellular appendages of bacteria: their role in contact-mediated cell-cell interactions in Myxococcus xanthus. BioEssays 2 1:590-595. Ellehauge, E., M. Norregaard-Madsen, and L. SsgaardAndersen. 1998. The FruA signal transduction protein provides a checkpoint for the temporal coordination of intercellular signals in M . xanthus development. Mol. Microbiol. 3 0:807-8 13. Gronewold, T. M. A., and D. Kaiser. 2001. The act operon controls the level and time of C-signal production for M . xanthus development. Mol. Microbiol. 40:744-756. Gronewold, T. M. A., and D. Kaiser. 2002. act operon control of developmental gene expression in Myxococcus xanthus. J. Bacteriol. 184:1172-1179. Hartzell, P., and D. Kaiser. 1991a. Upstream gene of the mgl operon controls the level of mglA protein in Myxococcus xanthus. J. Bacteriol. 173:7625-7635. Hartzell, P., and D. Kaiser. 1991b. Function of MglA, a 22kilodalton protein essential for gliding in Myxococcus xanthus. J. Bacteriol. 173:7615-7624. Hodgkin, J., and D. Kaiser. 1977. Cell-to-cell stimulation of movement in nonmotile mutants of Myxococcus. Proc. Natl. Acad. Sci. USA 74:2938-2942. Hodgkin, J., and D. Kaiser. 1979a. Genetics of gliding motility in M. xanthus (Myxobacterales):two gene systems control movement. Mol. Gen. Genet. 171:177-191. Hodgkin, J., and D. Kaiser. 197910. Genetics of gliding motility in M . xanthus (Myxobacterales):genes controlling movement of single cells. Mol. Gen. Genet. 171:167-176. Hoiczyk, E., and W. Baumeister. 1998. The junctional pore complex, a prokaryotic secretion organelle, is the molecular motor underlying gliding motility in cyanobacteria. Curr. Biol. 8:1161-1168. Igoshin, O., A. Mogilner, R. Welch, D. Kaiser, and G. Oster. 2001. Pattern formation and traveling waves in myxobacteria: theory and modeling. Proc. Natl. Acad. Sci. U S A 98~14913-14918. Igoshin, O., A. Goldbetter, D. Kaiser, and G . Oster. 2004. A biochemical oscillator explains the developmental progression
2 02 of myxobacteria. Proc. Natl. Acad. Sci. USA 101:1576015765. Jelsbak, L., and L. Ssgaard-Andersen. 1999. The cell-surface associated C-signal induces behavioral changes in individual M. xanthus cells during fruiting body morphogenesis. Proc. Natl. Acad. Sci. USA 965031-5036. Jelsbak, L., and L. Ssgaard-Andersen. 2000. Pattern formation: fruiting body morphogenesis in Myxococcus xanthus. Curr. Opin. Microbiol. 3:637-642. Jelsbak, L., and L. Ssgaard-Andersen. 2002. Pattern formation by a cell-surface associated morphogen in M. xanthus. Proc. Natl. Acad. Sci. USA 99:2032-2037. Jelsbak, L., and D. Kaiser. 2005. Regulating pilin expression reveals a threshold for type IV pilus assembly in Myxococcus xanthus. 1.Bacteriol. 187:2105-2112. Kaiser, A. D. 1979. Social gliding is correlated with the presence of pili in Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 76~5952-5956. Kaiser, A. D., and C. Crosby. 1983. Cell movement and its coordination in swarms of Myxococcus xanthus. Cell Motil. 3 :227-245. Kaiser, D. 2003. Coupling cell movement to multicellular development in myxobacteria. Nut. Rev. Microbiol. 1:45-54. Kaiser, D., and R. Welch. 2004. Dynamics of fruiting body morphogenesis. J. Bacteriol. 186:9 19-927. Kaiser, D., and R. Yu. 2005. Reversing cell polarity: evidence and hypothesis. Curr. Opin. Microbiol. 8:216-221. Kim, S. K., and D. Kaiser. 1990. Purification and properties of Myxococcus xanthus C-factor, an intercellular signaling protein. Proc. Natl. Acad. Sci. USA 87:3635-3639. Kim, S. K., and I). Kaiser. 1991. C-factor has distinct aggregation and sporulation thresholds during Myxococcus development. J. Bacteriol. 173:1722-1728. Kruse, T., S. Lobendanz, N. M. S. Bertheleson, and L. SsgaardAndersen. 2001. C-signal: a cell surface-associated morphogen that induces and coordinates multicellular fruiting body morphogenesis and sporulation in M. xanthus. Mol. Microbiol. 40:156-168. Kuhlwein, H., and H. Reichenbach. 1968. Swarming and Morphogenesis in Myxobacteria. Film C893/1965. Institut fur den Wissenschaftliche Film, Gottingen, Germany. Li, S., B. U. Lee, and L. Shimkets. 1992. csgA expression entrains Myxococcus xanthus development. Genes Dev. 6:401-410. Lobedanz, S., and L. Ssgaard-Andersen. 2003. Identification of the C-signal, a contact-dependent morphogen coordinating multiple developmental responses in Myxococcus xanthus. Genes Dev. 17:2151-2161. MacRae, T. H., and H. D. McCurdy. 1976a. Motility related fimbriae in the gliding organism Myxococcus xanthus. Can. J. Microbiol. 22:1589-1593. MacRae, T. H., and H. D. McCurdy. 1976b. Gliding motility mutants of Myxococcus xanthus. Can. J. Microbiol. 22:1282-1292. McBride, M. J. 2001. Bacterial gliding motility: multiple mechanisms for cell movement over surfaces. Annu. Rev. Microbiol. 5 5 :49-75. McCarter, L. L. 1995. Genetic and molecular characterization of the polar flagellum of Vibrio parahaemolyticus. 1.Bacterial. 1771595-1609.
DEVELOPMENT AND MOTILITY
102 McCleary, W. R., M. J. McBride, and D. R. Zusman. 1990. Developmental sensory transduction in Myxococcus xanthus involves methylation and demethylation of frzCD. J. Bacteriol. 172:48 77-4 887. McCleary, W. R., and D. R. Zusman. 1990a. Purification and characterization of the Myxococcus xanthus FrzE protein shows that it has autophosphorylation activity. J. Bacteriol. 172:6661-6668. McCleary, W. R., and D. R. Zusman. 1990b. FrzE of Myxococcus xanthus is homologous to both CheA and CheY of Salmonella typhimurium. Proc. Natl. Acad. Sci. USA 87~5898-5902. Merz. A. "1.., and K. T. Forest. 2002. Bacterial surface motility: slime trails, grappling hooks and nozzles. Curr. Biol. 12: R297-R303. Mignot, T.,J. P. Merlie, and D. Zusman. 2005. Regulated poleto-pole oscillations of a bacterial gliding motility protein. Science 310:855-857. Nudleman, E., and D. Kaiser. 2004. Pulling together with type IV pili. J. Mol. Microbiol. Biotechnol. 752-62. Nudleman, E., D. Wall, and D. Kaiser. 2005. Cell-to-cell transfer of bacterial outer-membrane lipoproteins. Science 309~125-127. Nudleman, E., D. Wall, and D. Kaiser. 2006. Polar assembly of the type IV pilus secretin in Myxococcus xanthus. Mol. Microbiol. 60:16-29. Ogawa, M., S. Fujitani, X. Mao, S. Inouye, and T. Komano. 1996. FruA, a putative transcription factor essential for the development of Myxococcus xanthus. Mol. Microbiol. 22~757-767. Reichenbach, H. 1984. Myxobacteria: a most peculiar group of social prokaryotes, p. 1-50. In E. Rosenberg (ed.), Myxobacteria, Springer-Verlag, New York, NY. Rodriguez, A. M., and A. M. Spormann. 1999. Genetic and molecular analysis of cglB, a gene essential for single-cell gliding in Myxococcus xanthus. J. Bacteriol. 181:438 14390. Sager, B., and D. Kaiser. 1993. Two cell-density domains within the Myxococcus xanthus fruiting body. Proc. Natl. Acad. Sci. USA 90:3690-3694. Sager, B., and D. Kaiser. 1994. Intercellular C-signaling and the traveling waves of Myxococcus. Genes Dev. 8:27932804. Shi, W., and D. R. Zusman. 1995. The frz signal transduction system controls multicellular behavior in Myxococcus xanthus, p. 419-430. In J. A. Hoch and T. J. Silhavy (ed.), Two-Component Signal Transduction. ASM Press, Washington, DC. Simon, M. A., D. L. Bowtell, G. S. Dodson, T. R. Laverty, and G. M. Rubin. 1991. Rasl and a putative guanine nucleotide exchange factor perform crucial steps in signaling by the sevenless protein tyrosine kinase. Cell 67:701-716. Simunovic, V., F. C. Gherardini, and L. J. Shimkets. 2003. Membrane localization of motility, signaling, and polyketide synthase proteins in Myxococcus xanthus. J. Bacteriol. 185:5066-5075. Skerker, J., and H. Berg. 2001. Direct observation of extension and retraction of type IV pili. Proc. Natl. Acad. Sci. USA 98:6901-6904. ~
Sliusarenko, O., J. Neu, D. Zusman, and G. Oster. 2006. Accordion waves in Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 103:1534-1539. Ssgaard-Andersen, L., and D. Kaiser. 1996. C-factor, a cellsurface-associated intercellular signaling protein, stimulates the cytoplasmic Frz signal transduction system in Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 93:2675-2679. Ssgaard-Andersen, L., F. Slack, H. Kimsey, and D. Kaiser. 1996. Intercellular C-signaling in Myxococcus xanthus involves a branched signal transduction pathway. Genes Dev. 10:740-754. Ssgaard-Andersen, L., M. Overgaard, S. Lobedanz, E. Ellehauge, L. Jelsbak, and A. A. Rasmussen. 2003. Coupling gene expression and multicellular morphogenesis during fruiting body formation in Myxococcus xanthus. Mol. Microbiol. 48:l-8. Sozinova, O., Y. Jang, D. Kaiser, and M. Alber. 2005. Threedimensional model of myxobacterial aggregation by contact-mediated interaction. Proc. Natl. Acad. Sci. USA 102:11308-1 1312. Sozinova, O., Y. Jang, D. Kaiser, and M. Alber. 2006. A three-dimensional model of myxobacterial fruiting body formation. Proc. Natl. Acad. Sci. USA 103:17255-17259. Spormann, A. M., and D. Kaiser. 1999. Gliding mutants of Myxococcus xanthus with high reversal frequencies and small displacements.J. Bacteriol. 181:2593-2601. Stephens, K., P. Hartzell, and D. Kaiser. 1989. Gliding motility in Myxococcus xanthus: the mgl locus, its RNA and predicted protein products. J. Bacteriol. 171:819-830. Stevens, A., and L. Ssgaard-Andersen. 2005. Making waves: pattern formation by a cell-surface-associatedsignal. Trends Microbiol. 13:249-252. Stock, J. B., M. G. Surette, M. Levit, and P. Park. 1995. Twocomponent signal transduction systems: structure-function relationships and mechanism of catalysis, p. 25-51. In J. A. Hoch and T. J. Silhavy (ed.), Two-Component Signal Transduction. ASM Press, Washington, DC. Sun, H., D. R. Zusman, and W. Shi. 2000. Type IV pilus of Myxococcus xanthus is a motility apparatus controlled by the frz chemosensory system. Curr. Biol. 10:1143-1146. Tieman, S., A. Koch, and D. White. 1996. Gliding motility in slide cultures of Myxococcus xanthus in stable and steep chemical gradients. J. Bacteriol. 178:3480-3485. Tzeng, L.-F., and M. Singer. 2005. DNA replication during sporulation in Myxococcus xanthus fruiting bodies. Proc. Natl. Acad. Sci. USA 102:14428-14433. Welch, R., and D. Kaiser. 2001. Cell behavior in traveling wave patterns of myxobacteria. Proc. Natl. Acad. Sci. U S A 98:14907-14912. Wolgemuth, C., E. Hoiczyk, D. Kaiser, and G. Oster. 2002. How myxobacteria glide. Curr. Biol. 12:369-377. Wu, S. S., and D. Kaiser. 1997. Regulation of expression of the pilA gene in Myxococcus xanthus. J. Bacteriol. 179:7748775 8. Wu, S. S., J. Wu, and D. Kaiser. 1997. The Myxococcus xanthus pilT locus is required for social gliding motility although pili are still produced. Mol. Microbiol. 23:109-121. Yu, R., and D. Kaiser. 2007. Gliding motility and polarized slime secretion. Mol. Microbiol. 63:454-467.
Myxobacteria: Multicellularity and Differentiation Edited by David E. Whitworth 0 2008 ASM Press, Washington, D.C.
Patricia Hartzell Wenyuan Shi Philip Youderian
Gliding Motility of Myxococcus xanthus
GLIDING MOTILITY DOES NOT DEPEND ON FLAGELLA Bacterial motility is an important physiological adaptation that facilitates the growth and survival of many different bacteria in their natural habitats and is a critical virulence determinant of bacterial pathogens. Many bacilli possess flagella that allow them to swim in aqueous environments. However, motility based on the rotation of flagellar filaments cannot function properly on semisolid or solid surfaces with limited water content. To move in such environments, some bacilli have developed mechanisms to amplify the number of flagella that can then be used to swarm over surfaces (for a review see Berg, 2005). Another group of bacteria, the spirochetes, have sheathed their flagella to permit corkscrew rotation along their long axes (Limberger, 2004). Other bacilli have developed very different motility mechanisms that function independently of flagella, also involving the movement of cells in the direction of their long axis on a surface, collectively termed “gliding motility.” Myxococcus xanthus is one of the diverse bacteria that display gliding motility, involving
6 two mechanisms, called adventurous (A) and social (S) gliding motility. The cells of the free-living soil bacillus M . xanthus are thin cylinders with rounded ends, about 1pm wide and 8 to 10 pm long, that can move across surfaces without the use of flagella (Burchard, 1981). Upon inoculation of a small sample of an M. xanthus culture into the center of a petri dish containing rich medium, M . xanthus glides outward from the point of inoculation and forms colonies with a starburst pattern, as shown in Fig. 1. Microscopic examination of the edge of a colony formed by wild-type cells on agar medium reveals both cells that venture away from the leading edge of the colony (“adventurous” cells) and cells that move as groups (“social” cells). When M. xanthus cells are starved for nutrients, tens of thousands of cells aggregate to form structures called fruiting bodies. This program of multicellular development is dependent on components involved in both systems of gliding motility. The genetic and phenotypic analysis of mutants defective in gliding motility has shown that gliding motility
Patricia Hartzell, Department of Microbiology, Molecular Biology and Biochemistry, University of Idaho, Moscow, ID 83844. Wenyuan Shi, Department of Oral Biology, School of Dentistry, Department of Microbiology, Immunology and Molecular Genetics, School of Medicine, University of California, Los Angeles, Los Angeles, CA 90095. Philip Youderian, Department of Biology, Texas A & M University, College Station, TX 83843.
103
DEVELOPMENT AND MOTILITY
104
Figure 1 Wild-type M . xanthus cells use A and S gliding simultaneously to move over surfaces. (A) After 3 days of growth on 0.3% agar supplemented with Casitone-Tris-potassium phosphate-magnesium (CTPM),colonies of the wild-type strain display a characteristic wavelike pattern on 0.3% agar surface. Cells of the wild-type strain DK1622 (1.5 X lo4cells) were spotted on rich medium and incubated at 32°C for 3 days. The photograph of a portion of the colony was taken using a Nikon SMZU microscope (P. Hartzell). (B) Three individual cells (I),groups of cells ( 2 ) ,and phase-bright slime trails (3) are visible at the edge of a colony growing on rich medium with 1.5% agar.
is more complex than flagellum-dependent motility. We are just beginning to understand how these molecular motors generate force and the detailed mechanisms of their action.
M. XANTHUS HAS TWO MECHANISMS OF GLIDING MOTILITY, A AND S The first molecular insights about the mechanisms of gliding motility were gleaned from the results of genetic studies performed by Hodgkin and Kaiser, who used chemical mutagens to generate mutations that affected gliding. Their studies show that mutations that impair the ability of M . xanthus to glide have one of three prototypical phenotypes (Hodgkin and Kaiser, 1979a, 197913). The wild-type strain, M . xanthus DK1622, produces colonies that spread over the agar surface. At the colony edge, groups of cells that are aligned close to one another as well as isolated cells are visible. In contrast, all three types of mutants form colonies that were smaller than those formed by the wild-type
strain on 1.5% agar plates with rich medium (Fig. 2). The vast majority of “point” mutants that affect gliding motility form colonies about one-half the size of a wild-type colony. Of this group, about one-third form small colonies with smooth edges. Compared with the wild type, these mutants are said to have mutations in A (adventurous)genes because they are defective in the ability of isolated cells to move away from a colony edge. “A” mutants are still able to glide using the alternative, S (social) motility system. About two-thirds of these point mutants form smaller colonies with flattened, dispersed edges. These mutants are said to have mutations in S genes because they are defective in the ability of groups of cells to move away from a colony edge. “S” mutants are still able to glide using the alternative, A motility system. A third class of mutations affects both motility systems simultaneously. Mutants with these mutations form colonies less than onequarter the size of a wild-type colony. Initially, these were found to have defects in a single gene named mglA, for mutual gliding.
6. GLIDINGMOTILITY OF M.XANTHUS
105
a inututiun ie iitgI.4 or nln24
A'S-
AS+
AS-, mglA, er nk24 Figure 2 Genetic studies identify different classes of gliding mutants. The wild-type strain uses A and S motility to produce large colonies (A+S+,top panel) that spread over the agar surface. A single mutation in an S gene, including pil, sgl, tgl, or eps, results in a colony (A'S-, middle panel, left) with reduced spreading compared with the wild-type colony, which typically has a glossy, mounded core. At higher magnification, isolated cells can be seen at the colony edge. A single mutation in an A gene, including agl, cgl, or agm, results in a colony (A-S+, middle panel, right) with reduced spreading compared with the wild-type colony. At higher magnification, no isolated cells can be seen at the colony edge. Introduction of a mutation in an A gliding gene into the S- mutant and vice versa yields a colony (A-S-, bottom panel) that is devoid of motility. Cells of the double mutants do not move when viewed by time-lapse videomicroscopy at 30-s intervals, and the colonies possess a smooth edge. Single mutations in mglA and nlu24 give a similar colony phenotype. Photographs were taken using a Nikon SMZU microscope (P. Hartzell).
One can deduce that this third class of mutations impairs both motility systems simultaneously, because when double mutants are constructed carrying any combination of a first mutation in an A gene paired with a second mutation in an S gene (A-S- mutants), they display the same small-colony phenotype as single mutants defective in mglA function (Hodgkin and Kaiser, 197913). The results of these genetic studies show that M. xanthus cells have two gliding systems, A-gliding, which controls
individual cell movement, and S-gliding, which controls the movement of groups of cells.
DEFECTS I N A O R s MOTILITY RESULT IN DIFFERENT ~vKJTANTPHENOTYPES The phenotypes of gliding mutants can be measured in terms of the differences in their velocities of gliding. Quantitative, high-resolution single-cell motility
106 assays have been used to calculate the average gliding velocities of wild-type and mutant M. xanthus cells. When viewed by videomicroscopy, individual wildtype cells glide with velocities that range between 1 and 20 pm/min (Spormann and Kaiser, 1995) with an average velocity of 4.4 pm/min (Spormann and Kaiser, 1999). The most common velocities range from 1.5 to 6 pm/min (Spormann and Kaiser, 1995). This observed variation in velocities is dependent on agar composition and whether a cell is separated or adjacent to another cell. Cells that are close to another cell, less than 0.5 pm, which is less than one cell diameter, move with an average velocity of 5.0 pm/min; often these cells were observed to glide transiently at velocities up to 20 p d m i n . In contrast, cells separated by more than one cell diameter move at about 3.8 pm/min. Wild-type M. xanthus cells exhibit several different behaviors during gliding. A cell may glide, stop for <9 s, and then reverse direction; glide, stop for >9 s, and then continue in the same direction; or glide, stop for >9 s, and then reverse direction (Spormann and Kaiser, 1999). Among wild-type cells, only 5% of the cells reverse direction after pausing for <9 s. These cells reverse the direction of movement at a frequency of once per 6 min. When viewed by time-lapse videomicroscopy at 30-s intervals, A-S- double-mutant cells are nonmotile (Spormann and Kaiser, 1999). Videomicroscopy reveals that mutant cells with defects in A-gliding genes (designated agl, agm, agi, and cgl) are unable to glide as isolated cells. Because these mutants cannot move as isolated cells, it is not possible to measure the average velocities of isolated cells. However, when in close proximity ( < 2 pm) to other cells, cglB mutants are motile, due to their S-motility, making it possible to measure their average velocities and frequency of cell reversals (Spormann and Kaiser, 1999). Individual cglB cells are able to move in both highreversal and low-reversal modes. Many cells of cglB mutants show an average number of reversals of 2.7 ( 2 1.4) per min. A subset of cells within the population can move for 150 s without reversing. The movement is punctuated by abrupt changes in velocities, as if the cells are stuck in forward glide, yet periodically attempt to brake. The average velocities of cglB cells in the highreversal mode is 2.5 (*1.7) pm per min, whereas that of cglB cells in the low-reversing mode is 4.7 (k3.0) pm per min (Spormann and Kaiser, 1999). The A and S mechanisms enable movement in different laboratory environments. Mutants with only Amotility (A+S-)glide like the wild-type strain over 1.5% agar surfaces but move more poorly than the wild type over 0.3 % agar. In general, the opposite is true for A-S+
DEVELOPMENT AND MOTILITY strains, which move more poorly over 1.5% agar but behave like the wild type on 0.3% agar (Shi and Zusman, 1993). These results, shown in Fig. 3, suggest that the A-motility genes encode products that enable cells to interact with a solid surface whereas the S-motility genes encode products that rely on cell-cell interactions to move over less viscous environments.
BOTH A AND S MOTILITY ARE DEPENDENT O N THE FUNCTIONS OF LARGE NUMBERS OF GENES DISPERSED O N THE M . XANTHUS GENOME The more detailed molecular genetic analysis of the two mechanisms of gliding motility in M. xanthus has been facilitated by the use of transposon mutagenesis. Screens were developed to identify insertions in genes for A and S motility. As described earlier, mutants with single mutations in A genes or single mutations in S genes form colonies about one-half the size of a wild-type colony whereas double mutants form small colonies. When an A or S mutant is mutagenized with a transposon, “nonmotile” (nonspreading) colonies are found that have transposon insertions in S or A genes, respectively. Because transposons are both genetic and physical markers, double mutants with a transposon insertion that inactivates an A or S gene can be used to identify the gene in which the second mutation, a transposon insertion, lies. The first mutants with transposon insertions in motility genes were made using the hybrid transposon, Tn5-lac (Kroos and Kaiser, 1984).This transposon carries an npt gene, which confers resistance to kanamycin (Kan) in both M. xanthus and Escherichia coli. The discovery that the bacteriophage P1 could be used to mediate the generalized transduction of genetic markers from E. coli into M . xanthus (Kaiser and Dworkin, 1975) enabled Kaiser and colleagues to use P1-mediated transduction of this transposon to generate mutants of 211. xanthus defective in multicellular development (Icross et al., 1986). A similar approach was used to make transposon insertions in genes required for gliding motility (MacNeil et al., 1994a). Insertions of Tn5-lac enabled the more rapid sequencing of genes involved in gliding motility, because the junctions between transposon insertions and their chromosomal targets could be recovered as inserts in plasmid subclones of mutant DNA that confer resistance to kanamycin (Kan‘). Determining the locations of motility genes on the physical map of the M. xanthus genome was made possible by probing blots from contour-clamped homogenous electric field (CHEF) gels of chromosomal DNA of transposon-carrying mutants
6.
GLIDINGMOTILITY OF M. XANTHUS
207 Spreading phenotypes
mglA A-S- (aglU, sgiK) al
n
r e
c,
A-S+(agmK)
l3
c
A-S+(aglU)
P
r A+S- (MxH1651) -.CI
s
c,
A+S- (SglK)
A+S+ (DK1622) =
A+S+ (DZ2)
0
100
200
300
1.9
400
Spreading area (mm2) after 5 days
Figure 3 The two gliding systems enable cells to adapt to different environments. The ability of cells to move over different agar surfaces supplemented with nutrients was quantified by measuring the spreading area (in square millimeters) after 5 days. Cells that lack A motility but retain S motility are able to spread on 0.3% agar, yet move poorly on 1.5% agar. The converse is true for cells that lack S motility, yet retain A motility. Black and gray bars indicate the colony surface area on 0.3 and 1.5% agar, respectively. The ratio of spreading on 0.3/1.5% agar surfaces for each strain is listed to the right of the bars. Data for DZ2 (on charcoal-yeast extract medium) are taken from Shi and Zusman (1993); all other values are from MacNeil et al., 1994b, and Youderian et al., 2003, for cells grown on CTPM medium.
that had been digested with AseI and SpeI with Tn5-lac (MacNeil et al., 1994a). About half of these insertions also place the expression of the lac2 reporter gene under the control of promoters for A and S gliding genes, which facilitates the analysis of the regulation of gliding motility during vegetative growth and development. Expression of a subset of gliding genes increases during starvation-induced development (MacNeil et al., 1994b).This result suggests that some gliding gene products play critical roles in fruiting body formation or sporulation. The analysis of pgalactosidase activity from Tn5-lac insertions reveals that transcription of motility genes does not require active gliding. P-Galactosidase activities were nearly identical from liquid-grown (where gliding is inoperative) and plate-grown cells (MacNeil et al., 1994a; Shi and Zusman, 1993). Expression of P-galactosidase from A and S promoters also was identical between transposons in the wild type (DK1622) background and the otherwise isogenic AmglBA background. This result shows that mglA does not play a role in global transcription of A and S genes. Analysis of Tn5-lac insertions in motility genes reveals that S-motility is dependent on type IV
pili (TFP) (Wu and Kaiser, 1995), 0-antigen biogenesis (Bowden and Kaplan, 1998),and a homolog of the chaperone, DnaK (Weimen et al., 1998). More recently, the transposon magellan-4, a derivative of the eukaryotic transposon Himar, which is active in M. xanthus, has been used to generate new mutants defective in A and S gliding (Youderian et al., 2003; Youderian and Hartzell, 2006). Nonmotile colonies obtained from magellan-4 transposon mutagenesis of a mutant with either an A or S motility defect have been found to carry transposon insertions in S and A genes, respectively, including all of the A and S genes identified previously by insertions made with Tn5-lac (Bowden and Kaplan, 1998; MacNeil et al., 1994a, 199413; Weimen et al., 1998; Wu and Kaiser, 1995). Transposon magellan4, like Tn5-lac, carries a selectable npt gene (Rubin et al., 1999). The use of magellan-4 offers three advantages over that of Tn5-lac. First, because it is much smaller than Tn5-lac, magellan-4 can be introduced into M . xanthus by electroporation to obtain a higher frequency of Kan' mutants. Second, magellan-4 has the plasmid R6Ky origin of replication, dependent on the activity of the T protein, facilitating a double selection for the recovery
108 of Kan‘ plasmid subclones with magellan-4 insertions and adjacent M . xanthus chromosomal DNA in E. coli. Third, and most important, the spectrum of insertions generated by magellan-4 in M . xanthus is broader than that of eubacterial transposons, including derivatives of Tn5. Mutagenesis with magellan-4 has made it possible to identify more than one-half of the genes known to be required for A and S gliding motility. The third genetic approach that has been used to identify 111. xanthus genes involved in motility, integrative disruption, is a technique borrowed from Saccharomyces cerevisiae. Most plasmids with origins of replication that function in E. coli cannot replicate autonomously in M . xanthus. If a fragment of DNA missing the 5’ and 3’ ends of a target gene is subcloned into a plasmid and the plasmid is introduced into M. xanthus, the integration of the plasmid following selection for an antibiotic resistance determinant on the plasmid results in a cointegrate in which plasmid vector DNA is flanked by two incomplete copies of the gene, one missing its 3’ end and one missing its 5’ end. Plasmids that disrupt a gene in this manner can excise from their chromosomal targets at a low frequency by homologous recombination. Thus, DNA extracted from M. xanthus mutants made in this manner can be used to electroporate E. coli to recover plasmids from libraries of subcloned M. xanthus DNA and thereby identify the target, disrupted genes rapidly by sequence analysis of their inserts and comparison of the insert sequences with the M. xanthus genome sequence. Using this approach, four groups have identified additional genes required for A and S motility. When PCR fragments resulting from the amplification of internal fragments of genes encoding 28 different response regulators were subcloned into a plasmid and integrated into the M . xanthus chromosome, 4 of the 28 recombinants resulting from integrative disruption were found to be defective in motility (Caberoy et al., 2003). A screen of Kan‘ mutants made by integrative disruption using plasmids with ca. 500-bp inserts of DNA were screened for a loss of the ability to bind calcofluor white, an indicator dye that binds exopolysaccharide (EPS), identified several genes required for the biogenesis of EPS, and demonstrated that the production of EPS is essential for S motility (Lu et al., 2005). In a more general screen for motility mutants, Lee et al. (2006) have confirmed independently that the aglT, agmK, agml’, agmU, and agmW genes are required for A-motility, and that the d i p , pilA, pilN, pilR, and tgl genes are required for Smotility. In addition, they identified a new gene required for A-motility, agiA (MXAN5319),which is predicted to encode a 787-amino-acid protein with multiple tetratricopeptide repeat (TPR) domains. Integrative disruption
DEVELOPMENT AND MOTILITY also has been used to demonstrate that the rasA (aka sgmO) gene, adjacent to frzS, is required for S-motility (Pham et al., 2005; Youderian and Hartzell, 2006). A comparison of the sets of mutants that have resulted from mutagenesis with transposon magellan-4 and by integrative disruption points out the fact that mutant hunts have yet to saturate the genes required for A and S motility-genes that have been identified by one method have yet to be identified by the other, and vice versa. Indeed, the spectra of magellan-4 insertions that result in a nonmotile phenotype starting with different A and S genetic backgrounds are different (P. L. Hartzell and P. Youderian, unpublished results), confirming that mutant hunts have yet to saturate the genes required for A and S motility. Both of these methods also have the limitation that genes disrupted by transposon or plasmid insertions must be nonessential for the growth of M . xanthus. As described below, an essential gene involved in motility has been identified by selecting for second-site suppressors of an mglA missense mutation (Thomasson et al., 2002), and a yeast two-hybrid screen has led to the identification of a gene required for A motility that has yet to be found using the methods of transposon mutagenesis and integrative disruption (Yang et al., 2004). Using these approaches, more than 30 different genes have been identified that are required for A motility, and more than different 80 genes have been identified that are required for S motility; summaries of these genes are shown in Tables 1 and 2. As is the case for most other gram-negative bacteria, many genes encoding products of related function are clustered as operons on the M. xanthus genome. In E. coli and Salmonella enterica serovar Typhimurium, the genes required for flagellar motility are clustered into several large cotranscription units that are regulated by a cascade of specific DNA-binding proteins. The organization of the A and S motility genes in M . xanthus differs from that of the flagellar genes in many bacteria, because they appear to be dispersed among many different operons on the M . xanthus genome.
THE MECHANISM OF ADVENTUROUS MOTILITY REMAINS A MYSTERY M . xanthus cells that have a functional A gliding system are able to move as isolated cells on a solid surface. M. xanthus cells leave “slime” trails behind when they glide, the exact composition of which is not yet known. The strong correlation between the presence of a slime trail and gliding has led to the speculation that the extrusion of slime might generate the force that propels cells using the A-gliding mechanism. Slime is thought
6 . GLIDING MOTILITY OF 211. XANTHUS
109
Table 1 Genes known to be required for A motility Assigned name
Gene
Coordinates
Related gene product or protein family
e value
aglR aglS aglV agl U agl W aglX agmR agmJ agmT agmK agm U
MXAN6862 MXAN6860 MXAN5754 MXAN6137 MXAN5756 MXAN5753 MXAN5818 MXAN62 59 MXAN6607 MXAN4863 MXAN48 70
8415360-8414623 8413986-8413402 7122477-7122938 3531405-3533108 7123727-7125022 7121748-7122458 7211395-7212510 7730848-7732758 8133818-8134837 6092480-6081012 6103539-6099883
le-10 0.60 4e-24 3e-4 le-52 8e-22 2e-55 6.2 le-25 2e-23 0.027
agiA agmA aglT
MXAN5319 MXAN3 886 MXAN4869
6623995-6621470 4672743-4670944 6099883-6098447
a g mH a g mI agmL; idh agm W agm 0
MXAN4638 MXAN3502 MXAN3537 MXAN5820 MXANS756 MXAN2538
5818631-5817792 4082356-4083270 4121250-4122548 7214074-7215156 7123727-7125022 2953965-2953411
agmF agmP agmQ agmS agmC
MXAN3352 MXAN2542 MXAN2923 MXAN6608 MXAN4798
3895929-3896969 2957914-2959197 3426566-3424470 8135638-8134847 6001965-5996770
agmX a g m D ; trpS agmE agmB; hrpB agmG aglZ agmN agm Z agm V cglB mglA mglB
MXAN4862 MXAN3842 MXANO635 MXAN3055 MXAN6519 MXAN353 6 MXANl673 MXAN2991 MXAN4866 MXAN3060 MXANl925 MXANl926
6080878-6078854 4621776-4620733 734257-73521 6 3585653-3583089 8039711-8041849 4120420-4121000 1979465-1977924 3508863-3504676 6095944-6095315 3589256-3590506 2258707-2258120 2259198-2258719
MotAITolQExbB proton channel ExbD/TolR ACC45288 ExbD/TolR AAM71873 p-Transducin-like protein HET-E2C AAL37299; TolB TolB AAM71875 TolQ biopolymer transport protein Putative ion transporting ATPase; CAB45559 ATP-binding cassette protein transporter BAB59028 Predicted periplasmic solute-binding protein AAM24479 TPR AAF31047 TPR-domain-containing protein AAM05026 and YbgF, CAC82711 787-amino-acid protein with TRP domains N-acetylmuramoyl-L-alanine amidase CAB73523 Predicted N-acetylglucosaminyltransferase (ARG99) AAH420 83 Lysophospholipase, ANN50803 Hypothetical protein ZP00079645 Isocitrate dehydrogenase BAB06878 Putative metalloprotease C6983 1 Carboxy-terminal protease BAC45699 Proto-oncogene tyrosine-protein kinase c-ABL CAA34438 Pseudouridylate synthase AAM24749 Multidrug resistance membrane protein Leucine aminopeptidase-related protein AAF96710 Enoyl-coenzyme A hydratase AEOll275 Putative hemagglutinin/hemolysin-related protein CAD18331 Putative DnaJ-domain-containing protein ACOl8929 tRNA synthetase CAB74224 ParA family; Soj/Par CAB 16134 ATP-dependent RNA helicase NP643917 Site-specificrecombinase, CAD14714 Myosin heavy chain Hypothetical protein BAB99656 No significant similarity No significant similarity No significant similarity GTPase Sarlp, CAA35978 RoadblocklLC7 domain
agmM
to play a role in elasticotaxis, which is the movement of cells oriented along stress lines in agar. Elasticotaxis is dependent on A-gliding and independent of S-gliding (Fontes and Kaiser, 1999). It has been proposed that the polyelectrolyte chains of slime that are extruded by Agliding cells allow them to align with the stressed agar. A-gliding motility may be the M. xanthus counterpart to secretion-mediated gliding in cyanobacteria such as Phormidium (Hoiczyk, 1998). The discovery of
6e-37 2e-15 3e-28 8e-5 e-157 2e-11 2e-81 9 3e-51 2e-33 5e-36 5e-33 2e-17 0.001 le-74 3e-33 e-105 2e-58 le-8
4e-4 1.5
nozzle-like structures in cyanobacteria that appear to release slime at the same rate at which cells were gliding led to the idea that slime secretion provided the propulsive force for gliding. Hoiczyk (2000) proposed that the steady secretion of mucilage through organelles called junctional pore complexes would be sufficient to generate thrust for locomotion in cyanobacteria. Rotation and forward motion would be directed by the flow of the mucilage through helical fibrils made of oscillin, a Ca2+
DEVELOPMENT AND MOTILITY
110 Table 2
Genes known to be required for S motility
Assigned name
Gene
Coordinates
Related gene product or protein family
e value
difA difB difC dip difE difG easA
MXAN6696 MXAN669.5 MXAN6694 MXAN6693 MXAN6692 MXAN6691 MXAN2293
8237133-8235892 8235895-8235158 8235114-8234692 8234621-8234253 8234199-8231626 8231629-8230799 2556513-2665704
1.00E-29 1.00E-05 6.00E-14 3.00E-19 3.00E-17 2.00E-17 1.00E-27
easB efP epsA
MXAN2294 MXA NS769 MXAN7451
2556760-2666518 7145107-7144529 9075393-9076208
epsB
MXAN7450
9073876-9075396
eps C
MXAN7449
9072562-9073098
epsD epsE epsF
MXAN7448 MXAN744.5 MXAN7444
9071518-9072531 9070740-9069502 9069342-9067687
epsG; mgtE epsH epsl; nla24 epsJ epsK epsL epsM epsN eps 0 epsP epsQ epsT eps U eps V
MXAN7443 MXAN7441 MXAN7440 MXAN7439 MXA N743 8 MXAN743 7 MXA N743 6 MXAN743S MXA N7433 MXAN7431 MXA N743 0 MXAN7426 MXAN7422 MXAN7421
9067641-9066332 9064818-9063364 9061999-9063354 9060497-9062002 9060367-9059264 9059242-9056069 9055621-9054281 9053040-9053837 9050769-9052871 9050527-9050108 9050081-9049782 9049152-9049568 9046164-9047408 9044670-9046148
eps W epsX epsY epsZ frzS gmd kdtA lpxK masK mglA mglB mmrA * nlal nlal9 nla23 pilA pilB pilC
MXAN7420 MXAN7418 MXAN7417 MXAN7415 MXAN4149 MXANS327 MXAN4714 MXAN4711 MXANl929 MXANl92S MXANl926 MXANS906 MXANS853 MXANlO78 MXANS 777 MXANS783 MXANS 788 MXANS 786
9044224-9044682 9042762-9044009 9042099-9042758 9038971-9040455 5096349-5098037 6633484-6634458 5908373-5907090 5905056-5903863 2263006-2264952 2258707-2258120 2259198-2258719 7326775-7325498 7262067-7263449 1256504-1257952 7153796-7152372 7159460-7158798 7166846-7165146 7163927-7162674
Methyl-accepting chemotaxis protein; Tar MmcQ Fibril biogenesis regulator Response regulator; A to C Histidine kinase; CheA Fibril biogenesis regulator; CheC Conserved hypothetical protein; DUFS74 protein family Hypothetical protein Translation elongation factor P; Efp UDP-N-acetylmannosamine transferase (EpsP); WecG protein family Putative glycosyl hydrolase; endo-l,4-P-glucanase precursor; AbfA protein family Transferase, hexapeptide repeat family; serine acetyltransferase; CysE Glycosyltransferase family 2 Glycosyltransferase family 1; RfaG PAS/PAC domain; response regulator/sensor histidine kinase Mg2+transporter; MgtE Glycosyltransferase family 1; RfaG Response regulator; AtoC Histidine kinase; BaeS Me2+resistance; AcrA Efflux CzcA family; AcrB Outer membrane efflux; TolC Hydrolase; MhpC vW factor type A; CnaB protein family Transposase Transposase Hypothetical protein Glycosyltransferase, group 2 Polysaccharide chain length determining protein; GumC Response regulator Hypothetical protein Polysaccharide biosynthesis/export; poly-export Sugar transferase Response regulator GDP-mannose-4,6-dehydratase KdtA; KDO addition Lipid A 4' kinase; LpxK Protein kinase Ras family monomeric GTPase RoadblockLC7 family Major facilitator family Response regulator; AtoC Response regulator; AtoC Response regulator; AtoC Pilin protein Pilus assembly ATPase PiIB; PulE Type 4 fimbrial assembly protein PilC; PulF
3.00E-62 5.OOE-55 9.00E-03 2.00E-26 2.00E-17 2.00E-28 1.00E-20 1.00E-67 4.00E-3 1 4.00E-115 2.00E-30 9.00E-09 1.00E-132 3.00E-14 9.00E-24 3.00E-09 5.00E-06 1.00E-23 2.00E-05 8.00E-22 2.00E-17 9.00E-03 2.00E-12 2.00E-140 7.OOE-82 3.00E-48 5.00E-27 2.00E-06 2.00E-11 7.00E-27 6.00E-134 3.00E-133 7.00E-126 8.00E-04 1.00E-157 5.OOE-101 (Continued)
6. GLIDINGMOTILITY OF M.XANTHUS Table 2
111
(Continued)
Assigned name
Gene
Coordinates
pilD; g s p 0 pilG
MXANS779 MXAN5782
7156015-7155008 7158778-7157783
pilH pill
MXANS781 MXANS780
7157786-7156806 7156806-7156039
pilM pilN pi10 pilP pi18 pilR pilS pilT purD purH
MXANS776 MXANS77S MXANS774 MXANS773 MXAN5772 MXANS784 MXANS785 MXANS787 MXAN2916 MXAN2914
7152261-7151074 7151046-7150369 7150354-7149737 7149732-7149130 7149072-7146367 7161083-7159647 7162657-7161080 7165103-7163985 3415613-3416878 3411223-3412767
rfaA rfa B rfaC rfbA rfbC rmd TPPA * scpA scpB selB sglK sgmA; ileS sgmB sgm C sgmD sgmE sgmF sgm G sgmH sgmI
MXAN4623 MXAN4622 MXAN462 1 MXAN4611 MXAN4610 MXAN5328 MXAN5907 MXAN3841 MXAN3840 MXANlO76 MXAN6671 MXAN03S8 MXAN0440 MXANllO6 MXANl641 MXANl795 MXAN2 128 MXAN22 03 MXAN2526 MXAN2S61 MXAN2921 MXAN2922 MXAN3506 MXAN3 759 MXAN3797 MXAN41 SO MXAN4613 MXAN4616 MXAN4639 MXAN4640 MXAN4707 MXANS333 MXANS592 MXANS766 MXANS770 MXANS831
5807239-5806457 5806448-5805135 5805093-5801266 5786671-5785784 5785787-5785236 6634459-6635385 7328957-7326786 4620569-4619700 4619713-4618724 1252102-1254018 8200793-8198970 420764-423664 502543-500069 1290271-1287593 1945852-1946811 2122220-2123269 2462214-2463125 2541175-2543460 2940096-2939569 2980677-2981636 3421643-3422788 3423007-3424401 4084582-4085541 4476390-4477931 4564786-4563644 5099222-5098089 5788612-5787587 5790997-5793249 5820518-5818644 5823072-5820562 5901165-5902136 6638572-6640812 6947304-6943193 7142238-7139056 7145604-7145242 7228546-7231050
SgmJ sgmK
sgmL sgmM; pccB2 sgmN s g m 0 ; rasA sgmP; rfbB SgmQ
sgmS sgmT sgm U sgm V sgm W sgmX sgm Y sgmZ; glgP
Related gene product or protein family Prepilin peptidase; PulO Putative efflux ATP-binding cassette transporter accessory factor PilG CcmA ATP-binding cassette Pultative efflux ATP-binding cassette protein, permease PiIM; TFP biogenesis protein PIIN; TFP biogenesis protein PilO; TFP pilus biogenesis protein PilP PilQ; TFP secretin Response regulator; AtoC Histidine kinase; NtrB PilT; twitching motility protein PurD; phosphoribosylamine glycine ligase Phosphoribosylaminoimidazolecarboxamide formyltransferase/IMP cyclohydrolase ATP-binding cassette protein ATP-binding cassette protein; TagH Glycosyltranferase family Glucose-1-phosphate thymidylyltransferase; RfbA dTDP-4-dehydrorhamnose 3,5-epimerase; RfbC GDP-4-dehydro-6-deoxy-o-mannose reductase Methyl-accepting chemotaxis protein; Tar ScpA, cell division ScpB, cell division; DUF387 Selenocysteine-specificelongation factor; SelB DnaK; heat shock protein 70 Ile-tRNA-synthetase Putative membrane protein DnaJ domaidTPR protein Hypothetical protein Putative lipoprotein Transcriptional activator; LysR family Hypothetical protein; hydrolase Hypothetical protein Fibronectin I11 Glucosyltransferase 1; RfaG NDP-sugar epimerase NAD-dep epimeraseldehydratase Acyl-CoA carboxyltransferase Acyl-CoA dehydrogenase; SCAD Hypothetical protein dTDP-glc 4,6-dehydratase; RfbB Glycosyltransferase;RfaG TPR repeat protein Histidine kinasehesponse regulator; KpdD Heptosyltransferase; RfaF Glycosyitransferase group; RfaG Response regulator; AtoC TPR repeat protein Conserved hypothetical protein Glycogen phosphorylase; GlgP . -
e value
9.00E-28
1.00E-63 4.00E-07 1.00E-53 1.00E-11 3.00E-20 2.00E-10 2.00E-59 9.00E-130 2.OOE-36 1.00E-130 2.00E-113 9.00E-77 9.00E-47 3.00E-82 6.00E-05 1.00E-116 3.00E-61 4.00E-41 5.00E-30 3.00E-36 2.00E-36 1.00E-48 0 8.00E-167 1.00E-31 4.00E-09
3.00E-26
1.00E-21 1.00E-21 5.OOE-33 6.OOE-154 5.00E-146 5.OOE-137 7.00E-20 2.00E-06 4.00E-28 2.00E-24 2.00E-18 1.00E-28 1.00E-07 5.OOE-05
(Continued)
DEVELOPMENT AND MOTILITY
112 Table 2
Genes known to be required for S motility (Continued)
Assigned name sgnA sgnB sgn C sgnD; p g i sgnE sgn G sgnH sgnl sigF
stk td wbgB wza
Gene
Coordinates
MXAN6225 MXAN6518 MXAN6627 MXAN6908 MXA N 7360 MXAN472 0 MXAN6679 MXAN72 03 MXAN0785 MXAN3474 MXAN3084 MXAN461 9 MXANl925
7572696-7571014 8039028-8039063 8155853-8154441 8460391-8461980 8982 802-8 9 83857 5903804-590281 8 8216055-8206615 8672540-8671500 894474-895295 4054907-4056529 3610985-3611746 5 799 15 3-5796 8 11 2245881-2245267
Related gene product or protein family Hypothetical protein Conserved hypothetical protein; Tol-Tol-Ttg2 Response regulator; A to C Glucose-6-phosphate isomerase Lipoprotein RfaE-like Hypothetical protein; RhsA CheR methyltransferase, SAM binding domain RNA polymerase sigma-32 factor family protein DnaK; heat shock protein 70 TPR repeat protein Glycosyltransferase; RfaG Polysaccharide export
binding protein (Hoiczyk and Baumeister, 1997),located on the cell surface. Support for this model comes from both the unicellular cyanobacterium Phormidium uncinatum and the multicellular cyanobacterium Anabaena. Wolgemuth et al. (2002) have proposed that nozzles located at the poles of the long cylindrical M. xanthus cell, similar to those described in Phormidium, are involved in secretion of polyelectrolyte.Using light and electron microscopy, the authors observed ribbons of mucilage (slime)being secreted from the ends of the cells. Up to 250 ring-shaped structures, or nozzles, shown in Fig. 4, may be observed at the poles of negatively stained isolated cell envelopes of wild-type M . xanthus (DK1622) by electron microscopy. Each cylindrically symmetric nozzle had an outer diameter of ca. 14 nm and a central opaque core of ca. 6.5 nm. These are consistent with the structures that were described in cyanobacteria. Active secretion of mucilage was studied by loading M. xanthus cells with the fluorescent dye acridine orange and then monitoring the release of acridine orangelabeled material from cells by fluorescence microscopy. The authors presented a model that propulsive force is generated by the hydration-driven swelling of the polyelectrolyte slime, similar in behavior to polyelectrolyte gel produced by snails, within the nozzle. The mucilage propulsion model is very attractive, and as described below, some of the A-gliding genes that have been characterized may contribute to the nozzle structure. However, it is possible that the release of slime is passive and occurs as a consequence of cells moving over a surface rather than the force generator. Full support for the model will depend on elucidation of the structure of the nozzle. While strong genetic and biochemical data that support the Wolgemuth model directly are lacking, some of the known A-gliding genes are postulated to form an inner-
e value
2.00E-09 1.00E-18 1.00E-83 3.00E-08 7.00E-51 1.00E-20 5.00E-35 3.00E-30 2.00E-120 2.00E-07 1.00E-07 2.OOE-25
outer membrane complex that may be related to the nozzles. At least 6 of the 35 genes (Table 1)that are specific to the A-gliding system encode proteins that are predicted to form an inner-outer membrane transport complex based on their similarity to the E. coli ToVTon proteins (White and Hartzell, 2002; Youderian et al., 2003). Moreover, at least four additional genes of the tol-ton family are near or within operons containing known A-gliding genes, suggesting that these genes also may be part of the A-gliding system. Two of the Tol-like proteins, AglU and AglW, are predicted to be outer-membrane lipoproteins that share identity with the enteric protein TolB. TolB has been shown to form a P-propeller structure, which is characteristic of proteins that interact with multiple partners (Bouveret et al., 1995; Ponting and Pallen, 1999). Both AglU and AglW contain multiple WD (Trp Asp) repeat motifs, which suggest that they also form P-propeller structures. Enteric TolB interacts with three other To1proteins, TolA, TolR, and TolQ, that reside in the inner membrane (Bouveret et al., 1999; Lazzaroni et al., 1999). The corresponding proteins in 111. xanthus are AglR, AglS, AglV, and AglX (Youderian et al., 2003). Together, these results suggest that M. xanthus requires one or two transport complexes for A-gliding. Other A-gliding genes, listed in Table 1, identified by transposon mutagenesis identify genes encoding proteins that might interact with Tol-like proteins to transport cargo. These include an ATPase, an ABC transporter and multiple TPRs. The TPR proteins are significant because there are numerous examples of interaction between (3-propeller and TPR proteins. AgmA, which is predicted to have acetylmuramoyl L-alanine amidase activity, may play a role in cell wall anchoring of an A-gliding motor protein (Youderian et al., 2003). Several proteases, encoded by agmM and agmW, are required
6. GLIDING MOTILITY OF M . XANTHUS
113
Figure 4 Slime secretion (through slime nozzles) may be important for A motility. (A) Electron micrograph of a negatively stained isolated cell envelope of M . xanthus DK1622, an A'S' strain, showing one of the cell poles. The nozzles are visible as ring-shaped structures, which are clustered at the poles (long arrow). Along the rest of the cell surface, the density of nozzles is much smaller (short arrows). The inset shows a higher magnification of the nozzle array in the region indicated by the long arrow. Scale bars are 0.2 pm and 50 nm (inset). (B) A gallery of electron micrographs of negatively stained isolated nozzles from M. xanthus DK1622. In these top views, each cylindrically symmetric nozzle has an outer diameter of 14 nm, with a central hole of 6 nm. The diameter is similar to those of corresponding structures found in cyanobacteria, suggesting that the remainder of the nozzle may be of similar size. (C) Schematic illustration of the arrangement and location of the different cellular structures involved in gliding motility in M. xanthus. Nozzles are clustered at the two cell poles and pili at one pole. S motility is generated by the pili, which extend, attach to nearby cells, and then retract, pulling the cells together. The authors propose that A motility is driven by the secretion of mucilage from the nozzles (indicated as small circles). As the mucilage adheres to the substrate, further secretion drives the cell in the opposite direction. The observed reversals of movement would be caused by alternation of the active polar nozzle cluster. (D) Cartoon illustrating the proposed layout of the nozzles in the polar region shown in panel A. The nozzle cross sections shown are drawn with the same geometry as those found in cyanobacteria. Figure reprinted from Current Biology with permission.
for A-gliding. They may play a role in processing other A-gliding proteins. It has been proposed that prior to cell reversal, proteolysis of motor protein complexes at one pole and reassembly of new complexes at the opposite pole occur. However, in light of the fact that cells pause for only about 10 s before reversing direction and
of the expense that this would incur, this proposal seems unlikely. The aglZ gene of M. xanthus was identified from a yeast two-hybrid assay in which MglA was used as bait (Yang et al., 2004). MglA is a 22-kDa cytoplasmic GTPase required for both A- and S-gliding motility
114 and sporulation (Hartzell, 1997; Hartzell and Kaiser, 1991a). Results suggest that MglA interacts with AglZ to regulate A motility because disruption or deletion of aglZ abolishes movement of isolated cells. The aglZ gene encodes a 153-kDa protein. The N terminus of AglZ shows similarity to the receiver domain of two-component response regulator proteins, while the C terminus contains heptad repeats (a b c d e f g), characteristic of coiled-coil proteins such as myosin. Consistent with this pattern, expression of AglZ in E. coli results in production of striated lattices in E. coli (Yang et al., 2004). As with myosin heavy chain, the purified C-terminal coiled-coil domain of AglZ forms filament structures in vitro.
THE MECHANISM OF S MOTILITY DEPENDS ON THE INTERACTION OF TFP WITH EPS, INVOLVED IN A CYCLE OF PILUS EXTENSION AND RETRACTION The majority of mutations that abolish S motility affect the production of TFP (Wu and Kaiser, 1995, 1996), the EPS component of fibrils (Lancer0 et al., 2004; Lu et al., 2005), or the lipopolysaccharide (LPS) moiety of 0-antigen (Bowden and Kaplan, 1998; Guo et al., 1996; Youderian and Hartzell, 2006) (Table 2). Polar TFP are absolutely required for S motility. TFP are polymers of the mature pilin protein, the product of the pilA gene (Wu and Kaiser, 1995), which is processed by a leader peptidase and secreted by a mechanism resembling that of the type I1 (general) secretory pathway (Peabody et al., 2003). The removal of pili either by genetic mutation (Wu and Kaiser, 1996)or by mechanical shearing in combination with blocking of protein biosynthesis (Rosenbluh and Eisenbach, 1992) confers a defect in S motility. The force-generating mechanism involved in S motility is likely the extension and retraction of pili (Nudleman and Kaiser, 2004), similar to the mechanism that drives twitching motility (for a review of twitching motility, see Burrows, 2005). When M. xanthus organisms are tethered perpendicularly to a glass or polystyrene surface in a TFP-dependent manner, cells display a jiggling motion during the retraction of a cell body to these surfaces, suggesting that social gliding motility involves a cycle of anterior pilus extrusion, adhesion of the pilus filament to a solid surface, and retraction (Sun et al., 2000). This cycle is dependent on the function of PilT, which is required for TFP retraction (Nudleman and Kaiser, 2004; Wu et al., 1997). A model showing a cycle of interaction between TFP and EPS that drives S motility is shown in Fig. 5. M. xanthus fibrils consist of extracellular matrix material
DEVELOPMENT AND MOTILITY composed of approximately equal parts carbohydrate and protein that surrounds the cells (Behmlander and Dworkin, 1991, 1994; Kearns et al., 2002; Kim et al., 1999; Li et al., 2003; Yang et al., 2000). Fibrils link neighboring cells to each other and to the substratum. Although fibrils contain both proteins and polysaccharides, it is the EPS component that is important for S motility (Li et al., 2005). The function of EPS in social motility has been elucidated by phenotypic analysis of mutants of M . xanthus lacking EPS, a secreted polymer comprised primarily of N-acetylglucosamine (GlcNAc) and glucosamine (GlcN) (Li et al., 2005; Lu et al., 2005). Mutants defective in EPS secretion are hyperpiliated, and the addition of protein-free fractions derived from matrix material stimulates pilus retraction and rescues this hyperpiliated phenotype. Interestingly, monomers of the amino sugar substituents of EPS including GlcN and GlcNAc have no effect on eps mutants but can cause hyperpiliation in wild-type cells. In contrast, chitin, a natural GlcNAc polymer, binds directly to TFP filaments and restores pilus retraction to hyperpiliated mutants. These data indicate that there is a close interaction between TFP and polysaccharides containing amino sugars, providing the first clue that EPS may provide attachment sites for the binding of TFP. The fact that the insoluble amino sugar polymer chitin triggers TFP retraction whereas soluble monomeric GlcNAc fails to do so leads to the hypothesis that TFP retraction may be a response to mechanical resistance provided by large amino sugar-containing polysaccharides, rather than to the binding of a simple ligand (Li et al., 2003). Because EPS is present both on cell surfaces and in slime trails, this interaction between TFP and amino-sugar-containing polysaccharides may be involved in slime-trailing behaviors (Fig. 5). These results may explain why certain motility mutants are able to move as isolated cells in solutions of 1% methylcellulose (Sun et al., 2000). Methylcellulose may bypass the need for EPS components needed for cell-cell contact during S motility. Many of the S motility genes listed in Table 2 are related to the synthesis, transport, or regulated expression of TFP and EPS. In addition, mutations that prevent the synthesis or assembly of the 0-antigen component of LPS also abolish S gliding (Bowden and Kaplan, 1998; Guo et al., 1996; Youderian and Hartzell, 2006). Why and how LPS is involved in the mechanism of S motility remain unanswered questions. Defects in 0-antigen production may result in pleiotropic defects in metabolism and/or cell wall structure, because 0antigen represents a significant fraction of the carbon present in gram-negative cells. Alternatively, 0-antigen
6 . GLIDINGMOTILITY OF 111. XANTHUS
115
A
Pili retract
4
B
\
Pili retract Cell body
Fibril material
- Pilus
Slime
Figure 5 The interaction between EPS and TFP stimulates retraction of pili. (A) The interaction between TFP and EPS in wild-type cells enables TFP retraction and S motility. (8)The absence of EPS in EPS mutants abolishes the EPS interaction with TFP, resulting in overpiliation and defects in S motility. (C) The interaction between TFP and EPS present in slime trails guides M. xanthus cells along these trails.
may participate directly in the process of pilus extension and retraction.
POLAR OSCILLATION OF TFP The frzS gene encodes a protein with an N-terminal response-regulator domain followed by a coiled-coil domain (Ward et al., 2000). Although frzS is linked genetically with the frz genes, which encode homologs of chemotaxis proteins, disruption of frzS blocks S motility. The phenotype of the frzS- mutant resembles that of a pilT mutant on agar (Ward et al., 2000), but in contrast with pilT mutants, the f ~ z Smutants can move as isolated cells in methylcellulose (Mignot et al., 2005). The frzS mutants are able to produce and retract pili, which shows that FrzS is not a part of the TFP motor (Mignot et al., 2005). To understand the role of FrzS in S motility, Mignot et al. (2005) constructed a FrzS-Gfp fusion and characterized the cellular location of the hybrid protein. FrzS-Gfp was found to accumulate preferentially at the leading end
of a gliding cell. The leading end of a gliding cell is the pole from which TFP are being extended and retracted to “pull” the cell forward. When movement of FrzS-Gfp was tracked in actively gliding cells, the position of the protein was observed to change upon cell reversal. In 86% of wild-type cells, FrzS was found to form a bright cluster at the leading pole of the cell. Only 2% of cells showed no symmetry for FrzS distribution. When an S gliding cell changes its direction by 180” (a cell reversal), the release of TFP switches to the opposite pole. What was the leading pole of the cell then becomes the lagging pole. Significantly, in 33 of 33 cells that were undergoing cell reversal, the localization of FrzS-Gfp was shown to switch to the opposite pole (Mignot et al., 2005). While proteolysis is known to govern polarity of proteins in other organisms, it does not appear to play a role in the FrzS polar exchange. During a reversal, FrzS-Gfp moves along what appears to be a spiral track in M. xanthus as shown in Color Plate 1. The precise nature of the track is yet to be determined, as is the signal that stimulates FrzS polar switching.
116
BOTH A A N D S MOTILITIES INVOLVE T H E EXCHANGE OF SURFACE LIPOPROTEINS BETWEEN CELLS The function of both motility systems depends on surface lipoproteins with the unusual property that they can be exchanged between cells in a process called extracellular complementation (Hodgkin and Kaiser, 1977; Nudleman et al., 2005). M. xanthus cells with a mutation in the cglB gene are defective in A motility when grown alone, but if cglB mutants are allowed to contact cells with a wild-type cglB gene, then the cglB mutants gain the ability to move as isolated cells transiently. The cglB gene encodes a 44-kDa protein with a typical aminoterminal lipoprotein signal sequence, consisting of two positively charged amino acids, followed by a stretch of 15 hydrophobic amino acids and then a cysteine to which lipid is attached (Rodriguez and Spormann, 1999).Antibodies against CglB have been used to confirm that the CglB lipoprotein is exposed on the surface of the cell. The “sharing” of CglB between cells is the mechanism that allows motility to be stimulated (Nudleman et al., 2005). During this unusual complementation in situ, no genetic material is exchanged and the CglB donor does not need to be motile for the transfer of CglB to occur. The CglB protein is unusually cysteine rich, which may be important for its function in A gliding. Similarly, cells with a mutation in a tgl gene lack S motility when grown alone, but tgl mutants gain the ability to move using S motility transiently in the presence of cells with a wild-type tgl gene (Nudleman et al., 2005). Like CglB, Tgl is a lipoprotein with TPR repeats that is transferred from a donor (tgl+)cell to a recipient (tgl-) cell when cells come into contact (Rodriguez-Soto and Kaiser, 1997). The exchange of Tgl protein and concomitant stimulation of S motility were shown by elegant mixing experiments shown in Fig. 6 . When tgl mutants expressing green fluorescent protein (GFP) were mixed with nonmotile donor tgl’ cells, the tgl mutant cells were shown to gain the ability to glide transiently. Tgl is needed to assemble PilQ, an ATPase required in turn for the activity of pili (Nudleman et al., 2006; Wall et al., 1999). At present, there are no known close homologs of the CglB and Tgl proteins predicted to be encoded by other microbial genomes.
THE MglA PROTEIN PLAYS A CENTRAL ROLE I N B O T H A A N D S MOTILITIES Early genetic studies showed that the mglA gene was uniquely involved in the function of both motility systems, because mutations in mglA block both A and S gliding simultaneously and the phenotype of mglA
DEVELOPMENT AND MOTILITY mutants is indistinguishable from that of A-S- double mutants (Hartzell and Kaiser, 1991a; Hodgkin and Kaiser, 1979b; Stephens et al., 1989; Stephens and Kaiser, 1987). This phenotype suggested that MglA might affect the transcription of gliding genes, be a common key component of the two different gliding motors, or play a role in coordinating the two motility systems. The mglA gene encodes a 22-kDa protein related to other members of the Ras superfamily of small, monomeric GTPases, and its function can be complemented, in part, by the expression of the small yeast SARlp GTPase, involved in vesicular transport (Hartzell, 1997). Although a subset of monomeric eukaryotic GTPases play roles in the regulation of gene expression, the function of MglA apparently does not play a role in regulation of the transcription of A and S genes. When the rates of expression of p-galactosidase from mutants with insertions of Tn.5-lac in A and S motility genes are compared in wildtype (mglA+)and mglA mutant backgrounds (MacNeil et al., 1994a), no significant differences are observed. Furthermore, the expression of many A and S genes also is unchanged in nonmotile A-S- cells. Hence, the regulation of transcription of the genes required for gliding motility in M. xanthus is remarkably different from the regulation of the majority of genes involved in flagellar biosynthesis, whose transcription depends on the state of flagellar assembly (for a review of flagellar regulation see Aldridge and Hughes, 2002). MglA may not be a common component of the two gliding motors. Although colonies formed by mglA mutants are indistinguishable from colonies formed by A-S- double mutants, time-lapse studies of individual cells show that mglA mutants reverse direction 17 times more often than cells of the wild-type strain (2.9/min for Amgl versus 0.17/min for DK1622) (Spormann and Kaiser, 1999). Hence, they are capable of gliding but are incapable of making net movement. However, this result is controversial, because some laboratories have been unable to detect movement of a AmglBA mutant even when viewed at 20-s intervals (Hartzell and Zusman labs, unpublished results). Mutants defective in mglA appear to produce the components, including pili, fibrils, and polyelectrolyte, that are required for A and S motility (Thomasson et al., 2002; R. Otto and P. Hartzell, unpublished data). Hence, the role of MglA may be to coordinate the two gliding motility systems or regulate the frequency with which cells reverse direction while gliding. This is consistent with the finding that MglA interacts with a component of each motility system (Fig. 7). MglA is a cytoplasmic protein that is predicted to interact with MglB, the product of the gene upstream of, and cotranscribed with, mglA (Hartzell and Kaiser,
6. GLIDING MOTILITY OF M. XANTHUS
117
R:
R:
Figure 6 Tgl stimulation. Stimulated recipient cells became motile and swarmed outward, while the donor strain remained nonmotile. The donor was found inside the original edge of the colony. A tgl mutant carrying an A- mutation (A-S- strain DK8602) was mixed with another nonmotile tgl' mutant strain (DK8601) and spotted on agar. (A) At 0 h, the mixed colony had a smooth edge (black arrowhead) because there were no motile cells (scale bar, 500 pm). (B) After several hours, the tgl' donor cells activated S motility in the tgl mutant recipient cells by stimulation. The outward swarming of the stimulated recipients after 4 days is indicated by the arrowhead. This motility was transient; it lasted only 1week. (C) Phase-contrast image of DK8601 (GFP- donor cells) mixed 1:l with a mixture of DK8607 (GFP+recipient cells) and DK8602 (GFP- recipient cells) at a 1 5 0 ratio. The original colony edge is indicated by the arrowhead. (D) Epifluorescent image of the field shown in panel C. The original colony edge is indicated by the dashed line. (E) Phase-contrast image of DK8602 (GFP- recipient cells) mixed 1:l with a mixture of DK8606 (GFP+donor cells) and DK8601 (GFP- donor cells) at a 1 3 0 ratio. (F) Epifluorescent image of panel E. Reprinted with permission from Science (Nudleman et al., 2005).
DEVELOPMENT AND MOTILITY
118
M a K , a prokin kinase required for S-motility
factor for MglA
Figure 7 The MglA protein interacts with multiple protein partners to regulate gliding. MglA is related to monomeric GTPases, which cycle between active, GTP-bound, and inactive, GDP-bound states. MglB (triangle)is predicted to regulate the activity of MglA, perhaps by acting as an exchange factor. MglA has been shown to interact with MasK and AglZ. A mutation in masK suppresses the nonmotile phenotype of the mglA8 mutation and restores S motility (Thomasson et al., 2002). AglZ, a coiled-coil protein required for A motility, was recovered from a yeast two-hybrid library probed with MglA as bait (Yang et al., 2004).
1991b; Stephens et al., 1989).Mutants defective in mglB have a reduced spreading rate and smaller colony size, most likely because they produce a reduced amount of MglA protein. This result suggests that MglB may stabilize or regulate the function of MglA. The growth rate of the mglB mutants is similar to that of the wild type, and the reduced colony size and gliding rate are thought to be due to the reduced activity of MglA resulting from loss of MglB. The MglB protein has an LC7hoadblock motif, conserved among a group of eukaryotic proteins that interact with ATPases and GTPases to regulate their activity (P. Hartzell, unpublished data). The identification of effectors of MglA has proven key to understanding the role of MglA in gliding motility. Genes encoding proteins that interact with MglA have been identified in each of the A and S motility systems that had not been identified previously, AglZ (Yang et al., 2004) and MasK (Thomasson et al., 2002), respectively. The masK gene was identified because it was found to carry a second-site mutation that suppresses a missense mutation in mglA, mglA8. Whereas the mglA8 mutant lacks both A and S motility, the masK mutation restores partial S-gliding, but not A-gliding, to the mglA8 mutant. Sequence analysis shows that MasK is a protein kinase in the family of serine-threoninetyrosine kinases. Antiphosphotyrosine antibody reacts with MasK produced in E. coli and with a protein of the size predicted for MasK in M. xanthus (Thomasson et al., 2002). When MasK is used as bait in a yeast twohybrid screen against a library of random M. xanthus inserts, independent clones containing the mglA gene
are recovered from yeast organisms that test positive for interaction. The partial S gliding restored by the masK mutation is associated with a hard, crunchy colony phenotype and the increased production of a protein that is associated with extracellular fibril material. When MglA was used as bait in a yeast two-hybrid screen, a fragment of the aglZ gene was recovered from a yeast clone that displayed a positive interaction (Yang et al., 2004). The aglZ gene encodes a protein of 150,000 Da that has an N-terminal receiver domain characteristic of response regulators, and a large C-terminal coiled-coil domain resembling those of myosins and related eukaryotic coiled-coil proteins (Yang et al., 2004). The yeast two-hybrid result provided initial evidence that MglA interacts with AglZ. Subsequently, purified AglZ was shown to bind specifically to purified MglA in vitro. Disruption or deletion of aglZ abolishes A-gliding, suggesting that MglA interacts with AglZ to control A-gliding. Purified AglZ forms filament-like structures in vitro in the absence of an exogenous energy source. This property of AglZ is similar to that of proteins that form intermediate filaments in eukaryotes.
THE TRANSCRIPTION OF A AND S MOTILITY GENES INVOLVES MULTIPLE REGULATORS Although the analysis of P-galactosidase activities produced from transcriptional fusions of known A and S genes with the lacZ reporter gene suggests that the levels of expression of many A and S genes do not change when
6. GLIDING MOTILITY OF 211. XANTHUS cells are grown on different substrates or when cells lack motility (MacNeil et al., 1994a; Shi and Zusman, 1993), several regulatory components for the S-gliding system have been identified recently. The sigF gene encodes a sigma factor that is required for S-gliding (Ueki and Inouye, 1998). SigF does not appear to affect expression of some of the known S-gliding genes, including pilA, rpoEl ,abcA, frgA, difA, or tgl, which has led to speculation that SigF regulates a subset of S-gliding genes. At least one two-component regulatory system regulates the transcription of the pilA gene, which encodes the prepilin protein. pilA is regulated positively by the PilR response regulator and negatively by PilS, the putative cognate kinase (Wu and Kaiser, 1997). PilR shares similarity with members of the NtrC family of response regulators, which are known to function with aS4. Consistent with this, analysis of the regulatory region of pilA identifies a promoter sequence related to that recognized by aS4in E. coli. In this regard, the regulation of the gene encoding the major pilin subunit in M. xanthus resembles that found in Pseudomonas aeruginosa. Until recently, few genes encoding proteins required for the expression of motility genes had been identified. However, the integrative disruption of 28 different NtrC-like activator (nla)genes has enabled the identification of four new response regulators required for gliding motility (Caberoy et al., 2003). Disruption of three of these genes, nlal, nla19, and nla23, causes defects in S-gliding,whereas disruption of the fourth, nla24, results in a defect in both A- and S-gliding. The NtrC-like activators are required for expression of genes that have aS4 promoters, by playing a critical role in the formation of an open complex that precedes the initiation of transcription of these genes. The activity of some NtrC-like proteins is modulated by phosphorylation. The nla24 mutant does not spread on 0.4 or 1.5% agar surfaces, nor does it show any movement when viewed by timelapse videomicroscopy. The mutant produces both LPS O-antigen and functional TFP, but is defective in the production of EPS (Lancer0 et al., 2004). The same gene was identified in a separate integrative disruption screen for genes involved in EPS biogenesis and was named epsI (Lu et al., 2005). Nla24 (EpsI) is a protein 437 amino acids long with a CheY-like response regulator domain, an AAA-type ATPase domain, and a DNA-binding domain. Expression of at least two essential A-gliding genes, aglU and cglB, is reduced threefold in an nla24 mutant, which might account for its effect on A-gliding. The proper function of components of the S-gliding motor also appears to depend on at least two of the five homologues of the chaperone DnaK that are predicted to be encoded by the M. xanthus genome. Mutations in
119 the sglK gene encoding one DnaK homologue abolish S motility (Weimer et al., 1998; Yang et al., 1998), whereas mutations in a second, stk, suppress defects in EPS biosynthesis (Kim et al., 1999). The different roles that these chaperones play in S-motility have yet to be determined.
A SUBSET OF GENES REQUIRED FOR GLIDING MOTILITY IS CRITICAL FOR MULTICELLULAR DEVELOPMENT M. xanthus has a complex life cycle. In the presence of adequate nutrients, the cells undergo vegetative growth and divide, but when the cells are starved of nutrients, they aggregate and form fruiting bodies containing myxospores. When nutrients become available, myxospores can germinate into vegetative cells. Gliding motility plays a critical role in this multicellular development of M. xanthus. When starved of nutrients, > l o s M. xanthus cells cooperate to form a three-dimensional fruiting body with an inner core that supports spore differentiation (Shimkets, 1999). During the first 10 h of development, cells exchange extracellular signals, including amino and fatty acids, which help coordinate cell movement and aggregation to produce a mound-shaped structure. Within this time period, large numbers of M. xanthus cells aggregate to form multicellular fruiting bodies, within which a subset of cells differentiate into refractile, heat-resistant spores. Motility is critical for fruiting body formation and sporulation. Nonmotile mutants, whether the result of a double mutation (one mutation in an A gene and one mutation in an S gene) or a single mutation in the mglA gene, fail to produce fruiting bodies and a full complement of mature spores and show reduced expression of developmentally regulated genes (Kroos et al., 1988). The production of heat-resistant spores is reduced about 105-foldin these mutants (Kroos et al., 1988). The role of motility in development is less clear when one examines the phenotypes of strains with single mutations in A or S genes. Mutations in most A genes, including aglZ and cglB, do not appear to affect fruiting body formation or sporulation (Yang et al., 2004). In contrast, mutations in the A-gliding genes that comprise the Tollike genes do not affect the timing or the formation of fruiting bodies but do affect the ability of M . xanthus to form heat-resistant spores (MacNeil et al., 1994b; White and Hartzell, 2000; Youderian et al., 2003). This subset of A genes encodes proteins that are predicted to form multisubunit transport complexes. Some of these proteins, including AglU and AglW, are predicted to be lipoproteins anchored to the outer membrane, whereas
DEVELOPMENT AND MOTILITY
120 others, including AglX, AglS, and AglR, are predicted to reside in the inner membrane. The sporulation phenotype of one mutant defective in a gene belonging to this group, aglU, has been studied in detail. The expression of agluincreases significantlyupon starvation-induced development (MacNeil et al., 1994b; Srinivasan et al., 2005; White and Hartzell, 2000). Vegetative rod-shaped cells of the aglU mutant differentiate into ovoid cells within the same time frame as the wildtype strain but fail to mature into refractile spheres, even with prolonged incubation, and yield only 0.01 % of the number of heat-resistant spores made by the wild-type strain. Microscopic examination of thin sections reveals that the spores formed within wild-type fruiting bodies have a thick, dense layer immediately surrounding the cytoplasmic core and an outermost layer composed of a loose, fibrous material. Although the outer, fibrous material is present in the spores formed by the aglU mutant, they have a less dense layer around the cytoplasmic core, which appears less organized than in wild-type spores (White and Harzell, 2000). This layer may contain the major spore coat proteins, proteins S and C, which are produced by the aglU mutant, but may not be localized properly. A subset of the genes required for S-motility are also required for fruiting body formation. Mutants listed in Table 2, particularly the ones defective in EPS production, are defective in fruiting body formation and sporulation, underscoring the importance of S motility for development. The role of TFP in the formation of fruiting bodies is complicated. A pilA mutant is still able to form fruiting bodies, but pilH mutants are defective in fruiting body formation (Bonner et al., 2006). Recently Lee et al. (2006)engineered a ApilA agiA double mutant, which is A-S- genetically, yet it retains social motility. Under certain conditions, M. xanthus, like P. aeruginosa (Durand et al., ZOOS), may form “pseudopili,” or alternative pili, that polymerize from a prepilin paralogue different from PilA. Like the P. aeruginosa genome, which contains multiple (four),potential prepilin genes, the M. xanthus genome has six, one of which may substitute for PilA during development.
THERE IS MUCH TO BE LEARNED ABOUT THE GENETIC BASIS AND MECHANISMS OF GLIDING Extensive genetic and genomic studies allow the identification and characterization of a large number of A and S motility genes. Significant advances in the past several years have shown that S gliding is powered by the extension and retraction of TFP. Retraction is likely
stimulated by components present in the extracellular matrix. The requirement of the extracellular matrix in retraction may explain why S-gliding is restricted to cells that are within one cell length of one another. The mechanism that allows movement of isolated (A) cells is still unclear. One model proposes that polar nozzles are used to generate thrust by secreting a polyelectrolyte from the rear pole of the cell. The wild-type cell appears to use both S and A gliding systems simultaneously, and MglA, a protein required for the function of both gliding systems, may help to coordinate the two motility systems so that the systems generate thrust in the same direction. We are just beginning to delve into the complexity of gliding motility in M. xanthus. The sequence of the M. xanthus genome reveals that many of the individual genes known to be required for gliding motility, such as pilA, have multiple paralogs. From a genetic point of view, only a few genes in the families of response regulators (28 of at least 148), TPR repeat proteins, and Tol/Ton exporters, to name a few of the gene families with known paralogs involved in gliding and very large numbers of paralogs predicted to be encoded by M. xanthus, have been analyzed. Indeed, 211. xanthus provides a unique model system in which we can dissect the roles of what appear to be partially redundant gene functions in the complex processes of motility. From a physiological point of view, M. xanthus is certain to yield novel insights into the mechanisms of cellular motility and the complex interplay between these mechanisms during the execution of its elegant program of multicellular development. We were supported by grants from the National Institutes of Health (GM.54666 to T S . and GM07.5242 to P.L.H.) and the National Science Foundation (MCB0242191 to P.L.H.).
References Aldridge, P., and K. T. Hughes. 2002. Regulation of flagellar assembly. Curr. Opin. Microbiol. 5:160-165. Behmlander, R. M., and M. Dworkin. 1991. Extracellular fibrils and contact-mediated cell interactions in Myxococcus xanthus. 1.Bacteriol. 173:7810-7821. Behmlander, R. M., and M. Dworkin. 1994. Integral proteins of the extracellular matrix fibrils of Myxococcus xanthus. J . Bacteriol. 176:6304-6311. Berg, H. C. 2005. Swarming motility: it better be wet. Curr. Biol. 15 :R599-R600. Bonner, P. J., W. P. Black, Z. Yang, and L. J. Shimkets. 2006. FibA and PilA act cooperatively during fruiting body formation of Myxococcus xanthus. Mol. Microbiol. 61: 1283-1 293. Bouveret, E., H. Benedetti, A. Rigal, E. Loret, and C. Lazdunski. 1999. In vitro characterization of peptidoglycanassociated lipoprotein (PAL)-peptidoglycan and PAL-TolB interactions. 1.Bacteriol. 15:6306-6311.
MOTILITY OF M. XANTHUS 6. GLIDING Bouveret, E., R. Derouiche, A. Rigal, R. Lloubes, C. Lazdunski, and H. Benedetti. 1995. Peptidoglycan-associated lipoprotein-TolB interaction. A possible key to explaining the formation of contact sites between the inner and outer membranes of Escherichia coli. J. Biol. Chem. 270: 11071-1 1077. Bowden, M. G., and H. B. Kaplan. 1998. The Myxococcus xanthus lipopolysaccharide 0-antigen is required for social motility and multicellular development. Mol. Microbiol. 30~275-284. Burchard, R. P. 1981. Gliding motility of prokaryotes: ultrastructure, physiology, and genetics. Annu. Rev. Microbiol. 35:497-529. Burrows, L. L. 2005. Weapons of mass retraction. Mol. Microbiol. 57: 8 78-8 8 8. Caberoy, N. B., R. D. Welch, J. S. Jakobsen, S. C. Slater, and A. G. Garza. 2003. Global mutational analysis of NtrC-like activators in Myxococcus xanthus: identifying activator mutants defective for motility and fruiting body development. J. Bacteriol. 185:6083-6094. Durand, E., G. Michel, R. Voulhoux, J. Kurner, A. Bernadac, and A. Filloux. 2005. XcpX controls biogenesis of the Pseudomonas aeruginosa XcpT-containing pseudopilus. J. Biol. Chem. 280:3 1378-3 1 389. Fontes, M., and D. Kaiser. 1999. Myxococcus cells respond to elastic forces in their substrate. Proc. Natl. Acad. Sci. USA 96~8052-8057. Guo, D., M. G. Bowden, R. Pershad, and H. B. Kaplan. 1996. The Myxococcus xanthus rfbABC operon encodes an ATPbinding cassette transporter homolog required for O-antigen biosynthesis and multicellular development. J. Bacteriol. 178~1631-1639. Hartzell, P., and D. Kaiser. 1991a. Function of MglA, a 22kilodalton protein essential for gliding in Myxococcus xanthus. J. Bacteriol. 173:7615-7624. Hartzell, P. L. 1997. Complementation of Myxococcus xanthus sporulation and motility defects by a eukaryotic RAS homolog. Proc. Natl. Acad. Sci. USA 979881-9886. Hartzell, l?L., and D. Kaiser. 1991b. Upstream gene of the mgl operon controls the level of MglA protein in M. xanthus. J. Bacteriol. 172:7625-7635. Hodgkin, J., and D. Kaiser. 1977. Cell-to-cell stimulation of movements in non-motile mutants of Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 74:2938-2942. Hodgkin, J., and D. Kaiser. 1979a. Genetics of gliding motility in Myxococcus xanthus (Myxobacterales): genes controlling movement of single cells. Mol. Gen. Genet. 171:167-171. Hodgkin, J., and D. Kaiser. 1979b. Genetics of gliding motility in Myxococcus xanthus (Myxobactererales):two gene systems control movement. Mol. Gen. Genet. 171:177-191. Hoiczyk, E. 2000. Gliding motility in cyanobacteria: observations and possible explanations. Arch. Microbiol. 174: 11-17. Hoiczyk, E. 1998. Structural and biochemical analysis of the sheath of Phormidium uncinatum. J. Bacteriol. 180:39233932. Hoiczyk, E., and W. Baumeister. 1997. Oscillin, an extracellular, Ca2+-bindingglycoprotein essential for the gliding motility of cyanobacteria. Mol. Microbiol. 26:699-708.
121 Kaiser, D., and M. Dworkin. 1975. Gene transfer to myxobacterium by Escherichia coli phage P1. Science 182653-654. Kearns, D. B., P. J. Bonner, D. R. Smith, and L. J. Shimkets. 2002. An extracellular matrix-associated zinc metalloprotease is required for dilauroyl phosphatidylethanolamine chemotactic excitation in Myxococcus xanthus. J. Bacteriol. 184:1678-1684. Kim, S. H., S. Ramaswamy, and J. Downard. 1999. Regulated exopolysaccharide production in Myxococcus xanthus. J . Bacteriol. 181:1496-1 507. Kroos, L., P. Hartzell, K. Stephens, and D. Kaiser. 1988. A link between cell movement and gene expression argues that motility is required for cell-cell signaling during fruiting body development. Genes Dev. 2:1677-1685. Kroos, L., and D. Kaiser. 1984. Construction of Tn5-lac, a transposon that fuses lacZ expression to exogenous promoters, and its introduction into Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 815816-5820. Kroos, L., A. Kuspa, and D. Kaiser. 1986. A global analysis of developmentally regulated genes in Myxococcus xanthus. Dev. Biol. 117252-266. Lancero, H., N. B. Caberoy, S. Castaneda, Y. Li, A. Lu, D. Dutton, X. Y. Duan, H. B. Kaplan, W. Shi, and A. G. Garza. 2004. Characterization of a Myxococcus xanthus mutant that is defective for adventurous motility and social motility. Microbiology 150:4085-4093. Lazzaroni, J. C., P. Germon, M. C. Ray, and A. Vianney. 1999. The To1 proteins of Escherichia coli and their involvement in the uptake of biomolecules and outer membrane stability. FEMS Microbiol. Lett. 15:191-197. Lee, C., J. Chung, J. Kim, and K. Cho. 2006. Identification of a gene required for gliding motility in Myxococcus xanthus. J. Microbiol. Biotechnol. 16:771-777. Li, Y., R. Lux, A. E. Pelling, J. K. Gimzewski, and W. Shi. 2005. Analysis of type IV pilus and its associated motility in Myxococcus xanthus using an antibody reactive with native pilin and pili. Microbiology 151:353-360. Li, Y., H. Sun, X. Ma, A. Lu, R. Lux, D. Zusman, and W. Shi. 2003. Extracellular polysaccharides mediate pilus retraction during social motility of Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 100:5443-5448. Limberger, R. J. 2004. The periplasmic flagellum of spirochetes. J. Mol. Microbiol. Biotechnol. E30-40. Lu, A., K. Cho, W. P. Black, X. Y. Duan, R. Lux, Z. Yang, H. B. Kaplan, D. R. Zusman, and W. Shi. 2005. Exopolysaccharide biosynthesis genes required for social motility in Myxococcus xanthus. Mol. Microbiol. 55:206-220. MacNeil, S. D., F. Calara, and P. L. Hartzell. 1994a. New clusters of genes required for gliding motility in Myxococcus xanthus. Mol. Microbiol. 14:61-71. MacNeil, S. D., A. Mouzeyan, and l? L. Hartzell. 1994b. Genes required for both gliding motility and development in Myxococcus xanthus. Mol. Microbiol. 14:785-795. Mignot, T., J. P. Merlie, Jr., and D. R. Zusman. 2005. Regulated pole-to-pole oscillations of a bacterial gliding motility protein. Science 310:855-85 7. Nudleman, E., and D. Kaiser. 2004. Pulling together with type IV pili. J. Mol. Microbiol. Biotechnol. 752-62.
122 Nudleman, E., D. Wall, and D. Kaiser. 2005. Cell-to-cell transfer of bacterial outer membrane lipoproteins. Science 309:125-127. Nudleman, E., D. Wall, and D. Kaiser. 2006. Polar assembly of the type IV pilus secretin in Myxococcus xanthus. Mol. Microbiol. 60:16-29. Peabody, C. R., Y. J. Chung, M. R. Yen, D. Vidal-Ingigliardi, A. P. Pugsley, and M. H. Saier, Jr. 2003. Type I1 protein secretion and its relationship to bacterial type IV pili and archaeal flagella. Microbiology 149:3051-3072. Pham, V. D., C. W. Shebelut, B. Mukherjee, and M. Singer. 2005. RasA is required for Myxococcus xanthus development and social motility. J. Bacteriol. 187:6845-6848. Ponting, C. P., and M. J. Pallen. 1999. A p-propeller domain within TolB. Mol. Microbiol. 31:739-740. Rodriguez, A. M., and A. M. Spormann. 1999. Genetic and molecular analysis of cglB, a gene essential for single-cellgliding in Myxococcus xanthus. J. Bacteriol. 181:4381-4390. Rodriguez-Soto, J. P., and D. Kaiser. 1997. The tgl gene: social motility and stimulation in Myxococcus xanthus. J. Bacteriol. 179:4361-4371. Rosenbluh, A., and M. Eisenbach. 1992. Effect of mechanical removal of pili on gliding motility of Myxococcus xanthus. J. Bacteriol. 1745406-54 13. Rubin, E. J., B. J. Akerley, V. N. No&, D. J. Lampe, R. N. Husson, and J. J. Mekalanos. 1999. In vivo transposition of marinerbased elements in enteric bacteria and mycobacteria. Proc. Natl. Acad. Sci. USA 96:1645-1650. Shi, W., and D. R. Zusman. 1993. The two motility systems of Myxococcus xanthus show different selective advantages on various surfaces. Proc. Natl. Acad. Sci. USA 90:33783382. Shimkets, L. J. 1999. Intercellular signaling during fruitingbody development of Myxococcus xanthus. Annu. Rev. Microbiol. 53:525-549. Spormann, A. M., and A. D. Kaiser. 1995. Gliding movements in Myxococcus xanthus. J. Bacteriol. 1775846-5852. Spormann, A. M., and D. Kaiser. 1999. Gliding mutants of Myxococcus xanthus with high reversal frequencies and small displacements. J. Bacteriol. 181:2593-2601. Srinivasan, B. S., N. B. Caberoy, G. Suen, R. G. Taylor, R. Shah, F. Tengra, B. S. Goldman, A. G. Garza, and R. D. Welch. 2005. Functional genome annotation through phylogenomic mapping. Nat. Biotechnol. 23:691-698. Stephens, K., P. L. Hartzell, and D. Kaiser. 1989. Gliding motility in Myxococcus xanthus: mgl locus, RNA, and predicted protein products. J. Bacteriol. 171:819-830. Stephens, K., and D. Kaiser. 1987. Genetics of gliding in Myxococcus xanthus: molecular cloning of the mgl locus. Mol. Gen. Genet. 207:256-266. Sun, H., D. R. Zusman, and W. Shi. 2000. Type IV pilus of Myxococcus xanthus is a motility apparatus controlled by the frz chemosensory system. Curr. Biol. 10:1143-1146. Thomasson, B., J. Link, A. G. Stassinopoulos, N. Burke, L. Plamann, and P. L. Hartzell. 2002. The GTPase, MglA, interacts with a tyrosine kinase to control type-IV
DEVELOPMENT AND MOTILITY pili-mediated motility of Myxococcus xanthus. Mol. Microbiol. 46: 1399-14 13. Ueki, T., and S. Inouye. 1998. A new sigma factor, SigD, essential for stationary phase is also required for multicellular differentiation in Myxococcus xanthus. Genes Cells 3:371-3 85. Wall, D., P. E. Kolenbrander, and D. Kaiser. 1999. The Myxococcus xanthus pilQ (sglA) gene encodes a secretin homolog required for type IV pilus biogenesis, social motility and development. J. Bacteriol. 181:24-33. Ward, M. J., H. Lew, and D. R. Zusman. 2000. Social motility in Myxococcus xanthus requires FrzS, a protein with an extensive coiled-coil domain. Mol. Microbiol. 37:13571371. Weimer, R. M., C. Creighton, A. Stassinopoulos, P. Youderian, and P. L. Hartzell. 1998. A chaperone in the HSP70 family controls production of extracellular fibrils in Myxococcus xanthus. J. Bacteriol. 1805357-5368. White, D. J., and P. L. Hartzell. 2000. AglU, a protein required for gliding motility and spore maturation of Myxococcus xanthus, is related to WD-repeat proteins. Mol. Microbiol. 36:662-678. Wolgemuth, C., E. Hoiczyk, D. Kaiser, and G. Oster. 2002. How myxobacteria glide. Curr. Biol. 12:369-377. Wu, S. S., and D. Kaiser. 1995. Genetic and functional evidence that Type IV pili are required for social gliding motility in Myxococcus xanthus. Mol. Microbiol. 18547-558. Wu, S. S., and D. Kaiser. 1996. Markerless deletions of pi1 genes in Myxococcus xanthus generated by counterselection with the Bacillus subtilis sacB gene. J. Bacteriol. 17858175821. Wu, S. S., and D. Kaiser. 1997. Regulation of expression of the pilA gene in Myxococcus xanthus. J. Bacteriol. 179:77487758. Wu, S. S., J. Wu, and D. Kaiser. 1997. The Myxococcus xanthus pilT locus is required for social gliding motility although pili are still produced. Mol. Microbiol. 23:109-121. Yang, R., S. Bartle, R. Otto, A. Stassinopoulos, M. Rogers, L. Plamann, and P. Hartzell. 2004. AglZ is a filament-forming coiled-coil protein required for adventurous gliding motility of Myxococcus xanthus. J. Bacteriol. 186:6168-6178. Yang, Z., Y. Geng, D. Xu, H. B. Kaplan, and W. Shi. 1998. A new set of chemotaxis homologues is essential for Myxococcus xanthus social motility. Mol. Microbiol. 30:1123-1130. Yang, Z., X. Ma, L. Tong, H. B. Kaplan, L. J. Shimkets, and W. Shi. 2000. Myxococcus xanthus dif genes are required for biogenesis of cell surface fibrils essential for social gliding motility. J. Bacteriol. 1825793-5798. Youderian, P., N. Burke, D. J. White, and P. L. Hartzell. 2003. Identification of genes required for adventurous gliding motility in Myxococcus xanthus with the transposable element mariner. Mol. Microbiol. 49:555570. Youderian, P., and P. L. Hartzell. 2006. Transposon insertions of magellan-4 that impair social gliding motility in Myxococcus xanthus. Genetics 172:1 397-14 10.
Myxobacteriu: Multicellularity and Differentiation Edited by David E. Whitworth 0 ZOOS ASM Press, Washington, D.C.
David R. Zusman Yuki F. I n c h Tiim Mignot
7
The Frz Chemosensory System of Myxococcus xanthus
MYXOCOCCUS XANTHUS EXHIBITS MANY SOCIAL BEHAVIORS Myxococcus xanthus has attracted much scientific interest because of its complex life cycle and morphogenetic potential. The bacteria grow in nature on complex organic material or prey upon other microorganisms. 111. xanthus cells are generally found in large groups (swarms) or biofilms. This social behavior facilitates predation and food gathering as large numbers of bacteria cooperate by producing antibiotics and digestive enzymes. When M. xanthus swarms are unable to find sufficient nutrients, they enter a developmental pathway in which they aggregate in a coordinated manner, forming raised pigmented mounds, 0.1 to 0.2 mm in height. Within the mounds, termed fruiting bodies, the cells differentiate to form spores. While the large majority of cells (80 to 90%)aggregate to form fruiting bodies, some cells follow a different developmental fate. These cells, called peripheral rods, remain as a monolayer of rod-shaped cells around and between fruiting bodies (O’Connor and Zusman, 1991). These cells do not aggregate or sporulate unless they are harvested and resuspended at high cell concentration on a fresh substrate. The peripheral rods move backwards and forwards in a rhythmic manner, forming “accordion waves” (Sliusarenko et al., 2006). The peripheral rods
/
have been hypothesized to be resting vegetative cells or scout cells, ready to feed if food or prey becomes available; spores are resting cells that cannot search for prey. The complex life cycle of 111. xantbus makes it an excellent bacterial system to study directed cell movements. The social behaviors of M . xanthus depend on cell motility. Individual bacterial cells move very slowly by gliding motility, about 2 to 4 p d m i n . While this slow rate of movement may be a disadvantage for cell dispersal, it may be advantageous for cell feeding, as it ensures that cells do not outrun their extracellular enzymes or their intercellular signals. Indeed, cells at the leading edge of a vegetative swarm venture outward but very quickly reverse direction, returning to the swarm (Reichenbach, 1999). Gliding motility is traditionally described as movement in the direction of the long axis of the cell at a solid-liquid or a solid-air interface without the aid of flagella (McBride, 2001). M . xanthus has two systems for gliding (Hodgkin and Kaiser, 1979).The first system is called adventurous (A)-motility and involves the movement of individual cells. A-motility is still not well understood, although many A-motility mutants have been isolated, and it is thought to require slime secretion (Youderian et al., 2003; Yu and Kaiser, 2007). Wolgemuth et al. hypothesized that directed extrusion of the slime, a polyelectrolyte gel, could
David R. Zusman, Yuki F. I n c h , and Tiim Mignot, Department of Molecular and Cell Biology, University of California, Berkeley, CA 94720-3204.
123
DEVELOPMENT AND MOTILITY
124 generate enough force to move cells forward (Wolgemuth et al., 2002),although recently Mignot et al. (2007)found evidence that suggests that A-motility is powered by the movement of specific adhesion complexes that track along helical cytoskeletal filaments. The second system is called social (S)-motility and involves the movement of cells in groups. S-motility requires type IV pili (Sun et al., 2000), lipopolysaccharide (LPS) O-antigen (Bowden and Kaplan, 1998), and extracellular matrix polysaccharide (EPS), which is a component of fibrils (Arnold and Shimkets, 1988). S-motility is similar to twitching motility in Pseudomonas aeruginosa and is powered by the extension and retraction of type IV pili. The pili are long, thin fibers that may be as long as 10 cell lengths (40 to 50 pm). They are extruded from one cell pole, where they adhere
to EPS or complex polysaccharides on another cell, the slime trail, or a prey microorganism. Retraction of the pili then pulls the cell in the direction of the adhering pili (Li et al., 2003; Sun et al., 2000).
THE frx GENES CONTROL CELLULAR REVERSALS The frz (frizzy) genes were discovered as part of a search for new mutants defective in cellular aggregation (Zusman, 1982). These mutants were defective at vegetative swarming and failed to aggregate into discrete mounds but instead formed “frizzy” filaments on starvation agar (Fig. 1).The first clue to the function of the frz genes came from observations of the motility pattern of single
A. Vegetative swarming on CYE agar (0.3%) Wild type, strain DZ2
Afrz (Z, A, B, CD, E or F)
B. Aggregation and development on CF starvation agar (1.5%) Wild type, strain DZ2
Afrz (Z, A, B, CD, E or F)
Figure 1 Developmental aggregation and vegetative swarming phenotypes of wild-type and frzCD mutants. Cells from M . xanthus wild-type strain DZ2 and Afrz ( Z ,A, B , CD, E , or F ) mutants were concentrated to 4 x l o 9 CFU ml-’ and spotted on CYE media containing 0.3% agar to analyze swarming or on CF agar (1.5%)to analyze fruiting body formation. The plates were examined under a dissecting microscope after 72 h of incubation at 32°C. The figure is modified from photographs published by Bustamante et al., 2004.
7. FRZ CHEMOSENSORY SYSTEMOF M.XANTHUS cells. When M. xanthus cells glide on an agar surface, cells reverse their direction of movement approximately every 7 to 8 min; net movement occurs since the interval between reversals can vary widely. The frz genes control the frequency at which cells reverse their direction of movement. For example, most frz mutants very rarely reverse direction; in contrast, some mutants in the receptor frzCD reverse much more frequently than the wild type, approximately every 2 min, and individual cells show no net movement (Blackhart and Zusman, 1985). These behavioral patterns suggested that the frz mutants might be similar to enteric chemotaxis mutants, as control of cellular reversals could affect directional movements. For example, enteric bacteria are known to direct their movements by undergoing a biased random walk. When the flagella rotate counterclockwise, the flagella form lefthanded helical bundles and the cells are pushed forward in a “run.” In contrast, when the flagella rotate clockwise, the flagellar bundles disperse and cells “tumble.” Tumbling results in randomly reorienting the bacteria. Thus, directed movements are achieved in these bacteria by controlling the run and tumble intervals. M. xanthus, in contrast, does not contain flagella and is nonmotile in a liquid medium. On a solid surface, cells predominantly move in existing slime trails, mostly in two dimensions. Since cells are flexible (individual cells can be seen bending), they require periodic directional corrections. Regulated cell reversals are proposed to be required for M. xanthus cells to undergo directed motility.
Frz PROTEINS SHOW SEQUENCE SIMILARITIESTO CHEMOTAXIS PROTEINS When the frz genes were sequenced, strong similaritieswere found between the Frz proteins and the major chemotaxis proteins of enteric bacteria (Fig. 2) (McBride et al., 1989).
~,
CheY-CheY
regulator
Chew
Chew
MCP
~~~~
125 In these bacteria, signals are recognized by chemoreceptors termed methyl-accepting chemotaxis proteins (MCPs). Receptors stimulate the activity of a histidine protein kinase (CheA)through interaction with a coupling protein, Chew (West and Stock, 2001). Typically an active CheA autophosphorylates and transfers a phosphoryl group to a single domain response regulator protein called CheY. In most flagellated bacteria, a change in direction is induced when phosphorylated CheY interacts with a switch component of the flagellar motor. The Frz system consists of FrzCD, a cytoplasmic chemoreceptor; FrzA and FrzB, Chew homologues; FrzE, a CheA-CheY-like fusion protein; FrzF, a methyltransferase; and FrzG, a methylesterase (McBride et al., 1989). FrzZ consists of two CheY-like domains connected by a linker region. frzZ is located 5’ to the frz operon but transcribed in the opposite orientation (Trudeau et al., 1996). Analysis of mutants containing inframe deletions showed that FrzCD (MCP), FrzA (Chew), and the CheA domain of FrzE constitute the core components of the Frz pathway, as they are essential for vegetative swarming, responses to repellents, and directed movement during development. FrzB (Chew), FrzF (CheR), FrzG (CheB),the CheY domain of FrzE, and FrzZ (CheY-CheY) are required for some but not all responses. Based on the Escherzchza coli paradigm, active FrzE should stimulate cellular reversals and inactive FrzE should inhibit cellular reversals (Ward and Zusman, 1990).
METHYLATION OF FrzCD IS CORRELATED WITH FACTORS AFFECTING CELL BEHAVIOR In enteric bacteria, methylation of receptors is required for adaptation to stimuli. Thus, the level of methylation of an MCP increases following the addition of an attractant and decreases following the addition of a
CheA-CheY
Response-
CheB
CheR
transferase
regulator
Figure 2 The frz operon contains genes that are homologous to proteins encoded by che genes from the enteric bacteria.
126 repellent. Methylated FrzCD, like the enteric MCPs, migrates as a ladder of bands that varies with the level of methylation during polyacrylamide gel electrophoresis. These MCP bands can be detected by Western immunoblot analysis using anti-FrzCD antibodies; FrzCD appears as multiple bands corresponding to the unmethylated (amidated and deamidated) and methylated forms of the receptor. Western blot analysis showed that vegetative cells are highly methylated (about 50% of FrzCD is methylated) but cells that are starved are relatively unmethylated (McBride et al., 1989). Developmental cells show an initial loss in methylation followed by an increase in the level of methylation so that by 72 h of development, when cells are mostly in fruiting bodies, FrzCD is about 70% methylated. The methylation changes in FrzCD suggest that during aggregation, FrzCD senses a chemical(s) produced by other cells that promotes cell movements towards aggregation centers (Geng et al., 1998). Geng et al. (1998) found that developmental mutants could be divided into two groups based on the level of FrzCD methylation: nonaggregating or abnormally aggregating mutants, including the asg, bsg, csg, and esg mutants, showed poor FrzCD methylation. Mutants blocked in late development and sporulation showed normal FrzCD methylation. Thus, the methylation of FrzCD defines a discrete step in the developmental program of M . xanthus. Indeed, Ssgaard-Andersen and Kaiser (1996) found that the csgA mutant, which does not show FrzCD methylation during development, can be stimulated to methylate FrzCD when cells are treated with C-factor and rescued for development. In an attempt to identify chemicals or nutrients that might be potential attractants or repellents for M. xanthus, numerous substances were tested to determine whether they affected the methylation state of FrzCD. Although the methylation of FrzCD was stimulated by the addition of rich media containing peptides (Casitone yeast extract [CYE] medium), yeast extract, or several defined chemicals including lauric acid and lauryl alcohol, individual amino acids, sugars, or nucleotides had no effect. Surprisingly, some phospholipids such as phosphatidyl ethanol did stimulate methylation of FrzCD (McBride et al., 1992). In contrast, several short-chain alcohols and dimethyl sulfoxide caused the demethylation of FrzCD.
DOES M. XANTHUS EXHIBIT CHEMOTAXIS ? The complex movements and behavior of M. xanthus suggested chemotaxis, but demonstrating chemotaxis and defining chemoeffectors were difficult to
DEVELOPMENT AND MOTILITY establish experimentally. For example, Dworkin and Eide (1983)tried but were unable to show chemotaxis of M . xanthus to a wide variety of potential chemoeffectors and suggested that chemotaxis may not be possible in M. xanthus since cells move so slowly, slower than the rate of diffusion of some small molecules. However, the rate of diffusion of peptides in agar is much slower than in buffer, even slower than the rate of movement of M. xanthus cells (R. Welch, personal communication). Shi et al. (1993) employed chambered petri dishes to establish sharp chemical gradients and found directed movement of M. xanthus cells towards yeast extract and Casitone and away from dimethyl sulfoxide and isoamyl alcohol. Furthermore, these movements were completely dependent on the Frz system, supporting the hypothesis that the Frz chemosensory system controls directed movements in this organism. Chemotaxis in M. xanthus was also documented by Kearns and Shimkets, who found that dilauroyl and dioleoyl phosphatidylethanolamine (PE) stimulated directed cell movement in a gradient on an agar surface (Kearns et al., 2002). They used changes in reversal period to show that M . xanthus responds to and adapts to these lipids. Unexpectedly, stimulation was not dependent on the Frz pathway although adaptation was dependent onthe pathway. They noted that directed movements towards these lipids were indeed chemotaxis since (i) biased movements were correlated with the suppression of reversals to achieve longer runs when exposed to PE, (ii)the responses were specific to PE molecules with particular fatty acids, and (iii) cells showed adaptation to the lipid attractants.
THE frz GENES REGULATE BOTH THE A- AND S-MOTILITY SYSTEMS Although it was known for many years that the f y z genes were required for normal aggregation during fruiting body formation, it was not known if they controlled the Amotility system, the S-motility system, or both. To investigate this point, the swarming ability of cells containing a single motility system was studied (V. H. Bustamante and D. R. Zusman, unpublished data). An A-S+ mutant (aglBI),which can move only by S-motility, swarms as well as the wild type on 0.3% CYE agar, but swarming is reduced and disorganized in an aglB2 frzCD double mutant. Thus, the Frz system is required for the organized spreading of M. xanthus colonies. In contrast, an A+S-mutant (pilA),which can move only by A-motility, fails to swarms at all on 1.5% CYE agar. However, a pilA fyzCD double mutant shows restored movement and spreads at about the same rate as an frzCD mutant. Thus, the Frz system inhibits the movement of A-motile
SYSTEMOF 111. XANTHUS 7. FRZ CHEMOSENSORY cells on rich media; it should be noted that this inhibition is nutrient dependent. The suppression of A-motility by the Frz system remains to be investigated.
THE f k z GENES REGULATE S-MOTILITY-DEPENDENT REVERSALS S-motility in M . xanthus, like twitching motility in P. aeruginosa, has been shown to involve the extension and retraction of type IV pili localized at the leading cell pole. The pili are extended from the cell pole, where they adhere to exopolysaccharides on the surface of another cell or the cell surface (slime trails) (Li et al., 2003). This interaction triggers pilus retraction, which pulls the cells forward in the direction of the adhering pili. Cellular reversals must therefore result from the sites of pilus extension switching from one cell pole to another. This switching is controlled by the f ~ chemosensory z system. Sun et al. (2000) developed an assay allowing single cells to move using only S-motility. Cells were placed on microscope slides and overlaid with 1% methylcellulose medium. Under these conditions, the pili presumably bind to the surface of slides coated with methylcellulose. The binding of pili to the slides “tether” the cells (they appear to be spherical instead of rod-shaped, from an end-on view). These tethered cells were followed by timelapse videomicroscopy. The cells remained bound to the surface for about 8 min, after which they were released, which corresponds to the reversal interval of wild-type cells when gliding on a solid surface. frzA-F mutants, which rarely reverse on an agar surface, remained tethered for extended periods of time; in contrast, constitutive signaling mutants, like the frzCD::TnSa224 mutant, remained tethered for only 2 min, which corresponds to the reversal period for these cells on a solid surface. The behavior of the tethered cells is consistent with the pili being extruded from one cell pole, adhering to a surface, and then retracting, pulling the cell in the direction of the adhering pili. This process is controlled by the frz chemosensory system.
ANALYSIS OF THE CYTOPLASMIC RECEPTOR f.xCD FrzCD, the Frz system chemoreceptor, contains a conserved C-terminal module present in MCPs; but in contrast to most MCPs, FrzCD is localized in the cytoplasm. Immunofluorescence and deconvolution microscopy showed that it is localized as an array of discrete clusters (D. P. Astling, E. M. F. Mauriello, and D. R. Zusman, unpublished data). Since the N-terminal region of FrzCD does not contain the canonical periplasmic
127
domain, a series of in-frame deletion mutants were constructed in fyzCD to determine the function of the various domains of FrzCD. Surprisingly, deletion of the N-terminal region of FrzCD (codons 6 to 130) showed only minor defects in swarming, and development was normal. Thus, the N-terminal region of FrzCD probably is not directly involved in sensing signals: signal input to the Frz system must be sensed by the conserved Cterminal module of FrzCD (Bowden and Kaplan, 1998). Perhaps the methyltransferase (FrzF) may be regulated to recognize different methylation sites in the C-terminal module of FrzCD. Alternatively, FrzCD may interact with another unidentified protein or MCP that could transduce a signal. Interestingly, deletion of about 25 amino acids from either end of the conserved C-terminal region of FrzCD resulted in a constitutive signaling state of FrzCD, which induces hyperreversals with no net cell movement. These deletions may result in FrzCD locked in a constitutively signaling conformation or deleted regions may contain potential interaction sites. Since the mechanisms governing methylation of a chemoreceptor can differ and could potentially provide a mechanism for differential responses by the bacteria to different stimuli, methylation site mutants of FrzCD were constructed and analyzed (Astling et al., 2006). For this study, potential methylation sites of FrzCD were systematically modified by site-directed mutagenesis, changing glutamine/glutamate pairs to alanines. Two of the sites, when mutated, had a stimulatory effect on the pathway (causing constitutive signaling), as evidenced by cells hyperreversing. In contrast, two other sites, when mutated, had an inhibitory effect on the pathway, causing cells to rarely reverse. This indicates that the methyltransferase can both activate and inhibit the Frz pathway, depending on which sites are modified by methylation. The stimulatory mutations blocked both vegetative swarming and developmental aggregation. The inhibitory mutations blocked developmental aggregation at low cell density, but not at high cell density, suggesting that specific methylation sites may be required for sensing low concentrations of developmental signals. The different phenotypes of the mutants observed in this study suggest that differential methylation could provide a potential signal input to the Frz chemosensory pathway.
ROLE OF THE METHYLTRANSFERASE FrzF I N FrzCD METHYLATION If the N-terminal domain of FrzCD is not required for sensing signals, how are these signals detected? Is regulated methylation/demethylation important for responses? To
128 address these questions, in-frame deletion mutants of frzG, which encodes a methylesterase, and frzF, which encodes a methyltransferase, were constructed. The frzG deletion mutant had only a minor impact on swarming and development. In contrast, the frzF mutant was defective in development (forming frizzy filaments) and in vegetative swarming. FrzF, unlike CheR from E. coli, is a large protein that contains three tetratricopeptide repeats (TPR) motifs, which are typically involved in protein-protein interactions. To see if the repeats were important for the regulation of methyltransferase activity, an in-frame deletion mutant lacking the TPR domains was constructed (I. Martinez-Flores, V. H. Bustamante, A. E. Scott, and D. R. Zusman, unpublished data). The mutant retained the ability to methylate FrzCD, albeit at a reduced level, but was unable to form fruiting bodies. However, the frzF,,, mutant was able to swarm on rich media. This indicates the importance of the TPR motifs for regulating FrzCD methylation during developmental aggregation. Proteins that might interact with the TPR domains are currently being investigated.
REGULATION OF THE Frz PATHWAY BY A NOVEL CheW-LIKE PROTEIN, FrzB In E. coli, the main apparent role of Chew is to facilitate the interactions of the receptor with the kinase, CheA. The Frz pathway has two Chew homologues, FrzA and FrzB. FrzB is a CheW-like protein with a novel regulatory role. Mutations in frzB give a phenotype similar to that of frzA-E mutants with respect to vegetative swarming and developmental aggregation; however, frzB mutants can still respond to repellents. This suggests that FrzB is not a part of the core Frz signal transduction pathway but acts as an accessory factor or regulator of the pathway, like FrzF and FrzG. We examined the interaction of both FrzA and FrzB with FrzCD and FrzE using the yeast two-hybrid interaction assay and pull-down assays with purified proteins (Astling, 2003). Using these assays, we found that FrzA can interact with both the receptor, FrzCD, and the CheA, FrzE. In addition, FrzA and FrzCD stimulate the kinase activity of FrzE in vitro. Thus, FrzA is a true Chew homolog. FrzB, on the other hand, interacted with FrzCD but not with FrzE, which is consistent with the observation that it lacks the CheAbinding domain. We used surface plasmon resonance spectroscopy to examine the kinetics of the interaction of FrzA and FrzB with FrzCD (D. P. Astling and D. R. Zusman, unpublished data). From this analysis, we obtained the kinetic parameters of both binding events. The data best fit a bivalent model where one receptor dimer can bind two molecules of FrzA or FrzB. FrzA has a greater
DEVELOPMENT AND MOTILITY affinity for FrzCD than does FrzB. We hypothesize that FrzB plays a role in receptor clustering or alternatively, in coupling the receptor to additional proteins. To test the latter hypothesis, we used the yeast two-hybrid interaction assay to search for proteins that interact with FrzB. We found several interacting proteins, two of which were MCPs. This is relevant because M. xanthus has 21 putative MCPs and eight CheA proteins. The significance of these interactions is currently being investigated.
ANALYSIS OF FrzE, A CheA-CheY HYBRID The Frz system regulates reversal frequency in both the A- and S-motility systems. How does a single chemotaxis-like pathway regulate coordinated signaling to two disparate motility systems? FrzE, a CheA-CheY fusion, plays a large role in this regulation. FrzE is essential for Frz signaling as shown by genetic studies (Blackhart and Zusman, 1985). The CheA domain of FrzE ( FrzEcheA)autophosphorylates and transfers the phosphate to downstream components, ultimately causing a reversal. Recent studies revealed that the activity of FrZECheAis regulated by the CheY domain of FrzE. Genetic analyses showed that the state of the CheY domain can independently influence the reversal period of the A- and S-motility systems (Bonner et al., 2005), and in vitro studies showed that this regulation occurs by inhibiting autophosphorylation activity of FrzECheA (Y. F. I n c h and D. R. Zusman, unpublished data) as described below. A deletion mutant of frzE, AfrzE, is unable to coordinate cellular movements and results in the typical Frz phenotype: reduced cellular reversals, decreased swarming, and frizzy aggregates on developmental media (Bustamante et al., 2004). However, a deletion of the CheY domain of FrzE, frzEAChcY,displayed differential cellular reversal periods with respect to A- and S-motility, defective vegetative swarming, and surprisingly, the formation of fruiting bodies on developmental media (Bonner et al., 2005). To further investigate the function of the CheY domain, point mutations of the conserved phospho-accepting aspartate residue in the CheY domain of FrzE (FrzEchey)were constructed such that the encoded proteins potentially mimic the constitutively active (FrzE-D709E) or constitutively inactive (FrzE-D709A)conformation observed for some response regulators (Klose et al., 1993). Single-cell motility analysis revealed that both frzE-D709A and frzE-D709E hyperreverse with respect to the A-motility system (Bonner et al., 2005). Because FrzECheycannot regulate FrzE autophosphorylation activity in these mutants, FrZEcheA
7. FRZ CHEMOSENSORY SYSTEMOF M.XANTHUS is most likely phosphorylated at increased levels compared to the wild type. Thus, phospho-signaling to the A-motility system is elevated, resulting in the observed hyperreversing phenotypes. Conversely, the frzE-D709A mutant displayed hyperreversals with respect to the Smotility system in contrast to the frzE-D709E mutant, which displayed hyporeversals. It was hypothesized that FrzECheydirectly regulates reversals of the S-modity system by a physical interaction that is enhanced in the frzE-D709A mutant. However, the double mutant frzEAChey AfrzZ hyporeversed with respect to both motility systems, suggesting that FrzZ regulates both systems downstream of FrzE. Further information as to how the CheY domain affects activity of the CheA domain came from in vitro analyses. To complement the genetic analyses, the Frz system was reconstituted in vitro to test biochemical activities by purifying FrzCD, FrzA, FrzE, and the independent domains of FrzE: FrzECheA and FrZECheY (Inclhn et al., 2007). FrzECheA autophosphorylated in the presence of FrzCD, FrzA, and ATP. FrzCD and FrzA were required for significant autophosphorylation. Surprisingly, the full-length FrzE protein did not autophosphorylate in the presence or absence of FrzCD and FrzA, suggesting that the CheY domain of FrzE inhibits autophosphorylation activity of the CheA domain. These genetic and biochemical data suggest that FrzE has divergent signaling effects on both the A- and S-motility systems and that FrzECheynegatively regulates FrzE autophosphorylation. It is unclear how this regulation is accomplished in vivo, and this is currently under investigation.
THE Frz PATHWAY OUTPUT The output pathway from the Frz system to the motility engines is still unknown. We predicted that additional unidentified proteins interact with FrzE and relay signals from the FrzCD, FrzA, and FrzE complex to the downstream motility components, but these downstream proteins remain elusive. To search for these downstream genes, I n c h et al. (2007) used a genetic screen for suppressors of a constitutively active FrzCD‘ mutant. Since the frzCDc colonies are very compact (cells hyperreverse and show no net translocation), suppressor mutants could easily be identified by their spreading phenotype. Presumably, only mutations in the frz genes or in genes that function downstream of the Frz chemosensory pathway would allow cells to escape the hyperreversing FrzCD‘ phenotype. Using this strategy, frzZ was found to suppress frzCDc,revealing that FrzZ is a downstream component of the Frz
129
pathway. FrzZ is a CheY-CheY fusion protein. frzZ deletion mutants show the typical Frz phenotype, including decreased cellular reversal frequencies in both the Aand S-motility systems (Bustamante et al., 2004). AfrzZ is also epistatic to frzEACheY, again suggesting that FrzZ acts downstream of FrzE. Both CheY domains of FrzZ contain the highly conserved Asp residues necessary for phosphotransfer from FrZECheA (Trudeau et al., 1996). In vitro experiments with [32P]FrzECheA demonstrated that both CheY domains of FrzZ can rapidly accept phosphate from FrzECheAand that phosphorylation occurs on the predicted aspartate residues (Inclhn et al., 2007). Based on these data, we propose that FrzZ mediates signals from FrzE to the downstream motility systems (Color Plate 2).
A DOWNSTREAM OSCILLATOR CONTROLS REVERSAL OF THE A- AND S-ENGINES Genetic analysis has shown that reversals for both the A- and S-motility engines are controlled by Frz signaling (Li et al., 2003). However, the link between the Frz pathway and the two motility engines and how it regulates reversal frequency remain major unsolved questions. The Frz system could be a branched pathway that signals each motility engine independently. This model would involve complex cross talk to integrate signaling output for two motility engines. Alternatively, the Frz system may communicate with a single protein that triggers coordinated reversals. The latter model predicts that disruption of the common output protein should disrupt both A- and S-motility. Furthermore, this model suggests that the output protein should interact with coupling proteins from each of the two motility systems. We have recently obtained evidence from the study of FrzS, an S-motility protein, and AglZ, an A-motility protein, that suggests that these similarly constructed proteins may be engine-specific output proteins for the two motility systems and that they regulate cell reversals. FrzS and AglZ have remarkably similar modular structures: a pseudoreceiver domain at their N terminus that is connected to an extended “coiled-coil’’ domain by a flexible alanine-proline-rich linker region. Interestingly, the function of each protein is related to a single motility system: FrzS and AglZ are specifically required for S- and A-motility, respectively (Ward et al., 2000). Microscopy and genetic studies have shown that FrzS is not a structural component of the S-engine, but more likely a regulator of S-motility (Ward et al., 2000). AglZ is structurally related to FrzS and may therefore also be a regulator of the A-engine rather than one of its structural components.
130
A strain that expressed a functional FrzS-green fluorescent protein (GFP) fusion was constructed and monitored in moving cells by fluorescence microscopy (Mignot et al., 2005). FrzS was observed to oscillate from pole to pole as cells reverse. Immediately after a reversal, FrzS accumulated at the leading, piliated pole; as cells moved forward, FrzS began to also accumulate at the lagging cell pole, until equivalent amounts of the FrzS protein were found at both poles. The cell then reversed, and FrzS rapidly relocalized from the old leading pole to the new leading pole. Importantly, the oscillations of FrzS were regulated by the signaling activity of the Frz system: pole-to-pole switching of FrzS was rarely observed in the frzE mutant, whereas a constitutive frzCDc mutation induced hyperreversals and concomitant hyperoscillations of FrzS (Mignot et al., 2005). The proposed mechanism of FrzS pole-to-pole trafficking is not discussed here, as it is described in detail in chapter 6. FrzS was also studied by constructing in-frame deletion mutants that express stable cryptic proteins; these mutants showed loss of function and/or distinct localization defects. Specifically, in-frame deletions of the pseudoreceiver domain, the coiled-coil domain, and a motif located at the very C terminus of the protein each resulted in aberrant localization and loss of function defects (Mignot et al., 2007a). Removal of the pseudoreceiver domain caused preferential targeting of the protein to the lagging end of the cells (normal localization is to the leading cell pole), whereas deletion of the C-terminal tail led to weak localization to the leading cell pole. Deletion of both the pseudoreceiver domain and the C-terminal tail resulted in complete loss of polar localization, a phenotype also observed when the coiled-coil domain was deleted. This analysis showed that pole-to-pole oscillations of FrzS result from complementary roles for the protein domains: the pseudoreceiver domain is essential for accumulation of FrzS at the leading pole, whereas the C-terminal tail is a polar anchoring factor that affects localization at both the leading pole and the lagging pole. Based on these experiments, we hypothesize that (i) signaling to the pseudoreceiver domain results in loss of affinity of FrzS for the leading pole, targeting it to the lagging pole; (ii) this targeting is facilitated by the C-terminal tail; and (iii) FrzS is transported from pole to pole via the coiled-coil domain. Since AglZ is similar in structure to FrzS, we studied the dynamic localization of AglZ, anticipating that it might provide information about cellular reversals mediated by the A-engine. A strain that expressed a functional AglZ-yellow fluorescent protein (YFP) fusion protein was constructed, and its localization was monitored by fluorescence microscopy (Mignot et al.,
DEVELOPMENT AND MOTILITY 2007b). We observed that the localization of AglZ was dependent on the activity of the A-engine: when the cells were actively moving, AglZ-YFP was localized in a series of clusters that spanned the cell length; in contrast, nonmoving cells showed AglZ-YFP localized at one cell pole or diffused in the cytoplasm. When moving cells reversed, the clusters dispersed and AglZ was rapidly relocalized to the new leading pole. As cell movement resumed, AglZ redistributed as ordered clusters from the leading pole. These AglZ oscillations were also regulated by the Frz system: AglZ never switched poles in the frzE mutant and the frzCD“ mutant displayed hyperoscillations of AglZ. These studies show that in M. xanthus, cellular reversals involve regulated oscillations of A- and S-motility proteins that are targeted to the new leading cell pole at the time of reversal.
HYPOTHESES AND FUTURE PERSPECTIVES The fact that proteins from both motility systems oscillate together from pole to pole during reversals suggests that a common regulator controls their dynamics. It is unlikely that FrzS and AglZ are directly phosphorylated by FrzE because FrzS and AglZ lack the critical aspartate residue that is phosphorylated in canonical receiver domains (Fraser et al., 2007) and FrzZ is known to be the cognate response regulator for phospho-FrzE. MglA is an ideal candidate for the common output regulator of the Frz pathway, as it is essential for both A- and S-motility (Spormann, 1999; see also chapter 6). Interestingly, MglA is homologous to small GTPases of the Ras family such as the yeast Sarl protein. In eukaryotic cells, small GTPases act as regulatory proteins often by recruiting factors to their site of action. It is therefore possible that MglA acts to recruit factors that are important for reversal of the two motility systems. Consistent with this hypothesis, MglA is essential for the localization of both FrzS and AglZ: in an mglA mutant, FrzS could localize to only one cell pole and AglZ was almost completely diffuse in the cytoplasm. This is likely a direct effect because protein interaction studies have shown that MglA can interact directly with FrzS and AglZ and MglA colocalizes with FrzS and AglZ. We speculate that the Frz pathway acts on MglA, which in turns recruits motility proteins to their sites of action to regulate cellular reversals. Cellular reversals are regulated by two molecular oscillators. The activator of the upstream oscillator consisting of the Frz chemosensory proteins and referred to as the “Frzilator” (Igoshin et al., 2004), is essential for operating the second oscillator, composed of structural components of the A- and S-motility proteins that periodically
7. FRZ CHEMOSENSORY SYSTEMOF M.
XANTHUS
oscillate from pole to pole (Color Plate 2). Indeed, core frz mutants are defective at cell reversals and pole-to-pole oscillations, showing that the downstream oscillator is not operational in these mutants. The periodicity of the downstream oscillator is probably regulated by signal flow from the Frz system: indeed, constitutive signaling from Frz leads to hyperoscillations of the polar proteins. How then does the Frz system regulate the oscillations of the downstream proteins? We hypothesize that Frz signaling somehow activates MglA, the common regulator of both motility systems (Color Plate 2). To validate this hypothesis, it will be critical to elucidate how the Frz system acts on MglA and how MglA then acts to regulate the downstream oscillator. For example, one could imagine a scenario where FrzZ acts as a shuttle to transduce signals directly from FrzE to MglA or MglB, a putative regulator of the GTPase activity of MglA. The other proteins recruited by MglA and the dynamics of their interactions are unknown. These proteins are the likely missing links that couple the Frz chemosensory system with the two engines of motility that propel M. xanthus. We are grateful to members of the Zusman laboratory, past and present, for many helpful discussions. The research in our laboratory was supported by a grant from the National Institutes of Health (GM20509).
References Arnold, J. W., and L. J. Shimkets. 1988. Cell-surface properties correlated with cohesion in Myxococcus xanthus. J. Bacterial. 170:5771-5777. Astling, D. P. 2003. Novel Regulatory Mechanisms of a Chemotaxis Pathway in the Gliding Bacterium Myxococcus xanthus. Ph.D. thesis. University of California, Berkeley. Astling, D. P., J. Y. Lee, and D. R. Zusman. 2006. Differential effects of chemoreceptor methylation-domain mutations on swarming and development in the social bacterium Myxococcus xanthus. Mol. Microbiol. 59:45-55. Blackhart, B. D., and D. R. Zusman. 1985. “Frizzy” genes of Myxococcus xanthus are involved in control of frequency of reversal of gliding motility. Proc. Natl. Acad. Sci. USA 82:8767-8770. Bonner, P. J., Q. Xu, W. P. Black, Z. Li, Z. Yang, and L. J. Shimkets. 2005. The Dif chemosensory pathway is directly involved in phosphatidylethanolamine sensory transduction in Myxococcus xanthus. Mol. Microbiol. 571499-1508. Bowden, M. G., and H. B. Kaplan. 1998. The Myxococcus xanthus lipopolysaccharide 0-antigen is required for social motility and multicellular development. Mol. Microbiol. 30~275-284. Bustamante, V. H., I. Martinez-Flores, H. C. Vlamakis, and D. R. Zusman. 2004. Analysis of the Frz signal transduction system of Myxococcus xanthus shows the importance of the conserved C-terminal region of the cytoplasmic chemoreceptor FrzCD in sensing signals. Mol. Microbiol. 53:15011513.
131 Dworkin, M., and D. Eide. 1983. Myxococcus xanthus does not respond chemotactically to moderate concentration gradients. J. Bacteriol. 154:437442. Fraser, J. S., J. P. Merlie, Jr., N. Nichols, S. R. Westfield, T. Mignot, D. E. Wemmer, D. R. Zusman, and T. Alber. 2007. An atypical receiver domain controls the dynamic polar localization of the Myxococcus xanthus social motility protein FrzS. Mol. Microbiol. 65:317-332. Geng, Y., Z. Yang, J. Downard, D. Zusman, and W. Shi. 1998. Methylation of FrzCD defines a discrete step in the developmental program of Myxococcus xanthus. J. Bacteriol. 180:5765-5768. Hodgkin, J., and D. Kaiser. 1979. Genetics of gliding motility in Myxococcus xanthus (Myxobactera1es)-two gene systems control movement. Mol. Gen. Genet. 171:177-191. Igoshin, 0. A., A. Goldbeter, D. Kaiser, and G. Oster. 2004. A biochemical oscillator explains several aspects of Myxococcus xanthus behavior during development. Proc. Natl. Acad. Sci. USA 101:15760-1 5 765. Inclh, Y. F., H. C. Vlamakis, and D. R. Zusman. 2007. FrzZ, a dual CheY-like response regulator, fuctions as an output for the Frz chemosensory pathway of Myxococcus xanthus. Mol. Microbiol. 65:90-102. Kearns,D.B.,P. J.Bonner,D.R. Smith, andL. J. Shimkets. 2002. An extracellular matrix-associated zinc metalloprotease is required for dilauroyl phosphatidylethanolamine chemotactic excitation in Myxococcus xanthus. J. Bacteriol. 184~1678-1684. Klose, K. E., D. S. Weiss, and S. Kustu. 1993. Glutamate at the site of phosphorylation of nitrogen-regulatory protein NTRC mimics aspartyl-phosphate and activates the protein. J. Mol. Biol. 232:67-78. Li, Y., H. Sun, X. Ma, A. Lu, R. Lux, D. Zusman, and W. Shi. 2003. Extracellular polysaccharides mediate pilus retraction during social motility of Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 1005443-5448. McBride, M. J. 2001. Bacterial gliding motility: multiple mechanisms for cell movement over surfaces. Annu. Rev. Microbiol. 55:49-75. McBride, M. J., T. Kohler, and D. R. Zusman. 1992. Methylation of FrzCD, a methyl-accepting taxis protein of Myxococcus xanthus, is correlated with factors affecting cell behavior. J. Bacteriol. 174:4246-4257. McBride, M. J., R. A. Weinberg, and D. R. Zusman. 1989. “Frizzy” aggregation genes of the gliding bacterium Myxococcus xanthus show sequence similaritiesto the chemotaxis genes of enteric bacteria. Proc. Natl. Acad. Sci. USA 86:424-428. Mignot, T., J. P. Merlie, and D. R. Zusman. 2005. Regulated pole-to-pole oscillations of a bacterial gliding motility protein. Science 310:855-857. Mignot, T., J. P. Merlie, Jr., and D. R. Zusman. 2007a. Two localization motifs mediate polar residence of FrzS during cell movement and reversals of Myxococcus xanthus. Mol. Microbiol. 65:363-372. Mignot, T., J. W. Shaevitz, P. L. Hartzell, and D. R. Zusman. 2007b. Evidence that focal adhesion complexes power bacterial gliding motility. Science 315:853-856. O’Connor, K. A., and D. R. Zusman. 1991. Development in Myxococcus xanthus involves differentiation into two cell
132 types, peripheral rods and spores. J. Bacteriol. 173:33183333. Reichenbach, H. 1999. The ecology of the myxobacteria. Enviyon. Microbiol. 1:15-21. Shi, W., T. Kohler, and D. R. Zusman. 1993. Chemotaxis plays a role in the social behaviour of Myxococcus xanthus. Mol. Microbiol. 9:601-611. Sliusarenko, O., J. Neu, D. R. Zusman, and G. Oster. 2006. Accordion waves in Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 103:1534-1539. Ssgaard-Andersen, L., and D. Kaiser. 1996. C factor, a cellsurface-associated intercellular signaling protein, stimulates the cytoplasmic Frz signal transduction system in Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 93:2675-2679. Spormann, A. M. 1999. Gliding motility in bacteria: insights from studies of Myxococcus xanthus. Microbiol. Mol. Biol. Rev. 63:621-641. Sun, H., D. R. Zusman, and W. Y. Shi. 2000. Type IV pilus of Myxococcus xanthus is a motility apparatus controlled by the frz chemosensory system. Curr. Biol. 10:1143-1146. Trudeau, K. G., M. J. Ward, and D. R. Zusman. 1996. Identification and characterization of FrzZ, a novel response regulator necessary for swarming and fruiting-body
DEVELOPMENT AND MOTILITY formation in Myxococcus xanthus. Mol. Microbiol. 20:645655. Ward, M. J., H. Lew, and D. R. Zusman. 2000. Social motility in Myxococcus xanthus requires FrzS, a protein with an extensive coiled-coil domain. Mol. Microbiol. 37:13571371. Ward, M. J., and D. R. Zusman. 1999. Motility in Myxococcus xanthus and its role in developmental aggregation. Curr. Opin. Microbiol. 2:624-629. West, A. H., and A. M. Stock. 2001. Histidine kinases and response regulator proteins in two-component signaling systems. Trends Biochem. Sci. 26:369-376. Wolgemuth, C., E. Hoiczyk, D. Kaiser, and G . Oster. 2002. How myxobacteria glide. Curr. Biol. 12:369-377. Youderian, P., N. Burke, D. J. White, and P. L. Hartzell. 2003. Identification of genes required for adventurous gliding motility in Myxococcus xanthus with the transposable element mariner. Mol. Microbiol. 4955.5-570. Yu, R., and D. Kaiser. 2007. Gliding motility and polarized slime secretion. Mol. Microbiol. 63:454-467. Zusman, D. R. 1982. “Frizzy” mutants: a new class of aggregation-defective developmental mutants of Myxococcus xanthus. J. Bacteriol. 150:1430-1437.
Regulatory Mechanisms
Myxobacteria: Multicellularity and Differentiation Edited by David E. Whitworth 0 2008 ASM Press, Washington, D.C.
John R. Kirby, James E. Berleman, Susanne Muller, Di Li, Jodie C. Scott, Janet M. Wilson
Chemosensory Signal Transduction Systems in Myxococcus xanthus
Many microbes have been analyzed for their capacity to utilize signal transduction in order to navigate their environment by responding to a multitude of stimuli including physical, chemical, and biological cues. The majority of the analyses have focused on two-component signal transduction (TCST) systems, even though these are likely to compose only a fraction of the total systems that have evolved to translate information derived from the environment (Ulrich et al., 2005). The prototype for the TCST system is composed of a sensor kinase (or phosphatase) and a response regulator. The sensor kinase is phosphorylated on a conserved histidine residue using ATP as the donor and subsequently passes the phosphoryl group to a conserved aspartate residue within the response regulator (Fig. 1).The domain architecture for TCST systems is highly variable but typically consists of an input sensor domain that is periplasmic (or extracellular in gram-positive bacteria), responds directly to environmental signals, and is covalently linked to the kinase domain. The prototypical response regulator contains the aspartyl receiver domain covalently linked to an output domain which binds DNA to affect transcription (Parkinson and Kofoid, 1992; Parkinson, 1993; Hoch and Silhavy, 1995; Stock et al., 2000; Hoch,
8
2000). Variable inputs and outputs allow these systems to respond to a great number of stimuli and subsequently regulate specific subsets of genes or operons. TCST systems in Myxococcus xanthus are discussed in more detail in chapter 10. The best-studied TCST system is the chemotaxis system (Fig. 1)that regulates flagellar motility in Escherichia coli. Forty years of research in the field of chemotaxis has led to a great understanding of the overall mechanism governing the behavioral responses to chemical stimuli. The most detailed description to date has been given for the response by E. coli to the chemoattractant aspartate that is mediated by Tar, a methyl-accepting chemotaxis protein (MCP) that also mediates the response to maltose (via the maltose binding protein) and two repellents, Ni2+ and Co2+.When the periplasmic ligand binding site is titrated with aspartate, the Tar homodimeric receptor undergoes a conformational change that influences the cytoplasmic Chew-CheA ternary signaling complex. Aspartate-bound Tar influences phosphotransfer from CheA to CheY such that the concentration of the phosphorylated CheY response regulator transiently decreases. Diminished levels of phospho-CheY (the tumble regulator) lead to prolonged duration of
John R. Kirby, James E. Berleman, Susanne Miiller, Di Li, Jodie C. Scott, and Janet M. Wilson, The University of Iowa, Department of Microbiology, 51 Newton Rd., Iowa City, IA 52242.
135
REGULATORY MECHANI sM s
136
B
C
Figure 1 Domain topology of TCST systems. (A) A prototypical TCST system is shown. A variable input (sensor) domain is covalently bound to the histidine kinase domain. Phosphorylation occurs on a conserved histidine residue. The phosphoryl group is transferred to a conserved aspartate residue within the receiver domain (Rec) in the response regulator. The prototypical RR output is a DNA-binding domain capable of influencing gene expression. The vertical bar represents the cytoplasmic membrane. (B) The specialized TCST system that controls chemotaxis is shown. The MCP transducer is depicted as transmembrane and is coupled by Chew to the CheA kinase. Only two methyl groups are shown to represent methylation of the receptor by CheR. Phosphorylated CheB can remove these methyl groups. Methylation is a hallmark feature of the chemotaxis TCST systems. Phosphotransfer to the response regulator CheY influences its ability to bind the FliM switch component at the flagellar motor. (C) A chemosensory system such as the Che3 system found in M . xanthus is depicted. Chemosensory systems represent a composite of the prototypical TCST and the specialized chemotaxis TSCT systems.
counterclockwise rotation of the flagellar motor, thereby producing a swimming event in response to the attractant aspartate. The flux of phosphoryl groups (or diminution) is attenuated by a second posttranslational modification, methylation, of the MCP chemoreceptor. CheR acts
constitutively as a methyltransferase (utilizing S-adenosylmethionine as the methyl donor) to methylate specific glutamate residues (generating glutamate-0-methyl esters) located within two cytoplasmic domains of the Tar receptor. CheB acts as a methylesterase to demethylate
8.
CHEMOSENSORY SIGNALTRANSDUCTION SYSTEMSIN M. XANTHUS
the glutamate methyl esters regenerating the glutamate residues within the Tar receptor. CheB is a response regulator and is regulated by phosphorylation by the CheA kinase. Therefore, CheA regulates two response regulators, CheY and CheB. Because CheB requires phosphorylation by CheA prior to becoming an active methylesterase, demethylation lags temporally with respect to the flux of phosphoryl groups transferred to CheY. Methylating the Tar receptor serves to upregulate the CheA kinase, thereby restoring the flux of phosphoryl group transfer to CheY. This results in adaptation to prestimulus levels even in the presence of the stimulating ligand, aspartate, and constitutes a feedback loop. Adaptation allows the cell to compare the chemical environment over time and adjust its behavior such that the cell migrates toward more favorable conditions (Falke et al., 1997; Falke and Hazelbauer, 2001; Bourret and Stock, 2002). Many other TCST systems have been analyzed including those that regulate responses to quorum sensing (Bassler, 2002), virulence (Krukonis and DiRita, 2003), protein folding (DiGiuseppe and Silhavy, 2003), osmolarity (Qin et al., 2003), competence (Tortosa and Dubnau, 1999), DNA uptake (Brencic and Winans, 2005), sporulation (Piggot and Hilbert, 2004), and others. Importantly, the vast majority of these systems have been shown to directly regulate transcription via a response regulator possessing a DNA-binding domain. Thus, the TCST system controlling chemotaxis whereby CheY interacts directly with switch components (FliM)at the base of the flagellar motor is an exception to the rule (Fig. 1). The majority of all work on chemotaxis systems up until the last decade focused on the control of flagellum-based motility. Recently several organisms that do not exclusively utilize flagellar motility but are known to encode multiple homologs for the chemotaxis proteins have been the focus of much work. The chemotaxis genes in these organisms are usually organized into multiple, discrete operons randomly dispersed on one or more chromosomes. Investigation into the roles of the various chemotaxis gene clusters in these organisms has revealed that not all chemotaxis systems regulate flagellar motility. Indeed, some of these chemotaxis-like systems do not regulate motility at all and have been termed “chemosensory” systems. Three important examples include the Che3 system in M. xanthus (Kirbyand Zusman, 2003),the Che3 system in Rhodospirillum centenum (Berleman and Bauer, 2005), and the Che4 (Wsp) system in Pseudomonas aeruginosa (Hickman et al., 2005). In these cases, it is clear that a chemotaxis-like system comprising MCPs, Chew, CheA, CheR, and CheB has been co-opted to regulate other outputs. In the case of the M. xanthus Che3 system, the data indicate that CrdA, a homolog of NtrC, directly regulates
137
genes that are necessary for the control of development. Similarly, the R. centenum Che3 system affects the timing and level of cyst formation. The Che4 (Wsp) system in P. aeruginosa affects biofilm formation by directly regulating the concentration of cyclic diguanylate (c-di-GMP) via the response regulator, WspR, that possesses a GGDEF domain. The conclusion drawn from these examples is that chemosensory systems are specialized TCST systems with built-in adaptation modules by virtue of utilizing an MCP chemoreceptor to process the stimulus (Fig. 1).Because the MCP chemoreceptors are not covalently linked to the kinase domain (as is the case with prototypical TCST systems), multiple chemoreceptors can interact with a given CheA kinase, thereby effectively increasing the repertoire for ligand binding and stimulus processing. Integration of mltiple, varied inputs therefore occurs at the level of the CheA kinase. CheA kinases dictate the flow of information from the receptors to the outputs governed by a given system. Therefore, the existence of a gene encoding a CheA homolog defines a chemosensory system. Several hundred genomes are now fully sequenced and publicly available, and it is apparent that most motile organisms utilize multiple chemosensory systems, with the average being either two or three (I. B. Zhulin, personal communication). It is worth mentioning that nonmotile organisms typically lack chemotaxis-like genes altogether. M . xanthus is unique in that is has eight full chemosensory systems and is currently the only species known to possess such a large number of these specialized TCST systems. Because these systems are classified as homologs based on sequence similarity but appear to carry out alternate functions, they are by definition paralogs and not orthologs. M. xanthus is therefore an ideal organism for analysis of chemosensory signal transduction systems in bacteria. Several major biological questions arise from the above observations: (i) What is the role of each chemosensory system in M. xanthus? (ii)How do these systems maintain molecular insulation in order to prevent cross talk? (iii) Do these systems display cross-regulation? (iv) Is there a relationship between the diversity of chemosensory signal transduction systems and the complex lifestyle displayed by M. xanthus in particular or for bacteria in general? Discussion of the M. xanthus chemosensory systems and the role of multiple paralogous systems follows.
THE EIGHT CHEMOSENSORY SYSTEMS IN M. XANTHUS Analysis of the completed 9.14-Mb 111. xanthus genome indicates that of nearly 7,400 putative open reading frames (ORFs), 605 genes encode putative
REGULATORY MECHANISMS
138 signal transduction proteins. Three hundred twentytwo of these genes are predicted to encode homologs that contain either histidine kinase domains or aspartyl receiver domains that compose 125 TCST systems to process environmentally derived stimuli (Ulrich and Zhulin, 2007; see chapter 10). Within this group of TCST systems are the eight CheA homologs that define eight chemosensory systems. Importantly, each cheA gene is located with a cluster that also encodes homologs to known chemotaxis genes, as well as some nonchemotaxis genes (Fig. 2 ) . Based on the paradigm for E . coli chemotaxis, we can predict with a high degree of
confidence that the chemotaxis homologs will behave in specific ways within each chemosensory system. Nevertheless, recent examples (mentioned above) have demonstrated that many systems deviate from the E. coli paradigm and therefore warrant full experimental investigation.
The M. xanthus Frz (Chel) and Dif (Che2) Systems The M . xanthus Frz (Chel) and Dif (Che2) systems are described in detail in chapters 7 and 13, respectively, and are not discussed here.
che3 crdA
crdB crdC cheW3 mcp3A
mcp3B
cheA3
cheY4cheW4b
cheA4
cheB3
cheR3
che4 cheW4a cheR4
mcp4
che5 cheYS cheVS
che6
V
cheRS
chew5
V
cheW6a cheR6
cheAS
cheBS
cheA6
cheB6
Y
cheW6b
mcp6
mcp5
V
V
socD
V
kefC
che7
che8
ybaD
ribD
ribE
nbH nusB
cheY7
cheA7
cheY8a
cheA8
cheW7
cpc7
mcp 7
cheW8a cheW8b
cpc8
cheR7
cheR8
cheB7
cheB8
Figure 2 Genetic organization of the eight chemosensory systems in M. xanthus. Each cluster or operon is defined by the existence of a gene encoding a homolog to CheA (black). Genes encoding homologs to Chew (dotted), MCPs (white), CheB (squares), CheR (circles), CheC (gray), and CheY-like response regulators (shaded) are shown. ORFs that do not display homology to any known chemotaxis gene are also shown (diagonal stripes).
des 7
cheY8b
8. CHEMOSENSORY SIGNALTRANSDUCTION SYSTEMSIN 211. XANTHUS The M. xanthus Che3 Chemosensory System Previous work demonstrated that the M. xanthus Che3 system affects the timing of development (Kirby and Zusman, 2003). Mutations generated in several of the che3 genes led to premature development on starvation (complement fixation [CF]) media and also appeared to influence the vegetative component of the life cycle. For example, the mcp3B mutant displayed a rippling phenotype on rich (charcoal-yeast extract [CYE]) media and was observed to form aggregates, indicative of starvation, at the outer edge of the growing colony (Fig. 3). Likewise the cheA3 mutant was shown to be premature for development. Although no obvious motility defects were identified through direct observation of individual cells, elevated gene expression was observed for known developmental markers including spi, tps, and mbhA. Because premature developmental gene expression was shown to correlate with premature aggregation and fruiting body formation for these mutants, the Che3 system was predicted to regulate developmental gene expression. Analysis of the che3 gene cluster indicated that a divergently transcribed NtrC homolog (CrdA [chemosensory regulator of development]) was the likely regulator for this system. A mutation in the crdA gene led to delayed development consistent with the prediction that CrdA is a homolog of NtrC, a known transcriptional activator. Because the cheA3 and crdA mutants have opposite phenotypes with respect to the timing of development, epistasis analysis was possible. The cheA3crdA double mutant was generated, analyzed, and found to be delayed for development. Thus, crdA is epistatic
DZ2 (wild type)
139
to cheA3 and allows us to generate a model in which CheA3 processes information via CrdA as its primary output (Fig. 4). Because cheA3 and crdA have opposite phenotypes, it is likely that CheA3 acts as a phosphatase (or an inhibitor of CrdA phosphorylation) during vegetative growth. The CheR3 methyltransferase and CheB3 methylesterase were shown to be critical for control of development as well. Based on the paradigm chemotaxis system in E. coli and the premature developmental phenotype displayed by the cheA3 mutant, we predicted that the cheR3 and cheB3 mutant phenotypes would show premature and delayed phenotypes, respectively. The cheR3 mutant cells displayed premature aggregation, as expected, but the cheB3 mutant also displayed prematurc‘aggregation. However, the timing of maturation of the cheB3 fruiting bodies and sporulation was found to be delayed. Thus, the cheB3 mutant displays a complex phenotype suggesting that the role of methylation may not follow the E. coli paradigm and is likely to be more complex, like that observed for Bacillus subtilis (Kirby et al., 1999; see “Summary and Conclusions” below). Another factor that may relate to the complex phenotype displayed by the cheB3 mutant is the presence of two MCPs, Mcp3A and Mcp3B, within the Che3 system. All MCPs studied to date are thought to form homodimers. The homodimers are thought to be highly organized within functional arrays (Maddock et al., 1993) and possibly form trimers of dimers that affect signal integration and processing (Ames et al., 2002). Although direct evidence of such higher-order structure
mcp3B
Figure 3 The che3 mutants display premature development on rich medium. Both the parent (DZ2) and mcp3B mutant were grown at 32°C on rich medium (CYE) for 3 days and photographed (Kirby and Zusman, 2003); both colonies are approximately 2 cm in diameter. The mcp3B mutant displays a rippling phenotype characteristic of development. Additionally, aggregates that resemble fruiting bodies are visible at the colony edge under higher magnification.
REGULATORY MECHANI sM s
140
as part of the mechanism for regulating CheA3 signal transduction to CrdA. CrdB Is a n Essential Lipoprotein That Senses Envelope Stress
CrdA
r+* 054
cheBR3
7
Figure 4 Model for Che3 signal transduction. The current model for Che3 chemosensory signal transduction is shown. CrdB is an outer membrane lipoprotein with a Germinal OmpA-like peptidoglycan binding domain and is predicted to sense envelope stress. A conformational change in CrdB transmits a signal to the MCP receptor complex affecting CheA kinase levels in a manner similar to that observed during chemotaxis. CrdC is homologous to Chew and may affect coupling of the receptors to the CheA3 kinase. Results indicate that CrdA is autoregulatory and regulates expression of che3, cheBR3, and other genes, thereby affecting development as described previously (Kirby and Zusman, 2003). The cytoplasmic and outer membranes (black lines) and the peptidoglycan layer (crosshatching) demarcate the periplasm.
has only been shown for E. coli, there is experimental evidence indicating that MCPs are organized in arrays in Caulobacter crescentus (Alley et al., 1992), Rhodobacter sphaeroides (Wadhams et al., ZOOO), B. subtilis (Kirby et al., 2000), and others. Because the ligands for MCP receptors in species other than E. coli are largely unknown, the precise function of receptor arrays in other bacteria has not been elucidated. We predict that Mcp3A and Mcp3B will form homodimers and function within an array of chemoreceptors. We also predict that the homodimers will display some level of interaction
The first gene in the che3 cluster is crdB (Fig. 2). This gene is predicted to encode a lipoprotein that also binds to peptidoglycan via its carboxy-terminal OmpA-like domain. A crdB-phoA fusion was created and assayed for activity on medium containing 5-bromo-4-chloro3-indolylphosphate (BCIP). The phoA gene encodes an alkaline phosphatase which is only active when exported. Subsequently, PhoA converts BCIP to a blue product within the medium. The CrdB-PhoA fusion construct was active both in E. coli and in M . xanthus. This analysisdlows us to conclude that CrdB is exported to the periplasm. Further analysis of the N-terminal sequence of CrdB indicates that the protein contains a signal peptide sequence and a lipobox (Sierakowska et al., 2003). CrdB contains a threonine residue immediately following the +1 cysteine anchor which is known to direct lipoproteins to the inner leaflet of the outer membrane (Seydel et al., 1999). The processing of lipoproteins occurs via the Signal Peptidase I1 pathway (Sankaran and Wu, 1995) and is specifically inhibited by the antibiotic globomycin. Globomycin was able to prevent the processing of the CrdB proprotein to the mature form, indicating that CrdB is exported via Signal Peptidase I1 to the periplasm, where it functions as a lipoprotein in M. xanthus. An insertion mutation previously generated in crdB was found to affect expression of downstream che3 genes, and thus, a new construct was created. Based on several failed attempts to delete this gene, we now believe that crdB is essential. Previous reports have indicated that certain lipoproteins affect outer membrane stability and are therefore critical for cell viability (Cascales et al., 2002). Multiple attempts to generate new insertions in crdB also failed. However, one insertion mutant was finally generated (crdB2303)but was found to be a merodiploid that allows for low level of expression of crdB and the downstream che3 genes. The crdB1303 mutant is delayed for development in contrast to the other che3 mutants, consistent with an overall membrane defect. Further analysis of the crdB1303 mutant indicated that it is sensitive to EDTA or ampicillin, both of which have the potential to affect membrane integrity. Both EDTA and ampicillin were able to lyse the crdB2303 mutant cells when present in the medium at concentrations that had no effect on the wild type.
8. CHEMOSENSORY SIGNALTRANSDUCTION SYSTEMSI N M. XANTHUS Based on these data, we hypothesize that CrdB senses membrane integrity or periplasmic intermediates that reflect the presence of environmental stress to initiate the developmental program. Our current working model is that CrdB transduces signals through one of the two MCPs in the Che3 signaling complex to either inhibit or stimulate CheA3 activity (Fig. 4).
141
information via Frz to affect TFP-based motility. It is worth noting that the data for the che4 mutants were obtained for single cells and not for cells within a population. Therefore it is not clear how these results apply to individuals within larger groups. Assessment of green fluorescent protein-labeled cells within populations is possible and will allow for more thorough analysis of the Che4 system.
CrdC is a Homolog of Chew The gene encoding CrdC lies between the chew3 gene and the first of the two MCP homologs (Fig. 2). Analysis of the ORF for CrdC indicates that crdC and mcp3A are translationally coupled, suggesting that these two proteins interact within the signal transduction complex. Recent sequence analysis suggests that CrdC is likely to be a highly diverged homolog of Chew. Chew3 and CrdC have been shown to interact in the yeast twohybrid assay while only CheW3 is able to bind CheA3. Therefore, our current model places CrdC in a position to affect Chew3 within the signaling complex. It is also possible that CrdC may act as an inhibitor of Chew3 to affecting coupling between the MCPs and CheA3.
The M. xanthus Che4 Chemosensory System The genes in the che4 operon were shown to compose an operon and encode homologs to one MCP, two Chews, CheA, a CheY-like response regulator, and CheR, but no CheB. Analysis of the Che4 system led to the conclusion that the system affects type IV pilus (TFP)-based motility. The link between the Che4 system and regulation of the TFP motor is not known. Phenotypes for the che4 mutants were only discernible in the uglBl mutant background which lacks adventurous (A) motility and were only apparent under specific developmental conditions. The observations are consistent with a model whereby the Che4 system perceives physical stimuli such as surface hydration and transduces information to the pilus-based machinery. Analysis of the parent and che4 system mutants led to the conclusion that cellular velocity is inversely correlated with reversal frequency for wild-type cells. This inverse correlation was eliminated in mcp4 and cheY4 mutants. The structural components required for TFP were not disrupted by any of the mutations in the che4 operon. Together the results allow us to conclude that the Che4 system affects reversal frequency of cells by modulating the function of the TFP (Vlamakis et al., 2004). Consistent with this model is the observation that CheA4 was found to interact with the N terminus of FrzCD, the cytoplasmic MCP receptor for the Frz system (D. R. Zusman, unpublished data). Because the Frz system is known to regulate cellular reversal frequency, it is possible that Che4 communicates specific
The M. xanthus CheS Chemosensory System The genes in the che.5 cluster encode homologs to one MCP, two Chews (one is CheV-like), CheA, a CheY-like response regulator, CheR, and CheB. A thorough analysis of the Che.5 system has not yet been performed and thus is not discussed in detail here. We do not yet know if tke che5 genes shown in Fig. 2 compose an operon. However, we have constructed a mutation in the cheA.5 gene and have observed premature development in the cheA5 mutant, similar to what was observed for cheA3 and cheA6. Sequence analysis indicates that the Che.5 system possesses a unique chemotaxis-like protein that is a receiver domain-Chew fusion, which we refer to as “CheV-like.” CheV proteins are currently described as Chew-receiver domain fusions (Fredrick and Helmann, 1994). Thus, the CheV-like homolog in CheS possesses both domains but in the opposite orientation relative to CheV. The domain orientation should have little impact in the overall function of the protein, as it is predicted to function within the signaling complex as a Chew homolog by coupling Mcp.5 to the CheA.5 kinase. No further details are available at this time.
The M. xanthus Che6 Chemosensory System There are two primary reasons for our investigation into the role of the Che6 chemosensory system. First, the gene order is consistent with that found in other organisms known to utilize TFP-based motility (Zhulin, personal communication) and we therefore hypothesized that the Che6 system would regulate TFP-based motility. Second, a mutant allele known to suppress the csgA mutant developmental defect was mapped to SOCD(suppressor of csgA) (Rhie and Shimkets, 1991),which we now know is cotranscribed as part of the che6 operon. In order to verify that the nonchemotaxis genes, especially socD, were part of the che6 operon, we performed reverse transcriptase PCR. The results indicated that the genes shown compose an operon (Fig. 2). As with the other che operons, the che6 cluster contains a full complement of chemotaxis genes encoding homologs to CheA, two Chew proteins, CheR, CheB, and one MCP. These chemotaxis genes, as well as SOCDand kefC (encoding a potassium efflux pump), are cotranscribed
142 as part of the che6 operon. Analysis of the start and stop codons and putative ribosome binding sites for each of the ORFs within the che6 cluster leads us to predict that each pair of these genes within the che6 operon (except socD-KefC) is translationally coupled and therefore likely to function within the same pathway. As is the case with all known chemosensory systems studied thus far, the CheA kinase is the central processor through which MCP-generated signals are integrated and processed. Multiple MCP homologs can affect one CheA kinase, as is the case for E. coli, B. subtilis, and Halobacterium salinarum. Likewise, each CheA typically has multiple targets including CheY and CheB. Thus, CheA is the central processor for each system and mutations made within CheA block its function and usually give distinct phenotypes. The cheA6 mutant was created and displays an obvious motility defect on all agar surfaces. However, the mutant will eventually form very small flares on 0.3% agar after several days, indicating that TFP-based motility is dramatically reduced. This TFP motility defect led to the hypothesis that Che6 affects pilA gene expression or pilus production. We measured the level of PilA monomer and production of surface pili by immunoblot analysis using anti-PilA antibody (Wu and Kaiser, 1997; Wall et al., 1998). Both the DZ2 parent and cheA6 mutant produced equal amounts of PilA monomer indicating that gene expression from the pi1 cluster is not affected in the cheA6 mutant. However, the cheA6 mutant cells were found to lack PilA normally detectable in cell surface preparations of TFP. Recent results from transmission electron microscopy confirm these results. In contrast to the wild type, the cheA6 mutant cells do not produce detectable pili under the conditions tested. Because the PilA monomer is produced at or near wild-type levels, we conclude that CheA6 is required for TFP assembly. Similar results were obtained for the mcp6 mutant, indicating that Mcp6 processes information via CheA6. The mechanism by which Che6 affects pilus assembly remains to be determined. The roles for SocD and KefC also remain unknown.
The M. xanthus Che7 Chemosensory System Similar to the case described above, the M . xanthus che7 gene cluster contains a full set of chemotaxis genes and additional ORFs that show no homology to chemotaxis proteins. In order to verify that these genes compose an operon, we performed reverse transcriptase-PCR and were able to show that the che7 genes are cotranscribed (Fig. 2). The che7 operon encodes homologs to CheA, Chew, CheY, CheR, CheB, and one cytoplasmic MCP. In
REGULATORY MECHANISMS addition, genes encoding a phycocyanobilin lyase (cpc) and a fatty acid desaturase (des)are expressed as part of the che7 operon. Analysis of the che7 ORFs allows us to predict that Mcp7 and Cpc7 are translationally coupled and are therefore thought to interact within the Che7 system. It has been demonstrated that Cpc phycocyanobilin lyase proteins carry out specific covalent modifications whereby phycobilins are attached to apophycobiliprotein subunits ( Fairchild et al., 1992). The phycobiliprotein subunits are part of the macromolecular phycobilisome light-harvesting complex. Because M . xanthus does not possess phycobilisomes, Cpc7 is not likely to function as a phycocyanobilin lyase per se but might function in another light-dependent adaptation process. Previous work has demonstrated that the 211. xanthus car locus regulates carotenoid production in a light-dependent fashion such that colonies become bright orange when exposed to light (Fontes et al., 1993). We therefore tested the che7 mutants for their ability to produce carotenoids with peak absorbance at 475 nm in response to light. The mcp7, cheB7, and cpc7 mutants each displayed enhanced production of carotenoids (475 nm) relative to the parent, while the cheA7 mutant was completely incapable of producing this carotenoid. The mechanism by which the Che7 system affects carotenoid production is unknown. However, it is worth noting that the cheA7 mutant displays a severe swarming defect when assayed on 0.3% agar surfaces, does not aggregate (Fig. 5 ) ,and does not produce viable spores under the conditions tested. The mcp7 and cpc7 mutants also display reduced motility, no aggregation, and no viable spore production. Together, these observations lead us to speculate that the Che7 system affects membrane composition.
The M. xanthus Che8 Chemosensory System The genes in the che8 cluster encode homologs to CheA, two Chews, two CheY-like receiver domain proteins, CheR, and CheB, but no MCP. There are 1 3 orphan MCP homologs encoded by the M. xanthus genome, and it is likely that one or more of these MCPs will transduce signals via the Che8 complex. Interestingly, the che8 cluster also encodes a homolog of the cpc phycocyanobilin lyase gene similar to that seen in the che7 operon. Even more striking is the presence of several genes that constitute a major portion of the riboflavin biosynthetic operon. Whether these genes are cotranscribed with the che homologs and compose an operon has not been clearly established, although preliminary evidence suggests that they are cotranscribed as depicted (Fig. 2).
8. CHEMOSENSORY SIGNALTRANSDUCTION SYSTEMSIN M. XANTHUS
143
Figure 5 The cheA7 mutant displays defective motility and aggregation. The DZFl parent and cheA7 mutant are shown. Cells were grown in rich medium (CYE), washed in buffer (MMC), and plated on either 0.3% agar containing CYE or on 1.5% agar with low nutrients (CF).DZFl cells swarm on rich medium (A) and develop on low-nutrient medium to produce fruiting bodies (B). The cheA7 mutant cells display major defects in swarming (C) and development (D) relative to the parent.
SUMMARY AND CONCLUSIONS The presence of multiple paralogous chemosensory systems in M. xanthus presents a unique opportunity to explore variations on a common theme for signal transduction, network integration, and genome evolution in a model organism for prokaryotic development. Analysis of each chemosensory system thus far indicates that each system plays a unique role during the vegetative and/or developmental aspects of the complex multicellular life cycle of M. xanthus (Table 1).Analysis of the timing and level of expression of each operon is currently under way. However, it is worth noting that mutations in all eight che systems display phenotypes during vegetative growth as well as during development, indicating that expression of all eight che systems is required for both vegetative and developmental aspects of the M. xanthus life cycle. The Frz chemosensory system appears to directly affect motility and most likely does so by regulating cell reversals via FrzS (Mignot et al., 2005; see chapter 7). Each
of the other chemosensory systems appears to have been co-opted to regulate functions other than chemotaxis per se, although the Dif system indirectly affects motility by regulating EPS production (Yang et al., 2000; Xu et al., 2005). Additionally, it is likely that Che4 affects cell reversal frequency but may do so indirectly by interacting with Frz. Furthermore, Che6 affects TFP assembly by an unknown mechanism. Thus, motility is affected by four of the eight chemosensory systems either directly or indirectly. Analysis of the other four chemosensory systems suggests that these specialized TCST systems have different outputs ranging from gene expression to biosynthesis of riboflavin (Table 1). One of the major conclusions from our analysis of the M. xanthus chemosensory systems is that the physiological role of each chemosensory system is specified by its nonchemotaxis genes. The genes in question encode proteins that may provide alternative inputs, alternative outputs, or regulators that impinge on the basic design of the chemosensory system. In several cases, more than
REGULATORY MECHANISMS
144 Table 1 Chemosensory systems in M . xanthus Che system
Proposed function
Reference(s)
Frz (Chel) Dif (Che2) Che3
Gliding reversal frequency EPS production and PE detection Developmental gene expression
Che4 CheS Che6
TFP-based motility TFP assembly
Che7
Carotenoid production
Che8
Riboflavin biosynthesis
Zusman lab; chapter 7 Yang and Shimkets; chapter 13 Kirby and Zusman, 2003; S. Mueller and J. R. Kirby, unpublished data; D. Li and J. R. Kirby, unpublished data Vlamakis et al., 2004 Kirby lab, unpublished data J. C. Scott and J. R. Kirby, unpublished data J. M. Wilson and J. R. Kirby, unpublished data Leclerc and Kirby, unpublished data
?
one permutation on the basic theme is evident (Fig. 4). There is at least one gene that is not homologous to any known chemotaxis gene in each operon (Fig. 2). The relationship between those gene products and the associated chemosensory system is the primary focus of our ongoing investigation. Another major conclusion that results from our analysis is that many of the chemosensory systems maintain molecular insulation (each system gives phenotypes specific to its system). However, it is possible and even probable that cross-regulation will emerge as part of the network design (e.g., Che4 and Frz). The level and specificity of integration remains largely unexplored. Several factors are likely to directly influence cross-regulation including the similarity and nature of specific functions such as receptor clustering and methylation systems. As mentioned above, C. CYescentus, E. coli, and R. sphaeroides MCP receptors are capable of forming large clusters either at the cell poles or in the cytoplasm. The topology of the receptor dictates localization, and the clustering is thought to affect the overall function of a given MCP within the receptor array (Bray et al., 1998). It is therefore likely that several of the M. xanthus MCP homologs will form an array and affect the mechanism of signal transduction for a given receptor. There are 13 orphan mcp chemoreceptor genes on the M. xanthus genome. The orphan MCPs are likely to influence the composition of a putative receptor array. Moreover, these MCPs are predicted to interact with at least one Chew homolog encoded within one of the eight che clusters. The sequence of each MCP and its gene neighborhood have not yet been correlated with the chemosensory systems, stimuli, or outputs. One important example of cross-regulation in chemotaxis has been demonstrated for the methylation system
in E. coli. In E. coli, the CheR methyltransferase and the CheB methylesterase have both been shown to interact with the carboxy-terminal pentapeptide NWETF motif on either Tar or Tsr, the serine receptor (Feng et al., 1999; Li and Hazelbauer, 2005). However, neither Tap (dipeptide receptor) nor Trg (ribose and galactose binding protein receptor) possesses the CheRB pentapeptide NWETF docking sequence. Nevertheless, both Tap and Trg require methylation and demethylation for proper functioning. The evidence indicates that Tap and Trg are methylated and demethylated much more efficiently if Tar and Tsr are present. The conclusion from these data is that CheR and CheB target the docking sites on Tar and Tsr and carry out modification of the neighboring Trg and Tap receptors. Thus, the receptor array allows for cross-regulation by CheR and CheB. Although there is no direct evidence suggesting this is the case in M. xunthus, it is likely that CheR and CheB from any given chemosensory system would be capable of modifying neighboring receptors if those receptors are found to lie within a clustered receptor array. Several observations support the notion that M. xunthus receptors exist within a signaling array and that methylation and demethylation reactions occur via one of many CheR and CheB homologs. First, the cheB3 mutant displayed a complex phenotype (premature aggregation and delayed sporulation) relative to the mcp3A and mcp3B mutants (see above). Furthermore, normal DifA function appears to require methylation by CheR and CheB homologs (Bonner et al., 2005) yet no genes for cheR or cheB are present in the difcluster. Finally, the Che4 system requires the CheR4 methyltransferase for proper functioning but lacks a cheB methylesterase gene (Vlamakis et al., 2004). The most likely scenario is that a CheB homolog from another
8. CHEMOSENSORY SIGNALTRANSDUCTION SYSTEMSIN M. XANTHUS system functions to modulate the overall methylation level of Mcp4. It is worth noting that even though the Dif system lacks CheR and CheB homologs, it does possess a CheC homolog likely to act as a phosphatase (Black and Yang, 2004; Szurmant et al., 2004). Therefore, the Dif chemosensory signal transduction system is regulated in a unique way relative to the other seven Che systems in M. xanthus. Another important point is that there is an orphan cheR-cheB gene cluster with associated ORFs that do not encode chemotaxis-like proteins. The role of this cluster is not yet known. In summary, the chemosensory systems found in M. xanthus appear to have evolved in order to regulate functions that need temporal control mechanisms. Temporal control is the hallmark feature of chemotaxis-like systems where methylation and demethylation of the input MCP chemoreceptor affects the rate of phosphotransfer from the CheA kinase to a response regulator. The role of methylation and demethylation for each of the eight M . xanthus chemosensory systems remains to be fully analyzed. The kinetics of modification have not been assessed for any specific residue in any MCP homolog in M. xanthus, although the overall rate of methylation changes for FrzCD was shown to occur over a 30-min to 2-h window (McBride and Zusman, 1993) consistent with the time domain in which M. xanthus makes decisions regarding cell reversals and commitment to sporulation. Lastly, it has been demonstrated that there is a relationship between the number of signal transduction systems, genome size, and the complexity of the environmental niche and corresponding lifestyle for a given microbe (Van Nimwegen, 2003; Konstantinidis and Tiedje, 2004; Ulrich et al., 2005). The M . xanthus genome does not represent a major deviation from this relationship with respect to the total number of signal transduction proteins. However, M. xanthus does possess a relatively large number of TCST systems (see chapter 10) and is unparalleled with respect to its eight chemosensory systems. It therefore appears that the eight chemosensory systems provide a particular advantage for M. xanthus. The most logical conclusion derived from these observations is that the chemosensory systems are being utilized for temporal regulation of a variety of functions necessary for survival by M. xanthus. Those systems listed in Table 1 are therefore predicted to be crucial for survival and temporally regulated during all aspects of the M. xanthus life cycle. It is well known that a variety of organisms use multiple paralogous chemosensory systems including P. aeruginosa, R. sphaeroides, R. centenum, and M . xanthus. Evolution of paralogous chemosensory systems
145
is strictly correlated with motility. For each organism with multiple chemosensory systems, at least one system has been shown to regulate motility. Whether or not additional chemosensory systems affect motility varies from one organism to another. For example, it appears that all chemosensory systems in R. sphaeroides affect flagellum-based motility (Armitage, 2003) while those in P. aeruginosa and R. centenum regulate distinct functions. In that regard M. xanthus may represent a case study encompassing both extremes where several systems regulate motility and several systems regulate alternative functions. Analysis of chemosensory signal transduction systems in each model organism is an excellent example of the modular nature of signal transduction and the evolution of bacterial genomes. We thank Aaron Buldoc, Carolin Groeger, Marion Leclerc, and Hsu-Ming Wen for work on the various Che systems. We also thank Larry Shimkets, Zhaomin Yang, Heidi Kaplan, Mitch Singer, David Zusman, Trish Hartzell, and Dale Kaiser for strains, plasmids, and antibodies. Support for this work was provided by Grant A1059682 from the National Institutes of Health to1.K.
References Alley, M. R., J. R. Maddock, and L. Shapiro. 1992. Polar localization of a bacterial chenioreceptor. Genes Dev. 6532.5836. Ames, P., C. A. Studdert, R. H. Reiser, and J. S. Parkinson. 2002. Collaborative signaling by mixed chemoreceptor teams in Escherichia coli. Proc. Natl. Acad. Sci. USA 99: 7060-7065. Armitage, J. P. 2003. Taxing questions in development. Trends Microbiol. 11:239-242. Bassler, B. L. 2002. Small talk. Cell-to-cell communication in bacteria. Cell 109:421-442. Berleman, J. E., and C. E. Bauer. 2005. Involvement of a Che-like signal transduction cascade in regulating cyst cell development in Rhodospirillum centenum. Mol. Microbiol. 56A4.57-1466. Black, W. P., and Z. Yang. 2004. Myxococcus xanthus chemotaxis homologs DifD and DifG negatively regulate fibril polysaccharide production. /. Bacteriol. 186: 1001-1008. Bonner, P. J., Q. Xu, W. P. Black, Z. Li, Z. Yang, and L. J. Shimkets. 2005. The Dif chemosensory pathway is directly involved in phosphatidylethanolamine sensory transduction in Myxococcus xanthus. Mol. Microbiol. 57: 1499-1508. Bourret, R. B., and A. M. Stock. 2002. Molecular information processing: lessons from bacterial chemotaxis. J. Biol. Chem. 27E9625-9628. Bray, D., M. D. Levin, and C. J. Morton-Firth. 1998. Receptor clustering as a cellular mechanism to control sensitivity. Nature 393:85-88.
146 Brencic, A., and S. C. Winans. 2005. Detection of and response to signals involved in host-microbe interactions by plantassociated bacteria. Microbiol. Mol. Biol. Rev. 69:155-194. Cascales, E., A. Bernadac, M. Gavioli, J.-C. Lazzaroni, and R. Lloubes. 2002. Pal lipoprotein of Escherichia coli plays a major role in outer membrane integrity. 1. Bacteriol. 184~754-759. DiGiuseppe, P. A., and T. J. Silhavy. 2003. Signal detection and target gene induction by the CpxRA two-component system. J. Bacteriol. 185:2432-2440. Fairchild, C. D., J. Zhao, J. Zhou, S. E. Colson, D. A. Bryant, and A. N. Glazer. 1992. Phycocyanin alpha-subunit phycocyanobilin lyase. Proc. Natl. Acad. Sci. USA 89:7017-7021. Falke, J. J., and G. L. Hazelbauer. 2001. Transmembrane signaling in bacterial chemoreceptors. Trends Biochem. Sci. 26~257-265. Falke, J. J., R. B. Bass, S. L. Butler, S. A. Chervitz, and M. A. Danielson. 1997. The two-component signaling pathway of bacterial chemotaxis: a molecular view of signal transduction by receptors, kinases, and adaptation enzymes. Annu. Rev. Cell Dev. Biol. 13:457-512. Feng, X., A. A. Lilly, and G. L. Hazelbauer. 1999. Enhanced function conferred on low-abundance chemoreceptor Trg by a methyltransferase-docking site.]. Bacteriol. 181:31643171. Fontes, M., R. Ruiz-Vazquez, and F. J. Murillo. 1993. Growth phase dependence of the activation of a bacterial gene for carotenoids synthesis by blue light. EMBO J. 12:12651275. Fredrick, K. L., and J. D. Helmann. 1994. Dual chemotaxis signaling pathways in Bacillus subtilis: a sigma D-dependent gene encodes a novel protein with both Chew and CheY homologous domains.]. Bacteriol. 176:2727-2735. Hickman, J. W., D. F. Tifrea, and C. S. Harwood. 2005. A chemosensory system that regulates biofilm formation through modulation of cyclic diguanylate levels. Proc. Natl. Acad. Sci. USA 102:14422-14427. Hoch, J. A. 2000. Two-component and phosphorelay signal transduction. Curr. Opin. Microbiol. 3:165-170. Hoch, J. A., and T. J. Silhavy (ed.). 1995. Two-Component Signal Transduction. ASM Press, Washington, DC. Kirby, J. R., M. M. Saulmon, C. J. Kristich, and G. W. Ordal. 1999. CheY-dependent methylation of the asparagine receptor, McpB, during chemotaxis in Bacillus subtilis. 1. Biol. Chem. 274:11092-11100. Kirby, J. R., T. B. Niewold, S. Maloy, and G. W. Ordal. 2000. CheB is required for behavioural responses to negative stimuli during chemotaxis in Bacillus subtilis. Mol. Microbiol. 35:44-57. Kirby, J. R., and D. R. Zusman. 2003. Chemosensory regulation of developmental gene expression in Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 100:2008-2013. Konstantinidis, K. T., and J. M. Tiedje. 2004. Trends between gene content and genome size in prokaryotic species with larger genomes. Proc. Natl. Acad. Sci. U S A 101:31603165. Krukonis, E. S., and V. J. DiRita. 2003. From motility to virulence: sensing and responding to environmental signals in Vibrio cholerae. Curr. Opin. Microbiol. 6:186-190.
REGULATORY MECHANISMS Li, M., and G. L. Hazelbauer. 2005. Adaptational assistance in clusters of bacterial chemoreceptors. Mol. Microbiol. 56:1617-1626. Maddock, J. R., M. R. Alley, and L. Shapiro. 1993. Polarized cells, polar actions. J. Bacteriol. 175:7125-7129. McBride, M. J., and D. R. Zusman. 1993. FrzCD, a methylaccepting taxis protein from Myxococcus xanthus, shows modulated methylation during fruiting body formation. J. Bacteriol. 175:4936-4940. Mignot, T., J. P. Merlie, Jr., and D. R. Zusman. 2005. Regulated pole-to-pole oscillations of a bacterial gliding motility protein. Science 310:855-857. Parkinson, J. S., and E. C. Kofoid. 1992. Communication modules in bacterial signaling proteins. Annu. Rev. Genet. 26~71-112. Parkinson, J. S. 1993. Signal transduction schemes of bacteria. Cell 73:857-871. Piggw P. J., and D. W. Hilbert. 2004. Sporulation of Bacillus subtilis. Curr. Opin. Microbiol. 7579-586. Qin, L., S. Cai, Y. Zhu, and M. Inouye. 2003. Cysteinescanning analysis of the dimerization domain of EnvZ, an osmosensing histidine kinase. J. Bacteriol. 185:34293435. Rhie, H., and L. J. Shimkets. 1991. Low-temperature induction of Myxococcus xanthus developmental gene expression in wild-type and csgA suppressor cells. J . Bacteriol. 173:2206-2211. Sankaran, K., and H. C. Wu. 1995. Bacterial prolipoprotein signal peptidase. Methods Enzymol. 248:169-180. Seydel, A., P. Gounon, and A. P. Pugsley. 1999. Testing the “ + 2 rule” for lipoprotein sorting in the Escherichia coli cell envelope with a new genetic selection. Mol. Microbiol. 34:810-821. Sierakowska, A., H. Willenbrock, G. von Heijne, H. Nielsen, S. Brunak, and A. Krogh. 2003. Prediction of lipoprotein signal peptides in Gram-negative bacteria. Protein Sci. 12:1652-1662. Stock, A. M., V. L. Robinson, and P. N. Goudreau. 2000. Two-component signal transduction. Annu. Rev. Biochem. 69: 183-2 15. Szurmant, H., T. J. Muff, and G. W. Ordal. 2004. Bacillus subtilis CheC and FliY are members of a novel class of CheY-Phydrolyzing proteins in the chemotactic signal transduction cascade.]. Biol. Chem. 279:21787-21792. Tortosa, P., and D. Dubnau. 1999. Competence for transformation: a matter of taste. Curr. Opin. Microbiol. 2588-592. Ulrich, L., E. V. Koonin, and I. B. Zhulin. 2005. Onecomponent systems dominate signal transduction in prokaryotes. Trends Microbiol. 1352-56. Ulrich, L. E., and I. B. Zhulin. 2007. MIST: a microbial signal transduction database. Nucleic Acids Res. 35:D386-D390. Van Nimwegen, E. 2003. Scaling laws in the functional content of genomes. Trends Genet. 19:479-484. Vlamakis, H. C., J. R. Kirby, and D. R. Zusman. 2004. The Che4 pathway of Myxococcus xanthus regulates type IV pilus-mediated motility. Mol. Microbiol. 52:17991811. Wadhams, G. H., A. C. Martin, and J. P. Armitage. 2000. Identification and localization of a methyl-accepting
8. CHEMOSENSORY SIGNALTRANSDUCTION SYSTEMS IN M. XANTHUS chemotaxis protein in Rhodobacter sphaeroides. Mol. Micro biol. 36:1222-1233. Wall, D., S. S. Wu, and D. Kaiser. 1998. Contact stimulation of Tgl and Type IV pili in Myxococcus xanthus.]. Bacteriol. 180:759-76 1. Wu, S. S., and D. Kaiser. 1997. Regulation of expression of the PilA gene in Myxococcus xanthus. J. Bacteriol. 179:77487758.
147
Xu, Q., W. P. Black, S. M. Ward, and Z. Yang. 2005. Nitratedependent activation of the Dif signaling pathway of Myxococcus xanthus mediated by a NarX-DifA interspecies chimera. J. Bacteriol. 187:6410-6418. Yang, Z., X. Ma, L. Tong, H. B. Kaplan, L. J. Shimkets, and W. Shi. 2000. Myxococcus xanthus dif genes are required for biogenesis of cell surface fibrils essential for social gliding motility. J. Bacteriol. 1825793-579 8.
Myxobacteria: Multicellularity and Differentiation Edited by David E. Whitworth 0 2008 ASM Press, Washington, D.C.
Lee ICroos Sumiko Inouye
Transcriptional Regulatory Mechanisms during Myxococcus xanthus Development'
Since the writing of Myxobacteria I1 there has been tremendous progress in understanding transcriptional regulation of gene expression in Myxococcus xanthus. Most of the progress has involved studies of genes induced during development or during light-induced carotenoid biosynthesis. The latter is described in chapter 12. In this chapter, we focus primarily on transcriptional regulation of developmental genes. While the identification of developmentally regulated M. xanthus genes continues, now on a comprehensive genome-wide scale with the use of DNA microarray expression profiling (see chapter 28), an understanding of the cis-acting DNA elements (promoters and transcription factor binding sites) and trans-acting proteins (RNApolymerase [RNAP] with particular sigma factors, activators, and repressors) has emerged for a handful of developmental genes. Much more work will be necessary in order to achieve an understanding of the transcriptional regulatory network comparable to that learned from studies of Bacillus subtilis sporulation (see chapter 21) and the Caulobacter cell cycle (see chapter 22). The task seems daunting because analysis of the M. xanthus genome sequence reveals copious potential for transcriptional regulation (see chapter 16).On the other hand, the
9
potential for discovery of novel regulatory mechanisms and strategies makes the task irresistible, not only to satisfy curiosity but also to provide paradigms for less tractable organisms and communities (i.e., biofilms) of practical importance.
SIGMA FACTORS-d4
AND THE u7OFAMILY
In prokaryotes, sigma factors of RNAP play a key role in the regulation of gene expression by recognizing specific promoters and initiating transcription. There are two structurally unrelated families of sigma factors, the as4 and a 7 0 families (Helmann and Chamberlin, 1988). As in many bacteria, M. xanthus has a single as4(Goldman et al., 2006), but unlike in other bacteria, as4is ' essential for M . xanthus growth (Keseler and Kaiser, 1997). This has made it difficult to determine the role of d4 in development. Nevertheless, a number of genes, some crucial for development, have been predicted to be transcribed by aS4-RNAP(Table 1).In one case (spi), mutational analysis of the promoter supports the prediction (Keseler and Kaiser, 1995). In the other cases, the prediction is supported by the sequence of the promoter and/or genetic dependence of expression on a predicted
Lee Kroos, Department of Biochemistry and Molecular Biology, Michigan State University, East Lansing, MI 48824. Sumiko Inouye, Department of Biochemistry, Robert Wood Johnson Medical School, Piscataway, NJ 08854.
149
REGULATORY MECHANISMS
150 Table 1 Putative d4-RNAP-transcribed genes Gene or operon actAB CDE orf2 asgE sdeK spi mbhA pilA
mrpc
Promoter sequence"
Induction time (h)b
-29 T a C A C AACCA T m T -24 TCGCGGAGTCCGGEA -28 C E T G C A C A A G G G E T -26 C E C G C T C T C A G C E G -28 GAGCACGCGTCTmT -27 T E C A C G C CA T C m T -29 T E C A T G C G T A G m T -29 T a C A C G A A C C T E G A
2-4 2-4 0-2 2 10 veg 0-2
Inferred EBP
SasR PilR MrpB
Reference(s) Gronewold and Kaiser, 2001 Garza et al., 2000 Garza et al., 2000 Garza et al., 1998 Keseler and Kaiser, 1995; Guo et al., 2000 Romeo and Zusman, 1991 Wu and Kaiser, 1997 Sun and Shi, 2001a; Nariya and Inouye, 2005
OEach putative promoter sequence is shown with the number to the left indicating the position relative to the predicted start site of transcription. Bold nucleotides match consensus sequence, TGGYRYRNNNNTTSA, and underlined nucleotides match those most highly conserved in the consensus (Thony and Hennecke, the E. coli us4 1989). bThe approximate time during development when expression begins to increase is listed. This has not been determined for the actABCDE operon. The pilA gene is expressed during vegetative growth (Veg), and its expression does not increase during development.
enhancer-binding protein (EBP) (see below), though in no case has transcription been reconstituted in vitro. d4RNAP appears to play a critical role primarily early in development, though it is also believed to transcribe at least one gene ( m b h A )later in development (Table 1). The a 7 0 family is predicted to include 47 members in M. xanthus, an unusually large number even considering the large genome size (Goldman et al., 2006). This includes the primary sigma factor, SigA, presumed to be responsible for the majority of transcription in growing cells; five smaller sigma factors, SigB through SigF (SigB-F), closely related in sequence to each other and to SigA (Fig. 1);SigG, which is less similar to SigA-F but is similar to Escherichia coli FIiA; 38 sigma factors that appear to belong to the extracytoplasmic function (ECF) subfamily; and 2 sigma factors with limited similarity to known sigma factors.
SigA-G SigA shows high sequence similarity to E. coli u70,except that SigA has an extra -100 residues at its amino-terminal end (Inouye, 1990). SigA was originally described as 01 (Rudd and Zusman, 1982) and later shown to be the most abundant sigma factor in RNAP holoenzyme purified from growing M. xanthus cells (Biran and Kroos, 1997). Biran and Kroos (Biran and Kroos, 1997) further demonstrated that reconstituted oA-RNAP could initiate transcription accurately in vitro from the vegA promoter, which is active in growing M. xanthus (Komano et al., 1987). Two other promoters active in growing cells, ugh11 (Biran and Kroos, 1997) and ZonD (Ueki and Inouye, 2002), also appear to be utilized by oARNAP in vitro. The lonD gene is also called bsgA, and this heat shock inducible gene (Ueki and Inouye, 2002) encodes an ATP-dependent protease that is essential for
development (Gill et al., 1993; Tojo et al., 1993). The only developmentally regulated promoter that has been shown to be recognized by a*-RNAP in vitro is the promoter of an operon identified by Tn5 lac insertion R4.514 (Ha0 et al., 2002). Expression of this operon is induced at about 9 h into development (Kroos et al., 1986) and appears to involve both negative autoregulation and positive control (Ha0 et al., 2002). Another indication that SigA is active during development comes from the finding that a missense mutation in the sigA gene causes a defect in A-signal production (Davis et al., 1995). SigA is highly similar to E. coli 0 7 0 and B. subtilis 043in the regions expected to recognize -10 and -35 promoter sequences (Inouye, 1990), and promoters recognized by M. xanthus oA-RNAPin vitro match the 070/u43 consensus quite well in their -35 regions (Table 2). However, their -10 regions match the consensus poorly and are more GC rich, leading to speculation that 111. xanthus a*-RNAP better tolerates GC-rich - 10 sequences (Biran and Kroos, 1997; Hao et al., 2002). By using a synthetic oligonucleotide corresponding to the highly conserved region 2.2 of SigA as a probe for Southern blot analysis with chromosomal DNA digests, genes sigB-E were identified and subsequently isolated and characterized (Apelian and Inouye, 1990,1993; Ueki and Inouye, 1998, 2001). Of these, SigD is most similar to SigA (Fig. 1).SigD shares 35 and 34% identity with M. xanthus SigA and E. coli RpoS, respectively (Ueki and Inouye, 1998). SigD lacks region 1.1,which is found only in primary sigma factors and RpoS homologues (Gruber and Bryant, 1997), and most of region 3.1, which is typically conserved in all sigma factors except those in the ECF subfamily (Lonetto et al., 1994). By analyzing protein expression patterns and cell viability of a sigD deletion mutant during the late log and stationary phases, as
9. 211. XANTHUS TRANSCRIPTIONAL REGULATORY MECHANISMS SigA S igD SigE SigC S igB SiqF
151
................ MEAINLNVSFESPELWP
18
RpoH box SigA SigD SigE SigC SigB SigF SigG
IETINIQIRTSRYLVQE ... I ~ R E P .T. . .P E E I ~ E K M E L P L D ~ K V .
DSHLG.. ETTFL . .
.................................................
:."QAHG
DSRTTRP DATHL . . GNSHV.. EATRL . .
RTRRE E G D A "... A EPE IK~ R L L KASE ~ R ~EME T E Q ~ TRRE EKF SGDA.AVNVDDI~R KPGE EME . . . . .WVL_Q- RRERS EARWGEGHPEVE.KRL,E GKREDE LAM
s
. . . . . . . . . . . .N A W G ~ ~ Y L G N S L D R E A G A ~ N R ~ S S F D D D ~ D I S D A V T G L I E G T ~ D T A G Y T
. . . . . . . .D E S L P ~ I R M E Q
630 167 202 216 216 185 161
(core binding) Sign SigD SigE SigC SigB SigF SigG
PSRSKRLRSFVES !GVSGHPGPFP LINRMRDFMREQIPDFDLVASPKA LMAEAGVDESTLNA LMAEVDPEAVAAOO
------4
708 246 280 295 295 264 239
I------(-35 recognition)
Figure 1 Alignment of parts of the amino acid sequences of M. xanthus SigA-G. Amino acids identical in more than 50% of the sequences are indicated by a black background. Conserved subregions of sigma factors and their functions are denoted under the sequences (Helmann and Chamberlin, 1988; Lonetto et al., 1992). well as its response to various stresses (osmotic, oxidative, heat, and cold), it was concluded that SigD shows characteristic features of stationary-phase sigma factors (Ueki and Inouye, 1998). The deletion of sigD also affects protein synthesis patterns during early development, resulting in a 24-h delay in the initiation of fruiting body formation and a reduced spore yield (26% of the parent strain). Therefore, SigD is a stationary-phase sigma factor that is also required for multicellular development. The sigD mutant responds to starvation by inducing (p)ppGppsynthesis normally but is impaired for production of A- and C-signals (Viswanathan et al., 2006b). SigD is needed for cellular responses to A-signal, but the sigD mutant can respond to C-signal from codeveloping wild-type cells by inducing a subset of late developmental genes. Other late genes require oD-RNAPor a gene under its control cell autonomously. M. xanthus appears to have expanded
the repertoire of its stationary-phase sigma factor to include a role in the decision whether to initiate fruiting body formation in response to starvation. As shown in Fig. 1, SigB, SigC, and SigE exhibit high similarity to M. xanthus SigA and SigD and have Table 2 oA-RNAP-transcribedgenes Gene or operon
-35 region"
a4514 vegA
TTGACA TAGACA TTGCCA TTGCCA
aphII lonD "old
Spacer (bp)b
-10 regionc
18
TACCTA TAAGGG TAAGGT TACGTT
17 17 16
nucleotides match the E. coli c7'O-B. strbtilis u+3consensus sequence of
TTGACA. 'Distance between the -35 and -10 regions.
TATAAT.
REGULATORY MECHANISMS
152 high similarity to a consensus sequence, Q(R/K)(R/K) LFFNLR, called the RpoH box, found in heat shock sigma factors (Nakahigashi et al., 1995).The RpoH box is known to be unique in heat shock sigma factors and to be located between regions 2 and 3. Although SigB, SigC, and SigE have the RpoH box with four, two, and one amino acid substitutions, respectively, they are not heat shock inducible or involved in the production of heat shock proteins (Apelian and Inouye, 1990; Apelian and Inouye, 1993; Ueki and Inouye, 2001). Analysis of the M. xanthus genome sequence failed to identify other sigma factors homologous to heat shock sigma factors, suggesting that M. xanthus utilizes a different mechanism to respond to heat shock. Indeed, a two-component signal transduction system, HsfNHsfB, has been shown to activate transcription of lonD (Ueki and Inouye, 2002). Further characterization of heat shock proteins identified by two-dimensional-gel analysis (Otani et al., 2001, 2005) should provide insights into molecular mechanisms of the M. xanthus heat shock response. Though not apparently involved in the heat shock response, SigB, SigC, and SigE are involved in M . xanthus development. Expression of sigB starts at the onset of sporulation, inside spores only, and SigB is essential for maturation of spores (Apelian and Inouye, 1990). In a sigB deletion mutant, expression of the spore-specificops gene (Teintze et al., 1985) is not observed (Apelian and Inouye, 1990). Expression of sigC begins early in development, and deletion of sigC results in earlier formation of fruiting bodies (Apelian and Inouye, 1993). Furthermore, the sigC deletion mutant, unlike the wild type, can form fruiting bodies and spores on semirich agar plates, suggesting that SigC may play a role in the expression of genes that negatively regulate the initiation of fruiting body development. Likewise, SigE might negatively regulate developmental genes, because deletion of sigE results in earlier fruiting body formation (Ueki and Inouye, 2001). However, unlike sigC, sigE is expressed during growth, and expression increases during stationary phase and at the onset of development. The different expression patterns of the two genes suggest that SigC and SigE differentially regulate developmental gene expression. Interestingly, the sigE deletion mutant exhibits defects in fruiting body formation at 37"C, a higher temperature than normal (30 to 32°C). A triple deletion mutant, AsigB AsigC AsigE, constructed by using positive-negative KG cassettes (Ueki et al., 1996), forms mounds earlier, but sporulates later, than the wild type. Its spores exhibit aberrant shape, and their viability is less than 0.001% when compared with the wild type (Ueki and Inouye, 2001). The dramatic sporulation defect of the triple mutant compared with that of the single mutants suggests that SigB,
SigC, and SigE have partially redundant functions during M. xanthus development. SigF is less similar in sequence than SigA-E, including in region 2.2 (Fig. l),which explains why it was not detected by the Southern blot approach used to identify sigB-E (Ueki and Inouye, 2001; Ueki et al., 2005). However, SigF is more similar to SigA-E than it is to SigG or ECF sigma factors found in the M. xanthus genome (D. Srinivasan, C. Wilkerson, and L. Kroos, unpublished data). An insertion mutant of sigF is not able to form fruiting bodies on nutrient-limited agar plates, whereas the mutant is still capable of forming spores as efficiently as the parental strain. Based on its poor motility on soft (0.3%) agar, the sigF mutant appears to be defective in S-motility (Ueki et al., 2005). Many genes involved in S-mstility have been identified (chapters 6 and 7). When expression of nine such genes or operons was examined in the sigF mutant by primer extension analysis, all were found to be expressed (Ueki et al., 2005). Therefore, it is possible that SigF directs transcription of an unidentified gene for social motility, which is also required for normal fruiting body formation. SigG shows low similarity to SigA-F (Fig. 1)but high similarity to E. coli FliA (T. Ueki and S. Inouye, unpublished data), which is involved in flagellar biosynthesis (Liu and Matsumura, 1995). A sigG deletion mutant shows no apparent defects in vegetative growth or fruiting body development (Ueki and Inouye, unpublished). Interestingly, analysis of DNA sequences surrounding the sigG gene revealed that it appears to be located in an operon encoding components of a type I11 secretion system. These systems are often found in pathogenic bacteria (Buttner and Bonas, 2002). While 111. xanthus is not known to be a plant or animal pathogen, it is reasonable to speculate that SigG regulates expression of a type 111 secretion system whose function remains to be elucidated.
ECF Sigmas As in other bacteria with a large number of sigma factors, such as Streptomyces coelicolor (Bentley et al., 2002) (see chapter 24), most of the expansion of the a 7 0 family in M. xanthus is due to members of the ECF subfamily (Goldman et al., 2006). ECF sigmas typically respond to external stimuli and direct transcription of genes whose products affect cell envelope functions such as transport, secretion, and homeostasis (Helmann, 2002). In Streptomyces species, at least one ECF sigma is required for aerial mycelium development (Bibb and Buttner, 2003; Yamazaki et al., 2000). While 38 ECF sigmas are predicted from analysis of the M. xanthus genome sequence (Goldman et al., 2006), only three have been
9. 211.
XANTHUS
TRANSCRIPTIONAL REGULATORY MECHANISMS
characterized in detail. CarQ regulates light-induced synthesis of carotenoids (Gorham et al., 1996), which protect cells from light damage (see chapter 12). RpoEl appears to regulate motility behavior (Ward et al., 1998). EcfA regulates expression of the early developmental gene identified by Tn5 lac a4445 (M. Esmaeiliyan, J. J. Rivera, and H. B. Kaplan, personal communication). An effort is under way to create an insertion mutation in each of the remaining 35 putative ECF sigma-encoding genes and examine the mutants for defects in motility, fruiting body development, and the production of certain secondary metabolites (S. Mittal, D. Srinivasan, R. Taylor, D. Krug, R. Welch, R. Muller, and L. Kroos, unpublished data). Hopefully, this will provide more insight into the role of ECF sigma factors in developmental gene transcription. ECF sigma activity is often regulated by a membraneembedded anti-sigma factor (Fig. 2) that is encoded by a gene directly downstream (Helmann, 2002). The antisigma forms a complex with the ECF sigma that sequesters it from the core subunits of RNAP or prevents the holoenzyme from interacting with its cognate promoters. This appears to be the case for CarQ, which is followed by CarR, an anti-sigma that becomes unstable in illuminated stationary-phase cells (Browning et al., 2003), providing a possible mechanism for induction of carotenogenesis by light (see chapter 12). Downstream of the ecfA gene is reaA, which encodes a negative regulator of 4445 expression that is likely to be an anti-sigma factor (Esmaeiliyan et al., personal communication). Like signal
1
1 1 target gene Figure 2 Typical a n t i d o regulatory circuit. A signal leads to destruction of the integral membrane anti-a, freeing the o to bind RNAP core subunits a,pp', and the resulting holoenzyme transcribes an operon encoding the u and its anti-o, as well as target genes of the regulon.
153
CarR, ReaA has a predicted transmembrane segment, but unlike CarR, ReaA has a predicted metal-binding motif (consensus sequence HXXXCXXC) like that found in S. coelicolor RsrA and related proteins (Paget and Buttner, 2003). RsrA is a zinc-containing anti-sigma factor that responds to oxidative stress, but not all RsrArelated proteins respond to oxidation (Helmann, 2002). Inspection of proteins encoded immediately downstream of anti-sigma factors in the M. xanthus genome revealed 17 proteins, in addition to EcfA, with a putative metalbinding motif (D. Srinivasan, J. J. Rivera, H. B. Kaplan, and L. Kroos, unpublished data).This is more than twice the number found in S. coelicolor (Bentley et al., 2002) and more than any other organism sequenced to date. Many questions remain about the ECF sigmas and their anti-sigmas. To what signal does each anti-sigma respond? How does the signal lead to anti-sigma inactivation? Which genes are under the control of each ECF sigma and what is the function(s) of each regulon? Important questions also remain about the organization of sigma factors into one or more cascades during M. xanthus development. Two sigma factor cascades govern differential gene expression in the forespore and mother cell during B. subtilis endospore formation (see chapter 21). So far, no sigma factor has been shown to be responsible for transcription of another sigma factor gene in M. xanthus. Many sigma factors direct transcription of the gene that encodes them, forming an autoregulatory loop, so it is conceivable that sigma cascades are not used during M . xanthus development.
TRANSCRIPTIONAL ACTIVATION AND REPRESSION Transcriptional activation, rather than relief from repression, appears to account for induction of most developmentally regulated M. xanthus genes studied so far, though some genes are subject to both positive and negative control. Most developmental activators studied so far are components of signal transduction pathways that involve two or more proteins. An exception appears to be MrpC, though its expression is controlled by the MrpAB two-component system (see below). A paucity of one-component regulators like MrpC (i.e., transcription factors likely controlled by binding of a small molecule) has been noted in the genome (Goldman et al., 2006). Considering the families of such regulators that are missing (IclR, LacI, ROK, and DeoR) or underrepresented (AraC, GntR, AsnC, and LuxR), which primarily regulate genes involved in sugar metabolism, this may reflect the inability of M. xanthus to utilize hexoses, pentoses, and polysaccharides and its preference for pyruvate or
154
amino acids as carbon and energy sources (Kaiser and Manoil, 1979). Beyond the apparent predominance of transcriptional activation likely mediated by sensor kinases that phosphorylate response regulators of twocomponent systems, which presumably couples developmental gene expression to extracellular and intracellular signals, our current knowledge is insufficient to discern much about the basic circuitry and fundamental logic of the regulatory network. Below, we describe first the few examples of negative regulation so far uncovered and then the numerous putative transcriptional activators.
Negative Regulation Only a few negative regulators have been implicated to change the timing or level of developmental gene expression, and in no case has repression of transcription been demonstrated in vitro. The first report of a putative repressor came from studies of expression of Tn5 lac 04514 (Ha0 et al., 2002). As noted above, the promoter of the operon identified by this insertion is the only developmentally regulated promoter that has been shown to be recognized by oA-RNAPin vitro. How, then, is expression of the operon prevented during growth and induced at about 9 h into development? There appear to be two mechanisms. One mechanism involves the product of the first gene of the operon, ORF1, acting negatively (i.e., negative autoregulation). ORFl has a putative helix-turn-helix (HTH)DNA-binding motif and is most similar to members of the TetR family of repressors. While the simple model that ORFl is a repressor of the 04514 promoter is attractive, recombinant ORFl produced in E. coli failed to bind to the 04514 promoter region in electrophoretic mobility shift assays (EMSAs). Whether direct or indirect, the role of ORFl is primarily to delay and reduce expression of the operon during development, rather than prevent transcription during growth. Deletion of orfl results in only slightly higher expression during growth, but during development expression increases earlier and reaches a higher level. This implies that a second mechanism regulates 04514 promoter activity in the absence of ORF1, and 5’ deletion analysis suggests positive regulation by one or more transcriptional activators, which have not been identified. The orfl mutant exhibits fourfold reduced sporulation, due either to misregulation of downstream genes in the operon or other genes. The two genes immediately downstream of orf2 are predicted to encode subunits of glutaconate coenzyme A-transferase, which is involved in glutamate fermentation. Null mutations in these genes do not reduce sporulation. A DNA-binding protein is implicated in negative regulation of fruA (Horiuchi et al., 2003), which is believed
REGULATORY MECHANISMS to encode a key activator of developmental genes (see below). EMSAs detected a protein that was designated Factor X, which bound specifically to a sequence (xbs) located downstream (+78 to +94) of the fruA transcriptional start site. Comparison of expression of lac2 fusions upstream or downstream of xbs supports the idea that Factor X binding to xbs negatively regulates fruA expression. Loss of xbs did not result in constitutive expression because like 04514, fruA is subject to positive control during development (see below). Factor X was present during growth and development, and placing xbs downstream of a vegetative promoter (uegA) greatly reduced expression of a lac2 reporter located farther downstream. The ability of xbs to mediate negative regulation of a different promoter might mean that Factor X blocks elongation rather than initiation of transcription. HthA is a putative DNA-binding protein involved in negative regulation of sdeK (Nielsen et al., 2004). SdeK is a histidine protein kinase (HPK)required for expression of developmental genes that also depend on C-signaling (Pollack and Singer, 2001). The hthA gene is cotranscribed with hthB (Nielsen et al., 2004). The sequence of HthB provides no clue to its function. Expression of sdeK was increased during growth and development of an hthA hthB double mutant, indicative of negative regulation by HthA and/or HthB, but it is unknown whether this is a direct or indirect effect. Intriguingly, an upstream DNA element negatively regulates activity of the 04406 promoter and this is relieved by C-signaling (Viswanathan et al., 2006a). Based on mutational and DNA sequence analyses, two 18-bp sequences located at bp -317 to -300 and -89 to -72, which match at 14 positions and are partially palindromic, are proposed to be bound by a repressor, forming a DNA loop that inhibits transcription. Many aspects of the model remain to be tested, and the putative repressor has not been identified. Nevertheless, it is clear that a 5’ deletion to bp -100 renders the 04406 promoter fourfold more active during development and independent of C-signaling. The deletion does not increase expression during growth. Like the 04514 and fruA promoters, the 04406 promoter appears to require positive regulation by one or more transcriptional activators (see below). The preceding examples of negative regulation, uncovered so far by studies of developmentally regulated M. xanthus genes, suggest that a combination of negative and positive control is used for some genes. None of the examples is well understood in terms of identifying the cis-acting DNA element and the trans-acting regulatory protein and its mechanism of action. Even more
9. M.
XANTHUS
TRANSCRIPTIONAL REGULATORY MECHANISMS
mysterious are two examples of negative regulation that are observed only when the promoter (04403 or 04406) is moved from its native location in the chromosome to the Mx8 phage attachment site (Fisseha et al., 1996; Loconto et al., 2005). In one of these cases, the negative DNA element is located 500 bp downstream of the promoter (Loconto et al., 2005).
155
signal
signal
1
1
Transcriptional Activators In eukaryotic model systems for the study of multicellular development, such as fruit flies, nematodes, sea urchins, frogs, and mice, the synthesis and activation of transcriptional activators in response to intracellular cues and extracellular signals are believed to be the principal mechanisms of gene regulation (Stathopoulos and Levine, 2005). Cascades of activators have been identified in the regulatory networks. While activators play an important role in gene regulation during B. subtilis sporulation, most of the regulation is accomplished by two cascades of sigma factors (see chapter 21). The two cascades are in different cell types and are connected to each other by signal transduction pathways that govern sigma factor activation. As noted above, there is so far no evidence of a sigma cascade operating during M. xanthus development. On the other hand, one apparent instance of a transcriptional activator (MrpC) binding to the promoter region of a gene encoding a later-acting activator (FruA) has been discovered (Ueki and Inouye, 2003), and there are indications of other activator cascades. By elucidating the direct connections in the M. xanthus regulatory network, we will learn whether it utilizes sigma cascades like B. subtilis, activator cascades like multicellular eukaryotes, or, perhaps most likely, a combination of these mechanisms. Bacterial activators likely to stimulate transcription by aS4-RNAPcan be recognized by their conserved ATPase domain (Buck et al., 2000). This domain is typically centrally located in the protein and followed by a C-terminal HTH DNA-binding domain. In M. xanthus, 52 genes predicted to encode this type of transcriptional activator have been found in the genome, and the predicted activator proteins have been called as4-activators(Gorski and Kaiser, 1998),NtrC-like activators (Caberoy et al., 2003), or EBPs (Jakobsen et al., 2004; Jelsbak et al., 2005) in large-scale gene knockout and expression studies. Most of these EBPs have a recognizable N-terminal sensory domain and presumably function in a signal transduction pathway (Fig. 3). Often, this domain resembles the receiver domain of response regulators that function in two-component systems. Such EBPs are one type of response regulator. Other response regulators do not have a central
Figure 3 Three types of transcriptional activators involved in M. xanthus development. The left part depicts an HPK or STPK in the inner membrane, undergoing autophosphorylation in response to an extracellular signal that has traversed the outer membrane (not shown) and is present in the periplasm. Transfer of phosphate from ATP to the FHA domain of an EBP by the STPK, or from the HPK to the receiver domain of a response regulator EBP, is proposed to facilitate DNA binding and/or oligomerization of the EBP. EBPs typically bind to DNA 70 to 150 bp upstream of the transcriptional start site (Buck et al., 2000). From the more distal sites, DNA looping is required for the EBP to interact with d4-RNAP.ATP hydrolysis by the EBP allows it to convert the d4-RNAP closed promoter complex to the open complex, activating transcription. The upper right part depicts a membrane-embedded HPK sensing a signal (e.g., C-signal) and transferring phosphate to FruA, a response regulator that is not an EBP. This is speculative since an HPK that phosphorylates FruA has not yet been identified. Phosphorylated FruA is shown interacting with RNAP containing a u70 family member (u’O-RNAP) whose identity also has not been established. Based on the sites of binding of the FruA DNA-binding domain mapped so far, DNA looping may not be required for FruA to interact with RNAP, which presumably facilitates recruitment or a subsequent step in transcription initiation. While FruA phosphoryiation likely involves at least one membrane-embedded HPK that responds to extracellular C-signal, the pathway might be more complex (e.g., one or more phosphotransfer proteins might function between the HPK and FruA) and FruA might also be phosphorylated by one or more HPKs that are not membrane embedded and respond to intracellular signals. Likewise, EBP phosphorylation pathways can involve more than two components and can respond to intracellular cues via cytoplasmic kinases. The lower right part depicts MrpC responding to an unknown cytoplasmic signal by binding to DNA and activating transcription, an example of a one-component system. For simplicity, MrpC is shown binding to the same promoter region as phosphorylated FruA, and the 04400 promoter region is the first example of such a promoter region (Yoder-Himes and Kroos, 2006; Mittal and Kroos, unpublished). o7O-RNAP denotes RNAP containing a sigma factor in the u70 family. The identity of the sigma factor responsible for 04400 promoter recognition is unknown.
REGULATORY MECHANISMS
156 ATPase domain and therefore are not EBPs that stimulate transcription by d4-RNAP. These response regulators have a predicted N-terminal receiver domain, and some have a C-terminal HTH DNA-binding domain (e.g., FruA) and likely stimulate transcription by RNAP containing a 0 7 0 family member (Fig. 3). Other putative transcriptional activators are neither response regulators nor EBPs (e.g., MrpC), but they have a predicted DNA-binding domain and in some cases one or more other domains found in proteins known to bind a small molecule and stimulate transcription by RNAP containing a 0 7 0 family member (Fig. 3). As noted above, these are called one-component regulators and are relatively scarce in the M. xanthus genome. Below, we describe the apparent involvement of EBPs in developmental gene transcription, based on mutational and expression studies; then, we describe MrpC and FruA, whose direct targets for activation are inferred from DNA-binding studies. Activated transcription in vitro has not yet been accomplished for any developmentally regulated M. xanthus gene.
Table 3
EBPs EBPs play a critical role in gene activation during M . xanthus development. Of the 52 genes in M . xanthus predicted to encode EBPs based on their conserved ATPase domain, 44 have been knocked out and 17 of the mutants exhibit defects in motility and/or development (Table 3 ) . Several EBPs appear to primarily affect motility. Since motility is crucial to bring cells into contact for C-signaling (chapter 4), the effects of mutations in these genes on development may be secondary to their motility defects. PilR is inferred to activate transcription of pilA (Table l),which encodes pilin, the protein used to build type IV pili necessary for S-motility (see chapter 6 ) . Defects in S-motility typically delay aggregation and may delay sporulation but do not necessarily reduce the firid spore number (e.g., nlal and nlu23 in Table 3 ) . Loss of both A- and S-motility, as in the nlu24 mutant, severely impairs sporulation (Table 3). The direct targets of activation by Nlal, Nla23, and Nla24 are unknown, but Nla23 is near genes involved in pilus biogenesis that might be its targets, and the nla24 mutant fails to
EBPs that affect motility and/or development
Gene (strain)
Mx no.
MXAN no.
Motility
Aggregation
pilR (Mxa15) nlal nla23 nla24 (Mxa296)
3013 3336 1973 2057 5565
5784 5853 5777 7440 0180
SSSA- SNormal
NRb Normal Normal <0.0002 NR
Gorski and Kaiser, 1998 Caberoy et al., 2003 Caberoy et al., 2003 Caberoy et al., 2003 Gorski and Kaiser, 1998
(DK1622::pLOJ3320)
3320
2902
NR
NR
Jakobsen et al., 2004
spdRc
1086
1078
S-
Delayed Delayed Delayed None Normal on TPM agar; none in submerged culture Normal on TPM agar and in submerged culture; incomplete on CF agar Accelerated or delayed for different mutants
Accelerated
crdA actB (Mxa259)
1467 4338
5153 3214
NR Normal
Delayed Delayed and incomplete
Normal <10-6
sas R mrpB nla4 nla6 nla28 (Mxa213) nlal8 Mx488.Y
0124 5602 0840 2063 1617 1598 0888 4885
1245 5124 2516 4042 1167 4020 3692 4899
Normal NR Normal Normal Normal Normal Normal Normal
None None Delayed and incomplete Delayed Delayed Larger aggregates Delayed Incomplete
NR None 0.2 0.2 2
Caberoy et al., 2003; Hager et al., 2001; Tse and Gill, 2002 Kirby and Zusman, 2003 Gorski and Kaiser, 1998; Gorski et al., 2000; Gronewold and Kaiser, 2001,2002 Guo et al., 2000 Sun and Shi, 2001a, 2001b Caberoy et al., 2003 Caberoy et al., 2003 Caberoy et al., 2003 Gorski and Kaiser, 1998 Caberoy et al., 2003 Jelsbak et al., 2005
"Numbers represent percentage relative to the wild type. "NR, not reported. 'This gene was called nlu19 in Caberoy et al., 2003.
Sporulation"
<0.0002 0.2
Reference(s)
9. M.
XANTHUS
TRANSCRIPTIONAL REGULATORY MECHANISMS
express genes required for A- and S-motility (Caberoy et al., 2003; Lancero et al., 2004). Interestingly, the nla24 mutant is defective for glycerol-induced sporulation, which does not require motility, so the targets of Nla24 might include genes required for cellular differentiation as well as motility genes. Two EBPs affect development only under certain conditions. Strain Mxa296 with a mutation in M X A N O l 8 0 develops normally on Tris-phosphate-magnesium (TPM) agar but fails to develop in submerged culture (Table 3). Both conditions invoke sudden, complete starvation. Submerged culture requires that cells adhere to polystyrene, so perhaps MXANO180 targets genes involved in that process or a subsequent step specific to these conditions. Strain DK1622::pLOJ3320 with a mutation in Mx3320 develops normally on TPM agar or in submerged culture, but development is impaired on clone fruiting (CF) agar (Table 3). Adding 1.5 mM (NH,),SO,, the concentration found in CF agar, to TPM agar resulted in a developmental defect similar to that seen on CF agar, suggesting that Mx3320 might be involved in the response to nitrogen availability during development (Jakobsen et al., 2004). Mx3320 was one of three genes in Table 3 that were found to be about threefold induced at 12 h into development in a microarray expression experiment (Jakobsen et al., 2004). The other two genes were pilR and spdR. Two EBPs, SpdR and CrdA, influence the timing of aggregation but do not prevent spore formation (Table 3 ) . Mutations in spdR allow aberrant expression of tps during the late log to early stationary phases of growth (Hager et al., 2001). Normally, t p s is not induced until about 6 h into development (Downard et al., 1984; Inouye et al., 1979a). It encodes protein S, which forms an outer coat of myxospores (Inouyeet al., 1979b).Other developmental genes are not induced aberrantly in spdR mutants; however, spdR mutations permit increased developmental expression of several genes when combined with a bsgA mutation (Hager et al., 2001). In addition, the spdR bsgA double mutant, unlike a bsgA mutant, constructs spore-filled fruiting bodies. BsgA is a protease similar to E. coli LonD that is involved in generating the B-signal (Gill et al., 1993). Neither the substrate of BsgA nor the molecular nature of the B-signal is known. Mutations in spdR bypass not only the need for B-signaling but also the requirement for A-signaling (Tse and Gill, 2002). Hence, SpdR is thought to activate genes that negatively regulate early developmental events, and A- and B-signalingmay normally overcome this inhibition (see chapter 3).Consistent with this idea, Hager et al. (Hager et al., 2001) reported accelerated aggregation and sporulation of an EMS-induced spdR mutant. However, Caberoy et al. (Caberoy et al., 2003)
157
reported delayed aggregation of an spdR (called nlal9 in their report) disruption mutant created by plasmid insertion and found that this mutation causes a defect in S-motility. Also, SpdR (Nla19) was found to interact with DifE, which is involved in the production of fibrils needed for S-motility (Lancero et al., 2005). Like SpdR, CrdA is an EBP that influences the timing of aggregation but not the ability to form spores (Table 3). The targets of these EBPs are unknown, but in the case of CrdA, regulatory input likely involves phosphorylation of its N-terminal receiver domain by the HPK CheA3, which appears to be part of a chemotaxis-like chemosensory system (Kirby and Zusman, 2003). This provided the first example of a chemotaxis-like pathway controlling activity of a transcriptional activator, a mechanism employed not only by M. xanthus but by other bacteria as well (see chapter 8). The nine EBPs in the lower part of Table 3 appear to primarily affect development. Mutations in these genes impair both aggregation and sporulation without detectably altering A- or S-motility on nutrient agar. Among these nine EBPs, the first six (beginning with ActB) listed in Table 3 are predicted to have N-terminal receiver domains typical of response regulators that function in two-component signal transduction systems, whereas the last three are predicted to have N-terminal forkheadassociated (FHA) domains (Jelsbak et al., 2005). FHA domains mediate phosphothreonine-dependent proteinprotein interactions (Durocher and Jackson, 2002). M. xanthus has a large family of putative Ser/Thr protein kinases (STPKs) (see chapter 11).It has been proposed that autophosphorylation of an STPK in response to a signal could result in association of an FHA-EBP and that phosphorylation of the FHA-EBP could allow it to activate transcription by d4-RNAP (Kroos, 2005) (Fig. 3). For the nine EBPs that primarily affect development, important questions remain about their cognate protein kinases and the signals to which they respond (i.e., input), as well as the target genes that they regulate (i.e., output). ActB is encoded by the second gene of an operon that regulates the level of CsgA (Gorski et al., 2000; Gronewold and Kaiser, 2001), the C-signaling protein (see chapter 4), during M. xanthus development. It has been postulated that ActB activates csgA transcription in response to C-signaling, creating a positive-feedback loop that boosts the level of CsgA during development (Gorski et al., 2000; Gronewold and Kaiser, 2001). However, the csgA promoter (Li et al., 1992) does not resemble those predicted to be transcribed by oS4-RNAP (Table 1).While there are precedents for EBP and EBPlike proteins activating transcription from promoters
REGULATORY MECHANISMS
158
(Guo et al., 2000). Immediately downstream of sasR in the 211. xanthus chromosome is sasN, which appears to encode a negative regulator of this signal transduction system (Xu et al., 1998).It is unlikely that SasN is a direct negative regulator of spi transcription since SasN has no apparent DNA-binding domain. On the other hand, SasR’s predicted C-terminal HTH may very well bind to an essential cis-acting element located between - 146 and -90 bp relative to the spi transcriptional start site, and SasR in extracts of developing 211. xanthus might account for retardation of DNA fragments containing the element in EMSAs (Gulati et al., 1995), though this hypothesis remains to be tested. Unless Sass phosphorylates another response regulator in addition to SasR, the phenotype of a sass null mutant is expected to be the same %s the phenotype of a sasR null mutant, which has not been reported. A sass null mutant forms defective fruiting bodies with 5 to 10% the wild-type number of spores (Yang and Kaplan, 1997). This implies that the Sass-SasR pathway regulates other developmental genes in addition to spi since Tn5 lac insertion a 4 5 2 1 in spi did not interfere with development (Kroos et al., 1986). MrpB is inferred to activate transcription of mrpC (Table l),a gene whose product binds to the frUA promoter region and likely activates transcription (Ueki and Inouye, 2003). If so, MrpB, MrpC, and FruA form a three-step transcription factor cascade that regulates genes essential for M. xanthus development (Fig. 4). The mrpB gene lies downstream of mrpA, and the two genes are cotranscribed (Sun and Shi, 2001a). MrpA is predicted to be an HPK that presumably phosphorylates MrpB. The signal(s) to which MrpA responds is unknown. A null mutation in mrpB blocks aggregation
transcribed by RNAP containing a sigma factor in the u70 family (Foster-Hartnett et al., 1994; Foster-Hartnett and Kranz, 1994; Kroos, 2005), it is possible that the connection between ActB and csgA is indirect. Recently, it was shown that expression of the act operon depends on csgA, actA, and actB (Gronewold and Kaiser, 2007). It remains to be determined whether dependence on csgA reflects a need for extracellular C-signaling. While it is attractive to think that ActB directly autoregulates the act operon, the DNA-binding domain of ActB does not appear to bind to the act promoter region (N. Caberoy and A. G. Garza, personal communication). An HPK that would phosphorylate ActB’s N-terminal receiver domain has not been identified. Neither has a cognate HPK been found for ActA’s predicted N-terminal receiver domain. The actA gene precedes actB in an operon, which appears to include at least five genes (Goldman et al., 2006). Interestingly, the C-terminal part of ActA is a GGDEF domain predicted to be in a novel class of guanylate cyclases that convert two molecules of GTP into cyclic diguanylic acid (c-di-GMP), which appears to be a second messenger widely used in bacteria (Paul et al., 2004). Hence, regulation of the CsgA level by Act proteins may be quite complex. SasR is inferred to activate transcription of spi, a developmentally regulated gene identified by Tn5 lac a 4 5 2 1 (Table 1).The spi gene is induced at about 2 h into development, and expression depends on A-signaling (Kuspa et al., 1986). Suppressor mutations that relieve the dependence of spi expression on A-signaling (Kaplan et al., 1991) led to the identification of Sass, an HPK that is thought to sense starvation and A-signal (Yang and Kaplan, 1997), and respond by phosphorylating SasR Starvation (P)PPGPP A-signal
........................................ .. .. .. i *+
v
1
mrpA
.1
@
L
I mrpc I
I J
.
f
?
I
@ ! @ ........... J . i . 1
-
C-signal (CsgA)
J
-
HPK112
~
r
~ J
C-signal-independent genes (tps, dofA)
~
-
~
\
C-signal-dependent genes (fdgA, sasA, dev, R4400, Q4403, R4406, C24499)
Figure 4 Model for a signaling and gene regulatory cascade leading to FruA-dependent gene expression. See the text for explanation.
+
~
9. M .
XANTHUS
TRANSCRIPTIONAL REGULATORY MECHANISMS
and sporulation, but an in-frame deletion in mrpA blocks only sporulation. This result, together with the phenotypes of mrpB mutants with single amino acid substitutions for the aspartate predicted to be phosphorylated in the receiver domain, suggests that MrpB phosphorylation by MrpA and at least one other (unidentified) HPK is essential for aggregation, and that MrpB dephosphorylation (possibly by MrpA) is required for sporulation. MrpA and MrpB function very early in development since a AmrpAB mutant fails to express the A-signalindependent sdeK gene (Sun and Shi, 2001b). Conceivably, MrpB could directly activate sdeK transcription from its putative os4-RNAP-transcribedpromoter (Table l),but this need not be the case since sdeK expression also depends on MrpC (Sun and Shi, 2001b), whose expression depends on MrpB (Sun and Shi, 2001a). Expression of the mrpAB operon is induced at the onset of development and depends to a small extent on (p)ppGpp- and A-signaling (Sun and Shi, 2001b). It will be interesting to see whether transcriptional activation plays a role in this induction, perhaps extending the putative transcription factor cascade to a fourth step that precedes MrpB, MrpC, and FruA (see below). Nla4 and Nla6 are EBPs of the response regulator type. Nla4 has no adjacent HPK in the chromosome. Nla6 is adjacent to a gene (MXAN4043) predicted to encode a protein resembling HPKs, but lacking two conserved domains found in most HPKs (A. G. Garza, personal communication). Neither the input nor the output of these response regulators is known. An nlu4 mutant is more defective in aggregation than an nlu6 mutant (Caberoy et al., 2003). Both mutants are defective in A-signal production, but again the nla4 mutant is more defective. Both mutants make about 500-fold fewer spores than the wild type during fruiting body development (Table 3), and neither could be rescued by codevelopment with wild-type cells (Caberoy et al., 2003). Also, both mutants fall into a rare class with reduced (about 20- to 40-fold) ability to form glycerol-induced spores. This combination of phenotypes suggests that Nla4 and Nla6 play not only an early role in A-signal production but also a later role in cellular differentiation in response to signals. Little more is known about Nla28 than about Nla4 and Nla6, except that Nla28 is not an orphan in the chromosome and the sporulation defects of an nla28 mutant are less severe. The predicted start codon of nla28 overlaps with the stop codon of MXANZ Z 66, which is predicted to encode an HPK (Goldman et al., 2006). An nlu28 mutant was about as defective as an nla6 mutant in A-signal production but made 10-fold more fruiting body spores (Table 3) and nearly as many glycerol spores as the wild type (Caberoy et al., 2003).
159
As noted above, the last three EBPs in Table 3 have N-terminal FHA domains that might be targets of STPIC signaling (Jelsbak et al., 2005). Consistent with this hypothesis, two of these FHA-EBPs have a predicted STPK adjacent or nearby (see chapter 11). However, the proximity of one of these, described in Table 3 as strain Mxa213, which has a mutation in MXAN4020, may be coincidental because MXAN4020 is separated by one gene from an operon that encodes both an STPK (Pkn4) and an enzyme, 6-phosphofructokinase (PFK), that is known to be a substrate of Pkn4 (Nariya and Inouye, 2002). Whether MXAN4020 is also a substrate of Pkn4 is an open question. If Pkn4-mediated phosphorylation of MXAN4020 was essential for its activity as a transcription factor during development, then apkn4 mutant would be expected to be as defective for development as the MXAN4020 mutant, but this is not the case. A pkn4 mutant sporulates at about 0.03% of wild-type efficiency (Nariya and Inouye, 2003), while the MXAN4020 mutant is at least 100-fold less efficient than the pkn4 mutant at forming myxospores (Table 3). Moreover, the sporulation defect of the pkn4 mutant can be accounted for by failure to phosphorylate PFK on Thr-226 (Nariya and Inouye, 2003), so MXAN4020 need not be a substrate of Pkn4. Nla18 is a predicted FHA-EBP adjacent to a divergent predicted STPK, MXAN3693, but it is unknown whether the two form a signal transduction system (Caberoy et al., 2003; Goldman et al., 2006; Jelsbak et al., 2005). Like Nla4 and Nla6, Nla18 appears to function both early and late in development. An early role is implied because an nlul8 mutant is defective in A-signal production and a late role is inferred from the mutant’s near inability (1,000-fold reduced) to form glycerol spores (Caberoy et al., 2003). However, the inability of the nlul8 mutant to make fruiting body spores (Table 3) can be partially rescued by codevelopment with wildtype cells, distinguishing this mutant from the nlu4 and nlu6 mutants (Caberoy et al., 2003). This result indicates that the nlaZ8 mutant retains some ability to respond to signals provided by wild-type cells during development. Further characterization of the nluZ8 mutant revealed a decrease in growth rate, altered expression of more than 700 genes in growing cells, and failure to accumulate (p)ppGppupon starvation (Diodati et al., 2006). Clearly, Nla18 is a key regulator during growth and early in development, whose direct targets will be important to elucidate. Mx488.5 is a predicted FHA-EBP that appears to be involved in the response to C-signal (Jelsbak et al., 2005). The FHA domain is crucial for Mx4885 function since an in-frame deletion of just this part of the gene caused the
REGULATORY MECHANISMS
160 same developmental defect as disruption of Mx4885. The
Mx4885 mutants fail to complete aggregation (Table 3 ) , much like csgA mutants unable to produce C-signal (Jelsbak et al., 2005).However, Mx4885 mutants produce normal levels of both C-signal and the protein that mediates cellular responses to C-signal, FruA. Also, unlike csgA or fruA mutants, Mx488.5 mutants respond to C-signaling by forming transient ridges between nascent fruiting bodies. This rippling behavior indicates the ability to respond to a low level of C-signaling (chapter 4). Moreover, Mx4885 mutants (Table 3 ) are less defective than csgA or fruA mutants with respect to sporulation and expression of Csignal-dependent genes (Jelsbak et al., 2005). This partial defect in responses to C-signaling, together with normal accumulation of C-signal and FruA in an Mx4885 null mutant, led Jelsbak et al. (Jelsbak et al., 2005) to propose that Mx4885 somehow facilitates FruA phosphorylation, which is believed to be a key event in the C-signal transduction pathway (Fig. 4). A simple model would be that Mx4885’s (unidentified) cognate STPK undergoes autophosphorylation in response to an early developmental signal; Mx4885 via its FHA domain interacts with, and is phosphorylated by, the STPK; and phosphorylated Mx4885 activates transcription by oS4-RNAPof a target gene(s) whose product (e.g., an unidentified HPK) facilitates phosphorylation of the FruA response regulator in response to C-signal (Kroos, 2005). Of the nine EBPs that primarily affect development, at least four (nla4,nla6, nla28, and nla18) are required for normal production of A-signal early in development, one (sasR) activates transcription in response to A-signal, one (actB)is required for normal production of C-signal later in development, and two (mrpB and Mx4885) enable cells to respond to C-signal. Strain Mxa213 with a mutation in MXAN4020 forms large aggregates at the normal time early in development, but these fail to progress to tight mounds, suggestive of a defect in C-signal production or response. Microarray expression profiling is being used to determine more precisely when and how developmental gene regulation goes awry in EBP mutants (Diodati et al., 2006) (Garza, personal communication). Several EBP-encoding genes are expressed at the time during development when the EBP is required for expression of other genes (Garza, personal communication). Tests of epistasis among the EBP-encoding genes suggest that Nla18 and Nla4 function first in a pathway, followed by Nla6, then Nla28, and finally MrpB, and in vitro DNA-binding studies support the idea that the steps in this pathway involve one EBP directly activating transcription of the next EBP-encoding gene (i.e., a transcription factor cascade) (Caberoy and Garza, personal communication).
The EBP cascade culminates in induction of the mrpAB operon, and the apparent transcription factor cascade continues with first MrpC and then FruA (Fig. 4). Evidence for parts of this cascade includes mapping mRNA 5’ ends to infer transcriptional start sites, mutational analysis of promoter regions, and EMSA and footprinting experiments to demonstrate DNA binding, but the ultimate evidence for direct effects on transcription will come from reconstitution of activated transcription in vitro with purified proteins.
MrpC MrpC is encoded by a gene that lies downstream of the mrpAB operon (Sun and Shi, 2001a) (Fig. 4). The mrpC gene~hasa putative as4-RNAP-transcribed promoter (Table 1)that is inferred to be activated by MrpB (Sun and Shi, 2001a). Also, MrpC positively autoregulates mrpC expression (Nariya and Inouye, 2005; Sun and Shi, 2001a), although the mechanism involved is a mystery since MrpC is not similar to EBPs. Rather, it is similar to cyclic AMP receptor protein (CRP)-like transcriptional regulators and is predicted to have a central cyclic nucleotide-binding domain and C-terminal HTH. Hence, transcriptional activation by MrpC might depend on a cyclic nucleotide or some other small-molecule effector (Fig. 3). Ueki and Inouye (Ueki and Inouye, 2003) noted that some residues of E. coli CRP that contact cyclic AMP are not conserved in MrpC and suggested that MrpC’s effector might instead be (p)ppGpp, an important signal early in M. xanthus development (see chapter 3 ) . They discovered that an N-terminally truncated form of MrpC in extracts of 12-h developing M. xanthus binds specifically to a cis-acting element located between bp -154 and -107 with respect to the fruA transcription initiation site. Within the element, two binding sites for a truncated form of MrpC, called MrpC2, were identified and found to contain inverted repeat sequences. Mutations in the binding sites for MrpC2 in the fruA promoter region impaired MrpC2 binding in vitro and fruA expression in vivo, providing strong evidence that an N-terminally truncated form of MrpC directly activates fruA transcription during M . xanthus development. As expected, fruA transcripts were not detectable in an mrpC mutant. Recently, it was shown that MrpC2 binds with higher affinity than MrpC to both the fruA and the mrpC promoter regions (Nariya and Inouye, 2006). Binding of MrpC2 to the mrpC promoter region suggests direct autoregulation and is consistent with the positive autoregulation observed in vivo (Nariya and Inouye, 2005; Sun and Shi, 2001a). It remains to be seen whether MrpC2 activates mrpC transcription by os4 RNAP or another holoenzyme.
9. M. X A N T H U S TRANSCRIPTIONAL REGULATORY MECHANISMS FruA FruA is a response regulator that responds to C-signal by governing motility behavior and activating transcription of genes required for sporulation (Ellehauge et al., 1998; Ogawa et al., 1996; Sogaard-Andersen et al., 1996; Ueki and Inouye, 2005a; Viswanathan et al., 2007b) (Fig. 4). The role of C-signaling and FruA in regulating the reversal frequency of gliding cell movements during developmental rippling and aggregation is discussed in chapters 4 and 5. Here, we focus on FruA as a transcriptional activator. Analysis of protein expression patterns in wild-type, AfruA, and AcsgA strains indicates that developmental genes under the control of FruA can be classified into two groups: csgA independent and csgA dependent (Horiuchi et al., 2002). FruA appears to directly activate dofA transcription, which does not depend on C-signaling, by binding to the upstream region of the dofA promoter from bp -57 to -42 and from bp -82 to -67 with respect to the transcription initiation site (Ueki and Inouye, 2005b) (a general model for activation of transcription by FruA is shown in Fig. 3). In addition, the fdgA gene, whose expression depends partially on csgA, was isolated as a likely transcription target of FruA from a genomic DNA library via in vitro selection in a DNAbinding assay using the DNA-binding domain of FruA (FruA-DBD) (Ueki and Inouye, 2005a). The fdgA gene encodes a protein similar to the outer membrane auxiliary family protein involved in the polysaccharide export system of Xanthomonas campestris. Polysaccharide has long been known to be crucial for M. xanthus development (Dworkin, 1993). FruA-DBD binds upstream of the fdgA P,, promoter from bp -89 to -64, a region required for the induction of fdgA expression during development (Ueki and Inouye, 2005a). The fdgA gene also appears to be transcribed from a second FruAdependent promoter, PD2,with an inferred initiation site 81 bp downstream of that for P,,. Transcription from P,, was normal in a csgA mutant, while that from P,, was reduced. In the same study, FruA was also shown to be required for developmental expression of sasA, which also depends partially on csgA. However, FruA-DBD did not bind detectably to the sasA promoter region, so it is not clear whether FruA directly activates sasA transcription. Proteins encoded at the sasA locus are involved in the biosynthesis of the lipopolysaccharide 0-antigen and are required for fruiting body development (Bowden and Kaplan, 1998; Guo et al., 1996; Kaplan et al., 1991). To establish a consensus sequence for FruA binding, in vitro selection was conducted with purified FruA protein and randomized oligonucleotides (C. Y. Xu and s. Inouye, unpublished data).The selected oligonucleotides were found to contain sequences similar to 5’TNUN(A/
161
C)CYNNAGGGCN3’, where U and Y represent G/A and C/T, respectively, and N can be any nucleotide. It may be possible to identify other genes regulated by FruA by searching the M. xanthus genome for sequences similar to this consensus. Analysis of the FruA sequence suggests that it is a response regulator of a two-component system (Ellehauge et al., 1998; Ogawa et al., 1996), but a cognate HPK has not been identified. D59 of FruA has been proposed to be the phosphorylation site, because mutant FruA with a D59A substitution fails to complement the developmental defect of a fruA deletion mutant, whereas FruA D59E, in which glutamate is expected to mimic phosphorylated aspartate, restored development (Ellehauge et al., 1998). D59 phosphorylation also appears to be important for FruA to function as a transcriptional activator, because expression of dofA- and tps-lacZ fusions in a mutant expressing only FruA D59A was as low as in a fruA deletion mutant, whereas their expression in a mutant expressing only FruA D59E was 2.5-fold higher than in the wild type (Ueki and Inouye, unpublished data). On the basis of these results, a model for a signaling and gene regulatory cascade leading to FruA-dependent gene expression is proposed (Fig. 4). Starvation and to a small extent (p)ppGpp and A-signaling are needed to activate the mrpAB operon (Sun and Shi, 2001a, 2001b). Expression of mrpC depends on the mrpAB operon, and most likely MrpB is an activator of mrpC transcription by aS4-RNAP.MrpC autoregulates its own expression. N-terminally truncated MrpC induces fruA transcription by binding upstream of the fruA promoter. FruA is activated by phosphorylation of D59. In the early stage of development, a small amount of FruA is phosphorylated by (unidentified) HPK1, resulting in the expression of C-signal-independent genes such as dofA and tps. As development proceeds and the concentration of C-signal (CsgA) increases, more FruA becomes phosphorylated, possibly by a different HPK, HPK2, resulting in the induction of C-signal-dependent genes such as fdgA, sasA, dev, 04400, 04403, a4406, and a4499 (see below). Alternatively, a single HPK might phosphorylate FruA both in response to an early (unidentified) signal and later in response to C-signal.
&-Acting DNA Elements in C-SignalDependent Promoter Regions Other than the MrpC and FruA-DBD binding sites mentioned in the preceding sections, very few transcription factor binding sites have been mapped in M. xanthus promoter regions. On the other hand, cis-acting DNA elements that are likely bound by unidentified transcription factors have been described for several genes. Those
REGULATORY MECHANISMS
162
CAYYCCY, in which Y is C or T), designated the C box, in the promoter regions (Brandner and Kroos, 1998; Fisseha et al., 1996; Fisseha et al., 1999). These studies provided a foundation for detailed mutational analyses aimed at identifying important cis-acting DNA elements in C-signal-dependent promoter regions. Mutational analysis of the 04403 promoter region identified three elements essential for developmental expression; the C box, a 5-bp element, and a 10-bp element (Viswanathan and Kroos, 2003). The sequence and position of these elements are shown in Fig. 5. Inspection of other C-signal-dependent promoter regions revealed sequences similar to the 5- and 10-bp elements, as well as the C box, located at similar positions. These observations guided mutational analyses of other promoter regions. The 04400 promoter region has exactly the same C box at the same position as the 04403 promoter region (Fig. 5). The two promoter regions also have a similar 5-bp elemeat located 6 bp upstream of their C box. If these sequence elements were recognized by the same transcription factor(s),mutations in these elements would
described for tps, ops, and mbhA were reviewed in Myxobacteria I I (Downard and Kroos, 1993). Recently, detailed mutational analyses of promoter regions of genes whose expression depends on C-signaling have revealed cis-acting DNA elements with unusual features, which are described below. Many developmentally regulated M. xanthus genes were identified by transposition of Tn.5 lac into the chromosome (Kroos et al., 1986). Most of these insertion mutations did not cause a detectable developmental defect, but the lac2 fusions created have been extremely useful markers of developmental progression. For example, Tn.5 lac reporters revealed that an asg mutant defective in A-signaling expresses only a few very early developmental genes (Kuspa et al., 1986),whereas a csgA mutant unable to produce C-signal is blocked later in development, exhibiting reduced or abolished expression of nearly all genes induced after 6 h into development (Kroos and Kaiser, 1987; Li and Shimkets, 1993). Cloning of DNA upstream of C-signal-dependent Tn5 lac insertions and RNA mapping to infer transcriptional start sites led to the identification of a conserved sequence (consensus
-10 region TACACC
r, R4403
TACAAC
r, 04400
~CATTCCC~ GAACTICATTCCT~ CGAAAT
r, 04499
10-bp element 5-bp element GGCATGTTCA GACCG
C box
-35 region
ICATCCC~ TTCATG I
I
I
-74.5
-6 1
-49
GTCGGG
GAACA
EATCCCTl
I
I
I
-61
-49
-83.5
AGGCGC
GCCGC
GAACA
I
I
I
I
I
-79
-69
-55
-46
-33
GGGGTGAGCCTTTGGGG,,GACGA I
\\
-91
1
-60 GAACC I
-62 GCACA
I C A T ~ C A I C A GTTGACG ~ TATCGA I
I
-51
-47
r, dev
TTCGTG
TCTCAT
r, R4406
~ A C T C C ~ TTCGCG
TAGGGT
r, fruA
ICATCGT~ I -52
I
I
-62
-51
Figure 5 Conserved regulatory elements in C-signal-dependent promoter regions and in the fruA promoter region. The promoter -10 and -35 regions are shown, except in the case of R4499, which has a C box centered at -33 bp relative to the transcriptional start site (rightangle arrow). The position and sequence of C boxes (boxed; matching the consensus sequence CAYYCCY, in which Y means C or T, except in the cases of the deu and R4406 promoter regions, which contain C-box-like sequences) and 5-bp elements (bold, matching the consensus sequence GAACA) are shown for each promoter region. An essential 10-bp element is shown for the R4403 promoter region, and sequences centered at -83.5 and -79 bp that exert a twofold or more positive effect on a4400 and a4499 expression, respectively, are also shown. See the text for references.
9. M.
XANTHUS
TRANSCRIPTIONAL REGULATORY MECHANISMS
be predicted to have similar effects on promoter activity. However, this was not the case (Yoderand Kroos, 2004a). Single-base-pair changes had very different effects on the activities of the two promoters. In the case of the a4400 promoter, the entire region from -63 to -31 bp was required for activity and a second region from -86 to -81 bp had a two- to fourfold positive effect that appeared to be responsible for the partial C-signal dependence of activity. In support of the latter observation, FruA-DBD binds to R4400 upstream DNA and mutations between -86 and -77 bp impair binding (Yoder-Himes and Kroos, 2006). Moreover, FruA is essential for a4400 promoter activity and based on chromatin immunoprecipitation (ChIP) experiments FruA is associated with the a4400 promoter region, both in the presence and in the absence of C-signaling. Hence, partial C-signal dependence of a4400 promoter activity may involve transcriptional activation by a certain level of phosphorylated FruA in the absence of C-signaling and more vigorous activation by a higher level of phosphorylated FruA upon C-signaling. Interestingly, it appears that MrpC2 binds in the vicinity of the 5-bp element in the a4400 promoter region (S. Mittal and L. Kroos, unpublished data). The a4499 promoter region has two C boxes with a 5-bp element 7 or 8 bp upstream (Fig. 5 ) . All these sequences are important for a4499 promoter activity (Yoder and Kroos, 2004b). Single-base-pair changes in the C box centered at -33 bp had very different effects on promoter activity than the corresponding changes in the C boxes centered at -49 bp in the a4403 and 04400 promoter regions. Mutations in the 5-bp elements and the sequences between the 5-bp elements and C boxes also had different effects on activity of the three promoters. These observations led to the hypothesis that the 5-bp elements and C boxes (and in some cases the DNA in between) constitute recognition sites for a family of transcription factors that interact in slightly different ways with similar sequences. It is too soon to judge the validity of this hypothesis, but some evidence suggests that MrpC2 binds to the 5-bp element centered at - 69 bp in the a4499 promoter region, suggesting at least one similarity with the a4400 promoter region (Mittal and Kroos, unpublished). The R4499 promoter region also includes an element from about - 8 1to -77 bp that exerts a twofold positive effect on transcription; however, this region does not appear to mediate the partial C-signal dependence of R4499 transcription, as does the corresponding region upstream of the a4400 promoter. Recently, the dev operon was shown to include at least eight genes and two repeats of a downstream set of clustered regularly interspaced short palindromic repeats (CRISPR) (Viswanathan et al., 2007a). The promoter
163
was identified, and cis-regulatory elements were shown to span a region of more than 1 kb surrounding the pro-
moter. Upstream and downstream regulatory elements seem to interact functionally. The FruA DNA-binding domain binds to a mirror-repeat sequence centered at -91 bp (Fig. 5 ) ,suggesting that FruA activates dev transcription (Viswanathan et al., 2007b). Also, a LysR-type transcriptional activator designated LadA binds about 350 bp downstream of the promoter to a positive regulatory element that is essential for expression unless downstream DNA is removed. Exactly how much downstream DNA must be removed is unknown. A deletion to +71 bp was sufficient to alleviate the need for the positive regulatory element, but it remains to be tested whether LadA is dispensable for dev transcription under these conditions or whether LadA binds elsewhere in the dev promoter region. The dev promoter region has a 5-bp element centered at -60 bp and C-box-like sequences centered at -51 and -47 bp (Fig. 5 ) , but these sequences have not yet been subjected to mutational analysis. In contrast to the partial C-signal dependence of the R4400, R4499, and dev promoters, transcription from the a 4 4 0 6 promoter depends absolutely on C-signaling, like that from the a 4 4 0 3 promoter (Kroos and Kaiser, 1987; Loconto et al., 2005). As described above, Csignal dependence of the 04406 promoter is mediated by an upstream negative regulatory element (Viswanathan et al., 2006a). In csgA mutant cells that fail to produce C-signal, the upstream element prevents $24406 promoter activity. In wild-type cells, C-signaling only partially overcomes the negative effect of the upstream element, since deletion of the element results in a fourfold increase in promoter activity. Another unusual feature of a 4 4 0 6 regulation is that DNA between 50 and 140 bp downstream of the transcriptional start site boosts expression of a downstream lucZ reporter about threefold. This could be due to binding of an activator protein that effects initiation of transcription, or it could be due to an effect on a postinitiation event (e.g., transcription termination, mRNA stability, or translation). More proximal to the a 4 4 0 6 promoter, a 5-bp element and a C-box-like sequence are important for expression (Fig. 5), as in the other C-signal-dependent promoter regions, but the effects of single-base-pair changes are unique, consistent with the idea that these sequences are bound by a different transcription factor(s). Although transcription of fruA does not depend on C-signaling, the fruA promoter region also has a 5-bp element and C box that are crucial for activity (Srinivasan and Kroos, 2004) (Fig. 5 ) . Again, single-basepair changes indicate that these sequences function uniquely compared to other 5-bp elements and C boxes.
REGULATORY MECHANISMS
164 Obviously, understanding how 5-bp elements and C boxes exert their positive effects on promoter activity is an important future goal.
CONCLUSIONS AND FUTURE DIRECTIONS Several transcription factors key to the M. xanthus developmental process have been identified. Some of these, like MrpC and FruA, emerged from transposon mutagenesis screens. Others, like uB-E and uS4,were identified by cross-hybridization with other sigma factor genes. Still others, like most of the EBPs in Table 3 , were identified in large-scale gene knockout studies made possible by the availability of the genome sequence. Additional large-scale gene knockout studies under way are expected to yield more key transcription factors. A limitation of genetic approaches, though, is that some factors may be redundant in function (e.g., uB,uc, and uEappear to have partially redundant functions). DNA affinity chromatography with a cis-acting regulatory element from the fruA promoter region was used to purify a sequence-specific DNA-binding protein, which turned out to be the previously identified MrpC (Ueki and Inouye, 2003). This approach recently yielded LadA, a novel transcription factor involved in regulation of the dev operon (Viswanathan et al., 2007b). Identification of transcription factors is one step toward understanding the role of transcriptional regulatory mechanisms during M . xanthus development. Another step is to elucidate the input and output functions of each transcription factor in the regulatory network. Input may include mechanisms that control expression and activity of the transcription factor. For example, MrpB, MrpC, and FruA likely form a cascade that operates at the transcriptional level (Fig. 4), and other EBPs extend this regulatory chain (Caberoy and Garza, personal communication). Phosphorylation likely controls the activity of EBPs, MrpB, and FruA, and MrpC likely responds to a small-molecule signal. Identifying signals that affect transcription factor activity directly or via HPKs or STPIG is a major challenge, and M . xanthus development is fertile territory for new discoveries. Sigma factor activity is regulated in many different ways. Anti-sigmas and proteolytic processing play important roles during B. subtilis sporulation (see chapter 21). It will be interesting to see whether these mechanisms or novel ones are employed by M. xanthus. Understanding the output functions of a transcription factor includes identifying all genes in its regulon and determining how it exerts its effects. Regulon identification is becoming easier. The microarray expression profiling of a transcription factor mutant can be compared
with that of the wild type to identify candidate genes that might be under direct control of the transcription factor. In the case of a sigma factor, it can be overproduced in growing cells and the genes transcribed can be identified by microarray analysis, though genes requiring a development-specific activator will be missed. Transcription factor binding sites can be identified on a genome-wide basis using ChIP followed by microarray analysis (ChIP on chip). Recently, this approach has been used to identify genes transcribed by RNAP with a particular sigma factor (Herring et al., 2005). Once several promoters or other transcription factor binding sites have been identified, bioinformatic approaches can be used to search the genome for additional sites. The combination of genomic approaches just described provides a preliminary view of a transcription factor’s regulon. Typically, some of the results are verified by gene-specific ChIP and RNA analysis (e.g., quantitative reverse transcriptase PCR). Further verification involves in vitro studies such as EMSAs and footprinting of DNA-bin ’ng proteins and transcription with particular sigma-containing RNAP holoenzymes. These studies provide a foundation for further investigation of the mechanism(s) by which the transcription factor exerts its effects. So far, in vitro studies have been performed for only a few developmental genes of M. xanthus. The potential for discovery of novel transcriptional regulatory mechanisms in M . xanthus is vast and largely untapped. Another way to deepen understanding of a transcription factor’s output is to perform functional studies on members of its regulon. Bioinformatic approaches including phylogenomics can provide clues to function (Srinivasan et al., 2005). The available partial genome sequence of Stigmatella aurantiaca and the anticipated complete genome sequence of Sorangium cellulosum will aid these efforts, as well as comparative genomic approaches to understand the transcriptional regulatory network in each organism (i.e., transcription factors and their target sequences). The ease of knocking out genes in M . xanthus facilitates functional studies. Also, methods have been devised for more detailed phenotypic analysis of development by making a time-lapse microscopic movie of each mutant created (Kaiser and Welch, 2004). While great progress has been made toward understanding transcriptional regulatory mechanisms during M. xanthus development, since the writing of Myxobacteria I I , we have only seen the tip of the iceberg. The genome sequence together with genomic and bioinformatic approaches will further accelerate the pace of discovery in the coming years. Investigators will be challenged to assimilate the avalanche of information
dt
9. M. XANTHUS TRANSCRIPTIONAL REGULATORY MECHANISMS and make wise decisions about where to dig deeper. To those in the field, happy digging! We are grateful to P. Viswanathan, S. Mittal, and T Ueki for helpful comments on the manuscript. We thank those who shared information prior to publication, especially A. G. Garza and H. B. Kaplan. Research on M. xanthus in the lab of L.K. was supported by NSF grant MCB-0416456, and L.K. was supported in part by the Michigan Agricultural Experiment Station. Research in the lab of S.I. was supported by a grant from the Foundation of the University of Medicine and Dentistry of New Jersey.
References Apelian, D., and S. Inouye. 1990. Development-specifica-factor essential for late-stage differentiation of Myxococcus xanthus. Genes Dev. 4:1396-1403. Apelian, D., and S. Inouye. 1993. A new putative sigma factor of Myxococcus xanthus. 1.Bacteriol. 175:3335-3342. Bentley, S. D., K. F. Chater, A. M. Cerdeno-Tarraga, G. L. Challis, N. R. Thomson, K. D. James, D. E. Harris, M. A. Quail, H. Kieser, D. Harper, A. Bateman, S. Brown, G. Chandra, C. W. Chen, M. Collins, A. Cronin, A. Fraser, A. Goble, J. Hidalgo, T. Hornsby, S. Howarth, C. H. Huang, T. Kieser, L. Larke, L. Murphy, K. Oliver, S. O’Neil, E. Rabbinowitsch, M. A. Rajandream, K. Rutherford, S. Rutter, K. Seeger, D. Saunders, S. Sharp, R. Squares, S. Squares, K. Taylor, T. Warren, A. Wietzorrek, J. Woodward, B. G. Barrell, J. Parkhill, and D. A. Hopwood. 2002. Complete genome sequence of the model actinomycete Streptomyces coelicolor A3(2). Nature 417:141-147. Bibb, M. J., and M. J. Buttner. 2003. The Streptomyces coelicolor developmental transcription factor aBldN is synthesized as a proprotein. J. Bacteriol. 185:2338-2345. Biran, D., and L. Kroos. 1997. In vitro transcription of Myxococcus xanthus genes with RNA polymerase containing a*, the major sigma factor in growing cells. Mol. Microbiol. 25~463-472. Bowden, M. G., and H. B. Kaplan. 1998. The Myxococcus xanthus lipopolysaccharide 0-antigen is required for social motility and multicellular development. Mol. Microbiol. 30:275-284. Brandner, J. P., and L. Kroos. 1998. Identification of the 04400 regulatory region, a developmental promoter of Myxococcus xanthus. J. Bacteriol. 180:1995-2004. Browning, D. F., D. E. Whitworth, and D. A. Hodgson. 2003. Light-induced carotenogenesis in Myxococcus xanthus: functional characterization of the ECF sigma factor CarQ and antisigma factor CarR. Mol. Microbiol. 48:237-251. Buck, M., M. T. Gallegos, D. J. Studholme, Y. Guo, and J. D. Gralla. 2000. The bacterial enhancer-dependent d4 (aN) transcription factor. J. Bacteriol. 182:4129-4136. Buttner, D., and U. Bonas. 2002. Port of entry-the type I11 secretion translocon. Trends Microbiol. 10:186-192. Caberoy, N. B., R. D. Welch, J. S. Jakobsen, S. C. Slater, and A. G. Garza. 2003. Global mutational analysis of NtrC-like activators in Myxococcus xanthus: identifying activator mutants defective for motility and fruiting body development. ]. Bacteriol. 185:6083-6094.
165
Davis, J., J. Mayor, and L. Plamann. 1995. A missense mutation in rpoD results in an A-signalling defect in Myxococcus xanthus. Mol. Microbiol. 18:943-952. Diodati, M. E., F. Ossa, N. B. Caberoy, I. R. Jose, W. Hiraiwa, M. M. Igo, M. Singer, and A. G. Garza. 2006. Nla18, a key regulatory protein required for normal growth and development of Myxococcus xanthus. ]. Bacteriol. 188:1733-1743. Downard, J., and L. Kroos. 1993. Transcriptional regulation of developmental gene expression in Myxococcus xanthus, p. 183-199. In M. Dworkin and D. Kaiser (ed.),Myxobacteria II. American Society for Microbiology, Washington, DC. Downard, J. S., D. Kupfer, and D. R. Zusman. 1984. Gene expression during development of Myxococcus xanthus: analysis of the genes for protein S. J. Mol. Biol. 175:469492. Durocher, D., and S. P. Jackson. 2002. The FHA domain. FEBS Lett. 513:58-66. Dworkin, M. 1993. Cell surfaces and appendages, p. 63-83. In M. Dworkin and D. Kaiser (ed.), Myxobacteria II. American Society for Microbiology, Washington, DC. Ellehauge, E., M. Norregaard-Madsen, and L. SegaardAndersen. 1998. The FruA signal transduction protein provides a checkpoint for the temporal co-ordination of intercellular signals in Myxococcus xanthus development. Mol. Microbiol. 30:807-817. Fisseha, M., M. Gloudemans, R. Gill, and L. Kroos. 1996. Characterization of the regulatory region of a cell interaction-dependent gene in Myxococcus xanthus. J. Bacteriol. 178:2539-2550. Fisseha, M., D. Biran, and L. Kroos. 1999. Identification of the a4499 regulatory region controlling developmental expression of a Myxococcus xanthus cytochrome P-450 system. J . Bacteriol. 1815467-5475. Foster-Hartnett, D., P. J. Cullen, E. M. Monika, and R. G. Kranz. 1994. A new type of NtrC transcriptional activator. 1.Bacteriol. 176:6175-6187. Foster-Hartnett, D., and R. G. Kranz. 1994. The Rhodobacter capsulatus glnB gene is regulated by NtrC at tandem rpoNindependent promoters. J. Bacteriol. 1765171-5176. Garza, A. G., J. S. Pollack, B. Z . Harris, A. Lee, I. M. Kaseler, E. F. Licking, and M. Singer. 1998. SdeK is required for early fruiting body development in Myxococcus xanthus. J. Bacteriol. 180:4628-4637. Garza, A. G., B. Z . Harris, B. M. Greenberg, and M. Singer. 2000. Control of asgE expression during growth and development of Myxococcus xanthus. J. Bacteriol. 182:66226629. Gill, R. E., M. Karlok, and D. Benton. 1993. Myxococcus xanthus encodes an ATP-dependent protease which is required for developmental gene transcription and intercellular signaling. J. Bacteriol. 175:453 8-4544. Goldman, B. S., W. C. Nierman, D. Kaiser, S. C. Slater, A. S. Durkin, J. Eisen, C. M. Ronning, W. B. Barbazuk, M. Blanchard, C. Field, C. Halling, G. Hinkle, 0. Iartchuk, H. S. Kim, C. Mackenzie, R. Madupu, N. Miller, A. Shvartsbeyn, S. A. Sullivan, M. Vaudin, R. Wiegand, and H. B. Kaplan. 2006. Evolution of sensory complexity recorded in a myxobacterial genome. Proc. Natl. Acad. Sci. USA 103:15200-15205.
166 Gorham, H., S. McGowan, P. Robson, and D. Hodgson. 1996. Light-induced carotenogenesis in Myxococcus xanthus: light-dependent membrane sequestration of ECF sigma factor CarQ by anti-sigma factor CarR. Mol. Microbiol. 19:171-186. Gorski, L., and D. Kaiser. 1998. Targeted mutagenesis of us4 activator proteins in Myxococcus xanthus. J. Bacteriol. 1 8 0 5896-5905. Gorski, L., T. Gronewold, and D. Kaiser. 2000. A us4activator protein necessary for spore differentiation within the fruiting body of Myxococcus xanthus. ]. Bacteriol. 182:24382444. Gronewold, T. M., and D. Kaiser. 2001. The act operon controls the level and time of C-signal production for Myxococcus xanthus development. Mol. Microbiol. 40:744-756. Gronewold, T. M., and D. Kaiser. 2002. act operon control of developmental gene expression in Myxococcus xanthus. J. Bacteriol. 184:1172-1179. Gronewold, T. M., and D. Kaiser. 2007. Mutations of the act promoter in Myxococcus xanthus. J. Bacteriol. 189:18361844. Gruber, T. M., and D. A. Bryant. 1997. Molecular systematic studies of eubacteria, using sigma70-type sigma factors of group 1 and group 2. J. Bacteriol. 179:1734-1747. Gulati, P., D. Xu, and H. Kaplan. 1995. Identification of the minimum regulatory region of a Myxococcus xanthus Asignal-dependent developmental gene. ]. Bacteriol. 177: 4645-4651. Guo, D., M. G. Bowden, R. Pershad, and H. B. Kaplan. 1996. The Myxococcus xanthus rfbABC operon encodes an ATP-binding cassette transporter homolog required for 0antigen biosynthesis and multicellular development. J. Bacteriol. 178: 1631-1639. Guo, D., Y. Wu, and H. B. Kaplan. 2000. Identification and characterization of genes required for early Myxococcus xanthus developmental gene expression. J. Bacteriol. 182:4/564-4571. Hager, E., H. Tse, and R. E. Gill. 2001. Identification and characterization of spdR mutations that bypass the BsgA protease-dependent regulation of developmental gene expression in Myxococcus xanthus. Mol. Microbiol. 39:765-780. Hao, T., D. Biran, G. J. Velicer, and L. Kroos. 2002. Identification of the a 4 5 1 4 regulatory region, a developmental promoter of Myxococcus xanthus that is transcribed in vitro by the major vegetative RNA polymerase. J. Bacteriol. 184~3348-3359. Helmann, J. D., and M. J. Chamberlin. 1988. Structure and function of bacterial sigma factors. Annu. Rev. Biochem. 57~839-872. Helmann, J. D. 2002. The extracytoplasmic function (ECF) sigma factors. Adv. Microb. Physiol. 46:47-110. Herring, C. D., M. Raffaelle, T. E. Allen, E. I. Kanin, R. Landick, A. Z. Ansari, and B. 0. Palsson. 2005. Immobilization of Escherichia coli RNA polymerase and location of binding sites by use of chromatin immunoprecipitation and microarrays. J. Bacteriol. 187:6 166-61 74. Horiuchi, T., M. Taoka, T. Isobe, T. Komano, and S. Inouye. 2002. Role of fruA and csgA genes in gene expression during development of Myxococcus xanthus: analysis by
REGULATORY MECHANISMS two-dimensional gel electrophoresis. J. Biol. Chem. 27226753-26760. Horiuchi, T., T. Akiyama, S. Inouye, and T. Komano. 2003. Regulation of FruA expression during vegetative growth and development of Myxococcus xanthus. J. Mol. Microbiol. Biotechnol. 5937-96. Inouye, M., S. Inouye, and D. R. Zusman. 1979a. Gene expression during development of Myxococcus xanthus: pattern of protein synthesis. Dev. Biol. 68579-591. Inouye, M., S. Inouye, and D. R. Zusman. 1979b. Biosynthesis and self-assembly of protein S, a development-specific protein of Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 76~209-213. Inouye, S. 1990. Cloning and DNA sequence of the gene coding for the major sigma factor from Myxococcus xanthus. J. Bacteriol. 172530-85. Jakobsen, J. S.,L. Jelsbak, R.D. Welch, C. Cummings,B. Goldman, E. Stark, S. Slater, and D. Kaiser. 2004. us4 enhancer binding proteins and Myxococcus xanthus fruiting body development. J. Bacteriol. 186:43 61-4368. Jelsbak, L., M. Givskov, and D. Kaiser. 2005. Enhancerbinding proteins with an FHA domain and the uS4 regulon in Myxococcus xanthus fruiting body development. Proc. Natl. Acad. Sci. USA 102:3010-3015. Kaiser, D., and C. Manoil. 1979. Myxobacteria: cell interactions, genetics and development. Annu. Rev. Microbiol. 33~595-639. Kaiser, D., and R. Welch. 2004. Dynamics of fruiting body morphogenesis. J. Bacteriol. 186:919-927. Kaplan, H. B., A. Kuspa, and D. Kaiser. 1991. Suppressors that permit A-signal-independent development gene expression in Myxococcus xanthus. J. Bacteriol. 173:1460-1470. Keseler, I., and D. Kaiser. 1995. An early A-signal-dependent gene in Myxococcus xanthus has a us4-likepromoter. J. Bacteriol. 1724638-4644. Keseler, I., and D. Kaiser. 1997. uS4,a vital protein for Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 94:1979-1984. Kirby, J. R., and D. R. Zusman. 2003. Chemosensory regulation of developmental gene expression in Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 100:2008-2013. Komano, T., T. Franceschini, and S. Inouye. 1987. Identification of a vegetative promoter in Myxococcus xanthus: a protein that has homology to histones. J. Mol. Biol. 196517524. Kroos, L., A. Kuspa, and D. Kaiser. 1986. A global analysis of developmentally regulated genes in Myxococcus xanthus. Dev. Biol. 112252-266. Kroos, L., and D. Kaiser. 1987. Expression of many developmentally regulated genes in Myxococcus depends on a sequence of cell interactions. Genes Dev. 1:840-854. Kroos, L. 2005. Eukaryotic-like signaling and gene regulation in a prokaryote that undergoes multicellular development. Proc. Natl. Acad. Sci. USA 102:2681-2682. Kuspa, A., L. Kroos, and D. Kaiser. 1986. Intercellular signaling is required for developmental gene expression in Myxococcus xanthus. Dev. Biol. 112267-276. Lancero, H., N. B. Caberoy, S. Castaneda,Y. Li, A. Lu, D. Dutton, X. Y. Duan, H. B. Kaplan, W. Shi, and A. G. Garza.
9. 211.
XANTHUS TRANSCRIPTIONAL REGULATORY MECHANISMS
2004. Characterization of a Myxococcus xantbus mutant that is defective for adventurous motility and social motility. Microbiology 150:4085-4093. Lancero, H. L., S. Castaneda, N. B. Caberoy, X. Ma, A. G. Garza, and W. Shi. 2005. Analysing protein-protein interactions of the Myxococcus xantbus Dif signalling pathway using the yeast two-hybrid system. Microbiology 151:1535-1541. Li, S.-F., B. Lee, and L. J. Shimkets. 1992. csgA expression entrains Myxococcus xantbus development. Genes Dev. 6:401-410. Li, S.-F., and L. J. Shimkets. 1993. Effect of dsp mutations on the cell-to-cell transmission of CsgA in Myxococcus xanthus. J. Bacteriol. 175:3648-3652. Liu, X., and P. Matsumura. 1995. An alternative sigma factor controls transcription of flagellar class-I11 operons in Escherichia coli: gene sequence, overproduction, purification and characterization. Gene 164:8 1-84. Loconto, J., P. Viswanathan, S. J. Nowak, M. Gloudemans, and L. Kroos. 2005. Identification of the 04406 regulatory region, a developmental promoter of Myxococcus xanthus, and a DNA segment responsible for chromosomal position-dependent inhibition of gene expression. J. Bacteriol. 187:4 149-4162. Lonetto, M., M. Gribskov, and C. A. Gross. 1992. The u70 family: sequence conservation and evolutionary relationships. ]. Bacteriol. 174:3843-3849. Lonetto, M., K. Brown, K. Rudd, and M. Buttner. 1994. Analysis of the Streptomyces coelicolor sigE gene reveals the existence of a subfamily of eubacterial polymerase u factors involved in the regulation of extracytoplasmic functions. Proc. Natl. Acad. Sci. USA 91:7573-7577. Nakahigashi, K., H. Yanagi, and T. Yura. 1995. Isolation and sequence analysis of rpoH genes encoding d2homologs from gram negative bacteria: conserved mRNA and protein segments for heat shock regulation. Nucleic Acids Res. b3:4383-4390. Nariya, H., and S. Inouye. 2002. Activation of 6-phosphofructokinase via phosphorylation by Pkn4, a protein Ser/Thr kinase of Myxococcus xantbus. Mol. Microbiol. 46:13531366. Nariya, H., and S. Inouye. 2003. An effective sporulation of Myxococcus xantbus requires glycogen consumption via Pkn4-activated 6-phosphofructokinase. Mol. Microbiol. 49517-528. Nariya, H., and S. Inouye. 2005. Identification of a protein Ser/ Thr kinase cascade that regulates essential transcriptional activators in Myxococcus xanthus development. Mol. Microbiol. 58:367-379. Nariya, H., and S. Inouye. 2006. A protein Ser/Thr kinase cascade negatively regulates the DNA-binding activity of MrpC, a smaller form of which may be necessary for the Myxococcus xanthus development. Mol. Microbiol. 60~120.5-1217. Nielsen, M., A. A. Rasmussen, E. Ellehauge, A. TreunerLange, and L. Ssgaard-Andersen. 2004. HthA, a putative DNA-binding protein, and HthB are important for fruiting body morphogenesis in Myxococcus xantbus. Microbiology 150:2171-2183.
167
Ogawa, M., S. Fujitani, X. Mao, S. Inouye, and T. Komano. 1996. FruA, a putative transcription factor essential for the development of Myxococcus xantbus. Mol. Microbiol. 22:757-767. Otani, M., J. Tabata, T. Ueki, K. Sano, and S. Inouye. 2001. Heat-shock-induced proteins from Myxococcus xantbus. J. Bacteriol. 183:6282-6287. Otani, M., T. Ueki., S. Kozuka, M. Segawa, K. Sano, and S. Inouye. 2005. Characterization of a small heat shock protein, Mx Hsp16.6, of Myxococcus xantbus. ]. Bacteriol. 1875236-5241. Paget, M. S., and M. J. Buttner. 2003. Thiol-based regulatory switches. Annu. Rev. Genet. 37:91-121. Paul, R., S. Weiser, N. C. Amiot, C. Chan, T. Schirmer, B. Giese, and U. Jenal. 2004. Cell cycle-dependent dynamic localization of a bacterial response regulator with a novel di-guanylate cyclase output domain. Genes Dev. 18:715-727. Pollack, J. S., and M. Singer. 2001. SdeK, a histidine kinase required for Myxococcus xantbus development. J. Bacteriol. 183:3589-3596. Romeo, J. M., and D. R. Zusman. 1991. Transcription of the myxobacterial hemagglutinin gene is mediated by a d4-like promoter and a cis-acting upstream regulatory region of DNA. J. Bacteriol. 173:2969-2976. Rudd, K. E., and D. R. Zusman. 1982. RNA polymerase of Myxococcus xanthus: purification and selective transcription in vitro with bacteriophage templates. J. Bacteriol. 151:89-105. Ssgaard-Andersen, L., F. Slack, H. Kimsey, and D. Kaiser. 1996. Intercellular C-signaling in Myxococcus xanthus involves a branched signal transduction pathway. Genes Dev. 10:740-754. Srinivasan, B. S., N. B. Caberoy, G. Suen, R. G. Taylor, R. Shah, F. Tengra, B. S. Goldman, A. G. Garza, and R. D. Welch. 2005. Functional genome annotation through phylogenomic mapping. Nat. Biotechnol. 23:691-698. Srinivasan, D., and L. Kroos. 2004. Mutational analysis of the fruA promoter region demonstrates that C-box and 5-basepair elements are important for expression of an essential developmental gene of Myxococcus xantbus. J. Bacteriol. 186~5961-5967. Stathopoulos, A., and M. Levine. 2005. Genomic regulatory networks and animal development. Dev. Cell 9:449-462. Sun, H., and W. Shi. 2001a. Genetic studies of mrp, a locus essential for cellular aggregation and sporulation of Myxococcus xantbus. J. Bacteriol. 183:4786-4795. Sun, H., and W. Shi. 2001b. Analyses of mrp genes during Myxococcus xantbus development. J. Bacteriol. 183:67336739. Teintze, M., R. Thomas, T. Furuichi, M. Inouye, and S. Inouye. 1985. Two homologous genes coding for spore-specific proteins are expressed at different times during development of Myxococcus xanthus. J. Bacteriol. 163:121-125. Thony, B., and H. Hennecke. 1989. The -24/-12 promoter comes of age. FEMS Microbiol. Rev. 5:341-357. Tojo, N., S. Inouye, and T. Komano. 1993. The lonD gene is homologous to the lon gene encoding an ATP-dependent protease and is essential for the development of Myxococcus xanthus. J. Bacteriol. 175:4545-4549.
168 Tse, H., and R. E. Gill. 2002. Bypass of A- and B-signaling requirements for Myxococcus xanthus development by mutations in spdR. J. Bacteriol. 184:1455-1457. Ueki, T., S. Inouye, and M. Inouye. 1996. Positive-negative KG cassettes for construction of multi-gene deletions using a single drug marker. Gene 183:153-157. Ueki, T., and S. Inouye. 1998. A new sigma factor, SigD, essential for stationary phase is also required for multicellular differentiation in Myxococcus xanthus. Genes Cells 3:371-385. Ueki, T., and S. Inouye. 2001. SigB, SigC, and SigE from Myxococcus xanthus homologous to u32are not required for heat shock response but for multicellular differentiation. J. Mol. Microbiol. Biotechnol. 3:28 7-293. Ueki, T., and S. Inouye. 2002. Transcriptional activation of a heat-shock gene, lonD, of Myxococcus xanthus by a two component histidine-aspartate phosphorelay system. J. Biol. Chem. 277:6170-6 177. Ueki, T., and S. Inouye. 2003. Identification of an activator protein required for the induction of fruA, a gene essential for fruiting body development in Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 10053782-8787. Ueki, T., and S. Inouye. 2005a. Identification of a gene involved in polysaccharide export as a transcription target of FruA, an essential factor for Myxococcus xanthus development. J , Biol. Chem. 280:32279-32284. Ueki, T., and S. Inouye. 2005b. Activation of a developmentspecific gene, dofA, by FruA, an essential transcription factor for development of Myxococcus xanthus. J. Bacteriol. 187: 8504-8 5 06. Ueki, T., C. Y. Xu, and S. Inouye. 2005. SigF, a new sigma factor required for a motility system of Myxococcus xanthus. J. Bacteriol. 18753537-854 1. Viswanathan, K., P. Viswanathan, and L. Kroos. 2006a. Mutational analysis of the Myxococcus xanthus R4406 promoter region reveals an upstream negative regulatory element that mediates C-signal dependence. J. Bacteriol. 188:$15-524. Viswanathan, P., and L. Kroos. 2003. cis elements necessary for developmental expression of a Myxococcus xanthus gene that depends on C signaling. J. Bacteriol. 185:1405-1414.
REGULATORY MECHANISMS Viswanathan, P., M. Singer, and L. Kroos. 2006b. Role of uD in regulating genes and signals during Myxococcus xanthus development. J. Bacteriol. 188:3246-3256. Viswanathan, P., K. Murphy, B. Julien, A. G. Garza, and L. Kroos. 2007a. Regulation of dev, an operon that includes genes essential for Myxococcus xanthus development and CRISPRassociated genes and repeats. J. Bacteriol. 189:3738-3750. Viswanathan, P., T. Ueki, S. Inouye, and L. Kroos. 2007b. Combinatorial regulation of genes essential for Myxococcus xanthus development involves response regulator and a LysRtype regulator. Proc. Natl. Acad. Sci. USA 104:7969-7974. Ward, M., H. Lew, A. Treuner-Lange, and D. Zusman. 1998. Regulation of motility behavior in Myxococcus xanthus may require an extracytoplasmic-function sigma factor. J. Bacterial. 18056684675. Wu, S. S., and D. Kaiser. 1997. Regulation of expression of the pilA gene in Myxococcus xanthus. J. Bacteriol. 179:77487758. Xu, D., C. Yang, and H. B. Kaplan. 1998. Myxococcus xanthus sasN encodes a regulator that prevents developmental gene expression during growth. J. Bacteriol. 180:6215-6223. Yamazaki, H., Y. Ohnishi, and S. Horinouchi. 2000. An A-factor-dependent extracytoplasmic function sigma factor (sigma(AdsA)) that is essential for morphological development in Streptomyces griseus. J. Bacteriol. 182:4596-4605. Yang, C., and H. B. Kaplan. 1997. Myxococcus xanthus sass encodes a sensor histidine kinase required for early developmental gene expression. J. Bacteriol. 179:7759-7767. Yoder, D., and L. ICroos. 2004a. Mutational analysis of the Myxococcus xanthus R4400 promoter region provides insight into developmental gene regulation by C signaling. J. Bacteriol. 186:661-671. Yoder, D., and L. Kroos. 2004b. Mutational analysis of the Myxococcus xanthus a 4 4 9 9 promoter region reveals shared and unique properties in comparison with other C-signaldependent promoters. J. Bacteriol. 186:3766-3776. Yoder-Himes, D., and L. Kroos. 2006. Regulation of the Myxococcus xanthus C-signal-dependent 0 4 4 0 0 promoter by the essential developmental protein FruA. J. Bacteriol. 188~5 167-5 176.
Myxobacteria: Multicellularity and Differentiation Edited by David E. Whitworth 0 2008 ASM Press, Washington, D.C.
David E. Whitworth Peter J. A. Cock
Two-Component Signal Transduction Systems of the Myxobacteria
INTRODUCTION Two-Component Signal Transduction Systems Myxobacteria live in an ever-changing environment and therefore require mechanisms to couple perception of environmental change with appropriate behavioral responses. Such signaling pathways can take diverse fofms, but they often share certain features enabling their grouping into families. The largest family of prokaryotic signaling pathways are collectively described as two-component systems (TCSs). TCSs are found across the tree of life; however, they are most abundant in the prokaryotes, particularly in the eubacteria (Hoch, 2000). The number of TCSs found in different organisms can vary widely, but they are exceptionally abundant in the myxobacteria. The evolutionary success of TCS signaling pathways apparently stems from their adaptability to the regulation of diverse physiological processes, and this feature is illustrated well by the known TCSs of myxobacteria. A typical TCS consists of a histidine protein kinase and a response regulator protein. Both histidine kinases and response regulators contain domains of defined
10
function (Kofoid and Parkinson, 1988), and possession of these domains typically indicates that a protein is a member of a TCS. The histidine kinase contains an input domain (typically N-terminal) that senses a feature of the environment and a transmitter domain (typically C-terminal) that has kinase activity and contains a conserved histidine residue. The response regulator consists of a receiver domain, which includes a conserved aspartate residue, and an output domain, which mediates an appropriate response to an environmental signal (Fig. 1A). When the histidine kinase senses a particular environmental change or signal, it becomes active and autophosphorylates on its conserved histidine residue. The phosphokinase then engages in a protein-protein association with its partner response regulator and directly transfers its phosphoryl group to the conserved aspartate residue within the response regulator receiver domain (Fig. 1A). The genes for a histidine kinase and response regulator pair are often found adjacent to one another in the genome (Fabret et al., 1999; Mizuno, 1997), and encountering such paired genes in the genome is very good evidence that they act together to form a TCS. In
David E. Whitworth, Department of Biological Sciences, University of Warwick, Coventry, CV4 7AL United Kingdom. Peter J. A. Cock, MOAC Doctoral Training Centre, University of Warwick, Coventry, CV4 7AL United Kingdom.
169
1 70
PhoR
-
REGULATORY MECHANISMS
Phosphate Metabolism
PhoB Flagella Rotation
MCP Methylation
Initiation of Sporulation
/
Figure 1 Typical TCSs and phosphorelays. Domain architecture graphics obtained using the SMART tool (http:Nsmart.embl-heidelberg.de). Phosphotransfer reactions are indicted with arrows. (A) The PhoBR and CheAY TCSs of E. coli. PhoR contains an orthodox transmitter domain comprising a HisKA phosphotransfer domain and a HATPase domain and is activated in response to low extracellular phosphate concentrations. This leads to phosphorylation of the DNA-binding response regulator PhoB, which activates transcription of phosphate-scavenging genes. CheA possesses an unorthodox transmitter domain, which contains a phospho-accepting Hpt domain and HATPase domain, with a vestigial form of the HisKA domain (lacking a phospho-accepting histidine residue) retained as a dimerization domain. Sensing of attractantshepellants by the Tar MCP activates CheA, which phosphorylates CheB and CheY. (B)The phosphorelay regulating initiation of sporulation in B. subtilis. SpoOF is phosphorylated by at least two kinases, including KinA and KinB. Phosphoryl groups are passed from SpoOF to SpoOA via a phosphotransfer protein, SpoOB, which is phosphorylated on a histidine residue and structurally resembles both Hpt domains and dimeric H-box/HisKA domains. SpoOA can also be phosphorylated directly by a third histidine kinase, KinC, and when phosphorylated promotes sporulation through changes in gene expression. For details see Stock et al. (1989) and Fabret et al. (1999).
a small proportion of cases the genes for partner histidine kinases and response regulators can be found fused together, resulting in a single polypeptide with both receiver and transmitter domains, which is then known as a hybrid kinase.
Myxobacterial Two-Component Systems Prior to the determination of the genome sequence, a large number of the TCS proteins of Myxococcus xanthus had been identified and characterized (Table 1). Most of these proteins are known to regulate events
Table 1 TCSs of M . xantbusa RR AglZ FrzS DotR
Family
RR domains
HK
Family
Hybrid?
HK Domains
TM?
P’ase
Bi
Orphan Orphan R>
A-motility S-motility Development
T
Bi
redCDEF
Development
R, T
T, R, R, R
Bi Bi
H>R> H>R> Orphan
T
Ntr
redCDEF
Yes
Tu, Chew, R Tu, Chew Tu, Chew, R
Mono Mono Mono
Orphan R>H> R>K> H>R>
Yes
Tu, Chew, R
Mono
H>R> Orphan
Development Development A-motility Development Development S-motility S-motility A- and S-motility Development Development
actAB
Development
CheY
R, coiled coil R, coiled coil R
TodK
PhoR
PAS, PAS, T
RedF
CheY
R
RedE
NtrB
MXAN2671 MXAN0732 FrzZ RedD FruA CheY4 DifD FrzG
CheY CheY FrzZ FrzZ NarL CheY CheY CheB
R R R, R R, R R, -HTH-LUXR R R R, Me esterase
AsgA RodK
? ?
RedC
NtrB
CheA4 DifE FrzE
CheA CheA CheA
Yes
CheB3 CrdA
CheB NtrC
R, Me esterase R, AAA, HTH-8
CheA3
CheA
ActB
NtrC
R, AAA, HTH-8
FrgC MrpB Nlal Nla4 Nla6 Nla23 Nla24
NtrC NtrC NtrC NtrC NtrC NtrC NtrC
R, AAA, H T H 8 R, AAA, H T H 8 R, AAA, HTH-8 R, AAA, HTH-8 R, AAA, HTH-8 R, AAA, H T H 8 R, AAA, HTH-8
FrgB MrpA MXAN5852
NtrB NtrB NtrB
TM
MXAN4043 MXANS778 MXAN7439
NtrB NtrB NtrB
TM TM
Nla28 PilR SasR SpdR HsfA
NtrC NtrC NtrC NtrC NtrC
R, AAA, HTH-8 R, AAA, H T H 8 R, AAA, HTH-8 R, AAA, H T H 8 R, AAA, HTH-8
MXANll66 PilS Sass SpdS HsfB
PhoR NtrB NtrB NtrB NtrB
MXAN5366
PleD
R, GGDEF
K>R> K>R> K>R> K>R> mxan5366> hsfBA> mxan53 66>
R, GGDEF
hsfBA> actAB
ActA
PleD
Locus phenotype
Genes
Yes Yes
TM
TM TM Yes
T T PutP, PAS, T
Ntr Ntr Bi
T T HAMP, T
Bi Bi Ntr
T PAS, T HAMP, T PAS, GAF, T R, T
Bi Ntr Ntr Ntr Bi
K>R> K>R> K>R> Orphan K>R> K>R> K>R>
?
Development S-motility Development Development S-motility A- and S-motility Development S-motility Development Development Heat shock
Reference Yang et al., 2004 Ward et al., 2000 Rasmussen and S.-Andersen, 2003 Higgs et al., 2005 Li and Plamann, 1996 Rasmussen et al., 2005 Trudeau et al., 1996 Higgs et al., 200.5 Ellehauge et al., 1998 Vlamakis et al., 2004 Yang et al., 1998 McCleary and Zusman, 1990 Kirby and Zusman, 2003 Kirby and Zusman, 2003 Gronewold and Kaiser, 2001 Cho et al., 2000 Sun and Shi, 2001 Caberoy et al., 2003 Caberoy et al., 2003 Caberoy et al., 2003 Caberoy et al., 2003 Lancer0 et al., 2004 Caberoy et al., 2003 Wu and Kaiser, 1997 Guo et al., 2000 Hager and Gill, 2001 Ueki and Inouye, 2002
?
Development
Gronewold and Kaiser, 2001
(Continued)
Table 1 TCSs of M. xanthusa (Continued)
RR
Family
RR domains
HK
Family
Hybrid?
HK Domains
TM?
P’ase
Genes
Locus phenotype
Reference
Ntr
Orphan
Development
Cho and Zusman, 1999a
Ntr
Orphan
Development
Lee et al., 2005
TM
FHA, PAS, PAS, T, R MASE1, PAS, T, R LBD,T, R
Ntr
Orphan
Kimura et al., 2003
EspA
NtrB
Yes
EspC
NtrB
Yes
TM
MokA
NtrB
Yes
PhoPl
PhoB
R, Trans-regC
PhoRl
PhoR
TM
T
Ntr
R>K>
Osmotic shock Development
PhoP2
PhoB
R, Trans-regC
PhoR2
PhoR
TM
T
Bi
K>R>
Development
PhoP3
PhoB
R, Trans-reLC
PhoR3
PhoR
TM
T
Bi
K>R>
Development
PhoP4
PhoB
R, Trans-reLC SdeK AsgD
PhoR
Bi Bi
Orphan Orphan asgD >
Development Development Development
MXAN6994
PhoR
PAS, T R,GAF, GAF, T T
Bi
asgD >
?
Yes
Carrero-Lerida et al., 2005 Moraleda-Muiioz et al., 2003 Moraleda-Muiioz et al., 2003 Pham et al., 2006 Pollack and Singer, 2001 Cho and Zusman, 1999b
mxan6994>
?
mxan6994>
SocD CyaB
R.CYCc
PhoR
PAS, GAF, T
Bi
Complex
Development
S. aurantiaca
?
Shimkets, 1999 Coudart-Cavalli et al., 1997
dExperimentallycharacterized TCS proteins of M. xanthus, grouped according to response regulator family (based on domain organization). Partner histidine kinases were assigned to response regulators on the basis of gene adjacency, unless additional evidence was available. Histidine kinase family membership is based on phylogenetic analyses, and the presence of transmembrane helices was determined using TMHMM v2.0 (Sonnhammer et al., 1998). The ability of a histidine kinase to act as a phosphatase (P’ase) was predicted using the method of Alves and Savageau (2003) as described in the text. In descriptions of domain architecture, some domain names were abbreviated as follows: R, receiver; T, orthodox transmitter; Tu, unorthodox transmitter; LBD, ligand binding domain. Arrowheads are used in descriptions of gene organization to represent relative directions of transcription. H, hybrid kinase; R, response regulator; K, histidine kinase.
10. TWO-COMPONENT SIGNALTRANSDUCTION SYSTEMS during fruiting body development (including regulation of motility), although several have been shown to act during vegetative growth. This situation contrasts with other bacteria, where most TCSs maintain homeostasis, and a minority are involved in complex regulated processes such as sporulation or cell cycle progression (for a review see Hoch and Silhavy, 1995). Such a distinction may reflect a bias in myxobacterial research (which largely focuses on development). However, myxobacterial TCSs are known which have roles in both vegetative growth and in development (Ueki and Inouye, 2002; Kimura et al., 2001). In a recent study a large number of M. xanthus NtrC-like response regulator genes were inactivated and their phenotypes were assessed (Caberoy et al., 2003). Of the 28 genes mutated, only 8 gave developmental and/or motility phenotypes, suggesting that most TCSs of M. xanthus either are not involved in development/motility or are redundant under laboratory conditions. The recent availability of genome sequences of four myxobacteria has enabled a comparative genomic analysis of TCS genes in myxobacterial genomes. According to an automated search protocol (Cock and Whitworth, 2007), the M. xanthus genome encodes 276 TCS proteins, and a similar number can be found in the genomes of Sorangium cellulosum and Stigmatella aurantiaca, with nearly 200 TCS proteins encoded by the smaller Anaeromyxobacter dehalogenans genome. For comparison, Escherichia coli and Bacillus subtilis possess 62 and 70 TCS proteins, respectively (Mizuno, 1997; Fabret et al., 1999).Figure 2 shows the number of TCS genes as a function of genome size for 228 bacterial genomes, including four myxobacteria. All four myxobacterial genomes possess significantly more TCS genes than would be predicted on the basis of their genome size alone and thus possess a high IQ as defined by Galperin (2005). Table 1summarizes the TCSs of M. xanthus that have been characterized experimentally. Pertinent features of each system are described, including their genomic organization, function, domain architecture, and predicted localization. Partnerships between histidine kinase and response regulators in Table 1 are described on the basis of gene adjacency unless further evidence is available in the literature. The large number of characterized M. xanthus TCS proteins precludes an exhaustive description of each system; however, general features of myxobacterial TCS properties can now be discerned and their description forms the basis for the remainder of this chapter.
Receiver and Transmitter Domains Amino acid sequences of transmitter and receiver domains are highly conserved, and several critical motifs
173 Sorangium cellulosum Stigmatella aurantiaca . . Myxococcus xanthus ....... . .
....
400
.
... ... ...... .. ,.
...
Anaeromyxobacter dehalogenans :,
.... ...
... ..
1
350
.. ...
..
. '... ..... ... ... ..... .... ... ... . .
300
d In 2
250
Nostoc
200
Geobactee
*-
2
n
E
150 100
3
50 0
r
0
1
2
4
6
8
10
12
14
Genome Size (Mbp)
Figure 2 Numbers of TCS genes found in different genomes as a function of genome size. Trend lines are shown for all bacteria (gray) and for four myxobacteria (black). The myxobacteria all possess an exceptionally large complement of TCS genes, given their genome size. Nostoc (PCC7120) and Geobacter sulfurreducens (PCA) also have relatively large numbers of TCS genes.
and residues within them have been identified (Parkinson and Kofoid, 1992). Receiver domains are characterized by a series of acidic residues and a lysine residue (D12, D13, D57, and K109 of E. coli CheY), while orthodox transmitter domains possess a series of sequence motifs called the H, N, D, F, and G boxes. The H box is also referred to as a HisKA domain (histidine kinase phosphoacceptor), while the N, D, F, and G boxes are together described as an HATPase domain (histidine kinase-type ATPase). Transmitter domains operate as dimers, with dimerization mediated by the H-box/HisKA region and autophosphorylation occurring in trans (for a review see Wolanin et al., 2002). Nearly all M. xanthus receiver domains (described in Table 1)contain the defining series of amino acids as found in E. coli CheY (D12/13, D57, and K109). However, ActA, AglZ, FruA, and FrzS contain only a single acidic residue equivalent to CheY D12/13 (FrzG lacks both acidic residues), while AglZ, FrzS, and MXAN5366 lack the phosphorylated aspartate equivalent to CheY D.57 (AglZ and MXAN5366 also lack a basic residue equivalent to CheY K109). Lack of a phospho-accepting aspartate residue suggests that the activity of AglZ, FrzS, and MXAN5366 might not be regulated by phosphorylation,
REGULATORY MECHANISMS
174 while the lack of conserved residues in ActA, FrzG, and FruA suggests that they might not be able to become phosphorylated either. None of these proteins have been shown to phosphorylate in vitro, though genetic evidence suggests that the putative phospho-accepting aspartate residue of FruA is required for function (Ellehauge et al., 1998).AglZ and FrzS share the same domain architecture, are both isolated in the genome, have no known partner kinase (although FrzS is probably regulated by the histidine kinase FrzE), and are both implicated in regulating directed motility. FrzS has recently been shown to dynamically relocalize from pole to pole during cell reversals (Mignot et al., 2005), and it would be interesting to determine whether AglZ exhibits similar dynamic properties. It has been proposed (Yang et al., 2004) that an alternative aspartate residue found in both FrzS and AglZ might be phosphorylated instead of the normal residue, corresponding to position 52 of CheY, instead of position 57. However, position 52 does not encode an aspartate residue in MXAN5366 (the other M . xanthus receiver domain lacking an aspartate at position 57), and by superimposing on known structures of receiver domains (for example, Feher et al., 1997),residue 52 would be predicted to lie on the face of the receiver domain opposite to position 57. It is tempting to speculate on possible alternative mechanisms responsible for regulating the activity of AglZ, FrzS, and MXAN5366. Phosphotransfer between histidine kinases and response regulators requires appropriate proteinprotein interactions, and it is possible that such interactions could be regulatory even in the absence of phosphotransfer (altering the conformation of the receiver domain to an "active" form equivalent to that typically resulting from phosphorylation). Various lines of evidence have allowed suggestions of potential partners for all three proteins; FrzE for FrzS (Bustamante et al., 2004), MglA for AglZ (Yang et al., 2004), and HsfB for MXAN.5366 (predicted by gene adjacency), two of which are histidine kinases (MglA is a GTPase). An alternative possibility is that these unusual proteins have evolved from response regulators and are no longer regulated by phosphorylation in the fashion suggested by the TCS paradigm. Phylogenetic analysis of receiver domains suggests that AglZ and FrzS are paralogues and that S. aurantiaca, S. cellulosum, and A. dehalogenans each possess orthologues of AglZ and FrzS. It is interesting that aglZ of S. cellulosum lacks a coiled-coil domain but is fused to a gene encoding a transmitter domain, which is particularly similar to SocD and MXAN5990 of M. xanthus. In orthodox transmitter domains, the H-box/HisI
domains of M. xanthus possess recognizable H boxes including the phospho-accepting histidine residue, while most transmitters also contain recognizable N, D, F, and G boxes. The exceptions are MrpA, which lacks N and D boxes; MXANll66, which does not have an obvious N box; and MXAN4043, which lacks N, D, F, and G boxes. It is known that some sequence boxes are dispensable for ATPase activity (Wolanin et al., 2002), so MrpA, MXAN1166, and MXAN4043 may still act as kinases, which seems likely given that all three genes are found adjacent to genes encoding NtrC homologues (mrpB, nla28, and nla6, respectively) and are themselves members of the NtrB family phylogenetically. Unorthodox transmitter domains are uncommon variants of transmitter domains and are exemplified by CheA of E. coli (Fig. 1A).They possess typical HATPase domains, but the H-box/HisKA region lacks the characteristic histidine residue and functions solely as a dimerization domain. Phosphorylation occurs instead on a histidine residue in an N-terminal Hpt domain, which structurally resembles a dimeric H-box/HisKA domain.
TCSs OF M . XANTHUS TCSs can be consistently grouped into particular subfamilies by applying several different assessment criteria, including gene organization, domain architecture, and phylogenetic relationships. The major families of TCSs are usually named after archetypal family members from E. coli. Thus, members of the four most common families are similar to CheAY, PhoBR, NtrBC, and NarXL of E . coli. Most myxobacterial TCS proteins fall into one of the four major TCS families; however, there are a remarkable number that do not (8 of the 33 described in Table 1).Common and unusual TCS response regulator domain architectures and their characterized members from M . xanthus are shown in Fig. 3 .
CheAY-LikeTCSs The M. xanthus genome sequence suggests the presence of eight TCSs resembling the chemotaxis system CheAY (Fig. 1A). The CheAY system of enteric bacteria regulates rotation of the flagella in response to attractive or repellent chemicals. The histidine kinase CheA causes phosphorylation of two response regulators, CheY and CheB. CheY and CheY-P interact with the flagellar motor to influence the direction of rotation, while CheB-P transiently alters the methylation state and sensitivity of chemoreceptors to attractants and repellants (e.g., Tar in Fig. 1A). CheA homologues can be distinguished from
10. TWO-COMPONENT SIGNALTRANSDUCTION SYSTEMS
I
Family
I
Domain Architecture
Examples in A4 xanthus
CheY
CheY4, D i m , DotR, RedF
PhoB
PhoP1, PhoP2, PhoP3, PhoP4
NarL
FruA
NtrC
ActB, CrdA, FrgC, HsfA, MrpB, Nlal, Nla4, Nla6, Nla23, Nla24, Nla28, PilR, SasR, SpdR
CheB
CheB3, FrzG
PleD
ActA
FrzZ
FrzZ, RedD
175
I
Figure 3 Domain architectures of response regulator families. Graphics of domain organization were obtained using the SMART tool (http:Nsmart.embl-heidelbergde). Families are named after archetypal members, and examples from M. xalzthus are indicated. fl
other histidine kinases as they contain unorthodox transmitter domains (Fig. 1A). CheY homologues possess a receiver domain without an output domain (Fig. 3 ) and are often found fused to CheA as a hybrid kinase (e.g., M. xanthus FrzE, CheA3, and CheA4). Four of the CheAY-like TCSs have been characterized in M. xanthus: the Che3, Che4, Dif, and Frz systems (Table 1).These systems regulate, respectively, timing of development, S-motility, fibril exopolysaccharide production, and Aand S-motility (Kirby and Zusman, 2003; McBride et al., 1989; McCleary and Zusman, 1990; Vlamakis et al., 2004; Yang et al., 1998). Intriguingly, in addition to regulating motility behavior, the CheAY-like systems of M. xanthus are thought to regulate gene expression through NtrC-like response regulators. The CheA3 histidine kinase has been shown to interact with the NtrC homologue CrdA, which is encoded close to cheA3 in the M. xanthus genome (Kirby and Zusman, 2003). NtrC homologues have DNA-binding output domains, and it is proposed that
the Che3 pathway inhibits developmentally regulated gene expression in the vegetative state, by acting through CrdA. A similar scenario is suggested by an interaction observed between the histidine kinase of the Dif pathway DifE, and another NtrC homologue, SpdR (referred to as Nla19 by Lancer0 et al., 2005). The adoption of chemotaxis-like signaling pathways to regulate gene expression appears to be unique to the myxobacteria and bears testimony to the versatility and flexibility of the basic TCS architecture. PhoBR-Like TCSs PhoBR-like systems are characterized by response regulators that possess a Trans-reg-C DNA-binding output domain (Fig. 3). In E. coli, the histidine kinase PhoR responds to low extracellular phosphate concentrations by phosphorylating the response regulator PhoB (Fig. 1A). PhoB contains a DNA-binding domain, and phosphorylation leads to altered gene expression-most notably of the pstSCAB and phoU genes, which encode
REGULATORY MECHANISMS
176 a high-affinity phosphate transporter and a phosphatedependent regulator, respectively. The laboratory of J. Muiioz-Dorado has described three Pho family TCSs in M . xanthus, which each partially regulate expression of overlapping sets of developmental phosphatase activities (Carrero-LCrida et al., 2005; Moraleda-Muiioz et al., 2003). Vegetative expression of the PhoPRl system is increased by extracellular phosphate, suggesting that it regulates phosphate uptake in addition to developmental phosphatase expression. This inference is supported by the close proximity of the phoPRl genes to the M. xanthus pstSCAB and phoU genes (Carrero-LCrida et al., 2005). The PhoRP2 and PhoRP3 TCSs are extremely similar in terms of sequence identity and appear to be the most recently duplicated of the contemporary TCSs in the M . xanthus genome. Indeed the S. aurantiaca and S. cellulosum genomes each encode only one set of orthologues of the PhoRP2/3 TCSs. The PhoRP2 and PhoRP3 systems are so similar that there are no differences in the PhoR2 and PhoR3 H-box/HisKA sequences or in the sequences of the PhoP2/PhoP3 DNA-binding helices, though the N-terminal regions of the PhoR2 and PhoR2 kinases exhibit similarity of only 55% (MoraledaMuiioz et al., 2003). It is predicted that the PhoR2 and PhoR3 proteins would interact with the PhoP2 and PhoP3 response regulators with equal affinity and that the PhoR2 and PhoR3 proteins would be expected to bind to identical stretches of DNA, regulating expression of the same genes, though possibly in response to different signals. Surprisingly the phenotypes of phoRP2 and phoRP3 mutants are not identical, suggesting that significant functional divergence of the two TCSs has occurred (Moraleda-Muiioz et al., 2003). A fourth PhoB-like response regulator (PhoP4) has recently been identified, encoded by a gene immediately upstream of the pstSCAB and phoU phosphate uptake genes. PhoP4 is required for proper expression of all developmental phosphatase activities and regulates expression of the pstSCAB and phoU genes (Pham et al., 2006). This observation suggests that PhoP4 probably acts upstream of the other PhoBR-like TCSs in the regulation of developmental phosphatase activities. Yeast two-hybrid assays detected a strong interaction between PhoR2 and PhoP4, suggesting that PhoR2 is a partner histidine kinase of PhoP4, but phenotypic differences between phoRP2 and phoP4 mutants imply that further histidine kinases of PhoP4 are yet to be identified. Mutants of phoPRl, phoRP2, phoRP3, and phoP4 all exhibit developmental defects and aberrant phosphatase expression. It is not currently understood how phosphatase
expression, phosphate acquisition, and development are related in M. xanthus. With four response regulators and three histidine kinases implicated, the M. xanthus Pho regulon appears to be the most complicated example yet described. In the majority of organisms one TCS suffices for the regulation of phosphatase expression and phosphate uptake-why then does M . xanthus require such an increase in complexity? Presumably the predatory and social aspects of myxobacterial physiology restrict the availability of phosphate to particular chemical forms and specific periods of the myxobacterial life cycle, which would then require a sophisticated ability to sense and assimilate phosphate. Alternatively, developmentally regulated phosphatase activities may have important roles during development quite apart from phosphate acquisition, and any link between phosphate acquisition and phosphatase expression may be purely an evolutionary relic.
NtrBC-Like TCSs RNA polymerase holoenzyme containing the alternative sigma factor d4 requires activator proteins to initiate promoter open complex formation (Sasse-Dwight and Gralla, 1990).Activator proteins possess an AAA ATPase domain with a HTH-8 DNA-binding domain (Fig. 3) and often also contain an N-terminal receiver domain (exemplified by NtrC of E. coli [Fig. 31).It has been shown that uniquely, os4 is essential in M . xanthus and that M . xanthus encodes a large number of NtrC homologues (Keseler and Kaiser, 1997; Jakobsen et al., 2004). Caberoy et al. (2003)undertook a global mutational analysis of os4-dependentactivator genes and found that of 28 mutants tested, 8 exhibited developmental defects. Of those eight genes, seven were ntrC homologues; nlal, nla4, nla6, nlal9 (spdR),nla23, nla24, and nla28. With seven previously identified ntrC homologues, the Caberoy et al. (2003)study has increased the number of characterized NtrC homologues in M. xanthus to 14 (Table l),most of which are known to regulate motility and/or development. The genome sequence suggests that M. xanthus encodes 28 NtrC homologues. This is an extremely large number compared to most other organisms: E . coli possesses just four NtrC homologues, while B. subtilis has none (Mizuno, 1997; Fabret et al., 1999). Why has M. xanthus expanded its complement of os4-dependent NtrC homologues, in preference to other families of TCS, which are generally found in numbers similar to those of other bacteria? That most NtrC homologues have roles in development and/or motility suggests that the use of enhancer binding proteins was specifically expanded during the evolution of multicellular development. Many enhancer binding proteins have N-terminal
10. TWO-COMPONENT SIGNALTRANSDUCTION SYSTEMS forkhead-associated domains (FHA) instead of receiver domains, including 12 in M . xanthus (Jelsbak et al., 2005). FHA domains bind phospho-threonine residues, and M . xanthus is also known to possess large numbers of Ser/Thr protein kinases associated with development. This suggests an extensive integration of the d4and Ser/ Thr kinase regulons in M . xanthus development. Presumably a core role for d4regulation in development also required appropriate integration with TCS networks, achieved by an expansion in the number of NtrC homologues.
NarXL-Like TCSs NarL family response regulators are characterized by a C-terminal DNA-binding domain similar to that of GerE and LuxR (Fig. 3). There are a surprisingly small number of NarXL-like TCS proteins in M. xanthus (four response regulators and two histidine kinases predicted in the genome); however, one of the prime regulators of development, FruA, is a member of this family. FruA is a critical control point in the progression of multicellular development. The f ~ u A gene is induced by starvation, an effect mediated by the A-signal (Ellehauge et al., 1998). It is thought that FruA is phosphorylated in response to C-signal, and as the activity of the C-signal-dependent positive-feedback loop increases, FruA-P progressively triggers population aggregation and then sporulation (Ogawa et al., 1996; Ellehauge et al., 1998).No histidine kinases have yet been identified in the literature as potential partners for FruA. In some ways the activity of FruA is analogous to that of SpoOA in the B. subtilis sporulatidn phosphorelay (Fig. lB), with both proteins serving as master checkpoints during cellular differentiation. Phosphorylation of SpoOA is under the control of several histidine kinases (directly and indirectly), and it is highly likely that the same is true for FruA, making identification of the FruA kinase(s) a challenging, but important task for the future.
PHOSPHORELAYS In addition to the typical configuration (Fig. lA), TCSs can also exist in an expanded form called a phosphorelay, exemplified by the sporulation phosphorelay of B. subtilis (Burbulys et al., 1991) (Fig. 1B). In the Spo pathway, histidine kinases KinA and KinB phosphorylate the response regulator SpoOF (consisting of a single receiver domain), which then sequentially passes the phosphoryl group to a histidine residue of SpoOB and then to a terminal response regulator SpoOA (Fig. 1B). SpoOB structurally resembles both H-box/HisKA and Hpt phosphotransfer domains,
177
though it shares no sequence relatedness to either domain. Phosphorelays of other organisms typically utilize an Hpt domain for phosphotransfer rather than SpoOB, and in such cases, the Hpt domain can be found as a free protein, or fused to a hybrid histidine kinase that also contains the initial transmitter domain and the first receiver domain. In M . xanthus, the only Hpt domains encoded in the genome are found as part of CheA-like histidine kinases, suggesting that the organism does not possess conventional phosphorelays (it also lacks any homologues of spoOB). However, there are features of TCS signaling in M . xanthus that are suggestive of complex TCS signaling pathways akin to phosphorelays. Most response regulators that are the final recipients of phosphorylation in a TCS exert their effects through C-terminal output domains. The most notable exceptions are the CheY homologues, which possess no output domain (Fig. 3) but act through protein-protein interactions with a target protein. However, response regulators consisting solely of receiver domains are also found within phosphorelays (for example, SpoOF in Fig. 1B). In these cases the response regulator does not need an output domain, as it merely acts as a shuttle to transfer phosphoryl groups between histidine residue phospho-acceptor proteins. There are 45 predicted response regulators in the M . xanthus genome that consist of a single receiver domain (including CheY4, DifD, DotR, MXAN0732, MXAN2671, and RedF). Of the 45, 4 are found within chemotaxis gene clusters and therefore probably represent true CheY homologues, leaving 41 non-CheY homologues. Most organisms have very few single receiver domain response regulators that are not CheY homologues: E. coli has none, B. subtilis has two including SpoOF, and Synechocystis PCC6803 has seven at most (Mizuno, 1997; Fabret et al., 1999; Mizuno et al., 1996).Does the large number of these proteins in M . xanthus suggest their involvement within phosphorelays, and if so, which other proteins are acting as the phosphohistidine shuttle proteins analogous to Hpt domains/SpoOB? Intriguingly, the M. xanthus genome has four genes encoding proteins which possess H-bodHisKA domains without the associated HATPase domains (including two that are located adjacent to other TCS genes). Potentially these could act as phosphoryl group shuttle proteins in phosphorelays. If M. xanthus does not use single domain response regulators in phosphorelays, then presumably it has adopted the use of response regulators that operate by protein-protein interaction with effector proteins, rather than DNA-binding, to an extent not seen in other organisms.
REGULATORY MECHANISMS
1 78 The M. xanthus genome also contains genes for four response regulators that contain two receiver domains with no output domains, including FrzZ and RedD (Table 1).Such domain architecture is not found in any organism outside the myxobacteria. FrzZ is required for A-motility (Trudeau et al., 1996), and the two receiver domains of FrzZ have been shown to interact with different cellular proteins. The first receiver domain interacted with an ABC transporter in a yeast two-hybrid assay (Ward et al., 1998a), while the second domain interacted with an extracytoplasmic function-type sigma factor (Ward et al., 1998b). FrzZ is probably phosphorylated by FrzE, but how FrzZ phosphorylation affects its behavior and the nature of its relationship with FrzE are not clear. A second protein of 211. xanthus consisting solely of two receiver domains is RedD, which affects the timing of development (Higgs et al., 2005). The redD gene is found within a cluster of four contiguous TCS genes, redC through redF (redC-F). RedC and RedE are both histidine kinases: RedD contains two receiver domains, while RedF consists of a single receiver domain (Table 1).
In gene organization, the four red genes resemble a typical phosphorelay, except that the SpoOF equivalent has duplicated receiver domains (Fig. 4). Epistasis analysis suggests that the RedCDEF proteins form a single linear signaling pathway akin to a phosphorelay, with RedCD acting upstream of RedEF. This was supported by yeast two-hybrid assay data, which demonstrated RedC-D, RedD-E, and RedE-F interactions (Higgs et al., 2005). However, it appears that RedC and RedE interact with different receiver domains of RedD, suggesting that information flow through the Red pathway is not as simple as the sequential transfer of a single phosphoryl group. It is possible that the phosphorylation state of one RedD receiver domain affects the ability of the other domain to act as a substrate for RedE phosphotransfer, modulating phosphotransfer from RedE to RedF. If the Red pathway does operate using a phosphoryl group shuttle mechanism, then presumably the H-box/HisKA region of RedE would receive phosphoryl groups from RedD before passing them on to RedF, as well as becoming phosphorylated by the RedE HATPase domain.
,
h
Transmitter Domain
MXAN229-30
n RedC-F
n
I
n
Receiver Domain
n
/ I /
/,
n
n [\
MXAN7362-4
DV
MXAN4444-5
H
D V D
D
b
H
D
D I /
I
I
D
H
D D /
H
MXAN6734-5
Figure 4 Complex TCS gene clusters of M. xanthus. Genes are represented by arrows pointing in the direction of transcription. Domains encoded within each gene are shown as SMART graphics (http://smart.embl-heidelberg.de), with conserved histidine and aspartate residues predicted to be involved in phosphotransfer indicated below each gene.
10. TWO-COMPONENT SIGNALTRANSDUCTION SYSTEMS GENOME CONTEXT Gene Organization Information regarding the partnerships between TCS proteins can be inferred by assessing the relative location of their genes. For typical TCSs, the genes encoding partner histidine kinases and response regulators are found adjacent in the genome. However, often a single TCS gene is found isolated in the genome (orphaned) or multiple TCS genes are found together in complex gene clusters. In complex gene clusters, the genomic organization is sometimes reminiscent of the path of phosphotransfer (as appears to be the case for the Red pathway), but this is not always the case. The physical distribution of TCS genes found in the M. xanthus genome appears to be random (Fig. 5). Some regions of the genome seem to have a particularly high density of TCS genes, particularly at -5.2 Mbp (including the fm genes) and at -8.5 Mbp (including asgD, espC, dotR, socD, and todK). There also appears to be one large region of relatively low TCS gene density, with a notable coding strand bias
1 79
(from -1.5 to -2.8 Mbp). The identified TCS genes of M. xanthus do not generally appear to be clustered; however, there are regions which appear to be particularly densely occupied (at around 8.6 and 1.3 Mbp in Fig. 5). Of the characterized TCSs described in Table 1 the majority of TCS genes are found in pairs, with one histidine kinase (often a hybrid kinase) and one response regulator gene (40 of 63 genes, 6 3 % ) ,and it can be assumed that such pairs constitute an entire TCS. The gene pairs are carried on the same strand of DNA (with the exception of todKldotR) and are probably expressed as polycistronic mRNAs. Of the remaining TCS genes described in Table 1, 11 are orphaned, while 12 lie in complex gene clusters or in pairs of two histidine kinase or two response regulator genes. In these cases it is difficult to identify signaling partnerships from genome locality, although some orphan genes encode hybrid kinases containing single transmitter and receiver domains, presumably encoding a TCS within a single polypeptide (EspA, EspC, and MokA). Several key regulators of motility/
8 Mbp
7 Mbp
pilRS, nla23
MbP
Myxococcus xanfhus 9,139,763 bp
rn
nla4
phoPR1 phoP4
8 Mbp
4 Mbp Figure 5 A map of the M. xanthus genome showing the location of TCS genes. The origin of the genome is defined as the start of the dnaA gene (position 1).Inner and outer rings represent genes coded on the + and - strands, respectively. Figure produced using GenomeDiagram (Pritchard et al., 2006).
REGULATORY MECHANISMS
180 Table 2 Organization of TCS genes of M. xanthus and other bacteriag Organism
E. coli B. subtilis Synechocystis PCC6803 S. coelicolor M. xanthus
No. (%) orphan
No. (%)
No. (%)
paired
complex
8 (13) 9(13) 52(60)
42 (68) 60(85) 22(26)
12 (19) 2(3) 12(14)
192
55 (29)
276
105 (38)
112 (58) 94 (34)
25 (13) 77 (28)
Total TCS 62 71 86
“TCS genes were classified as either orphan (no other TCS gene within 5 kbp), paired (one histidine kinase gene adjacent to one response regulator gene with no other TCS genes within 5 kbp), or in complex gene clusters (at least two TCS genes within 5 kbp of one another, but not paired). Particularly large numbers or proportions of TCSs in different categories are highlighted in bold. Numbers quoted for nonmyxobacteria derived from Fabret et al. (1999),Mizuno (1997), Mizuno et al. (1996),and Hutchings et al. (2004).
development are encoded by orphaned genes, including AglZ, FruA, FrzS, FrzZ, PhoP4, and SdeK, and in each case partner TCS proteins remain to be identified conclusively (although FrzS and FrzZ are probably phosphorylated by FrzE, while PhoP4 is probably phosphorylated by PhoR2). A survey of the TCS genes in the M. xanthus genome showsthat 105areorphaned (38%),94arepaired (34%), while 77 (28%) lie in complex gene clusters (Table 2). It is interesting that the proportion of paired genes is much higher among the characterized TCS genes described in Table 1 than is found in the genome sequence, perhaps suggesting that in general paired systems give stronger phenotypes than orphanedkomplex TCSs. The number pf TCS genes found in complex clusters is unparalleled by any currently sequenced bacterial genome.
Synteny of TCS Genes In most organisms, the relative order of paired histidine kinase and response regulator genes in pairs is maintained within particular TCS families. For instance in B. subtilis, every PhoBR-like TCS has the response regulator gene upstream of the histidine kinase gene, while every NarXL-like TCS has the gene for the histidine kinase preceding that of the response regulator (Fabret et al., 1999). In E. coli the same general trends are observed, with the vast majority of PhoBR-like TCSs having response regulator genes upstream of the histidine kinase gene (Mizuno, 1997). In the paired NtrBC-like TCS genes of M. xanthus, synteny is strongly maintained, with all paired examples in Table 1 having the histidine kinase gene 5 ’ relative to that of the response regulator (only 1of 21 NtrBC-like TCS gene pairs in the genome has the inverse orientation). The PhoBR-like systems exhibit less
conservation of synteny. Of 10 paired phoB R-like gene pairs in the M. xanthus genome, 3 are found with the histidine kinase gene upstream of the response regulator gene (including phoPRZ), while 7 have the inverse orientation (includingphoRP2 and phoRP3). The significance of such observations is uncertain; however, strong maintenance of synteny within a TCS family may reflect relatively recent lineage-specific gene duplication. Thus, the notable maintenance of synteny within the ntrBC-like family supports the notion of myxobacterium-specific amplification of that TCS family, as described above.
Gene Neighborhood The tendency for genes of related biological function to be housed together in bacterial genomes can sometimes be exploited to suggest potential functions for regulatory genes. A good example previously mentioned is the phoP4 gene, which is immediately upstream of the phosphate uptake pstSCAB, phoU operon, and is responsible for its correct expression (Pham et al., 2006). Other TCS genes of M. xanthus have similarly been found to regulate adjacent genedoperons. The PilRS system is responsible for regulating expression of pili and the pilSR genes are found immediately upstream of the pilin gene, pilA, and other genes involved in pilus assembly (Wu and Kaiser, 1995). The ActA and ActB response regulators are encoded in an operon together with the genes for two further proteins, ActC and ActD. While ActA and ActB regulate the expression level of CsgA (the C-signal), ActC and ActD regulate the timing of CsgA production (Gronewold and Kaiser, 2001). The genes for EspA and EspB are also housed together in the genome. An espA mutant exhibits early sporulation, while an espB mutant shows delayed sporulation (Cho and Zusman, 1999a). The sass gene is required for expression of the a 4 5 2 1 developmental reporter and is located close to that of its partner response regulator, sasR. However, the two genes are separated by three open reading frames, including one encoding a putative methyl-accepting chemotaxis protein (MCP). Immediately downstream of sasR is sasN, which inhibits expression of the a 4 5 2 1 reporter. sass, sasR, and sasN are found on separate transcription units (Guo et al., 2000). A similar gene organization is shown by the mrpABC locus, which includes genes for the MrpA histidine kinase, the MrpB response regulator, and MrpC. MrpB is essential for the production of MrpC, which binds to the fruA promoter and is required for fruA promoter activity (Sun and Shi, 2001; Ueki and Inouye, 2003). Chemotaxis genes are also typically found together in large clusters, and the situation is no different in M. xanthus. The Che3, Che4, Dif, and Frz systems are encoded within gene clusters, and each
10. TWO-COMPONENT SIGNAL TRANSDUCTION SYSTEMS contains at least one chew, cheA, cheY, and mcp gene. The che3, che4, and frz clusters also include cheR homologues, while the che3 and frz clusters possess additional cheB homologues (Kirby and Zusman, 2003; McBride et a1.,1989; McCleary and Zusman, 1990; Vlamakis et al., 2004; Yang et al., 1998). The advent of the annotated genome sequence for M. xanthus has enabled an inspection of genes potentially cotranscribed with the TCS genes described in Table 1. In many cases the TCS genes were found to be upstream of hypothetical genes with no predicted function or were in such an orientation that cotranscription with neighboring genes would be impossible. However, in a few cases neighboring genes had been ascribed a predicted function in the genome annotation. The hsfBA genes were found to be upstream of a putative polyphosphate kinase gene; however, it is known that the Hsf system regulates expression of the LonD protease, the gene for which is found separated from the hsf genes. The gene for Nla4 was found to lie upstream of a large cluster of general secretory pathway genes. A mutation generated by insertion of DNA upstream of the first secretory pathway gene gave a similar phenotype to that of an nla4 insertion mutant (Caberoy et al., 2003). However, expression of the secretory genes appears to be unaffected by disruption of nZa4, implying that they are not subject to Nla4 regulation (A. G. Garza, personal communication). AsgA is encoded downstream of a cluster of pyruvate dehydrogenase genes, while the SOLDgene is found adjacent to an uncharacterized chemotaxis gene cluster, and separated from todK by only one gene (encoding a potassium efflux protein, KefC). The nla6 gene is located upstream of an anion transport gene, but an insertion in this gene did not give the same phenotype as an nla6 mutant (Caberoy et al., 2003). Finally, the gene encoding Nla23 was found to reside upstream of a large cluster of pilus biogenesis genes, implying a role in pilus production. Whether there is any significance to the juxtaposition of the genes described above remains to be determined, but they may represent candidates for genes under the control of the respective TCSs.
SIGNALS AND FUNCTIONS In general the signals to which TCSs respond and the responses that they bring about are poorly understood and often difficult to characterize. For several myxobacterial TCSs a biological factor that induces signaling can be proposed (for example, the HsfBA system is probably activated by heat shock); however, there are no cases in which the actual biochemistry of signal perception is understood or the chemical nature of the signal is defined.
181
Similarly, while the broad role of many response regulators is known, a detailed understanding of response elicitation is lacking. This is particularly true for most of the TCSs involved in the regulation of development, where at best a position in the hierarchy of regulatory processes during development is known.
Soluble and Transmembrane Histidine Kinases For the >200 complete bacterial genomes sequenced, 74% of bacterial histidine kinases are found to contain transmembrane helices and would be predicted to possess regions extending into the periplasmic space. The percentage of transmembrane hybrid kinases is slightly lower (52%). Of the TCSs described in Table 1, only 45% of histidine kinases and 20% of hybrid kinases are predicted or known to be transmembrane (or 54 and 8%, respectively, if we consider all histidine and hybrid kinases encoded in the genome of M. xanthus). Figure 6 shows the numbers of histidine kinases in different genomes, compared with the number of those histidine kinases predicted to be transmembrane. Most myxobacteria (with the exception of A. dehalogenans) possess a surprisingly small proportion of transmembrane histidine kinases, implying an unusual degree of intracellular sensing, matched only by members of the cyanobacteria (Galperin, 2005). The presence of transmembrane helices in a histidine kinase suggests that the signal being sensed is extracellular or periplasmic, while those histidine kinases lacking transmembrane regions would most likely be sensing a cytoplasmic signal. There are well-known exceptions to this rule of thumb. CheA homologues are known to associate with membranebound receptor complexes that detect extracellular signals and lack their own transmembrane helices. Similarly, the transmembrane helices of the histidine kinase KdpD have been shown not to be involved in sensing (Heermann et al., 2003). AsgA, EspA, FrgB, HsfB, MrpA, RedE, RodK, SdeK, SocD, SpdS, and TodK do not possess transmembrane helices, implying that they respond to internal signals (Hsfl3responds indirectly to heat shock). Conversely, EspC, MokA, PhoR1, PhoR2, PhoR3, PilS, RedC, and Sass all contain transmembrane helices, implying that they respond to extracytoplasmic stimuli.
Input Functions Many histidine kinases possess conserved N-terminal domains, which appear to be important for signal perception, acting as input domains. Some input domains are commonly found in large numbers of histidine kinases, whereas others are relatively rare. The commonest input domain is the PAS domain, which usually
REGULATORY MECHANISMS
182 120 100 -
80 In 8 In
m K
60
E
E
40 20 0
0
50
100
150
200
Total kinases Figure 6 Transmembrane histidine kinases (includinghybrid kinases) found in different genomes as a function of total numbers of histidine kinases (including hybrids). Trend lines are shown for all bacteria (gray) and for four myxobacteria (black).A d , Anaeromyxobacter dehalogenans; M x , Myxococcus xanthus; Sa, Stigmatella aurantiaca; Sc, Sorangium cellulosum. Most myxobacteria (with the exception of A. dehalogenans) have exceptionally low proportions of TM histidine kinases, implying an unusual degree of sensing of intracellular conditions. Two genomes of Nostoc sp. also exhibit evidence of significant intracellular sensing (see Galperin, 2005).
senses redox potential (Taylor and Zhulin, 1999). Two PAS domains are found in EspA and TodK, while EspC, MXAN5852, PilS, SdeK, and SocD each possess a single PAS domain. Other frequently found input domains are HAMP domains, which act as signal transducing linker domains (found in MXAN7439 and Sass) and GAF domains, which bind cyclic GMP (Galperin et al., 2001). GAF domains are found in AsgD (2 copies), SocD, and S/pdS. CheA homologues frequently possess CheW-like input domains, including the four CheA homologues of M. xanthus described in Table 1. Several histidine kinases of M. xanthus also possess relatively rare input domains. EspA contains an N-terminal FHA domain and is one of only four TCS proteins in GenBank to do so. The presence of an FHA domain in EspA and the lack of transmembrane helices suggest that EspA is activated by binding to specific phosphothreonine residues of a protein(s) within the cytoplasm. EspC also contains an unusual input domain (MASE1), an integral membrane sensor domain, which is found only 13 times in nonmyxobacterial TCS proteins in GenBank. MXAN5852 possesses an N-terminal Na+/proline symporter domain (COG0591j, while MokA contains a predicted periplasmic ligand-binding sensor domain (COG3292j. Several histidine kinases (andhybrid kinases) identified in the M . xanthus genome sequence possess uncommon
input domains. In one protein (MXAN4053) a Ser/Thr kinase domain, an ATPase domain, and a GAF domain precede the transmitter domain, which is a relatively unusual domain architecture (around 50 cases in GenBank). Other input domains found among the histidine kinases include a CHASE domain in MXAN6941 (predicted to be a ligand-binding domain) and a short coiled-coil domain in MXAN2606 (an unusual protein containing two transmitter domains separated by a receiver domain). One histidine kinase (MXAN0168) possesses a KdpD input domain and is preceded by a series of kdp gene orthologues (kdpFABC) implying a role in osmosensitive K+ channel regulation. Intriguingly, the putative kdpD orthologue is not adjacent to the usual partner response regulator gene (KdpEj. The M . xanthus genome also encodes two hybrid kinases, MXAN0712 and MXAN673.5 (each containing a single transmitter domain, upstream of three receiver domains), with a large number of tandemly repeated HAMP/Tar domains (10 and 14 HAMP domains, respectively), a geometry which again is relatively unusual (around 50 cases in GenBank).
Output Functions Most response regulator output domains are DNAbinding domains, and the proteins typically exhibit
10. TWO-COMPONENT SIGNALTRANSDUCTION SYSTEMS phosphorylation-dependent regulation of gene expression. The PhoB, NtrC, and NarL families of response regulators each possess different types of DNA-binding domains (Trans-reg-C, HTH-8, and HTH-LUXR, respectively), and several examples of each are found in the M. xanthus genome. However, despite the large number of myxobacterial DNA-binding response regulators described in the literature (Table l),the genes directly regulated by such proteins are known in very few cases. The NarL-like response regulator FruA has been shown to be required for the production of at least seven developmental proteins, including protein S and DofA (Horiuchi et al., 2002). The DNA-binding domain of FruA bound in vitro to the dofA and fdgA promoters, indicating that they are direct targets of FruA regulation (Ueki and Inouye, 2005a, 2005b). The NtrC homologue HsfA binds to the promoter region of the lonD heat shock gene and is able to initiate transcription from that promoter in vitro (Ueki and Inouye, 2002). As a final example, the pilA gene, encoding pilin, requires the NtrC homologue PilR for expression, and a d4-like promoter sequence has been identified upstream of pilA, suggesting direct regulation by PilR (Wu and Kaiser, 1997). Three non-DNA-binding output domains have also been identified in myxococcal TCS proteins: GGDEF, methyl esterase, and coiled-coil domains (Table 1). GGDEF domains are found in ActB and MXAN5366 and typically have diguanylate cyclase activity, producing cyclic-di-GMP as a signaling molecule. The genome sequence of M. xanthus encodes at least 11TCS proteins containing GGDEF domains, suggesting an extensive integration of TCS and cyclic-di-GMP signaling networks in the myxobacteria. However, none of the predicted TCS proteins of M. xanthus contain EAL domains which possess cyclic-di-GMP phosphodiesterase activity ( Galperin et al., 2001). CheB homologues are characterized by a Cterminal methyl esterase domain and are responsible for modulating chemotactic receptor sensitivity by reversible demethylation. Two orthologues of CheB are known in M. xanthus. One operates within the Frz system (FrzG), while the other is encoded within the che3 chemotactic gene cluster (CheB3). Coiled-coil output domains are found within the paralogous proteins FrzS and AglZ, which regulate motility. It has been suggested that the coiled-coil domain allows interaction with components of the motility systems (Yang et al., 2004). A survey of the TCS genes in M. xanthus identified several response regulators with unusual output domains. Two proteins, MXAN4257 and MXAN5053, contained GAF and GGDEF domains C-terminal to a receiver domain. Four cases with such domain architecture were found in GenBank, all from members
183
of the Deltaproteobacteria. One response regulator (MXAN5592) contained a putative C-terminal HTHXre domain, while another (MXAN4717) contained C-terminal DnaJ and TPR domains, both domain architectures without examples in GenBank, although with orthologues in other myxobacteria. One protein (MXAN6032) contains an N-terminal receiver domain and a Germinal Chew domain. While the combination of single Chew and receiver domains is common, the domains are usually found in the opposite orientation. In fact the case with an N-terminal receiver domain does not appear in GenBank. In one response regulator (MXAN2807),GSPII-E-N and HDc domains are found N-terminal to a receiver domain. GSPII-E-N domains are found in proteins typically involved with type IV pilus biogenesis, suggesting a role in motility, while HDc domains are thought to act as metal-dependent phosphohydrolases. Seven examples of proteins with similar domain architectures appear in GenBank, all from members of the Deltaproteobacteria.
COMPLEX SYSTEMS A typical TCS comprises a single histidine kinase that brings about phosphorylation of a partner response regulator through a His-to-Asp phosphotransfer reaction. However, well-characterized systems suggest that TCS pathways do not always act in such a straightforward and simple manner. The TCSs shown in Fig. 1 exhibit phosphorylation of multiple response regulators by a single histidine kinase, phosphorylation of a single response regulator by multiple histidine kinases, and flow of phosphoryl groups from Asp to His as well as from His to Asp. Such phenomena are possible because of the structural similarity of all receiver domains and all transmitter domains. Consequently TCS proteins can interact in a multitude of ways to create a diverse set of signaling pathway structures, of which the typical TCS is the simplest case. There are many factors which suggest that M. xanthus has complex TCS pathways. First, it possesses an exceptionally large number of response regulators that consist solely of a receiver domain. These response regulators most probably act as intermediaries in phosphotransfer schemes, shuttling phosphoryl groups between different histidine phospho-accepting TCS proteins. Second, an extremely large number of TCS genes are orphaned or in complex TCS gene clusters in the M. xanthus genome. Genes for histidine kinases and response regulators that have single partners are typically found to be adjacent, so the lack of partner genes suggests that orphan TCSs are likely to have multiple partners. By a similar argument,
184 it might be expected that the proteins encoded in complex gene clusters act together, as exemplified by the RedCDEF system (Higgs et al., 2005), necessarily forming complex pathways. A third phenomenon that is suggestive of complex pathways is the failure to find partner proteins for TCS proteins with major phenotypes. For instance, even with much effort partner proteins have not been identified for the response regulator FruA or the histidine kinase SdeK, despite both proteins being critical for multicellular development. If each protein had a single partner, it would be expected that the partner would be found relatively easily in screens of randomly generated mutations. However, if each protein had multiple “redundant” partners, each of which contributed to signaling, then deletion of a single partner might be insufficient to generate the expected phenotype. It seems plausible that proteins such as FruA and SdeK, which regulate key stages of development, might have multiple partners, by analogy to the sporulation phosphorelay of B. subtilis, where multiple kinases affect the phosphorylation state of the master regulator SpoOA (Fig. 1B). A further argument for complex TCSs can be obtained by addressing the frequency with which TCS gene deletions give rise to phenotypes. If we are correct in our assumption that the relative abundance of TCSs found in 111.xanthus is due to requirements for the regulation of motility, development, and/or sporulation, then we might expect the majority of TCS gene disruptions to generate a developmental phenotype. During a systematic disruption of NtrC homologues, Caberoy et al. (2003) found that less than one-third of gene disruptions gave an apparent phenotype. The low proportion of phenotypes observed could be indicative of redundancy in those TCS signaling pathways and thus of multiple partnerships between the TCS proteins. Finally, supporting evidence for complex TCS pathways is also provided by an enrichment for paired TCSs in the list of characterized systems (63%), in comparison to those identified in the genome sequence (34%). This implies that paired systems generally give stronger phenotypes than those TCS proteins encoded by orphan genes or in TCS gene clusters, which would be a likely consequence of genetic redundancy in the pathways encoded by orphans and gene clusters. The domain architecture of individual TCS proteins can also be suggestive of complexity beyond that of the typical TCS. For instance the hybrid histidine kinase RodK has a single transmitter domain and three receiver domains. The most plausible model for RodK action is that the three receiver domains compete as substrates for phosphoryl group transfer from the RodK transmitter domain and that the effector activity of RodK is determined by the phosphorylation state of its receiver
REGULATORY MECHANISMS domains (Rasmussen et al., 2005). An example where a similar mechanism is known to operate is the Vir system of Agrobacterium tumefaciens. VirA is a hybrid kinase containing single transmitter and receiver domains. The response-eliciting component of the Vir system is VirG, a PhoB-like response regulator. The receiver domain of VirA inhibits phosphotransfer from VirA to VirG; however, phosphorylation of the VirA receiver domain relieves the inhibition, allowing phosphorylation of VirG (Chang et al., 1996). Competition between multiple receiver domains to indirectly modulate the phosphorylation state of a response-eliciting receiver domain has also been observed in the chemotactic system of Helicobacter pylori. In this system, retrophosphorylation of CheAY2 occurs by phosphotransfer from the CheY1 receiver domain, and it is consequently thought that the CheY2 receiver domain acts as a phosphate sink to modulate the half-life of phosphorylated CheY1 ( CheYl-P) (J‘1menezPearson et al., 2005). Similar mechanisms can be invoked to provide a simple model of signal transduction through the RedCDEF system, as described above, which contains a response regulator with two receiver domains. There are 16 TCS genes in the M. xanthus genome that are predicted to encode multiple receiver domains (13 in S. cellulosum and 19 in S. aurantiaca), while one gene encodes multiple transmitter domains ( 5 in S. cellulosum and 7 in S. aurantiaca).It seems probable that the signaling pathways in which these proteins act will operate by mechanisms that are dependent upon competition between multiple transmitter-receiver domain interactions and presumably will exhibit dynamic behaviors very different from those of typical TCSs.
DOWN-REGULATION OF PHOSPHORYLATION The phosphorylation state of the proteins within a TCS dictates the strength of the signal passing through that pathway. Generally TCSs become activated, upon receipt of appropriate environmental cues, by a regulated increase in kinase activity. However, in many TCSs regulated phosphatase activities and/or the regulated inhibition of kinase activity is just as important in modulating signal strength. TCS phosphatase activities can act upon phosphorylated histidine or aspartate residues and are found to occur within histidine kinases and response regulators and as separate modulatory proteins. An inhibitor of kinase activity, KipI, has been identified in the B. subtilis sporulation phosphorelay. KipI inhibits KinA kinase activity, preventing sporulation (Wang et al., 1997). A gene encoding a putative KipI homologue is
10. TWO-COMPONENT SIGNALTRANSDUCTION SYSTEMS present in myxobacterial genomes, though it is not found in the vicinity of any TCS genes.
Phosphoaspartate Phosphatases Phosphorylated response regulators typically possess intrinsic autophosphatase activity. In some cases this can be very fast, but it is often slow, depending on the dynamic requirements of the process being regulated. For example, E. coli CheB-P has a half-life of 2 s (Stewart, 1997), while KdpD of E. coli has no significant autophosphatase activity (Puppe et al., 1996). Large variations in phosphatase activity have been observed for different response regulators using in vitro phosphorylation assays (Skerker et al., 2005). This suggests that in some cases, failure to observe phosphorylation of TCS proteins in vitro might be due to high intrinsic phosphatase activities of the proteins rather than an inability to phosphorylate. It is intriguing that for most M. xanthus TCS proteins (with the exception of FrzE and HsfA [Acuna et al., 1995; Ueki and Inouye, 2002]),there is no evidence of aspartate phosphorylation in vitro, though genetic and sequence analyses often suggest the importance of phosphorylation site residues (see, for example, Li and Plamann, 1996, and Ellehauge et al., 1998). Phosphoaspartate residues in response regulators can also be hydrolyzed by extrinsic phosphatases. For example the phosphorelay governing sporulation in B. subtilis contains two receiver domains, which can each be dephosphorylated by three phosphatases: SpoOE, YisI, and YnzD act on SpoOA-P, while RapA, RapB, and RapE act upon SpoOF-P (Perego, 1998). In addition to sporulation pathways, response regulator phosphatases are also important components of chemotactic signaling pathways in enteric bacteria (CheZ orthologues), and a putative CheY-P phosphatase has been identified in the 211. xanthus Dif pathway (DifG [Yang and Li, 2005]), though not in the Frz pathway (McBride et al., 1992). Surprisingly, there are no homologues of the Rap, SpoOE, YisI, YnzD, or CheZ phosphatases encoded in the M. xanthus genome, suggesting that modulation of signal flow by regulated phosphoaspartate phosphatase activity is not generally adopted by the myxobacteria.
Phosphohistidine Phosphatases In contrast to response regulators, phosphorylated histidine kinases appear to be relatively long-lived species in isolation; however, they typically each have at least one specific phosphatase in vivo-their partner response regulator(s).However, phosphohistidine phosphatase proteins are known, the best-studied example being SixA of E. coli. SixA is known to dephosphorylate an Hpt domain within the Arc phosphorelay (Matsubara and Mizuno,
185
2000). There are two homologues of SixA encoded in each available myxobacterial genome (except for a single homologue in A. dehalogenans), but they are typically found to be separated from any TCS genes. The exception to this generalization is M. xanthus, where one of the SixA homologues is encoded close to the phoRPl genes.
Bifunctional Histidine Kinases In a typical histidine kinase, the presence/absence of appropriate environmental signals acts as an on/off switch regulating kinase activity of the transmitter domain. However, some histidine kinases also possess phosphatase activity towards their partner response regulators when they are inactive as kinases. In these bifunctional proteins, both kinase and phosphatase activities reside within the transmitter domain, but phosphatase activity exists at a discrete site from kinase activity, indicating that phosphatase activity is more than just a consequence of retrophosphorylation (Hsing and Silhavy, 1997; Kramer and Weiss, 1999).Phosphorylation of response regulators can occur independently of the partner histidine kinase by interactions with noncognate histidine kinases or small molecule phospho-donors such as acetylphosphate. It is thought that phosphatase activity in bifunctional transmitter domains acts to reduce the phosphorylation of the partner response regulator by noncognate sources, while the partner response regulators of monofunctional kinases are efficient at integrating signals from multiple sources (Alves and Savageau, 2003). Alves and Savageau (2003) have devised an approach to predict whether a transmitter domain is monofunctional or bifunctional by modeling transmitter domain sequences onto known monofunctional (CheA) or bifunctional (EnvZ) transmitter domain structures. The results of such an analysis are shown in Table 1 for experimentally characterized histidine kinases of M . xanthus, which are consequently described as Mono, Bi, or Ntr. Ntr-type transmitter domains would be expected to be bifunctional in the presence of a PI1 protein, but otherwise monofunctional (Alvesand Savageau, 2003). As M. xanthus encodes no obvious PI1 homologue, it is likely that the Ntr-type proteins are monofunctional. Nevertheless, one example suggests that this assumption is invalid. Expression of pilA is dependent on the PilRS TCS, and pilA expression is increased in a pilS mutant, despite being reduced in a pilR background (Wu and Kaiser, 1997).Such behavior suggests that PilS has a bifunctional transmitter domain, despite being predicted to be Ntr-type (Table l),which may in turn imply that there is a PI1 analogue for the Pi1 system in M . xanthus. It would be extremely interesting to know definitively whether particular histidine kinases do have phosphatase
REGULATORY MECHANISMS
186 activity, as such properties have a fundamental impact on the dynamic properties of a TCS, and such knowledge would enable inferences to be drawn about the integration of multiple signals by different TCS pathways.
TCSs OF OTHER MYXOBACTERIA Only one myxobacterial TCS protein has been described from a species other than M. xantbus. The CyaB protein of S. aurantiaca contains an N-terminal receiver domain and a C-terminal CyaA adenylate cyclase domain (CoudartCavalli et al., 1997). Site-directed mutagenesis of the phospho-accepting aspartate residue in the CyaB receiver domain significantly reduced adenylate cyclase activity of the protein, implying that phosphorylation of the receiver domain stimulates cyclic AMP production. CyclicAMP is known to transiently accumulate during early development in S. aurantiaca (Coudart-Cavalli et al., 1997) and M . xanthus (Yajko and Zusman, 1978). An orthologue of cyaB exists in the M . xanthus genome alongside a histidine kinase gene, while S. cellulosum possesses two cyaB orthologues (one orphan gene and one adjacent to a hybrid kinase). The myxobacteria seem to possess multiple TCS proteins that contain cyclic nucleotide metabolism domains. In addition to a cyaB orthologue, the M . xantbus genome includes the gene for a response regulator with an N-terminal Ser/Thr kinase domain, a central receiver domain, and a C-terminal CyaA domain. Orthologues of this gene are apparent in the S. cellulosum genome and in S. aurantiaca (three paralogues). Additionally, M . xantbus possesses the gene for a histidine kinase which has an N-terminal cyclic nucleotide binding domain, implying that TCSs of M . xanthus are responding to changes in cyclic nucleotide levels, as well as regulating them. With the sequencing of multiple myxobacterial genomes it has become possible to use comparative genomics to gain novel insights into the TCSs of myxobacteria. Genomes can be assessed for the presence or absence of specific TCS homologues, lineage-specific changes in TCS properties can be identified, and in some cases changes in gene organization can guide searches for partner proteins. The sequences of multiple myxobacterial genomes can also provide important insights into the evolution of TCSs, from which inferences can be made regarding evolutionary changes in TCS network connectivity and the properties of contemporary TCSs.
PERSPECTIVES The myxobacteria possess exceptionally large numbers of TCS genes, which is not merely due to their large genome sizes (Fig. 2). Many TCS proteins of M . xantbus
have been characterized experimentally, with most being involved with the regulation of motility and/or development (Table 1).There is accumulating evidence that the TCS pathways of M. xantbus are relatively complex. The large number of response regulators that lack output domains, an absence of conventional phosphorelays, large numbers of TCS genes found as orphans or in complex gene clusters (Table 2; Fig. 4), and the complex domain architectures of myxobacterial TCS proteins all suggest that the TCSs of myxobacteria operate in significantly different ways from most other bacteria (with the notable exception of the cyanobacteria). One particularly unusual feature of myxobacterial TCSs is the adoption of chemotactic signaling pathways to regulate gene expression, a behavior which so far has been observed only in M. xanthus. The low proportion of sensor kinases with transmembrane helices implies that myxobacteria are unusually sensitive to changes in their internal state. This observation and their large numbers of TCS proteins suggests that the myxobacteria are complex introverts with high IQs, making them excellent role models as well as model organisms for cellular signaling studies.
References Acuna, G., W. Shi, K. Trudeau, and D. R. Zusman. 1995. The ‘CheA’ and ‘CheY’ domains of Myxococcus xanthus FrzE function independently in vitro as an autokinase and a phosphate acceptor respectively. FEBS Lett. 16:3 1-33. Alves, R., and M. A. Savageau. 2003. Comparative analysis of prototype two-component systems with either bifunctional or monofunctional sensors: differences in molecular structure and physiological function. Mol. Microbiol. 48:25-51. Burbulys, D., K. A. Trach, and J. A. Hoch. 1991. Initiation of sporulation in B. subtilis is controlled by a multicomponent phosphorelay. Cell 64545-552. Bustamante, V. H., I. Martinez-Flores, H. C. Vlamakis, and D. R. Zusman. 2004. Analysis of the Frz signal transduction system of Myxococcus xanthus shows the importance of the conserved C-terminal region of the cytoplasmic chemoreceptor FrzCD in sensing signals. Mol. Microbiol. 53:15011513. Caberoy, N. B., R. D. Welch, J. S. Jakobsen, S. C. Slater, and A. G. Garza. 2003. Global mutational analysis of NtrC-like activators in Myxococcus xanthus: identifying activator mutants defective for motility and fruiting body development. ]. Bacteriol. 185:6083-6094. Carrero-LCrida, J., A. Moraleda-Muiioz, R. Garcia-Herniindez, J. PCrez, and J. Muiioz-Dorado. 2005. PhoR1-PhoP1, a third two-component system of the family PhoRP from Myxococcus xanthus: role in development. ]. Bacteriol. 187:4976-4983. Chang, C. H., J. Zhu, and S. C. Winans. 1996. Pleiotropic phenotypes caused by genetic ablation of the receiver module of the Agrobacterium tumefaciens VirA protein. ]. Bacteriol. 178:4710-4716.
10. TWO-COMPONENT SIGNALTRANSDUCTION SYSTEMS Cho, K., and D. R. Zusman. 1999a. Sporulation timing in Myxococcus xanthus is controlled by the espAB locus. Mol. Bacteriol. 34:7 14-725. Cho, K., and D. R. Zusman. 1999b. AsgD, a new two-component regulator required for A-signalling and nutrient sensing during early development of Myxococcus xanthus. Mol. Microbiol. 34:268-281. Cho, K., A. Treuner-Lange, K. A. O’Connor, and D. R. Zusman. 2000. Developmental aggregation of Myxococcus xanthus requires frgA, an frz-related gene. J. Bacteriol. 182: 6614-6621. Cock. P. J., and D. E. Whitworth. 2007. Evolution of gene overlaps: relative reading frame bias in prokaryotic twocomponent system genes. J. Mol. Evol. 64:457-462. Coudart-Cavelli, M. P., 0. Sismeiro, and A. Danchin. 1997. Bifunctional structure of two adenylyl cyclases from the myxobacterium Stigmatella aurantiaca. Biochimie 79:757-767. Ellehauge, E., M. Norregaard-Madsen, and L. SsgaardAndersen. 1998. The FruA signal transduction protein provides a checkpoint for the temporal co-ordination of intercellular signals in Myxococcus xanthus development. Mol. Microbiol. 30:807-817. Fabret, C., V. A. Feher, and J. A. Hoch. 1999. Two-component signal transduction in Bacillus subtilis: how one organism sees its world. J. Bacteriol. 181:1975-1983. Feher, V. A., J. W. Zapf, J. A. Hoch, J. M. Whiteley, L. P. McIntosh, M. Rance, N. J. Skelton., F. W. Dahlquist, and J. Cavanagh. 1997. High-resolution NMR structure and backbone dynamics of the Bacillus subtilis response regulator, SpoOF: implications for phosphorylation and molecular recognition. Biochemistry 36: 10015-1 0025. Galperin, M. Y., A. N. Nikolskaya, and E. V. Koonin. 2001. Novel domains of the prokaryotic two-component signal transduction systems. FEMS Microbiol. Lett. 203:ll-21. Galperin, M. Y. 2005. A census of membrane-bound and intracellular signal transduction proteins in bacteria: bacterial IQ, extroverts and introverts. BMC Microbiol. 5:35. Gronewold, T. M. A., and D. Kaiser. 2001. The act operon controls the level and time of C-signal production for Myxococcus xanthus development. Mol. Microbiol. 40:744-756. Guo, D., Y. Wu, and H. B. Kaplan. 2000. Identification and characterization of genes required for early Myxococcus xanthus developmental gene expression. J. Bacteriol. 182:4564-4571. Hager, E., and R. E. Gill. 2001. Identification and characterization of spdR mutations that bypass protease-dependent regulation of developmental gene expression in Myxococcus xanthus. Mol. Microbiol. 39:765-780. Heermann, R., A. Fohrmann, K. Altendorf, and K. Jung. 2003. The transmembrane domains of the sensor kinase KdpD of Escherichia coli are not essential for sensing K+ limitation. Mol. Micro biol. 47: 83 9-84 8. Higgs, P. I., K. Cho, D. E. Whitworth, L. S. Evans, and D. R. Zusman. 2005. Four unusual two-component signal transduction homologs, RedC to RedF, are necessary for timely development in Myxococcus xanthus. J. Bacteriol. 187~8191-8195. Hoch, J. A. 2000. Two-component and phosphorelay signal transduction. Curr. Opin. Microbiol. 3:165-170.
187
Hoch, J. A., and T. J. Silhavy. 1995. Two-Component Signal Transduction. American Society for Microbiology, Washington, DC. Horiuchi, T., M. Taoka, T. Isobe, T. Komano, and S. Inouye. 2002. Role of fruA and csgA genes in gene expression during development of Myxococcus xanthus. Analysis by two-dimensional gene electrophoresis. J. Biol. Chem. 277~26753-26760. Hsing, W., and T. J. Silhavy. 1997. Function of conserved histidine-243 in phosphatase activity of EnvZ, the sensor for porin osmoregulation in Escherichia coli. J. Bacteriol. 179:3729-373 5. Hutchings, M. I., P. A. Hoskisson, G. Chandra, and M. J. Buttner. 2004. Sensing and responding to diverse extracellular signals? Analysis of sensor kinases and response regulators of Streptomyces coelicolor A3(2). Microbiology 150:2795-2806. Jakobsen, J. S., L. Jelsbak, L. Jelsbak, R. D. Welch, C. Cummings, B. Goldman, E. Stark, S. Slater, and D. Kaiser. 2004. Sigma54 enhancer binding proteins and Myxococcus xanthus fruiting body development. J. Bacteriol. 186:4361-4368. Jelsbak, L., M. Givskov, and D. Kaiser. 2005. Enhancer-binding proteins with a forkhead-associated domain and the d4regulon in Myxococcus xanthus fruiting body development. Proc. Natl. Acad. Sci. USA 102:3010-3015. Jimenez-Pearson., M. A., I. Delany, V. Scarlato, and D. Beier. 2005. Phosphate flow in the chemotactic response system of Helicobacter pylori. Microbiology 151:3299-3311. Keseler, I. M., and D. Kaiser. 1997. Sigma54, a vital protein for Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 94:19791984. Kimura, Y., H. Nakano, H. Terasaka, and K. Takegawa. 2001. Myxococcus xanthus m o k A encodes a histidine kinaseresponse regulator hybrid sensor required for development and osmotic tolerance. J. Bacteriol. 183:1140-1146. Kirby, J. R., and D. R. Zusman. 2003. Chemosensory regulation of developmental gene expression in Myxococcus xanthus. Proc. Natl. Acad. Sci USA 100:2008-2013. Kofoid, E. C., and J. S. Parkinson. 1988. Transmitter and receiver modules in bacterial signaling proteins. Proc. Natl. Acad. Sci. USA 85:4981-4985. Kramer, G., and V. Weiss. 1999. Functional dissection of the transmitter module of the histidine kinase NtrB in Escherichia coli. Proc. Natl. Acad. Sci. USA 96604-609. Lancero, H., N. B. Caberoy, S. Castaneda, Y. Li, A. Lu, D. Dutton, X.-Y. Duan, H. B. Kaplan, W. Shi, and A. G. Garza. 2004. Characterization of a Myxococcus xanthus mutant that is defective for adventurous motility and social motility. Microbiology 150:4085-4093. Lancero,H., S. Castaaneda,N. B. Caberoy,X. Ma,A. G. Garza, and W. Shi. 2005. Analysing protein-protein interactions of the Myxococcus xanthus Dif signalling pathway using the yeast two-hybrid system. Microbiology 151:1535-1541. Lee, B., P. I. Higgs, D. R. Zusman, and K. Cho. 2005. EspC is involved in controlling the timing of development in Myxococcus xanthus. J. Bacteriol. 1875029-5031. Li, Y., and L. Plamann. 1996. Purification and in vitro phosphorylation of Myxococcus xanthus AsgA protein. J. Bacteriol. 178:289-292.
188 Matsubara, M., and T. Mizuno. 2000. The SixA phosphohistidine phosphatase modulates the ArcBA phosphorelay signal transduction in Escherichia coli. FEBS Lett. 470:118124. McBride, M. J., T. Kohler, and D. R. Zusman. 1992. Methylation of FrzCD, a methyl-accepting taxis protein of Myxococcus xanthus, is correlated with factors affecting cell behavior. J. Bacteriol. 174:4246-4257. McBride, M. J., R. A. Weinberg, and D. R. Zusman. 1989. “Frizzy” aggregation genes of the gliding bacterium Myxococcus xanthus show sequence similarities to the chemotaxis genes of enteric bacteria. Proc. Natl. Acad. Sci. USA 86~424-428. McCleary, W. R., and D. R. Zusman. 1990. FrzE of Myxococcus xanthus is homologous to both CheA and CheY of Salmonella typhimurium. Proc. Natl. Acad. Sci. USA 875898-5902. Mignot, T., J. P. Merlie, Jr., and D. R. Zusman. 2005. Regulated pole-to-pole oscillations of a bacterial gliding motility protein. Science 310:855-857. Mizuno, T., T. Kaneko, and S. Tabata. 1996. Compilation of all genes encoding bacterial two-component signal transducers in the genome of the cyanobacterium, Synechocystis sp. strain PCC 6803. D N A Res. 3:407-414. Mizuno, T. 1997. Compilation of all genes encoding twocomponent phosphotransfer signal transducers in the genome of Escherichia coli. D N A Res. 4:161-168. Moraleda-Muiioz, A., J. Carrero-LCrida, J. PCrez, and J. Muiioz-Dorado. 2003. Role of two novel two-component regulatory systems in development and phosphatase expression in Myxococcus xanthus. J. Bacteriol. 185:1376-1383. Ogawa, M., S. Fujitani, X. Mao, S. Inouye, and T. Komano. 1996. FruA, a putative transcription factor essential for the development of Myxococcus xanthus. Mol. Microbiol. 22~757-767. Parkinson, J. S., and E. C. Kofoid. 1992. Communication modules in bacterial signalling proteins. Annu. Rev. Genet. 26:71-112. Perego, M. 1998. Kinase-phosphatase competition regulates Bacillus subtilis development. Trends Microbiol. 6366-370. Pham, V. D., C. W. Shebelut, I. R. Jose, D. A. Hodgson, D. E. Whitworth, and M. Singer. 2006. The response regulator PhoP4 is required for late developmental events in Myxococcus xanthus. Microbiology 152:1609-1620. Pollack, J. S., and M. Singer. 2001. SdeK, a histidine kinase required for Myxococcus xanthus development. J. Bacteriol. 183:35 89-3 596. Pritchard, L., J. A. White, P. R. J. Birch, and I. K. Toth. 2006. GenomeDiagram: a python package for the visualisation of large-scale genomic data. Bioinformatics 22:616-617. Puppe, W., M. Jung, M. Lucassen, and K. Altendorf. 1996. Characterization of truncated forms of the KdpD protein, the sensor kinase of the K+-translocating Kdp system of Escherichia coli. J. Biol. Chem. 271:25027-25034. Rasmussen, A. A., and L. Ssgaard-Andersen. 2003. TodK, a putative histidine protein kinase, regulates timing of fruiting body morphogenesis in Myxococcus xanthus. J. Bacteriol. 185~5452-5464. Rasmussen, A. A., S. L. Porter, J. P. Armitage, and L. SsgaardAndersen. 2005. Coupling of multicellular morphogenesis
REGULATORY MECHANI sM s and cellular differentiation by an unusual hybrid histidine protein kinase in Myxococcus xanthus. Mol. Microbiol. 56:1358-1372. Sasse-Dwight, S., and J. D. Gralla. 1990. Role of eukaryotictype functional domains found in the prokaryotic enhancer receptor factor sigma 54. Cell 62:945-954. Shimkets, L. J. 1999. Intercellular signalling during fruitingbody development of Myxococcus xanthus. Annu. Rev. Microbiol. 53525-549. Skerker, J. M., M. S. Prasol, B. S. Perchuk, E. G. Biondi, and M. T. Laub. 2005. Two-component signal transduction pathways regulating growth and cell cycle progression in a bacterium: a system-level analysis. PLoS Biol. 3:e334. Sonnhammer, E. L., G . von Heijne, and A. Krogh. 1998. A hidden Markov model for predicting transmembrane helices in protein sequences. Proc. Int. Conf. Intell. Syst. Mol. Biol. 6~175-182. Stewart, R. C. 1997. Activating and inhibitory mutations in the regulatory domain of CheB, the methylesterase in bacterial chemotaxis. J. Biol. Chem. 268:1921-1930. Stock, J. B., A. J. Ninfa, and A. M. Stock. 1989. Protein phosphorylation and regulation of adaptive responses in bacteria. Microbiol. Rev. 53:450-490. Sun, H., and W. Shi. 2001. Genetic studies of mrp, a locus essential for cellular aggregation and sporulation of Myxococcus xanthus. J. Bacteriol. 183:4786-4795. Taylor, B. L., and I. B. Zhulin. 1999. PAS domains: internal sensors of oxygen, redox potential, and light. Microbiol. Mol. Biol. Rev. 63:479-506. Trudeau, K. G., M. J. Ward, and D. R. Zusman. 1996. Identification and characterization of FrzZ, a novel response regulator necessary for swarming and fruiting-body formation in Myxococcus xanthus. Mol. Microbiol. 20:645-655. Ueki, T., and S. Inouye. 2005a. Activation of a developmentspecific gene, dofA, by FruA, an essential transcription factor for development of Myxococcus xanthus. J. Bacteriol. 187~8504-8506. Ueki, T., and S. Inouye. 2005b. Identification of a gene involved in polysaccharide export as a transcription target of FruA, an essential factor for Myxococcus xanthus development. J. Biol. Chem. 280:32279-32284. Ueki, T., and S. Inouye. 2003. Identification of an activator protein required for the induction of fruA, a gene essential for fruiting body development in Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 10023782-8787. Ueki, T., and S. Inouye. 2002. Transcriptional activation of a heat-shock gene, lonD, of Myxococcus xanthus by a two component histidine-aspartate phosphorelay system. J. Biol. Chem. 2 7 7 6 170-61 77. Vlamakis, H. C., J. R. Kirby, and D. R. Zusman. 2004. The Che4 pathway of Myxococcus xanthus regulates type IV pilus-mediated motility. Mol. Microbiol. 52:17991811. Wang, L., R. Grau, M. Perego, and J. A. Hoch. 1997. A novel histidine kinase inhibitor regulating development in Bacillus subtilis. Genes Deu. 11:2569-2579. Ward, M. J., K. C. Mok, D. P. Astling, H. Lew, and D. R. Zusman, 1998a. An ABC transporter plays a developmental aggregation role in Myxococcus xanthus. J. Bacteriol. 1805697-5703.
10. TWO-COMPONENT SIGNALTRANSDUCTION SYSTEMS Ward, M. J., H. Lew, A. Treuner-Lange, and D. R. Zusman. 1998b. Regulation of motility behavior in Myxococcus xanthus may require an extracytoplasmic-function sigma factor. J. Bacteriol. 180:5668-5675. Ward, M. J., H. Lew, and D. R. Zusman. 2000. Social motility in Myxococcus xanthus requires FrzS, a protein with an extensive coiled-coil domain. Mol. Microbiol. 37:1357-1371. Wolanin, P. M., E A. Thomason, and J. B. Stock. 2002. Histidine protein kinases: key signal transducers outside the animal kingdom. Genome Biol. 3:REVIEWS3013. Wu, S. S., and D. Kaiser. 1995. Genetic and functional evidence that Type IV pili are required for social gliding motility in Myxococcus xanthus. Mol. Microbiol. 18547-55 8. Wu, S. S., and D. Kaiser. 1997. Regulation of expression of the pilA gene in Myxococcus xanthus. 1. Bacteriol. 179:77487758.
189
Yajko, D. M., and D. R. Zusman. 1978. Changes in cyclic AMP levels during development in Myxococcus xanthus. J. Bacteriol. 133:1540-1542. Yang, R., S. Bartle, R. Otto, A. Stassinopoulos, M. Rogers, L. Plamann, and P. Hartzell. 2004. AglZ is a filament-forming coiled-coil protein required for adventurous gliding motility of Myxococcus xanthus. 1. Bacteriol. 186:61686178. Yang, Z., and Z. Li. 2005. Demonstration of interactions among Myxococcus xanthus Dif chemotaxis-like proteins by the yeast two-hybrid system. Arch. Microbiol. 183:243252. Yang, Z., Y. Geng, D. Xu, H. B. Kaplan, and W. Shi. 1998. A new set of chemotaxis homologues is essential for Myxococcus xanthus social motility. Mol. Microbiol. 30:11231130.
Myxobacteria: Mlrlticelltrlarity and Differentiation Edited by David E. Whitworth 02008 ASM Press, Washington, D.C.
Sumiko Inouye Hirofumi Nariya Jos6 Mufioz-Dorado
Protein Ser/Thr Kinases and Phosphatases in
11
Myxococcus xanthus
Protein Ser/Thr/Tyr kinases and protein phosphatases function as biological switches that turn on and off signal transduction pathways, where they participate by phosphorylation and dephosphorylation. Recently, whole bacterial genome sequencing has revealed that they exist in a wide variety of pathogenic and developmental bacteria ( Av-Gay and Everett, 2000; Wang et al., 2002; Petrickova and Petricek, 2003).The physiological roles of the protein Ser/Thr kinase (PSTK) signaling systems in prokaryotes are beginning to be understood at the molecular level. PSTKs are now known to play important roles in the secondary metabolism of Streptomyces coelicolorA3(2)(Lee et al., 2002), the survival of Mycobacterium tuberculosis within macrophages (Walburger et al., 2004), activation of a myxobacterial enzyme for the consumption of glycogen (Nariya and Inouye, 2002, 2003), and regulation of gene expression during Myxococcus xanthus development (Nariya and Inouye, 2006). The first PSTK in prokaryotes was identified and physiologically characterized because of its role in M. xanthus development (Mufioz-Dorado et al., 1991). During the past 15 years since their discovery, several PSTICs have been cloned and investigated for their roles in the M . xanthus life cycle.
In the course of these investigations, three multikinase-associated proteins, MkapA, MkapB, and MkapC, each of which contains well-known protein-protein interaction domains, have been identified and studied for their roles in the PSTK signaling systems in M. xanthus (Nariya and Inouye, 2005a). Recently, the first functional PSTK cascade in prokaryotes was discovered (Nariya and Inouye, 2005b). This regulatory cascade regulates the expression of two transcription activators, MrpC and FruA, which are essential for multicellular fruiting body formation and sporulation (Nariya and Inouye, 2006). Based on analyzing the recently complete M. xanthus genome sequence database (Goldman et al., 2006), 102 genes that encode putative PSTKs and 34 genes of putative protein phosphatases (PPs) have been identified. In contrast to PSTKs, only one protein Ser/ Thr phosphatase has been characterized (Treuner-Lange et al., 2001). The PSTK signal transduction systems function differently from the two-component His-Asp phosphorelay system (TCST) that plays predominant roles in prokaryotic signal transduction. There are two important differences. In response to a signal, PSTICs use ATP to phosphorylate
Sumiko Inouye, Department of Biochemistry, Robert Wood Johnson Medical School, Piscataway, NJ 08854. Hirofumi Nariya, Department of Biochemistry, Robert Wood Johnson Medical School, Piscataway, NJ 08854. Jose Mufioz-Dorado, Departamento de Microbiologia, Facultad de Ciencias, Universidad de Granada, E-18071 Granada, Spain.
191
REGULATORY MECHANISMS
192 their own Ser/Thr residues for activation of their kinase activities. An activated PSTK phosphorylates its substrate using ATP, in contrast to TCST, where that high-energy phosphogroup is transferred from His residues of kinases to Asp residues of response regulators (Stock et al., 2000). Thus, the signal received by a PSTK system can be greatly amplified and maintained for long time periods, because the phosphogroups of Ser/Thr/Tyr are much more stable than those of His/Asp, and a specific phosphatase is required for the inactivation of PSTKs. In this chapter, we first describe PSTKs and then PPs in M. xanthus.
EUKARYOTIC-LIKE PROTEIN Ser/Thr KINASES Identification and Classification of PSTKs in the Genome M. xanthus contains 102 PSTKs, the largest number so far found in a bacterial genome, based on an analysis of the genome sequence database (http://www.tigr.org) (Goldman et al., 2006), and the M. xanthus kinome is roughly equivalent to that of Saccharomyces cerevisiae, which has 130 PSTKs (Manning et al., 2002). The catalytic domains of PSTKs consist of 11 subdomains, each of which contains highly conserved sequences (Fig. 1) (Hanks et al., 1988). All but three myxobacterial PSTKs are typical eukaryotic protein kinases (ePKs) containing most of the conserved essential residues in subdomains I through XI of the kinase catalytic domain (CD) (Hanks and Hunter, 1995).In addition to ePKs, M . xanthus contains three atypical protein kinases (aPKs) that belong to the RIO and ABCl (RIO-1 and ABC1-1 and 1-2, respectively) families, and have weak similarity to ePKs (reviewed by Laronde-LeBlanc and Wlodawer, 2005). I
II
Ill IV
V
Vla
Although phosphorylation of Tyr has been shown with cell extracts from M. xanthus (Frasch and Dworkin, 1996), as in the case of yeast, no typical protein Tyr kinases have been detected in the Myxococcus kinome, based on sequence analysis. Classification of PSTKs can be made based upon amino acid substitutions in the CD, and five groups of ePKs have been identified due to substitutions in subdomains VIb (HRDxK) and VIII (TxxxxAPE) as shown in Fig. 1. Most ePKs fall into one of two groups, 56 Pkt and 27 Psk, with all three remaining groups making up less than 20% of the ePKs (7 Pab, 5 Psd, and 4 Pdd). Classification of ePK from other bacteria shows that the Psk, Psd, and Pdd groups are almost exclusively found in M . xanthus. Phylogenetic analysis, using the neighborjoining method on the catalytic domain sequences, allows additional classification consistent with the structural classification- As shown in Fig. 2, the Pkt group is further divided into six subgroups, 14 PktA, 10 PktB, 9 PktC, 12 PktD, 4 PktE, and 7 PktF, while the Psk group is divided into two subgroups, 17 PskA and 10 PskB. Among the 11 ePK-branches, PktE and Pdd seem to be recently diverged from PktF and Psd, respectively. The 102 PSTKs identified were named according to the classification described above, disregarding their previous characterization (Table 1). The classification and catalytic domain substitution patterns of some ePKs are particularly interesting. In the Psk group, the conserved K residue in subdomain VIb that forms the catalytic loop facilitating the phosphotransfer reaction by its positive charge, is replaced by a Ser residue. In addition, the Thr residue in subdomain VIII known as the activation site of ePK (Adams et al., 1995), is replaced by a Lys residue. PskA2/Pkn6 and PskASIPknl4 have been demonstrated to be active Vlb
VII
Vlll
IX
x
XI
Group
Pkt (56) Psk(27) PSd (5) Pdd(4) Pab(7)
G G A G E R
G T G G
G V A K E A V V K E G L V/L K E G S L K E G V/LV# #
ATP binding
Catalytic loop
- -
Active Acti- Interact Stabilize Interact center vation with catalytic with loop R(XI) loop E(VIII)
Figure 1 Classification of 99 eukaryotic-like PSTKs. The amino acid residues used for classification are highlighted by gray boxes. Invariant residues in the catalytic domain (Hanks and Hunter, 1995) are in boldface. Symbols: #, at least one residue is substituted or deleted at the position of each Pab; x, the residue is not well conserved.
11. PROTEINSER/THR KINASESAND PHOSPHATASES IN M . XANTHUS
193
Figure 2 Phylogenetic tree of 99 eukaryotic-like PSTKs. The phylogenetic tree was built by the neighbor-joining method (CLUSTALW [http://align.genome.jp/] and NJplot [http://pbil. univ-lyon.fr/software/njplot.html])using manually aligned kinase catalytic domains (subdomain from I to XI [Hanks et al., 19881) and illustrated using Tree View programs (http:// taxonomy.zoology.gla.ac.uk/rod/treeview.html). The Pkt group is shown by black lines in a medium-gray background except for the PktC subgroup, which is shown with dotted lines. The Psk group is shown by gray lines in a light-gray background, and the Pab group is indicated by dotted lines. Psd and Pdd groups are shown with dotted and large dotted lines in light- and dark-gray backgrounds, respectively. PSTKs grouped by open ovals have homologies not only in the catalytic domain but also in the regulatory domain.
ePKs, phosphorylating at Ser and Thr residues, implying that the Ser residue in subdomain VIb may function as the activation site in the Psk group (Zhang et al., 1996; Nariya and Inouye, 2005b). The three rarer ePK groups have striking substitutions. Pab are abnormal ePKs having substitutions at essential residues in subdomains I1 (AxK), VIb (HRD), and VII (DFG) in addition to substitutions in VIb and VIII (Fig. 1). Therefore, Pab PSTKs may be catalytically inactive ePKs that act as modulators, substrates for active kinases, and scaffolds for assembly of signaling complexes as in eukaryotes (reviewed by Manning et al.,
2002). The Psd group has a negative-charged Asp residue at the Lys residue position in subdomain VIII of the Psk group, and Pdd has another Asp residue substitution at the Ser residue in subdomain VIb of the Psd group. In contrast, the Asp residue in the HRD sequence of subdomain VIb is replaced with an Ala residue. However, an Asp residue two residues downstream (HRAxD)may compensate for the role of the Asp residue in HRD.
Analysis of Domain Structure of 102 PSTKs Most M . xanthus PSTKs have CDs at the N-terminal end and regulatory domains (RDs) at the C-terminal end,
REGULATORY MECHANISMS
194 Table 1 List of PSTK genes" Gene
MXAN
Type
pktAl A2/pknl A3 A4/pkn9 AS/p knD 1 A6 A7 A8Ipkn3 A9 A10 All A12 A13 A14
6500 1467 614 755 930 4338 3094 3202 4591 3338 6043 2680 6317 1297
s2 s2 s2 M1 s2 s2 s2 M1 M1 M1 M1 M1 M3
pktBl B2 B3 B4 BS B6 B7 B8IpknD2 B9 B10
642 8 5045 882 2255 6183 871 5176 933 4337 3092
M1 M1 M1 M1 M1 M3 s2 s2 s2 s2
pktCl C2/pkn8 c3 c4
70 1710 273 8 1480 7162 117 6561 6009 7082
M1 M2 M2 M1 M1 M1 M1 M1 M2
4017 2156
M1 M1
cs C6 c7 C8 c9 pktDl/pkn4 D2
s1
Gene
MXAN
Type
906 525 1577 5886 2549 7251 6420 3955 1163 4049
M1 M1 s1 s2
pktEl E2 E3 E4
3182 2176 1233 4842
M1 M1 M1 M1
pktFl F2 F3 F4 FS F6 F7
4373 4482 1896 7370 2840 7269 2399
M1 M1 M1 M1 M1
pskAl A21pkn6 A3/masK A4 AS/p knl4 A6 A7 A8 A9 A10 A1 1/pkn7 A12 A13/pknll A14
2059 2550 1929 2596 5116 5696 4700 5976 552 2586 2910 4557 291 1 6545
M1 M1 M1 M1 s2 M1 M1 s1 M1 M1 M1 M1 M1 M3
D3 04 DS D6 D7/pknS DUpkn12 D9/pkn2 D1 OIpknlO Dll D12
s1 s2 M1 M1 M1 s2
s1 M1
Gene
MXAN
Type
23 18 3099
M1 M1 M1
pskBl B2 B3 B4 BS B6 B7 B8 B9 B10
396 7208 6570 265 3693 5620 6669 6312 960 2980
M3 M1 M1 M1 M1 M1 51 M1 51 51
psdl 2 3 4
3272 7371 1892 4479 4371
51 51 51 51 51
pddl 2 3 4
3183 2177 1234 4841
51 51 51 51
pub1 2 3 4 6 7
5517 40.53 2077 1088 3710 4434 724
M3 M2 M2 51 52 M1 51
riol abcl-1 abcl-2
2315 725 3899
52 51 M3
A1 5lpknl3 A16 A17
S
S
"Abbreviations: M1, type I receptor type kinase; M2, type I1 receptor type kinase; M3, membrane embedded kinase; S1, small cytoplasmic kinase; S2, large cytoplasmic kinase.
like PSTKs in other bacteria, according to CDD-BLAST searches at http://www.ncbi.nlm.nih.gov/Structure/cdd/ cddshtml. Prediction of hydrophobic sequences using the TMHMM program (http://www.cbs.dtu.dk/services/ TMHMM-2.0/) indicates diverse PSTK subcellular localization (Table 1).The largest group consists of 55 type I receptor kinases with an exposed RD in the periplasmic space (Ml). Two moderate-sized groups are made up of cytoplasmic kinases, 19 small PSTKs with fewer
than 150 residues in the C-terminal domain (Sl) and 17 PSTKs with a longer RD at the C-terminal end (S2).The two smallest groups are of five type I1 receptor kinases that have two hydrophobic sequences resulting in exposure of both CD and RD in the cytoplasmic space (M2) and a group of six membrane-anchored kinases (M3). Despite the diverse localization, over one-half of the PSTKs are receptor-type kinases. The frequency of receptor-type kinases suggests that these PSTKs are involved in
11. PROTEINSER/THRKINASESAND PHOSPHATASES IN 211. XANTHUS signaling systems required to regulate the complex M. xanthus life cycles together with the sensor histidine kinases in TCST. The majority of RDs have no similarity to other proteiddomains based upon the CDD-BLAST search. However, some RDs have recognizable functional domains. PktD9/Pkn2 has a Class I11 adenylyl cyclase (AC) domain (PF00211) and PktD12 consists of a Class I11 AC domain and a typical receiver domain of TCST. These domains suggest that M. xanthus may have unique regulatory modes for signal transduction by cyclic AMP (cAMP)/cGMP via both PSTK and TCST systems. Six PSTKs, PktBWPktD2, PktB9, PktB10, PktD6, PktD8/ Pknl2, and Pab2, are predicted to contain an ATPase domain with P-loop (COG3899). Pab2 contains a histidine kinase domain with a cNMP binding domain. Thirteen homologues of Pab2 are found in the cyanobacterium Nostoc PCC7120, a filamentous heterocyst former, but not in the unicellular cyanobacterium Synechocystis PCC6803 (Wang et al., 2002). Three PSTKs, PktD3, PktD4, and Pab3, contain a conserved domain with unknown function (DUF323)in their RDs. The RDs of PktB1, B2, B8, B9, and B10, and PskB5 and B8 contain a TPR (tetratricopeptide repeat), a repetitive domain that is generally responsible for protein-protein interactions and assembly of multiprotein complexes. The RDs of all PSTKs in the PktC subgroup except PktCl contain a kinesin light chain-like domain similar to TPR. The RD of PktA12 contains the C-terminal domain of TonB that is known to function as a dimerization domain and to contact with TonB receptor (Chang et al., 2001), implying that PktA12 may form a homodimer or a heterodimer with TonB to regulate iron transportation. The diverse properties of PSTK RDs indicate that they have a broad importance for M. xanthus physiological activity.
195
pktE-pdd and pktF-psd Duplications Nested gene duplication, divergence, and rearrangements have occurred with p k t E , pktF, pdd, and psd PSTKs. Analysis of the PSTK phylogenetic tree indicates that the PktE and PktF groups recently diverged, as did the Pdd and Psd groups (Fig. 2). Each pair of groups retains weak similarity not only in their CDs but also in their RDs. We first focus on p k t E and pdd. Based on genome linkage analysis, every pdd is located downstream of a pktE: p k t E l - p d d l , pktE2-pdd2, pktE3-pdd3, and pktE4-pdd4 (Fig. 3a). Upstream of each pktE-pdd, there
a PkE-Pdd 1
ll"_".." ." . I.*_ . __ _"""._._ ". .I"
2 3 4
....."_". _.. . -.
orf309 -like
pktE
b
Multiple-Gene Duplications of PSTK Gene duplication is a potent mode of evolutionary adaptation (Ohno, 1999).Numerous examples of gene duplication followed by evolutionary modification of one or both of the duplicates are known throughout eubacteria, archaea, and eukaryotes (reviewed by Pao et al., 1998; Gilsdorf et al., 2004; and Irish and Litt, 2005). However, the PSTK family in M. xanthus is a unique example of diversity and size for a recognizable functional family of duplicated genes within a single genome. Determination of the diverse PSTK functions in M. xanthus is providing fundamental information on cell function and will be important in understanding how M. xanthus gained a large genome, with many genes involved in PSTK signaling systems together with TCST signaling systems. Below we describe the best-investigated duplication events.
pktF4 psdl Figure 3 Multiple gene duplication of pktE-pdd and psd-pktF. (a) Highly conserved regions in pktE-pdd duplications are shown by bars. A gray bar in pdd2 indicates a local duplication. Black, dark-gray, and light-gray arrows are p k t E , pdd, and an orf309-like ORF (having homology with orf309 in psd4-pktF2 in panel b), respectively. (b) Thick black, dark-gray, and light-gray arrows represent genes for PktF, Psd, and ORF309 homologues, respectively. Thin black and light-gray arrows indicate the genes for a putative phagerelated transcriptional regulator with H T H 3 type DNA binding domain (PF01381) and an ORF with unknown function, respectively. Putative hydrophobic sequences are shown by open ovals.
196 is another gene with similarity to orf309 (Akiyama and Komano, 2004) having weak similarity to the C-terminal half of bacterial small Ras-like GTP-binding proteins based on BLAST searches. The four trios of orf309-like open reading frame (ORF), PktE, and Pdd have over 95% amino acid and DNA identity, and each appears to form an operon. Based on the sequence analysis of the pktE-pdd regions, the region containing pktE3pdd3 appears to retain the most ancestral structure of the pktE-pdd duplications. Although we do not know the mechanism(s) by which these duplications occurred, the upstream region of pktE2-pdd2 contains a transposase gene while integrase and transposase genes are upstream of pktE4-pdd4. Similar duplication was also observed in the pktF and psd groups, although the gene organization of pktF-psd duplicates is more complex than the pktE-pdd duplication (Fig. 3b). Four of seven pktFs exist as a pair with four psds, while three pktFs are found alone. Interestingly, whereas orf309 identified previously by Akiyama and Komano (2004) was found downstream of psd4pktF2, its homologue, orf309-like ORF, was also found downstream of pktF7 and two pktF-psd pairs, psd5pktFl and psd3-pktF3, and upstream of pktF5. In addition to the orf309-like ORF, a putative transcriptional regulator with helix-turn-helix type 3 of DNA-binding domain (PF01381) is present in these regions with local duplications. In contrast to the pktE-pdd pairs, the RDs of PktF4 and PktF6 have apparently been lost by deletion and the other PktF RDs share less than 30% similarity at the amino acid level. The distribution of ORF309 homol o p e indicates that the pktE-pdd duplication family is likely derived from one of the pktF-psd duplicates. As in the case of pktE-pdd duplications, an integrase gene upstream of psd4-pktF2, and transposase and resolvase genes downstream of psd3-pktF3 were observed. Structurally, PktE and PktF are type I receptor PSTKs ( M l ) while Pdd and Psd are cytoplasmic PSTKs with short RDs ( S l ) . These PSTK pairs likely form kinase cascades, in a manner similar to that of PktC2/ Pkn8 to PskASIPknl4 (Nariya and Inouye, 2005b). In addition to the nine ORF309 homologues associated with PktE-Pdd and PktF-Psd, no other ORF309 homologue has been detected in M. xanthus or other representative bacterial genomes. ORF309 may play a specific role in M. xanthus physiology.
pktC Duplication Similar duplication events are also evident in the PktC group, which consists of 9 PSTKs (Fig. 2). PSTKs belonging to the PktC group have 8 to 10 tandem repeats of KLC-type TPR domain in their RDs downstream of
REGULATORY MECHANISMS the hydrophobic sequence, except for PktC1, in which the TPR domain seems to be truncated. Gene organization analysis shows that each member of the pktC gene family forms a gene cluster with a gene encoding an extracytoplasmic-function-typesigma factor, RpoE-like homologue (Erickson and Gross, 1989). Homologues of rpoE are located downstream of pktC2 and C3, upstream of pktC5, C8, and C9, and two genes upstream of C1. The rpoE homologue associated with pktC5 seems to be a pseudogene having a frame shift mutation by addition of a guanine residue in the middle, and there is no rpoE homologue associated with pktC4, C6, and C7. Thus, PktC may form a signaling pathway with an extracytoplasmic function sigma factor to regulate gene expression in specific circumstances. As in the pktE-pdd and pktF-psd duplications, transposase and integrase genes are also found in the upstream region of pktC2. One of the nine PktC has been shown to form a PSTK-PSTK cascade with a cytoplasmic PSTK. PktC2/ Pkn8, in conjunction with PskAS/Pknl4, regulates MrpC function, an essential transcriptional regulator for fruiting body formation and sporulation (Nariya and Inouye, 2006; see following section). The other PktCs may also form specific PSTK-PSTK cascades and/or modulate the PktC2-PskA5 cascade by forming heterodimer complexes with PktC2 via their TPR domains.
Gene Organization Adjacent to pstk Genes The role of PSTK genes and their importance can be investigated by assessing adjacent genes. We have observed that genes with diverse functions are present immediately adjacent to or near pstk genes, forming gene clusters. p s t k s Adjacent to p s t k Genes We previously found that pskA13/pknll is only 20 bp downstream of pskAl llpkn7 in the same orientation, forming an operon (Inouye et al., 2000). Similarly, pub7 and pktB9 are 76 and 78 bp upstream of abcl-1 and pktA6, respectively, in the same orientation and hence may form operons. In contrast, pktD7/pkn.5 and pskA2/pkn6 are found with opposite orientations and share a 128-bp common promoter region (Zhang et al., 1996). Several pstk pairs are separated by one to three genes; pktBl0 and pktB8 are found one and two genes upstream of pktA7 and pktA5, respectively. Another example is pab2, which has a histidine kinase domain and is three genes away from pktDl2, which contains the receiver domain of a TCST. In addition, we have already described sets of PktF-Psd and PktE-Pdd PSTK pairs arising from multiple-gene duplications.
11. PROTEINSER/THR KINASESAND PHOSPHATASESIN M . XANTHUS
pstks Adjacent to Phosphatase ( p p ) Genes Four p p genes that encode protein phosphatases in the phosphoprotein phosphatase (PPP) superfamily (see “Protein Phosphatases” below) are located close to pstk genes. One of them, MXAN7163, is located immediately downstream of pktC.5, forming an operon. Two genes, MXAN6S43 and MXANOSSS, are separated from pskB5 and pskA9 by one and two genes, respectively, and may form operons. Two genes located between MXANO.555 and pskA9 encode ATPase and ABC permease activities of a putative ABC transporter, activities that are possibly regulated by pskA9 and MXAN0.555 by reversible phosphorylation and dephosphorylation. Another gene, MXAN0267, is located in the opposite orientation to pskB4, separated by one unknown gene. In addition, pskA12 is located in the same orientation and probably in the same operon as a SpoIIE-like protein phosphatase (see below).
pstks Adjacent to oS4-DependentEBPs with FHA Domains As described in chapter 9, 12 enhancer-binding proteins (EBPs) have been predicted to contain the forkheadassociated (FHA) domain at their N terminus, and 6 of them are located adjacent to or near PSTKs (Jelsbak et al., 2005). Based on a genome analysis, a new gene ( M X A N 0 9 0 7 ) that encodes FHA-EBP was identified (Fig. 4), and this gene’s termination codon overlaps with the initiation codon of pktD3. The regulatory mechanism of a PSTK-EBP mediated by its FHA domain is described in chapter 9.
197
pstks Adjacent to Genes for Carbohydrate Metabolism Several genes required for carbohydrate metabolism are adjacent to or near the pstks. The pfk gene ( M X A N 4 0 16 ) that encodes 6-phosphofructokinase (PFK), a key enzyme for glycolysis, is located 18 bp upstream of pktDllpklz4, forming an operon. PFK activity was activated upon phosphorylation by PktD 1/Pkn4, and effective sporulation of M . xanthus requires glycogen consumption by activated PFK during development (see below) (Nariya and Inouye, 2002, 2003, 2 0 0 5 ~ )Simi. larly, glgA (glycogen synthase; M X A N 1 2 9 6 ) is located 35 bp upstream of pktA14, while the poly-3-hydroxybutyrate (PHB) depolymerase gene ( M X A N 6 3 1 3 ) is terminated by overlapping 4 bp with the initiation codon of pskB8. Another PHB depolymerase gene ( M X A N 0 0 1 6 ) is also located upstream of pskA16 in the same orientation, separated by one gene. PHB is another reservoir carbohydrate in prokaryotes (reviewed by Steinbiichel et al., 1992). Moreover, a gene for a-lY4-glucanase( M X A N 3694) for glycogen consumption is located 91 bp downstream of pskB.5. pktA1 is located between the mannose-1-phosphate guanylyltransferase (MXAN6SO1) and phosphomannomutase ( M X A N 6 4 9 9 ) genes for lipopolysaccharide synthesis in the same orientation, probably forming an operon. Since M . xanthus cannot utilize carbohydrates as a carbon source, sugar metabolites and their reservoir carbohydrates seem to be synthesized via gluconeogenesis using glucogenic amino acids and lipids (Bretscher and Kaiser, 1978). Thus, M. xanthus appears to uniquely develop the regulation of carbohydrate metabolism through PSTK signaling pathways.
Roles of PSTKs in the M. xunthus Life Cycle pktA6 Mx1288 pktBl0
pktA7
1 kP
-
Mx4901
MAXN0907 pktD3
Figure 4 FHA-EBP genes adjacent to or near pstks. Thick black and dark-gray arrows represent pstk and the gene for FHA-EBP, respectively. Thin light-gray arrows represent the genes for PFK, a-l,4-glucanase, and unknown functions.
After PktA2/Pknl was discovered and characterized for its role in the M. xanthus life cycle (Mufioz-Dorado et al., 1991), eight additional PSTKs were studied among the 13 PSTKs described by Inouye et al. (2000), PskA3/ MasK (Thomasson et al., 2002), and PktASIPknDl and PktBWPknD2 (Stein et al., 2006). Eight eukaryote Ser/ Thr/Tyr protein kinase inhibitors whose specificities are well documented have been tested for growth, motility, and developmental effects (Jain and Inouye, 1998).None of them had any effect on vegetative growth, while several inhibited development and sporulation to various degrees. The protein kinase C inhibitors, staurosporine, K-252c, and chelerythrin, and a Tyr kinase inhibitor, tyrphostin B52, are found to inhibit fruiting body development. Furthermore, sporulation is completely inhibited by K-252c, chelerythrin, and tyrphostin B52, but not by staurosporine, suggesting that staurosporine specifically
REGULATORY MECHANISMS
198 inhibits one branch required for the formation of the fruiting bodies but does not affect other branches leading to sporulation.
PktA2Pknl The first PSTK identified not only in M. xanthus but in all prokaryotes was pktA2/pknl (Mufioz-Dorado et al., 1991). The expression of pktA2/pkn1 is developmentally regulated and starts immediately prior to spore formation. Strains in which pktA2/pknl have been deleted show premature fruiting body formation and a 35% spore yield relative to the parent strain. PktA2/Pknl purified from E. coli is a Mg2+-dependentkinase and is autophosphorylated at both Ser and Thr.
PktD9mkn2 PktD9/Pkn2 is a transmembrane receptor-type PSTK ( M l ) with a cytoplasmic CD and a 207-residue C-terminal domain outside the cytoplasmic membrane. Disruption of pktD9/pkn2 has no effect on vegetative growth but causes faster development and reduces spore yield to 50 to 70% of that of the parent strain (Udo et al., 1995). In contrast to the pktD9/pkn2 disruption strain, a pktD9/pkn2 overexpression strain forms fruiting bodies more slowly than the parent strain but with a similar reduction in spore yield (Udo et al., 1996). PktD9/Pkn2 has a 28-fold-higher activity in the presence of Mn2+than Mg2+,and its activity was inhibited by H-7, a eukaryotic PSTK inhibitor, but not by genistein (Udo et al., 1997). Interestingly, when PktD9/Pkn2 was produced in E. coli, p-lactamase was found to serve as an effective substrate for PktD9/Pkn2 (Udo et al., 1995). Furthermore, a novel method to identify substrates using the toxic effect of PktD9/Pkn2 expression in E. coli was developed and found that E. coli histonelike proteins HUa and HUP act as suppressors that are phosphorylated at Thr-59 (Udo et al., 2000). As mentioned above, PktD9/Pkn2 contains a Class I11 AC domain (PF00211) in the cytoplasmic region followed by the CD. Class I11 ACs are commonly found in eukaryotes and some developmental and pathogenic bacteria. Class I11 ACs are evolutionally different from Class I ACs, exclusively found in enterobacteria (Linder and Schultz, 2003). AC synthesizes the most prevalent signaling molecule, CAMP, which plays important roles in regulating various cellular processes under the influence of diverse environmental stimuli in both prokaryotes and eukaryotes. Although the regulation and cellular functions of mammalian ACs have been well studied (reviewed by Linder and Schultz, 2003), those of prokaryotic Class I11 ACs are still unclear, especially their cellular functions. In M. xanthus, the intercellular level
of CAMPis known to rapidly increase during early development (Yajko and Zusman, 1978; Kimura et al., 2002). Biochemical studies revealed that AC activity of PktD9/ Pkn2 is greatly activated upon autophosphorylation at Thr residues in the Ala-Ser-Gly-Thr-rich region between the CD and AC domains. The activation of AC by phosphorylation occurs during development and is required for normal fruiting body formation with effective sporulation (Nariya and Inouye, unpublished results). M. xanthus contains another PSTK with a Class I11 AC domain, PktD12, which has a unique domain architecture found only in M . xanthus. PktD12 is located in the cytoplasm. It has a CD at the N terminus followed by the receiver domain of a TCST and then by a Class I11 AC domain. The receiver domain of PktD12 was found to associate with the histidine kinase domain of Pab2 by a genomic yeast two-hybrid screen (Nariya and Inouye, unpublished results). This suggests the intriguing possibility that the regulation of AC occurs by TCST and PSTK. If the receptor-type histidine kinase in Pab2 receives an environmental signal, it may activate AC by phosphorylating the receiver domain of PktD12, and Pab2 activity may also be regulated positively or negatively by the Ser/Thr kinase domain of PktD12.
PktD7/PknS and PskA2/Pkn6 The two pstk genes, pktD7/pknS and pskA2/pkn6, are oriented in opposite directions but share a 128-bp promoter region between their transcription initiation sites (Zhang et al., 1996). PktD7mkn5 consists of 380 amino acid residues with the insertion of 113 amino acid residues between subdomain V and VIa. It is a soluble PSTK in the cytoplasm, while PskA2/Pkn6 has 710 amino acid residues and is a transmembrane receptor-type PSTK. Purified from Escherichia coli, PktD7/Pkn5 is autophosphorylated at Ser while PskA2/Pkn6 is autophosphorylated at both Ser and Thr. Both genes are expressed constitutively throughout the M. xanthus life cycle, with slight increases early in development. PktD7Pkn5 and PskA2Pkn6 have reciprocal roles in M. xanthus growth and development, because a pktD7/pkn5 deletion strain forms fruiting bodies much faster than the parent strain, while a pskA2/kn6 deletion strain develops slower than the parent strain. The pktD7/pknS deletion strain is able to form fruiting bodies on semirich media, suggesting that PktD7/PknS negatively regulates M. xanthus development.
PktA4/Pkn9 PktA4/Pkn9 is a transmembrane receptor-type PSTK exposing a CD in the cytoplasmic space and a C-terminal RD in the periplasmic space. It is constitutively expressed during vegetative growth and upregulated during the
11. PROTEINSER/THRKINASES AND PHOSPHATASES IN M. XANTHUS aggregation stage of early development. The deletion of pktA4/pkn9 causes severely reduced development progression and spore formation. Two-dimensional gel analysis reveals that the deletion of pktA4/pkn9 prevents the expression of four unidentified membrane proteins, KREP9-1-4 (Hanlon et al., 1997).
PskAYMasK Isolation of pskA3ImasK was as a suppressor gene for an mglA mutant (Thomasson et al., 2002). The mglA gene encodes a 22-kDa GTPase critical for single-cell (A)gliding, type IV pilus-mediated (S) gliding, and M. xanthus development. S-motility and starvation-induced development can be restored by mas81 5, an allele-specific extragenic suppressor of mglA8, but it is unable to restore A-motility. pskA3/masK encodes a protein Ser/Thr/Tyr kinase and is in an operon immediately upstream of the mglBA operon. The interaction between PskA3lMasK and MglA was observed using a genomic yeast twohybrid screen. Tyr phosphorylation of PskA3lMasK in E. coli extracts has been shown by Western blot analysis using the anti-PskA3/MasK antibody and phosphoTyr antibody. However, PskA3lMasK purified from E. coli has autokinase activity to its own Ser/Thr residue(s), but not to its Tyr residue (Nariya and Inouye, unpublished results).
PktDl/Pkn4-PFK Cascade PktDUPkn4 is a PFK kinase, and the pfk gene forms an operon with pktDZlpkn4 (Nariya and Inouye, 2002). PFK is a key enzyme for glycogen metabolism in prokaryotes and eukaryotes, and its activity in eukaryotes is regulated by phosphorylation (Kemp et al., 1981; Foe and Kemp, 1982). By mutational analysis, PktDUPkn4 purified from E. coli is found to phosphorylate M. xanthus PFK at Thr-226, located in the putative allosteric effector site. Phosphorylation of PFK by PktDllPkn4 enhanced its activity 2.7-fold, and phosphorylation of PFK by PktDl/Pkn4 is almost completely inhibited by phosphoenolpyruvate, an allosteric inhibitor for PFK. The association of PFK with the regulatory domain of Pkn4 was demonstrated by immunoprecipitation using anti-Pkn4 immunoglobulin G, and this association is inhibited by phosphoenolpyruvate. M . xanthus accumulates glycogen during stationary phase and early in development (Nariya and Inouye, 2003). A pfk-pktDllpkn4 deletion strain accumulates glycogen at a higher level than the parent strain but is unable to consume glycogen during developmental progression and exhibits a poor spore yield. Based on a genetic complementation analysis of the pfk-pktD1l pkn4 deletion strain with the pfk and pktDllpkn4
199
genes, glycogen consumption and a high spore yield require not only the pfk gene but also the pktDllpkn4 gene. Furthermore, phosphorylation is critical for glycogen consumption because the pfk (Thr226Ala) gene mutated at the phosphorylation site did not complement a pfk mutant, suggesting that glycogen metabolism in M . xanthus is regulated in a manner similar to that in eukaryotes, requiring a PSTK (Nariya and Inouye, 2003).
p ktAS/pk n D 1 and p ktB 8/pknD2 Two pstk genes, pktASlpknD1 and pktB81pknD2, were identified during the course of sequencing the espAB region (Cho and Zusman, 1999). pktAUpknD2 is located upstream of the espAB operon and is oriented in the opposite direction. pktBWpknD2 is located immediately downstream of the espAB operon and is also oriented in the opposite direction. Based on hydrophobic sequence analysis, both PSTKs appear to be cytoplasmic kinases belonging to the S2 group (Table 1).The espAB operon regulates the timing of aggregation and sporulation. espA encodes a sensor histidine kinase with an FHA domain at the N terminus, and espB encodes a putative peptide transporter with 12 or 13 hydrophobic sequences (Cho and Zusman, 1999). The espA deletion mutant aggregates and sporulates much earlier than the wild type (DZ2)and forms a large number of small-sized fruiting bodies containing mature spores. Interestingly, numerous individual spores are found outside the fruiting bodies. espB deletion causes delays in aggregation and translucent mound formation, indicating defective sporulation. The espA-espB deletion mutant reveals that EspA is epistatic to EspB. The pktASlpknD2- or pktB8/ pknD2-deficient mutant forms translucent mounds and produces low spore yields, similar to the espB mutants (Stein et al, 2006). Double-mutant analysis reveals that espA is epistatic to pktA.5 and pktB8 to aggregation and fruiting body morphology, while pktA.5 and pktB8 are epistatic to sporulation efficiency. Expression of both pktASlpknD2 and pktB8lpknD2 is observed in vegetative growth and development. PktASIPknDl is phosphorylated at the Thr residue(s) in vitro, but PktB8lPknD2 is not autophosphorylated. However, PktB8/PknD2(Aspl62Asn), a phosphorylation-defective mutant, reveals a phenotype similar to that of the pktB8l pknD2-deficient mutant, indicating that PktB8lPknD2 functions as an active PSTK in vivo. The interaction of Tap-tagged PktASIPknDl or PktB8lPknD2 with EspA has been demonstrated by immunoprecipitation during development, suggesting that both PSTKs regulate fruiting body morphology and sporulation by interacting with EspA and EspB.
REGULATORY MECHANISMS
200
Mkaps and PktDl/Pkn4-PFK Cascade Three new factors, multikinase associated protein A (MkapA), MkapB, and MkapC, each of which contains well-known protein-protein interaction domains, and their associated PSTKs have been identified using a genomic yeast two-hybrid screen (Nariya and Inouye, 2005a). MkapA contains a Zn-finger-like domain that consists of CX,CX,HX,H with nine residues in a finger loop. Zn-fingers function as molecular recognition elements and are extremely common protein domains; perhaps 1% of all mammalian genes encode Zn-finger proteins (Mackay and Crossley, 1998). Interestingly, no MkapA homologues are found in the M. xanthus genome database or in any prokaryote genome sequence database so far determined. MkapB contains eight tandem repeats of a TPR domain. MkapB inhibits PFK activation in a phosphorylation-dependent manner by interfering with the association between MkapB and PFK. MkapC contains three repeats of a fibronectin type 3-like domain that is commonly found in mammalian proteins. A schematic diagram of the complex PSTK networks including the common modulating factors, MkapA, MkapB, and MkapC, is shown in Fig. 5. MkapA associates not only with PktDl/Pkn4 but also with other membrane-associated PSTKs, PktA2Rkn1, PktD9/Pkn2, PktC2/PltnSY and PktA4/Pkn9. MkapA may function as a regulator for the autokinase activity of PSTKs or as an adapter molecule for recruiting downstream factors in their signaling pathways. MkapB also associates with PktC2/PknS containing a TPR domain and PktA4/Pkn9
without a TPR domain; therefore, MkapB also functions as an adapter molecule for recruiting factors in the PktC2/Pkn8 and PktA4IPkn9 signaling pathway. MkapC associates with PktC2/Pkn8 in addition to PktDUPkn4. K9apl is an FHA protein with unknown function. Thus, environmental signals transmitted by membrane receptor-type PSTKs form complex networks with common modulating factors and regulate physiological functions of M. xanthus.
PktC2/PknS-PskAS/Pkn14 Cascade and Regulation of Essential Transcription Activators MrpC and FruA In addition to the Mkaps, PktC2IPkn8 associates with PskA5/Pknl4, a cytoplasmic PSTK. PktC2/Pkn8 phosphorylates PskA5IPknl4, suggesting that PktC2IPkn8 forms a kinase cascade with PskASIPknl4. In addition, PskASIPknl4 associates with MrpC (Nariya and Inouye, 2005a), an essential transcription factor for fruA expression (Ueki and Inouye, 2003). PskASIPknl4 is an MrpC kinase, with Thr-21 and/or Thr-22 as the likely site(s) of MrpC phosphorylation (Nariya and Inouye, 2005b). Both MrpC and FruA are transcriptional activators that are essential for multicellular fruiting body formation and sporulation (see chapter 9). Importantly, MrpC binding activity is greatly reduced upon its phosphorylation by PskASIPknl4. MrpC binds to at least eight sites in the upstream region of its promoter region (Nariya and Inouye, 2006). Based on analysis of MrpC binding sites in the mrpC and fruA promoter regions,
PSTK cascade
Figure 5 A signaling network of PSTKs sharing Mkaps in M. xanthus. Bars with arrowheads at both ends indicate interactions identified by the yeast two-hybrid screens. The T-bar is the inhibition of PFK phosphorylation by MkapB. Gray arrows are characterized phosphorylation pathways.
11. PROTEINSER/THR KINASESAND PHOSPHATASES IN M. XANTHUS there are two types of MrpC-specific binding sequences, A/GTTTC/GAA/G and GTGTCNNNNNNNGACAC. We have proposed a model for regulation of mrpC and fruA expression by a eukaryotic-like protein Ser/ Thr kinase cascade and prokaryotic two-component His-Asp phosphorelay system (Fig. 6 ) (Nariya and Inouye, 2006). As described by Sun and Shi (2001), mrpC is located downstream of mrpAB, encoding MrpA (histidine kinase) and MrpB (response regulator belonging to the NtrC family). A typical -24/-12 box for crS4is present in the upstream region of the mrpC promoter. During vegetative growth, the mrpC gene is likely transcribed at low levels by RNA polymerase with oS4 in the presence of basal levels of phosphorylated MrpB. The MrpC that is produced is phosphorylated by PskASIPknl4, activated by the PktC2IPkn8. The PktC2/ Pkn8-PskASIPknl4 kinase cascade negatively regulates mrpC expression by phosphorylation to prevent the
201
untimely initiation of development during vegetative growth. In early development, mrpAB expression is activated by environmental signals and produces MrpA and MrpB. MrpB, essential for mrpC expression, is activated by phosphorylation. Activated MrpB induces mrpC expression, and MrpC autoregulates its own expression. Since PskASIPkn14 expression is observed mainly during vegetative growth, the newly synthesized MrpC is likely not phosphorylated but instead processed to MrpC2, which has a higher affinity for the mrpC and fruA promoter regions. MrpC2 lacks the N-terminal 25 residues of MrpC and exhibits four- and eightfold-greater binding promoter regions, respecactivity to the mrpC and f r ~ A tively. Importantly, PskASIPknl4 is not able to phosphorylate MrpC2. Since MrpC2 was not detected in lonD mutant cells, LonD may play a role in the proteolytic processing of
I Vegetative growth I
I+
Developmental gene expression
I
I
Development
I
Fruiting body formation Sporulation
Figure 6 Regulation of mrpC and fruA expression in M . xanthus. PktC2JPkn8 and PskASJ Pknl4, forming a kinase cascade, and MrpA, a TCST histidine kinase, are highlighted in dark and light gray, respectively. See the text for details.
REGULATORY MECHANISMS
202 MrpC. LonD is the only protease known at present to be essential for fruiting body formation (Gill et al., 1993; Tojo et al., 1993). The accumulation of MrpC2 activates the expression of fruA, and FruA activates downstream genes involved in the regulation of aggregation, fruiting body formation, and sporulation. Therefore, fruiting body development and sporulation of 111. xanthus are achieved not only by the PSTK signaling system and a TCST system but also by PSTK networks and Mkaps.
Mutational Analysis of 102 PSTKs for Motility and Development In addition to the PSTKs described above, four PSTKs, PktA8mkn3, PskAl 1lPkn7, PskAl3lPkn11, and PskAlSI Pknl3, have been also characterized (S. Inouye, W. Zhang, M. Y. Hsu, R. Jain, and E. Farez-Vidal, unpublished results). To investigate the roles of the remaining 89 PSTKs, their plasmid-insertion mutants were constructed using plasmids carrying DNA fragments of the kinase catalytic core domain (subdomains I11 to IX) in Fig. 1. The disruption mutants were made in DZF1, which is proficient at fruiting body formation but has a partially impaired or leaky S-motility (+/- in Table 2). All 89 disruption mutants could be isolated, suggesting that all 102 PSTKs are not required during vegetative growth at 30°C in Casitone-yeast extract (CYE) medium. All mutants were examined for A- and S-motilities, fruiting body formation, and sporulation and were found to have A-motility, while three mutants, pktF4, pskA9, and pskB7, were defective in S-motility (-), forming a smooth edge at the end of colonies on 0.3% CYE agar plates (Table 2). Whereas the parent strain completes fruiting body formation in 2 days, the majority of the mutants showed defective phenotypes to various degrees including delayed fruiting body formation. pskA12 and abcl-2 mutants are stopped at the aggregation stage, and pktASlpknD1, pktB9, pktD6, pktE2, and pdd3 mutants are able to aggregate but cannot form mature fruiting bodies. pktA.5, pktD6, pskAl2, pdd3, and abcl-2 mutants are also defective in sporulation. The fruiting body morphology and sporulation of the pktA.5 mutant differ from those of the pktA.5-deficient mutant in DZ2 (A+S+)as described by Stein et al. (2006). These differences appear to be caused by the partially impaired Smotility of DZFl (A+S+’-).Six mutants, (pktB4, pktB6, pktD8/pkn12, pktF4, pktF6, and pab4) are able to form fruiting bodies with abnormal patterns. While the majority of mutants delay fruiting body formation, four mutants, pktA2/pknl, pktC2lPkn8, pktD7lpkn.5, and pskASIPknl4, accelerate fruiting body formation (Mufioz-Dorado et al., 1991; Zhang et al., 1996; Nariya and Inouye, 2005b). Since the pktD7lpkn.5 mutant is
Table 2 Developmental phenotypes and motility of pstk mutants Strain DZFl PktAS PktD6 PskA12 Pdd3 ABC1-2 PktA7 PktAl PktBl PktB4 PktB6 PktB8 PktB9 PktC7 PktC8 PktD8 PktE2 PktF4 PktF6 PskA9 PskA15 PskB7 PskB8 Pab4
FB formation (days)”
Sporulation (%)
2 def def def def def 5 4 4 def def 7 def 4 2 def def def def 7 7 4 7 def
100 0 0 0 0 0 10 7 3 2 2 1 0.1 2 6 1 0.3 7 10 80 40 85 40 2
S-motility +/-‘2
+/f/-
+/+/+/+/+/ff-
+/-
+/+/+/+/+/+/+/-
+/-
f/-
+/+/-
“Fruiting body formation. bSee text. Spore numbers were counted a t 7 days of development. ‘Deficient in FB formation.
able to form fruiting bodies on semirich medium, PktD71 PknS has been proposed to negatively regulate fruiting body development (Zhang et al., 1996).
PROTEIN PHOSPHATASES The finding of a large family of PSTKs in 111. xanthus raised the question of the existence of PPs. PPs are the counterpart of PSTKs, since they are required to dephosphorylate the substrates phosphorylated by the kinases. Identification of PPs in myxobacteria has been difficult; the first gene encoding a phosphatase was not cloned until 2001 (Treuner-Lange et al., 2001), 1 0 years after the report of the cloning of the first PSTK in M. xanthus. However, five patterns of phosphatase activity on p-nitrophenyl phosphate (PNPP) were detected in 1990 in this bacterium: two during vegetative growth and three during development. In addition to their expression profiles, the five activities could be differentiated by their dependency on magnesium, optimum pH, and inhibition by dithiothreitol (Weinberg and Zusman, 1990). None of these activities have been correlated with a
11. PROTEINSER/THR KINASESAND PHOSPHATASES IN 211. XANTHUS specific protein, so far. Neither is it known how many proteins are responsible for each activity. In an attempt to clone genes that encode proteins with phosphatase activity, three two-component regulatory systems of the family PhoRP have been identified in M. xanthus ( Carrero-Lirida et al., 2005; Moraleda-Muiioz et al., 2003). These three systems seem to be involved in the regulation of the expression of putative Mg2+-independent acid and neutral phosphatases, which are induced during development. Single- and double-deletion mutants for these systems exhibit reduced levels of these phosphatase activities and severe defects in both aggregation and sporulation in phosphate-free media (Carrero-LCrida et al., 2005; Moraleda-Muiioz et al., 2003), as described below.
The Protein Phosphatase Pphl Pphl is the only PP from M. xanthus that has been characterized to date (Treuner-Lange et al., 2001). It is a protein of 254 residues with a molecular weight of 28,308. It corresponds to MXAN2044 in the genome. This phosphatase belongs to the PP2C family and exhibits all the characteristics of this type of protein, such as dependency on Mn2+ and MgZ+,resistance to okadaic acid, and an ability to dephosphorylate phosphothreonine and phosphoserine. The gene that encodes this phosphatase was cloned in a search of proteins that interact with FrzZ by using the yeast two-hybrid system. The Frz system regulates the directed movement of cells during growth and development (seechapter 7 of this book). It consists of several proteins that are homologues of chemotaxis proteins of enteric bacteria. FrzZ contains two CheY-like domains connected by a linker rich in alanine and proline (Trudeau et al., 1996). As a result of the interaction between FrzZ and Pphl, a pphl mutant exhibits defects in cell motility as it spreads 10 to 30% less than the wild type on soft agar. This reduction in swarming seems to be originated by a defect in pilin transport, low levels of methylation of FrzCD, and low frequency of cell reversals. The pphl mutation also causes defects in late exponential growth and aggregation. Thus, mutant cells enter into stationary phase at low cell densities and originate abnormal fruiting bodies with a reduced number of myxospores. In addition to FrzZ, Pphl interacts with the protein kinase Pkn5l PktD7 in the yeast two-hybrid system, and this opens the door to the elucidation of the signal transduction pathways in which this kinase and phosphatase participate. To date, although there have been reported in M. xanthus several signal transduction pathways for kinases and phosphatases, in none of them have the two partners of the reversible phosphorylation been identified. Even in the case of Pphl, in spite of the reported interaction between
203
Pphl and FrzZ and PknSIPktD7, it remains unknown whether this interaction is mediated through the addition or removal of a phosphate group.
The Two-Component Systems of the Family PhoRP Three two-component systems of the family PhoRP were cloned while searching for genes that encode PPs in M . xanthus (Carrero-LCrida et al., 2005; Moraleda-Muiioz et al., 2003). These three systems are partially responsible for the expression of Mg2+-independent acid and neutral phosphatase activities, which are detected only during development. In addition, a fourth response regulator (PhoP4) has been found in the M. xanthus genome, which regulates the expression of the three developmentspecific phosphatase activities. This response regulator is orphan in the genome, but it has been demonstrated by using the yeast two-hybrid system that it interacts with the histidine kinase PhoR2 (Pham et al., 2006). It is so far unknown which genes encode the proteins responsible for phosphatase activities in M. xanthus extracts during vegetative growth and development. However, analysis of the genome seems to indicate that at least part of the activity on PNPP must be originated by specific PPs. Only one putative nonspecific acid phosphatase and two alkaline phosphatases are encoded by M. xanthus. If acid and neutral phosphatase activities on PNPP are partially due to PPs, the PhoRP systems would regulate the expression of these types of proteins. Nonetheless, it is known that pphl is not under control of any PhoRP system (CarreroLCrida et al., 2005; Moraleda-Muiioz et al., 2003). Single- and double-deletion mutants have been constructed in the PhoRP systems, and analyses of the mutants have revealed no effect on vegetative growth. On the contrary, these mutations cause severe defects in both aggregation and/or sporulation. Thus, single phoRP2 and phoRP3 mutants and the double-deletion mutant in these two systems originate flat fruiting bodies in phosphate-free media. However, cells are able to sporulate in these flat aggregates almost at the same level as the wild-type strain, although some myxospores fail to reshape completely, remaining as rod cells instead of becoming coccoids (Moraleda-Muiioz et al., 2003). A phoRPl deletion also causes defects in development, although they are not so dramatic. Thus, the mutant originates lower numbers of fruiting bodies than the wild-type strain, although this smaller number is correlated with a larger size. As a consequence, the number of spores is similar in both strains (Carrero-LCrida et al., 2005). On the contrary, a phoP4 mutant only originates a reduction in spore viability (Pham et al., 2006). It has still to be elucidated whether these phenotypes are the
REGULATORY MECHANISMS
204
result of the lower levels of expression of phosphatase activities or they are originated through defects in the expression levels of unknown genes that could be under the control of these systems.
PPs in the Genome PPs are more diverse in sequence than kinases. In fact, four major superfamilies of phosphatases exist (Shi et al., 1998): phosphoprotein phosphatases (PPP), Mn2+-or Mg2+-dependentprotein phosphatases (PPM), conventional phosphotyrosine phosphatases (CPTP), and low-molecular-mass phosphotyrosine phosphatases (LMMPTP).While PPP and PPM dephosphorylate phosphoserine and phosphothreonine residues, PTP dephosphorylate mainly phosphotyrosine residues. A group of PTP can dephosphorylate the three phosphoamino acids, and they have been named dual-specificity phosphatases. In addition, the four families of phosphatases exhibit differences in catalytic mechanism. Members of the PPP and PPM families, in spite of their sequence differences, are metalloenzymes which dephosphorylate their substrate in a single step through the activation of a water molecule by the metal. On the contrary, phosphatases belonging to the PTP families do not require metal, and dephosphorylation is performed in two steps, with the formation of a cysteinyl-phosphate enzyme intermediate (Barford et al., 1998). An inspection of the M . xanthus genome has revealed the presence of the four superfamilies of PPs, a fact that is observed only in a few prokaryotes, members of the PPP family being the most abundant. The total number of genes that encode PPs is 34, which corresponds to approximately one-third of the genes for kinases. However, this lower number of phosphatases than kinases seems to be a general rule among living organisms, prokaryotes and eukaryotes, rather than an exception. Among the 34 PPs, only four seem to be forming operons with protein kinases: three are PPPs, and the fourth one belongs to the PPM superfamily. The rest are either far from genes that encode kinases or encoded by complementary strands (see above).
The PPM Superfamily The PPM superfamily includes PP2C phosphatases, which are abundant in eukaryotes, and SpoIIE-like phosphatases, which have been reported only in the prokaryotes, with the exception of three found in Saccharomycetes (http://www.sanger.ac.uk/cgi-bin/Pfam/
speciesdist.pl?acc=PFO7228&id=SpoIIE&depth=all). The proteins of this superfamily consist of 11 subdomains, which contain the residues that are involved in the coordination of the metal ions of the active site (Shi, 2004). There are four genes in the M . xanthus genome
containing the Pfam for PP2C protein phosphatases (PF00481), including Pphl (Treuner-Lange et al., 2001). Three of these proteins (Table 3 ) have a number of amino acids that ranges from 247 to 265, approximately the size of the PP2C domain. On the contrary, M X A N 4 3 9 8 encodes a multidomain protein, with a cNMP-binding motif located in the C-terminal portion of the protein, and the PP2C consensus sequences in the amino terminus. This architecture has not been found in any other phosphatase and points to M X A N 4 3 9 8 participating in a signal transduction pathway where adenylyll guanylyl cyclases are implicated, responding to changes in the levels of cyclic nucleotides. PP2C phosphatases are not found in all the bacteria whose genomes have been sequenced. In those bacteria where PP2C phosphatases are encoded, their number ranges from one to three, with Anabaena sp. PCC 7120 (Kaneko et al., 2001) being the only bacterium, along with M . xanthus, where four PP2C phosphatases are present. In contrast, only one gene ( M X A N 4 5 6 2 ) has been found to encode a SpoIIE-like phosphatase (PF07228), a number that is extremely low if we compare M. xanthus with other bacteria that undergo developmental cycles, such as Bacillus subtilis, which carries 4 such genes (Duncan et al., 1995; Vijay et al., 2000; Yang et al., 1996), or S. coelicolor and Streptomyces avermitilis, which carry 48 and 4 7 such genes, respectively (Zhang and Shi, 2004b). The M . xanthus SpoIIE phosphatase exhibits a multidomain architecture, since it contains a HAMP domain in the middle portion of the protein and a SpoIIE domain in the carboxyl terminus (Table 3). This domain organization has also been reported in the phosphatase IcfG from Synechocystis (Beuf et al., 1994). M . xanthus SpoIIE seems to be in the same operon as the protein kinase, PskA12, so it is possible that they participate in the same signal transduction pathway, controlling the same process.
The PPP Superfamily The PPP superfamily of phosphatases has been defined by three domains, with the consensus sequences GDXHG, GDXXDRG, and GNHDIE, where X can be any amino acid (Zhang and Shi, 2004a; Shi et al., 1998). The search for these signature motifs in M . xanthus has revealed the existence of only two genes ( M X A N Z 5 0 9 and M X A N 5 4 6 7 ) . However, all phosphatases belonging to the PPP family contain the Pfam for metallophosphatases (PF00149, also named calcineurin-like phosphoesterases), which exhibits a logo that is not exactly identical to the sequence signatures defined by Shi and coworkers (http://www.sanger.ac.uk/cgi-bid Pfam/getacc?PF00149). The residues that are conserved in metallophosphatases are involved in metal binding,
11. PROTEINSEF~THR KINASESAND PHOSPHATASES IN M. XANTHUS
205
Table 3 List of protein phosphatases Superfamily PPM
Family and gene PP2C M X AN1412 MXAN2044 MXAN43 98 MXAN5349 SpoIIE MXAN4562
Size (aa)
Pfam
Location
247 254 442 265
PP2C PP2C PP2C PP2C
Cytoplasm Cytoplasm Cytoplasm Cytoplasm
521
SpoIIE
Membrane
PPP
MXANO149 M X ANO2 67 M X ANO344 MXAN0414 MXANO.555 MXAN0888 MXAN12 72 MXAN1509 MXAN2 613 MXAN3577 M X AN3 722 M X AN4086 MXAN4207 MXAN4.514 M X AN4 779 MXAN4 8 82 MXAN513 1 MXAN5467 MXAN6076 MXAN6105 MXAN6383 MXAN6.543 MXAN6890 MXAN6972 MXAN7163
417 367 245 3 79 276 465 290 235 310 485 260 398 399 3 75 316 234 256 339 217 276 426 336 389 300 230
Metallophosphatase Metallophosphatase Metallophosphatase Metallophosphatase Metallophosphatase Metallophosphatase Metallophosphatase Metallophosphatase Metallophosphatase Metallophosphatase Metallophosphatase Metallophosphatase Metallophosphatase Metallophosphatase Metallophosphatase Metallophosphatase Metallophosphatase Metallophosphatase Metallophosphatase Metallophosphatase Metallophosphatase Metallophosphatase Metallophosphatase Metallophosphatase Metallophosphatase
Cytoplasm Cytoplasm Cytoplasm Membrane Cytoplasm Membrane Cytoplasm Cytoplasm Cytoplasm Cytoplasm Cytoplasm Membrane Membrane Cytoplasm Cytoplasm Cytoplasm Cytoplasm Cytoplasm Cytoplasm Cytoplasm Membrane Cytoplasm Cytoplasm Cytoplasm Cytoplasm
CPTP
MXAN0419 MXAN0448 MXANl665
214 193 237
Dual-specificity PTP Dual-specificity PTP Dual-specificity PTP
Cytoplasm Cytoplasm Membrane
LMMPTP
M X ANO575
135
Low-molecular-mass PTP
Cytoplasm
which is required for dephosphorylation (Egloff et al., 1995). A search of the M. xanthus genome has revealed 33 genes encoding proteins that contain the Pfam for metallophosphatases. As the signature for this family of proteins is also found in proteins such as nucleotidases, sphingomyelin phosphodiesterases, and 2'-3' CAMP phosphodiesterases, as well as nucleases, it is difficult to estimate the exact number of PPP-type phosphatases in M. xanthus. However, as 6 of the 33 proteins exhibit
an architecture where the metallophosphatase domain is found with a nucleotidase domain, at least these six genes must be considered to encode proteins with nucleotidase activity (MXAN2236, 2661, 4313, 5325, 5361, and 6266). Another one (MXAN2579) encodes a multidomain protein, with a PKD signature sequence in the C-terminal region. PKD domains received their name because they were first identified in the polycystic kidney disease protein PKDl (Bycroft et al., 1999).
REGULATORY MECHANISMS
206 They are involved in protein-protein and proteincarbohydrate interactions and are usually located extracellularly. This location matches with the fact that the 111. xanthus protein exhibits a signal peptide sequence, which indicates that it most likely will be secreted to the periplasmic space. For this reason we have also excluded this protein as a PPP. Finally, gene MXAN09.58 contains only the metallophosphatase domain, but it shows a great similarity with the C subunit of the exonuclease Sbc. In addition, the gene is located before another one that encodes a protein with similarities to the D subunit of the same type of exonuclease. For these two reasons, MXANO958 has not been considered to be a PPP either. The other 25 genes can be considered to encode putative serinehhreonine phosphatases (Table 3 ) . The fact that the 25 proteins possess most of the residues that are involved in binding the catalytic metals also suggests that
they will most likely exhibit phosphoprotein phosphatase activity. A logo of the three domains defined for PPPs obtained with the sequences of the 25 M . xanthus proteins is shown in Fig. 7 (Schuster-Bockler et al., 2004). Only experimental characterization of these proteins will shed light on their activity and function. If all these genes really encode PPPs, this number will be the highest so far reported in a bacterium. Even if we consider the density of genes, expressed as the number of PPPs per megabase of genome, M . xanthus will exhibit the highest density in all living organisms, prokaryotes and eukaryotes. Only 5 of the 25 proteins exhibit transmembrane domains in the N-terminal portion of the protein, which indicates that they must be anchored to the membrane, with the catalytic domain located in the cytoplasm. This topology makes these proteins very interesting, as they may somehow sense signals and function as the first
Domain I dq
Domain I 1
Domain 111 A4
Figure 7 Logo of the three domains of the 25 putative PPP superfamily protein phosphatases from M. xanthus.
11. PROTEINSER/THRKINASESAND PHOSPHATASES IN M. XANTHUS component of the signal transduction pathways in which they participate. The rest of the proteins are all located in the cytoplasm. T h e PTPs As mentioned above there are two superfamilies of proteins with the ability to dephosphorylate phosphotyrosine residues, CPTP and LMMPTP. These two superfamilies share the sequence signature CX,R. However, they are distinguished by the position of a catalytic aspartate, which is found 25 to 50 residues in the N-terminal direction from the active site cysteine in CPTP and 80 to 110 residues in the C-terminal direction from the catalytic cysteine in LMMPTP (Shi et al., 1998). There are three members in the M. xanthus genome with the consensus sequences of the CPTP family (Table 3 ) that exhibit the Pfam for dual-specificity phosphatases (PF00782). Therefore, they will most likely dephosphorylate not only phosphotyrosine residues, but also phosphoserine and phosphothreonine. On the contrary, there is only one gene that encodes a protein with the signature sequence for LMMPTPs (PF01451).As a protein with the same signature sequence has been experimentally shown to confer arsenate resistance (Bennett et al., 2001), the capacity of the M . xanthus protein to desphosphorylate phosphotyrosine remains questionable. Two of the CPTPs and the LMMPTP are located in the cytoplasm, while the third CPTP is anchored to the membrane through two transmembrane domains located in the N-terminal region of the protein. The four PTPs are not found scattered along the chromosome, as it can be observed with the PPs of the other families. On the contrary, all of them are clustered in the first quarter of the genome. Characterization of these PPs will undoubtedly shed some light on the significance of tyrosine phosphorylation in M . xanthus.
CONCLUDING REMARKS The physiological roles of the PSTK signaling systems in M . xanthus are beginning to be understood at the molecular level. Their roles appear to be similar to those of the protein Ser/Thr and Tyr kinases in eukaryotes, known to regulate diverse cellular functions by forming kinase cascades with scaffold and adapter proteins. In M . xanthus, a receptor-type PSTK, PktC2/Pkn8, forms a kinase cascade with a cytoplasmic PSTK, PskASIPknl4, that negatively regulates mrpC expression during vegetative growth. This cascade prevents the untimely initiation of development (Nariya and Inouye, 2005b). Expression of mrpC during development is activated by MrpA and MrpB, a histidine kinase and a response regulator of a two-component His-Asp phosphorelay system (Sun and
207
Shi, 2001). Accumulation of MrpC induces fruA expression, producing FruA, a key transcription factor for both C-signal-independent and -dependent gene expression (Fig. 6; see also Fig. 4 in chapter 9). FruA is likely to activate directly dofA and fdgA expression and, indirectly, tps and sasA expression. However, they are not essential for fruiting body formation and sporulation, suggesting that FruA possibly regulates the expression of other essential genes for development (see chapter 9). Thus, a PSTK signaling cascade, together with a twocomponent His-Asp phosphorelay system, regulates the timely expression of developmentally essential genes, mrpC and fmA, to achieve a fruitful development. The PSTK signaling pathways in M. xanthus are also modulated by the formation of PSTK networks mediated by MkapA, B, and c, which associate with multiple PSTKs (Fig. 5 ) . The activation of PFK by PktDl/Pkn4 is inhibited by MkapB, which also associates with PktC21 Pkn8 and PktA4/Pkn9 (Nariya and Inouye, 2005a). MkapA associates with the kinase catalytic domain of PktDl/Pkn4 and PktC2/Pkn8, suggesting that PFK activation by PktDl/Pkn4 can be modulated at a specific period of vegetative growth and in late development, during which mkapA expression is observed (Nariya and Inouye, unpublished results). Therefore, M . xanthus PSTKs appear to regulate cellular functions by forming complex signaling networks with adapter proteins via protein-protein interaction domains. As described in chapter 9, M . xanthus contains 1 3 FHA-EBPs. In addition, 3 1proteins containing the FHA domain have been identified in the genome database (Nariya and Inouye, unpublished results). A majority of them have no known function, but two are associated with Class I11 adenylyl/guanylyl cyclases (MXAN3956 and 6571) and one with EspA (MXAN0931 [Cho and Zusman, 19991). This is another clue that PSTKs regulate protein function by forming complexes mediated by well-known protein-protein interaction domains. To understand how PSTKs regulate the function of FHAproteins, identification of their upstream PSTKs is essential. Various methods are now available to detect these protein-protein interactions. Using PSTKs as baits in genomic yeast two-hybrid screens has been well established as PknSapl, a FHA-protein, was identified using PktA4/Pkn9 as bait (Nariya and Inouye, 2005a) (see also Fig. 5 ) . A genomic yeast two-hybrid screen can be used to identify the upstream PSTKs of FHA-proteins. Proteomic and phosphoproteomic approaches are other choices that show promise but are thus far unproven with M . xanthus PSTKs. Interestingly, a putative physiological substrate with the FHA domain, GarA, of PknB of M. tuberculosis was identified using the
208 proteomic approach (Villarino et al., 2005). Moreover, three physiological substrates of PknA and PknB of 111. tuberculosis were identified based on peptide library screening for their preferred phosphorylation motives (Kang et al., 2005). The roles of the upstream PSTKs can be addressed by identifying the downstream genes regulated by FHA-EBPs. The microarray expression profiling of nlal8, an FHA-EBP mutant (see Table 3 in chapter 9), has been demonstrated (Diodati et al., 2006). Analysis of the downstream genes’ expression and gene products will elucidate how PSTKs regulate FHA-EBP by phosphorylation and modulate cellular functions. Although mutational analysis showed that the majority of PSTKs were not essential for fruiting body formation and sporulation under the conditions used, they must be important in their natural habitats. The uniquely large number of PSTKs containing various functional domains in M . xanthus suggests an unusually flexible regulatory capacity in their signaling systems. Furthermore, the pattern and level of pstk gene duplications suggests that PSTKs have acquired many new functions, which have allowed refinement of preexisting functional capacities. Although a large number of PPs also exist in M . xanthus and they are known to play critical roles in PSTK signaling pathways, the interactions between PPs and PSTKs have not been well established in M. xanthus and their reversible phosphorylation remains to be elucidated. Further study of the PSTK signaling networks with Mkaps, PSTK-associated proteins, and PPs will lead us to understand the complicated signaling pathways, mediated not only by PSTKs but also by His-Asp phosphorelay systems, which govern the unique physiology of M . xanthus. We are grateful to M. Travisano for comments on the first part the manuscript. This work has been supported by grants from the Foundation of University of Medicine and Dentistry of New Jersey for S.I. and from the Ministerio de Ciencia y Tecnologia, Spain, for J.M.-D. (grant BMC2003-02038). of
References Adams, J. A., M. L. McGlone, R. Gibson, and S. S. Taylor. 1995. Phosphorylation modulates catalytic function and regulation in the CAMP-dependent protein kinase. Biochemistry 34:2447-2454. Akiyama, T., and T. Komano. 2004. Analysis of fruE, a novel developmental gene of Myxococcus xanthus. J. Mol. Microbiol. Biotechnol. 6:164-173. Av-Gay, Y., and M. Everett. 2000. The eukaryotic-like Serl Thr protein kinases of Mycobacterium tuberculosis. Trends Microbiol. 8:238-244. Barford, D., A. K. Das, and M.-P. Egloff. 1998. The structure and mechanism of protein phosphatases: insights into catalysis and regulation. Annu. Rev. Biophys. Biomol. Struct. 27:133-164.
REGULATORY MECHANISMS Bennett, M. S., Z. Guan,M. Laurber, andX.-D. Su. 2001. Bacillus subtilis arsenate reductase is structurally and functionally similar to low molecular weight protein tyrosine phosphatases. Proc. Natl. Acad. Sci. USA 98:13577-13582. Beuf, L., S. Bedu, M. C. Durand, and F. Joset. 1994. A protein involved in co-ordinated regulation of inorganic carbon and glucose metabolism in the facultative photoautotrophic cyanobacterium Synechocystis PCC6803. Plant Mol. Biol. 25:855-864. Bretscher, A. P., and D. Kaiser. 1978. Nutrition of Myxococcus xanthus, a fruiting myxobacterium. J. Bacteriol. 133:763768. Bycroft, M., A. Bateman, J. Clarke, S. J. Hamill, R. Sandford, R. L. Thomas, and C. Chothia. 1999. The structure of a PKD domain from polycystin-1: implications for polycystic kidney disease. EMBO J. 18:297-305. Carrero-Lkrida, J., A. Moraleda-Muiioz, R. Garcia-Hernandez, J. Perez, and J. Muiioz-Dorado. 2005. PhoR1-PhoP1, a third two-component system of the family PhoRP from Myxococcus xanthus: role in development. J. Bacteriol. 187:4976-49 83. Chang, C., A. Mooser, A. Pluckthun, and A. Wlodawer. 2001. Crystal structure of the dimeric C-terminal domain of TonB reveals a novel fold. J. Biol. Chem. 276:27535-27540. Cho, K., and D. R. Zusman. 1999. AsgD, a new twocomponent regulator required for A-signalling and nutrient sensing during early development of Myxococcus xanthus. Mol. Microbiol. 34:268-281. Diodati, M. E., F. Ossa, N. B. Caberoy, I. R. Jose, W. Hiraiwa, M. M. Igo, M. Singer, and A. G. Garza. 2006. Nla18, a key regulatory protein required for normal growth and development of Myxococcus xanthus. J. Bacteriol. 188:1733-1743. Duncan, L., S. Alper, F. Arigoni, R. Losick, and P. Stragier. 1995. Activation of cell-specific transcription by a serine phosphatase at the site of asymmetric division. Science 270:641-644. Egloff, M. P., P. T. W. Cohen, P. Reinemer, and D. Barford. 1995. Crystal structure of the catalytic subunit of human protein phosphatase 1 and its complex with tungstate. J. Mol. Biol. 254~942-959. Erickson, J. W., and C. A. Gross. 1989. Identification of the sigma E subunit of Escherichia coli RNA polymerase: a second alternate sigma factor involved in high-temperature gene expression. Genes Dev. 3:1462-1471. Foe, L. G., and R. G. Kemp. 1982. Properties of phospho and dephospho forms of muscle phosphofructokinase. J. Biol. Chem. 257:6368-6372. Frasch, S. C., and M. Dworkin. 1996. Tyrosine phosphorylation in Myxococcus xanthus, a multicellular prokaryote. J. Bacteriol. 178:4084-4088. Gill, R. E., M. Karlok, and D. Benton. 1993. Myxococcus xanthus encodes an ATP-dependent protease which is required for developmental gene transcription and intercellular signaling. J. Bacteriol. 175:4538-4544. Gilsdorf, J. R.,C. F. Mars, and B. Foxman. 2004. Haemophilus influenzae: genetic variability and natural selection to identify virulence factors. Infect. Immun. 72:2457-2461. Goldman, B. S., W. C. Nierman, D. Kaiser, S. C. Slater, A. S. Durkin, J. A. Eisen, C. M. Ronning, W. B. Barbazuk, M. Blanchard, C. Field, C. Halling, G. Hinkle, 0. Iartchuk,
11. PROTEINSEF~THR KINASESAND PHOSPHATASES IN M. XANTHUS H. S. Kim, C. Mackenzie, R. Madupu, N. Miller, A. Shvartsbeyn, S. A. Sullivan, M. Vaudin, R. Wiegand, and H. B. Kaplan. 2006. Evolution of sensory complexity recorded in a myxobacterial genome. Proc. Natl. Acad. Sci. USA 103:15200-15205. Hanks, S. K., A. M. Quinn, and T. Hunter. 1988. The protein kinase family: conserved features and deduced phylogeny of the catalytic domains. Science 241:42-52. Hanks, S. K., and T. Hunter. 1995. The eukaryotic protein kinase superfamily: kinase domain structure and classification. FASEBJ. 9576-596. Hanlon, W. A., M. Inouye, and S. Inouye. 1997. Pkn9, a Ser/ Thr protein kinase involved in the development of Myxococcus xanthus. Mol. Microbiol. 23:459-471. Inouye, S., R. Jain, T. Ueki, H. Nariya, C. Y. Xu, M. Y. Hsu, B. A. Fernandez-Luque, J. Muiioz-Dorado, E. Farez-Vidal, and M. Inouye. 2000. A large family of eukaryotic-like protein Ser/Thr kinases of Myxococcus xanthus, a developmental bacterium. Microb. Comp. Genomics 5:103-120. Irish, V. F., and A. Litt. 2005. Flower development and evolution: gene duplication, diversification and redeployment. Curr. Opin. Genet. Dev. 15:454-460. Jain, R., and S. Inouye. 1998. Inhibition of development of M~XOCOCCUS xanthus by eukaryotic protein kinase inhibitors. J. Bacteriol. 180:6544-6550. Jelsbak, L., M. Givskov, and D. Kaiser. 2005. Enhancer-binding proteins with a forkhead-associated domain and the sigma54 regulon in Myxococcus xanthus fruiting body development. Proc. Natl. Acad. Sci. USA 102:3010-3015. Kaneko, T., Y. Nakamura, C. P. Wolk, T. Kuritz, S. Sasamoto, A. Watanabe, M. Iriguchi, A. Ishikawa, K. Kawashima, T. Kimura, Y. Kishida, M. Kohara, M. Matsumoto, A. Matsuno, A. Muraki, N. Nakazaki, S. Shimpo, M. Sugimoto, M. Takazawa, M. Yamada, M. Yasuda, and S. Tabata. 2001. Complete genomic sequence of the filamentous nitrogenfixing cyanobacterium Anabaena sp. strain PCC 7120. D N A Res. 8:205-213. Kang, C. M., D. W. Abbott, S. T. Park, C. C. Dascher, L. C. Cantley, and R. N. Husson. 2005. The Mycobacterium tuberculosis serinelthreonine kinases PknA and PknB: substrate identification and regulation of cell shape. Genes Dev. 19:1692-1704. Kemp, R.G., L. G . Foe, S. P. Latshaw, R. A. Poorman, and R. L. Heinrikson. 1981. Studies on the phosphorylation of muscle phosphofructokinase. J. Biol. Chem. 256:7282-7286. Kimura, Y., Y. Mishima, H. Nakano, and K. Takegawa. 2002. An adenylyl cyclase, CyaA, of Myxococcus xanthus functions in signal transduction during osmotic stress. J. Bacterial. 184:3578-3585. Laronde-LeBlanc, N., and A. Wlodawer. 2005. The RIO kinases: an atypical protein kinase family required for ribosome biogenesis and cell cycle progression. Biochim. Biophys. Acta 1754:14-24. Lee, P. C., T. Umeyama, and S. Horinouchi. 2002. afsS is a target of AfsR, a transcriptional factor with ATPase activity that globally controls secondary metabolism in Streptomyces coelicolor A3(2).Mol. Microbiol. 43:1413-1430. Linder, J. U., and J. E. Schultz. 2003. The class I11 adenylyl cyclases: multi-purpose signalling modules. Cell. Signal. 15~1081-1089.
209
Mackay, J. P., and M. Crossley. 1998. Zinc fingers are sticking together. Trends Biochem. Sci. 23:l-4. Manning, G., D. B. Whyte, R. Martinez, T. Hunter, and S. Sudarsanam. 2002. The protein kinase complement of the human genome. Science 298:1912-1934. Moraleda-Muiioz, A., J. Carrero-Ltrida, J. Perez, and J. Muiioz-Dorado. 2003. Role of two novel two-component regulatory systems in development and phosphatase expression in Myxococcus xanthus. J. Bacteriol. 185:1376-1383. Muiioz-Dorado, J., S. Inouye, and M. Inouye. 1991. A gene encoding a protein serinekhreonine kinase is required for normal development of M . xanthus, a gram-negative bacterium. Cell 67:995-1006. Nariya, H., and S. Inouye. 2002. Activation of 6-phosphofructokinase via phosphorylation by Pkn4, a protein Ser/Thr kinase of Myxococcus xanthus. Mol. Microbiol. 46:13531366. Nariya, H., and S. Inouye. 2003. An effective sporulation of Myxococcus xanthus requires glycogen consumption via Pkn4-activated 6-phosphofructokinase. Mol. Microbiol. 49:5 17-528. Nariya, H., and S. Inouye. 2005a. Modulating factors for the Pkn4 kinase cascade in regulating 6-phosphofructokinase in Myxococcus xanthus. Mol. Microbiol. 56:1314-1328. Nariya, H., and S. Inouye. 2005b. Identification of a protein SeriThr kinase cascade that regulates essential transcriptional activators in Myxococcus xanthus development. Mol. Microbiol. 5 8:3 67-3 79. . that modulate the Nariya, H., and S. Inouye. 2 0 0 5 ~ Factors Pkn4 kinase cascade in Myxococcus xanthus. J. Mol. Microbiol. Biotechnol. 9:147-153. Nariya, H., and S. Inouye. 2006. A protein SerKhr kinase cascade negatively regulates the DNA-binding activity of MrpC, a smaller form of which may be necessary for the Myxococcus xanthus development. Mol. Microbiol. 60:1205-1217. Ohno, S. 1999. Gene duplication and the uniqueness of vertebrate genomes circa 1970-1999. Semin. Cell Dev. Biol. 10~517-522. Pao, S. S., I. T. Paulsen, and M. H. Saier, Jr. 1998. Major facilitator superfamily. Microbiol. Mol. Biol. Rev. 62:l-34. Petrickova, K., and M. Petricek. 2003. Eukaryotic-type protein kinases in Streptomyces coelicolor: variation on a common theme. Microbiology 1491609-1621. Pham, V. D., C. W. Shebelut, I. R. Jose, D. A. Hodgson, D. E. Whitworth, and M. Singer. 2006. The response regulator PhoP4 is required for late developmental events in Myxococcus xanthus. Microbiology 152:1609-1620. Schuster-Bockler, B., J. Schultz, and S. Rahmann. 2004. HMM Logos for visualization of protein families. BMC Bioinformatics 5:7-15. Shi, L., M. Potts, and P. J. Kennelly. 1998. The serine, threonine, and/or tyrosine-specific protein kinases and protein phosphatases of prokaryotic organisms: a family portrait. FEMS Microbiol. Rev. 22:229-253. Shi, L. 2004. Manganese-dependent protein 0-phosphatases in prokaryotes and their biological functions. Front. Biosci. 9: 1382-1 397. Stein, E. A., K. Cho, P. I. Higgs, and D. R. Zusman. 2006. Two Ser/Thr protein kinases essential for efficient
210 aggregation and spore morphogenesis in Myxococcus xanthus. Mol. Microbiol. 60:1414-1431. Steinbiichel, A., E. Hustede, M. Liebergesell, U. Pieper, A. Timm, and H. Valentin. 1992. Molecular basis for biosynthesis and accumulation of polyhydroxyalkanoic acids in bacteria. FEMS Microbiol. Rev. 9:217-230. Stock, A. M., V. L. Robinson, and P. N. Goudreau. 2000. Two-component signal transduction. Annu. Rev. Biochem. 69~183-215. Sun, H., and W. Shi. 2001. Analyses of mrp genes during Myxococcus xanthus development. J. Bacteriol. 183:67336739. Thomasson, B., J. Link, A. G. Stassinopoulos, N. Burke, L. Plamann, and P. L. Hartzell. 2002. MglA, a small GTPase, interacts with a tyrosine kinase t o control type IV pilimediated motility and development of Myxococcus xanthus. Mol. Microbiol. 46:1399-1413. Tojo, N., S. Inouye, and T. Komano. 1993. The lonD gene is homologous to the lon gene encoding an ATP-dependent protease and is essential for the development of Myxococcus xanthus.J. Bacteriol. 175:4545-4549. Treuner-Lange, A., M. J. Ward, and D. R. Zusman. 2001. Pphl from Myxococcus xanthus is a protein phosphatase involved in vegetative growth and development. Mol. Microbiol. 40~126-140. Trudeau, K. G., M. J. Ward, and D. R. Zusman. 1996. Identification and characterization of FrzZ, a novel response regulator necessary for swarming and fruiting-body formation in Myxococcus xanthus. Mol. Microbiol. 20:645-655. Udo, H., C. K. Lam, S. Mori, M. Inouye, and S. Inouye. 2000. Identification of a substrate for Pkn2, a protein Ser/ Thr kinase from Myxococcus xanthus by a novel method for substrate identification. J. Mol. Microbiol. Biotechnol. 2557-563. Udo, H., J. Muiioz-Dorado, M. Inouye, and S. Inouye. 1995. Myxococcus xanthus,a gram-negative bacterium, contains a transmembrane protein serinekhreonine kinase that blocks the secretion of beta-lactamase by phosphorylation. Genes Dev. 9:972-983. Udo, H., M. Inouye, and S. Inouye. 1996. Effects of overexpression of Pkn2, a transmembrane protein serinekhreonine kinase, on development of Myxococcus xanthus. J. Bacterial. 178:6647-6649. Udo, H., M. Inouye, and S. Inouye. 1997. Biochemical characterization of Pkn2, a protein Ser/Thr kinase from Myxococcus xanthus, a Gram-negative developmental bacterium. FEBS Lett. 400:188-192.
REGULATORY MECHANISMS Ueki, T., and S. Inouye. 2003. Identification of an activator protein required for the induction of fruA, a gene essential for fruiting body development in Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 100:8782-8787. Vijay, K., M. S. Brody, E. Fredlund, and C. W. Price. 2000. A PP2C phosphatase containing a PAS domain is required to convey signals of energy stress to the sigmaB transcription factor of Bacillus subtilis. Mol. Microbiol. 35:180-188. Villarino, A., R. Duran, A. Wehenkel, P. Fernandez, P. England, P. Brodin, S. T. Cole, U. Zimny-Arndt, P. R. Jungblut, C. Cervenansky, and P. M. Alzari. 2005. Proteomic identification of M. tuberculosis protein kinase substrates: PknB recruits GarA, a FHA domain-containing protein, through activation loop-mediated interactions. J . Mol. Biol. 350:953-963. Walburger, A., A. Koul, G. Ferrari, L. Nguyen, C. PrescianottoBaschong, K. Huygen, B. Klebl, C. Thompson, G. Bacher, and J. Pieters. 2004. Protein kinase G from pathogenic mycobacteria promotes survival within macrophages. Science 304:1800-1804. Wang, L., Y. P. Sun, W. L. Chen, J. H. Li, and C. C. Zhang. 2002. Genomic analysis of protein kinases, protein phosphatases and two-component regulatory systems of the cyanobacterium Anabaena sp. strain PCC 7120. FEMS Microbiol. Lett. 21E155-165. Weinberg, R. A., and D. R. Zusman. 1990. Alkaline, acid, and neutral phosphatase activities are induced during development in Myxococcus xanthus. J. Bacteriol. 172:2294-2302. Yajko, D. M., and D. R. Zusman. 1978. Changes in cyclic AMP levels during development in Myxococcus xanthus. J. Bacteriol. 133:1540-1542. Yang, X., C. M. Kang, M. S. Brody, and C. W. Price. 1996. Opposing pairs of serine protein kinases and phosphatases transmit signals of environmental stress to activate a bacterial transcription factor. Genes Dev. 10:22652275. Zhang, W., and L. Shi. 2004a. Comparative analysis of eukaryotic-type protein phosphatases in two streptomycete genomes. Microbiology 150:2247-2256. Zhang, W., and L. Shi. 2004b. Evolution of the PPM-family protein phosphatases in Streptomyces: duplication of catalytic domain and lateral recruitment of additional sensory domains. Microbiology 150:4189-4197. Zhang, W., M. Inouye, and S. Inouye. 1996. Reciprocal regulation of the differentiation of Myxococcus xanthus by Pkn5 and Pkn6, eukaryotic-like Ser/Thr protein kinases. Mol. Microbiol. 20:435-447.
Myxobacteria: Mtrlticellularity and Differentiation Edited by David E. Whitworth 02008 ASM Press, Washington, D.C.
Montserrat Elias-Arnanz Marta Fontes S. Padmanabhan
Carotenogenesis in Myxococcus xanthus:a Complex Regulatory Network
CAROTENOIDS Myxobacteria frequently occur as brightly colored colonies and sporangioles, due to the presence of carotenoids and/or other pigments (Reichenbach and Kleinig, 1984; Hodgson and Murillo, 1993; Hodgson and Berry, 1998). The carotenoid pigments, which range in color from light yellow to deep red, form a major class of lipophilic isoprenoids that include the hydrocarbon carotenes with acyclic, monocyclic, or bicyclic ends and their oxygenated (hydroxy, aldehydic, keto, carboxyl, methoxy, oxy, epoxy, and glycosidic) derivatives known as xanthophylls. Most natural carotenoids are C,, terpenes derived from eight isoprenoid units (although some C3,, C45, and C,, have been reported), the colorless C40 phytoene being the universal progenitor in their biosynthesis. The color of a given carotenoid is determined by the number of conjugated double bonds, which exist mostly in the all-trans conformation except for phytoene, which often occurs as the 15,15’-cis isomer. The carotenoids serve important biological roles and are widely distributed in nature, being synthesized de novo in anoxygenic photosynthetic bacteria, cyanobacteria, some nonphotosynthetic bacteria, some fungi, and all algae and plants, but need to be provided as dietary supplements in animals
12
( Goodwin, 1980). Being relatively hydrophobic, carotenoids are typically associated with membranes and may or may not be specifically bound to proteins. Their primary function is to protect cells against photo-oxidative damage by quenching the triplet excited states of photosensitizer molecules such as chlorophyll and porphyrins and the singlet excited state of oxygen that result from absorption of light energy. They also play roles in light harvesting and as redox intermediates in the shuttling of electrons in photosynthesis (Frank and Brudvig, 2004). Carotenoids also serve as precursors for molecules required for photoreception and hormonal action, the classic example being the production of retinoids from carotenes. The chemistry and biosynthesis of carotenoids, their species-specific distribution, and their cellular functions and localization have been extensively discussed in various other reviews, including those with an emphasis on carotenoids in eubacteria in general (Armstrong, 1997) or specifically on myxobacteria (Reichenbach and Kleinig, 1984; Hodgson and Murillo, 1993; Hodgson and Berry, 1998). Almost 1 decade has passed since the last review on carotenogenesis in Myxococcus xanthus was published, undoubtedly one of the most important
Montserrat Elias-Arnanz, Marta Fontes, and S. Padmanabhan, Departamento de GenCtica y Microbiologia, Facultad de Biologia, Universidad de Murcia, 30100 Murcia, Spain.
211
212
model systems for the genetic analysis of light-induced carotenogenesis. The aim of the present review is to provide an update on the considerable amount of new information that has since been acquired with respect to the inducing signals, their reception and transduction, and the transcriptional regulation of the structural genes involved in carotenoid biosynthesis in M. xanthus.
CAROTENOID SYNTHESIS IN M . XANTHUS-AN INDUCIBLE RESPONSE A Brief Overview of the Known Genetic Elements Involved Early work showed that carotenoid production in M. xanthus is a response induced on exposure to blue light (Burchard and Dworkin, 1966). No carotenoids or their precursors are found in dark-grown M. xanthus cultures (Martinez-Laborda et al., 1990), which appear yellow due to noncarotenoid pigments, named DKxanthenes, that have only recently been characterized (Meiser et al., 2006). In the light, cells turn red due to production of carotenoids, and this wild-type, light-inducible phenotype has been designated Car+.Loss of the ability to synthesize carotenoids would maintain the yellow color under all conditions (Car- phenotype), while their constitutive production would keep cells always red ( Carcphenotype). By means of this conspicuous color change, spontaneous as well as chemical-, W-,and transposon-induced mutants were isolated. While Car' mutants would be affected in the regulation of light-induced carotenogenesis, Carmutants could result either from the loss or from the inactivation of an enzyme essential for a step leading to the synthesis of a colored carotene or of a positive regulator of carotenogenesis. Analysis of Car- and Carc mutants obtained primarily by transposon insertions allowed the early mapping and identification of structural and regulatory genes, and their epistatic analysis. These were later extended by chemical analysis of the carotenoid content of the mutants, gene expression assays, and cloning and sequencing of the affected loci. More recently, in silico analysis of the M. xanthus genome or the study of proteins that physically interact with those previously identified has allowed the identification of new elements involved in the light induction of carotenoid synthesis. Also, a number of the corresponding gene products have been subjected to detailed molecular analysis of their domain organization, structures, and modes of action. The designation crt has been employed for structural genes that encode the enzymes with known or putative functions in the carotenoid biosynthetic pathway, and the cur notation has been used for regulatory genes and for all operons, including those for structural genes. Figure 1
REGULATORY MECHANISMS summarizes our present knowledge of the genetic elements involved and their interactions. Structural genes for carotenoid synthesis are located at two unlinked loci: crtlb and carBA, the latter of which is constituted by two contiguous operons (curl3 and curA). The carB operon groups six structural genes, and its expression is controlled by a light-inducible promoter (Martinez-Laborda et al., 1986, 1989, 1990; Balsalobre et al., 1987; Ruiz-VAzquez et al., 1993; Botella et al., 1995). Immediately downstream of carB lies the carA operon, made up of structural and regulatory genes which are expressed in a light-independent manner (Martinez-Laborda et al., 1986; Hodgson, 1993; Botella et al., 1995). The unlinked crtIb gene is under the control of a light-inducible promoter but is fully activated only in stationary-phase cells (Balsalobre et al., 1987; Fontes et al., 1993). The light-dependent expression of carB and crtIb is controlled by the products of the carQRS operon, constituted by three translationally coupled genes whose expression is also induced by light (Hodgson, 1993; McGowan et al., 1993). CarQ and CarR form an ECF (extracytoplasmic function) sigma-antisigma pair that acts in regulating the expression of carQRS itself and crtlb (Gorham et al., 1996; Martinez-Argudo et al., 1998; Browning et al., 2003; Whitworth et al., 2004). Cars activates carB expression in the light by counteracting the action of CarA, which is a repressor of carB in the dark and is encoded by a gene in the carA operon (Whitworth and Hodgson, 2001; Cervantes and Murillo, 2002; L6pez-Rubio et al., 2002, 2004; Perez-Marin et al., 2004; Navarro-AvilCs et al., 2007). Other regulatory factors, all expressed independent of light, include (i)CarF, which may be a receiver or transmitter of the light signal (Fontes et al., 2003); (ii) CarD, a global regulator of transcription similar to the eukaryotic high-mobility group A proteins (Nicolh et al., 1994; 1996; Padmanabhan et al., 2001; Cayuela et al., 2003); (iii) CarG, a recently identified protein that acts in concert with CarD (Pefialver et al., 2006); and (iv) the product of ihfA, the ortholog of the Escherichia coli integration host factor (Moreno et al., 2001). Our present understanding of each of these structural and regulatory genes and the specific functions of their products will be discussed in the following sections.
The Structural Genes and Their Functions As mentioned earlier, structural genes for carotenoid synthesis cluster at the carBA locus, with the exception of crtlb. Their activitieswere assigned by analyzing the carotenoids accumulated in different mutants, from sequence homology to previously characterized genes, and by heterologous expression in E. coli (Martinez-Laborda et al.,
12. CAROTENOGENESIS IN 211. XANTHUS
213 copper
blue-light
h crtE-la-6-D-C orf6 )IcrtYc-Yd orf9 carA orfll car6operon
Y
carA operon
Figure 1 Known genetic elements involved in carotenogenesis and their interactions. Squat arrows show photoinducible genes (crtlb)or operons (carQRS and the carB-carA cluster with the constituent genes indicated) implicated in the carotenogenic pathway. Genes for other proteins involved whose expression is not light dependent are not shown. Labeled ovals are the regulatory factors that have been identified and characterized, with continuous arrows indicating positive regulation, and blunt-ended lines indicating negative regulation. Carotenoid biosynthesis enzymes are encoded by crtlb, all of the genes in the carB operon, and crtYc and crtYd in the carA operon. The latter operon also codes for the regulatory factors CarA and O r f l l . See the text for further explanation.
1990; Ruiz-VBzquez et al., 1993; Botella et al., 1995; Cervantes, 2000; Iniesta, 2003). Genes at the carBA region are organized as crtE-crtla-crtB-crtD-crtC-orf6-crtYccrtYd-orf9-carA-orfl l, where the ones designated as orf are genes whose functions remain to be fully assigned, and the two most downstream are regulatory genes (Fig. 1). Figure 2 shows the proposed pathway for carotenoid biosynthesis in M . xanthus and the genes involved in each step. The pathway represents the basic steps for the synthesis of myxobacton, the primary carotenoid end product in lightgrown M . xanthus (large amounts of phytoene, the first C,, precursor in the biosynthetic pathway, are also found in light-grown M. xanthus, as well as lower amounts of other end products not shown in Fig. 2). Myxobacton is a monocyclic carotenoid with a keto group on the ring at one end of the molecule and a glycosyl group usually esterified to a straight-chain fatty acid at the other (for a detailed review on the structures and biosynthesis of myxobacterial carotenoids see Reichenbach and Kleinig, 1984). CrtE, the product of the first gene of the carB operon, catalyzes the conversion of farnesyl pyrophosphate into geranylgeranyl pyrophosphate (GGPP).Condensation of
two GGPP molecules to yield the colorless C,, phytoene is carried out by CrtB. Four successive dehydrogenation steps introduce conjugated double bounds into the chain to produce lycopene, a colored intermediate. Unlike other bacteria, M. xanthus requires two dehydrogenases (CrtIa and CrtIb) to transform phytoene into lycopene, as has been shown by heterologous expression of CrtIa and/or crtlb in E. coli strains that accumulate phytoene (Iniesta, 2003). Among bacteria, only cyanobacteria are known to use two different enzymes for this conversion, but their action is sequential and not synergetic as in M . xanthus. The cyclization at one end of lycopene to produce y-carotene is catalyzed by the combined action of CrtYc and CrtYd, so denoted from their amino acid sequence homology to heterodimeric lycopene cyclases from gram-positive bacteria, and to the N-terminal domain of bifunctional fungal lycopene cyclases, whose C-terminal regions also show phytoene synthase activity (Krubasik and Sandmann, 2000). Heterologous gene expression of crtYc or crtYd, or the two together, in E. coli strains overproducing lycopene showed that their products act in concert to catalyze cyclization of lycopene but at only one end of the molecule, unlike their
REGULATORY MECHANISMS
214
*cH20pP FP P m
C
4 H
CrtE 2
O
P
P
GGPP
CrtB
1
dehydrogenases, catalyzes an additional desaturation reaction. Gene orf6, located at the 3‘ end of the carB operon, codes for a protein similar to bacterial glycosyltransferases. So, Orf6 may catalyze the addition of the sugar moiety to the chain. At present, it is not known if the product of orf9 plays a structural or a regulatory role. While some data point to Orf9 participating in the regulation of the light-inducible promoters (Cervantes and Murillo, 2002), its sequence similarity to bacterial acyltransferases makes it tempting to assign a role for Orf9 in myxobacton esterification.
Crtla + Crtlb Molecular Aspects of Signal Reception and Transduction in Carotenogenesis The Signals-Blue Light and Copper
Lycopene
1
CrtYc + CrtYd
y-carotene
crtc
1 1-OH-1’2’dihydroy-carotene
1 I-OH-3’4’dehydro1‘2’d ihyd roy-carote ne
Orf6? Orf9?
Fatty acid
Figure 2 Carotenoid biosynthetic pathway and the genes involved in each step. The proposed pathway for the biosynthesis of myxobacton, the primary carotenoid in M. xanthus, is shown. The roles of Orf6 and Orf9 remain to be defined but may be linked to the glycosyl- and acyltransferase activities, respectively, that are essential for production of myxobacton. counterparts mentioned above that do so at both ends (Iniesta, 2003). CrtC catalyzes the hydroxylation of y-carotene, while CrtD, a protein similar to carotene
Burchard and Dworkin (1966) demonstrated that accumulation of carotenoids in M. xanthus was dependent on the extent of illumination and the growth phase. Carotenoid levels, whose maximum build-up occurred for stationaryphase cells irradiated with blue light (405to 410 nm), were found to correlate directly with increasedprotection against photolysis (Burchard and Hendricks, 1969). The observation that the action spectra for photolysis and carotenogenesis in M. xanthus were similar and corresponded closely to the absorption spectrum of the iron-containing protoporphyrin IX, led to the proposal that this compound was the photosensitizer that linked the processes of photolysis and carotenogenesis (Burchard et al., 1966; Burchard and Hendricks, 1969). This highly hydrophobic compound accumulates in cell membranes, with an almost 16-fold increase in its levels as cell cultures approach stationary phase. Protoporphyrin IX absorbs strongly at 410 nm to form the relatively long-lived yet highly reactive “excited triplet state” species, which can directly cause cell damage and photolysis by modifying cellular components such as lipids, proteins, and nucleic acids. The excited triplet-state protoporphyrin IX can also react with oxygen to generate another highly reactive and diffusible singlet oxygen (‘02), species, which is significantly long-lived in hydrophobic environments such as membranes. Both of these reactive species are quenched by carotenoids. The major site for accumulation of the hydrophobic carotenoids, as with protoporphyrin IX and lo2,is the cytoplasmic membrane (Kleinig, 1972), which is thus the cellular location where the initial signal reception and transmission, as well as the final quenching by carotenoids, take place. Blue light is the principal environmental agent required for the induction of carotenogenesis in M. xanthus. It has been reported that the need for light can be partially bypassed by hyperoxygenation in the dark, suggesting that an oxygen-related species could be the actual
12. CAROTENOGENESIS IN 211. XANTHUS switch that triggers the cascade culminating in carotenogenesis (Robson, 1992; Hodgson and Murillo, 1993). Indications that this effector was '0, came from the observation that expression of the carQRS operon could be induced by agents capable of generating lo2,such as the dye toluidine blue in the presence of both oxygen and red light (Robson, 1992). On their own, neither the dye nor red light was capable of inducing carQRS expression. Moreover, quenchers of '0,other than carotenoids also greatly diminished the effect of light. It has very recently been found that carotenoid synthesis in wild-type M . xanthus in the absence of light can be induced by copper (Moraleda-Muiioz et al., 2005). Like many essential transition elements, copper must be maintained at specific levels in cells but is toxic at elevated concentrations. At metal concentrations below the lethal threshold, copper-induced carotenogenesis is observed under conditions suboptimal for growth like lower temperatures and diminished nutrient levels. Every regulatory element implicated in the light-triggered induction of carotenoid synthesis is found to also respond to copper, with the one exception discussed below. While the chemistry of the copper response and whether its action is direct or indirect remain to be worked out, it is noteworthy that can be generated from the membranelocated phosphatidylcholine hydroperoxide if copper is present (Takayama et al., 2001). Thus, lo2could provide the common link between the light- and the copperinduced pathways to carotenogenesis in M . xanthus.
Role of CarF and CarR in Signal Reception or Transduction Available data suggest that induction by light and copper converge to a common route very early in carotenoid synthesis in M . xanthus, as all but one of the known regulatory elements involved respond to both light and copper, except for the factor CarF (Moraleda-Muiioz et al., 2005). CarF, whose expression is not affected by illumination, is the factor that comes into play earliest in the response to light (Fontes et al., 2003). Its precise mechanism of action remains to be elucidated, but two alternatives have been considered. Either CarF is the primary photoreceptor working directly or through an associated chromophore, or it is a transmitter of the light signal from the primary to-be-identified photoreceptor to the next junction in the cascade. Sequence analysis failed to reveal any similarity of CarF to known blue light receptors, prokaryotic or eukaryotic. Furthermore, the only proteins showing any resemblance to the CarF sequence are members of a family of proteins of undefined functions named Kua (see Fontes et al., 2003). Whatever may be the exact function of CarF, its role is in the response to light and not in
215
that to copper. If the responses to both light and copper are, as suggested, dependent on the same inducer (singlet oxygen), then CarF would be specifically required in the photosensitizing mechanism to generate that inducer. Epistatic analysis established that CarF is followed in the light-induced cascade by CarR. Mutations in carR were among the earliest to be mapped in this light-induced process (Martinez-Laborda et al., 1986; Balsalobre et al., 1987). These cause constitutive synthesis of carotenoids, indicating that CarR is a negative regulator of light-induced carotenogenesis. carR is the second of three translationally coupled genes of the light-inducible carQRS operon, which plays a central role in the induction of carotenogenesis (McGowan et al., 1993).Its expression increases almost 80-fold when exposed to light (Hodgson, 1993). CarR is an inner membrane protein that is unstable when exposed to light, particularly when cells enter the stationary phase (Gorham et al., 1996; Browning et al., 2003). It is worth noting that maximal levels of carotenoids are also produced in the early stationary phase. These, however, do not appear to afford any protection to the lightmediated elimination of CarR (Browning et al., 2003). These observations have led to the proposal that light induces carotenogenesis by bringing about degradation of CarR, to thereby relieve the CarR-mediated repression of the process. CarF would then be expected to be involved in channeling this light-driven inactivation of CarR. The functional link between CarR and CarF appears to be via direct physical interactions between them, suggesting that CarF is an anti-anti-sigma factor (Fontes et al., 2003; Galbis-Martinez, 2005; also see below). The molecular basis for light- or copper-induced CarR destruction is, however, unknown. Whether these agents chemically modify CarR to lower its stability or whether it is modifications of the proteins associated with CarR that cause the release and the consequent destruction of an unstable CarR remains an intriguing question. Besides CarF, at least one other key factor is known to be physically associated with CarR, as will be discussed next.
Specific and Global Transcriptional Factors in the Regulation of the curQRS Operon
CarQ Cells bearing lack-of-function mutations in carQ or cars, the first and third genes of the carQRS operon, are Car- (McGowan et al., 1993; Gorham et al., 1996).Thus, CarQ and Cars, in contrast to CarR, are positive regulators of carotenogenesis. Mutations at carQ are epistatic over those at carR and block the activation of the crtIb structural gene and of the carQRS operon itself (Fontes et al., 1993; McGowan et al., 1993; Gorham et al., 1996).
REGULATORY MECHANISMS
216 The CarQ sequence suggested it to be a member of the extracytoplasmic function (ECF) subfamily of bacterial ~'O-likefactors (Lonetto et al., 1994). ECF cr factors act in cellular responses to different extracytoplasmic stimuli, and their activity is typically modulated by their association with membrane-bound anti-a factors (Hughes and Mathee, 1998). In vitro transcription runoff assays using E. coli core RNA polymerase indicated that CarQ could specifically initiate transcription at PQRS,the curQRS promoter (but not at PI, the crtIb promoter-see below), and that it physically interacted with CarR (Browning et al., 2003). CarR would then be an anti-o factor. As with many other ECF a-anti-cr factor pairs the genes for CarQ and CarR are translationally coupled. The membraneassociated CarR sequesters CarQ in a transcriptionally inactive complex. Inactivation of CarR, possibly via degradation, in response to specific environmental signals (light and copper) and the consequent release of CarQ appear to be the mechanism by which carotenogenesis is induced. The -10 region of PQRs (Fig. 3 ) , like other ECF crfactor-dependent promoters in M. xanthus, shares little
P,S
or no sequence identity with the consensus sequences for known cr factors (Whitworth et al., 2004). On the other hand, the -35 region of PQRsshows a greater resemblance to the consensus for ECF cr-factor-dependent promoters (Lonetto et al., 1994). Using a series of nested deletions, the minimal carQRS promoter was mapped to a 145-bp stretch extending upstream from the transcriptional start point, which also includes the promoter for the divergent gufA (for gene of unknown function). Mutational analysis of this minimal promoter identified several positions important for promoter activity and suggested transcriptional coupling between the gufA and carQRS promoters (Whitworth et al., 2004). It was also proposed that DNA between positions +40 and +66 may influence mRNA stability or have enhancer activity, just as reported earlier with P, (Martinez-Argudo et al., 1998; see below). As mentioned previously, a minimal heterologous in vitro transcription system with CarQ and E. coli core RNA polymerase alone could be made to initiate transcription at Pqns (Browning et al., 2003). However, the rather large size of the promoter region for
-35 I
-10 I
TCACCGAACCTTGAG~GCGCGAGCGCCG~~CTTTCGCAGGTGGCCCGTAGAGGAGTCG AGTGGCTTGGAACTCTTCGCGCTCGCGGCCTTTGTG~GCGTC~CCGGGCATCTCCTCAGC
CarD-binding
PI3
-35
-10
I
I
-35
-10 I
TGGACGCAAACGCTACCTCTAGGAAA ACCTGCGTTTGCGATGGAGATCCTTT
CarA-binding
Figure 3 Design of the light-inducible promoters identified in M. xanthus. Sequences show the promoter regions of PQRS,P,, and P,, the three M. xanthus light-inducible promoters. The -10 and -35 bases are marked for all three promoters. In PQRS,the CarD-binding site is boxed (with the two AT-rich tracts underlined). Two bases at the -35 region of PQRS,which on mutation remove light-induced promoter activity, are underlined (Whitworth et al., 2004). The bases underlined in the P, promoter sequence are critical for promoter activity (MartinezArgudo et al., 1998). In the P, promoter region, the inverted repeats of the bipartite CarA operator, PI and pII, are boxed and marked by arrows.
12. CAROTENOGENESIS IN 211. XANTHUS
217
carQRS suggests that accessory transcriptional regulators come into play in activating this promoter and, as described next, several factors have indeed been found.
The CarD-CarG complex The carD gene was identified in a screen for Carmutants among a large collection of strains bearing Tn.5 insertions (Nicolb et al., 1994). carD is actively and continuously transcribed during vegetative growth in a light-independent manner. It was established that the critical role of CarD in carotenogenesis was in the lightinduced activation of the carQRS operon and of crtIb (see below). The sequence of carD revealed it to code for a very novel transcriptional factor in prokaryotes, with attributes more akin to eukaryotic transcriptional factors (Nicolhs et al., 1996).Thus, the 136-residue C-terminal segment of CarD contains four repeats of the so-called "AT-hook" motif (a conserved RGRP sequence embedded in a cluster of basic residues and proline) linked to a highly acidic region, an arrangement very similar to that in eukaryotic high-mobility group A (HMGA) proteins (Fig. 4). Eukaryotic HMGA, of which the most extensively studied are the mammalian proteins, are small (I, 107 residues), relatively abundant nonhistone components of chromatin that function as DNA architectural factors to remodel chromatin and prime it for the assembly of specific nucleoprotein complexes essential in transcription, replication, recombination, and repair (Reeves, 2001,2003). The HMGA-like module in CarD is autonomously stable and monomeric, with an intrinsically random structure that exhibits specific minor-groove DNAbinding to appropriately spaced AT-rich tracts characteristic of eukaryotic HMGA (Padmanabhan et al., 2001).
1
180 183
Pfam 02559 4
228229
316
HMGA-like - b
Interaction with CarG
Interaction with DNA
Figure 4 Domain organization of CarD. The independent domain formed by the first 180 N-terminal residues of CarD, which defines the family Pfam 02559 of CarD-like proteins, is schematically represented by an unfilled rectangle. The Cterminal HMGA-like domain is made up of two stretches: a highly acidic one represented by the filled rectangle (residues 183 to 228) and a basic one which contains the four AT-hook repeats represented by the four stippled boxes (residues 229 to 316). The specific CarD domains that interact with CarG and DNA are as indicated.
Moreover, like the latter, CarD plays multifunctional roles in vivo. Besides being required for light-induced carotenogenesis, CarD is necessary for multicellular development and for the regulation of various vegetatively expressed genes (Nicolhs et al., 1994; Galbis-Martinez et al., 2004; see below). CarD has an N-terminal stretch of around 180 amino acids that is absent in eukaryotic HMGA (Fig. 4). This segment of CarD defines a family of proteins (Pfam 02559) each of whose members exist as a stand-alone module in a diverse array of bacterial species, including 211. xanthus. This domain of well-defined structure is essential for CarD function in vivo, as its absence results in a phenotype identical to that caused by a complete deletion of carD. Acquisition of the eukaryotic HMGA segment via lateral gene transfer, followed by fusion to a module existing only in bacteria, has been invoked to account for the unique domain architecture of CarD (Cayuela et al., 2003). Activation of the light-inducible PQRs promoter depends on a DNA segment which includes two appropriately spaced AT-rich tracts at -63 to -77 bp relative to the transcription start site (Fig. 3; Nicolhs et al., 1996). In vitro gel shift assays show that CarD binds to DNA probes containing these tracts via its HMGAlike C-terminal domain with the expected minor-groove binding specificity (Padmanabhan et al., 2001; Cayuela et al., 2003), and DNase I footprinting maps CarDbinding to the above two AT-rich tracts (Peiialver-Mellado et al., 2006). Intriguingly, it has been reported that only mutation of the more upstream AT-rich tract completely inactivates PQRS,and it was thus suggested that the two AT-rich tracts have different roles in vivo (Whitworth et al., 2004). Recent studies have identified a new factor, CarG (the product of the gene directly downstream of carD), which is required in every CarD-dependent process analyzed (Pefialver-Mellado et al., 2006). Although CarG shows no significant overall similarity to any protein in the database, it does contain a conserved H/C-rich segment, HEx,Hx,Gx,HCX,CXMX~~CX~C (where x is any amino acid) that is found in predicted zinc-dependent metalloproteases found in Archaea (hence named archaemetzincins), the hyperthermophilic bacterium Aquifex aeolicus, and two recently reported cases in humans (Diaz-Perales et al., 2005). The motif in CarG has the conserved G replaced by E, and even more significantly a Q replaces the invariant E, a change known to eliminate protease activity but not zinc binding. Purified CarG has been shown to lack protease activity and to contain two equivalents of zinc; also, a crucial structural role for the conserved Cs has been demonstrated by sitedirected mutagenesis. CarG thus appears to be a novel
218 transcriptional factor in which a metalloprotease motif has evolved to fulfill a purely structural role. Although no DNA-binding activity has been detected for the zincbound CarG, it associates with DNA through physical interaction with the N-terminal domain of CarD (Peiialver-Mellado et al., 2006). That CarD and CarG form a complex rationalizes not only how they simultaneously regulate different processes but also why they always coexist in the bacteria in which they occur. Analysis of microbial genomes for the presence of proteins similar to CarD and CarG indicates that these exist only in myxobacteria and that when one is present in a myxobacterium, so is the other: carD and carG exist in Stigmatella aurantiaca and Anaeromyxobacter dehalogenans, but both are absent in Sorangium cellulosum (Cayuela et al., 2003; Peiialver et al., 2006). What role CarG provides to the complex in transcriptional regulation is currently under study. One working hypothesis is that CarG serves as an adaptor to bridge the DNA-bound CarD with the basal transcriptional machinery. A direct interaction of CarG, CarD, or the CarD-CarG complex with CarQ, or M . xanthus RNA polymerase holoenzyme, has so far not been observed. It is therefore likely that other factors also participate in transcriptional regulation orchestrated by CarD and CarG, and these remain to be identified.
IhfA One other factor that has been experimentally shown to play a role in the initiation of transcription at PQRs is the nucleoid-associated integration host factor (IHF) (Moreno et al., 2001). It was identified in a screen for Car- mutants as a mutation unlinked to any of the known genes involved in carotenogenesis. The mutation mapped to ihfA, which encodes the or-subunit of the IHF heterodimer, a histone-like global transcriptional factor. Mutations in ihfA are epistatic over those in carR, and genetic data suggest that its gene product participates directly in activating PQRS.The IHF heterodimer and the related HU (heat unstable nucleoid protein) homodimer are DNAbinding architectural factors whose ability to bend DNA facilitates the assembly of higher-order nucleoprotein complexes essential in diverse processes including recombination, transcription, and replication (Nash, 1996).In contrast to nonspecific DNA-binding for HU, IHF binds preferentially to the consensus WATCAANNNNTTR (Friedman, 1988). Although no exact match to this consensus has been found at the PQRs promoter region, we cannot discard the possibility that IHF does interact with this region, as the strong “HU-like” character of M. xanthus IHF could confer it with the ability to bind DNA less specifically than, or differently from, E. co2iIHF (Moreno
REGULATORY MECHANISMS et al., 2001). Thus, IHF may activate PQRsby directly participating in the assembly of an essential transcriptionally competent complex together with CarD and CarG or by controlling the expression of an as yet unknown gene.
Transcriptional Regulation of the Structural Gene crtIb As mentioned earlier, the monocistronic crtlb codes for one of the two dehydrogenases involved in the conversion of phytoene to lycopene. Activity at the crtlb promoter, PI, is stimulated by light, but maximal induction (-400-fold) occurs only when cells reach stationary phase or are starved of a carbon source. Genetic evidence indicated that expression of PI requires CarQ (Fontes et al., 1993). As with PQRs and other ECF o-factordependent promoters in M. xanthus, the -35 region of PI but not the -10 region resembles the consensus (Fig. 3; Martinez-Argudo et al., 1998; Whitworth et al., 2004). Mutational analysis of P, revealed that promoter activity is critically dependent on a 5-bp stretch around position -31, of which four are conserved in PQRS,and three contiguous base pairs at - 10, which are also present in PQRs (Fig. 3; Martinez-Argudo et al., 1998).However, in contrast to PQns, no transcription was observed at PI with a minimal heterologous in vitro transcription system employing CarQ and E. coli RNA polymerase (Browning et al., 2003), suggesting the involvement of additional factors. As with PQRS,the minimal crtIb promoter region is rather large and spans positions -59 and +120 relative to the transcription start, with a putative enhancer-like element between +30 and +120 (Martinez-Argudo et al., 1998). This could also reflect the need for accessory factors in regulating transcription from P,. Although CarD is also absolutely required for the activation of crtlb (Nicolis et al., 1994), the PI promoter region does not reveal a sequence that could be a CarD-binding site, nor is there any evidence yet for CarD binding to this region (Martinez-Argudo et al., 1998). Thus, while CarD is critical for the activation of PI, its regulatory role may be indirect. Mutations at ihfA also impair expression of crtlb, but this is overcome if carQ is expressed from a heterologous, ihfA-independent promoter (Moreno et al., 2001). Thus, the role of IHF in activating PI appears to be indirect and linked to PQRs activation and the consequent production of CarQ.
Transcriptional Regulation of the Structural Genes in the curl3 Operon Other than crtlb, all of the structural genes for carotenoid synthesis in M. xanthus are grouped at the carBcarA cluster. The carB locus was identified by Tn5 insertion mutations which generated a Car- phenotype
12. CAROTENOGENESIS IN M. XANTHUS (Martinez-Laborda et al., 1986; Balsalobre et al., 1987; Ruiz-Vizquez et al., 1993). Subsequent sequencing revealed the presence of six structural genes at this locus, and the enzymes they encode were deduced by analyzing predicted amino acid sequences (Botella et al., 1995) and characterizing the carotenoids accumulated on heterologous expression in E. coli (Cervantes, 2000; Iniesta, 2003). Gene expression analyses indicate that most or all of the genes in the carB cluster are transcribed from its light-inducible promoter, P,. The carA locus was originally defined by the point mutation carA 2 that resulted in constitutive, lightindependent expression of carotenoids, just as with mutations in the unlinked carR locus (Balsalobre et al., 1987; Martinez-Laborda and Murillo, 1989). However, the carAl mutant cells were orange and so less intensely pigmented than the deep red of cells with carR mutations, because the latter also activate crtlb (Martinez-Laborda and Murillo, 1989: Fontes et al., 1993).There is evidence for a light-independent promoter, PA,located within orf6, which would drive expression of the five genes that are further downstream and encompass the carA operon. Nevertheless, expression of the genes in the carA operon is also increased in the light presumably due to transcriptional readthrough from P, (Ruiz-Vizquez et al., 1993; Botella, 1996). The mechanism of light-induced expression at P, differs from those at PQRs and PI. Light enhances P, activity about 20-fold (monitored by the expression of reporter lacZ fusions [Balsalobre et al., 1987]), compared to 400-fold for PI (Fontes et al., 1993) or 80-fold for PqRs (Hodgson, 1993). Whereas maximal light activation of P, required cells to reach stationary phase, P, could be photoinduced at any time during the growth cycle. In contrast to PQRs and PI, the -35 region of P, perfectly matches the TTGACA of E. coli promoters dependent on the major vegetative u70-RNA polymerase holoenzyme but, as is often the case in GC-rich M. xanthus, the - 10 hexamer in P, (TACCTC) diverges considerably from the AT-rich consensus (TATAAT)in E. coli (Fig. 3) (Botella et al., 1995).Purified M . xanthus containing the major vegetative uA(MxRNAP) does indeed bind specifically to P, to form stable, open complexes capable of transcription in vitro (L6pez-Rubio et al., 2004). P,, but not PQRs or PI, is also affected by mutations a t two unlinked loci. One of these is c a d , the third gene of the carQRS operon, and the other is carA, the fourth gene of the carA operon. Absence of cars blocks activation of P, by light, while expressing cars from a lightindependent promoter strongly activates P, even in the dark (McGowan et al., 1993), indicating that Cars is a positive regulator of P,. The first known CUTSmutation
219
identified (carS1)was nonetheless a gain-of-function one that led to a light-independent, constitutive expression at P, (Balsalobre, 1989). It corresponded to a stop codon that produced CarS1, a truncated version of Cars lacking the last 25 amino acids (McGowan et al., 1993).As discussed later, the molecular basis of Cars action can rationalize this gain-of-function carS2 phenotype. Constitutive expression at P, was observed for the carAl mutation that originally defined the carA locus, which suggested that this locus houses elements for the negative regulation of P, in the dark (MartinezLaborda and Murillo, 1989). DNA sequence analysis of the carA2 clone revealed two closely linked nucleotide changes: one was a single nucleotide deletion at the 3’ end of orf9, which changes the last six amino acids of Orf9 and adds 76 residues at its C terminus; the second mutation was an A-to-T transversion at the fifth codon of carA that causes an I-to-F substitution (Botella et al., 1995). Nonpolar deletions in each of orf9, carA, and the downstream o r f l l showed that orf9 may play a role in positively regulating PQRs and thereby crtlb, while the negative regulator of P, is encoded by carA (Cervantes and Murillo, 2002). It was also shown that CarA, which does not participate in the light activation of PI or of PQRS, is a transcriptional repressor that binds specifically to the DNA region encompassing P,, while Cars interacts specifically with CarA and not DNA to derepress P, (Whitworth and Hodgson, 2001; Lopez-Rubio et al., 2002). CarA and Cars thus form a repressorantirepressor pair in regulating P,.
CarA Operator Design and Model for Regulation of P, The CarA operator design and the mechanism underlying the repression-antirepression switch of P, have been subjected to detailed molecular analysis (Fig. 5 ) (LopezRubio et al., 2004). The operator has a bipartite design and is made up of a high-affinity CarA-binding site (PI), which is a perfect interrupted palindrome located between positions -46 and -63 relative to the transcription start site, and a low-affinity one (pII)corresponding to an imperfect interrupted palindrome spanning positions -25 to -40 (Fig. 3). This is manifested in vitro by the stepwise binding of dimeric CarA, first to PI and then cooperatively to pII. Binding of CarA to pII, which overlaps partially with the -35 promoter region, effectively occludes a*-RNA polymerase from P, and thereby downregulates expression of the carB cluster. On the other hand, CarA bound to pII is efficiently dislodged by Cars, when present, to activate P,. The bipartite operator design thereby enables a rapid and effective response to light and provides the operative mechanism for the
REGULATORY MECHANISMS
220
f 2arS I )
LIGHT
CarA dimer
-35
e c=acsQ
PI
-10
car5operon
PI1
Figure 5 Model for the action of CarA and Cars in the regulation of the P, promoter. The dimeric CarA repressor is shown with its N- and C-terminal domains represented by the small and large spheres, respectively. In the dark (left panel), two CarA dimers bind via their Nterminal domains and in a cooperative fashion to the bipartite CarA operator. The two sites in the operator, palindromes PI and pII, are each shown as a pair of convergent arrows. Occupancy of pII by CarA blocks promoter access to the RNA polymerase holoenzyme (shown by the object labeled RNAP) leading to repression of curB. On exposure to light (right panel) CarS, shown by the dark ellipsoids, is produced, and its interaction with the N-terminal domain of CarA readily dismantles CarA-pII complexes. The RNA polymerase holoenzyme thereby gains access to the promoter, leading to the derepression of carB. repression of the carB operon by CarA and the derepression of P, by CarS. T h e CarA-Cars Repressor-Antirepressor Pair While Cars has no known sequence homologs, CarA has an N terminus which shows sequence similarity to the Nterminal, DNA-binding domains of bacterial MerR transcriptional factors (Botella et al., 1995). MerR factors act in different stress responses and, depending on the union of specific cofactors like heavy metals or drugs to their Cterminal domains, they repress or activate transcription by inducing DNA conformational changes (Brown et al., 2003). Their DNA-binding sites, in contrast to the CarA operator, fall entirely within a suboptimal 19-bp (not the usual 17-bp) spacer region that separates the -35 and -10 regions. As with MerR proteins, the CarA C-terminal domain was predicted to bind a specific cofactor by sequence analysis. However, the putative ligand, vitamin B,, or a related cobalamin derivative, is a novel one for a transcriptional cofactor (Cervantes and Murillo, 2002). Recent studies confirm the predicted structuralfunctional domain organization for CarA. Its 78-residue N-terminal segment is indeed a stable, autonomously folded unit that contains the binding determinants not only for the operator DNA but also for the antirepressor Cars (PCrez-Marin et al., 2004). That the domain adopts the winged-helix topology of MerR family DNAbinding domains has been confirmed by nuclear magnetic resonance. These studies, together with site-directed
mutagenesis, show that common structural elements in Car.A mediate specific interactions with DNA as well as with Cars (Navarro-AvilCset al., 2007). The findings also suggest that the CarA-binding site on Cars may mimic operator DNA in size, shape, and electrostatic complementarity, so that this could form the physical-structural basis for antirepression by CarS. Like reported DNA mimics, Cars is a highly acidic protein. Significantly, the truncated form of CarS, CarS1, is even more acidic and its greater affinity for CarA would explain the gainof-function carSl phenotype mentioned earlier (L6pezRubio et al., 2002). That the CarA C-terminal domain is an independently stable domain that binds vitamin B,, and mediates CarA dimerization has been shown experimentally (PCrezMarin et al., 2004; M. C. P6rez-Marin, S. Padmanabhan, M. C. Polanco, F. J. Murillo, and M. Elias-Arnanz, unpublished data). Moreover, B,, appears to be involved in the regulation of P, (Cervantes and Murillo, 2002). However, the findings that the N-terminal domain of CarA is capable of repressing P, in vivo when expressed at sufficiently high levels (Ptrez-Marin et al., 2004) and that mutating key H residues of the B,,-binding motif does not affect CarA function in vivo (Pkrez-Marin et al., unpublished), suggest that it may not be through CarA that B,, acts in regulating P,. In fact, detailed genetic analyses have indicated that it is O r f l l , the product of the gene directly downstream of CMA,that is involved in repressing P, in a B12-dependentmanner. O r f l l , which
12. CAROTENOGENESIS IN 211. XANTHUS
22 1
shares significant sequence similarity and domain organization with CarA (Botella et al., 1995; Cervantes and Murillo, 2002), would thus be yet another transcriptional factor in the gene regulation cascade for M. xanthus carotenogenesis. Its binding to the CarA operator and to Cars via the N-terminal domain has been experimentally confirmed, and the molecular details of how its activity is modulated by B,, are being worked out (Pkrez-Marin et al., unpublished data). The reasons for B,, recruitment in this light response is still an open question. A possible link to photoreception is suggested by the fact that at least one protein-bound form of BI2, cob(I)alamin, exhibits strong blue-light absorbance with a maximum near 400 nm (Jarrett et al., 1997), the same blue-light region where maximal photoinduction of carotenogenesis occurs. Whether this or some other mode of action underlies the role of B,, in M. xunthus carotenogenesis, however, remains to be experimentally examined.
THE CarD-CarG COMPLEX LINKS MULTICELLULAR DEVELOPMENT AND CAROTENOGENESIS The most studied phenomenon in M . xanthus is undoubtedly its striking ability to form multicellular fruiting bodies on starvation, a process that has served as a prokaryotic model for the study of cell-cell interactions and cellular differentiation (Kaiser, 2003, 2004). A genetic link between fruiting body development and the blue-light response in M. xanthus was first provided by the identification of gene carD. The carD2 mutation, a Tn5 insertion at curD originally isolated because it impaired light-induced carotenogenesis, was found to affect fruiting body formation as well. The mutant exhibited only weak signs of aggregation, as if blocked at an early stage of development (Nicolis et al., 1994).The same defective phenotype was observed for a strain bearing a complete in-frame deletion of curD
WT
(Fig. 6) (Cayuela et al., 2003). This developmental phenotype of carD mutants was not due to lack of carotenogenesis, since normal differentiation and development was observed for M . xanthus bearing other Car- mutations such as those in curB, crtlb, or carQ (Nicol6s et al., 1994). CarD was shown to be essential for activating the expression of genes involved in both the early and late stages of development mediated by the A-factor and C-factor intercellular signals, respectively (Nicolis et al., 1994). As mentioned earlier, CarG is a recently identified factor that works with CarD in the regulation of multicellular development and carotenogenesis in M. xanthus. Strains with in-frame deletions at carG are Car- and are blocked at an early stage of fruiting body formation, just as with carD mutant strains (Fig. 6). Like CarD, CarG appears to be required for expression of a set of A- and C-signaldependent developmental markers (Pefialver et al., 2006). Thus, CarD and CarG might be required in the production of the C- and A- signals or in the pathways that these signals mediate. Light and lack of nutrients, respectively, trigger carotenogenesis and multicellular development in M. xanthus, and both entail the transcriptional activation of specific genes. But of the carotenoid genes whose expression is stimulated by light, only that for crtlb is also dependent on starvation, the cue for activating the developmental process. Thus, two distinct environmental signals converge in their requirement for the CarD-CarG pair but diverge in the set of genes that are ultimately triggered.
LIGHT-REGULATED CAROTENOGENESIS IN OTHER BACTERIA Carotenoids are present in all photosynthetic and some nonphotosynthetic bacteria. Most prokaryotes known to produce carotenoids do so in a constitutive manner. However, besides M. xunthus, some other species
carDA
carGA
Figure 6 Developmental phenotype of the carD and carG mutants. Photographs were taken after 5-day incubation of 10-pl droplets of cells (1.25 X lo8 cells/ml) spotted on CF agar. The wild-type control strain is DK1622.
222 of nonphotosynthetic bacteria such as Mycobacterium marinum, Flavobacterium dehydrogenans, Brevibacterium sulfureum, and Streptomyces, have been reported to produce carotenoids in a light-dependent manner (Bramley and Mackenzie, 1988; Armstrong, 1997). As with algae, fungi, and plants, light is the most influential environmental agent in bacterial carotenoid synthesis. Here, we focus on those bacteria for which the effect of blue light on carotenogenesis has been best characterized at a genetic and molecular level, highlighting the parallels with M . xanthus. In anoxygenic photosynthetic bacteria, light and oxygen have long been known to control accumulation of carotenoids (Armstrong, 1997). The seven structural carotenogenic genes (crtA through crtF and crtl) in Rhodobacter capsulatus and Rhodobacter sphaeroides are clustered with those for bacteriochlorophyll and light-harvesting proteins. Carotenoids accumulate preferentially under low-oxygen and low-light conditions. Regulation includes repression by PpsWCrtJ, a soluble tetrameric protein with a helix-turn-helix DNA-binding motif that recognizes a conserved 18-bp interrupted palindrome sequence. Repressed genes or gene clusters always contain two binding sites, one located within the promoter region and another either straddling the -35 promoter region or lying 100 to 150 bp upstream of the transcription start site (Bauer et al., 2002; Movskin et al., 2005). The two sites and the requirement for cooperative binding between them for effective repression by PpsR are reminiscent of the operator design and mode of action of M . xanthus CarA. Moreover, at least in R. sphaeroides, PpsR-DNA binding is inhibited by its antirepressor AppA, a flavin-containing blue-light photoreceptor, which both mediates breakage of a disulfide bond in PpsR and forms a stable AppA-PpsR, complex (Bauer et al., 2002; Masuda and Bauer, 2002). In this case, blue-light excitation of AppA deters its ability to complex with PpsR,, thereby permitting it to bind DNA and to downregulate carotenogenesis under high-light conditions. In the nonphotosynthetic Streptomyces, several species have been found to synthesize carotenoids on illumination, but this ability is lost at a relatively high frequency by many strains (see Kato et al., 1995).Genetic studies on carotenogenesis in Streptomyces had been reported only for S. griseus and S. setonii (Kato et al., 1995; Schumann et al., 1996; Lee et al., 2001). In these, however, carotenogenesis occurs in a cryptic manner: a crt gene cluster is found, but the condition under which carotenoid synthesis takes place is unknown. Involvement of the stress response sigma factor CrtS has been suggested, though it is unclear how its expression or activation is controlled
REGULATORY MECHANISMS (Kato et al., 1995; Lee et al., 2001). The recent construction of a strain that did not produce two pigmented antibiotics revealed light-induced carotenogenesis in Streptomyces coelicolor A(2) and prompted its genetic analysis (Takano et al., 2005). The structural genes for carotenogenesis in S. coelicolor A(2)exist as two convergent operons, crtEIBV and crtYTU, whose expression is induced on illumination. Their photodependent transcription has been linked to at least two genes, litS and litR, located in two divergent operons that flank crtYTU: litRQ and litSAB. Genetic analysis suggested an essential role for litS in the activation of the crt genes by light. A complete deletion of litS impaired carotenoid synthesis, while its overexpression off a light-independent promoter led to constitutive carotenogenesis. Moreover, the activities of the photoinducible PcrtE, PcrtY, and Pli, promoters were completely abolished in the litS mutant. LitS, a 22-kDa protein, has 21% sequence identity to 211. xanthus ECF sigma factor CarQ. By in vitro runoff transcription analysis, LitS was indeed shown to be a (T factor necessary for transcriptional activation of PcrtE and PcrtY. However, specific transcriptional initiation could not be reproduced in vitro with the P,, DNA probe, suggesting that factors other than LitS and core RNA polymerase may be required in this case. As exemplified by the CarQ-CarR pair, many ECF sigma factors use membrane-bound antisigma factors to recognize the extracytoplasmic inducing cues. Of the two other genes in the litSAB operon, LitB shows weak similarity to RsrR, an S. coelicolor antisigma factor. However, inactivating LitB (or the putative lipoprotein LitA) did not affect carotenoid synthesis. LitR was originally proposed to work as an essential positive regulator in the light induction of Plits, since all photoinducible promoters were inactivated in a litR mutant background (Takano et al., 2005). However, adding purified LitR to the in vitro runoff reactions with the LitS-RNA polymerase holoenzyme did not result in production of specific transcripts from Plits. Nor did LitR, a 35-kDa protein with a MerR-like DNA-binding domain, show in vitro binding to the litS promoter region (which does contain an interrupted palindrome). Interestingly, sequence analysis suggested a domain organization for LitR (Takano et al., 2006a) that resembles that for M . xanthus CarA (Cervantes and Murillo, 2002): a predicted DNA-binding motif linked to a putative BIZbinding C-terminal domain. In their first study (Takano et al., ZOOS), a positive regulatory role was attributed to LitR. More recently, photoinduction of PcrtYon heterologous expression of litS and litR in S. griseus and constitutive expression from this promoter on deleting litR from the plasmid led to the conclusion that LitR, like CarA,
12. CAROTENOGENESIS IN 111. XANTHUS acts as a repressor in the dark, rather than as an activator (Takano et al., 2006a). Moreover, the authors speculated that B,, may play a role in the light-dependent regulation by LitR, as was proposed earlier for M. xanthus CarA (Cervantes and Murillo, 2002). Nonetheless, LitR binding to DNA and/or B,, remains to be demonstrated. Although in a divergent orientation, rather than convergent as in s. coelicolor A(2),the carotenogenic crtEIBV and crtYTUoperons are also found in S. avermitilis,which appears to produce carotenoids in a light-dependent manner (Takano et al., 2005). Furthermore, sequence conservation of the litQRS gene cluster in S. avermitilis could be indicative of a regulatory mechanism that is similar to the one described for S. coelicolor A(2). The increasing number of newly available microbial genome sequences continues to provide new insights into the existence and arrangement of crt and possible regulatory genes in different bacteria. Thus, until recently CarA and O r f l l in M. xanthus were the only known transcriptional regulators made up of a MerR-type DNA-binding domain linked to a vitamin B,,-binding domain. Together with the recently reported S . coelicolor and S . avermitilis LitR, microbial genome data reveal new examples of proteins with a domain organization similar to that of C a r N Orfl 1 in several other nonphotosynthetic bacteria such as Bdellovibrio bacteriovorus, Dechloromonas aromatica, Exiguobacterium sp., Magnetococcus sp., Nocardia farcinica, Pseudomonas putida, Ralstonia eutropha, and Thermus thermophilus. Of these, only T. thermophilus has been reported to produce carotenoids (Hoshino et al., 1993),possibly in a light-dependent manner (Takano et al., 2006b). The genome sequence of T. thermophilus reveals some noteworthy features with regard to carotenogenesis (Henne et al., 2004). First, the crt genes involved in the early steps of the biosynthetic pathway are found in the chromosome, while those involved in the subsequent steps are plasmid encoded (though not organized in a single cluster, they are found in close proximity to one another). The T. thermophilus CarA analog (TtCarA) gene is also located in the megaplasmid and lies immediately upstream of the phytoene synthase gene but is transcribed in the opposite sense. This suggests a link between TtCarA and carotenogenesis in T. thermophilus, perhaps as in M. xanthus. Interestingly, genes involved in the early steps of cobalamin biosynthesis are located in the chromosome, whereas those participating in the later steps of the pathway occur in the megaplasmid. This genomic organization mirrors that for the crt genes, and its significance, if any, is an open question. It is nonetheless tempting to speculate that such an arrangement might reflect a relationship between cobalamin synthesis, carotenogenesis, and CarA function.
223
CONCLUDING REMARKS Studies with M . xanthus continue to provide insights into the general principles underlying the complexity of the biosynthetic pathways and regulatory mechanisms involved in eubacterial carotenogenesis. They reveal how an ingenious circuitry for receiving the inducing signal and transmitting it meshes with the gene regulatory machinery. This machinery involves, among others, transcriptional factors with novel domain architectures, and a design of cis-regulatory elements that together lead to an exquisite control of gene expression. A rapid and an effective response to the environmental cues that stimulate the process is thereby accomplished. W e thank Francisco Murillo for continual support, and him as well as all past and present members of our group for their many contributions. Our current understanding of lightinduced carotenogenesis in M. xanthus also owes considerably t o work from David Hodgson and his group at the University of Warwick (United Kingdom). W e benefited from sequence data provided to us by The Institute for Genomic Research (http://www.tigr.org) for M. xanthus and S. aurantiaca; Rolf Miiller (Universitat des Saarlandes) for S. cellulosum; and the U.S. Department of Energy Joint Genome Institute (http:// www.jgi.doe.gov) for A. dehalogenans. We are grateful to the Ministerio de Educacidn y Ciencia (Spain) for uninterrupted research funding.
References Armstrong, G. A. 1997. Genetics of eubacterial carotenoid synthesis: a colourful tale. Annu. Rev. Microbiol. 51:629-659. Balsalobre, J. M. 1989. Induccidn por la luz de la expresi6n ge'nica y la carotenoginesis en Myxococcus xanthus. Ph.D. thesis. University of Murcia, Murcia, Spain. Balsalobre, J. M., R. M. Ruiz-Vazquez, and F. J. Murillo. 1987. Light induction of gene expression in Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 84:2359-2362. Bauer, C., S. Essen, L. R. Swem, D. L. Swem, and S. Masuda. 2002. Redox and light regulation of gene expression in photosynthetic prokaryotes. Philos. Trans. R. SOC. Lond. B 358:147-154. Botella, J. A., R. M. Ruiz-Vazquez, and F. J. Murillo. 1995. A cluster of structural and regulatory genes for light-induced carotenogenesis in Myscococcus xanthus. Eur. J. Biochem. 2 2 3 ~ 28-248. 3 Bramley, P. M., and A. Mackenzie. 1988. Regulation of carotenoid biosynthesis. Curr. Top. Cell. Regul. 29:291-343. Brown, N. L., J. V. Stoyanov, S. P. Kidd, and J. L. Hobman. 2003. The MerR family of transcriptional regulators. FEMS Microbiol. Rev. 27:145-163. Browning, D. F., D. E. Whitworth, and D. A. Hodgson. 2003. Light-induced carotenogenesis in Myxococcus xanthus: functional characterization of the ECF sigma factor CarQ and antisigma factor CarR. Mol. Microbiol. 48:237-251. Burchard, R. P., and M. Dworkin. 1966. Light-induced lysis and carotenogenesis in Myxococcus xanthus. J. Bacteriol. 9 1535-545.
224 Burchard, R. P., S. A. Gordon, and M. Dworkin. 1966. Action spectrum for the photolysis of Myxococcus xanthus. J. Bacteriol. 9 1:896-8 97. Burchard, R. P., and S. P. Hendricks. 1969. Action spectrum for carotenogenesis in Myxococcus xanthus. J. Bacteriol. 971665-1668. Cayuela, M. L., M. Elias-Arnanz, M. Peiialver-Mellado, S. Padmanabhan, and F. J. Murillo. 2003. The Stigmatella aurantiaca homolog of Myxococcus xanthus high-mobilitygroup A-type transcription factor CarD: insights into the functional modules of CarD and their distribution in bacteria. J. Bacteriol. 185:3527-3537. Cervantes, M. 2000. Accidn de genes estructurales y reguladores en la respuesta a la luz azul de Myxococcus xanthus. Ph.D. thesis. University of Murcia, Murcia, Spain. Cervantes, M., and F. J. Murillo. 2002. Role for vitamin B,, in light induction of gene expression in the bacterium Myxococcus xanthus. J. Bacteriol. 184:2215-2224. Diaz-Perales, A., V. Quesada, J. R. Peinado, A. P. Ugalde, J. Alvarez, M. F. SuLrez, F. X. Gomis-Ruth, and C. Lopez0 t h . 2005. Identification and characterization of human archaemetzincin-1 and -2, two novel members of a family of metalloproteases widely distributed in Archaea. J. Biol. Chem. 280:3 0367-3 0375. Fontes, M., L. Galbis-Martinez, and F. J. Murillo. 2003. A novel regulatory gene for light-induced carotenoid synthesis in the bacterium Myxococcus xanthus. Mol. Microbiol. 47~561-571. Fontes, M., R. M. Ruiz-Vazquez, and F. J. Murillo. 1993. Growth-phase dependence of the activation of a bacterial gene for carotenoid synthesis by blue light. E M B O J. 12~1265-1275. Frank, H. A,, and G. W. Brudvig. 2004. Redox functions of carotenoids in photosynthesis. Biochemistry 43536078615. Friedman, D. I. 1988. Integration host factor: a protein for all reasons. Cell 5554.5-554. Galbis-Martinez, L. 2005. Analisis genitico y molecular de un transductor de la seiial luminosa en la bacteria Myxococcus xanthus. Ph.D. thesis. University of Murcia, Murcia, Spain. Galbis-Martinez, M., M. Fontes, and F. J. Murillo. 2004. The high-mobility group A-type protein of the bacterium Myxococcus xanthus as a transcription factor for several distinct vegetative genes. Genetics 167:15 85-1 595. Goodwin, T. W. 1980. The Biochemistry of Carotenoids, vol. 1. Plants. Chapman and Hall, Ltd., London, United Kingdom. Gorham, H. C., S. J. McGowan, P. R. H. Robson, and D. A. Hodgson. 1996. Light-induced carotenogenesis in Myxococcus xanthus: light-dependent membrane sequestration of ECF sigma factor CarQ by anti-sigma factor CarR. Mol. Microbiol. 48:237-251. Henne, A., et al. 2004. The genome sequence of the extreme thermophile Thermus thermophilus. Nut. Biotechnol. 22: 547-553. Hodgson, D. A. 1993. Light-induced carotenogenesis in Myxococcus xanthus: genetic analysis of the carR region. Mol. Microbiol. 7:471-488. Hodgson, D. A., and A. E. Berry. 1998. Light regulation of carotenoid synthesis in Myxococcus xanthus, p. 185-211.
REGULATORY MECHANISMS In M. X. Caddick, S. Baumberg, D. A. Hodgson, and M. K. Phillips-Jones (ed.),Microbial Responses to Light and Time, Society for General Microbiology Symposium, vol. 56. Cambridge University Press, Cambridge, United Kingdom. Hodgson, D. A., and F. J. Murillo. 1993. Genetics of regulation and pathway of synthesis of carotenoids, p. 157-181. In M. Dworkin and D. Kaiser (ed.), Myxobacteria II. American Society for Microbiology, Washington, DC. Hoshino, T., R. Fujii, and T. Nakahara. 1993. Molecular cloning and sequence analysis of the crtB gene of Thermus thermophilus HB27, an extreme thermophile producing carotenoid pigments. Appl. Enuiron. Microbiol. 58:93-98. Hughes, K. T., and K. Mathee. 1998. The anti-sigma factors. Annu. Rev. Microbiol. 52:231-286. Iniesta, A. A. 2003. Expresidn de genes carotenogdnicos de la bacteria Myxococcus xanthus en Escherichia coli. Ph.D. thesis. University of Murcia, Murcia, Spain. Jarrett, J. T., C. W. Goulding, K. Fluhr, S. Huang, and R. G. Matthews. 1997. Purification and assay of cobalamin-dependent methionine synthase from Escherichia coli. Methods Enzymol. 28 1:196-2 13. Kaiser, D. 2003. Coupling cell movement to multicellular development in myxobacteria. Nut. Rev. Microbiol. 1:45-54. Kaiser, D. 2004. Signaling in myxobacteria. Annu. Rev. Microbiol. 58:75-98. Kato, F., T. Hino, A. Nakaji, M. Tanaka, and Y. Koyama. 1995. Carotenoid synthesis in Streptomyces setonii ISP539.5 is induced by the gene crtS, whose product is similar to a sigma factor. Mol. Gen. Genet. 247:387-390. Kleinig, H. 1972. Membranes from Myxococcus fuluus (Myxobacterales) containing carotenoid glucosides. Isolation and composition. Biochim. Biophys. Acta 274:489-498. Krubasik P., and G. Sandmann. 2000. Molecular evolution of lycopene cyclases involved in the formation of carotenoids with ionone end groups. Biochem. Soc. Trans. 28906-810. Lee, H. S., Y. Ohnishi, and S. Horinouchi. 2001. A sigma B-like factor responsible for carotenoid biosynthesis in Streptomyces griseus. J. Mol. Microbiol. Biotechnol. 3:95-101. Lonetto, M. A., K. L. Brown, K. E. Rudd, and M. J. Buttner. 1994. Analysis of the Streptomyces coelicolor sigE gene reveals the existence of a subfamily of eubacterial RNA polymerase u factors involved in the regulation of extracytoplasmic functions. Proc. Natl. Acad. Sci. USA 91:75737577. Lopez-Rubio, J. J., M. Elias-Arnanz, S. Padmanabhan, and F. J. Murillo. 2002. A repressor-antirepressor pair links two loci controlling light-induced carotenogenesis in Myxococcus xanthus. J. Biol. Chem. 2777262-7270. Lopez-Rubio, J. J., S. Padmanabhan, J. M. Lazaro, M. Salas, F. J. Murillo, and M. Elias-Arnanz. 2004. Operator design and mechanism for CarA repressor-mediated down-regulation of the photoinducible carB operon in Myxococcus xanthus. J. Biol. Chem. 279:28945-28953. Martinez-Argudo, I., R. M. Ruiz-Vazquez, and F. J. Murillo. 1998. The structure of an ECF-u-dependent, light-inducible promoter from the bacterium Myxococcus xanthus. Mol. Microbiol. 302383-893. Martinez-Laborda, A., J. M. Balsalobre, M. Fontes, and F. J. Murillo. 1990. Accumulation of carotenoids in structural
12. CAROTENOGENESIS IN M.
XANTHUS
and regulatory mutants of the bacterium Myxococcus xanthus. Mol. Gen. Genet. 223:205-210. Martinez-Laborda, A., M. Elias, R. Ruiz-Vazquez, and F. J. Murillo. 1986. Insertion of Tn5 linked mutations affecting carotenoid synthesis in Myxococcus xanthus. Mol. Gen. Genet. 205:107-114. Martinez-Laborda, A., and F. J. Murillo. 1989. Genic and allelic interactions in the carotenogenic response of Myxococcus xanthus to blue light. Genetics 122:48 1-490. Masuda, S., and C. E. Bauer. 2002. AppA is a blue light photoreceptor that antirepresses photosynthesis gene expression in Rhodobacter sphaeroides. Cell 110:613-623. McGowan, S. J., H. C. Gorham, and D. A. Hodgson. 1993. Light-induced carotenogenesis in Myxococcus xanthus: DNA sequence analysis of the carR region. Mol. Microbiol. 10:713-735. Meiser, P., H. B. Bode, and R. Muller. 2006. The unique DKxanthene secondary metabolite family from the myxobacterium Myxococcus xanthus is required for developmental sporulation. Proc. Natl. Acad. Sci. USA 103:19128-19133. Moraleda-Muiioz, A., J. PCrez, M. Fontes, F. J. Murillo, and J. Muiioz-Dorado. 2005. Copper induction of carotenoid synthesis in the bacterium Myxococcus xanthus. Mol. Microbiol. 56:1159-1168. Moreno, A. J., M. Fontes, and E J. Murillo. 2001. ihfA gene of the bacterium Myxococcus xanthus and its role in activation of carotenoid genes by blue light. J. Bacteriol. 183557569. Moskvin, 0.V., L. Gomelsky, and M. Gomelsky. 2005. Transcriptome analysis of the Rhodobacter sphaeroides PpsR regulon: PpsR as a master regulator of photosystem development. J. Bacteriol. 187:2148-2156. Nash, H. A. 1996. The HU and IHF proteins: accessory factors for complex DNA assemblies, p. 149-179. In E. C. C. Lin and A. S. Lynch (ed.), Regulation of Gene Expression in Escherichia coli. R. G. Landes Company, Austin, TX. Navarro-AvilCs, G., M. A. JimCnez, M. C. PCrez-Marin, C. Gonzalez, M. Rico, F. J. Murillo, M. Elias-Amanz, and S. Padmanabhan. 2007. Structural basis for operator and antirepressor recognition by Myxococcus xanthus CarA repressor. Mol. Microbiol. 63:980-994. Nicolas, F. J., M. L. Cayuela, I. M. Martinez-Argudo, R. M. Ruiz-Vazquez, and F. J. Murillo. 1996. High mobility group I(Y)-like DNA-binding domains on a bacterial transcription factor. Proc. Natl. Acad. Sci. USA 93:6881-6885. Nicolas, F. J., R. M. Ruiz-Vazquez, and F. J. Murillo. 1994. A genetic link between light response and multicellular development in the bacterium Myxococcus xanthus. Genes Dev. 8~2375-2387. Padmanabhan, S., M. Elias-Arnanz, E. Carpio, P. Aparicio, and F. J. Murillo. 2001. Domain architecture of a high mobility group A-type bacterial transcriptional factor. J. Biol. Chem. 276:4156641575.
225 Peiialver-Mellado, M., F. Garcia-Heras, S. Padmanabhan, D. Garcia-Moreno, F. J. Murillo, and M. Elias-Arnanz. 2006. Recruitment of a novel zinc-bound transcriptional factor by a bacterial HMGA-type protein is required for regulating multiple processes in Myxococcus xanthus. Mol. Microbiol. 61:9 10-926. PCrez-Marin, M. C., J. J. Lopez-Rubio, F. J. Murillo, M. EliasArnanz, and S. Padmanabhan. 2004. The N-terminus of Myxococcus xanthus CarA repressor is an autonomously folding domain that mediates physical and functional interactions with both operator DNA and antirepressor protein. J. Biol. Chem. 279:33093-33103. Reeves, R. 2001. Molecular biology of HMGA proteins: hubs of nuclear function. Gene 27763-81. Reeves, R. 2003. HMGA proteins: flexibility finds a nuclear niche. Biochem. Cell Biol. 81:185-195. Reichenbach, H., and H. Kleinig. 1984. Pigments of myxobacteria, p. 128-137. In E. Rosenberg (ed.), Myxobacteria: Development and Cell Interactions. Springer-Verlag, New York, NY. Robson, P. R. H. 1992. Towards a Mechanism of Carotenogenesis in Myxococcus xanthus. Ph.D. thesis. University of Warwick, Coventry, England. Ruiz-Vazquez, R. M., M. Fontes, and F. J. Murillo. 1993. Clustering and co-ordinated activation of carotenoid genes in Myxococcus xanthus by blue light. Mol. Microbiol. 10:25-34. Schumann, G., H. Nurnberger, G. Sandmann, and H. Krugel. 1996. Activation and analysis of cryptic crt genes for carotenoid biosynthesis from Streptomyces griseus. Mol. Gen. Genet. 252:65 8-666. Takano, H., D. Asker, T. Beppu, and K. Ueda. 2006a. Genetic control for light-induced carotenoid production in nonphototrophic bacteria. J. Ind. Microbiol. Biotechnol. 33: 88-93. Takano, H., T. Beppu, and K. Ueda. 2006b. The CarMLitRfamily transcriptional regulator: its possible role as a photosensor and wide distribution in non-phototrophic bacteria. Biosci. Biotechnol. Biochem. 70:2320-2324. Takano, H., S. Obitsu, T. Beppu, and K. Ueda. 2005. Lightinduced carotenogenesis in Streptomyces coelicolor A3(2): identification of an extracytoplasmic function sigma factor that directs photodependent transcription of the carotenoid biosynthesis gene cluster. J. Bacteriol. 187:1825-1832. Takayama, F., T. Egashira, and Y. Yamanaka. 2001. Singlet oxygen generation from phosphatidylcholine hydroperoxide in the presence of copper. Life Sci. 68:1807-1815. Whitworth, D. E., S. J. Bryan, A. E. Berry, S. J. McGowan, and D. A. Hodgson. 2004. Genetic dissection of the lightinducible carQRS promoter region of Myxococcus xanthus. J. Bacteriol. 186:7836-7846. Whitworth, D. E., and D. A. Hodgson. 2001. Light-induced carotenogenesis in Myxococcus xanthus: evidence that Cars acts as an anti-repressor of CarA. Mol. Microbiol. 425309419.
Structure and
Myxobacteria: Multicellularity and Differentiation Edited by David E. Whitworth 0 2008 ASM Press, Washington, D.C.
Zhaomin Yang, Xue-yan Duan, Mehdi Esmaeiliyan, Heidi B. Kaplan
Composition, Structure, and Function of the Myxococcus xanthus Cell Envelope
The cell envelope functions as the boundary and interaction surface between the bacterial cell and its environment. All cell-cell interactions and environmental responses are sensed and transduced through the cell envelope. A typical gram-negative bacterium cell envelope consists of the inner membrane, the periplasmic region containing the peptidoglycan (PG) and soluble proteins, the outer membrane, the lipopolysaccharide (LPS), and in some cases an extracellular matrix (ECM) of capsular exopolysaccharide (EPS) and proteins (Raetz and Whitfield, 2002). The myxobacteria are gram-negative soil bacteria that exhibit a variety of social behaviors during all stages of their life cycle. Growing cells exhibit at least two group behaviors: cooperative feeding and social motility. Cells that are starved at high density on a solid surface initiate the complex multicellular behavioral response of fruiting body development in which approximately 100,000 cells aggregate into haystack-shaped mounds and differentiate into environmentally resistant spherical myxospores. Coordination of these social behaviors requires the exchange of a number of extracellular signals that
13
stimulate signal transduction pathways. Since Myxococcus xunthus serves as the premier model bacterium for the study of social behaviors, the investigation of its cell envelope through which the signals are sensed and transduced has been an active research area. This chapter covers in detail the polysaccharide-containing components of the M. xanthus cell envelope including the PG, LPS, ECM, and EPS.
PEPTIDOGLYCAN The cell wall PG forms the network of glycans and amino peptides that determines cell shape, protects the cell from the internal turgor pressure applied by the cytoplasmic contents, and plays a key role in cell division. The PG subunits (muropeptides)are synthesized in the cytoplasm and transported to the outer surface of the cytoplasmic membrane, where they are polymerized by transglycosylases that link the glycan moieties and transpeptidases that cross-link the amino peptides (Atrich et al., 1999). Specifically, PG consists of glycan strands of alternating p-1,4-linked N-acetylglucosamine (GlcNAc) and
Zhaomin Yang, Department of Biology, Virginia Polytechnic and State University, Blacksburg, VA 24060. Xue-yan Duan, Department of Molecular and Cellular Biology, Baylor College of Medicine, Houston, TX 77030. Mehdi Esmaeiliyan, Department of Natural Sciences, University of Houston/Downtown, Houston, TX 77002. Heidi B. Kaplan, Department of Microbiology and Molecular Genetics, University of Texas Medical School, Houston, TX 77030.
229
230 N-acetylmuramic acid (MurNAc) disaccharides that are cross-linked by short peptides. The structure of the Escherichia coli PG is typical of gram-negative bacteria. The PG monomer consists of the two amino sugars GlcNAc and MurNAc, with a 1,6 anhydro-N-acetylmuramic acid end (Vollmer and Holtje, 2004). The pentapeptide [L-ala-D-glu-(y)-m-dap-D-ala-D-ala] is attached to the MurNAc with cross-links that are formed between the D-alanine (ala) at position 4 of one stem peptide and mdiaminopimelic acid (dap) at position 3 of a second stem peptide of a neighboring glycan strand (Vollmer and Holtje, 2004). There has been no comprehensive analysis of the M . xanthus PG structure in more than 3 decades. White et al. (1968) found that the murein components of M. xanthus PG were similar to those of other gram-negative bacteria. The overall composition of the PG was 1.0 glutamic acid (glu), 1.0 dap, 1.7 ala, 0.75 GlcNAc, and 0.75 MurNAc. This analysis of the M. xanthus murein components revealed two additional findings. First, the PG was associated with substantial amounts of glycine, serine, and glucose. Second, the vegetative cell wall PG was suggested to be discontinuous in that whole sacculi were not isolated and trypsin and sodium dodecyl sulfate were able to completely disassociate the PG (White et al., 1968). Since the shape of the bacterial cell is dictated by the PG, it would be expected that during differentiation the M . xanthus cells would remodel their PG as they change from rod-shaped vegetative cells to spherical spores. It is unclear if this transformation involves de novo synthesis and/or breakage and restructuring of the cross-links between the muropeptide polymers (O’Connor and Zusman, 1999). White et al. (1968)and Johnson and White (1972) compared the PG of vegetative cells to starvationindependent spores (often referred to as glycerol spores). Although the overall composition and proportion by weight (0.9%) of both cell types were similar, there appeared to be an increase in muropeptide cross-linking in the spherical spores. Interestingly, O’Connor and Zusman (1997)noted that M . xanthus P-lactamase activity was induced by the same compounds that induce starvation-independent spore formation. These compounds are cell-wall-damaging agents, including 0.5 M glycerol, lysozyme, p-lactam antibiotics, D-amino acids, glycine, dimethyl sulfoxide, D-cycloserine, phosphomycin, phenyl ethanol, and ethylene glycol (O’Connor and Zusman, 1997; Dworkin and Gibson, 1964). This connection prompted them to propose that these two processes share common pathway components. It is likely that muropeptides function as the intermediate signal, since Jacobs et al. (1994) found that
STRUCTUREAND METABOLISM P-lactamase activity in E. coli can be controlled by alternations in the cytoplasmic levels of muropeptides (Park, 1995). O’Connor and Zusman (1999) also showed that P-lactamase activity is part of the normal developmental program, as an increase in its activity can be detected as soon as the developmental program is initiated. Other evidence suggests that external PG components have a direct role in the developmental program of M . xanthus. Shimkets and Kaiser (1982a, 1982b)showed that exogenously added PG and its components, specifically a mixture of GlcNAc, MurNAc, dap, and ala at 2.5 mM each, can initiate rippling, which is the organized movement of cells during early development, and can rescue sporulation of a csgA mutant. The mechanism of this rescue is unknown; however, Janssen and Dworkin (1985) showed that various sugars including glucosamine and mannosamine could also rescue sporulation of a csgA mutant. Shimkets and Kaiser (1982a, 198213) suggested that the lysis of cells during development releases a necessary amount of PG and other components required for the developmental process. O’Connor and Zusman (1999) noticed that Plactamase induction seems to influence the course of development, as the addition of P-lactamase inducers expedites the onset of aggregation and sporulation in a developing population of cells. They proposed that the induction of P-lactamase is likely to play a role in aggregation and in the restructuring of PG that occurs during differentiation into spores. However, it should also be considered that P-lactamase induction might reflect an alteration in the balance of cell wall synthesis and degradation that is anticipated to accompany the entrance into development as the cells arrest their growth. This would be expected to lead to the accumulation of muropeptides that would induce p-lactamase and serve as an indicator of cell wall integrity. A number of genes would .be predicted to be directly or indirectly expressed in response to a muropeptide signal. One group would be the early developmental genes controlled by the Che3 chemosensory system due to the activity in this pathway of the CrdB regulator that is predicted to have a PG-interaction domain (Kirby and Zusman, 2003). Other potential responsive genes would be those expressed early in development, such as L2444.5, which is regulated by an extracytoplasmic function (ECF) sigma factor-anti-ECF sigma factor pathway that senses envelope stress (Rivera, 2002). It is anticipated that further investigations of the biosynthesis and assembly of M . xanthus PG and its role in developmental signaling will lead to important insights into the structure of the cell envelope and the control of the developmental program.
13. M.XANTHUS CELLENVELOPE
PERIPLASMIC PROTEINS In addition to the cell-wall peptidoglycan and the enzymes involved in its biogenesis, the periplasmic space of gramnegative bacteria contains many enzymes and proteins of structural and functional importance. Among them are lipoproteins, hydrolytic enzymes, binding proteins important for nutrient acquisition, folding factors, and chaperons, and proteins that perform signaling/sensory functions and energy metabolism. Readers are directed to recent reviews on these topics (Behrens, 2003; Bos and Tommassen, 2004; Dwyer and Hellinga, 2004; Lomovskaya and Totrov, 2005; Mogensen and Otzen, 2005; Schlieker et al., 2004). Although little is known about the M. xanthus periplasm specifically, there are a few developmentally relevant periplasmic proteins including the glucose inhibition division (GidA) protein (White et al., 2001) and the periplasmic heat shock proteins (HSPs) (Nelson and Killeen, 1986). M. xanthus GidA, a flavin adenine nucleotide binding protein, was localized to the periplasm by cell fractionation and immunoelectron microscopy (White et al., 2001). M. xanthus gidA mutants appear to be unstable but are proficient in development. Stable derivatives, designated gidA*, arise after passage of the original mutants on solid media. The gidA* derivatives are defective in development and produce small heatstable and protease-resistant extracellular molecules that inhibit the development of wild-type cells. Intriguingly, the disruption of aglU, which maps immediately downstream of, and is transcribed with, gidA, eliminates the presence of the gzdA* developmentally inhibitory molecules. In contrast to the aglU mutants that are defective in adventurous (A)-motility, the gidA mutants are not (White and Hartzell, 2000; White et al., 2001). In other bacteria, GidA has been shown to be important for virulence and may function as a global regulator of gene expression (Kinscherf and Willis, 2002; Sha et al., 2004). White et al. (2001) suggest that GidA may play an important role in sensing or mediating redox-related phenomena and possibly couple replication and growth with cell division and differentiation. Heat-shocked developing M. xanthus cells produce at least two periplasmic HSPs that are not made by vegetative cells (Nelson and Killeen, 1986). The identities and the functions of these periplasmic HSPs remain to be determined; however, heat shock treatment prior to starvation or glycerol addition accelerates myxospore formation (Killeen and Nelson, 1988). Only one of the two HSPs is produced by heat-shocked starvationindependent spores (Nelson and Killeen, 1986), supporting the differences between starvation-independent and
23 1 starvation-dependent spores (Downard and Zusman, 1985; Shimkets and Seale, 1975).
IS THERE A PERIPLASMIC MOTILITY APPARATUS? Are there M. xanthus proteins in the periplasm that are involved in gliding motility? Chain-like aggregates or strands have been observed and isolated from the M. xanthus periplasm (Freese et al., 1997; Lunsdorf and Reichenbach, 1989). The isolated strands appear to be composed of ring-like and centrally elongated elements (Fig. l a ) . It is proposed that these strands align and form ribbon-like structures (Fig. 1b) with a periodicity visible by electron microscopy (EM). It was further suggested that these periplasmic structures could be part of the gliding machinery that may utilize membrane potential to power adventurous (A) gliding motility of myxobacteria (Freese et al., 1997; Lunsdorf and Schairer, 2001). However, it is important to note that there is no direct evidence linking these structures to gliding and the identity of the structural components remains unknown.
OUTER MEMBRANE PROTEINS Among the outer membrane proteins of M. xanthus two lipoproteins, CglB and Tgl, are important for A and social (S) gliding motility, respectively (see chapter 6). The motility defects of both cglB and tgl mutants can be rescued by direct contact with cells that produce wildtype CglB and Tgl, respectively (Hodgkin and Kaiser, 1979a, 197910; Rodriguez and Spormann, 1999; Wall et al., 1998).The underlying mechanism for the stimulation of motility by physical contact, in the case of Tgl, is the transfer of this lipoprotein from Tgl+ to Tgl- cells (Nudleman et al., 2005, 2006). It remains to be determined whether CglB and Tgl are anchored to the outer or inner leaflets of the outer membrane. The mechanism of lipoprotein transfer also remains unknown (Bayan et al., 2006). MBHA is a developmentally regulated lectin that accumulates to its highest level during the aggregation phase of fruiting body formation (Cumsky and Zusman, 1979). Although the bulk of MBHA appears to be cell surface associated, as it can be easily washed off cells, osmotic shock treatment of M. xanthus cells reveals that 20% of the MBHA protein is localized within the periplasm (Nelson et al., 1981). At least two factors regulate MBHA accumulation. First, m b h A transcription, which is 0-54 dependent, increases during development (Romeo and Zusman, 1991). Second, the stability of the m b h A mRNA is increased during development (Romeo
STRUCTUREAND METABOLISM
232
and Zusman, 1992). It is suspected that MBHA plays a role in M. xanthus cell cohesion (Romeo and Zusman, 1987),which is known to be important for fruiting body development (Shimkets, 1986a). Consistent with a role in cell cohesion, MBHA has four highly conserved domains that are predicted to form a multivalent structure (Romeo et al., 1986). MBHA-deficient strains are delayed in development but are otherwise able to aggregate and sporulate (Romeo and Zusman, 1987). It was suggested that there might be two systems for cell cohesion during development, of which only one requires magnesium (Romeo and Zusman, 1987). MBHA is predicted to function in a magnesium-independent cell cohesion system, since the developmental defects of mbhA mutants are more noticeable in the absence of magnesium (Romeo and Zusman, 1987).
LPS In gram-negative bacteria, the lipid component of the outer layer of the cell envelope is composed of LPS, which forms a selective barrier between the environment and the periplasmic region of the cell. The M. xanthus LPS is similar in general structure to the LPS of other gramnegative bacteria (Fink and Zissler, 1989).Each LPS molecule is composed of three parts: a hydrophobic lipid A,
which comprises the lipid portion of the outer leaflet of the outer membrane; a covalently attached nonrepeating core oligosaccharide region; and a distal repeating polysaccharide termed 0 antigen (Raetz and Whitfield, 2002). This general structure of the M. xanthus LPS was revealed by studies of LPS biosynthesis mutants isolated in genetic screens using monoclonal antibodies (MAbs) generated against the cell surface of M. xanthus cells (Gill et al., 1985; Gill and Dworkin, 1986). One set of MAbs (2600, 1733, 1514, 1412, and 783) was determined to recognize LPS 0 antigen based on their reactivity to the ladder of bands of crude LPS preparations separated on polyacrylamide gels (Fink and Zissler, 1989). Another MAb, 2254, recognizes LPS core, as its reactivity is confined to the lowest band on these gels (Fink and Zissler, 1989). Five Tn.5 transposon mutants were identified, which are nonreactive to MAbs that recognize the LPS 0 antigen, suggesting that the insertion mutations altered the biosynthesis or assembly of the LPS 0 antigen (Fink et al., 1989).When analyzed in detail, these mutants were determined to have a number of phenotypes including a defect in S-motility, whereas A-motility remained unaffected (Bowden and Kaplan, 1998). Kaplan et al. (1991) identified the sasA locus using UV mutagenesis in a genetic suppressor screen designed to identify elements in a pathway linking extracellular A
Figure 1 Chain-like strand in the M. xantbus periplasm and the structural model. (a) Single strand fragment associated with a membrane fragment (MF).The ring elements (arrowheads) stand oblique to the plane of the carbon support. Inset: ring elements (circle) and central elongated elements (dots) are shown at higher magnification; arrow indicates the spoke-like central mass. Bar, 50 nm (inset, 30 nm). (b). Three-dimensional model of the location of strands within the cell wall of M. xanthus. These strands are proposed to be inserted between the flexible peptidoglycan sacculus and the outer membrane. RE, ring element, framed; EE, elongated element; CM, cytoplasmic membrane; OM, outer membrane; cema, central mass; pm, peripheral mass. Adapted from Freese et al., 1997.
13. 211. XANTHUS CELLENVELOPE signal to its responsive gene, 4521. One of the sasA alleles mapped to the wzm gene (formally rfbA).The wzm gene and its downstream gene wzt (formally rfaB) encode an ABC transporter, which is predicated to be required for the export of LPS 0 antigen into the periplasm (Guo et al. 1996). Recently, a similar genetic screen with the newly developed mini-Himarl transposon identified many more LPS biosynthesis genes, which encode a majority of the M. xanthus sugar biosynthesis enzymes (X.-Y. Duan, M. Esmaeiliyan, and H. B. Kaplan, unpublished data). Youderian and Hartzell (2006) conducted an extensive screen for S-motility mutants with the magellan-4 transposon and also identified many genes involved in LPS biosynthesis. Interestingly, all of the LPS mutants identified by our two laboratories mapped to three loci: two LPS O-antigen loci and one LPS core locus. All the LPS mutants that have been identified have developmental defects. They form defective fruiting bodies and sporulate at reduced efficiencies (Fink et al., 1989; Guo et al., 1996; Bowden and Kaplan, 1998; Youderian and Hartzell, 2006; Duan et al., unpublished). Further analysis of some LPS mutants showed that they are defective in S-motility,whereas they are wild type for A-motility (Bowden and Kaplan, 1998; Duan et al., unpublished). These data support the concept that different mechanisms control A- and S-motilityand indicate that S-motilityplays a major role in M. xanthus fruiting body formation. The carbohydrate composition of wild-type LPS consists of glucose, mannose, rhamnose, arabinose, xylose, galactosamine, glucosamine, KDO (2-keto-3-deoxyoctulosonic acid), and 3-O-methylpentose and 6-O-methylgalactosamine (Ashton, 1993). Interestingly, the 111. xanthus EPS is proposed to contain five of these monosaccharides: galactose, glucosamine, glucose, rhamnose, and xylose (Behmlander and Dworkin, 1994b). It is possible that the LPS preparations analyzed were contaminated with EPS, which would also partition to the aqueous phase with the LPS hot-phenol extraction method. Currently comprehensive analyses of both the LPS and EPS structures are being pursued (H. B. Kaplan and W. Shi, personal communication). Genomic and genetic analyses to identify all of the genes required for M. xanthus LPS biosynthesis and assembly are certain to be pursued in the future. In addition, a thorough biochemical analysis of the structures of mutant M. xanthus LPS molecules and the activities of the biosynthetic enzymes will be critical to elucidate the pathways involved in the production of the individual sugars and the assembly of the LPS. Furthermore, an analysis of the mechanisms underlying the increase in expression of early developmental genes, such as 4521, in the absence of LPS 0 antigen should reveal interesting
233 signaling pathways. It is likely that the LPS-dependent effects on 4522 developmental expression, which is controlled by the SasS/R/N histidine kinase three-component system (Yang and Kaplan, 1997; Xu and Kaplan, 1998; Guo et al., 2000), are responsive to envelope stress.
ECM STRUCTURE AND FUNCTION M . xanthus cells are covered by an ECM comprised of approximately equal amounts by weight of protein and polysaccharide (Kim et al., 1999; Merroun et al., 2003; Behmlander and Dworkin, 1994b), which was previously referred to as fibrils (Fig. 2). The word “fibril” has been used to describe the apparent filamentous fibers on the cell surface of M. xanthus cells observed by scanning EM (SEM) (Dworkin, 1999) (Fig. 2C and D). The fiber-like structures observed using SEM are most likely the result of dehydration of the polysaccharide during sample preparation. The polysaccharide component of the ECM is now referred to as EPS (Lancer0 et al., 2004; Lu et al., 2005; Xu et al., 2005; Yang and Li, 2005). M . xunthus ECM is required for cellular cohesion, Smotility, and fruiting body morphogenesis. Early genetic studies indicated that dsp mutants, which grow dispersed and are defectivein cellular cohesion, lack S-motilityand are unable to form fruiting bodies (Shimkets, 1986a, 198610). It was established later that dsp mutants are altered in their surface properties and lack ECM when examined by EM (Arnold and Shimkets, 198810; Behmlander and Dworkin, 1991). The correlation between the presence of ECM and cellular cohesion was also demonstrated using the diazo dye Congo red that can disrupt the formation of ECM and inhibited both cellular cohesion and fruiting body development (Arnold and Shimkets, 1988a, 198813).These observations provided the evidence that cohesive forces may be necessary to maintain the integrity of cell groups, which is important for S-motility and fruiting body formation. The ECM function in S-motility has become clearer in recent years. Yang et al. (2000) suggested that the ECM provides more than a cohesion force for S-motility (Yang et al., 2000). It was postulated that the ECM is involved in the activation of the S-motility motor in a cell-proximity-dependent manner (Yang et al., 2000). Li et al. (2003)suggested that EPS provides the anchor for the retraction of type IV pili (Tfp),which are now widely accepted as the motor for M. xanthus S-motility and bacterial twitching motility (see chapter 6 ) . In essence, EPS from neighboring cells may anchor Tfp at the distal end and this attachment or anchoring may in turn activate the retraction of Tfp at the base. It has been suggested that the signal for tethering or attachment at the distal end may be transmitted to the cell body mechanically
STRUCTUREAND METABOLISM
234 through structural restraints or shifts (Li et al., 2003; Black et al., 2006).Interestingly, EPS may not be the only material that can trigger or activate the S-motility motor. An increase in the viscosity of the medium by the addition of methylcellulose can stimulate the S-motility-dependent movement of individual cells (Sun et al., 2000). The EPS component of M. xanthus ECM is necessary for the structural integrity of the ECM (Behmlander and Dworkin, 199413). Whereas digestion of carbohydrate by periodic acid essentially destroys the observed ECM structure, EM observations indicate that protein removal by protease does not significantly alter the structure of isolated ECM. Furthermore, the ECM proteins are not required for M. xanthus S-motility as treatment with
proteases does not appreciably affect the activity of ECM in causing Tfp to disappear from cell surfaces (Li et al., 2003), and mutations in fibA, which encodes the only known ECM protein, result in no obvious defects in ECM structure and S-motility (Kearns et al., 2002). The ECM may be considered to be a layer of EPS associated with nonspecific outer membrane proteins.
REGULATION OF EPS PRODUCTION The production of EPS is a highly regulated process. Early work suggested that cellular cohesion and surface properties are likely regulated by cell density (Arnold and Shimkets, 1988b; Behmlander and Dworkin,
Figure 2 ECM visualized by different EM techniques. (A and B) Transmission EM micrographs prepared either by spray-freezing freeze substitution (A) or by staining with lanthanum and fixation with glutaraldehyde (B). (C and D) SEM images at different magnifications. Samples in panels C and D were fixed with glutaraldehyde and coated with platinum. Fig. 2A from Kim et al., 1999; Fig. 2B from Merroun et al., 2003; Fig. 2C and D from Behmlander and Dworkin, 1991.
13. 111. XANTHUS CELLENVELOPE 1991), and more recent work indicates a nutritional dependency of the two different motility systems (Hillesland and Velicer, 2005). Genetic evidence provides strong support for EPS regulation in M. xanthus. The dsp mutants were the first mutants known to display defects in cellular cohesion (Arnold and Shimkets, 1988b). Another group of mutants, the sticky (stk) mutants, show an increased ability to agglutinate and a strong tendency to adhere to surfaces (Dana and Shimkets, 1993). Both groups show aberrant ECM production with dsp mutants producing no detectable ECM and stk mutants overproducing ECM (Arnold and Shimkets, 1988b; Dana and Shimkets, 1993). In addition, mgl, tgl, and sgl mutants, which are all defective in S-motility, were found to have reduced levels of ECM by dye binding and/or EM (Dana and Shimkets, 1993). ECM biosynthesis and assembly are regulated by the dif locus, which was identified by mutants defective in development and encodes a set of proteins that are homologs of the well-studied chemotaxis proteins of the enteric bacteria and Bacillus subtilis (Black and Yang, 2004; Yang et al., 1998b) (see chapter 8). DifA is a homolog of the methyl-accepting chemotaxis proteins, DifC is a homolog of the coupling protein Chew, DifD is a homolog of the response regulator CheY, DifE is a homolog of the histidine kinase CheA, and DifG is a homolog of the protein phosphatase CheC. The Dif proteins both positively and negatively regulate EPS production. Mutations in difA, difC, and difE virtually eliminate EPS production, whereas those in difD and difG result in EPS overproduction (Bellenger et al., 2002; Black and Yang, 2004; Xu et al., 2005; Yang et al., 1998b; Yang et al., 2000). Genetic epistasis tests indicate that DifD does not function downstream of DifE in the regulation of EPS (Black et al., 2006). Based on these observations, a model is proposed for the regulation of EPS by a pathway consisting of Dif proteins (Fig. 3) (Black et al., 2006). In this model, DifA, DifC, and DifE are proposed to form a signaling complex anchored to the membrane by the MCP homolog DifA. This is analogous to the classical chemotaxis pathway where MCPs, Chew, and CheA form a membrane signaling complex (Webre et al., 2003). It is further proposed that stimulation of the DifA-DifC and DifE complex leads to increased DifE kinase activity. DifE would modulate EPS production by a phosphorylation-dependent mechanism through components yet to be identified. DifD is proposed as a phosphate sink and would be analogous in function to the CheY2 of Rhizobium rneliloti chemotaxis (Sourjik and Schmitt, 1996, 1998). DifG is proposed to accelerate the dephosphorylation of DifD
235 phosphate (DifD-P) (Black and Yang, 2004). Moreover, DifG must somehow inhibit the central pathway independently of DifD because mutations in d i p and difG show certain additive effects. The formation of a ternary complex by DifA, DifC, and DifE is supported by yeast two- and three-hybrid experiments (Yang and Li, 2005), although the proposed phosphorylation reactions between the Dif proteins have not been biochemically demonstrated. It was shown recently that signal perception by the Dif pathway involves Tfp (Black et al., 2006). First, Tfp was found to be required for EPS production in M. xanthus. Second, Dif proteins function downstream of Tfp as demonstrated by genetic epistasis tests. Finally, Tfp do not appear to function as either exogenous or endogenous signals for the Dif pathway. It can therefore be inferred that 111. xanthus Tfp function as a sensor or part of a sensory apparatus for the perception of signals for the Dif pathway in the regulation of EPS production. It was further proposed that S-motility involves a regulatory loop in which EPS triggers Tfp retraction
Figure 3 Model depicting the regulation of EPS production in M. xanthus by Tfp and the Dif pathway. Demonstrated interactions are indicated by solid lines, and proposed interactions are indicated by dashed lines. Arrows and bars indicate positive and negative regulation, respectively. See the text for details of the model. Adapted.from Black et al., 2006.
STRUCTUREAND METABOLISM
236 and Tfp provide proximity signals to the Dif pathway to modulate EPS production (Black et al., 2006). This
proposal is based on the following concepts: Tfp are the likely motors for S-motility, EPS appears to regulate Tfp retraction, and S-motility requires cell proximity for normal function. Besides the Dif proteins and Tfp, other proteins are likely to play regulatory roles in M. xanthus EPS production. Two of the best known are the DnaK homologs StkA (described above) and SglK. Whereas stkA mutants are more cohesive and overproduce EPS (Dana and Shimkets, 1993), sglK null mutants lack EPS (Weimer et al., 1998; Yang et al., 1998a) and show virtually no group movement on soft-agar plates. Surprisingly, an sglK insertion mutant at the 3’ end of the gene forms fractal patterns on soft-agar plates (MacNeil et al., 1994). The fibR gene, which is linked to the sglK locus, seems to encode a negative regulator of ECM, because a fibR mutant produces an increased amount of the ECM protein FibA (Weimer et al., 1998). The possible regulatory function of FibR is further supported by the fact that it is a homolog of the repressors of alginate production in Pseudomonas spp. It is likely that stkA, sglK, and fibR are all regulators of M . xanthus EPS biosynthesis based on the phenotypes of their corresponding mutants and that they are all homologs of regulatory proteins. There are a number of other potential or confirmed regulators of M . xanthus EPS biosynthesis. The NtrClike activator Nla24 that regulates both A- and S-motility (Caberoy et al., 2003; Lancero et al., 2004) is thought to regulate S-motility due to its control of EPS production (Lancero et al., 2004). The nla24 mutant lacks EPS production. Similarly, three additional NtrC-like activators, Nlal, Nla19, and Nla23, are potential regulators of M . xanthus EPS biosynthesis (Caberoy et al., 2003). Another regulator of M. xanthus EPS is the tyrosine kinase MasK (Thomasson et al., 2002). The masK815 mutant allele, which harbors two missense mutations, was isolated as a partial suppressor of the S-motility defect resulting from an mglA missense mutation. A masK81.S mglA8 double mutant overproduced both EPS and the ECM protein FibA. Distinct from other EPS regulators, masK appears to be an essential gene in M. xanthus. Other less well characterized proteins are involved in M. xanthus EPS biosynthesis in a regulatory or biosynthetic capacity. The protein pair RppA and MmrA are homologs of a sensory transducer and a multidrugresistant protein, respectively, that appear to be important for M. xanthus EPS or polysaccharide biosynthesis (Kimura et al., 2004). It is puzzling that although single mutations in either rppA or mmrA showed little effect on polysaccharide production, an rppA mmrA double
mutant showed a decreased amount of polysaccharide production. It is unclear how a transducer and a multidrug transporter could perform redundant or synergistic functions in either EPS regulation or biosynthesis. Other proteins with ambiguous functions in EPS production include RasA, EsgA, and MglA (Dana and Shimkets, 1993; Pham et al., 2005; Ramaswamy et al., 1997). Further investigation is necessary to fully understand the complexity of EPS regulation in M. xanthus.
ECM BIOGENESIS Preliminary biochemical studies indicated that M . xanthus EPS contains at least five monosaccharides: galactose, glucosamine, glucose, rhamnose, and xylose (Behmlander and Dworkin, 1994b). From the newly sequenced M . xanthus genome, pathways for producing all these sugars in either UDP-, GDP- or dTDP-activated forms can be constructed ( Z . Yang and H. B. Kaplan, unpublished data). The activated monosaccharides may be used for EPS production by synthetic enzymes, some of which are likely encoded by the eps locus (Lu et al., 2005). The deduced eps products include homologs of various polysaccharide synthesis proteins such as glycosyltransferases and polysaccharideexport proteins. Mutations in some of the esp genes and the two eas genes have been shown to result in defects in EPS production (Lu et al., 2005; Barbu et al., unpublished). The Nla24 protein (described above) is encoded by the eps locus (Lancero et al., 2004; Lu et al., 2005). The eps locus likely encodes additional regulators of EPS production because many of the open reading frames are homologs of regulatory proteins such as histidine kinases and response regulators (Lu et al., 2005). The exact functions of these potential biosynthetic enzymes and regulatory proteins remain to be elucidated. As described above, the only known ECM protein is FibA. It was identified by its reactivity with a monoclonal antibody (MAb2105) raised against cell surface antigens of M . xanthus (Behmlanderand Dworkin, 1994a; Gill et al., 1985).It was initially puzzling that MAb2105 recognized proteins with apparently different molecular weights or electrophoretic mobility. Behmlander and Dworkin (1994a) suggested that these proteins arose from the same protein product. Amino acid sequencing of the 31-kDa protein (Behmlander and Dworkin, 1994a) and the availability of M. xanthus genome sequence led to the discovery of the fibA gene (Kearns et al., 2002). The deduced polypeptide sequence suggests that FibA is a zinc metalloprotease. The immunoblot results suggest that FibA undergoes autoproteolytic processing that generates multiple polypeptides
23 7
13. M. XANTHUS CELLENVELOPE recognizable by MAb2105 (Kearns et al., 2002). The fibA mutants are defective in tactic responses to dilauroyl (12:O) phosphatidylethanolamine (PE) but have normal responses to dioleoyl (18:l) PE. The fruiting bodies formed by fibA mutants are elongated and irregularly shaped. Otherwise, fibA mutants are proficient in cellular cohesion and in both A- and S-motility, further suggesting that ECM proteins are not critical for the structure of ECM or its function in cohesion and S-motility. The mechanisms by which FibA is involved in PE taxis are not understood. FibA may be involved in sensory functions or responsible for the proteolytic processing of proteins that participate in the M. xanthus PE response.
PILI M. xanthus cells possess polar pili that can be visualized by negative staining and transmission EM (Dobson and McCurdy, 1979; Dobson et al., 1979; Kaiser, 1979). The pili are less than 10 nm in diameter and are usually present only at one of the two cell poles. Recent observations with the atomic force microscope confirm the dimensions (5 to 8 nm) and polar localization of M. xanthus pili (Pelling et al., 2005). Consistent with the polar localization, the sequence of M. xanthus pilA, which encodes the pilin subunit of the polymeric pilus fiber, indicates that M. xanthus pili belong to the class of Tfp which are found in gram-negative bacteria (Wu and Kaiser, 1995).Bacterial Tfp are important for many processes including surface movement, competency, and host colonization by pathogens. Readers are directed to recent reviews on Tfp (Craig et al., 2004; Mattick, 2002; Meibom et al., 2005; Pizarro-Cerda and Cossart, 2006; Wall and Kaiser, 1999). The atomic structures of pilins from a few organisms have been solved by X-ray crystallography or nuclear magnetic resonance (Craig et al., 2004; Ramboarina et al., 2005). Studies of M . xanthus pili focus on their involvement in S-motility. Dale Kaiser was the first to observe a tight association between pili and S-motility (Kaiser, 1979). This was confirmed by the observation that mechanical removal of M. xanthus Tfp was accompanied by the loss of S-motility (Rosenbluh and Eisenbach, 1992). Further studies provided additional support for the requirement of Tfp for S-motility (Nudleman et al., 2005,2006; Wall et al., 1998; Wall et al., 1999; Wu and Kaiser, 1995, 1996; Wu et al., 1997, 1998; Youderian and Hartzell, 2005). It is generally accepted that Tfp retraction provides the force for M. xanthus S-motility and the related surface motility known as bacterial twitching (Mattick, 2002; Nudleman and Kaiser, 2004). Tfp are proposed
to attach to their targets either on cells or other surfaces and pull the cells forward by retracting. The pulling force of Tfp retraction was measured by laser tweezers (Maier et al., 2002, 2004; Merz et al., 2000), and the retraction was observed directly using a novel combination of staining with amino-specific Cy3 fluorescent dye and total internal reflection microscopy (Skerker and Berg, 2001). There is also indirect evidence for pilus retraction in M. xanthus (Sun et al., 2000). Readers are directed to chapter 6 in this book for more information on the function of Tfp in M . xanthus S-motility.
References Arnold, J. W., and L. J. Shimkets. 1988a. Inhibition of cell-cell interactions in Myxococcus xanthus by Congo red. J. Bacteriol. 1705765-5770. Arnold, J. W., and L. J. Shimkets. 1988b. Cell surface properties correlated with cohesion in Myxococcus xanthus. J. Bacteriol. 1705771-5777. Ashton, A. 1993. Structural Studies of the Lipopolysaccharide from Myxococcus xanthus and Lipopolysaccharide Mutants. Ph.D. thesis. University of Minnesota, Minneapolis. Atrich, A., G. Bacher, G. Allmaier, M. P. Williamson, and S. J. Foster. 1999. Analysis of peptidoglycan structure from vegetative cells of Bacillus subtilis 168 and role of PBP 5 in peptidoglycan maturation. J. Bacteriol. 181:3956-3966. Bayan, N., I., Guilvout, and A. I?. Pugsley. 2006. Secretins take shape. Mol. Microbiol. 60:1-4. Behmlander, R. M., and M. Dworkin. 1991. Extracellular fibrils and contact-mediated cell interactions in Myxococcus xanthus. J. Bacteriol. 173:7810-7820. Behmlander, R. M., and M. Dworkin. 1994a. Integral proteins of the extracellular matrix fibrils of Myxococcus xanthus. J. Bacteriol. 176:6304-63 11. Behmlander, R. M., and M. Dworkin. 199413. Biochemical and structural analyses of the extracellular matrix fibrils of Myxococcus xanthus. J. Bacteriol. 176:6295-6303. Behrens, S. 2003. Periplasmic chaperones-preservers of subunit folding energy for organelle assembly. Cell 113556-557. Bellenger, K., X. Ma, W. Shi, and Z. Yang. 2002. A Chew homologue is required for Myxococcus xanthus fruiting body development, social gliding motility, and fibril biogenesis. J. Bacteriol. 1845654-5660. Black, W. P., and Z. Yang. 2004. Myxococcus xanthus chemotaxis homologs DifD and DifG negatively regulate fibril polysaccharide production. J. Bacteriol. 186:1001-100 8. Black, W. P., Q. Xu, and Z. Yang. 2006. Type IV pili function upstream of the Dif chemotaxis pathway in Myxococcus xanthus EPS regulation. Mol. Microbiol. 61:447-456. Bos, M. P., and J. Tommassen. 2004. Biogenesis of the Gramnegative bacterial outer membrane. Curr. Opin. Microbiol. 7~610-616. Bowden, M. G., and H. B. Kaplan. 1998. The Myxococcus xanthus lipopolysaccharide 0-antigen is required for social motility and multicellular development. Mol. Microbiol. 30:275-284.
238 Caberoy, N. B., R. D. Welch, J. S., Jakobsen, S. C., Slater, and A. G. Garza. 2003. Global mutational analysis of NtrC-like activators in Myxococcus xanthus: identifying activator mutants defective for motility and fruiting body development.]. Bacteriol. 185:6083-6094. Craig, L., M. E. Pique, and J. A. Tainer. 2004. Type IV pilus structure and bacterial pathogenicity. Nat. Rev. Microbiol. 2~363-378. Cumsky, M., and D. R. Zusman. 1979. Myxobacterial hemagglutinin: a development-specific lectin of Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 765505-5509. Dana, J. R., and L. J. Shimkets. 1993. Regulation of cohesiondependent cell interactions in Myxococcus xanthus. J. Bacteriol. 175:3636-3647. Dobson, W. J., and H. D. McCurdy. 1979. The function of fimbriae in Myxococcus xanthus. I. Purification and properties of M . xanthus fimbriae. Can. J. Microbiol. 25:1152-1160. Dobson, W. J., H. D. McCurdy, and T. H. MacRae. 1979. The function of fimbriae in Myxococcus xanthus. 11. The role of fimbriae in cell-cell interactions. Can. J. Microbiol. 25~1359-1372. Downard, J. S., and D. R. Zusman. 1985. Differential expression of protein S genes during Myxococcus xanthus development. J. Bacteriol. 161:1146-1155. Dworkin, M. 1999. Fibrils as extracellular appendages of bacteria: their role in contact-mediated cell-cell interactions in Myxococcus xanthus. Bioessays 21590-595. Dworkin, M., and S. M. Gibson. 1964. A system for studying microbial morphogenesis: rapid formation of microcysts in Myxococcus xanthus. Science 146:243-244. Dwyer, M. A., and G. J. Hellinga. 2004. Periplasmic binding proteins: a versatile superfamily for protein engineering. Curr. Opin. Struct. Biol. 14:495-504. Fink, J. M., and J. F. Zissler. 1989. Characterization of lipopolysaccharide from Myxococcus xanthus by use of monoclonal antibodies. /. Bacteriol. 171:2028-2032. Fink, J. M., M. Kalos, and J. F. Zissler. 1989. Isolation of cell surface antigen mutants of Myxococcus xanthus by use of monoclonal antibodies. J. Bacteriol. 171:2033-2041. Freese, A., H. Reichenbach, and H. Lunsdorf. 1997. Further characterization and in situ localization of chain-like aggregates of the gliding bacteria Myxococcus fulvus and Myxococcus xanthus. J. Bacteriol. 179:1246-1252. Gill, J., E. Stellwag, and M. Dworkin. 1985. Monoclonal antibodies against cell-surface antigens of developing cells of Myxococcus xanthus. Ann. Inst. Pasteur. Microbiol. 136A:ll-18. Gill, J. S., and M. Dworkin. 1986. Cell surface antigens during submerged development of Myxococcus xanthus examined with monoclonal antibodies. J. Bacteriol. 168505-5 11. Guo, D., Y. Wu, and H. B. Kaplan. 2000. Identification and characterization of genes required for early Myxococcus xanthus developmental gene expression. J. Bacteriol. 182~4564-4571. Guo, D., M. G. Bowden, R. Pershad, and H. B. Kaplan. 1996. The Myxococcus xanthus rfbABC operon encodes an ATPbinding cassette transporter homolog required for 0-antigen biosynthesis and multicellular development. J. Bacteriol. 178~1631-1639.
STRUCTUREAND METABOLISM Hillesland, K. L., and G. J. Velicer. 2005. Resource level affects relative performance of the two motility systems of Myxococcus xanthus. Microb. Ecol. 49558-566. Hodgkin, J., and D. Kaiser. 1979a. Genetics of gliding motility in Myxococcus xanthus: two gene systems control movement. Mol. Gen. Genet. 171:177-191. Hodgkin, J., and D. Kaiser. 1979b. Genetics of gliding motility in Myxococcus xanthus (Myxobacterales):genes controlling movement of single cells. Mol. Gen. Genet. 171:167-176. Jacobs, C., L. J. Huang, E. Bartowsky, S. Normark, and J. T. Park. 1994. Bacterial cell wall recycling provides cytosolic muropeptides as effectors for p-lactamase induction. EMBO J. 13:4684-4694. Janssen, G. R., and M. Dworkin. 1985. Cell-cell interactions in developmental lysis of Myxococcus xanthus. Dev. Biol. 112:194-202. Johnson, Y. R., and D. White. 1972. Myxospore formation in Myxococcus xanthus: chemical changes in the cell wall during cellular morphogenesis. J. Bacteriol. 1123349-855. Kaiser, D. 1979. Social gliding is correlated with the presence of pili in Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 7659.52-5956. Kaplan, H. B., A. Kuspa, and D. Kaiser. 1991. Suppressors that permit A-signal-independent developmental gene expression in M ~ X O C O C C xanthus. US J. Bacteriol. 173:1460-1470. Kearns, D. B., P. J. Bonner, D. R. Smith, and L. J. Shimkets. 2002. An extracellular matrix-associated zinc metalloprotease is required for dilauroyl phosphatidylethanolamine chemotactic excitation in Myxococcus xanthus. J. Bacteriol. 184:1678-1684. Killeen, K. P., and D. R. Nelson. 1988. Acceleration of starvation- and glycerol-induced myxospore formation by prior heat shock in Myxococcus xanthus. J. Bacteriol. 17052005207. Kim, S. H., S. Ramaswamy, and J. Downard. 1999. Regulated exopolysaccharide production in Myxococcus xanthus. J. Bacteriol. 181:1496-1507. Kimura, Y., S. Ishida, H. Matoba, and N. Okahisa. 2004. RppA, a transducer homologue, and MmrA, a multidrug transporter homologue, are involved in the biogenesis and/or assembly of polysaccharide in Myxococcus xanthus. Microbiology 150:631-639. Kinscherf, T. G., and D. K. Willis. 2002. Global regulation by gidA in Pseudomonas syringae. J. Bacteriol. 184:22812286. Kirby, J. R., and D. R. Zusman. 2003. Chemosensory regulation of developmental gene expression in Myxococcus xanthus. Proc. Natl. Acad. Sci. U S A 100:2008-2013. Lancero, H., N. B. Caberoy, S. Castaneda, Y. Li, A. Lu, D. Dutton, X. Y. Duan, H. B. Kaplan, W. Shi, and A. G. Garza. 2004. Characterization of a Myxococcus xanthus mutant that is defective for adventurous motility and social motility. Microbiology 150:4085-4093. Li, Y., H. Sun, X. Ma, A. Lu, R. Lux, D. Zusman, and W. Shi. 2003. Extracellular polysaccharides mediate pilus retraction during social motility of Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 1005443-5448. Lomovskaya, O., and M. Totrov. 2005. Vacuuming the periplasm. J. Bacteriol. 187:1879-1883.
13. M . XANTHUS CELLENVELOPE Lu, A., K. Cho, W. P. Black, X. Y. Duan, R. Lux, Z. Yang, H. B. Kaplan, D. R. Zusman, and W. Shi. 2005. Exopolysaccharide biosynthesis genes required for social motility in Myxococcus xanthus. Mol. Microbiol. 55:206-220. Lunsdorf, H., and H. Reichenbach. 1989. Ultrastructural details of the apparatus of gliding motility of Myxococcus fulvus (Myxobacterales).Microbiology 135:1633-1 64 1. Lunsdorf, H., and H. U. Schairer. 2001. Frozen motion of gliding bacteria outlines inherent features of the motility apparatus. Microbiology 147:939-947. MacNeil, S. D., A. Mouzeyan, and P. L. Hartzell. 1994. Genes required for both gliding motility and development in M y x o coccus xanthus. Mol. Microbiol. 14:785-795. Maier, B., L. Potter, M. So, C. D. Long, H. S. Seifert, and M. P. Sheetz. 2002. Single pilus motor forces exceed 100 pN. Proc. Natl. Acad. Sci. U S A 99:16012-16017. Maier, B., M. Koomey, and M. P. Sheetz. 2004. A forcedependent switch reverses type IV pilus retraction. Proc. Natl. Acad. Sci. U S A 101:10961-10966. Mattick, J. S. 2002. Type IV pili and twitching motility. Annu. Rev. Microbiol. 56:289-314. Meibom, K. L., M. Blokesch, N. A. Dolganov, C. Y. Wu, and G. K. Schoolnik. 2005. Chitin induces natural competence in Vibrio cholerae. Science 310:1824-1827. Merroun, M. L., K. Ben Chekroun, J. M. Arias, and M. T. Gonzalez-Munoz. 2003. Lanthanum fixation by Myxococcus xanthus: cellular location and extracellular polysaccharide observation. Chemosphere 52:113-120. Merz, A. J., M. So, and M. P. Sheetz. 2000. Pilus retraction powers bacterial twitching motility. Nature 407:98-102. Mogensen, J. E., and D. E. Otzen. 2005. Interactions between folding factors and bacterial outer membrane proteins. Mol. Microbiol. 57326-346. Nelson, D. R., M. G. Cumsky, andD. R. Zusman. 1981. Localization of myxobacterial hemagglutinin in the periplasmic space and on the cell surface of Myxococcus xanthus during developmental aggregation. J. Biol. Chem. 256:12589-12595. Nelson, D. R., and K. I? Killeen. 1986. Heat shock proteins of vegetative and fruiting Myxococcus xanthus cells. J. Bacterial. 168:1100-1 106. Nudleman, E., and D. Kaiser. 2004. Pulling together with type IV pili. J . Mol. Microbiol. Biotechnol. 752-62. Nudleman, E., D. Wall, and D. Kaiser. 2005. Cell-to-cell transfer of bacterial outer membrane lipoproteins. Science 309~125-127. Nudleman, E., D. Wall, and D. Kaiser. 2006. Polar assembly of the type IV pilus secretin in Myxococcus xanthus. Mol. Microbiol. 60:16-29. O’Connor, K. A., and D. R. Zusman. 1997. Starvationindependent sporulation in Myxococcus xanthus involves the pathway for p-lactamase induction and provides a mechanism for competitive cell survival. Mol. Microbiol. 24~839-850. O’Connor, K. A., and D. R. Zusman. 1999. Induction of plactamase influences the course of development in Myxococcus xanthus. J . Bacteriol. 181:6319-6331. Park, J. T. 1995. Why does Escherichia coli recycle its cell wall peptides? Mol. Microbiol. 17:421-426.
239 Pelling, A. E., Y. Li, W. Shi, and J. K. Gimzewski. 2005. Nanoscale visualization and characterization of Myxococcus xanthus cells with atomic force microscopy. Proc. Natl. Acad. Sci. U S A 102:6484-6489. Pham, V. D., C. W. Shebelut, B. Mukherjee, and M. Singe. 2005. RasA is required for Myxococcus xanthus development and social motility. J. Bacteriol. 187:6845-6848. Pizarro-Cerda, J., and P. Cossart. 2006. Bacterial adhesion and entry into host cells. Cell 124:715-727. Raetz, C. R., and C. Whitfield. 2002. Lipopolysaccharide endotoxins. Annu. Rev. Biochem. 71:635-700. Ramaswamy, S., M. Dworkin, and J. Downard. 1997. Identification and characterization of Myxococcus xanthus mutants deficient in calcofluor white binding.]. Bacteriol. 179:28632871. Ramboarina, S., P. J. Fernandes, S. Daniell, S. Islam, P. Simpson, G. Frankel, F. Booy, M. S. Donnenberg, and S. Matthews. 2005. Structure of the bundle-forming pilus from enteropathogenic Escherichia coli. J. Biol. Chem. 280:40252-40260. Rivera, J. J. 2002. An Extracytoplasmic Function Sigma Factor Operon Regulates Myxococcus xanthus Developmental Gene Expression. Ph.D. thesis. University of Texas, Houston. Rodriguez, A. M., and A. M. Spormann. 1999. Genetic and molecular analysis of cglB, a gene essential for single-cell gliding in Myxococcus xanthus.J. Bacteriol. 181:4381-4390. Romeo, J. M., B. Esmon, and D. R. Zusman. 1986. Nucleotide sequence of the myxobacterial hemagglutinin gene contains four homologous domains. Proc. Natl. Acad. Sci. U S A 83:6332-63 36. Romeo, J. M., and D. R. Zusman. 1987. Cloning of the gene for myxobacterial hemagglutinin and isolation and analysis of structural gene mutations. J. Bacteriol. 169:3801-3808. Romeo, J. M., and D. R. Zusman. 1991. Transcription of the myxobacterial hemagglutinin gene is mediated by a sigma 54-like promoter and a cis-acting upstream regulatory region of DNA. J. Bacteriol. 173:2969-2976. Romeo, J. M., and D. R. Zusman. 1992. Determinants of an unusually stable mRNA in the bacterium Myxococcus xanthus. Mol. Microbiol. 6:2975-2988. Rosenbluh, A., and M. Eisenbach. 1992. Effect of mechanical removal of pili on gliding motility of Myxococcus xanthus. J. Bacteriol. 17454064413. Schlieker, C., A. Mogk, and B. Bukau. 2004. A PDZ switch for a cellular stress response. Cell 117417-419. Sha, J., E. V. Kozlova, A. A. Fadl, J. I? Olano, C. W. Houston, J. W. Peterson, and A. K. Chopra. 2004. Molecular characterization of a glucose-inhibited division gene, gidA, that regulates cytotoxic enterotoxin of Aeromonas hydrophila. Infect. Imrnun. 72:1084-1095. Shimkets, L., and T. W. Seale. 1975. Fruiting-body formation and myxospore differentiation and germination in M x y o coccus xanthus viewed by scanning electron microscopy. J. Bacteriol. 121:711-720. Shimkets, L. J., and D. Kaiser. 1982a. Induction of coordinated cell movement in Myxococcus xanthus. J. Bacteriol. 152:451-461. Shimkets, L. J., and D. Kaiser. 1982b. Murein components rescue developmental sporulation of Myxococcus xanthus. J. Bacteriol. 152:462-470.
240 Shimkets, L. J. 1986a.Role of cell cohesion in Myxococcusxanthus fruiting body formation. J. Bacteriol. 1 6 6 9 4 2 4 4 8 . Shimkets, L. J. 1986b. Correlation of energy-dependent cell cohesion with social motility in Myxococcus xanthus. J. Bacteriol. 1665337-841. Skerker, J. M., and H. C. Berg. 2001. Direct observation of extension and retraction of type IV pili. Proc. Natl. Acad. Sci. USA 98:6901-6904. Sourjik, V., and R. Schmitt. 1996. Different roles of CheYl and CheY2 in the chemotaxis of Rhizobium meliloti. Mol. Microbiol. 22:427-436. Sourjik, V., and R. Schmitt. 1998. Phosphotransfer between CheA, CheY1, and CheY2 in the chemotaxis signal transduction chain of Rhizobium meliloti. Biochemistry 37: 232 7-23 35. Sun, H., D. R. Zusman, and W. Shi. 2000. Type IV pilus of Myxococcus xanthus is a motility apparatus controlled by the frz chemosensory system. Curr. Biol. 10:1143-1146. Thomasson, B., J. Link, A. G. Stassinopoulos, N. Burke, L. Plamann, and P. L. Hartzell. 2002. MglA, a small GTPase, interacts with a tyrosine kinase to control type IV pili-mediated motility and development of Myxococcus xanthus. Mol. Microbiol. 46: 1399-141 3. Vollmer, W., and J. V. Holtje. 2004. The architecture of the murein (peptidoglycan) in gram-negative bacteria: vertical scaffold or horizontal layer(s).J. Bacteriol. 1865978-5987. Wall, D., S. S. Wu, and D. Kaiser. 1998. Contact stimulation of Tgl and type IV pili in Myxococcus xanthus. J. Bacteriol. 180:759-76 1. Wall, D., and D. Kaiser. 1999. Type IV pili and cell motility. Mol. Microbiol. 32:l-10. Wall, D., P. E. Kolenbrander, and D. Kaiser. 1999. The Myxococcus xanthus pilQ (sglA) gene encodes a secretin homolog required for type IV pilus biogenesis, social motility, and development. J. Bacteriol. 181:24-33. Webre, D. J., P. M. Wolanin, and J. B. Stock. 2003. Bacterial chemotaxis. Cum Biol. 13:R47-R49. Weimer, R. M., C. Creighton, A. Stassinopoulos, P. Youderian, and P. L. Hartzell. 1998. A chaperone in the HSP70 family controls production of extracellular fibrils in Myxococcus xanthus. J. Bacteriol. 18053574368. White, D., M. Dworkin, and D. J. Tipper. 1968. Peptidoglycan of Myxococcus xanthus: structure and relation to morphogenesis. J. Bacteriol. 95:2186-2197. White, D. J., and P. L. Hartzell. 2000. AglU, a protein required for gliding motility and spore maturation of Myxococcus
STRUCTURE AND METABOLISM xanthus, is related to WD-repeat proteins. Mol. Microbiol. 36:662-678. White, D. J., R. Merod, B. Thomasson, and P. L Hartzell. 2001. GidA is an FAD-binding protein involved in development of Myxococcus xanthus. Mol. Microbiol. 42503517. Wu, S. S., and D. Kaiser. 1995 Genetic and functional evidence that Type IV pili are required for social gliding motility in Myxococcus xanthus. Mol. Microbiol. 18547-558. Wu, S. S., and D. Kaiser. 1996. Markerless deletions of pi1 genes in Myxococcus xanthus generated by counterselection with the Bacillus subtilis sacB gene. J. Bacteriol. 178: 58 17-5 82 1. Wu, S. S., J. Wu, and D. Kaiser. 1997. The Myxococcus xanthus pilT locus is required for social gliding motility although pili are still produced. Mol. Microbiol. 23:109-121. Wu, S. S., J. Wu, Y. L. Cheng, and D. Kaiser. 1998. The pilH gene encodes an ABC transporter homologue required for type IV pilus biogenesis and social gliding motility in Myxococcus xanthus. Mol. Microbiol. 29:1249-1261. Xu, D., C. Yang, and H. B. Kaplan. 1998. Myxococcus xanthus sasN encodes a regulator that prevents developmental gene expression during growth. J. Bacteriol. 180:6215-6223. Xu, Q., W. P. Black, S. M. Ward, and Z. Yang. 2005. Nitratedependent activation of the Dif signaling pathway of Myxococcus xanthus mediated by a NarX-DifA interspecies chimera. ]. Bacteriol. 187:6410-6418. Yang, C., and H. B. Kaplan. 1997. Myxococcus xanthus sass encodes a sensor histidine kinase required for early developmental gene expression. ]. Bacteriol. 179:7759-7767. Yang, Z., Y. Geng, and W. Shi. 1998a. A DnaK homolog in Myxococcus xanthus is involved in social motility and fruiting body formation.]. Bacteriol. 180:218-224. Yang, Z., Y. Geng, D. Xu, H. B. Kaplan, and W. Shi. 1998b. A new set of chemotaxis homologues is essential for Myxococcus xanthus social motility. Mol. Microbiol. 30:1123-1130. Yang, Z., X. Ma, L. Tong, H. B. Kaplan, L. J. Shimkets, and W. Shi. 2000. Myxococcus xanthus dif genes are required for biogenesis of cell surface fibrils essential for social gliding motility. J. Bacteriol. 18257934798. Yang, Z., and Z. Li. 2005. Demonstration of interactions among Myxococcus xanthus Dif chemotaxis-like proteins by the yeast two-hybrid system. Arch. Microbiol. 183:243-252. Youderian, P. A., and P. Hartzell. 2006. Transposon insertions of magellan-4 that impair social gliding motility in Myxococcus xanthus. Genetics 172:1397-1410.
Myxohacteria: Multicellularity and Differentiation Edited by David E. Whitworth 0 2008 ASM Press, Washington, D.C.
Patrick D. Curtis Lawrence J. Shimkets
Metabolic Pathways Relevant to Predation, Signaling, and Development
Genome analysis offers the ability to examine the central metabolism of organisms from a perspective that, while holistic in information content, lacks the accuracy of conclusions derived from sound biochemical and genetic analysis. Shadows of possibilities emerge that only silhouette necessary experimentation. Conversely, biochemical and genetic analyses are incomplete without the genomic knowledge. Here, previously determined biochemical and genetic data are analyzed through the scope of genomic analysis, with particular attention to catabolic and anabolic pathways involved in the basic biology of the myxobacteria. Genomic annotation was based upon the completed Myxococcus xanthus genome (Goldman et al., 2006). Metabolic pathways were examined by homology searching using the amino acid sequence for well-characterized enzymes. Metacyc (http://metacyc.org/) contains a compilation of most known pathways, enzymes, and genes and has a pathway analysis for 211. xanthus which was often used as a reference point (Caspi et al., 2006). A limitation of this analysis is its reliance on the annotation conducted by The Institute for Genomic Research (TIGR). To reduce problems with annotation errors, all pathways in the chapter were also annotated by hand
14
using BLASTp. Two considerations were used in assigning a function to a putative protein sequence, amino acid identity with known members of the EC class (>30%) and orthologous gene (COG) determination by TIGR. A strong hit satisfies both criteria. Gene and protein names adopted below are derived primarily from homologs found in Escherichia coli K-12 when applicable. This policy deviates from TIGR, which names the genes/ proteins after the closest named homolog in the database and does not specify the origin of the name. As with any homology-based search, the results should be considered “putative” until gene knockouts and enzyme assays confirm the identity and function of the homolog. The word putative is avoided for the sake of brevity but it should be understood that the proposed pathways represent little more than the best candidates. The first portion of this chapter, entitled “Catabolic Pathways,” deals with the catabolism of amino acids and lipids, as they are the principal carbon and energy sources derived from prey bacteria. The second portion of this chapter, entitled “Anabolic Pathways,” highlights the synthesis of lipids because of their unusual chemical structures in myxobacteria, and also the spore-specific components trehalose and ether lipids.
Patrick D. Curtis and Lawrence J. Shimkets, Department of Microbiology, University of Georgia, Athens, GA 30602.
241
STRUCTURE AND METABOLISM
242
CATABOLIC PATHWAYS Most myxobacteria, including M . xanthus, can catabolize prey microorganisms. M. xanthus utilizes amino acids and lipids as carbon and energy sources, incorporates purines and pyrimidines via salvage pathways, and fails to utilize sugars (Bretscher and Kaiser, 1978; Hemphill and Zahler, 1968a, 196810; Lau et al., 2002; Loebeck and Klein, 1956). The literature is extensive and is not reviewed here. Rather, genomic evidence for specific pathways involved in assimilation and catabolism is provided. Most of the research examining the sources of energy for the myxobacteria has focused on the assimilation of amino acids, not without reason (Bretscher and Kaiser, 1978; Dworkin, 1962). The average E. coli cell is composed of roughly 55% (dry weight) protein (Neidhardt and Umbarger, 1996), by far the largest component of the cell. Though lipids represent a much smaller fraction of the total cell mass (9% [Neidhardt and Umbarger, 1996]),their energetic content is considerable. The catabolism of serine to CO, yields approximately 11 ATP, while the catabolism of a 16:O fatty acid to CO, yields approximately 80 ATP. Therefore, both cellular fractions represent rich sources of energy.
Amino Acid Catabolism In most cases there is excellent agreement between the presence of a particular amino acid catabolic pathway and the ability of that amino acid to stimulate growth in defined and minimal media. The pathways are listed below according to amino acid, roughly in alphabetical order.
Alanine M . xanthus uses alanine dehydrogenase for catabolism of L-alanine (Fig. 1, reaction 1).L-Alanine can also be converted to D-alanine for use in peptidoglycan biosynthesis using alanine racemase (EC 5.1.1.1, Alr, MXAN7160). D-Alanine may be catabolized to pyruvate using the ironcontaining alcohol dehydrogenase MXAN5629. Arginine M. xanthus contains many pathways for catabolizing arginine, so it is surprising that arginine is not a major component in minimal and defined media (Bretscher and Kaiser, 1978). Collectively these pathways produce L-glutamate, putrescine (and hence other polyamines), L-proline, and succinate. The bulk of the arginine degradation in E. coli is carried out by the arginine succinyltransferase pathway, which is found in many Proteobacteria that use arginine as a sole carbon source. This pathway is not present
in M . xanthus, which may explain the lack of importance for arginine in minimal media. Several arginine catabolic pathways use arginase to hydrolyze L-arginine. Different organisms have variations of the arginase pathway depending on the fate of the catabolic products. The classical arginase pathway yields L-glutamate. There is weak homology for arginase, (26 to 31% amino acid identity), but beyond that the pathway is conserved in M. xanthus (Fig. 1, reaction 2). There is a branch in this pathway as ornithine cyclodeaminase (EC 4.3.1.12, MXAN7463) converts L-ornithine to L-proline. The arginine decarboxylase pathway produces putrescine as an intermediate for synthesis of other polyamines and succinate as an end product (Fig. 1,reaction 3).E. coli has two forms of arginine decarboxylase. Constitutively expressed SpeA is used for the biosynthesis of putrescine, while inducibly expressed AdiA is used for arginine catabolism. Interestingly, there is no homolog for AdiA but there is a strong homolog of SpeA in M . xanthus. All the other elements of the pathway to degrade arginine to succinate exist, suggesting that SpeA plays a dual role in the biosynthesis of putrescine and catabolism of arginine. M . xanthus contains two homologs of E. coli transaminase YgiG (MXAN3014 and MXAN7377). E. coli uses at least three different enzymes to catalyze the last reaction, Sad, AldA, and GabD. There are many homologs of each in M . xanthus, and though MXAN2844 is annotated as succinate semialdehyde dehydrogenase, there are other possibilities.
AsparagineIAspartate E. coli K-12 has three different L-asparaginases to deaminate asparagine, two of which are present in M . xanthus. E. coli aspartase (EC 4.3.1.1) removes the nitrogen from aspartate to produce fumarate, but there is no evidence for this enzyme in M. xanthus. Instead 111.xanthus appears to use a version of this pathway found in some gram-positive bacteria and mammals to produce phosphoenolpyruvate (Fig. 1, reaction 4).
Cysteine Two pathways are known for the catabolism of L-cysteine to pyruvate; at least one may exist in M . xanthus. In E. coli two desulfhydrases convert L-cysteine to pyruvate in a single step. Tryptophanase (TnaA, also used in tryptophan catabolism) does not appear to have a homolog in M . xanthus. MetC, P-cystathionase (also used in methionine synthesis), has four homologs in M . xanthus: MXAN2035, MXAN0969, MXAN0970, and MXAN1955 (Fig. 1, reaction 5). Of these, MXAN2035
14. PREDATION,SIGNALING,AND DEVELOPMENT (1)
+
L-alanine + NAD++ H,O
243
pyruvate + NADH + ammonia
EC 1.4.1.1 alanine dehydrogenase MXAN4146
(2)
L-arginine
+
L-ornithine EC 3.5.3.1 arginase MXAN4431
L-glutamatey-semialdehyde
EC 2.6.1.13 ornithine amino transferase MXAN7377
+
L-glutamate y-semialdehyde
pyrroline 5-carboxylate
Spontaneous
(3)
L-arginine
+
agmatine
EC 4.1.1.19 arginine decarboxylase MXAN2742
+
+
L-glutamate
EC 1.5.1.12 1-pyrroline-5-carhoxylatedehydrogenase MXAN5891
putrescine
+
4-amino-butyraldehyde
EC 3.5.3.11 EC 2.6.1.29 agmatinase putrescine transaminase MXAN4431 MXAN3014 or MXAN7377
4-amino-butyraldehyde
+
4-aminobutyrate
9 succinate semialdehyde
EC 1.2.1.19 EC 2.6.1.19 y-aminohutyraldehydedehydrogenase 4-aminobutyrate transaminase MXAN0921 MXAN3014
succinate semialdehyde
+
succinate
EC 1.2.1.16 succinate semialdehyde dehydrogenase MXAN2844
(4)
L-asparagine
+
L-aspartate + a-ketoglutarate
EC 3.5.1.1 L-asparaginase I MXAN5198 L-asparaginase I1 MXAN1160
Oxaloacetate + GTP
+
+
oxaloacetate + L-glutamate
EC 2.6.1.1 aspartate aminotransferase MXAN3386
phosphoenolpyruvate+ GDP + C 0 2
EC 4.1.1.32 phosphoenolpyrnvatecarboxykinase MXAN1264
(5)
L-cysteine + H,O
3 pyruvate + ammonia + H2S
EC 4.4.1.1 L-cysteine desulihydrase MXAN2035
Figure 1 Amino acid catabolic reactions 1 through 5 (alanine to cysteine). See the text.
has the most homology with the bacterial and yeast genes. The oxidation of cysteine to 3-sulfinoalanine and eventually to pyruvate is the major route of cysteine catabolism in mammals. The first step in this pathway involves cysteine dioxygenase (EC 1.13.11.20). There is weak evidence for a homolog in M . xantbus (MXAN4718). The remaining enzyme in the pathway, aspartate amino transferase (MXAN3386), is likely to be present (see catabolism of asparagine/aspartate).
Glutamine/Glutamate Two enzymes convert L-glutamine to L-glutamate: glutaminase and glutamate synthase. The biochemical properties of the two major E. coli glutaminases (EC 3.5.1.2) have been studied in detail, but the genes encoding them have not been identified. Two putative glutaminase genes have been described in E. coli, ybaS and yneH; however, neither has a homolog in M. xantbus. Nor is there a homolog of human glutaminase C. A number of amidotransferases, such as anthranilate synthetase, have
STRUCTUREAND METABOLISM
244 glutaminase activity and may contribute to glutamine catabolism. The biochemistry and genetics of glutamate synthase are known from a variety of bacteria. Glutamate synthase (EC 1.4.1.13) catalyzes the transamination of aketoglutarate to produce two glutamates. The Klebsiella aerogenes enzyme consists of a single 175-kDa protein, whereas the E. coli enzyme consists of 53- and 135-kDa subunits. M. xanthus has adjacent genes encoding two subunits like E. coli, MXAN3917 and MXAN3918. One of the major catabolic pathways for glutamate involves deamination to a-ketoglutarate by glutamate dehydrogenase. This is a key reaction in the catabolism of arginine, glutamine, histidine, and proline, all of which produce glutamate as an intermediate. Glutamate dehydrogenase catalyzes a reversible reaction that can be either anabolic or catabolic depending on the conditions and the organism. M . xanthus glutamate dehydrogenase (GdhA) is MXAN5873. Mammalian glutamate dehydrogenase uses both NAD’ and NADP+ as cofactors (EC 1.4.1.3). Most prokaryotic enzymes can use either NAD+ (EC 1.4.1.2) or NADP’ (EC 1.4.1.4). The TIGR annotation gives the M. xanthus enzyme as EC 1.4.1.3, suggesting that this enzyme may use both cofactors. The primary pathway for the use of L-glutamate as a carbon source in E. coli is transamination of oxaloacetate to form L-aspartate and a-ketoglutarate by AspC (Fig. 2, reaction 6). Aspartate aminotransferase (EC 2.6.1.1) resembling that of a Bacillus species is found in the M. xanthus genome, MXAN3386, and is dramatically different from the E. coli AspC. AspA (EC 4.3.1.1), which produces fumarate from L-aspartate by deamination, appears to be absent, suggesting that the aspartate is directed toward the synthesis of the aspartate family of amino acids or catabolized to oxaloacetate and phosphoenolpyruvate (see “Asparagine/Aspartate” above). Glutamate can also enter the tricarboxylic acid TCA cycle as succinate using the three-step glutamate decarboxylase pathway (Fig. 2, reaction 7) in which the final two steps are identical to those for putrescine degradation (see “Arginine” above). There is also evidence for a second bacterial pathway for catabolizing glutamate to succinate (Fig. 2, reaction 8).
Glycine Glycine plays a key role in C1 anabolism through the generation of the C1 donor Nlo-formyl-tetrahydrofolate. All elements of the pathway are present in M . xanthus with the exception of the anaerobic formate dehydrogenase (Fig. 2, reaction 9). These results suggest that the pathway is strictly anabolic unless another route exists for oxidation of formate. Four polypeptides are involved
in the glycine cleavage pathway, GcvHPT and LpdA, with the first three genes forming an operon in both E. coli and M . xanthus.
Histidine M . xanthus may catabolize L-histidine to L-glutamate (Fig. 3, reaction 10).The first three enzymes in the pathway have strong M. xanthus homologs, but here the trail runs dry. Bacillus subtilis uses EC 3.5.3.8 for the last step, but evidence for this enzyme in M. xanthus is lacking. Pseudomonas uses two enzymes for the last step, EC 3.5.3.13 and EC 3.5.1.68, to remove ammonia and formate sequentially. The first of these has a homolog in the M. xanthus genome (formiminoglutamate deiminase, MXANlOlO), but a homolog of the formate hydrolase is not apparent.
IsoleucinelLeucineNaline The branched-chain amino acids are unique in that they are essential for growth in all M. xanthus isolates that have been examined and are also essential for secondary metabolite production in myxobacteria and other organisms (Bretscher and Kaiser, 1978). Catabolism of these amino acids appears to be directed primarily at their use as primers for the synthesis of branchedchain fatty acids (see “Lipid Biogenesis” below). The key enzyme in this process is the branched-chain keto acid dehydrogenase (BCKAD) complex, also known as Esg in M. xanthus, which catalyzes the oxidative decarboxylation of the branched-chain a-keto acids derived from leucine, isoleucine, and valine (Fig. 3, reaction 11). In mammals the complex consists of 12 branched-chain a-ketoacid dehydrogenase ( E l )subunits and 6 dihydrolipoyl dehydrogenase (E3) subunits noncovalently associated with a core of 24 dihydrolipoyl transacylase (E2) components. A BCKAD kinase inactivates the complex by phosphorylation of alpha subunits of the heterotetrameric (a2p2) E l component; BCKAD phosphatase removes phosphates to activate the complex. The complex is present in M. xanthus, though there is no evidence for regulation by covalent modification.
Lysine At least nine pathways are known for the catabolism of lysine that vary in the initial products. The genes are known for only three of these pathways. In E. coli, lysine is decarboxylated to cadaverine by IdcC or CadA, neither of which is present in M . xanthus. In plants and animals, lysine is oxidized to saccharopine by using a unique dehydrogenase that is not present in M. xanthus. In fungi, lysine is degraded to glutarate following acetylation of the six-amino group with a unique lysine Wacetyltransferase,
14. PREDATION,SIGNALING,AND DEVELOPMENT (6)
L-glutamine + a-ketoglutarate
+
L-glutamate
+ oxaloacetate
EC 1.4.1.13 glutamate synthase MXAN3917, large subunit MXAN3918. small sununit
(7)
L-glutamate
(8)
L-glutamate
+
succinyl-CoA 3
L-aspartate + a-ketoglutarate
+
succinate semialdehyde
EC 2.6.1.19 4-aminobutyrateaminotransferase MXAN3014
a-ketoglutarate 3
EC 2.6.1.1 aspartate aminotransferase MXAN3386
3
EC 2.6.1.1 aspartate aminotransferase MXAN3386
3 4-aminobutyrate
EC 4.1.1.15 glutamate decarboxylase MXAN6783
245
+
succinate
EC 1.2.1.16 succinate semialdehyde dehydrogenase MXAN2844
S-succinyl-dihyrolipoamide 3 succinyl-Cod
EC 1.2.42 2-oxoglutarate dehydrogenase MXAN6035, E l component
EC 2.3.1.61 2-oxoglutarate dehydrogenase MXAN6036, E2 component
succinate
EC 6.2.1.5 succinyl-CoAsynthase MXAN3542, a subunit MXAN3541, p subunit
(9)
Glycine + tetrahydrofolate 3 5,lO-methylene-tetrahydrofolate EC 1.4.4.2 and EC 2.1.2.10 glycine dehydrogenase,MXAN3042 tetrahydrofolateaminomethyltransferase MXAN3041, MXAN3040, MXAN6341
5,lO-methylene-tetrahydrofolate 3
5,lO-methenyl-tetrahydrofolate
EC 1.5.1.15 methylenetetrahydrofoiate dehydrogeanse MXAN1095
5,lO-methenyl-tetrahydrofolate
+
N1o-formyl-tetrahydrofolate
EC35.4.9 methenyl tetrahydrofolatecyclobydrolase MXAN2226
N1o-formyl-tetrahydrofolate+ phosphate + ADP
+
formate + tetrahydrofolate + ATP
EC 6.3.4.3 formate tetrahydroformateligase MXAN0175
Figure 2 Amino acid catabolic reactions 6 through 9 (glutamine to glycine). See the text.
which is also missing in M. xanthus. Therefore, it is not clear whether lysine is catabolized.
Methionine The methionine catabolic pathway is similar to that observed in mammals and involves the synthesis of Sadenosyl-L-methionine for transmethylation reactions and then hydrolysis of adenosine to produce homocysteine (Fig. 3 , reaction 12).Homocysteine forms a branch point in the pathway and may either be recycled to Lmethionine, using the vitamin B,,-dependent methionine synthase (not shown), or degraded to succinate.
Phenylalanine/Tyrosine Only a single aerobic phenylalanine/tyrosine catabolic pathway is known. In this pathway phenylalanine is catabolized to tyrosine by phenylalanine 4-hydroxylase (EC 1.14.16.1, MXAN5127) and eventually to succinate. While homologs for genes involved in some later steps are found in M . xanthus, no homolog for the second step in the catabolic pathway (EC 2.6.1.5, tyrosine aminotransferase) is found. In M. xanthus, 14C-labeled phenylalanine is converted to tyrosine and the excess tyrosine is secreted into the growth medium (Hemphill and Zahler, 1968a), suggesting that this pathway is,
STRUCTURE AND METABOLISM
246 (10)
L-histidine -9
-9
urocanate
EC 4.3.1.3 histidine ammonia lyase MXAN3465
4-imidazolone-5-propionate
EC 4.2.1.49 urocanase MXAN4343
4-imidazolone-5-propionate -9
N-formimine-L-glutamate -9
EC 3.5.2.7 imidazolone-5-propionatase MXAN4345
(1 1)
L-isoleucine L-leucine L-valine
2-keto-3-methyl-valerate 2-keto-4-methyl-pentanoate 2-keto isovalerate
+
S-adenosyl-L-methionine -9
L-methionine -9
EC 2.5.1.6 methionine adenosyltransferase MXAN6517
S-adenosyl-homocysteine -9
homocysteine
cystathionine -9
EC 43.1.22 cystathionine p-synthase MXAN2041
propionyl CoA -9
S-adenosyl-homocysteine
EC 2.1.1.73 DNA methyltransferase MXAN3598
EC 3.3.1.1 S-adenosylhomocysteinehydrolase MXAN6516
Homocysteine -9
2-methylbutyr yl-CoA Isovaleryl-CoA Isobutyryl-CoA
-9
BCKAD Ela, MXAN4564 BCKAD Elp, MXAN4565 BCKAD E2, MXAN4217 BCKAD E3, MXAN4219
EC 2.6.2.42 branched chain amino acid aminotransferase MXAN2987
(12)
L-glutamate
??
-9
L-methionine
EC 2.1.1.13 methionhe synthase MXAN1971
2-oxobutanoate -9
EC 4.4.1.1 cystathionine y-lyase MXAN3917
propionyl CoA
no EC number or gene
(S)-methyl-malonyl-CoA -9 (R)-methyl-malonyl-CoA
EC 6.4.1.3 propionyl-CoA carboxylase MXAN1111, a subunit MXAN1113, p subunit
(R)-methyI-malonyl-CoA -9
EC 5.1.99.1 no gene associated with activity
succinyl-CoA
EC 5.4.99.2 methylmalonyl-CoAmntase MXAN2263, a subunit MXAN2264, p subunit
Figure 3 Amino acid catabolic reactions 10 through 12 (histidine to methionine). See the text.
indeed, blocked at the first step of tyrosine catabolism (Fig. 4, reaction 13). Proline There is a single pathway for degradation of proline that involves the sequential action of proline dehydrogenase and 1-pyrroline-5-carboxylate dehydrogenase to produce glutamate (Fig. 4,reaction 14). In bacteria the two enzyme domains are usually encoded by a single gene, whereas in eukaryotes they are separate genes. M. xanthus appears to have separate and unlinked genes.
Serine L-Serine is deaminated to pyruvate and ammonia in E. coli by three homologous serine deaminases, but only one is found in M. xanthus (Fig. 4, reaction 15). Threonine L-Threonine can be converted to many metabolites by pathways whose biochemistry has far surpassed the genetics. All the known pathways begin with either deamination to 2-oxobutanoate or oxidation to 2amino-3-oxobutanoate. In M. xanthus, the former reaction appears to be present while the latter reaction is
14. PREDATION,SIGNALING,AND DEVELOPMENT (13)
+
L-phenylalanine
247
L-tyrosine
EC 1.14.16.1 phenylalanine 4-hydroxylase MXAN5127
(14)
L-proliie
3
l-pyrroline 5-carboxylate
EC 15.99.8 proline dehydrogenase MXAN7405
(15)
L-serine
+
+
L-glutamate
EC 1.5.1.12 l-pyrroline-5-carboxylate dehydrogenase MXAN5891
pyruvate + ammonia
EC 4.3.1.17 serine deaminase MXAN6186
(16)
L-threonine
+
2-oxobutanoate + cysteine + ammonia 3
EC 4.3.1.19 threonine dehydratase MXAN5874 threonine deaminase MXAN6186
cystathionine
EC 4.4.1.1 cystathionine y-lyase MXAN2035
Figure 4 Amino acid catabolic reactions 13 through 16 (phenylalanine to threonine). See the text.
doubtful (Fig. 4, reaction 16). Threonine is then catabolized to cystathionine, an intermediate in methionine catabolism that can be used to generate succinate, methionine, or pyruvate (see “Methionine” above).
Tryptophan There are seven known pathways for the catabolism of tryptophan. The biochemistry has been examined more extensively than the genetics due to the fact that various products of tryptophan are plant hormones, dyes, or putrid by-products during cheese production. The simplest pathway is the conversion of tryptophan to pyruvate and indole in a single step by tryptophanase. This pathway is intriguing, as indole induces spore formation in Stigmatella and may be a cell-cell signal (Stamm et al., 2005). Neither a homolog of this enzyme nor any enzymes in the other pathways were detected in 111. xanthus. Additionally, no tryptophanase homolog could be found in the Stzgmatella aurantiaca genome. Some parts of a eukaryotic catabolic pathway are found, but several enzymes in the middle of the pathway are missing. In summary, genome evidence suggests that M. xanthus is missing pathways for catabolism of seven amino acids: leucine, isoleucine, valine, phenylalanine, tyrosine, tryptophan, and lysine. The absence of catabolic pathways for leucine, isoleucine, and valine is not surprising given that M. xanthus is auxotrophic for these amino acids (Bretscher and Kaiser, 1978) and also uses them in fatty acid biosynthesis (see “Fatty Acid Primer Synthesis”
below). Catabolism of these amino acids would reduce branched-chain fatty acid synthesis, which appears to be necessary for development (Toal et al., 1995). The absence of phenylalanine/tyrosine, tryptophan, and lysine catabolic pathways is a mystery. It is interesting that of the mixture of amino acids that comprise the A signal (an early developmental signal), leucine, isoleucine, phenylalanine, tyrosine, and tryptophan account for 63% of the A signal activity (Kuspa et al., 1992).
Purine and Pyrimidine Salvage A variety of labeling and analog toxicity studies indicate that purines and pyrimidines are efficiently salvaged in M. xanthus, and these studies are not reviewed here (Hemphill and Zahler, 1968a; Tsai and Westby, 1978). M. xanthus has a suite of purine and pyrimidine salvage pathways comparable to what is found in E. coli with several options for converting nucleobases and nucleosides into nucleotides. The preferred method of nucleoside and deoxynucleoside salvage in E. coli is to first remove the (deoxy)ribose moiety from the (deoxy)nucleoside and then use a phosphoribosyltransferase to create the nucleotide monophosphate. E. coli can also transport the (deoxy)nucleosides and then phosphorylate them. Adenosine and deoxyadenosine are salvaged through deamination to inosine/deoxyinosine (EC 3.5.4.4, Add, MXAN1519) and hydrolysis to hypoxanthine (purine nucleoside phosphorylase, DeoD, MXAN2306). Guanosine and deoxyguanosine are hydrolyzed to guanine
248 (purine nucleoside phosphorylase, DeoD, MXAN2306). Phosphoribosyltransferases were found for adenine (EC 2.4.2.7, Apt, MXAN5352) and uracil (EC 2.4.2.9, Upp, MXANO124). E. coli has two additional phosphoribosyltransferases, both of which function on hypoxanthine and guanine. The M. xanthus homolog appears to be more closely related to one with a bias for hypoxanthine (EC 2.4.2.-, Hyp, MXAN5070), but it should be assumed from labeling studies that M. xanthus can utilize guanine as well. Uridine and cytidine are phosphorylated to UMP/ CMP by uridine kinase (EC2.7.1.48, Udk, MXAN4159). Thymidine and thymine nucleotides are deoxy compounds with no ribonucleotide counterparts; thymidine is phosphorylated to TMP by thymidine kinase (EC 2.7.1.21, Tdk, MXAN5072). Deoxycytidine is deaminated to deoxyuridine and either degraded to uracil and/or phosphorylated to deoxyUMP by thymidine kinase (EC 2.7.1.21, Tdk, MXAN5072) and then converted to TMP by thymidylate synthase (EC 2.1.1.45, ThyA, MXAN.5942). It should be noted from the genomic studies here and from previous labeling studies (Hemphill and Zahler, 1968a) that the main fate of exogenous purinedpyrimidines is incorporation into nucleic acids and not degradation for energy generation.
Lipid Catabolism Lipid oxidation has been demonstrated by 14C-labeling experiments in Myxococcus virescens (Loebeck and Klein, 1956) and methyloleate feeding in M. xanthus (Lau et al., 2002). The principal, but not sole, source of lipids in prey bacteria is the phospholipids, which may be hydrolyzed by four classes of lipases in M. xanthus. Phospholipase D (MXAN6753) removes the head group, leaving phosphatidic acid at the sn-3 position on the glycerol backbone. Fatty acids are located at the sn-1 and sn-2 positions in phospholipids. Three classes of lipases generate glycerol and fatty acids (MoraledaMuiioz and Shimkets, 2007). Both are robust carbon sources, although glycerol utilization by 111. xanthus has not been reported. Interestingly, most lipase homologs identified here are not predicted to have sn positionspecific activity, perhaps indicating their importance for catabolism over lipid signaling, Lipases Patatins were originally identified as the major potato storage protein, but biochemical characterization revealed fatty esterase activity, usually on compounds containing a single fatty acyl chain. MXAN3852 contains the conserved active site, the oxyanion hole,
STRUCTURE AND METABOLISM and other features that suggest it may be catalytically functional. Three other patatin homologs are present in M. xanthus but may be missing motifs necessary for catalytic function. MXAN3852 is expressed only during starvation, and deletion results in no phenotype with regards to aggregation and sporulation on TPM agar (Tris-phosphate-magnesium starvation agar; MoraledaMuiioz and Shimkets, 2007). On clone fruiting (CF) agar there is a 24-h delay to both aggregation and sporulation, and the fruiting bodies are unusually large and amorphous. The a / p hydrolases contain an eight-stranded p sheet (stabilized by intervening a helices) formed of two antiparallel p strands followed by six parallel p strands (Ollis et al., 1992). Many a/p hydrolases are proteases, but at least two putative a / p hydrolases in M . xanthus are likely lipases. MXAN5522 (a triacyl glycerol lipase acting on all three sn positions) is located directly upstream of a lipase chaperone homolog (MXAN5523). Lipases often have chaperones to prevent catalytic activity until they are secreted. This lipase is expressed during both vegetative growth and development (MoraledaMufioz and Shimkets, 2007). Interestingly, it shows a sharp spike of expression 24 h into development. Deletion of MXAN5522 results in threefold-increased spore yield on both TPM and CF. While there is no defect in aggregation on TPM, this strain aggregates 24 h faster than the wild type on CF. A possible explanation for the increased rate of sporulation in this strain may be that it is unable to utilize storage lipids for energy, thereby starving faster than normal. MXAN4638 is a putative lysophospholipase, which removes the fatty acid from lysophospholipids. Interestingly, disruption of the gene upstream and operonic with MXAN4638 generates an A-motility defect (Youderian et al., 2003). M. xanthus contains two GDSL lipase homologs (MXAN.5500 and MXAN4569), which are general bacterial lipases that liberate fatty acids from either sn position in phospholipids. The C-terminal domain is a @-barrelporin which forms a pore in the outer membrane through which the N-terminal catalytic domain is secreted and ultimately anchors the lipase facing away from the membrane. Both M. xanthus homologs lack the C-terminal anchoring domain. There is one report of a GDSL lipase anchored to the membrane by an Nterminal acylation (Klingsbichel, 1996), and though MXAN4569 appears to have an acylation signal, MXANSSOO does not. The lack of an anchoring domain may be an indication that MXAN5.500 diffuses away from the cell. Deletion of MXAN4569 has no effect on aggregation or sporulation on TPM (Moraleda-Muiioz and Shimkets, 2007). On CF, aggregation shows a mild
14. PREDATION,SIGNALING,AND DEVELOPMENT fruiting body morphology defect and sporulation proceeds approximately 24 h faster than the wild type. The lipases from MXAN3852, MXAN5522, and MXAN4569 were all tested in vitro against various tagged lipid derivatives to elucidate fatty acyl chain specificity. All lipases showed preferences for short acyl chains and had the highest activity for two carbon chain lengths.
p Oxidation Fatty acids are usually degraded by P oxidation, where two carbon acetate units are sequentially removed from the carboxyl end of the fatty acid, also known as the A terminus, as opposed to the methyl end or w terminus (for a review, see Clark and Cronan, 1996). This process resembles fatty acid elongation in reverse. FadL (MXAN7040) translocates the free fatty acid across the outer membrane, which is then translocated across the inner membrane and esterified to a coenzyme A (CoA) moiety by FadD (Weimar et al., 2002). M. xanthus contains as many as 10 FadD homologs (EC 6.2.1.3; the most likely homologs are MXAN1573, MXAN7148, MXAN0216, and MXAN0225). Next, a trans double bond is introduced two carbons from the A terminus (A2)by FadE. While the M. xanthus FadE (EC 1.3.99.3) homologs are not clear, MXAN3795 and MXAN3797 are conspicuous possibilities as they appear to be in an operon with MXAN3791, an AtoB homolog. Also interesting is MXAN6989, which encodes a peptide with strong homology to the C terminus of FadE and appears to be in an operon with FadA and FadB homologs. Next, water is substituted into the double bond to create a P-hydroxy fatty acyl-CoA which is oxidized to a p-keto group. Both these reactions are performed by the multifunctional fatty acid oxidation complex P subunit FadB. Additionally, FadB epimerizes D-P-hydroxy fatty acyl CoA to L-P-hydroxy fatty acyl-CoA for further processing. M. xanthus contains two FadB homologs (EC 1.1.1.35, MXAN5371 and MXAN6987). Upon generation of the p-keto fatty acyl-CoA, the fatty acid oxidation complex 01 subunit (FadA) cleaves the chain at the P-keto group by adding a CoA moiety, creating acetylCoA and the fatty acyl-CoA truncated by two carbons. M. xanthus contains two FadA homologs (EC 2.3.1.16), MXAN5372 and MXAN6988, that are located next to FadB homologs. This P oxidation cycle continues until there are only four carbons left on the fatty acid chain when the chain is hydrolyzed to two acetyl-CoA units by AtoB (EC 2.3.1.9, MXAN3791 and MXAN5135). For fatty acids with unsaturations at odd numbers of carbons from the carboxyl terminus, FadB isomerizes the
249
cis-A3double bond to trans-A2.Fatty acids with unsaturations at even numbers of carbons from the carboxyl terminus are reduced by 2,4-dienoyl-CoA reductase (EC 1.3.1.34, FadH, MXAN3389). This enzyme reduces the second double bond of the trans-A2,&-A4 intermediate to trans-A2 fatty acyl-CoA, which is further oxidized by the FadE and the unsaturated fatty acid degradation pathway (Hubbard et al., 2003). Like E. coli, M. xanthus contains two sets of both major P oxidation pathway enzymes, FadA and FadB. In E. coli, one set functions under aerobic conditions while the other set works under anaerobic conditions (Campbell et al., 2003). While M . xanthus is a strict aerobe, it is possible that the cells may encounter periods where energy generation is needed under oxygen-limiting conditions, such as the interior of a fruiting body, where free fatty acids may be a plentiful energy source. Expression studies would help determine if and when each set of P oxidation enzymes is expressed. a Oxidation 01 oxidation removes a single carbon from the fatty acid (Caspi et al., 2006) and has been demonstrated in S. aurantiaca (Dickschat et al., 2005). Molecular oxygen is added to a free fatty acid to create P-hydroperoxyfatty acid by the fatty acid 01 dioxygenase (EC 1.11.13 .-, MXAN5217). This molecule spontaneously degrades, though it can be facilitated by the dioxygenase (Hamberg et al., 2005), to either a @-hydroxy-fattyacid, or a fatty aldehyde with a loss of CO, and H,O. The fatty aldehyde dehydrogenase (EC 1.2.1.3, MXAN6986) oxidizes the aldehyde to the acid and in the process reduces NAD to NADH. a oxidation only generates one NADH as opposed to the several generated as a result of P oxidation. Despite the two homologs, 01 oxidation was not observed in radiolabeling studies with M. xanthus (Bode et al., 2005). In peas, a oxidation is observed only during seed germination, indicating a situational role for this pathway (Saffert et al., 2000). By analogy, perhaps 01 oxidation is important during M . xanthus spore germination. w Oxidation
During w oxidation the last carbon in a chain is converted to a carboxyl group, creating fatty acids from alkanes and dicarboxylates from fatty acids. M. xanthus does not appear to have a homolog of the w fatty acid oxidase.
Carbohydrate Utilization M . xanthus and most myxobacteria (with the exception of Sorangiurn and Byssophaga species) cannot grow
250 on carbohydrates (Bretscher and Kaiser, 1978; Watson and Dworkin, 1968). Prey bacteria are not particularly enriched in carbohydrates (E. coli contains 2.5% glycogen [Neidhardt and Umbarger, 1996]),yet carbohydrates are energy rich and are available in the rhizosphere. The ability of M . xanthus to utilize carbohydrates is examined in two parts: carbohydrate assimilation by the phosphotransferase system and carbohydrate metabolism by glycolysis. Phosphotransferase System While sugars can be internalized through each of the major types of transport systems, the phosphotransferase system (PTS) is specific to carbohydrates and should be an indicator of carbohydrate utilization. The PTS pathway begins with the transfer of a phosphoryl group from phosphoenolpyruvate (PEP) to Enzyme I (EI).This phosphoryl group is then transferred to HPr, which then transfers the phosphoryl group, sometimes through another carrier protein, to the permease composed of IIA, IIB, IIC, and sometimes IID subunits, which phosphorylate the transported carbohydrate. M . xanthus has homologs of all the PTS components neatly located in an operon (EI, MXAN6530; HPr, MXAN6.531; and IID-A, MXAN6532-5). In E. coli, EI and HPr are common components of all PTS and substrate specificity is conferred by the sugar-specificI1 proteins. Unlike E. coli, which has over a dozen sets of I1 proteins, M. xanthus has only one, but it is homologous with the E. coli system that has the broadest substrate range. The M. xanthus system is a Class 4 PTS (for a review see Postma et al., 1993), due to the presence of a IID homolog, most closely resembling the mannose PTS in E. coli. The M . xanthus system has separate IIA and IIB subunits like Klebsiella pneumoniae and B. subtilis (Postma et al., 1993).The M . xanthus IIA and IIB homologs have the conserved His10 and His175 residues involved in the phosphorelay (Erni et al., 1987). Therefore, at this level of analysis M . xanthus has all the necessary components for a functional PTS system. The most homologous system is the E. coli mannose PTS system, which transports eight different sugars: mannose, N-acetylglucosamine, glucosamine, fructose, 2-deoxyglucose,glucose, trehalose,and methyl ct-glucoside (Postma et al., 1996). Many of these sugars have been examined as growth substrates in defined and minimal media for M . xanthus without success (Bretscher and Kaiser, 1978). N-acetylglucosamine has been used in peptidoglycan labeling studies with poor incorporation (L. Shimkets, unpublished data). As PTS is dependent on the availability of PEP, perhaps the cellular level of PEP is in short supply under the conditions examined in transport
STRUCTUREAND METABOLISM studies. Another possibility is that the PTS system may be expressed only under specific conditions. As the system is predicted to transport trehalose, a possible link with germination exists as the spore-specific sugar trehalose is first secreted and then disappears (see “Trehalose Biosynthesis” below). Glycolysis Monosaccharides are used for exopolysaccharide, peptidoglycan, and lipopolysaccharide biosynthesis. Additionally, M . xanthus produces glycogen (a homopolysaccharide of glucose monomers joined with 0l-1~4linkages and ct-1,6 branches) during early to middle stationary phase (Nariya and Inouye, 2003). Glycogen is a common carbon and energy storage polymer in many bacteria. Monosaccharides are produced by gluconeogenesis, which uses many of the glycolysis enzymes in reverse. Glycolysis has three kinase reactions not involved in gluconeogenesis: those performed by glucokinase, phosphofructokinase, and pyruvate kinase. For the first step in glycolysis, glucose must be phosphorylated to glucosel-phosphate. Extracellular glucose can be phosphorylated by the PTS system (see above), though activity has not been demonstrated in M . xanthus. While intracellular glucose is phosphorylated by glucokinase, there are no recognizable glucokinase homologs in the genome, and glucokinase activity was not observed in M . xanthus (Watson and Dworkin, 1968). Glycogen is consumed in E. coli by removing glucose monomers from the nonreducing end of the chain as glucose-l-phosphate by glycogen phosphorylase GlgP (Alonso-Casajus et al., 2006). N o glycogen phosphorylase homolog is found in the genome. The second unique kinase step, that performed by phosphofructokinase, has been carefully studied (Nariya and Inouye, 2002). M . xanthus has a phosphofructokinase (EC 2.7.1.11, Pfk, MXAN6373). The activity of Pfk is modulated by phosphorylation of Thr-226 by the protein serinekhreonine kinase Pkn4 (Nariya and Inouye, 2002). Additionally, the kinase activity of Pkn4 is controlled by a regulatory protein MkapB and may be involved in a larger signaling network (Nariya and Inouye, 2005). Deletion of phosphofructokinase leads to elevated glycogen levels and decreased spore production (Nariya and Inouye, 2003). Spore production can be restored by supplying the mutant with exogenous pyruvate, which is then presumably catabolized by the tricarboxylic acid cycle. The third unique kinase step is missing. Although M . xanthus contains two pyruvate kinase homologs (EC 2.7.1.40, Pyk, MXAN3514 and MXAN6299), enzymatic activity was not found in cell or spore extracts
14. PREDATION,SIGNALING,AND DEVELOPMENT (Watson and Dworkin, 1968). Pyruvate kinase is necessary for development in S. aurantiaca and binds indole, a known sporulation inducer in that organism (Stamm et al., 2005). Glycogen produced during stationary phase is consumed prior to sporulation, consumption of glycogen is necessary for efficient sporulation, and consumption of glycogen is linked to the regulated activity of phosphofructokinase (Nariya and Inouye, 2003). These results indicate that glycolysis occurs during development. However, the source of phosphorylated glucose for glycolysis is unclear as no glycogen phosphorylase homolog is found in the genome, and the later and critical glycolysis reaction perfomed by pyruvate kinase has not been detected. Additionally, during development monosaccharides are needed for trehalose (see “Trehalose Biosynthesis” below) and exopolysaccharide biosynthesis, suggesting competing destinations for glucose in the cell. One hypothesis is that the glycolytic pathway functions incompletely during development. The reaction following phosphofructokinase cleaves fructose-1, 6-bisphosphate into glyceraldehyde-3-phosphate and dihydroxyacetone phosphate. Dihydroxyacetone phosphate may be used in ether lipid biosynthesis (see “Ether Lipid Biosynthesis” below), compounds found only in M. xanthus during development. This unconventional use of glycolysis may explain the unusual method of enzymatic regulation of phosphofructokinase. Further testing, specifically during development, is required to determine if M. xanthus uses the full glycolysis pathway.
ANABOLIC PATHWAYS Lacking space in this chapter to focus on all aspects of anabolism, we chose to focus on unusual cell envelope molecules and spore-specific products. Over the years the structural components of the myxobacterial cell envelope have been examined. Lipids are by far the most structurally divergent group of macromolecules relative to other Proteobacteria and may perform roles in cell signaling (Curtis et al., 2006; Downard and Toal, 1995; Kearns et al., 2001). In the following sections we discuss phospholipids and ceramides. Steroids and polyketides are discussed in chapter 15. Additionally the synthesis and fate of the spore-specific carbohydrate trehalose are examined.
Lipid Biogenesis While phospholipids perform a basic structural role, the fatty acid diversity of myxobacteria is extraordinary and far exceeds that necessary for structural purposes. While most Proteobacteria have 3 to 5 different fatty acids (Cronan and Rock, 1996), vegetative M. xanthus
251 cells contain at least 18 different fatty acids (Curtis et al., 2006; Kearns et al., 2001). Within M. xanthus phospholipids, fatty acids can have straight chains or branched chains, and either type can be saturated or unsaturated. The principal phospholipid in 211. xanthus, phosphatidylethanolamine (PE), deviates markedly from PE species of other Proteobacteria in that it contains unsaturated fatty acids at the sn-1 position, which appear to have a role in cell signaling (Curtis et al., 2006). In this section, phospholipid (and ceramide) biosynthesis is examined with attention to sources of structural diversity.
Fatty Acid Primer Synthesis The biosynthesis of fatty acids begins with the generation of a primer that is extended in a series of repetitive cycles to form the fatty acid. For straight-chain fatty acids with even numbers of carbons the primer is acetyl-CoA. The branched-chain fatty acids are the most abundant fatty acids in M. xanthus. The primers for branched-chain fatty acids are derived from the branched-chain amino acids (see “Isoleucine/Leucine/Valine” in “Amino Acid Catabolism” above for detail). Fatty acids derived from leucine constitute the iso-odd fatty acid family, those derived from isoleucine constitute the anteiso-odd family, and those derived from valine constitute the iso-even family. In the first step of branched-chain primer synthesis, the amine group is removed, producing the aketoacid branched-chain derivative (EC 2.6.1.42, IlvE, MXAN2987). The a-ketoacids are decarboxylated and attached to CoA by the BCKAD complex to form the primer. Myxobacterial BCKAD enzymes may have more pronounced substrate specificities than usual. Deuterium-labeled leucine was incorporated into fatty acids of both M. xanthus and S. aurantiaca, whereas little labeled valine was incorporated (Bode et al., 2005); concordantly, the most abundant branched-chain fatty acids in these two organisms are the iso-odd fatty acids (Dickschat et al., 2005; Kearns et al., 2001; Ware and Dworkin, 1973). In Streptomyces, incorporation of valine into iso-even fatty acids was equal to that of leucine into isoodd fatty acids (Cropp et al., 2000). M . xanthus transposon insertions in the genes encoding Elci and E1P subunits (esg locus) block development within the first 5 h (Downard et al., 1993).These mutants show decreased amounts of branched-chain fatty acids and increased levels of the fatty acid 16:105c (Curtis et al., 2006; Kearns et al., 2001). It is possible that branched-chain fatty acids are necessary for the generation of a chemical signal required for development or for extracellular matrix production (Kim et al., 1999),which is essential for fruiting body formation (Behmlander and Dworkin, 1991; Lu et al., 2005; Yang et al., 2000).
STRUCTURE AND METABOLISM
252
52
ACP-S-C-CHa CH -(CH2),-CH3
FabUA H20<
B
?H ACP-S-C-CH2-CH -(CH2)x-CH3
Figure 5 Fatty acid biosynthesis is a cyclic process. Condensation of malonyl-ACP with the growing chain by FabB, FabF, or FabH results in chain extension by two carbons. The p-keto group is reduced to a P-hydroxyl group by FabG, which is then dehydrated to a trans-A2 unsaturated bond by FabZ or FabA. This bond is reduced to full saturation by FabI or isomerized to &-A3 and preserved in later chain extension to create unsaturated fatty acids.
Both M. xantbus and S. aurantiaca have a second, novel system for branched-chain primer synthesis. Acetate is incorporated into isovaleryl-CoA, the primer for iso-odd fatty acids, using a shunt from the mevalonate pathway for isoprenoid biosynthesis (Bode et al., 2006a; Mahmud et al., 2002). In the mevalonate pathway, three acetate units are used to construct HMG-CoA, which is reduced to mevalonic acid. In the proposed shunt, HMG-CoA is decarboxylated and dehydrated to 3-methylbut-3-enol-CoA, which is then isomerized to 3,3-dimethylacrylyl-CoA and finally reduced to isovaleryl-CoA. HMG-CoA decarboxylase and dehydratase activities have been demonstrated in cell extracts of the S. aurantiaca BCKAD mutant (Mahmud et al., 2005), and labeled 3,3-dimethylacrylate was shown to be incorporated into the branched-chain fatty acids of both S. aurantiaca and M. xanthus BCKADdeficient strains (Mahmud et al., 2002). No evidence has been found that this pathway functions in the wild type under vegetative conditions. However, this pathway does function during fruiting body formation, presumably to supply isovaleryl-CoA when leucine is limited (Bode et al., 2006b).
Fatty Acid Elongation Fatty acid elongation is a repetitive process (Fig. 5 ) that extends the fatty acid two carbons per cycle (Cronan and Rock, 1996).The initial step is the condensation of the primer, or the primer extended by previous cycles, with malonyl-ACP. The condensation step can be performed by ketoacyl-ACP synthase I (KAS I, FabB), KAS I1 (FabF), or KAS I11 (FabH). In the process, the terminal carboxylic acid of malonyl-ACP leaves, resulting in a
p-ketoacyl-ACP. FabB is necessary for initiating extension of growing unsaturated fatty acid chains but can also extend saturated chains. FabF is required for thermal regulation of fatty acid saturation. FabH performs the first condensation of the primer with malonyl-ACP. In the next step the p-carboxyl group is reduced to a hydroxyl group with FabG. Then, the molecule is dehydrated with FabA or FabZ, resulting in a trans-A2double bond. The sole product of FabZ-mediated dehydration is the trans-A2 compound. While FabA generates transA2 double bonds as well, it can also isomerize the double bond to &-A3, which is preserved in later cycles by FabB to create unsaturated fatty acids. FabA, while active on saturated chains, is inactive on unsaturated chains. Therefore, while FabA initiates unsaturated fatty acid biosynthesis by creating the cis double bond, it is FabZ that extends unsaturated fatty acids (Heath and Rock, 1996). Finally, the trans-A2 double bond is reduced with FabI to create a saturated carbon chain. The M. xantbus fatty acid elongation machinery is spectacularly redundant. There are multiple homologs for almost every enzyme. MXAN4772-4768 contains homologs of plsX, fabD (malonyl transacetylase, which transfers the malonyl group from a CoA moiety to an ACP), fabG, acpP (acylcarrierprotein),and fabF (Fig. 6). This cluster is organized similarly to the E. coli fatty acid biosynthetic cluster (plsX,fabH, fabD, fabG, acpP, and fabF) with the notable absence of fabH. Also, the rpmF gene is upstream of the E. coli cluster but transcribed in the same orientation. A second operon (MXAN63926401) contains homologs of acpP, fab2, a P-lactamase, fabF, fabF, f a b z , fabG, fabG, fabF, and fabF (Fig. 7).
14. PREDATION, SIGNALING, AND DEVELOPMENT
found in M. xanthus fatty acids can be introduced by just two desaturases (A5 and All, [Curtis et al., 2006]), and the M. xanthus genome contains many desaturase homologs. Only trace amounts of 16:106/7 contradict the desaturase hypothesis. Fatty acid diversity in S . aurantiaca is due to nontraditional combinations of pathways. Supplementing the S. aurantiaca BCKAD mutant with isovalerate enriched for both iso-odd and iso-even fatty acids, when it would be predicted to enrich for only iso-odd (Dickschat et al., 2005), consistent with the use of OL oxidation during de novo fatty acid synthesis. S. aurantiaca fed with labeled isol7:O produced labeled isol6:0, isolS:O, isol4:0, isol3:0, and isol1:O products, consistent with the use of a and p fatty acid oxidation. Labeled isol7:O was also found as isol5:1, suggesting that unsaturation is introduced into the preformed fatty acid by a desaturase. It is unclear whether desaturase activity accounts for the major portion of unsaturated fatty acids, as the points of unsaturation would require many different desaturases. For example, isol7:l includes 04, 0 5 , 06, and 07 isomers, which would require the action of four separate desaturases. Alternatively, 01 and p oxidation of fatty acids may increase diversity. Clearly, fatty acid biosynthesis of S. aurantiaca is not a unidirectional, cyclic pathway but involves both extension and retraction with possible intermediate desaturation.
In total there are at least three FabZ/A homologs (two in the clusters and one elsewhere) and five FabF/B homologs. Homology alone cannot distinguish FabZ from FabA or FabF from FabB, as the M. xanthus proteins diverge greatly from other characterized homologs. The extraordinary redundancy may relate to the immense fatty acid diversity. Though many of these enzymes have yet to be characterized, the function for one fatty acid biosynthetic enzyme has been elucidated. The enzyme encoded by a fubH gene, MXAN0853, has been shown to be the major FabH for initiating straight-chain fatty acids (Bode et al., 2006a). A mutant of MXAN083 has severely reduced levels of straight-chain fatty acids, while branched-chain fatty acids collectively increase in compensation. Residual levels of straight-chain fatty acids are produced through the activity of an unknown FabH by using butyrylCoA as the primer. Other fabH genes, MXANO215 and MXAN7353, were also disrupted but had no effect on fatty acid composition. In E. coli, unsaturated fatty acids are made as part of the fatty acid elongation cycle. FabA introduces cis double bonds at many places during fatty acid extension, but FabB extends a specific molecule, thereby creating only one unsaturated fatty acid. It is unclear how unsaturated fatty acids are made in M. xanthus. The multiple FabNZ and FabF/B homologs in M . xanthus could generate the different unsaturated fatty acids (Curtis et al., 2006; Kearns et al., 2001). In cyanobacteria and B a d lus, unsaturated fatty acids are generated by desaturases that introduce double bonds into fatty acids at specific points from the A end. Most of the points of unsaturation
mpPV fabZI/P-lactam!se
6395
6396
fdF
fdF
253
Phospholipid Biosynthesis During phospholipid biosynthesis, the newly formed fatty acyl chains are attached to glycerol-3-phosphate to create phosphatidic acid, and then the head group is
\-&)l61pIsI fab
6399
fdG
1 '
fdGv
6400
6401
fdF
fdF
Figure 7 A second putative fatty acid biosynthesis cluster operon in M . xanthus. The operon contains 10 genes (MXAN6392 through 6401): acpP, fubz, a 6-lactamase gene, fubF, fabF, fubz, fubG, fabG, fabF, and fu6F.
254
added (Cronan and Rock, 1996). First, a fatty acid is esterified to the sn-1 position by glycerol-3-phosphate acyltransferase (PlsB). M . xanthus contains two homologs of this essential enzyme (EC 2.3.1.15, MXAN3288 [PlsBl] and MXAN1675 [PlsB2]), neither of which is essential, suggesting that both are functional (Curtis et al., 2006). While the sn-1 position in E. coli is occupied solely by saturated fatty acids (unless the cells are cold shocked) (Cronan and Rock, 1996),M . xanthus has predominantly unsaturated fatty acids at sn-1 (Curtis et al., 2006). It would appear that both PlsBl and PlsB2 can use unsaturated fatty acids, as elimination of either enzyme does not eliminate sn-1 unsaturated fatty acids from PE. PlsB1 appears to have a slight preference for longer-chain fatty acids, as PE species containing 17 carbon fatty acids decrease in a mutant lacking this enzyme (Curtis et al., 2006). In the second step, 1-acyl-sn-glycerol-3-phosphate acyltransferase (PlsC) adds a fatty acid to the sn-2 position of 1-acyl-sn-glycerol-3-phosphate to generate phosphatidic acid. Five PlsC homologs have been identified (EC 2.3.1.51, PlsC1, MXAN3330; PlsC2, MXAN3969; PlsC3, MXAN5578; PlsC4, MXANO955; PlsC5, MXAN0147). Disruption of the genes encoding PlsC2, PlsC4, and PlsC5 resulted in almost no detectable change to the membrane, suggesting either that those genes are not necessary for phospholipid biosynthesis or that there is functional redundancy (Curtis et al., 2006). Finally the head groups are synthesized on phosphatidic acid. Radiolabeling studies indicate that 76% of the M. xanthus phospholipid is PE, while phosphatidylglycerol (PG) constitutes 9% and cardiolipin 1 % (Orndorff and Dworkin, 1980). The relative amounts of PE, PG, and cardiolipin in the M. xanthus membrane fall along predicted lines for the Proteobacteria. However, relative to E. coli, the outer membrane of M. xanthus is unusually rich in PE, resulting in a lower buoyant density (Orndorff and Dworkin, 1980; Simunovic et al., 2003). PE is derived by first charging phosphatidic acid with CMP to create CDP-diacyl glycerol and then exchanging CDP with serine to create phosphatidylserine, which is decarboxylated to PE. The first step is catalyzed by phosphatidate cytidylyltransferase (EC 2.7.7.41, CdsA, MXAN2556). While no clear phosphatidylserine synthase homolog was found to mediate the serine substitution, there is a homolog for the terminal enzyme phosphatidylserine decarboxylase (EC 4.1.1.65, Psd, MXAN3724). The CDP-diacylglycerol intermediate is also used for PG synthesis. A glycerol-3-phosphate molecule is substituted for CDP to create phosphatidylglycerolphosphate, which is then dephosphorylated to create PG. Two PG molecules
STRUCTUREAND METABOLISM are condensed (with one glycerol as a leaving group) to synthesize cardiolipin. Glycerol-3-phosphate substitution is performed by phosphatidylglycerolphosphate synthase (EC 2.7.8.5, PgsA, MXAN4626). Homologs for the PgpA and PgpB phosphatidylglycerolphosphate phosphatases were not evident, but M . xanthus contains a cardiolipin synthase homolog (EC 2.7.8.-, Cls, MXAN0537). Phosphatidylinositol (PI) has not been detected in M. xanthus despite extensive examination (Orndorff et al., 1980) but has been found in other myxobacteria. PI accounts for 18 to 25% of S. aurantiaca phospholipid (Caillon et al., 1983) and 7% of M . fulvus phospholipid (Kleinig, 1972).Phospholipase C-mediated hydrolysis of PI generates phosphorylated inositol signaling molecules in eukaryotic organisms. Interestingly, release of phosphorylated inositol was observed during aggregation of S. aurantiaca (Benaissa et al., 1994).111. xanthus appears to have some elements of inositol metabolism. Three genes in a cluster are predicted to involve inositol in some fashion. The protein predicted by MXAN0450 has >30% identity to a CDP-diacylglycerol-inositol3-phosphatidyltransferase from Bacteroides thetaiotaomicron but does not match a COG. MXAN045 1 may encode an inositophosphorylceramide synthase, which would add inositol to a ceramide. The predicted protein has 28% identity to the enzyme from Issatchenkia orientalis but also does not match a COG. The clearest homolog of the cluster is MXAN0452, encoding a myo-inositol-l-phosphate synthase (EC 5.5.1.4, MXAN0452) that converts glucose-6-phosphate to inositol-1-phosphate. However, the production and fate of inositol in M . xanthus are unknown.
Ceramide and Sphingolipid Biosynthesis Ceramides are long-chain 1,3-dihydroxy-2-amino bases with amide-attached fatty acids. While they have not yet been examined in M . xanthus, multiple forms have been found in Cystobacter fuscus (Eckau et al., 1984) and Myxococcus stipitatus (Stein and Budzikiewicz, 1988). Ceramides regulate many cellular functions such as cell proliferation, differentiation, apoptosis, and the inflammatory response in eukaryotes (for a review, see Hannun, 1994). The structural diversity among the myxobacterial ceramides is immense. There is variation in the length, branching, and unsaturation in the acyl backbones of both the sphinganine base and amidelinked fatty acid. The head groups also offer another level of structural diversity, as the ceramides found in M . stipitatus had phosphoethanolamine at the sphinganine-1 position while those found in C. fuscus had no additional attachment. How this diversity is produced
14. PREDATION, SIGNALING, AND DEVELOPMENT and what role it plays in the life cycle of these organisms is presently unclear but potentially relevant to the developmental cycle.
Spore-Specific Products The culmination of fruiting body development is the production of dormant, heat-resistant spores. Little is known about the structure of the spore, and even less is known about the contribution of the envelope components to dormancy and germination. Aside from the spore coat proteins, which have been studied in some detail and are not discussed here, two other spore-specific products are known: ether lipids and trehalose. Annotation of the genome revealed some clear candidates for the biosynthetic pathways for both spore-specific products. It should now be possible to make mutations in the relevant genes and assess their contributions to the construction and durability of the spore.
Ether Lipid Biosynthesis Though ester linkages are the most common linkages in PE of the Bacteria, myxobacterial phospholipids also contain alkyl ether linkages, with an ether bond between the acyl chain and glycerol backbone, and alkyl-l-enyl linkages in which the ether link is followed by a double bond between the first two carbons on the acyl chain (Fig. 8). Phospholipids containing ether-linked chains (ether lipids) are prevalent in myxobacteria. Only 28% of S. aurantiaca PE is the traditional diacyl form, while 3 8 % is alkyl-acyl form and 34% is dialkyl. S. aurantiaca PG is 36% diacyl, 50% alkyl-acyl, and 14% dialkyl, while all of the PI is in the dialkyl form (Caillon et al., 1983). M. fulvus PE is 84% diacyl, 2 % alkyl-acyl, and 14% alk-lenyl-acyl (Kleinig, 1972).The PI in this organism is 71% diacyl, 22% alkyl-acyl, and 7% alk-l-enyl-acyl. M. xanthus produces two ether lipids during development (Ring et al., 2006). Perhaps more surprising given the immense fatty acid diversity of M. xanthus PE, only the branched-chain fatty acid isol5:O is found in ether-linked lipids. The first is PE with isol5:O at the Ester (acyl)
Ether (aikjti)
Ak-I -enyl
Figure 8 Acyl chains can be linked to the glycerol backbone of certain lipids in three ways. First i s the ester linkage; this i s the most common linkage type. Second is the ether linkage, requiring a fatty alcohol instead of a fatty acid. Third is the alk-l-enyl linkage, where an unsaturation immediately follows the ether bond.
255 sn-1 position in an alk-l-enyl linkage and isol5:O esterlinked at the sn-2 position (this compound is referred to as VEPE). The second compound is an unusual neutral lipid, with isol5:O ester-linked to the sn-1 and sn-2 positions, while isol5:O fatty acid is ether-linked to the sn-3 position (this compound is referred to as TG-1). Both VEPE and TG-1 are highly enriched in myxospores. Ether lipids are more stable to environmental stresses and may improve the spore resistance properties. Mutants unable to synthesize branched-chain fatty acids are developmentally defective but can be complemented by adding exogenous TG-1. Interestingly, several mutants blocked at multiple points of development accumulate VEPE to the same levels as the wild type but have reduced levels of TG-1, suggesting that TG-1 synthesis may be tied to developmental signaling. Ether lipid synthesis in eukaryotic organisms begins by transferring a fatty acyl chain to the sn-1 position of dihydroxyacetone phosphate (DHAP) (for a review, see Nagan and Zoeller, 2001). A fatty alcohol is then exchanged for the fatty acid, creating the ether linkage. The sn-2 keto group is reduced to an alcohol, and the second fatty acid is attached. The phosphate group is removed, and the PE head group is attached from CDPethanolamine. M. xanthus contains homologs of some of the eukaryotic enzymes. The first few steps of ether lipid synthesis in M. xanthus may be accomplished by MXAN1675 and MXAN1676. MXAN1675 (PlsB2) is a unique gene fusion containing a fatty acid reductase at the N terminus and a glycerol3-phosphate acyltransferase at the C terminus (Curtis et al., 2006). As such, the C-terminal domain could create l-acyl-glycerol-3-phosphateand the N-terminal domain could generate the fatty alcohol from a fatty acyl-CoA. Interestingly, while bacterial fatty acid reduction requires two separate enzymes (Reiser and Somerville, 1997), eukaryotic fatty acid reductases are able to reduce fatty acids to fatty alcohols with one enzyme (Metz et al., 2000; Mot0 et al., 2003), and the N-terminal portion of PlsB2 more closely resembles eukaryotic enzymes. The fatty alcohol exchange step may be performed by the 1-alkyldihydroxyacetone 3-phosphate synthase (EC 2.5.1.26) homolog MXAN1676 (45% amino acid identity to the human counterpart). As the human enzyme uses a DHAP backbone as opposed to the bacterial G3P backbone, oxidation of the G3P sn-2 alcohol to a keto group may be necessary for l-alkydihydroxyacetone 3-phosphate synthase function. While there is a reductase in the cluster (MXAN1674), another possibility is M. xanthus SocA (MXAN5208), which is the first enzyme reported to reduce the sn-2 alcohol of a lysophospholipid (Avadhani et al., 2006).
STRUCTUREAND METABOLISM
256 Trehalose Biosynthesis Trehalose is composed of two glucose monomers with an a 1-1 linkage and confers resistance to osmotic stress (Styrvold and Strom, 1991), heat, and desiccation (Crowe et al., 1984). Given these properties, it is not surprising that trehalose is found in the spores and cysts of a variety of organisms, including M. xanthus (Crowe et al., 1984; McBride and Ensign, 1987a, 1987b; McBride and Zusman, 1989). There are many pathways for trehalose synthesis. In one pathway, glucose-6-phosphate is condensed with UDPD-glucose by trehalose-6-phosphate synthase (EC 2.4.1.15, OtsA, MXAN1192) to create trehalose-6phosphate (Caspi et al., 2006). Then, the phosphate is removed by trehalose-6-phosphate phosphatase. There is no clear M. xanthus homolog of this enzyme, though another phosphatase may perform this function. Trehalose accumulation in M. xanthus requires SigD, a sigma factor necessary for stationary-phase survival and development (Ueki and Inouye, 1998). In the absence of SigD, trehalose synthesis is severely limited in myxospores. The fate of trehalose in M. xanthus is a mystery. In both glycerol spores and fruiting body spores, 75% of the trehalose disappears within the first 2.5 h of germination (McBride and Zusman, 1989).Approximately 25% of trehalose is released into the medium but eventually consumed by the germinating spores. In E. coli, trehalose is degraded to glucose by a periplasmic trehalase; no homolog exists in M. xanthus. Trehalase activity was not detected in spore extracts, and the glucose concentration was not elevated. In fact, the germinating spores were unable to utilize exogenously added glucose (McBride and Zusman, 1989).
SUMMARY The complexity of the myxobacteria is not limited to patterns of behavior but is etched into the fabric of the large and diverse genome. The small sampling of metabolic pathways examined here lead to the following conclusions.
1. Despite the fact that 111. xanthus is thought to subsist on the protein component of prey cells, catabolic pathways for seven amino acids are absent. 2. Pathways exist to salvage purines and pyrimidines for incorporation into nucleic acids. 3 . M. xanthus contains many lipases, some of which have relevance in aggregation and sporulation, and released fatty acids are likely catabolized by j3 oxidation in M. xanthus.
4. Although M. xanthus has homologs for a functional PTS system and both produces and consumes glycogen, it appears that there is no fully functioning glycolysis pathway. Genome analysis has deepened the mystery of M. xanthus sugar metabolism. 5. Lipid biogenesis is spectacularly complex, and the source of unsaturated fatty acids in M. xanthus is unknown, though there is evidence that desaturases may be involved. Fatty acid biogenesis in S. aurantiaca is even more astounding as it uses a and j3 oxidation to generate fatty acid diversity. 6. Myxobacteria produce ether lipids, which is unusual for prokaryotes. 7. While M. xanthus produces the spore-protectant trehalose, it is unclear how this compound is synthesized or degraded. We thank Nilesh Patel, James Henriksen, and Zhengchang Su for help with the genome annotation. This material is based upon work supported by the National Science Foundation under Grant No. 0343874 to L.S.
References Alonso-Casajus, N., D. Dauvillee, A. M. Viale, F. J. Munoz, E. Baroja-Fernandez, M. T. Moran-Zorzano, G . Eydallin, S. Ball, and J. Pozueta-Romero. 2006. Glycogen phosphorylase, the product of the glgP Gene, catalyzes glycogen breakdown by removing glucose units from the nonreducing ends in Escherichia coli. J. Bacteriol. 1885266-5272. Avadhani, M., R. Geyer, D. C. White, and L. J. Shimkets. 2006. Lysophosphatidylethanolamine is a substrate for the shortchain alcohol dehydrogenase SocA from Myxococcus xanthus. J. Bacteriol. 188:8543-8550. Behmlander, R. M., and M. Dworkin. 1991. Extracellular fibrils and contact-mediated cell interactions in Myxococcus xanthus. J. Bacteriol. 173:7810-7820. Benaissa, M., J. Vieyres-Lubochinsky, R. Odeide, and B. Lubochinsky. 1994. Stimulation of inositide degradation in clumping Stigmatella aurantiaca. J. Bacteriol. 176:13901393. Bode, H. B., J. S. Dickschat, R. M. Kroppenstedt, S. Schulz, and R. Miiller. 2005. Biosynthesis of iso-fatty acids in myxobacteria: iso-even fatty acids are derived by alphaoxidation from iso-odd fatty acids. ]. Am. Chem. Soc. 127:5 32-5 3 3, Bode, H. B., M. W. Ring, D. Kaiser, A. C. David, R. M. Kroppenstedt, and G . Schwar. 2006a. Straight-chain fatty acids are dispensable in the myxobacterium Myxococcus xanthus for vegetative growth and fruiting body formation. J. Bacteriol. 1885632-5634. Bode, H. B., M. W. Ring, G. Schwar, R. M. Kroppenstedt, D. Kaiser, and R. Muller. 2006b. 3-Hydroxy-3-methylglutarylcoenzyme A (CoA) synthase is involved in biosynthesis of isovaleryl-CoA in the myxobacterium Myxococcus xanthus during fruiting body formation. J. Bacteriol. 188:65246528.
14. PREDATION,SIGNALING,AND DEVELOPMENT Bretscher, A. P., and D. Kaiser. 1978. Nutrition of Myxococcus xanthus, a fruiting myxobacterium. J. Bacteriol. 133:763768. Caillon, E., B. Lubochinsky, and D. Rigomier. 1983. Occurrence of dialkyl ether phospholipids in Stigmatella aurantiaca DW4. J. Bacteriol. 153:1348-1351. Campbell, J. W., R. M. Morgan-Kiss, and J. E. Cronan, Jr. 2003. A new Escherichia coli metabolic competency: growth on fatty acids by a novel anaerobic beta-oxidation pathway. Mol. Microbiol. 47:793-805. Caspi, R.,H. Foerster, C. A. Fulcher, R. Hopkinson, J. Ingraham, P. Kaipa, M. Krummenacker, S. Paley, J. Pick, S. Y. Rhee, C. Tissier, P. Zhang, and P. D. Karp. 2006. MetaCyc: a multiorganism database of metabolic pathways and enzymes. Nucleic Acids Res. 34:D511-D516. Clark, D. P., and J. E. Cronan. 1996. Two-carbon compounds and fatty acids as carbon sources, p. 343-357. In F. C. Neidhardt, R. Curtiss, J. L. Ingram, E. C. C. Lin, K. B. Low, and B. Magasanik (ed.), Escherichia coli and Salmonella. ASM Press, Washington, DC. Cronan, J. E., and C. 0. Rock. 1996. Biosynthesis of membrane lipids, p. 612-636. In F. C. Neidhardt, R. Curtiss, J. L. Ingram, E. C. C. Lin, K. B. Low, and B. Magasanik (ed.), Escherichia coli and Salmonella. ASM Press, Washington, DC. Cropp, T. A., A. A. Smogowicz, E. W. Hafner, C. D. Denoya, H. A. McArthur, and K. A. Reynolds. 2000. Fatty-acid biosynthesis in a branched-chain alpha-keto acid dehydrogenase mutant of Streptomyces avermitilis. Can. J. Microbiol. 46~506-514. Crowe, J. H., L. M. Crowe, and D. Chapman. 1984. Infrared spectroscopic studies on interactions of water and carbohydrates with a biological membrane. Arch. Biochem. Biophys. 232:400-407. Curtis, P. D., R. Geyer, D. C. White, and L. J. Shimkets. 2006. Novel lipids in Myxococcus xanthus and their role in chemotaxis. Environ. Microbiol. 8:1935-1949. Dickschat, J. S., H. B. Bode, R. M. Kroppenstedt, R. Miiller, and S. Schulz. 2005. Biosynthesis of iso-fatty acids in myxobacteria. Org. Biomol. Chem. 3:2824-2831. Downard, J., S. V. Ramaswamy, and K. S. Kil. 1993. Identification of esg, a genetic locus involved in cell-cell signaling during Myxococcus xanthus development. J. Bacteriol. 175:7762-7770. Downard, J., and D. Toal. 1995. Branched-chain fatty acids: the case for a novel form of cell-cell signalling during Myxococcus xanthus development. Mol. Microbiol. 16:171-175. Dworkin, M. 1962. Nutritional requirements for vegetative growth of Myxococcus xanthus. J. Bacteriol. 84:250-257. Eckau, H., D. Dill, and H. Budzikiewicz. 1984. Novel ceramides from Cystobacter fuscus (Myxobacterales). 2. Naturforsch. C J. Biosci. 39:l-9. Erni, B., B. Zanolari, and H. P. Kocher. 1987. The mannose permease of Escherichia coli consists of three different proteins. Amino acid sequence and function in sugar transport, sugar phosphorylation, and penetration of phage lambda DNA. J. Biol. Chem. 26252384247. Goldman, B. S., W. C. Nierman, D. Kaiser, S. C. Slater, A. S. Durkin, J. A. Eisen, C. M. Ronning, W. B. Barbazuk,
25 7 M. Blanchard, C. Field, C. Halling, G. Hinkle, 0. Iartchuk, H. S. Kim, C. Mackenzie, R. Madupu, N. Miller, A. Shvartsbeyn, S. A. Sullivan, M. Vaudin, R. Wiegand, and H. B. Kaplan. 2006. Evolution of sensory complexity recorded in a myxobacterial genome. Proc. Natl. Acad. Sci. USA 103:15200-15205. Hamberg, M., I. Ponce de Leon, A. Sanz, and C. Castresana. 2002. Fatty acid alpha-dioxygenases. Prostaglandins Other Lipid Mediat. 68-69:363-374. Hannun, Y. A. 1994. The sphingomyelin cycle and the second messenger function of ceramide. J. Biol. Chem. 269:31253128. Heath, R. J., and C. 0. Rock. 1996. Roles of the FabA and FabZ beta-hydroxyacyl-acyl carrier protein dehydratases in Escherichia coli fatty acid biosynthesis. J. Biol. Chem. 271~27795-27801. Hemphill, H. E., and S. A. Zahler. 1968a. Nutrition of Myxococcus xanthus FBa and some of its auxotrophic mutants. J. Bacteriol. 95:lOll-1017. Hemphill, H. E., and S. A. Zahler. 1968b. Nutritional induction and suppression of fruiting in Myxococcus xanthus FBa. J. Bacteriol. 95:1018-1023. Hubbard, P. A., X. Liang, H. Schulz, and J. J. Kim. 2003. The crystal structure and reaction mechanism of Escherichia coli 2,4-dienoyl-CoA reductase. J. Biol. Chem. 278:3755337560. Kearns, D. B., A. Venot, P. J. Bonner, B. Stevens, G.-J. Boons, and L. J. Shimkets. 2001. Identification of a developmental chemoattractant in Myxococcus xanthus through metabolic engineering. Proc. Natl. Acad. Sci. USA 98:13990-13994. Kim, S. H., S. Ramaswamy, and J. Downard. 1999. Regulated exopolysaccharide production in Myxococcus xanthus. J. Bacteriol. 181:1496-1507. Kleinig, H. 1972. Membranes from Myxococcus fulvus (Myxobacterales) containing carotenoid glucosides. I. Isolation and composition. Biochim. Biophys. Acta 274:489-498. Klingsbichel, E. 1996. Esterase EstA from Pseudomonas marginata: Heterologous Expression, Biological, Biochemical, and Biocatalytical Characterization. Ph.D. thesis. Technical University of Graz, Graz, Austria. Kuspa, A., L. Plamann, and D. Kaiser. 1992. Identification of heat-stable A-factor from Myxococcus xanthus. J. Bacteriol. 174~3319-3326. Lau, J., S. Frykman, R. Regentin, S. Ou, H. Tsuruta, and P. Licari. 2002. Optimizing the heterologous production of epothilone D in Myxococcus xanthus. Biotechnol. Bioeng. 78:280-288. Loebeck, M. E., and H. P. Klein. 1956. Substrates for Myxococcus virescens with special reference to eubacterial fractions. J. Gen. Microbiol. 14:281-289. Lu, A., K. Cho, W. P. Black, X. Y. Duan, R. Lux, Z. Yang, H. B. Kaplan, D. R. Zusman, and W. Shi. 2005. Exopolysaccharide biosynthesis genes required for social motility in Myxococcus xanthus. Mol. Microbiol. 55:206-220. Mahmud, T., H. B. Bode, B. Silakowski, R. M. Kroppenstedt, M. Xu, S. Nordhoff, G. Hofle, and R. Miiller. 2002. A novel biosynthetic pathway providing precursors for fatty acid biosynthesis and secondary metabolite formation in myxobacteria. J. Biol. Chem. 277:32768-32774.
258 Mahmud, T., S. C. Wenzel, E. Wan, K. W. Wen, H. B. Bode, N. Gaitatzis, and R. Muller. 2005. A biosynthetic pathway to isovaleryl-CoA in myxobacteria: the involvement of the mevalonate pathway. Chembiochem 6:322-330. McBride, M. J., and J. C. Ensign. 1987a. Effects of intracellular trehalose content on Streptomyces griseus spores. J. Bacteriol. 169:4995-5001. McBride, M. J., and J. C. Ensign. 198710. Metabolism of endogenous trehalose by Streptomyces griseus spores and by spores or cells of other actinomycetes. J. Bacteriol. 1695002-5007. McBride, M. J., and D. R. Zusman. 1989. Trehalose accumulation in vegetative cells and spores of Myxococcus xanthus. J. Bacteriol. 171:6383-6386. Metz, J. G., M. R. Pollard, L. Anderson, T. R. Hayes, and M. W. Lassner. 2000. Purification of a jojoba embryo fatty acyl-coenzyme A reductase and expression of its cDNA in high erucic acid rapeseed. Plant Physiol. 122:635-644. Moraleda-Muiioz, A., and L. J. Shimkets. 2007. Lipolytic enzymes in Myxococcus xanthus. J. Bacteriol. 189:30723080. Moto, K., T. Yoshiga, M. Yamamoto, S. Takahashi, K. Okano, T. Ando, T. Nakata, and S. Matsumoto. 2003. Pheromone gland-specific fatty-acyl reductase of the silkmoth, Bombyx mori. Proc. Natl. Acad. Sci. USA 100:9156-9161. Nagan, N., and R. A. Zoeller. 2001. Plasmalogens: biosynthesis and functions. Prog. Lipid Res. 40:199-229. Nariya, H., and S. Inouye. 2002. Activation of 6-phosphofructokinase via phosphorylation by Pkn4, a protein Ser/Thr kinase of Myxococcus xanthus. Mol. Microbiol. 46:13531366. Nariya, H., and S. Inouye. 2003. An effective sporulation of Myxococcus xanthus requires glycogen consumption via Pkn4-activated 6-phosphofructokinase. Mol. Microbiol. 4 9 5 1 7-528. Nariya, H., and S. Inouye. 2005. Modulating factors for the Pkn4 kinase cascade in regulating B-phosphofructokinase in Myxococcus xanthus. Mol. Microbiol. 56:13141328. Neidhardt, F. C., and H. E. Umbarger. 1996. Chemical composition of Escherichia coli, p. 13-16. In F. C. Neidhardt, R. Curtiss, J. L. Ingram, E. C. C. Lin, K. B. Low, and B. Magasanik (ed.), Escherichia coli and Salmonella. ASM Press, Washington, DC. Ollis, D. L., E. Cheah, M. Cygler, B. Dijkstra, F. Frolow, S. M. Franken, M. Harel, S. J. Remington, I. Silman, J. Schrag, et al. 1992. The alpha/beta hydrolase fold. Protein Eng. 5 :197-2 11. Orndorff, P. E., and M. Dworkin. 1980. Separation and properties of the cytoplasmic and outer membranes of vegetative cells of Myxococcus xanthus. J. Bacteriol. 141:914-927. Postma, P. W., J. W. Lengeler, and G. R. Jacobson. 1996. Phosphoeno1pyruvate:carbohydrate phosphotransferase systems, p. 1149-1174. In F. C. Neidhardt, R. Curtiss, J. L. Ingram, E. C. C. Lin, K. B. Low, and B. Magasanik (ed.), Escherichia coli and Salmonella. ASM Press, Washington, DC. Postma, P. W., J. W. Lengeler, and G. R. Jacobson. 1993. Phosphoenolpyruvate: carbohydrate phosphotransferase systems of bacteria. Microbiol. Rev. 57543-594.
STRUCTUREAND METABOLISM Reiser, S., and C. Somerville. 1997. Isolation of mutants of Acinetobacter calcoaceticus deficient in wax ester synthesis and complementation of one mutation with a gene encoding a fatty acyl coenzyme A reductase. J. Bacteriol. 179:2969-2975. Ring, M. W., G . Schwar, V. Thiel, J. S. Dickschat, R. M. Kroppenstedt, S. Schulz, and H. B. Bode. 2006. Novel isobranched ether lipids as specific markers of developmental sporulation in the myxobacterium Myxococcus xanthus. J. Biol. Chem. 281:36691-36700. Saffert, A., J. Hartmann-Schreier, A. Schon, and P. Schreier. 2000. A dual function alpha-dioxygenase-peroxidase and NAD( +) oxidoreductase active enzyme from germinating pea rationalizing alpha-oxidation of fatty acids in plants. Plant Physiol. 123:1545-1552. Simunovic, V., F. C. Gherardini, and L. J. Shimkets. 2003. Membrane localization of motility, signaling, and polyketide synthetase proteins in Myxococcus xanthus. J. Bacteriol. 185:5066-5075. Stamm, I., F. Lottspeich, and W. Plaga. 2005. The pyruvate kinase of Stigmatella aurantiaca is an indole binding protein and essential for development. Mol. Microbiol. 56:1386-1395. Stein, J., and H. Budzikiewicz. 1988. Bacterial components .36. ceramide-1-phosphoethanolaminesfrom Myxococcus stipitatus. Z. Naturforsch. B J. Chem. Sci. 43:1063-1067. Styrvold, 0. B., and A. R. Strom. 1991. Synthesis, accumulation, and excretion of trehalose in osmotically stressed Escherichia coli K-12 strains: influence of amber suppressors and function of the periplasmic trehalase. J. Bacteriol. 173~1187-1192. Toal, D. R., S. W. Clifton, B. A. Roe, and J. Downard. 1995. The esg locus of Myxococcus xanthus encodes the E l a and E l p subunits of a branched-chain keto acid dehydrogenase. Mol. Micro biol. 16: 177-1 89. Tsai, W. C., and C. A. Westby. 1978. Synthesis and salvage of purines during cellular morphogenesis of Myxococcus xanthus. J. Bacteriol. 136582-587. Ueki, T., and S. Inouye. 1998. A new sigma factor, SigD, essential for stationary phase is also required for multicellular differentiation in Myxococcus xanthus. Genes Cells 3:371-385. Ware, J. C., and M. Dworkin. 1973. Fatty acids of Myxococcus xanthus. J. Bacteriol. 115:253-261. Watson, B. F., and M. Dworkin. 1968. Comparative intermediary metabolism of vegetative cells and microcysts of Myxococcus xanthus. J. Bacteriol. 96:1465-1473. Weimar, J. D., C. C. DiRusso, R. Delio, and P. N. Black. 2002. Functional role of fatty acyl-coenzyme A synthetase in the transmembrane movement and activation of exogenous long-chain fatty acids. Amino acid residues within the ATPI AMP signature motif of Escherichia coli FadD are required for enzyme activity and fatty acid transport. J. Biol. Chem. 277~29369-29376. Yang, Z., X. Ma, L. Tong, H. B. Kaplan, L. J. Shimkets, and W. Shi. 2000. Myxococcus xanthus dif genes are required for biogenesis of cell surface fibrils essential for social gliding motility. J. Bacteriol. 1825793-5798. Youderian, P., N. Burke, D. J. White, and P. L. Hartzell. 2003. Identification of genes required for adventurous gliding motility in Myxococcus xanthus with the transposable element mariner. Mol. Microbiol. 49555-570.
Myxobacteria: Multicellularity and Differentiation Edited by David E. Whitworth 0 2008 ASM Press, Washington, D.C.
Helge B. Bode Rolf Miiller
Secondary Metabolism in Myxobacteria
MYXOBACTERIAL SECONDARY METABOLITES
Myxobacteria as Proficient Producers of Biologically Active Metabolites-a Brief Introduction In the past 30 years myxobacteria have been established as proficient producers of various secondary metabolites (Gerth et al., 2003; Bode and Miiller, 2006) and are regarded today as one of the few important sources for microbial natural products besides actinomycetes and fungi. Mainly due to the efforts of the research groups of Hans Reichenbach and Gerhard Hofle at the German Research Centre of Biotechnology (GBF), more than 7,500 myxobacterial strains have been isolated and more than 100 different basic structures with approximately 500 structural variants have been described, as has been discussed in several recent reviews (Reichenbach, 2001; Gerth et al., 2003; Hofle and Reichenbach, 2005). Accordingly, the goal of this chapter is not to give detailed information about the past research regarding myxobacterial secondary metabolism (which additionally is described in chapter 19, focusing on the genus Sorungium) but to highlight recent developments which
15
add to, and also significantly change, our approach to exploit the enormous chemical diversity that is found in myxobacteria. The advanced exploitation of this resource is of special interest because the mode of action of natural products from myxobacteria is often unusual, as they target cellular structures which are only seldom or not at all hit by other secondary metabolites. Epothilone from Sorungium cellulosum recently successfully finished phase I11 clinical trials as an anticancer agent, as it represents a paclitaxel mimetic. Furthermore, this compound can be used to treat paclitaxel-resistant tumors, shows good water solubility, and can be produced by fermentation (Reichenbach, 2001; Hofle and Reichenbach, 2005). Since the isolation of epothilone, additional myxobacterial compounds have been found that show the opposite mode of action by destabilizing microtubuli (i.e., disorazol [Elnakady et al., 20041 and tubulysin [Sasse et al., 20001) or act on the actin skeleton (i.e., rhizopodin [Gronewold et al., 19991 and chondramid [Sasse et al., 19981) (Fig. l a ) . Interestingly, secondary metabolites from myxobacteria often include structural elements which are rarely found in other sources. From the biosynthetic point of view, most of
Helge B. Bode and Rolf Miiller, Institut fur Pharrnazeutische Biotechnologie, Universitat des Saarlandes, Postfach 15 11 50, 66041 Saarbriicken, Germany.
259
STRUCTUREAND METABOLISM
260 a.
Chivosazol A
OH
Disorazol A,
H
Chondramid A b
Aurafuron A
OH
Cycloartenol
Leupyrrin A,
Figure 1 Myxobacterial secondary metabolites that act on the eukaryotic cytoskeleton (a) and examples of novel secondary metabolites from myxobacteria (b).
the isolated compounds represent hybrids of polyketides (PKs)and structural elements derived fromnonribosomally made peptides (NRPs) (Silakowskiet al., 2001a), whereas pure PKs are rarely found. Examples for simple PKs are the
aurafurons (Kunze et al., ZOOS), tuscolid, tuscuron (Niggemann et al., 2004), and dawenol (Soker et al., 2003) (Fig. lb). Only fungi and myxobacteria have been described as producers of P-methoxyacrylates, which have been
15. SECONDARY METABOLISM IN MYXOBACTERIA described as potent inhibitors of the mitochondria1respiratory cytochrome bc, complex. Synthetic derivatives of the fungal strobilurins are used as fungicides in agriculture (H. Sauter, 8 Irseer Naturstofftage der Dechema e.V., 1996). Recently the cyrmenins (Fig. l b ) were found as a new peptide/polyketide hybrid of this class of compounds lacking the bisthiazol moiety found in other myxobacterial P-methoxyacrylates (e.g., myxothiazol, melithiazol, and cystothiazol) (Sasse et al., 2003; Leibold et al., 2004). In actinomycetes and fungi, the two most potent secondary metabolite-producing microbial groups, PKs and NRPs are often modified furthermore by glycosylation (e.g., erythromycin and vancomycin), oxygenation (almost all type I1 PKs), or halogenation (e.g., vancomycin), and other so-called post-PK synthase (PKS) or post-NRP synthase (NRPs) biochemical reactions increase the possible chemical diversity even more. Such postassembly modifications have also been described in myxobacteria (e.g., stigmatellin and chondramid) but seem to be not as common as in actinomycetes and fungi (see “Secondary Metabolism in Myxococcus xanthus” below) Examples for post-PK modified compounds are the cytotoxic chivosazols bearing a 6-deoxyglucose moiety attached to the aglycon (Irschik et al., 1995; Jansen et al., 1997; Perlova et al., 2006b) or theleupyrrins, which are derived from a mixed polyketide/peptide/isoprenoid biosynthesis. The latter compounds are furthermore modified, giving a perfect example for nature’s capabilities in combining different biochemical principles (Bode et al., 2003a, 2004). Four myxobacterial compounds (apicularen Wansen et al., 20001, saframycin [Irschik et al., 19881, chondramide [Jansen et al., 19961, and rhizopodin [Sasse et al., 19931) are known to be highly similar to secondary metabolites isolated from higher marine organisms (e.g., sponges and molluscs) (Bode and Miiller, 2005). These findings raise the question of the origin of these marine compounds, and in fact there is increasing evidence that marine myxobacteria exist (Iizuka et al., 1998; Fudou et al., 2001; Iizuka et al., 2003a, 2003b) and may thus in fact be responsible for the formation of these products. It is clear today that some other compounds isolated from higher organisms are indeed produced by bacterial symbionts within the eukaryote (Piel, 2004; Schmidt et al., 2005). The biotechnological implications of these findings are discussed below. A biochemical curiosity in myxobacterial metabolism is the presence of true steroids, which is highly unusual for prokaryotes (Kohl et al., 1983; Bode et al., 2003b). Besides myxobacteria, only the proteobacterium Methylococcus capsulutus (Bird et al., 1971) and the planctomycete Gemmata obscuriglobus (Pearson et al., 2003)
261
are known for the biosynthesis of these compounds typical for eukaryotes. Whereas Methylococcus and Gemmata species produce only very early intermediates of steroid biosynthesis, almost all intermediates of the human cholesterol biosynthesis except for cholesterol itself have been identified in Nunnocystis exedens. Furthermore, de novo steroid biosynthesis was shown and the first prokaryotic steroid cyclase was cloned from Stigmutella uuruntiucu (Bode et al., 2003b). However, in contrast to their important role in eukaryotes, no biological function could be assigned to steroids in myxobacteria or prokaryotes in general. Loss of steroid production did not result in a specific phenotype in S. aurantiuca.
The Scent of Myxobacteria: Identification of Volatile Compounds Myxobacteria have been known for years for the earthy smell of their cultures, which is due to the production of geosmin (Trowitsch et al., 1981), a degraded sesquiterpene that is also produced by several other organisms and that is responsible for the typical odor of freshly broken soil. A detailed analysis of the emitted volatiles of selected myxobacteria grown on agar plates confirmed the production of geosmin and enabled the elucidation of its biosynthesis, which differs from the biosynthesis in plants and streptomycetes (Dickschat et al., 2005). Interestingly, a plethora of additional compounds from different structural classes have been collected using a closed-loop-stripping apparatus or solid-phase microextraction followed by detailed gas chromatography coupled with mass spectroscopy (GUMS) (Dickschat et al., 2004; Schulz et al., 2004). Besides additional sesquiterpenes, several previously unknown pyrazines, acylphenylketones, and fatty acid derivatives have been identified (Fig. 2a). This diversity is remarkable because only five different myxobacterial strains have been analyzed so far. Several of the detected compounds seemed to be derived from branched-chain amino acids and/or fatty acids, which was confirmed by biosynthetic studies using a combination of classical feeding experiments and GUMS analysis of the produced volatiles as shown in Fig. 2b. This observation once more underlines the importance of fatty acids (see “Biosynthesis of Unusual Precursors for Myxobacterial Secondary Metabolites” below) and branched-chain amino acids for myxobacteria, with the latter ones even being essential for M. xanthus (Bretscher and Kaiser, 1978). Notably, some of these compounds are very similar in structure to stigmolone, which has been described as a developmental signal in S. aurantiaca (Plaga et al., 1998). Because some compounds are produced in relatively high amounts and require a complex biosynthesis,
STRUCTUREAND METABOLISM
262 a) from C. crocatus
po 2-(l-hydroxy-l-methylethyl)3-methoxypyrazine
2,5-bis(l methylethyl) pyrazine
(6R,IOR)-6,1O-dimethylbicyclo [4.4.0]deGl -en-3-one
from S. aurantiaca
0"""u
0
/
methyl 9methyldecanoate
N-(2-phenylethylidene) isopentylamine
zonarene
p ROH OH
geosmin
(1 (10)€,5€)germacradien-1l-ol
b)
A 2\. leucine
@OH
C02H
OH H02CaSCoA 3-hydroxy-3-methylg Iutaryl-CoA
phenylalanine
cinnamic acid
OASACP
@
I
H02C&
Y
U
A
C
OASCoA
I
w 0
S
\
9-methyldecan-3-one
[F] 9-methyl-I-phenyldecan-I-one
in S. aurantiaca Sg a15
in M. xanthus OK1622
P
15. SECONDARY METABOLISM IN MYXOBACTERIA they most likely fulfill a currently unknown biological function and are not just “biochemical accidents.” One might speculate that some of these volatiles might participate in long-distance communication or defense mechanisms.
Secondary Metabolism in M. xanthus The best-studied myxobacterium, M. xanthus DK1622, is the model organism for studying fruiting body formation and motility in myxobacteria. Despite the yellow appearance of M . xanthus DK1622, it was thought to be a nonproducer of secondary metabolites based on investigations in several laboratories in the last 30 years. In contrast, a recent detailed analysis of the genome sequence revealed at least 18 different biosynthesis gene clusters for the production of secondary metabolites, most of them harboring PKS genes and NRPS genes as was previously shown for other myxobacteria (Silakowski et al., 2001a). Of these gene clusters, 14 are located between 3.2 and 5.8 Mb of the genome. However, no evidence for plasmid or chromosome fusion leading to the observed superclustering has been found (Goldman et al., 2006). More than 8.5% of the genome is dedicated to secondary metabolism, which is a higher percentage than observed in the well-known secondary metabolite producers Streptomyces coelicolor (Bentley et al., 2002) (4.5%) and S. avermitilis (Omura et al., 2001; Ikeda et al., 2003) (6.6%). Some of the identified biosynthetic genes show striking homology to such genes encoded in known gene clusters from other myxobacteria, and subsequently the corresponding compounds have been identified after large-scale cultivation, extraction, and purification (H.B. Bode, P. Meiser, and R. Muller, unpublished results). The major compounds produced in liquid media are the myxalamides, with myxalamide A (Jansen et al., 1983; Silakowski et al., 2001b) being the main component. Minor products are the myxalamides B and C, with all myxalamides found as all-trans isomers. Furthermore, the iron siderophores myxochelins A and B (Kunze et al., 1989; Silakowski et al., 2000; Gaitatzis et al., 2001), the myxovirescins A and C (Trowitzsch Kienast et al., 1989; Simunovic et al., 2006), and known and novel myxochromides (Wenzel et al., 2006) have been isolated and characterized from strain DK1622 (Fig. 3). Additionally, completely new compounds have been isolated. The structure of the pigment that gives M . xanthus
263
its typical yellow appearance has been elucidated and named DKxanthene (Meiser et al., 2006). Several members of this mixed polyketide/peptide structure have been identified that show differences in length of the polyene chain, methylation pattern, and/or hydroxylation of the terminal asparagine moiety. DKxanthenenegative mutants show the expected loss of the yellow color and instead have a beige appearance as already described for tan mutants in the literature (Burchard et al., 1977; Laue and Gill, 1995). Interestingly, almost no viable spores can be obtained from these mutants whereas no difference in fruiting body formation and total number of spores can be observed compared to the wild type. Additionally, no difference between the wild type and DKxanthene-negative mutants in the number of glycerol-induced spores (Dworkin and Gibson, 1964) was found, which adds further evidence that the two spore types are biochemically different (Rosenbluh and Rosenberg, 1989). Although the sporulation defect of tan mutants was described previously (Laue and Gill, 1995), nothing was known about the underlying principles. Addition of pure DKxanthene A to DKxanthene-negative mutants restored the sporulation defect at least partially, indicating that the loss of the secondary metabolite itself and not loss of any hypothetical additional factor is responsible for the tan phenotype (Meiser et al., 2006). The direct biological function of these compounds is currently unknown, but they are very unstable in the light, forming numerous cisltrans isomers. This might indicate a role in the adsorbtion of UV light and therefore enable UV protection which might be important for the bacteria, especially for the spores. Additionally, DKxanthenes show antioxidative properties which might contribute to their possible function as cell protectants. The increased production of this family of compounds during fruiting body formation and the close association with the cells support this hypothesis (Meiser et al., 2006). Detailed analysis of 10 different wild-type M . xanthus isolates of various origins showed that all of them produce myxalamides and DKxanthene, whereas myxochromides, myxovirescin, and myxochelin are produced by only some of these strains (D. Krug, G. Velicer, R. Miiller, unpublished results). Therefore, myxalamides and DKxanthenes might be useful as chemotaxonomic markers for M. xanthus species.
Figure 2 (a)Selected volatiles from Chondromyces crocutus and S. uuruntiacu. (b) Biosynthesis of the main volatile compounds 9-methyldecane-3-one and (S)-9-rnethyldecane-3-01from M. xanthus DK1622 and of 9-methyl-1-phenyldecan-1-one from S. auruntiaca Sg a l 5 .
STRUCTUREAND METABOLISM
264
OH
Myxovirescin A Myxovirescin C
Myxalamid A Myxalarnid B Myxalamid C
R = isobutyl R = isopropyl R = ethyl
&I R=0 R = H, H
0
OH HO Myxochelin A Myxochelin B
R = OH R = NH,
R
0
Myxochromide A, R = Et Myxochromide A, R = CH=CH-Me Myxochrornide A4 R = CH=CH-Et
name DKxanthene-492 DKxanthene-504 DKxanthene-508 DKxanthene-518 DKxanthene-520 DKxanthene-534 DKxanthene-544 DKxanthene-548 DKxanthene-560 DKxanthene-574 DKxanthene-5 86
n 0 1 0 1 1 1 2
1 2 2 3
R' CH3 H CH3 CH3 H CH3 CH3 CH3 CH3 CH3 CH3
R2 H H H H H H H CH3 H CH3 H
R3 H H OH H OH OH H OH OH OH OH
Figure 3 Secondary metabolites isolated from M. xanthus DK1622. DKxanthene numbers indicate their molecular weight.
As discussed above, only five structurally different compound classes have been identified from M. xunthus DK1622 to date, whereas the potential biosynthesis of 18 different compounds is encoded in the genome.
The reason for this discrepancy might be that some of the biosynthetic gene clusters are expressed only under certain conditions which have not been identified yet. Besides a broad variation of the culture conditions to
15. SECONDARYMETABOLISM IN MYXOBACTERIA induce the biosynthesis of additional compounds (see, in comparison, Bode et al., 2002), we have also analyzed the secondary metabolite profile during fruiting body formation. Interestingly, several secondary metabolites are produced exclusively during this highly complex process and are currently being isolated and characterized (P. Meiser, H. B. Bode, and R. Muller, unpublished results). In a recent proteome study, the overall expression of the biosynthetic gene clusters was addressed during vegetative growth. Notably, using two-dimensional high-pressure liquid chromatography (HPLC)-MS/MS analysis, proteins corresponding to genes from 11 different biosynthetic gene clusters were detected during exponential growth (Schley et al., 2006). It can therefore be concluded that most of the 18 gene clusters identified are indeed transcribed and translated, which leads to the speculation that indeed further currently unknown and novel secondary metabolites can be found in the future.
MOLECULAR BIOLOGY OF MYXOBACTERIAL SECONDARY METABOLISM A General Introduction to PKSs and NRPSs Most of the myxobacterial secondary metabolites that have been investigated with respect to the underlying biosynthetic mechanisms by genetic and/or (bio)chemical methods are mixtures of PKs and NRPs. Because these natural products are derived from highly complex enzymatic systems called PKSs and NRPSs, these enzymes and the resulting secondary metabolites are introduced briefly (Fig. 4). Nevertheless, it should be mentioned that many bacterial natural products are derived from other biochemical pathways, such as isoprenoids (e.g., geosmin) (Dewick, 2002) or shikimate-derived compounds (e.g., benzoate-derived natural products) (Knaggs, 2003). There are three different types of bacterial PKSs, all of which employ the biochemical principles of the wellknown fatty acid biosynthesis pathway but differ in several aspects and therefore finally produce an enormous variety of low-molecular-weight compounds. Bacterial type I PKSs (Fig. 4a) are multifunctional enzymes that are organized into modules, each of which harbors a set of distinct, noniteratively acting activities (domains) responsible for several biosynthetic steps during the catalysis of one chain elongation cycle of the PK (Staunton and Weissman, 2001). Type I PKSs are involved in the biosynthesis of macrolides (e.g., erythromycin [Fig. 4b]), polyethers (e.g., monensin), or polyenes (e.g., amphotericin), to name just a few. They are also involved in the biosynthesis of polyether compounds from dinoflagellates
265
(Snyder et al., 2003), which are currently the largest secondary metabolites known (their molecular mass can exceed 3,000 Da; e.g., brevetoxin). Malonyl-coenzyme A (CoA) is used almost exclusively for the chain elongation of fatty acids, which after completion of the reductive cascade results in the production of saturated fatty acids (Michal, 1999; Heath and Rock, 2002). PK chain elongation can use a variety of alternative substrates like methylmalonyl-CoA or methoxymalonyl-CoA. Furthermore, every new chain extension cycle can start beginning with any intermediate of the reductive cascade leading to an immense variety of possible compounds whose structures can be predicted to some extent from the order of the domains and the module organization of the corresponding enzymes. Type I1 PKSs are multienzyme complexes carrying a single set of iteratively acting proteins (Fig. 4c); they are involved in the biosynthesis of aromatic and often polycyclic PKs (Shen, 2000). Examples are several antibiotics and anticancer compounds in clinical use like tetracycline or daunorubicin. Type I11 PKSs, also known as chalcone synthase-like PKSs, are homodimeric enzymes which iteratively act as condensing enzymes, as exemplified by the formation of flaviolin (Austin and Noel, 2003) (see also Fig. 10 in chapter 19). They are widespread in plants because they are required for the biosynthesis of various pigments and other plant natural products (e.g., flavonoids, chalcones, and stilbenes) but recently have been shown to be quite common in bacteria as well (Funa et al., 1999) and have also been found recently in myxobacteria (Gross et al., 2006a) (see also chapter 19). It may be mentioned at this point that several exceptions to the presented classification system regarding the different PKSs have already been described (Muller, 2004; Wenzel and Muller, 2005b). With the increasing genomic information of all kinds of species and subsequent biochemical studies, the number of elucidated unusual mechanisms will increase even more, most likely leading to the description of all kinds of transitions between the above-described main principles. NRPSs are similar to the type I PKSs in that they are multifunctional enzymes, organized into modules with sets of noniteratively acting activities for the incorporation and processing of one amino acid per module (Fig. 4d) (Sieber and Marahiel, 200.5). Peptides with chain lengths between 2 and 20 amino acids have been identified as nonribosomally made, including some pharmaceutically important molecules (e.g., cyclosporin and vancomycin [Fig. 4el). The chemical diversity found in the peptides isolated to date is a result of the different ways in which amino acids can be processed and altered during the biosynthesis. Additionally, even
module-2
module-4
module-3
module-5
0
OH
0
b) Propionyl-CoA 6 Methylmalonyl-CoA
c)
@
9 Tetracenomycin C
0
0
[wo 1 R
oxidation hydroxylation methylation
r
Acetyj-CoA
TcmyNr
- 10 CoA - 9 co, module-’ (loading)
-
0 COEnz
9 Malonyl-CoA
d)
1
module-2
0
0
module-3
0
“
O
w
/
/
OH
OH
o
H C02H
OH
module-4
OH
0
HO,C
266
II
oxvaenation
15. SECONDARYMETABOLISM I N MYXOBACTERIA
267
Table 1 Completely sequenced biosynthesis gene clusters identified from myxobacteria Compound
Producer
Type
Y
Reference(s) Pospiech et al., 1995, 1996 Schupp et al., 1995; Ligon et al., 2002 Silakowski et al., 1999 Silakowski et al., 2000 Julien et al., 2000; Molnar et al., 2000 Silakowski et al., 2001b Gaitatzis et al., 2002 Weinig et al., 2003 Feng et al., 2005 Sandmann et al., 2004 Carvalho et al., 2005; Kopp et al., 2005 Perlova et al., 2006b Wenzel et al., 200% Wenzel et al., 2006 Rachid et al., 2006a Simunovic et al., 2006 Meiser et al., 2006 Sandmann et al., 2007
Saframycin Soraphen
NRPS PIZS
M. xanthus DSM504I15 S. cellulosum So ce26
1995/1996 1995/2002
Myxothiazol Myxochelin Epothilone
PKSMRPS NRPS PKSMRPS
S. aurantiaca DW4/3-1 S. aurantiaca Sg a15 S. cellulosum So ce90
1999 2000 2000
Myxalamid Stigmatellin MelithiazoU Cystothiazol Tubulysin Disorazol
PKSMRPS PKS PKSMRPS PKSMRPS PIZSMRPS
S. aurantiaca Sg a15 S. aurantiaca Sg a15 Melittangium lichenicola Me 146 C. fuscus AJ-13278 Angiococcus disciformis An d48 S. cellulosum So ce12
2001 2002 2003 2005 2004 2005
Chivosazol Myxochromide S Myxochromide A Chondramide Myxovirescin DKxanthene Aurachin
PKSMRPS PKSMRPS PKSMRPS PKSINRPS PKSMRPS PKSMRPS PIZS/isoprenoid
S. cellulosum So ce56 S. aurantiaca DW4/3-1 M. xanthus DK1622 C. crocatus Cm c5 M . xanthus DK1622 M . xanthus DK1622 S. aurantiaca Sg a15
2005 2005 2006 2006 2006 2006 2006
nonproteinogenic amino acids can be generated and incorporated by these enzyme systems leading to almost endless possible combinations. Type I PKS, NRPS, hybrids thereof, and type I11 PKS have been identified in several gram-positive and gramnegative bacteria, and recently also type I1 PKSs have been found in myxobacteria (Sandmann et al., 2007).
Identification of Secondary Metabolite Biosynthetic Gene Clusters In the last 10 years 17 myxobacterial biosynthetic gene clusters have been identified (Table 1);some of them were
even reported twice (Julien et al., 2000; Molnar et al., 2000; Tang et al., 2000; Weinig et al., 2003; Carvalho et al., 2005; Feng et al., 2005; Kopp et al., 2005). In general, the biosynthesis gene cluster of a secondary metabolite is identified by generating a cosmid or bacterial artificial chromosome library, which is then screened using gene probes based on the significant similarity of PKS or NRPS genes. However, for myxobacteria which carry numerous different biosynthesis gene clusters (Silakowski et al., 2001a; BodeandMiiller, 2005)this approach results in the identification of a high number of positive hits, which leaves the desired gene cluster unidentified.
Figure 4 Mechanisms of bacterial PKSs and NRPSs and examples of resulting natural products. Post-PKS and/or NRPS modifications are highlighted in gray. (a) Generalized example of a type I PKS consisting of noniteratively acting domains. Abbreviations: AT, acyltransferase; ACP, acyl carrier protein; KS, ketosynthase; KR, ketoreductase; DH, dehydratase; ER, enoylreductase; an a or a p inside the circles indicates the specificity of AT domains for malonylCoA or methylmalonyl-CoA, respectively (Staunton and Weissman, 2001). (b) Biosynthesis of erythromycin A is a well-studied example of the action of a type I PKS, 6-deoxyerythronolide B synthase (DEBS) (Staunton and Weissman, 2001). (c) Type I1 PKS consisting of iteratively acting subunits as exemplified for the biosynthesis of tetracenomycin C: KS,, ketosynthase KS,, chain length factor (Shen, 2000). (d) Generalized example of NRPS biosynthesis: A, adenylation (the specificity of A domains is indicated according to standard amino acid abbreviations); T, peptidyl carrier protein; C, condensation; E, epimerization; HC, heterocyclization; Ox, oxidation (Sieber and Marahiel, 2005). ( e ) Formation of glycopeptide antibiotics like vancomycin provides well-documented examples of NRPS-directed biosynthesis, which are then further modified to give the active compounds (Recktenwald et al., 2002).
STRUCTUREAND METABOLISM
268
targeted approach
random approach
- construction of transposon library
- amplify PCR fragments of all gene clusters - cloning of these fragments into integrative plasmids
- disruption of all gene clusters
screening / \
via HPLClMS
I
fl
DW413-1 wild type
via bioassay
myxothiazol
I
myxochrornide
aurafuron
10
30
20
I
40
1
Next Page
15. SECONDARY METABOLISM IN MYXOBACTERIA An alternative strategy is a targeted approach that requires the construction of knockout mutants in the producing strain, the success of which cannot be taken for granted (Kopp et al., 2004). Initially, as many distinct fragments of specific PKS or NRPS genes as possible are obtained using degenerate PCR. The resulting fragments are then used to generate insertions in the corresponding gene clusters, and the obtained mutants are compared to the wild type with respect to the production of the analyzed compound by use of bioassays or high-throughput chromatography coupled to mass and/or UV spectroscopy (see also chapter 19). This approach even enabled the correlation of biosynthetic gene clusters of unknown function to completely novel secondary metabolites produced under the conditions used for cultivation as was described for aurafuron and myxochromid in S. aurantiaca Sg a15 (Kunze et al., 2005; Wenzel et al., 2005b) (Fig. 5 , left). An even more randomized approach is the construction of a transposon library followed by an appropriate screening of the mutants. However, if this approach cannot be coupled to a simple bioassay, several thousand mutants have to be analyzed using analytical chemistry methods. The gene cluster encoding the core biosynthetic machinery of the potent cytotoxin tubulysin was identified by this approach (Sandmann et al., 2004). After transposon library construction, four tubulysin-negative mutants were identified out of 1,200 tested strains by using a very sensitive and highly specific cell nucleus fragmentation assay. In this case, no HPLC or thin-layer chromatography-based screening would have been useful because of the very low amount of tubulysin produced (less than 1 mgliter), whereas as little as 50 ng/ ml can be detected using the bioassay. Finally, transposon recovery and sequencing allowed the identification of the gene cluster in a previously generated cosmid library (Fig. 5, right). Another alternative strategy is the heterologous expression of the constructed cosmid or bacterial artificial chromosome library in suitable hosts followed by a screening for the production of the desired compound. This strategy has been used in some metagenomic
269
approaches (Ferrer et al., 2005; Lorenz and Eck, 2005). There are several pitfalls in using this method with myxobacterial gene clusters due to differences in codon usage, promoter activity, substrate supply, and the length of the required DNA fragment, which are discussed in more detail in “Heterologous Expression. . .” (also see Wenzel and Muller, 2005).
Molecular Biology and Biochemistry of Myxobacterial Secondary Metabolites One goal of modern natural product research is the targeted modification of selected secondary metabolites in order to obtain compounds with altered and superior properties (e.g., better or more-specific biological activity, stability, and water solubility). The progress in understanding the biochemical principles of the enzymes involved in biosyntheses leads to achievements towards this goal by manipulation of the corresponding genes, a process which is called combinatorial biosynthesis (Walsh, 2002; Weissman and Leadlay, 2005). The huge diversity of known secondary metabolites derived from only a few different types of biochemical reactions implies that Nature is in fact already employing a similar approach during the course of evolution, indicating that it may be possible to speed up the process in the laboratory. Other methods that have been used to alter structures in vivo are feeding techniques using slightly modified precursors which are added to the culture broth of wildtype strains (precursor-directed biosynthesis) (Thiericke and Rohr, 1993) or mutants unable to produce the natural precursor (mutasynthesis) (Weist and Siissmuth, 2005). Although these methods have not been used extensively for myxobacterial compounds, the unique features of several secondary metabolites from myxobacteria make them attractive targets for these approaches. Since the first molecular biological studies on myxobacterial secondary metabolism in S. cellulosum (Schupp et al., 1995), M . xanthus (Pospiech et al., 1995; Paitan et al., 1999), and S. aurantiaca (Beyer et al., 1999; Silakowski et al., 1999) were published, almost every publication described highly unusual genetic and biochemical features (Wenzel and Muller, 2005b). Studies on the
Figure 5 Identification of myxobacterial biosynthesis gene clusters which allows the correlation between compounds and genes. Loss of production is easily detected by analytical methods (Silakowski et al., 2001a) or in compound-specific bioassays (Sandmann et al., 2004). Genomic DNA carrying different gene clusters responsible for the production of known and unknown compounds is simplified by a line. Extracts are made from mutants obtained via the targeted or the random approach and are either screened directly via HPLC/MS or used in a bioassay. In the example shown the destruction of the tubulin network is assayed after incubation of eukaryotic cells with the extracts followed by fixation and immunofluorescene microscopy of the cells.
Previous Page
2 70 biosynthesis of stigmatellin in S. aurantiaca Sg a15 revealed the first example of an exclusively iterative acting module in a type I PKS (Gaitatzis et al., 2002; Moss et al., 2004), whereas studies dealing with the biosynthesis of myxochromides in S. aurantiaca DW4/3-1 and M . xanthus DK1622 showed the skipping of a complete NRPS module which had never been observed before (Wenzel et al., 2005b, 2006). The understanding of the underlying mechanisms is the prerequisite to use these and other biochemical principles in a targeted manner in the future. Besides its interesting biological activity as an acetylCoA carboxylase inhibitor, soraphen from S. cellulosum is one of the few natural products in general that use benzoyl-CoA as a starting unit resulting in a phenylsubstituted macrolide (Bedorf et al., 1993; Hill et al., 2003a). Phenyl substituents are interesting starting points for chemical synthesis in order to modify structure and/or properties. Consequently, the biosynthetic genes for soraphen were identified (Schupp et al., 1995; Ligon et al., 2002). A DNA fragment encoding the loading module of the soraphen PKS was fused with elongation modules from an actinomycete gene cluster resulting in the production of novel phenyl-substituted products after feeding of benzoic acid to the newly constructed strain (Wilkinson et al., 2001). In addition, feeding of chemically modified benzoic acid derivatives led to the formation of the expected final compounds in the same system (Garcia-Bernard0 et al., 2004) and in the original producer (Hill and Thompson, 2003). In vitro studies also play an important role in determining enzyme specificity and promiscuity, leading to information which can be used to generate modified analogues of selected secondary metabolites. Exemplarily, the Walsh group (O’Connor et al., 2003) generated several nonnatural starting units of epothilone by incubation of various activated thioesters with the required recombinant enzymes (Fig. 6). However, it needs to be shown whether the observed broad in vitro promiscuity can be used to obtain modified epothilones in vivo. In vitro experiments with purified enzymes have also been used to confirm or elucidate complete biochemical pathways involved in myxobacterial secondary metabolism. The iron siderophore myxochelin was synthesized in vitro after production of the recombinant enzymes in Escherichia coli and incubation with the required substrates resulting in direct evidence for the postulated biosynthetic mechanism (Silakowski et al., 2000) including an unusual reductive chain termination mechanism (Gaitatzis et al., 2001) (Fig. 7a and b). Additionally, another unprecedented chain termination mechanism and methyl ester formation in the
STRUCTURE AND METABOLISM biosynthesis of myxothiazol and melithiazol has been confirmed in vitro and in vivo using mutants that are deficient in the formation of the required enzymes (Miiller and Muller, 2006; Miiller et al., 2006). The terminal amide found in both structures is derived from glycine extension and subsequent oxidation which results in the formation of an unstable intermediate that releases the amide upon degradation. In vitro studies show that the methyl ester found in myxothioazol Z and melithiazol is then made by MelJ-catalyzed hydrolysis of the corresponding amide to the free acid and subsequent methylation by the 0-methyltransferase MelK (Fig. 7c). An unusual cyclization domain was postulated to be involved in the formation of the chromone ring moiety of stigmatellin (Gaitatzis et al., 2002). Therefore, the terminal enzyme lacks a thioesterase domain and hydrolysis of the thioester bond occurs during the cyclization leading to an aromatic chromone (Fig. 7d).
Biosynthesis of Unusual Precursors for Myxobacterial Secondary Metabolites The branched-chain carboxylic acids isovalerate, isobutyrate, and 2-methylbutyrate are common building blocks for various bacterial secondary metabolites, as well as iso- or anteiso-fatty acids. According to textbook biochemistry they are incorporated as their CoAthioesters to start the biosynthesis of either fatty acids or PKs or PK/NRP hybrids as in the case of avermectin from S. avermitilis or myxothiazol and myxalamide in different myxobacteria. These thioesters are derived from the degradation of the branched-chain amino acids leucine, valine, and isoleucine, which are transaminated to the corresponding a-ketocarboxylic acids (Michal, 1999). Subsequently, these intermediates are degraded by the branched-chain keto acid dehydrogenase (BKD) complex, giving rise to the mentioned thioesters. In streptomycetes, disruption of the genes encoding the BKD complex results in a complete loss of all three branched-chain carboxylic acids and therefore also results in the loss of all compounds derived thereof (secondary metabolites, iso- and anteiso-fatty acids) (Hafner et al., 1991). However, disruption of the bkd locus in myxobacteria leads only to a reduction of isovalerylCoA (IV-CoA)-dependent compounds, which was first shown for 211. xanthus (Toal et al., 1995). Detailed analysis of the bkd mutants of 211. xanthus and S. aurantiaca revealed the presence of an alternative biosynthetic pathway to IV-CoA as the reason for the described phenomenon. This pathway branches from the wellknown mevalonate-dependent isoprenoid biosynthesis pathway and is induced under vegetative conditions in bkd mutants only (Mahmud et al., 2002,2005) (Fig. 8 ) .
15. SECONDARY METABOLISM IN MYXOBACTERIA
271
b)
a) EpoA
EpoB
EpoC
cxi:l)7&&)
NADPH MMCoA
n
R~<&s-Epoc
+
EpoC NADPH MMCoA
EpoB
HN ,
Ep°C
X = S, R = Me X = OIS, R = Ph X = S, R = 2-OH-Ph
EpoC
X
I
R
lhydrolysis
X = OIN, R = Me, R'= H X = 0, R = Me, R'= Me
I
R
R'= H X = OIS, R = Me X = OIS, R = CH,NH, X = 0, R = CH(CH,), X = S, R = CH,-NH-Boc
Figure 6 Biosynthesis of epothilone intermediates with alternate starter units. (a) Natural start of epothilone biosynthesis (Molnar et al., 2000). (b) Generation of alternative starting units using EpoC and different N-acetylcysteamine thioesters (SNAC) (which serve as substrate mimics of the intermediary enzyme-bound thioesters) of heterocyclic carboxylic acids. Products were detected bound to EpoC (Hicks et al., 2004). (c) Similar work using EpoA, EpoB, and EpoC. In this case the products were detected after hydrolysis from the biosynthetic enzyme (Schneider et al., 2002). Abbreviations: MMCoA, methylmalonyl-CoA; Boc, tButyloxycarbony1; ER, domains for enoylreductase; HC, heterocyclization; Ox, oxidation; for other abbreviations, see the legend to Fig. 4. Reprinted from the Journal of Industrial Microbiology and Biotechnology (Bode and Miiller, 2006) with kind permission of the publisher (0 Springer-Verlag, Berlin, Germany).
Additionally, feeding experiments performed under various conditions indicated that the alternative pathway is also activated during fruiting body formation (Bode et al., 2006) when cells are starving for nutrients and especially for leucine. Therefore, this novel and alternative pathway found in myxobacteria might serve as a backup to maintain the IV-CoA concentrations required
for development. The reason for the existence of this backup mechanism might be that 70 to 75% of all fatty acids in 111. xanthus are derived from IV-CoA and that these fatty acids seem to be required for development (Downard and Toal, 1995; Toal et al., 1995). Moreover, a spore-specific compound which is also derived from IV-CoA is produced as the main lipid during fruiting
STRUCTUREAND METABOLISM
272 a)
MxcE
MxcF
MxcG
?@ SH
SH
OH
Myxochelin A
OH
OH
R = see (Silakowski et al. 2001b) b)
R' = see (Weinig et al. 2003)
MxaA
R" = see (Silakowski et al. 1999) R"'= see (Gaitatzis et al. 2002) + NAD(P)H
c) MelFlMtaF
* L
MelGlMtaG
gMe P S
$H
MeOi~~ RIR' MeOIa.
OMe
RIR'
R'
4
StiJ
15. SECONDARYMETABOLISM IN MYXOBACTERIA body formation (Ring et al., 2006). Consequently, bkd mutants are deficient in aggregation and sporulation, which led to their alternative description as esg mutants because it was thought that they are deficient in the production of the E-signal, which itself is postulated to be an iso-fatty acid (Downard et al., 1993; Downard and Toal, 1995). Other unusual starter moieties used in the biosynthesis of myxobacterial natural products are benzoate and cinnamate, which are actually regarded as typical plant metabolites (Bode and Miiller, 2003) and which are described in more detail in chapter 19.
Heterologous Expression of Complex Biosynthetic Gene Clusters as a Tool To Access the Diversity of Myxobacterial Secondary Metabolites A genetic system is the limiting factor for identification and manipulation of secondary metabolite gene clusters in many organisms, including myxobacteria. Myxobacteria often grow slowly, with doubling times between 16 and 24 h; they are naturally multiresistant against most commonly used antibiotics; and often genetic methods established for one strain cannot be applied to others, even if they are phylogenetically closely related (Kopp et al., 2004). Given these disadvantages and the fact that gene clusters for secondary metabolites can easily reach sizes of 80 to 100 kb, manipulation of the natural producer and heterologous expression are challenging tasks. However, with myxobacterial strain collections available (for example, see the DSMZ website at www.dsmz. de) and given the potency of the produced compounds, there is an urgent need to access the genomic potential of these bacteria on the molecular level. A preferred way to reach this goal would be to establish an efficient method of cloning the desired gene clusters and their subsequent heterologous expression in a suitable host (Fig. 9). The epothilone gene cluster was identified and analyzed by scientists at NOVARTIS
2 73
(Molnar et al., 2000) and KOSAN biosciences (Julien et al., 2000; Tang et al., 2000) in parallel. The latter group cloned the biosynthetic genes onto two plasmids and expressed them in S. coelicolor (Tang et al., 2000). Although the construction of the expression plasmids included several rounds of cloning and the resultant strain produced epothilone only in trace amounts (50 &liter compared to 20 mg/liter in the original natural producer), this work showed that expression of myxobacterial gene clusters in other hosts is possible in general. Similar results were obtained for the production of soraphen A from S. cellulosum So ce26 in Streptomyces lividans ZX7 (Zirkle et al., 2004). Here a productivity of one-tenth of the amount made by the natural producer (0.3 mg/liter) was achieved, but the cloning procedures were even more laborious and time-consuming. In order to obtain better production titers of epothilone, the KOSAN group used M. xanthus as expression system and stepwise integrated the gene cluster for epothilone biosynthesis into the chromosome (Julien and Shah, 2002). The initial production titers were much better than those in S. coelicolor (0.3 mg/liter) and could be improved to the level of the original natural producer by optimization of the fermentation conditions (Lau et al., 2002). However, it is noteworthy that in the meantime the titer in the natural producer has been increased dramatically by classical mutagenesis and media optimization similar to the production of soraphen (from 0.3 mg/liter to 1.5 g/liter) (Gerth et al., 2003; see also chapter 19). Furthermore, the KOSAN group constructed an epothilone-producing M . xunthus mutant in which the cytochrome P450-dependent epoxidase epoK gene was inactivated, which resulted in the exclusive production of the reduced epothilones C and D instead of the epoxide forms A and B, respectively (Julien and Shah, 2002). The advantages of M . xanthus as host for heterologous expression of other myxobacterial biosynthetic gene
Figure 7 Unusual chain termination mechanisms found in myxobacteria. Active domains are shown in gray. (a) The biosynthesis of myxochelin as confirmed by in vitro studies (Gaitatzis et al., 2001). (b) A similar mechanism was postulated for the biosynthesis of the myxalamids (Silakowski et al., 2001b). The reduction of the thioester to the aldehyde might also take place in a two-step mechanism with an enzyme-bound 0,s-hemiacetal as intermediate. (c) Postulated final steps in the biosynthesis of myxothiazol (Silakowski et al., 1999) and melithiazol (Weinig et al., 2003) are in agreement with results from recent in vitro studies (Muller et al., 2006). (d) Postulated chromone ring formation in stigmatellin biosynthesis. Acetate and propionate units as determined from labeling experiments are shown in bold (Gaitatzis et al., 2002). Abbreviations: IC, isochorismate synthase; ArCP, aryl carrier protein; Red, reduction; Mox, monooxygenase; TE, thioesterase; for other abbreviations, see the legend to Fig. 4. Reprinted from the Journal of Industrial Microbiology and Biotechnology (Bode and Miiller, 2006) with kind permission of the publisher (0 Springer-Verlag, Berlin, Germany).
STRUCTUREAND METABOLISM
2 74
0
acetyl-CoA
acetoacetyl-CoA
3-hydroxy-3-methylglutaryl-CoA
3-methylbut-3-enoyl-CoA
mevalonate
dimethylallylpyrophosphate
I uSC~A 3,3-dimethylacryly I-CoA
iso-odd fatty acids secondary metabolites .c-(e.g. myxothiazol)
L O P P isopentenylpyrophosphate
isoprenoids isovaleryl-CoA
COZH leucine
2- ketoisocaproate
Figure 8 Alternative pathway to IV-CoA branching from the mevalonate-dependent isopre-
noid biosynthesis. clusters are quite obvious: the codon usage and the physiology are very similar, the cluster-specific promoters are likely to be active in M. xanthus, and posttranslational activation of the PKS or NRPS modules should be very efficient (Wenzel and Muller, 2005a). However, no plasmids are available and all required genes have to be integrated into the chromosome. Additionally, the doubling time of 4 to 5 h is still slow compared to that of other bacteria, and 111. xanthus has a strong tendency to produce high levels of ammonium during growth, making fermentations difficult to perform (Gerth et al., 2003).
A systematic search for fast-growing myxobacteria at the GBF (Gesellschaft fur Biotechnologische Forschung) led to the isolation of several moderately thermophilic strains belonging to almost all known species which are characterized by doubling times of less than 2 h (Gerth and Miiller, 2005). These strains are currently being developed as expression hosts for selected gene clusters (0.Perlova, K. Gerth, and R. Miiller, unpublished data). Pseudomonads have recently been shown to be an attractive alternative to myxobacteria as heterologous hosts (Wenzel et al., 2005). They can grow almost as
15. SECONDARYMETABOLISM IN MYXOBACTERIA
275
3)
Cosmid 1
Insertion of oriT and Pseudornonas gene fragment (trpf)
PseudornonasputiddCMch37
2
I
Completion of genec
Promotor exchange and deletion of nonessentialgenes 4
Icm-xyls-pmI
Final construct (CMch37)
Figure 9 Heterologous expression of myxobacterial biosynthesis gene clusters exemplified for myxochromides from S. uuruntiuca DW4/3-1 (Wenzelet al., 2005a). (1)Construction of cosmid library; (2)identification and analysis of the desired gene cluster; (3) “stitching” of the cluster and construction of the expression construct CMch37, using Red/ET technology; (4) transformation of the final cluster into P. putidu; and (5)production of myxochromide after induction.
fast as E. coli, they show a codon usage very similar to that of myxobacteria, and plasmids harboring inducible promoters are available. Furthermore, it was shown that PKSs and NWSs are efficiently posttranslationally
activated by intrinsic phosphopantetheinyl transferases in different Pseudomonas sp. (Gross et al., 2005). The remaining obstacles in using pseudomonads as expression hosts are the required exchange of the regulatory elements
2 76 and the enormous size of the biosynthetic gene clusters, which would obviously hamper their efficient cloning and modification. Both problems have been solved recently using RedET recombineering, a restrictionenzyme-independent cloning technique that allows the reassembly (“stitching”) of a complete gene cluster from different cosmids plus the exchange of the promoter(s) as shown for myxochromide S production in Pseudomonas putida (Wenzel et al., 2005a). The simplified process is shown in Fig. 9. Whereas the natural production in S. aurantiaca is only 8 mg/liter after 7 days of fermentation, more than 40 mg/liter has been obtained from P. putida after 2 days of growth. Additionally, this comparably high titer resulted in the production of previously unknown derivatives. Furthermore, the heterologous expression of this biosynthetic gene cluster unambiguously showed that the PKS is acting iteratively and that NWS module 4 is skipped during the biosynthesis, a process that has not been described previously (Fig. 9; see also Wenzel et al., 2006). Additional heterologous production in P. putida has been obtained for myxothiazol, which is normally produced by S. aurantiaca. In these experiments, genes encoding the production of the required extender unit methylmalonyl-CoA had to be coexpressed in the heterologous host (Gross et al., 2006b; Perlova et al., 2006a). Furthermore, a “silent” gene cluster responsible for the biosynthesis of an unknown product in S. cellulosum So ce56 was expressed in P. putida, resulting in the production of flaviolin, a pigment known from bacteria and fungi, which is derived from a type I11 PKS (Gross et al., 2006a) (see also chapter 19). The powerful combination of advanced cloning techniques and an advantageous expression host (which is currently far from being optimized) will allow the production of several interesting myxobacterial secondary metabolites in the future. It might even allow the production of new compounds directly from environmental DNA samples, eliminating the time-consuming or sometimes impossible isolation of pure cultures of any kind of putative producer of natural products.
Regulation of Myxobacterial Secondary Metabolism and Outlook The previous sections show some examples of secondary metabolites from myxobacteria and provide information about the elucidated underlying principles of their biosynthesis. Whereas some knowledge has been gained regarding this area of research, almost nothing is known about the regulation of secondary metabolite production in myxobacteria. In streptomycetes and fungi secondary metabolites are often produced at the
STRUCTUREAND METABOLISM onset of the stationary growth phase and are therefore regarded as a defense mechanism to allow progression of the life cycle and protection of the produced spores (Chater and Bibb, 1997).This seems to be true for some myxobacterial compounds as well (see “Secondary Metabolism in M. xanthus” above). However, several myxobacterial compounds known today are produced during vegetative growth even at low cell densities (our unpublished results). Because of their unusual lifestyle as organisms preying on bacteria and fungi, most of the known secondary metabolites from myxobacteria might be involved in killing prey, which is then made accessible as a nutrient source by different exoenzymes that are released by the cells (Petit and Guespin-Michel, 1992). However, the produced amounts of some secondary metabolites might be too low for detection under vegetative or developmental conditions, and therefore, the long-term goal must be to understand the regulation of myxobacterial secondary metabolite formation in order to increase or induce their production. One example of a study dealing with the regulation of secondary metabolite formation is the identification of a positive regulator of stigmatellin biosynthesis in Cystobacter fuscus. After transposon mutagenesis, 1,200 mutants have been screened using bioassays enabling the detection of decrease or loss of either argyrin or stigmatellin biosynthesis. Several mutants were identified showing a strongly reduced or even completely abolished stigmatellin production. In addition to the identification of the stigmatellin biosynthesis gene cluster, the novel positive regulator StiR related to sigmatellin production was identified after transposon recovery from the chromosome of a mutant (Rachid et al., 2006b). StiR shows some similarity to two-component sensor histidine kinases, and its role in regulating stigmatellin biosynthesis was confirmed by reconstruction of a genotypically identical mutant by double-crossover experiments. However, the regulatory cascade involving StiR has not yet been elucidated in detail. In other bacteria several two-component regulatory systems have been described as being involved in secondary metabolite production (Sola-Landa et al., 2003), and interestingly the number of this type of genes is comparably high in M. xanthus and S. cellulosum, the two sequenced myxobacterial model strains. A second example of a study dealing with the regulation of secondary metabolite formation is related to the positive regulator ChiR required for chivosazol biosynthesis in S. cellulosum. Using the putative chivosazol promoter region as bait, several DNA-binding proteins were isolated after incubation with a crude protein extract of
15. SECONDARY METABOLISM IN MYXOBACTERIA S . cellulosum So ce56. Mass spectrometric analysis of these proteins revealed several expected DNA-binding proteins such as RNA polymerases but also some regulator proteins. The corresponding genes were identified in the genome of strain So ce56 and disrupted, and the
277
resulting mutants were characterized phenotypically (Rachid et al., 2007). A >2O-fold reduction in the transcription of chivosazol biosynthesis genes could be observed, and hardly any chivosazol was found in a chiR mutant (Fig. 10a). Additionally, no fruiting body
200
6
+Growth +wt
+chiR-
9
150 -
5
- chiR+++
_1
4
5
g 1 100 -
8
0
/
Q
50
3 n
-I
0
0
200
100
Time
Oh
b)
300
400
of incubation [h]
24h
72h
h
Figure 10 Effect of chiR disruption (chiR-)and overexpression (chiR+++)on chivosazol production (a) and fruiting body formation in S. cellulosum So ce.56 (b).
2 78 formation could be observed in this mutant (Fig. lob). Overexpression of ChiR in S. cellulosum So ce56 did not influence development of the strain but led to a significant overproduction of chivosazol (Fig. 10a). This indicates that ChiR is pleiotropically involved in regulation of secondary metabolism and development in S. cellulosum as was already described for several regulators in streptomycetes (Sprusansky et al., 2003,2005). Myxobacterial functional genomics has only just started with the recently finished genome sequences of M . xanthus and S . cellulosum (see also chapter 19). Furthermore, the genomes of S . aurantiaca, which has been sequenced to a fourfold coverage (www.tigr.org; see chapter 18) and Anaeromyxobacter dehalogenans (genome.jgi-psf.org) are available. In the future, it is assumed that this information in combination with the possibility to clone whole biosynthesis gene clusters into optimized expression hosts will greatly speed up myxobacterial natural product research.
References Austin, M. B., and J. P. Noel. 2003. The chalcone synthase superfamily of type I11 polyketide synthases. Nat. Prod. Rep. 20:79-110. Bedorf, N., D. Schomburg, K. Gerth, H. Reichenbach, and G. Hofle. 1993. Isolation and structure elucidation of soraphen A,, a novel antifungal macrolide from Sorangium cellulosum. Liebigs Ann. Chem. 1993:1017-1021. Bentley, S. D., K. F. Chater, A. M. Cerdeno-Tarraga, G. L. Challis, N. R. Thomson, K. D. James, D. E. Harris, M. A. Quail, H. Kieser, D. Harper, A. Bateman, S . Brown, G. Chandra, C. W. Chen, M. Collins, A. Cronin, A. Fraser, A. Goble, J. Hidalgo, T. Hornsby, S. Howarth, C. H. Huang, T. Kieser, L. Larke, L. Murphy, K. Oliver, S. O’Neil, E. Rabbinowitsch, M. A. Rajandream, K. Rutherford, S. Rutter, K. Seeger, D. Saunders, S. Sharp, R. Squares, S. Squares, K. Taylor, T. Warren, A. Wietzorrek, J. Woodward, B. G. Barrell, J. Parkhill, and D. A. Hopwood. 2002. Complete genome sequence of the model actinomycete Streptomyces coelicolor A3(2). Nature 417:141-147. Beyer, S., B. Kunze, B. Silakowski, and R. Miiller. 1999. Metabolic diversity in myxobacteria: identification of the myxalamid and the stigmatellin biosynthetic gene cluster of Stigmatella aurantiaca Sg a15 and a combined polyketide(po1y)peptide gene cluster from the epothilone producing strain Sorangium cellulosum So ce90. Biochim. Biophys. Acta 1 4 4 5 185-195. Bird, C., J. Lynch, F. Pirt, W. Reid, and C. Brooks. 1971. Steroids and squalene in Methylococcus capsulatus grown on methane. Nature 230:473-474. Bode, H. B., B. Bethe, R. Hofs, and A. Zeeck. 2002. Big effects from small changes: possible ways to explore nature’s chemical diversity. Chembiochem 3:619-627. Bode, H. B., H. Irschik, S. C. Wenzel, H. Reichenbach, R. Miiller, and G. Hofle. 2003a. The leupyrrins: a structurally unique family of secondary metabolites from the
STRUCTUREAND METABOLISM myxobacterium Sorangium cellulosum. J. Nut. Prod. 66:1203-1206. Bode, H. B., and R. Miiller. 2003. Possibility of bacterial recruitment of plant genes associated with the biosynthesis of secondary metabolites. Plant Physiol. 132:1153-1161. Bode, H. B., and R. Miiller. 2005. The impact of bacterial genomics on natural product research. Angew. Chem. Int. Ed. Engl. 44:6828-6846. Bode, H. B., and R. Muller. 2006. Analysis of myxobacterial secondary metabolism goes molecular. J. Ind. Microbiol. Biotechnol. 33577-58 8. Bode, H. B., M. W. Ring, G. Schwar, R. M. Kroppenstedt, D. Kaiser, and R. Miiller. 2006.3-Hydroxy-3-methylglutarylcoenzyme A (CoA) synthase is involved in the biosynthesis of isovaleryl-CoA in the myxobacterium Myxococcus xanthus during fruiting body formation. J. Bacteriol. 188:65246528. Bode, H. B., S. C. Wenzel, H. Irschik, G. Hofle, and R. Miiller. 2004. Unusual biosynthesis of leupyrrins in the myxobacterium Sorangium cellulosum. Angew. Chem. Int. Ed. Engl. 43:4163-4167. Bode, H. B., B. Zeggel, B. Silakowski, S. C. Wenzel, H. Reichenbach, and R. Miiller. 2003b. Steroid biosynthesis in prokaryotes: identification of myxobacterial steroids and cloning of the first bacterial 2,3(S)-oxidosqualene cyclase from the myxobacterium Stigmatella aurantiaca. Mol. Microbiol. 47:471-481. Bretscher, A. P., and D. Kaiser. 1978. Nutrition of Myxococcus xanthus, a fruiting myxobacterium. J. Bacteriol. 133:763768. Burchard, R. P., A. C. Burchard, and J. H. Parish. 1977. Pigmentation phenotype instability in Myxococcus xanthus. Can. J. Microbiol. 23:1657-1662. Carvalho, R., R. Reid, N. Viswanathan, H. Gramajo, and B. Julien. 2005. The biosynthetic genes for disorazoles, potent cytotoxic compounds that disrupt microtubule formation. Gene 359:91-98. Chater, K. F., and M. J. Bibb. 1997. Regulation of bacterial antibiotic production p. 149-182. In H.-J. Rehm and D. W. Reed (ed.), Biotechnology. VCH, Mannheim, Germany. Dewick, P. M. 2002. The biosynthesis of C5-C25 terpenoid compounds. Nut. Prod. Rep. 19:181-222. Dickschat, J. S., H. B. Bode, T. Mahmud, R. Miiller, and S. Schulz. 2005. A novel type of geosmin biosynthesis in myxobacteria. /. Org. Chem. 705174-5182. Dickschat, J. S., S. C. Wenzel, H. B. Bode, R. Muller, and S. Schulz. 2004. Biosynthesis of volatiles by the myxobacterium Myxococcus xanthus. Chembiochem 5:778-787. Downard, J., and D. Toal. 1995. Branched-chain fatty acids: the case for a novel form of cell-cell signalling during Myxococcus xanthus development. Mol. Microbiol. 16:171-175. Downard, J., S. V. Ramaswamy, and K. S. Kil. 1993. Identification of esg, a genetic locus involved in cell-cell signaling during Myxococcus xanthus development. ]. Bacteriol. 175:7762-7770. Dworkin, M., and S. M. Gibson. 1964. A system for studying microbial morphogenesis: rapid formation of microcysts in Myxococcus xanthus. Science 146:243-244.
15. SECONDARY METABOLISM IN MYXOBACTERIA Elnakady, Y., F. Sasse, H. Liinsdorf, and H. Reichenbach. 2004. Disorazol A,, a highly effective antimitotic agent acting on tubulin polymerization and inducing apoptosis in mammalian cells. Biochem. Pharmacol. 62927-935. Feng, Z., J. Qi, T. Tsuge, Y. Oba, T. Kobayashi, Y. Suzuki, Y. Sakagami, and M. Ojika. 2005. Construction of a bacterial artificial chromosome library for a myxobacterium of the genus Cystobacter and characterization of an antibiotic biosynthetic gene cluster. Biosci. Biotechnol. Biochem. 69:1372-1380. Ferrer, M., F. Martinez-Abarca, and P. N. Golyshin. 2005. Mining genomes and ‘metagenomes’ for novel catalysts. Curr. Opin. Biotechnol. 16588-593. Fudou, R., T. Iizuka, S. Sato, T. Ando, N. Shimba, and S. Yamanaka. 2001. Haliangicin, a novel antifungal metabolite produced by a marine myxobacterium. 2. Isolation and structure elucidation. J. Antibiot. 54:153-156. Funa, N., Y. Ohnishi, I. Fujii, M. Shibuya, Y. Ebizuka, and S. Horinouchi. 1999. A new pathway for polyketide synthesis in microorganisms. Nature 4005397-899. Gaitatzis, N., B. Kunze, and R. Miiller. 2001. In vitro reconstitution of the myxochelin biosynthetic machinery of Stigmatella aurantiaca Sg a15: biochemical characterization of a reductive release mechanism from nonribosomal peptide synthetases. Proc. Natl. Acad. Sci. USA 98:11136-11141. Gaitatzis, N., B. Silakowski, B. Kunze, G. Nordsiek, H. Blocker, G. Hofle, and R. Miiller. 2002. The biosynthesis of the aromatic myxobacterial electron transport inhibitor stigmatellin is directed by a novel type of modular polyketide synthase.]. Biol. Chem. 277:13082-13090. Garcia-Bernardo, J., L. Xiang, H. Hong, B. S. Moore, and P. F. Leadlay. 2004. Engineered biosynthesis of phenyl-substituted polyketides. Chembiochem 5:1129-1131. Gerth, K., and R. Miiller. 2005. Moderately thermophilic myxobacteria: novel potential for production of natural products. Environ. Microbiol. 7:874-8 80. Gerth, K., S. Pradella, 0. Perlova, S. Beyer, and R. Miiller. 2003. Myxobacteria: proficient producers of novel natural products with various biological activities-past and future biotechnological aspects with the focus on the genus Sorangium. J. Biotechnol. 106:233-253. Goldman, B. S., W. C. Nierman, D. Kaiser, S. C. Slater, A. S. Durkin, J. Eisen, C . M. Ronning, W. B. Barbazuk, M. Blanchard, C. Field, C. Halling, G. Hinkle, 0. Iartchuk, H. S. Kim, C. Mackenzie, R. Madupu, N. Miller, A. Shvartsbeyn, S. A. Sullivan, M. Vaudin, R. Wiegand, and H. B. Kaplan. 2006. Evolution of sensory complexity recorded in a myxobacterial genome. Proc. Natl. Acad. Sci. USA 103:15200-15205. Gronewold, T. M., F. Sasse, H. Lunsdorf, and H. Reichenbach. 1999. Effects of rhizopodin and latrunculin B on the morphology and on the actin cytoskeleton of mammalian cells. Cell Tissue Res. 295:121-129. Gross, F., D. Gottschalk, and R. Miiller. 2005. Posttranslational modification of myxobacterial carrier protein domains in Pseudomonas sp. by an intrinsic phosphopantetheinyl transferase. Appl. Microbiol. Biotechnol. 68:66-74. Gross, F., N. Luniak, 0. Perlova, N. Gaitatzis, H. JenkeKodama, K. Gerth, D. Gottschalk, E. Dittmann, and R. Miiller. 2006a. Bacterial type I11 polyketide synthases:
2 79
phylogenetic analysis and potential for the production of novel secondary metabolites by heterologous expression in pseudomonads. Arch. Microbiol. 185:28-38. Gross, F., M. W. Ring, 0.Perlova, J. Fu, S. Schneider, K. Gerth, S. Kuhlmann, F. Stewart, Y. Zhang, and R. Miiller. 2006b. Metabolic engineering of Pseudomonas putida for methylmalonyl-CoA biosynthesis to enable complex heterologous secondary metabolite formation. Chem. Biol. 13:12531264. Hafner, E. W., B. W. Holley, K. S. Holdom, S. E. Lee, R. G. Wax, D. Beck, H. A. McArthur, and W. C. Wernau. 1991. Branched-chain fatty acid requirement for avermectin production by a mutant of Streptomyces avermitilis lacking branched-chain 2-0x0 acid dehydrogenase activity. J. Antibiot. 44:349-356. Heath, R. J., and C. 0. Rock. 2002. The Claisen condensation in biology. Nut. Prod. Rep. 19581-596. Hicks, L. M., S. E. O’Connor, M. T. Mazur, C. T. Walsh, and N. L. Kelleher. 2004. Mass spectrometric interrogation of thioester-bound intermediates in the initial stages of epothilone biosynthesis. Chem. Biol. 11:327-335. Hill, A., B. Thompson, J. Harris, and R. Segret. 2003. Investigation of the early stages in soraphen A biosynthesis. Chem. Commun. (Cambridge)2003:1358-1359. Hill, A. M., and B. L. Thompson. 2003. Novel soraphens from precursor directed biosynthesis. Chem. Commun. (Cambridge) 2003:1360-1361. Hofle, G., and H. Reichenbach. 2005. Epothilone, a myxobacterial metabolite with promising antitumor activity, p. 413450. In G. M. Cragg, D. G. Kingston, and D. J. Newman (ed.), Anticancer Agents from Natural Products. Taylor & Francis, Boca Raton, FL. Iizuka, T., Y. Jojima, R. Fudou, A. Hiraishi, J. W. Ahn, and S. Yamanaka. 2003a. Plesiocystis pacifica gen. nov., sp. nov., a marine myxobacterium that contains dihydrogenated menaquinone, isolated from the Pacific coasts of Japan. Int. J. Syst. Evol. Microbiol. 53:189-195. Iizuka, T., Y. Jojima, R. Fudou, M. Tokura, A. Hiraishi, and S. Yamanaka. 2003b. Enhygromyxa salina gen. nov., sp. nov., a slightly halophilic myxobacterium isolated from the coastal areas of Japan. Syst. Appl. Microbiol. 26:189-196. Iizuka, T., Y. Jojima, R. Fudou, and S. Yamanaka. 1998. Isolation of myxobacteria from the marine environment. FEMS Microbiol. Lett. 169:3 17-322. Ikeda, H., J. Ishikawa, A. Hanamoto, M. Shinose, H. Kikuchi, T. Shiba, Y. Sakaki, M. Hattori, and S. Omura. 2003. Complete genome sequence and comparative analysis of the industrial microorganism Streptomyces avermitilis. Nut. Biotechnol. 21526-531. Irschik, H., R. Jansen, K. Gerth, G. Hofle, and H. Reichenbach. 1995. Chivosazol A, a new inhibitor of eukaryotic organisms isolated from myxobacteria. J. Antibiot. (Tokyo) 48~962-966. Irschik, H., W. Trowitzsch-Kienast, K. Gerth, G. Hofle, and H. Reichenbach. 1988. Saframycin M x l , a new natural saframycin isolated from a myxobacterium. J. Antibiot. (Tokyo) 41:993-998. Jansen, R., H. Irschik, H. Reichenbach, and G. Hofle. 1997. Antibiotics from gliding bacteria, LXXX. Chivosazoles A-F: novel antifungal and cytotoxic macrolides from Sorangium
280 cellulosum (Myxobacteria).Liebigs Ann. Chem. 1997:17251732. Jansen, R., B. Kunze, H. Reichenbach, and G. Hofle. 1996. Chondramides A-D, new cytostatic and antifungal cyclodepsipeptides from Chondromyces crocatus (myxobacteria): isolation and structure elucidation. Liebigs Ann. Chem. 1996:285-290. Jansen, R., B. Kunze, H. Reichenbach, and G. Hofle. 2000. Antibiotics from gliding bacteria LXXXVI, Apicularen A and B, cytotoxic 10-membered lactones with a novel mechanism of action from Chondromyces species (myxobacteria): isolation, structure elucidation, and biosynthesis. Eur. J. o r g . Chem. 6:913-919. Jansen, R., G. Reifenstahl, K. Gerth, H. Reichenbach, and G. Hofle. 1983. Antibiotika aus Gleitenden Bakterien, XV: Myxalamide A, B, C und D, eine Gruppe homologer Antibiotilta aus Myxococcus xanthus M x x12 (Myxobacterales). Liebigs Ann. Chem. 7:1081-1095. Julien, B., and S. Shah. 2002. Heterologous expression of epothilone biosynthetic genes in Myxococcus xanthus. Antimicrob. Agents Chemother. 46:2772-2778. Julien, B., S. Shah, R. Ziermann, R. Goldman, L. Katz, and C. Khosla. 2000. Isolation and characterization of the epothilone biosynthetic gene cluster from Sorangium cellulosum. Gene 249:153-160. Knaggs, A. R. 2003. The biosynthesis of shikimate metabolites. Nut. Prod. Rep. 20:119-136. Kohl, W., A. Gloe, and H. Reichenbach. 1983. Steroids from the myxobacterium Nannocystis exedens. J. Gen. Microbiol. 129:1629-1635. Kopp, M., H. Irschik, F. Gross, 0. Perlova, A. Sandmann, K. Gerth, and R. Miiller. 2004. Critical variations of conjugational DNA transfer into secondary metabolite multiproducing Sorangium cellulosum strains So ce12 and So ceS6: development of a mariner-based transposon mutagenesis system. 1. Biotechnol. 10E29-40. Kopp, M., H. Irschik, S. Pradella, and R. Miiller. 2005. Production of the tubulin destabilizer disorazol in Sorangium cellulosum: biosynthetic machinery and regulatory genes. Chembiochem 6:1277-1286. Kunze, B., N. Bedorf, W. Kohl, G. Hofle, and H. Reichenbach. 1989. Myxochelin A, a new iron-chelating compound from Angiococcus disciformis (Myxobacterales).Production, isolation, physico-chemical and biological properties. J. Antibiot. (Tokyo) 42:14-17. Kunze, B., H. Reichenbach, R. Miiller, and G. Hofle. 2005. Aurafuron A and B, new bioactive polyketides from Stigmatella aurantiaca and Archangium gephyra (myxobacteria). J. Antibiot. (Tokyo) 58:244-251. Lau, J., S. Frykman, R. Regentin, S. Ou, H. Tsuruta, and l? Licari. 2002. Optimizing the heterologous production of epothilone D in Myxococcus xanthus. Biotechnol. Bioeng. 78:280-288. Laue, B. E., and R. E. Gill. 1995. Using a phase-locked mutant of Myxococcus xanthus to study the role of phase variation in development. J. Bacteriol. 177:4089-4096. Leibold, T., F. Sasse, H. Reichenbach, and G. Hofle. 2004. Cyrmenins, novel antifungal peptides containing a nitrogenlinked beta-methoxyacrylate pharmacophore: isolation and structural elucidation. Eur. J. Org. Chem. 2004:43 1-435.
STRUCTURE AND METABOLISM Ligon, J., S. Hill, J. Beck, R. Zirkle, I. Monar, J. Zawodny, S. Money, and T. Schupp. 2002. Characterization of the biosynthetic gene cluster for the antifungal polyketide soraphen A from Sorangium cellulosurn So ce26. Gene 285:257-267. Lorenz, P., and J. Eck. 2005. Metagenomics and industrial applications. Nut. Rev. Microbiol. 3510-516. Mahmud, T., H. B. Bode, B. Silakowski, R. M. Kroppenstedt, M. Xu, S. Nordhoff, G. Hofle, and R. Miiller. 2002. A novel biosynthetic pathway providing precursors for fatty acid biosynthesis and secondary metabolite formation in myxobacteria. 1. Biol. Chem. 277:32768-32774. Mahmud, T., S. C . Wenzel, E. Wan, K. W. Wen, H. B. Bode, N. Gaitatzis, and R. Miiller. 200.5. A novel biosynthetic pathway to isovaleryl-CoA in myxobacteria: the involvement of the mevalonate pathway. Chembiochem 6:322-330. Meiser, P., H. B. Bode, and R. Miiller. 2006. The unique DKxanthene secondary metabolite family from the myxobacterium Myxococcus xanthus is required for developmental sporulation. Proc. Natl. Acad. Sci. USA 103:19128-19133. Michal, G. 1999. Biochemical Pathways. Spektrum Akad. Verlag, Heidelberg, Germany. Molnar, I., T. Schupp, M. Ono, R. Zirkle, M. Milnamow, B. Nowak-Thompson, N. Engel, C. Toupet, A. Stratmann, D. D. Cyr, J. Gorlach, J. M. Mayo, A. Hu, S. Goff, J. Schmid, and J. M. Ligon. 2000. The biosynthetic gene cluster for the microtubule-stabilizing agents epothilones A and B from Sorangium cellulosum So ce90. Chem. Biol. 7:97-109. Moss, S. J., C. J. Martin, and B. Wilkinson. 2004. Loss of co-linearity by modular polyketide synthases: a mechanism for the evolution of chemical diversity. Nut. Prod. Rep. 211575-593. Miiller, I., and R. Miiller. 2006. Biochemical characterization of MelJ and MelK. FEBSJ. 273:3768-3778. Miiller, I., S. Weinig, H. Steinmetz, B. Kunze, S. Veluthoor, T. Mahmud, and R. Miiller. 2006. A unique mechanism for methyl ester formation via an amide intermediate found in myxobacteria. Chembiochem 7:1197-1205. Miiller, R. 2004. Don’t classify polyketide synthases. Chem. Biol. 11:4-6. Niggemann, J., M. Herrmann, K. Gerth, H. Irschik, H. Reichenbach, and G. Hofle. 2004. Tuscolid and tuscoron A and B: isolation, structural elucidation and studies on the biosynthesis of novel Furan-3(2H)-one-containingmetabolites from the myxobacterium Sorangium cellulosum. Eur. J. Org. Chem. 2004:487-492. O’Connor, S. E., C. T. Walsh, and F. Liu. 2003. Biosynthesis of epothilone intermediates with alternate starter units: engineering polyketide-nonribosomal interfaces. Angew. Chem. Int. Ed. Engl. 42:3917-3921. Omura, S., H. Ikeda, J. Ishikawa, A. Hanamoto, C. Takahashi, M. Shinose, Y. Takahashi, H. Horikawa, H. Nakazawa, T. Osonoe, H. Kikuchi, T. Shiba, Y. Sakaki, and M. Hattori. 2001. Genome sequence of an industrial microorganism Streptomyces avermitilis: deducing the ability of producing secondary metabolites. Proc. Natl. Acad. Sci. USA 98: 12215-12220. Paitan, Y., G. Alon, E. Orr, E. Z. Ron, and E. Rosenberg. 1999. The first gene in the biosynthesis of the polyketide antibiotic TA of Myxococcus xanthus codes for a unique PKS module coupled to a peptide synthetase. J. Mol. Biol. 286:465-474.
15. SECONDARYMETABOLISM IN MYXOBACTERIA Pearson, A., M. Budin, and J. J. Brocks. 2003. Phylogenetic and biochemical evidence for sterol synthesis in the bacterium Gemmata obscuriglobus. Proc. Natl. Acad. Sci. USA 100: 15352-15357. Perlova, O., J. Fu, S. Kuhlmann, D. Krug, F. Stewart, Y. Zhang, and R. Miiller. 2006a. Reconstitution of myxothiazol biosynthetic gene cluster by Red/ET recombination and heterologous expression in Myxococcus xanthus. Appl. Environ. Microbiol. 72:7485-7494. Perlova, O., K. Gerth, A. Hans, 0.Kaiser, and R. Miiller. 2006b. Identification and analysis of the chivosazol biosynthetic gene cluster from the myxobacterial model strain Sorangium cellulosum So ce56. J. Biotechnol. 121:174-191. Petit, F., and J. F. Guespin-Michel. 1992. Production of an extracellular milk-clotting activity during development in Myxococcus xanthus. J. Bacteriol. 1745136-5140. Piel, J. 2004. Metabolites from symbiotic bacteria. Nut. Prod. Rep. 2 1:5 19-53 8. Plaga, W., I. Stamm, and H. U. Schairer. 1998. Intercellular signaling in Stigmatella aurantiaca: purification and characterization of stigmolone, a myxobacterial pheromone. Proc. Natl. Acad. Sci. USA 95~11263-11267. Pospiech, A., J. Bietenhader, and T. Schupp. 1996. Two multifunctional peptide synthetases and an 0-methyltransferase are involved in the biosynthesis of the DNA-binding antibiotic and antitumour agent saframycin M x l from Myxococcus xanthus. Microbiology 142(Pt.4):741-746. Pospiech, A., B. Cluzel, H. Bietenhader, and T. Schupp. 1995. A new Myxococcus xanthus gene cluster for the biosynthesis of the antibiotic saframycin M x l encoding a peptide synthetase. Microbiology 141:1793-1803. Rachid, S., D. Krug, I. Kochems, B. Kunze, M. Scharfe, H. Blocker, M. Zabriski, and R. Muller. 2006a. Molecular and biochemical studies of chondramide formation-highly cytotoxic natural products from Chondromyces crocatus Cm c5. Chem. Biol. 13:667-681. Rachid, S., F. Sasse, S. Beyer, and R.Miiller. 2006b. Identification of StiR, the first regulator of secondary metabolite formation in the myxobacterium Cystobacter fuscus Cb f17.1. J. Biotechnol. 121:429-441. Rachid, S., K. Gerth, I. Kochems, and R. Miiller. 2007. Deciphering regulatory mechanisms for secondary metabolite production in the myxobacterium Sorangium cellulosum So ce56. Mol. Microbiol, 63:1783-1796. Recktenwald, J., R. Shawky, 0. Puk, F. Pfennig, U. Keller, W. Wohlleben, and S. Pelzer. 2002. Nonribosomal biosynthesis of vancomycin-type antibiotics: a heptapeptide backbone and eight peptide synthetase modules. Microbiology 148:1105-1118. Reichenbach, H. 2001. Myxobacteria, producers of novel bioactive substances. J. Ind. Microbiol. Biotechnol. 27:149156. Ring, M. W., G . Schwar, V. Thiel, J. S. Dickschat, R. M. Kroppenstedt, S. Schulz, and H. B. Bode. 2006. Novel isobranched etherlipids as specific markers of developmental sporulation in the myxobacterium Myxococcus xanthus. J. Biol. Chem. 268:36691-36700. Rosenbluh, A., and E. Rosenberg. 1989. Sporulation of Myxococcus xanthus in liquid shake flask cultures. J. Bacteriol. 171~4521-4524.
281
Sandmann, A., J. S. Dickschat, H. Jenke-Kodama, B. Kunze, E. Dittmann, and R. Miiller. 2007. Aurachin alkaloid biosynthesis in the Gram-negative myxobacterium Stigmatella aurantiaca: involvement of a type I1 polyketide synthase. Angew. Chem. Int. Ed. Engl. 46:2712-2716. Sandmann, A., F. Sasse, and R. Miiller. 2004. Identification and analysis of the core biosynthetic machinery of tubulysin, a potent cytotoxin with potential anticancer activity. Chem. Biol. 11:1071-1079. Sasse, F., B. Kunze, T. M. Gronewold, and H. Reichenbach. 1998. The chondramides: cytostatic agents from myxobacteria acting on the actin cytoskeleton. J. Natl. Cancer Inst. 90~1559-1563. Sasse, F., T. Leibold, B. Kunze, G. Hofle, and H. Reichenbach. 2003. Cyrmenins, new betamethoxyacrylate inhibitors of the electron transport. Production, isolation, physicochemical and biological properties. J. Antibiot. (Tokyo) 56~827-831. Sasse, F., H. Steinmetz, J. Heil, G. Hofle, and H. Reichenbach. 2000. Tubulysins, new cytostatic peptides from myxobacteria acting on microtubuli: production, isolation, physicochemical and biological properties. J. Antibiot. (Tokyo) 53: 879-8 85. Sasse, F., H. Steinmetz, G. Hofle, and H. Reichenbach. 1993. Rhizopodin, a new compound from Myxococcus stipitatus (myxobacteria) causes formation of rhizopodia-like structures in animal cell cultures. Production, isolation, physicochemical and biological properties. J. Antibiot. (Tokyo) 46~741-748. Schley, C., M. 0. Altmeyer, R. Swart, R. Muller, and C. G. Huber. 2006. Proteome analysis of Myxococcus xanthus by off-line two-dimensional chromatographic separation using monolithic poly-(styrene-divinylbenzene) columns combined with ion-trap tandem mass spectrometry. J. Proteome Res. 5:2760-2768. Schmidt, E. W., J. T. Nelson, D. A. Rasko, S. Sudek, J. A. Eisen, M. G. Haygood, and J. Ravel. 2005. Patellamide A and C biosynthesis by a microcin-like pathway in Prochloyon didemni, the cyanobacterial symbiont of Lissoclinum patella. Proc. Natl. Acad. Sci. USA 102:7315-7320. Schneider, T. L., C. T. Walsh, and S. E. O'Connor. 2002. Utilization of alternate substrates by the first three modules of the epothilone synthetase assembly line. J. Am. Chem. SOL. 124:11272-11273. Schulz, S., J. Fuhlendorff, and H. Reichenbach. 2004. Identification and synthesis of volatiles released by the myxobacterium Chondromyces crocatus. Tetrahedron 60:3 863-3 872. Schupp, T., C. Toupet, B. Cluzel, S. Neff, S. Hill, J. J. Beck, and J. M. Ligon. 1995. A Sorangium cellulosum (myxobacterium) gene cluster for the biosynthesis of the macrolide antibiotic soraphen A: cloning, characterization, and homology to polyketide synthase genes from actinomycetes. J. Bacteriol. 1723673-3679. Shen, B. 2000. Biosynthesis of aromatic polyketides, p. 1-53. In A. Meijere, K. Houk, H. Kessler, J. Lehn, S. Ley, S. Schreiber, and J. Thiem (ed.), Topics in Current Chemistry. Springer Verlag, Berlin, Germany. Sieber, S. A., and M. A. Marahiel. 2005. Molecular mechanisms underlying nonribosomal peptide synthesis: approaches to new antibiotics. Chem. Rev. 105:715-738.
282 Silakowski, B., B. Kunze, and R. Miiller. 2001a. Multiple hybrid polyketide synthase/nonribosomal peptide synthetase gene clusters in the myxobacterium Stigmatella aurantiaca. Gene 275:233-240. Silakowski, B., B. Kunze, G. Nordsiek, H. Blocker, G. Hofle, and R. Miiller. 2000. The myxochelin iron transport regulon of the myxobacterium Stigmatella aurantiaca Sg a15. Eur. J. Biochem. 267:6476-6485. Silakowski, B., G. Nordsiek, B. Kunze, H. Blocker, and R. Miiller. 2001b. Novel features in a combined polyketide synthasehon-ribosomal peptide synthetase: the myxalamid biosynthetic gene cluster of the myxobacterium Stigmatella aurantiaca Sgal5. Chem. Biol. 859-69. Silakowski, B., H. U. Schairer, H. Ehret, B. Kunze, S. Weinig, G. Nordsiek, P. Brandt, H. Blocker, G. Hofle, S. Beyer, and R. Miiller. 1999. New lessons for combinatorial biosynthesis from myxobacteria: the myxothiazol biosynthetic gene cluster of Stigmatella aurantiaca DW4/3-1. J. Biol. Chem. 274:37391-37399. Simunovic, V., J. Zapp, S. Rachid, D. Krug, P. Meiser, and R. Miiller. 2006. Myxovirescin biosynthesis is directed by an intriguing megasynthetase consisting of hybrid polyketide synthasestnonribosomal peptide synthetase, 3-hydroxy-3methylglutaryl CoA synthases and trans-acting acyltransferases. Chembiochem 21206-1220. Snyder, R. V., P. D. Gibbs, A. Palacios, L. Abiy, R. Dickey, J. V. Lopez, and K. S. Rein. 2003. Polyketide synthase genes from marine dinoflagellates. Mar. Biotechnol. (NY) 5:l-12. Soker, U., B. Kunze, H. Reichenbach, and G. Hofle. 2003. Dawenol, a new polyene metabolite from the myxobacterium Stigmatella aurantiaca. Z. Naturforsch. B 58:10241026. Sola-Landa, A., R. S. Moura, and J. F. Martin. 2003. The twocomponent PhoR-PhoP system controls both primary metabolism and secondary metabolite biosynthesis in Streptomyces lividans. Proc. Natl. Acad. Sci. U S A 100:6133-6138. Sprusansky, O., K. Stirrett, D. Skinner, C. Denoya, and J. Westpheling. 2005. The bkdR gene of Streptomyces coelicolor is required for morphogenesis and antibiotic production and encodes a transcriptional regulator of a branched-chain amino acid dehydrogenase complex. J. Bacteriol. 182664-671. Sprusansky, O., L. Zhou, S. Jordan, J. White, and J. Westpheling. 2003. Identification of three new genes involved in morphogenesis and antibiotic production in Streptomyces coelicolor. J. Bacteriol. 185:6 147-6 157. Staunton, J., and K. J. Weissman. 2001. Polyketide biosynthesis: a millennium review. Nut. Prod. Rep. 18:380-416. Tang, L., S. Shah, L. Chung, J. Carney, L. Katz, C. Khosla, and B. Julien. 2000. Cloning and heterologous expression of the epothilone gene cluster. Science 287:640-642. Thiericke, R., and J. Rohr. 1993. Biological variation of microbial metabolites by precursor-directed biosynthesis. Nut. Prod. Rep. 10:265-289.
STRUCTUREAND METABOLISM Toal, D. R., S. W. Clifton, B. A. Roe, and J. Downard. 1995. The esg locus of Myxococcus xanthus encodes the E l alpha and E l beta subunits of a branched-chain keto acid dehydrogenase. Mol. Microbiol. 16:177-189. Trowitsch, W., L. Witte, and H. Reichenbach. 1981. Geosmin from earthly smelling cultures of Nannocystis exedens (Myxobacterales).FEMS Microbiol. Lett. 12:257-260. Trowitzsch Kienast, W., K. Schober, V. Wray, K. Gerth, H. Reichenbach, and G. Hofle. 1989. Zur Konstitution der Myxovirescine B-T und Biogenese des Myxovirescins A. Liebigs Ann. Chem. 1989:345-355. Walsh, C. T. 2002. Combinatorial biosynthesis of antibiotics: challenges and opportunities. Chembiochem 3: 125-134. Weinig, S., H. J. Hecht, T. Mahmud, and R. Miiller. 2003. Melithiazol biosynthesis: further insights into myxobacterial PKS/NRPS systems and evidence for a new subclass of methyl transferases. Chem. Biol. 10:939-952. Weissman, K. J., and P. F. Leadlay. 2005. Combinatorial biosynthesis of reduced polyketides. Nut. Rev. Microbiol. 3: 925-936. Weist, S., and R. D. Sussmuth. 2005. Mutational biosynthesisa tool for the generation of structural diversity in the biosynthesis of antibiotics. Appl. Microbiol. Biotechnol. 68:141-150. Wenzel, S., and R. Miiller. 2005a. Recent developments towards the heterologous expression of complex bacterial natural product biosynthetic pathways. Curr. Opin. Biotechnol. 16594-606. Wenzel, S. C., F. Gross, Y. Zhang, J. Fu, F. A. Stewart, and R. Miiller. 2005a. Heterologous expression of a myxobacterial natural products assembly line in pseudomonads via red/ET recombineering. Chem. Biol. 12:349-356. Wenzel, S. C., B. Kunze, G. Hofle, B. Silakowski, M. Scharfe, H. Blocker, and R. Miiller. 2005b. Structure and biosynthesis of myxochromides S1-3 in Stigmatella aurantiaca: evidence for an iterative bacterial type I polyketide synthase and for module skipping in nonribosomal peptide biosynthesis. Chembiochem 6:375-385. Wenzel, S. C., P. Meiser, T. Binz, T. Mahmud, and R. Miiller. 2006. Nonribosomal peptide biosynthesis: point mutations and module skipping lead to chemical diversity. Angew. Chem. Int. Ed. 45:2296-2301. Wenzel, S. C., and R. Muller. 2005b. Formation of novel secondary metabolites by bacterial multimodular assembly lines: deviations from text book biosynthetic logic. Curr. Opin. Chem. Biol. 9:447-458. Wilkinson, C. J., E. J. Frost, J. Staunton, and P. F. Leadlay. 2001. Chain initiation on the soraphen-producing modular polyketide synthase from Sorangium cellulosum. Chem. Biol. 8:1197-1208. Zirkle, R., J. M. Ligon, and I. Molnar. 2004. Heterologous production of the antifungal polyketide antibiotic soraphen A of Sorangium cellulosum So ce26 in Streptomyces lividans. Microbiology 150:2761-2774.
Myxobacterial Genornics and Postgenornics
Myxobacteria: Multicellularity and Differentiation Edited by David E. Whitworth 0 2008 ASM Press, Washington, D.C.
Catherine M. Ronning William C. Nierman
The Genomes of Myxococcus xanthus and Stigmatella aurantiaca
Myxobacteria are unique among bacterial organisms in that they are able to move, or glide, without flagella; they form “social” multicellular hunting and feeding swarms; and in response to starvation, they form multicellular spore-forming fruiting body aggregates. This lifestyle, together with the ability to distinguish species by fruiting body morphology, implies a tightly coordinated, heritable system of signaling between individual cells. In this chapter we discuss and compare the genomic sequences of Myxococcus xanthus DK1622 (hereafter referred to as DK1622 or strain DK1622) and Stigmatella uurantiaca DW4/3-1 (hereafter referred to as DW4/3-1 or strain DW4/3-l), two related aerobic, fruiting-body-forming myxobacteria. The structure and complexity of the S. aurantiaca fruiting body are the primary characteristics distinguishing it from M. xanthus DK1622, as well as the production of the signaling pheromone stigmolone, which may be analogous to the M . xanthus DK1622 quorum-sensing A-signal. Having the genome sequence of both organisms will greatly facilitate research into these and other myxobacteria-specific phenotypic traits.
16
SEQUENCING AND FINISHING
M. xanthus DK1622 The M. xanthus DK1622 genome, initially sequenced to 4.5X coverage by Monsanto, was completed at The Institute for Genomic Research by additional random sequencing (Goldman et al., 2006). Genomic DNA was randomly sheared to create two libraries containing either small (-2-kb) or large (15-to 20-kb) inserts. After verification of the randomness of the libraries, each was subjected to high-throughput DNA sequencing of both ends, until 8X coverage was attained. Fragments were assembled using Celera Assembler (Myers et al., 2000). Sequence ambiguities and frameshifts were edited manually. The finished (or i.e., a single, ordered, contiguous DNA sequence of high quality and low error rate) genomjc sequence was then annotated to identify putative coding regions. Both auto- and manual annotation were performed on the closed genome. Predicted open reading frames (ORFs) were identified with the program GLIMMER (Salzberg et al., 1998). A number of other programs were utilized
Catherine M. Ronning and William C. Nierman, J. Craig Venter Institute, 9712 Medical Center Dr., Rockville, MD 20850.
285
MYXOBACTERIAL GENOMICS AND POSTGENOMICS
286 to identify putative genes, including BLAST-ExtendRepraze (http://ber.sourceforge.net), a modified SmithWaterman algorithm (Waterman, 1988) which aligns protein-protein matches; BLASTp similarity searches; BLASTx searches against a database of nonredundant bacterial proteins; and hidden Markov model (HMM) alignments constructed from PFAMs (Bateman et al., 2004) and TIGRFAMs (Haft et al., 2003). Outputs from these similarity searches were manually curated (Fraser et al., 1995), and role categories (Riley, 1993) were assigned. The DK1622 genome is composed of a single circular chromosome containing 9,139,763 bp (Table 1) (Goldman et al., 2006). The 7,380 identified genes represent a coding density of 91.1%, suggesting that the genome has undergone a long period of refined selection. Nearly one-half of the genes (48.9%) were assigned a putative function.
S. aurantiaca DW4/3-1 Whereas M. xanthus DK1622 was sequenced to full closure with manually curated annotation, S. aurantiaca DW4/3-1 is unfinished. The genome was sequenced with the whole-genome random sequencing method to 5 x coverage. Overlapping contigs and contigs linked by spanning clones with each of their end sequences in adjacent contigs were assembled into scaffolds (i.e., contigs were placed in correct order and orientation on the chromosome) but not fully closed (i.e., sequencinggaps remain). Autoannotation
using only informatics tools without manual review was then performed on the assembled scaffolds. The 5 X assembly of the strain DW4/3-1 genome consists of 61 scaffolds spanning 10.1 Mb and has a GC content of 67.1% (Table 1).The number of genes in the genome is predicted to be 8,586, with an average gene length of 1,098 nucleotides. Included in this number are 623 partial genes, i.e., genes whose 5' or 3' end extends beyond the end of the scaffold. Like M. xanthus DK1622, nearly one-half of the genes (48.1%) were assigned a putative function. The 5 X draft sequence of S. aurantiaca DW4/3-1 is of consequential value, and comparison to the gene content of M. xanthus DK1622 suggests that it provides good visibility of most of the genes. Identification of certain housekeeping genes, including all known ribosomal large subunits, tRNA ligases, and members of the isopentenyl diphosphate biosynthetic pathway, provides further evidence for this near completeness. In addition to the 623 partial genes, other genes are undoubtedly missing from the sequence. Based on the span of the scaffolds compared to the bases in the contigs in the scaffolds, -100 kb of sequence is entirely missing from the genome sequence within the scaffolds. There is no way to estimate the amount of sequence missing between the scaffolds. Other limitations derive from the early draft status of this genome sequence. Regions of the genome present within the sequence are represented by variable coverage, even though on average the coverage is 5 X I Low-coverage
Table 1 Selected features of the genomes of M. xanthus DK1622 and S. uuruntiucu DW4/3-1 Features ORFs Total no. of ORFs Assigned function Conserved hypothetical Unknown function Hypothetical proteins Unique proteins' Chromosomes Size (bp) No. of ORFs Average gene length (nt) GC content ei
M. xunthusa 7,380 3,608 (48.9%) 688 (9.3%) 1,119 (15.2%) 1,965 (26.6%) 1,566 (21.2%) 9,139,763 7,380 1,128 68.90%
S. auruntiacah 8,586 4,271 (48.1%) 913 (10.6%) 909 (10.6%) 2,450 (28.5%) 2,748 (32.0%) 10,158,519d 8,656 1,098 67.10%
8X coverage, manual annotation.
"X coverage, autoannotation. 'Number of proteins unique to one species relative to the other, based on reciprocal BLASTp analysis with cutoff value for P of s e - ' " . dTotal span of 61 scaffolds.
16. THEGENOMES OF hl. XANTHUS
AND
s. AURANTIACA
regions have lower sequence accuracy, confounding accurate gene annotation and PCR primer design. At the other end of the resolution scale, the fragmentation of the DW4/3-1 genome sequence into 61 scaffolds will eliminate the ability to perform chromosomescale structural comparisons. Comparative analysis of synteny relative to other myxobacterial genomes cannot be undertaken with such a fragmented genome sequence. Since the large chromosome size is a focus of study with these genomes, the DW4/3-1 sequence cannot be used in comparative studies of chromosome organization and stability in its present 5 X sequence coverage status. These limitations notwithstanding, most genes in S. aurantiaca DW4/3-1 could be identified using M. xanthus DK1622 as the reference genome since sequence identity between the two species is high. Thus, comparisons between the two genomes can be made but must be interpreted with caution.
COMPARATIVE GENOMICS Phylogeny Unlike most other prokaryotes, the myxobacteria exhibit social behavior and multicellular development. Such complex behavior is correlated with their large genomes. While fruiting body morphology has classically been the basis for species classification within the myxobacteria, the molecular classification methods of DNA sequencing and analysis have also been used. An analysis of the 16s rRNA sequence from 12 different myxobacteria showed that they form a monophyletic group within the purple bacteria 6 subdivision and that three distinct subgroups (Myxococcus, Chondromyces, and Nannocystis) are contained within the group (Shimkets and Woese, 1992). Classification by this method agreed with most morphological, behavioral, and metabolic differences between members but not with fruiting body complexity. The two fruiting-body-forming species whose genomes have been sequenced, M. xanthus DK1622 and S. aurantiaca DW4/3-1, lie within the same subgroup (Myxococcus) as determined by this method. Some of the features distinguishing this subgroup from the other two are the production of reddish-pigmented monocyclic carotenoid glucosides and structurally distinctive antibiotics (Shimkets and Woese, 1992). A more in-depth analysis of 16s rRNAs from 54 myxobacterial strains representing 21 species agreed with the trifurcation of the Myxococcales order (Sproer et al., 1999).
287 direct comparison inherently difficult; DK1622 has been completely closed and finished and manually curated, while DW4/3-1 has been sequenced only to 5X, assembled but not closed, and autoannotated. Bearing these limitations in mind, however, we can make some comparisons of gene content between the two and provide some inferences as to their function. Putative gene numbers and other statistics on the genomes are given in Table 1. These numbers have been discussed individually for the two species in the previous section. To identify putative orthologs between the two genomes, the 8,586 predicted DW4/3-1 proteins were compared against the 7,380 predicted DK1622 proteins by reciprocal BLASTp with a cutoff value for P of I e - 1 0 . One hundred eight of the predicted DK1622 proteins are partial (i.e., contain frameshifts and/or point mutations and thus could not be translated); these were analyzed using their nucleotide sequences and BLASTx. DK1622 was found to have 1,566 proteins (21.2%) that were unique relative to DW4/3-1 (Table l), 1,400 of which were annotated as hypothetical (annotation based solely on gene prediction program[s]) or conserved hypothetical (based on similarity to a hypothetical protein from a different organism) or were of unknown function; 2,748 (32.0%) of the DW4/3-1 proteins were unique, of which 1,845 fell into one of these three categories. Additionally in DW4/3-1 there are 54 unclassified genes, or genes for which a function could not be automatically assigned. It is these proteins of hypothetical or unknown function that are unique to one or the other species that may define the morphogenetic differences between the two species and which require further analysis and experimentation. The unusually large size of the M. xanthus DK1622 genome is reportedly due to expansion by lineage-specific duplications of specific categories of genes, particularly those involved in cell-cell signaling, small-molecule sensing, and multicomponent transcriptional control, allowing the evolution of the complex molecular machinery required for development of a multicellular lifestyle (Goldman et al., 2006). Since S. aurantiaca DW4/3-1 is similar in size to M . xanthus DK1622, we compared the numbers of genes in these specific roles. The analysis revealed that the relative abundance of such genes in DW4/3-1 is similar to that of DK1622, thus supporting the hypothesis of genome expansion by gene duplication in at least these two myxobacterial species.
Motility
Features of the Genornes of M. xanthus DK1622 and S. aurantiaca DW4/3-1 As mentioned previously, the different closure status of M. xanthus DK1622 and S. aurantiaca DW4/3-1 makes
Gliding motility has also been extensively studied in M. xanthus DK1622, which, rather than swimming, hunts by gliding over the soil surface as a coordinated aggregation of thousands of cells, an essential element
MYXOBACTERIAL GENOMICS AND POSTGENOMICS
288 of its complex lifestyle (Spormann, 1999). This S (social) motility system is regulated by several large gene clusters, including the che4 operon (Vlamakis et al., 2004); the type IV pili genes (Wu and Kaiser, 1995; Wu et al., 1997,1998; Wallet al., 1999);the dsp-diflocus (Lancer0 et al., 2002); and the sasA locus (formerly rfbABC), encoding the lipopolysaccharide 0-antigen biosynthesis genes (Guo et al., 1996; Bowden and Kaplan, 1998). Additionally, myxobacterial cells can move individually, as regulated by the adventurous (A)system (Hodgkin and Kaiser, 1979). There are at least 30 genes involved in A-motility (Youderian et al., 2003) as well as the “frizzy” chemosensory system implicated in single-cell reversals (McBride et al., 1989; McCleary et al., 1990; McCleary and Zusman, 1990; Trudeau et al., 1996; Ward et al., 2000). All of the M. xanthus DK1622 genes involved in Amotility except agmJ, which has similarity to carbohydrate kinases, and agmN, a hypothetical protein, have orthologs in S . aurantiaca DW4/3-1. Similarly, the Smotility genes and several other genes involved in gliding mostly have orthologs in DW4/3-1. The DK1622 dsp-dif locus is mostly present in DW4/3-1, with the exception of difG and 5 of the 21 genes identified within the locus (Table 2). difG, which is homologous to the Bacillus subtilis chemotaxis gene cheC (Black and Yang, 2004), is one of two genes involved in the difBG mechanism of self-recognition (Bonner et al., 2005). The five genes missing in DW4/3-1 are annotated as lipoprotein, putative (two genes), conserved hypothetical protein (two genes), and glyoxalase family protein in DK1622.
The M. xanthus DK1622 che3 locus, che4 operon, frizzy aggregation genes, type IV pili genes, and sasA locus appear to have been conserved in S. aurantiaca DW4/3-1 as well, with only a small rearrangement in the type IV pili gene cluster (Table 2). In contrast, only 21 of the possible 26 to 28 putative genes of the DK1622 exopolysaccharide (EPS) synthesis region and 1 of the 2 genes in the EPS-associated (EAS) region (Lu et al., 2005), which are essential for EPS biogenesis, a requirement for S-motility, have orthologs in DW4/3-1. Many of these putative orthologs have weak identities, and only seven are clustered. Three uncharacterized putative chemotaxis clusters were identified in DK1622 by the presence of cheA-like genes (Table 2). All have orthologs in DW4/3-1, where they are at least partially clustered.
Secondary Metabolite Production Myxobacteria feed by lysing cells of other bacteria and yeasts. Bioactive compounds synthesized and secreted through secondary metabolite biosynthetic gene clusters may aid predation and inhibit competition (Chater, 1989). Both M. xanthus DK1622 and S. aurantiaca DW4/3-1 are prolific producers of biologically active secondary metabolite compounds, the genes for which often occur in hybrid polyketide synthase/ nonribosomal peptide synthetase (PKS/NRPS) gene clusters (Silakowski et al., 2001a). We have identified 26 PKS genes, 24 NRPS genes, and 8 hybrid PKS/NRPS genes in the DK1622 genome, and 23 PKS genes, 18 NRPS genes, and 1 NRPS/PKS hybrid gene in DW4/
Table 2 Putative chemotaxis gene clusters identified in M. xanthus DK1622 and their S. aurantiaca DW4/3-1 orthologs, identified by the presence of cheA-like genes“ S. aurantiaca DW4l3-1
Loci
No. of orthologs/total no. of M. xanthus genes in cluster
MXAN-268-MXAN-2686 MXAN-413 8-MXAN-4150 M X AN-4 751-MXA N-4 759 MXAN-5144-MXAN-5155 M X AN-5 771-MXAN-58 04 MXAN-6012-MXAN-6033 M X AN-6683-MXAN-6711 M X AN-693 8-MXAN-6966
6/6 13/13 819 9/12 3 2/34 21/22 23/29 25/29
M. xanthus DK1622 Cluster che4 locusb “frizzy” locusc Putative chemotaxis cluster 1 che3 locusd Type IV pili gene cluster@ Putative chemotaxis cluster 2 dsp-dif locusf Putative chemotaxis cluster 3
“Orthologs were identified by reciprocal BLASTp with‘ I <e-Io. “Vlamakis et al., 2004. <McCleary et al., 1990; McCleary and Zusrnan, 1990; McBride et al., 1989; Trudeau et al., 1996. “Kirby and Zusrnan, 2003. ‘Wu et al., 1998; Wallet al., 1999. ‘Lancer0 et al.. 2002.
Clustered? Yes ( STIA U-4 791-STIA U-4 798 ) Yes (STIAU-6832-STIA U-6847) Yes (STIAU-5413-STIAU-5420) Yes (STIAU-1046-STIAU-1056) Partially (STIAU-7815-STIAU-7844) Partially (STIAU-8085-STIAU-8106) Partially (STIAU-8661-STIAU-8676) Partially (STIAU-0562-STIA U-0584)
16. THEGENOMES OF M .
XANTHUS AND
S. AURANTIACA
3-1. These genes were used to locate regions of putative secondary metabolite biosynthetic gene clusters on both genomes. Twelve such clusters have been identified in the DK1622 genome, and 14 have been identified in DW4/3-1 (Tables 3 and 4). Most genes within the DK1622 clusters have orthologs in DW4/3-1, but occur in clusters in the latter S. uuruntzuca DW4/3-1 (either wholly or partially) only five times (Table 3). Similarly, most genes contained within the 14 DW4/3-1 clusters have orthologs to genes in DK1622 but are clustered in only four DK1622 regions (Table 4). Of these, DK1622 cluster 4/DW4/3-1 cluster 10 and DK1622 cluster 9/DW4/3-1 cluster 4 are the only two pairs of putatively identified secondary metabolite biosynthetic gene clusters that appear to be orthologous (Tables 3 and 4). Secondary metabolite gene clusters are often species specific, e.g., clusters within various Aspergillus species exhibit little orthology (Nierman et al., 2005). This diversity in natural products among various soildwelling organisms may provide a unique set of chemical weapons, providing each species with a competitive edge in such an environment. Most of these putative secondary metabolite biosynthetic gene clusters, in both S . aurantiacu DW4/3-1 and M . xanthus DK1622, were identified through bioinformatics techniques; further research is critical to confirm the functionality of these clusters as well as to characterize their products. Secondary metabolite gene clusters have been most extensively studied in S. aurantiaca DW4/3-1, with the elucidation of the pathways and/or isolation of several
Table 3
289 biologically active compounds. Myxochromide S is encoded by a 30-kb cluster in DW4/3-1 containing three PKS and NRPS genes (Wenzel et al., 2005) (Table4, cluster 14). The DW4/3-1 myxothiazol biosynthetic gene cluster (Silakowski et al., 1999) (Table 4, clusters 2 and 9), which encodes an electron transport inhibitor, contains seven genes (mtaA through mtaG) that correspond strongly to five genes in M. xanthus DK1622, indicating that two genes may have become duplicated. DW4/3-1 mtaC and mtaD (STIAU-027.5 and STIAU-0276) are both highly similar to the gene encoding MXAN-4299, a hybrid NRPS/PKS ( P values, 4.1 e-13*and 0, respectively). Similarly, both DW4/3-1 mtuE and mtuF (STIAU-4029 and STIAU-4028) have identities to the DK1622 PKS gene MXAN-4527 (P values, 2.9 e-229 and 3.7 e-265, respectively).The fact that this single, published cluster is split among two different clusters at different locations in our analysis is another indication of the unfinished status of the S . aurantiaca DW4/3-1 genome. While the genes discussed above were first identified in the sequenced strain DW4/3-1, several secondary metabolite gene clusters have been elucidated using S. aurantiaca Sgal5. Three genes encoding the biosynthetic pathway of the quinoline antibiotic and electron transport inhibitor aurachin (aroA,,, , aroAA2,aroA,,), first identified in strain Sgal5 (Silakowski et al., 2000a), were annotated in strain DW4/3-1 (STIAU-7537, STIAU-4733, and STIAU-4992). aroAoo, is type I and uroAA2and u ~ o A A ,are type I1 3-deoxy-~-arabino-heptulosonate-7-phosphate (DAHP) synthase genes. S. aurun-
Secondary metabolite biosynthetic gene clusters in M. xanthus and putative orthology (if present) in S. aurantiaca
DW4/3- 1a S. aurantiaca DW4/3-1 ortholog M. xanthus DK1622 Cluster
Loci
No. of orthologsltotal no. of M. xanthus genes in cluster
Clustered? (loci)
10/18
Partially (STIAU-3537-STIAU-3544) No No Partially (STIAU-4986-STIA U-4999; cluster 10 below) Partially (STIAU-6538-STIA U-6572) No No Partially (STIAU-6757-STIA U-6764) Yes (STIAU-0873-STIAU-0897; cluster 4 below) No No No
1 2 3 4
MXAN-2 283-MXAN-1300 MXAN-1588-MXAN-162 6 MXAN-2792-MXAN-2 799 MXAN-3618-MXAN-3651
26/29 619 29/35
5 6" 7 8 9
MXAN-3 777-MXAN-3800 M X A N-3 93 1 -MXA N-3 953 M X A N-3 994-MXA N-4003 M X A N-4063-MXA N-4080 M X A N-4290-MXA N-43 0.5
21/24 18/23 5110 14/18 14/16
10 11 12
MXAN-4398-MXAN-4416 M X AN-4523-MXA N-4534 M X AN-4592-MXA N-4607
13/18 10/12 12/16
"Orthologs were identified by reciprocal BLASTp with a P cutoff value of <1.0e-l0, or by BLASTx in the case of partial proteins. "Antibiotic TA biosynthetic cluster, M . xanthus strain ER-15 (Paitan et al., 1999).
MYXOBACTERIAL GENOMICS AND POSTGENOMICS
290
Table 4 Secondary metabolite biosynthetic gene clusters in S. aurantiaca DW4/3-1 and putative orthology (if present) in M. xanthus" M. xantbus DK1662 ortholog S. aurantiaca DW4/3-1 Loci
No. of orthologsltotal no. of M. xanthus genes in cluster
1 2b 3 4
STIA U-0239-STIA U-0251 STIAU-0266-STIAU-0283 STIA U-0 734STIA U-0 750 STIA U-0867-STIA U-0897
8/12 14/18 15/16 26/30
5 6 7 8 9" 1O d
STIA U-1156-STIA U-1165 STIA U-1833STIA U-1869 STIA U-1968-STIAU-1 984 STIA U-2975-STIA U-2 986 STIA U-4026STIA U-4049 STIA U-4986STIA U-5000
5/10 32/37 12/17 7/12 16/24 11/15
11 12 13 14
STIA U-6385-STIA U-6404 STIA U-7278STIA U-7295 STIAU-73 1 7-STIA U-73 63 STIA U-8749STIA U-8757
Cluster
14/19 16/17 33/47 517
Clustered? (loci)
No No No Partially (MXAN-4290-MXAN-4305; cluster 9 above) No Partially (MXAN-3 51 3-MXA N-3 543) No No Partially (MXAN-0915-MXAN-0925) Partially (MXAN-3639-MXAN-3647; cluster 4 above) No No No No
"Orthologs were identified by reciprocal BLASTp with a P cutoff value of Sl.Oe-'", or by BLASTx in the case of partial proteins. 6Myxothiazol biosynthetic cluster, S. aurantiaca strain DW4/3-1 (Silakowski et al., 1999); possibly stigmolone (Plaga and Ulrich, 1999). <Myxothiazolbiosynthetic cluster, S. auruntiaca strain DW4/3-1 (Silakowski et al., 1999). dMyxochelin iron transport cluster, S. uurantiucu strain Sgal5 (Silakowski et al., 2000). <Myxochromide S, S . uurantiaca strain DW4/3-1 (Wenzel et al., 2005).
tiaca Sgal.5 aroAA5 and to a lesser extent aroA,, have one counterpart in M. xanthus DK1622, MXAN-3642 (3-deoxy-7-phosphoheptulonate synthase); the third S. aurantiaca Sgal5 gene, aroAool, was not found in M. xanthus DK1622. The S. aurantiaca Sgal5 steroid biosynthesis genes cas (cycloartenol synthase) and sqs (squalene synthase) (Bode et al., 2003) have been identified as STIAU-7491 and STIAU-7492, respectively, in strain DW4/3-1. Neither of the S. aurantiaca Sgal5 steroid biosynthesis genes was found in M. xanthus DK1622. The catecholate siderophore myxochelin iron transport cluster (mxcA-I,K,L),also described in strain Sgal.5 (Silakowski et al., 2000b) has been identified in strain DW4/3-1 (Table 4, cluster 10). This gene cluster also contains the aurachin biosynthetic gene aroA,, (STIAU4992). Most of this cluster is conserved in M. xanthus DK1622 (Table 3, cluster 4), which contains orthologs to all of the genes except mxcA. These partially clustered M. xanthus genes appear to be slightly rearranged relative to S. aurantiaca DW4/3-1. Some of the secondary metabolite gene clusters previously described in other strains were only partially identified in S . aurantiaca DW4/3-1, however. The 65kb aromatic electron transport inhibitor stigmatellin
biosynthetic cluster consists of 20 genes identified in Sgal5 (stiA-L, plus additional ORFs) (Gaitatzis et al., 2002). All of the sti genes except stiK had orthologs in strain DW4/31,as did four of the ORFs of unknown function (ORF1, 7, 8, and 9), for a total of 11putative orthologs in strain DW4/3-1. BLASTp analysis indicates that strain Sgal5 stiC, stiF, and stiH may have arisen as duplications of STIAU-0879 from DW4/3-1, and strain Sgal5 stiA, still, and stiE may be duplications of STIAU-7283. However, a more conservative and possibly more plausible explanation may be that these missing or duplicated strain Sgal.5 genes are within the estimated 100 kb or more of missing genome sequence from within or between strain DW4/31 scaffolds. Comparison of the strain Sgal5 stigmatellin gene cluster with M. xanthus DK1622 reveals that 14 of the 20 genes have orthologs in strain DK1622, but strain Sgal.5 stiC, stiF, and stiH appear to be duplications of PKS MXAN-4298, and stiB and sti] both exhibit strong similarity to MXAN-4527, also a PKS. Myxalamid, another electron transport inhibitor found in Sgal.5, is encoded by an 11-gene, combined PKS/NRPS cluster (mxaA-F and ORF1-4) (Silakowski et al., 2001b). All 11 genes had distinct orthologs in strain DW4/3-1, although two (ORF3 and ORF4) had much weaker identities. All have orthologs in
16. THEGENOMES OF M .
XANTHUS AND
S . AURANTIACA
M . xanthus DK1622, but mxaB2, mxaD, and mxaE are all similar to MXAN-4.528. Another interesting observation when comparing these two secondary metabolite clusters from strain Sgal5, stigmatellin and myxalamid, with M. xanthus DK1622 is that although in strain Sgal5 these are two distinct clusters composed of a total of 31 different genes, some of the orthologs in strain DK1622 are shared between both clusters; e.g., MXAN-4527 and MXAN-4528 are homologous to genes in both the myxalamid and stigmatellin biosynthetic pathways in strain SgalS. Since both compounds are electron transport inhibitors it would be interesting to see if and how SgalS has specialized to have evolved two separate genetic clusters from M . xanthus DK1622, or vice versa. The antibiotic gene cluster, saframycin M x l , has been identified in the M. xanthus strain DMS04/15 (Pospiech et al., 1995). Two of the three genes in the cluster, safA and safB, have orthologs in M. xanthus DK1622 (MXAN-3634 and MXAN-44 14, respectively), although the strain DK1622 safA ortholog contains a disrupted reading frame. Both ORFs are annotated as NRPSs and are in regions that also include several other NRPS and PKS genes. safC has a much weaker similarity to the strain DK1622 gene encoding MXAN-5983, an 0-methyltransferase family protein. M. xanthus DM504/1S safA and safB also exhibit significant similarity to STIAU-6395 and STIAU-1156, respectively. The former is an NRPS associated with myxochromide S biosynthesis in that species (Wenzel et al., ZOOS), while the latter is annotated as saframycin M x l synthetase B. As in strain DK1622, strain DM504/15 safC is only weakly similar to STIAU-5543, annotated as O-methyltransferase mdmC. M. xanthus DK1622 also produces the polyketide antibiotic TA, encoded by a gene cluster identified in M. xanthus ER-15 (Paitan et al., 1999) (Table 4). Orthologs of all genes have been identified in strain DK1622, although two (taB and taE) have only weak similarities. Additionally, gene order within the cluster appears to be conserved between the two strains. BLASTp analysis of the M. xanthus ER-15 gene cluster and S. aurantiaca DW4/3-1 revealed that four of the genes are missing in strain DW4/3-1 (taA, taB, taD, and taE) and four (taC, taF, taG, and taK) only weakly resemble the M. xanthus ER-15 counterparts. The strongest similarity lies between the putative regulator genes taR1, taR2, and taR3 (GenBank accession numbers AY376594 and AY388473, Y. Paitan) and the S. aurantiaca DW4/3-1 genes encoding STIAU-5987 (Tar1), STIAU-6643 (mutant Ntr C-like activator), and STIAU-6642 (metallo-beta-lactamase family protein), respectively.
291 The differences between the two S. aurantiaca strains in secondary metabolite genes may be due to the unfinished status of the DW4/3-1 genome. However, such species-to-species differences in secondary metabolism biosynthetic gene clusters have also been shown to occur in Aspergillus fumigatus (Nierman et al., 2005), and our findings with S. aurantiaca DW4/3-1 as well as M. xanthus DK1622 may be indicative of true genotypic variation between strains of these two species of myxobacteria. Such differences in secondary metabolite repertoire may provide each species and perhaps distinct strains within species with unique chemical weapons which may provide competitive advantage. Clearly more research is indicated.
Signaling and Chemotaxis in Development The myxobacteria have a large capacity for signal transduction, and chemotactic-like signal transduction has been extensively studied in M. xanthus DK1622. While the strain DK1622 genome encodes a plethora of signaling components, the density of one-component transcriptional regulators is relatively low compared to that of other soil bacteria (Goldman et al., 2006). Several such regulatory proteins are either missing (e.g., IclR, LacI, ROK, and DeoR) or significantly reduced in number (e.g., AraC, GntR, AsnC, and LuxR) in strain DK1622 (Goldman et al., 2006). Feeding and fruiting body development in the myxobacteria are coordinated by the regulatory A- and C-signal genes. The quorum-sensing A-signal senses the approach of starvation and induces cellular aggregation (Kaplan et al., 1991). In M. xanthus DK1622, A-signal production is controlled by five asg genes (asgA-E)that may function together in response to the nutritional state of the cell (Kaiser, 2004). These genes, M X A N 2670, MXAN-2913, MXAN-5204, MXAN-6996, and MXAN-1010, all have high identities to S. aurantiaca DW4/3-1 genes STIAU-4780, STIAU-7250, STIAU1092, STIAU-0817, and STIAU-8408 based on reciprocal best matches. The sasNlsasRlsasS genes then respond to the A-signal to regulate normal growth of the fruiting body (GUOet al., 2000). These three strain DK1622 genes (MXAN-1244, MXAN-1245, and MXAN-1249) also are orthologous to strain DW4/3-1 genes (STIAU3620, STIAU-3617, and STIAU-3603). As mentioned earlier, eight chemotaxis clusters have been putatively identified in M. xanthus DK1622, each containing a cheA-like gene (Table 2). The function of at least three of these clusters is yet to be ascertained. Orthologs to all of these clusters, complete or nearly so, have been found in S. aurantiaca DW4/3-1 (Table 2) and correspond to strain DW4/3-1 clusters also identified by
292 the presence of a cheA-like gene, suggesting that all of these chemotaxis-like clusters are critical in some way to the myxobacterial lifestyle. S. aurantiaca DW4/3-1 produces the pheromone stigmolone, a signal molecule that may be comparable in function to the quorum-sensing A-signal in M. xanthus DK1622 (Kaiser, 2004). Stigmolone has been purified, its activity as a cell aggregation trigger has been demonstrated (Plaga et al., 1998), and its structure has been determined (Hull et al., 1998). The genes involved in stigmolone biosynthesis and fruiting body formation, fbfAB C D (STIAU-0266 through STIA U-0269) were identified by complementation analysis with a mutant S. aurantiaca DW4/3-1 strain as a cluster (Plaga and Ulrich, 1999). Two of these, fbfA and f b p , had previously been sequenced and have homologies to chitin synthase and galactose oxidase, respectively (Silakowski et al., 1996, 1998).fbfC and fbfD are putatively identified as STIAU0267 and STIAU-0266 from the S. aurantiaca DW4/3-1 genome sequence, both annotated as hypothetical proteins. Three of the four M. xanthus DK1622 orthologs (MXAN-3482 through MXAN-3484) are clustered. After development is initiated, the species-specific shape of the fruiting body is determined by the morphogenetic C-signal, whose increase triggers the time-ordered expression of specific developmental genes (Gronewold and Kaiser, 2001). The extracellular C-signal is the product of the csgA gene (Kim and Kaiser, 1990), identified as MXAN-1294. An ortholog of csgA exists in S. aurantiaca DW4/3-1 (STIAU-3543). csgA is regulated by the act operon, four genes (actA-D) that create a positivefeedback loop to control the time course of csgA expression (Gronewold and Kaiser, 2001) (MXAN-3213 through MXAN-3217). The S. aurantiaca DW4/3-1 orthologs are STIAU-5047 through STIAU-505 1.These genes are likely to encode the DW4/3-1 C-signal biosynthetic genes. There are many other genes that are associated with fruiting body development and may be associated with the C-signal, including devRS (Thony-Meyer and Kaiser, 1993), fruA (Ueki and Inouye, 2003), frgABC, and the socA locus (Lee and Shimkets, 1994). All except devRS (discussed in “Development” below) have orthologs in S. aurantiaca DW4/3-1. Three more classes of genes have been shown to be essential for normal development, bsg, dsg, and esg, but since signal molecules for these have yet to be found they are referred to as inferred signal molecules (Kaiser, 2004). The lonD/bsgA gene, identified as MXAN-3993 (ATP-dependent protease La), is required for development and for intercellular signaling (Gill et al., 1993; Tojo et al., 1993). MXAN-0581 encodes a translation
MYXOBACTERIAL GENOMICS AND POSTGENOMICS initiation factor, IF-3, identified as dsg/infC (Cheng et al., 1994). The esg locus, encoding branched-chain keto acid dehydrogenase E l alpha and beta subunits, constitutes the fifth cell-cell signaling system in the myxobacteria (Downard et al., 1993; Toal et al., 1995). The two genes are identified as MXAN-4564 (alpha subunit) and MXAN-4565 (beta subunit). All have orthologs in S. aurantiaca DW4/3-1. Thus, the M . xanthus DK1622 genome contains evidence of an abundance of the intercellular and nutritional signals that coordinate feeding and fruiting body development, as well as allowing the cell to adjust its metabolism in response to the dense cellular environment within the fruiting body. All of the genes known to be involved in signaling of development in M. xanthus DK1622 that we examined had orthologs in S. aurantiaca DW4/3-1, with the exception of devRS, which may be a consequence of the unfinished status of strain DW4/3-1 annotation. The control of development in these two species is likely to be very similar based on the presence of DW4/3-1 orthologs to the genes controlling development in strain DK1622. Further experimentation will be required to determine if interspecific differences are due to as yet undiscovered genes or to subtle variation in the proteins encoded by these genes or in their regulation.
Gas Vesicle Genes It is interesting that a total of 10 putative gas vesicle proteins have been identified in the M . xanthus DK1622 genome (Table 5 ) . Nine of these proteins were identified by orthology to protein domains associated with gas vesicle proteins, and eight are localized within a cluster (MXAN-2765 through MXAN-2772). BLASTp analysis of the 14-gene gas vesicle protein cluster from Bacillus megaterium (Li and Cannon, 1998) identified a 10th putative gas vesicle protein in M. xanthus DK1622, MXAN-2257. Gas vesicles provide buoyancy and have been found mostly in aquatic prokaryotes, but also in some terrestrial species (van Keulen et al., 2005). Eight of the 14 proteins encoded by the B. megaterium gas vesicle cluster had weak similarity to seven strain DK1622 proteins. A minimal complement of eight gvp genes were found to be required for the formation of functional gas vesicles in halophilic archaea, and homologs to all eight were found to be present in B. megaterium (Offner et al., 2000). All but two of these eight necessary genes (gvpG and gvpR) had B. megaterium orthologs in strain DK1622. One putative ortholog was found in the S. aurantiaca DW4/3-1 genome (STIAU-2003), and possibly one other, weakly similar gene (STIAU-1144) (Table 5 ). It would be interesting to see if any of the DK1622 orthologs are functional, perhaps providing a survival
16. THEGENOMES OF M. Table 5
XANTHUS AND
S.
293
AURANTIACA
Gas vesicle genes identified in M. xanthus DK1622 and putative orthologs in S. aurantiaca DW4/3-1“
S. auralztiaca DW4/3-1 homologs
M. xanthus DK1622 Annotation
Locus
Annotation
P value
MXAN-1551 MXAN-2257
Gas vesicle protein, GvpUGvpF family FHA domain protein
STIA U-2003 STIA U-1144
C-gvpF Mutant NtrC-like activator, putative
5.50E-57 9.90E-20
MXAN-2765 MXAN-2766 MXAN-2767 MXAN-2768 MXAN-2 769 MXAN-2770 MXA N-2 771 MXA N-2 772
Gas vesicle protein, GvpA family Putative gas vesicle protein GvpK Gas vesicle protein, GvpA family Putative gas vesicle protein GvpG Gas vesicle protein, GvpL/GvpF family
STIA U-2003
C-gvpF
5.30E-20
Gas vesicle structural protein GvpA Gas vesicle protein, GvpL/GvpF family Gas vesicle protein, GvpL/GvpF family
STIA U-2003 STIA U-2003
C-€YPF C-gvpF
1.lOE-11 6.70E-20
Locus
“Orthologs were identified by reciprocal BLASTp with a P cutoff value of ~ 1 . 0 ~ ’ ~
mechanism to the organism in flooded soils or being involved in other cellular processes (Li and Cannon, 1998; van Keulen et al., 2005). The differential representation of these genes between DK1622 and DW4/3-1 may reflect specific adaptations to their respective ecological niches. Development Development has been extensively studied in M. xanthus DK1622. Of genes involved specifically in fruiting body development, no significant similarities were found to the M. xanthus DK1622 devRS genes in S. aurantiaca DW4/3-1. These two genes, which may be transcribed as an operon, are unique and essential for normal fruiting body development in strain DK1622, possibly acting as autoregulators controlling the timing of gene expression (Thony-Meyer and Kaiser, 1993). The DK1622 genes hthA, hthB, sdeK, and tps have homologs in strain DW4/3-1. The che3 gene cluster (Kirby and Zusman, 2003) appears to be orthologous to a cluster in DW4/3-1 (see“Signalingand Chemotaxis” above).Most of the other developmental genes studied, except the DK1622 fruE and pru genes, have orthologs in the DW4/3-1 genome. The FruE protein is unique to DK1622 and is required for normal development (Akiyama and Komano, 2003). pru encodes protein U, a secreted spore coat protein produced late in development (Gollop et al., 1991). The main feature distinguishing S. aurantiaca DW4/ 3-1 from M. xanthus DK1622 is the development and complexity of the fruiting body (Kaiser, 2004). Strain DW4/ 3-1 fruiting bodies are composed of a stalk, branches, and multiple cysts, in contrast to strain DK1622,which does not exhibit such structures. The DW4/3-1 sigma factor genes sigA (Skladny et al., 1994) (STIAU-1092), sigB (Skladny
et al., 1992) (STIAU-2459), and sigC (Coudart, 1998) (STIAU-7997) contribute to the specificintercellular interactions and biosynthesis of development-specific proteins involved in fruiting body morphogenesis. The RNA polymerases sigA and sigB have nearly identical counterparts in the M. xanthus DK1622 rpoD and sigB genes, with similarities of 93.6 and 95.9%, respectively. Two other DK1622 sigma factors involved in fruiting body development also have orthologs in DW4/3-1: DK1622 sigC is 90.8% similar to the DW4/3-1 rpoHgene, and DK1622 sigE is 94.2% similar to DW4/3-1 sigE. The DK1622 szgD gene is only 30.4% similarto aDW4/3-1 hypothetical protein; however, sigD is annotated as containing a frameshiftin this strain of M. xanthus. hspA (STIAU-651 l), which encodes the stressinduced low-molecular-weight heat shock protein SP2 1 that is expressed late in the morphogenesis of the S. auruntiaca DW4/3-1 fruiting body, is more similar to plant than to bacterial HSPs (Heidelbach et al., 1993). An ortholog to the DW4/3-1 hspA gene was also identified in M. xanthus DK1622 as MXAN-4268, annotated as a heat shock protein. Two genes involved in late (fbfA,STIAU-0268) (Silakowski et al., 1996)and early (fbp,STIAU-0269) (Silakowski et al., 1998)stalk formation are also implicated in the biosynthesis of the pheromone signaling compound stigmolone (discussed previously). Recently, a pyruvate kinase gene encoding indole binding protein 2 ( p y k A , STIAU-8263/STIAU-8264) was shown to be essential for fruiting body formation (Stamm et al., 2005). The carDSagene (STIAU-6095) exhibits orthology to the M. xanthus DK1622 high-mobility-group A-type transcription factor carD (Cayuela et al., 2003), which is involved in both light-induced carotenogenesis and
294 fruiting body formation (Nicolas et al., 1994) (also discussed in “Light-Induced Carotenogenesis” below). Deletion of carD in DK1622 is fully complemented by carD, (Cayuela et al., 2003). It is interesting that fruiting body morphogenesis in S. aurantiaca DW4/3-1, in contrast to strain DK1622, requires light (Qualls et al., 1978; Shimkets, 1990), particularly blue light between 400 and 500 nm (White et al., 1980), and that exposure of strain DW4/3-1 to light results in a distinct pattern of protein synthesis during development (Inouye et al., 1980).
Light-Induced Carotenogenesis Carotenoids accumulate in M. xanthus DK1622 upon illumination with blue light (Burchard and Hendricks, 1969) or exposure to copper (Moraleda-Munoz et al., 2005) and may act as a photoprotectant against photolysis by quenching singlet oxygen and other radicals (Rau, 1988). Three unlinked loci are involved in the light-induced synthesis and regulation of carotenoids in strain DK1622. The DK1622 carA-carB operons are present in a single cluster consisting of 11structural and regulatory genes (Botella et al., 1995) (MXAN-0894 through MXAN-0904). The six genes in the structural carB operon, crtEBDC and two of unknown function, all have orthologs in S. auruntiacu DW4/3-1 (STIAU8489 through STIAU-8494), while only two of the five carA operon genes, ORFlO (MXAN-0903) and ORF11 (MXAN-0904), have S. aurantiaca DW4/3-1 orthologs (STIAU-8485 and STIA U-8486, respectively). These two genes, both MerR-like transcriptional activators, may have resulted from duplication of an ancestral gene (Cervantes and Murillo, 2002). A third carA gene, ORF8 (MXAN-0901) has some similarity to STIAU-8487. Therefore, while the entire structural carB operon appears to be preserved in S. aurantiaca DW4/3-1, the carA locus is missing two or three genes of unknown function. Carotenoid pigments were observed in S. aurantiaca DW4/ 3-1 nearly 40 years ago (Kleinigand Reichenbach, 1969; Kleinig et al., 1970), but to our knowledge these are the only reports of pigment development in Stigmatella. A second locus, the carQRS locus, regulates lightinducible carotenoid biosynthesis (McGowan et al., 1993) (MXAN-4088 through MXAN-4090). Orthologs of two of the three genes were found in S. aurantiaca DW4/31 (carQ, STIAU-3827, and carR, STIAU-3826), while there was no ortholog to cars. carQ and carR encode positive and negative regulators, respectively, of the lightinducible promoter encoded by the carQRS locus and are essential for its expression, while the cars gene, which activates the carB promoter, is not required (McGowan et al., 1993).
MYXOBACTERIAL GENOMICS AND POSTGENOMICS The third locus is a single gene, crtl, another structural gene encoding phytoene dehydrogenase (formerly designated carC) which is involved in an early step in the carotenoid pathway (Fontes et al., 1993). crtI has been identified as MXAN-7517 and STIAU-0585. Several other genes involved with light induction or perception have been identified in M. xanthus DK1622 and have orthologs in S. aurantiaca DW4/3-1, including carF (MXAN-5750; STIA U-7779), which inactivates carR (Fontes et al., 2003), and ihfA (MXAN-3596; STIAU-1959), encoding the a subunit of the integration host factor protein, which is required for the blue-light response (Moreno et al., 2001), as well as carD, which in addition to its blue-light response also acts in response to starvation to form fruiting bodies (discussed in “Development” above) and is similar to eukaryotic high-mobility group A transcription factors (Galbis-Martinez et al., 2004) (MXAN-5622). A carD ortholog has been identified in S. aurantiaca DW4/3-1 (STIAU-6095) (Cayuela et al., 2003). In addition to the production of carotenoid pigments as photoprotectants, M. xanthus DK1622 also contains a gene encoding DNA photolyase (phrA;MXAN-4136) which acts by repairing pyrimidine dimers resulting from ultraviolet irradiation (Sancar, 1990). The strain DK1622 phrA gene was found to be more similar in amino acid sequence to eukaryotic photolyases than to those of other eubacteria, suggesting a very ancient evolutionary divergence of the two forms of this gene (O’Connor et al., 1996). The S. aurantiaca DW4/3-1 ortholog is STIAU-2132.
CONCLUSIONS Myxobacteria are unique in the prokaryotic world in that they engage in “social” behavior involving the interaction and cooperation of thousands of individual cells, requiring a complex communication network. Some of this complexity may be reflected in the unusually large genome sizes of M. xanthus DK1622 and S. aurantiaca DW4/3-1. Indeed, it has been suggested that at least part of the genome expansion in strain DK1622 is due to nonrandom, lineage-specific duplications ( Goldman et al., 2006), and this hypothesis is supported by analysis of these particular genes in strain DW4/3-1. Many similarities between these two genomes exist, including genes for chemotaxis, signaling, and motility, and genes involved in development (except the critical DK1622 devRS operon) and carotenogenesis. Similarly, many of the secondary metabolite genes in both DK1622 and DW4/3-1 are orthologous; however, the cluster structure of these genes is often disrupted in the other species. The major
16. THEGENOMES OF 211.
XANTHUS AND
S. AURANTIACA
morphological difference between DK1622 and DW4/31 is the complexity of the fruiting body. Some but not all of the genes identified as necessary for DK1622 fruiting body development have orthologs in DW4/3-1 , leaving open speculation into the genetic mechanisms of regulation. Finally, we found no DW4/3-1 genes orthologous to the DK1622 gas vesicle operon, nor did we identify any DK1622 genes resembling the two DW4/3-1 steroid biosynthesis genes. In this chapter we have attempted to use wholegenome sequence data and comparative genome analysis from two species of myxobacteria, M. xanthus DK1622 and S. aurantiaca DW4/3-1, to provide insight into such mechanisms. The information here and in the genome sequence of these two strains of myxobacteria is a starting point for researchers to further study the complex genetic and environmental interactions inherent in this very intriguing bacterial lifestyle.
References Akiyama, T., and T. Komano. 2003. Analysis of fruE, a novel developmental gene of Myxococcus xanthus. J. Mol. Microbiol. Biotechnol. 6:164-173. Bateman, A., L. Coin, R. Durbin, R. D. Finn, V. Hollich, S. Griffiths-Jones, A. Khanna, M. Marshall, S. Moxon, E. L. Sonnhammer, D. J. Studholme, C. Yeats, and S. R. Eddy. 2004. The Pfam protein families database. Nucleic Acids Res. 32(Database issue):D138-Dl41. Black, W. P., and Z. Yang. 2004. Myxococcus xanthus chemotaxis homologs DifD and DifG negatively regulate fibril polysaccharide production. J. Bacteriol. 186:lOOl-1008. Bode, H. B., B. Zeggel, B. Silakowski, S. C. Wenzel, H. Reichenbach, and R. Muller. 2003. Steroid biosynthesis in prokaryotes: identification of myxobacterial steroids and cloning of the first bacterial 2,3(S)-oxidosqualene cyclase from the myxobacterium Stigmatella aurantiaca. Mol. Microbiol. 47471-4 8 1. Bonner, P. J., Q. Xu, W. P. Black, Z. Li, Z. Yang, and L. J. Shimkets. 2005. The Dif chemosensory pathway is directly involved in phosphatidylethanolamine sensory transduction in Myxococcus xanthus. Mol. Microbiol. 57:1499-1508. Botella, J. A., F. J. Murillo, and R. Ruiz-Vazquez. 1995. A cluster of structural and regulatory genes for light-induced carotenogenesis in Myxococcus xanthus. Eur. J. Biochem. 233:238-248. Bowden, M. G., and H. B. Kaplan. 1998. The Myxococcus xanthus lipopolysaccharide 0-antigen is required for social motility and multicellular development. Mol. Microbiol. 30~275-284. Burchard, R. P., and S. B. Hendricks. 1969. Action spectrum for carotenogenesis in Myxococcus xanthus. J. Bacterial. 97~1165-1168. Cayuela, M. L., M. Elias-Arnanz, M. Penalver-Mellado, S. Padmanabhan, and F. J. Murillo. 2003. The Stigmatella aurantiaca homolog of Myxococcus xanthus high-mobility-
295 group A-type transcription factor CarD: insights into the functional modules of CarD and their distribution in bacteria. J. Bacteriol. 185:3527-3537. Cervantes, M., and F. J. Murillo. 2002. Role for vitamin B( 12) in light induction of gene expression in the bacterium Myxococcus xanthus. J. Bacteriol. 184:2215-2224. Chater, K. F. 1989. Multilevel regulation of Streptomyces differentiation. Trends Genet. 5:372-377. Cheng, Y. L., L. V. Kalman, and D. Kaiser. 1994. The dsg gene of Myxococcus xanthus encodes a protein similar to translation initiation factor IF3. J. Bacteriol. 176:1427-1433. Coudart, M. P. 1998. Independent patterns of expression of two alternative sigma factors, sigB and sigC, of the myxobacterium Stigmatella aurantiaca during development. Mol. Biol. Rep. 25:183-188. Downard, J., S. V. Ramaswamy, and K. S. Kil. 1993. Identification of esg, a genetic locus involved in cell-cell signaling during Myxococcus xanthus development. ]. Bacteriol. 175:7762-7770. Fontes, M., L. Galbis-Martinez, and F. J. Murillo. 2003. A novel regulatory gene for light-induced carotenoid synthesis in the bacterium Myxococcus xanthus. Mol. Microbiol. 47:561-571. Fontes, M., R. Ruiz-Vazquez, and F. J. Murillo. 1993. Growth phase dependence of the activation of a bacterial gene for carotenoid synthesis by blue light. EMBO J. 12:1265-1275. Fraser, C. M., J. D. Gocayne, 0. White, M. D. Adams, R. A. Clayton, R. D. Fleischmann, C. J. Bult, A. R. Kerlavage, G. Sutton, J. M. Kelley, R. D. Fritchman, J. F. Weidman, K. V. Small, M. Sandusky, J. Fuhrmann, D. Nguyen, T. R. Utterback, D. M. Saudek, C. A. Phillips, J. M. Merrick, J. F. Tomb, B. A. Dougherty, K. F. Bott, P. C. Hu, T. S. Lucier, S. N. Peterson, H. 0. Smith, C. A. Hutchison 111, and J. C. Venter. 1995. The minimal gene complement of Mycoplasma genitalium. Science 27Ck397-403. Gaitatzis, N., B. Silakowski, B. Kunze, G. Nordsiek, H. Blocker, G. Hofle, and R. Muller. 2002. The biosynthesis of the aromatic myxobacterial electron transport inhibitor stigmatellin is directed by a novel type of modular polyketide synthase. 1. Biol. Chem. 27713082-13090. Galbis-Martinez, M., M. Fontes, and F. J. Murillo. 2004. The high-mobility group A-type protein CarD of the bacterium Myxococcus xanthus as a transcription factor for several distinct vegetative genes. Genetics 167:1585-1595. Gill, R. E., M. Karlok, and D. Benton. 1993. Myxococcus xanthus encodes an ATP-dependent protease which is required for developmental gene transcription and intercellular signaling. J. Bacteriol. 175:4538-4544. Goldman, B. S., W. C . Nierman, D. Kaiser, S. C. Slater, A. S. Durkin, J. A. Eisen, C. M. Ronning, W. B. Barbazuk, M. Blanchard, C. Field, C. Halling, G. Hinkle, 0. Iartchuk, H. S. Kim, C. Mackenzie, R. Madupu, N. Miller, A. Shvartsbeyn, S. A. Sullivan, M. Vaudin, R. Wiegand, and H. B. Kaplan. 2006. Evolution of sensory complexity recorded in a myxobacterial genome. Proc. Natl. Acad. Sci. USA 103:15200-15205. Gollop, R., M. Inouye, and S. Inouye. 1991. Protein U, a latedevelopmental spore coat protein of Myxococcus xanthus, is a secretory protein. J. Bacteriol. 173:3597-3600.
296 Gronewold, T. M., and D. Kaiser. 2001. The act operon controls the level and time of C-signal production for Myxococcus xanthus development. Mol. Microbiol. 40:744-756. Guo, D., M. G. Bowden, R. Pershad, and H. B. Kaplan. 1996. The Myxococcus xanthus rfbABC operon encodes an ATPbinding cassette transporter homolog required for O-antigen biosynthesis and multicellular development. J. Bacteriol. 178:1631-1639. Guo, D., Y. Wu, and H. B. Kaplan. 2000. Identification and characterization of genes required for early Myxococcus xanthus developmental gene expression. J. Bacteriol. 182: 45 64-457 1. Haft, D. H., J. D. Selengut, and 0. White. 2003. The TIGRFAMs database of protein families. Nucleic Acids Res. 3 1:371-3 73. Heidelbach, M., H. Skladny, and H. U. Schairer. 1993. Heat shock and development induce synthesis of a low-molecularweight stress-responsive protein in the myxobacterium Stigmatella aurantiaca. J. Bacteriol. 175:7479-7482. Hodgkin, J., and D. Kaiser. 1979. Genetics of gliding motility in Myxococcus xanthus (Myxobacterales): genes controlling movement of single cells. Mol. Gen. Genet. 171:167176. Hull, W. E., A. Berkessel, and W. Plaga. 1998. Structure elucidation and chemical synthesis of stigmolone, a novel type of prokaryotic pheromone. Proc. Natl. Acad. Sci. USA 95: 11268-1 1273. Inouye, S., D. White, and M. Inouye. 1980. Development of Stigmatella aurantiaca: effects of light and gene expression. J. Bacteriol. 141:1360-1365. Kaiser, D. 2004. Signaling in myxobacteria. Annu. Rev. Microbiol. 58:75-98. Kaplan, H. B., A. Kuspa, and D. Kaiser. 1991. Suppressors that permit A-signal-independent developmental gene expression in Myxococcus xanthus. J. Bacteriol. 173:1460-1470. Kim, S. K., and D. Kaiser. 1990. C-factor: a cell-cell signaling protein required for fruiting body morphogenesis of M. xanthus. Cell 61:19-26. Kirby, J. R., and D. R. Zusman. 2003. Chemosensory regulation of developmental gene expression in Myxococcus xunthus. Proc. Natl. Acad. Sci. USA 100:2008-2013. Kleinig, H., and H. Reichenbach. 1969. Carotenoid pigments of Stigmatella aurantiaca (Myxobacterales). I. The minor carotenoids. Arch. Mikrobiol. 68:210-217. Kleinig, H., H. Reichenbach, and H. Achenbach. 1970. Carotenoid pigments of Stigmatella aurantiaca (Myxobacterales). 11. Acylated carotenoid glucosides. Arch. Mikrobiol. 74~223-234. Lancero, H., J. E. Brofft, J. Downard, B. W. Birren, C. Nusbaum, J. Naylor, W. Shi, and L. J. Shimkets. 2002. Mapping of Myxococcus xanthus social motility dsp mutations to the dif genes. J. Bacteriol. 184:1462-1465. Lee, K., and L. J. Shimkets. 1994. Cloning and characterization of the socA locus which restores development to Myxococcus xanthus C-signaling mutants. J. Bacteriol. 176: 2200-2209. Li, N., and M. C. Cannon. 1998. Gas vesicle genes identified in Bacillus megaterium and functional expression in Escherichia coli. J. Bacteriol. 180:2450-2458.
MYXOBACTERIAL GENOMICS AND POSTGENOMICS Lu, A., K. Cho, W. P. Black, X. Y. Duan, R. Lux, Z. Yang, H. B. Kaplan, D. R. Zusman, and W. Shi. 2005. Exopolysaccharide biosynthesis genes required for social motility in Myxococcus xanthus. Mol. Microbiol. 55:206-220. McBride, M. J., R. A. Weinberg, and D. R. Zusman. 1989. “Frizzy” aggregation genes of the gliding bacterium Myxococcus xanthus show sequence similarities to the chemotaxis genes of enteric bacteria. Proc. Natl. Acad. Sci. USA 86:424-428. McCleary, W. R., M. J. McBride, and D. R. Zusman. 1990. Developmental sensory transduction in Myxococcus xanthus involves methylation and demethylation of FrzCD. J. Bucteriol. 172:4877-4887. McCleary, W. R., and D. R. Zusman. 1990. FrzE of Myxococcus xunthus is homologous to both CheA and CheY of Salmonella typhimurium. Proc. Natl. Acad. Sci. USA 8758985902. McGowan, S. J., H. C. Gorham, and D. A. Hodgson. 1993. Light-induced carotenogenesis in Myxococcus xanthus: DNA sequence analysis of the carR region. Mol. Microbiol. 10:713-735. Moraleda-Munoz, A., J. Perez, M. Fontes, F. J. Murillo, and J. Munoz-Dorado. 2005. Copper induction of carotenoid synthesis in the bacterium Myxococcus xanthus. Mol. Microbiol. 56:1159-1168. Moreno, A. J., M. Fontes, and F. J. Murillo. 2001. ihfA gene of the bacterium Myxococcus xanthus and its role in activation of carotenoid genes by blue light. J. Bacterial. 183557-569. Myers, E. W., G. G. Sutton, A. L. Delcher, I. M. Dew, D. P. Fasulo, M. J. Flanigan, S. A. Kravitz, C . M. Mobarry, K. H. Reinert, K. A. Remington, E. L. Anson, R. A. Bolanos, H. H. Chou, C. M. Jordan, A. L. Halpern, S. Lonardi, E. M. Beasley, R. C. Brandon, L. Chen, P. J. Dunn, Z. Lai, Y. Liang, D. R. Nusskern, M. Zhan, Q. Zhang, X. Zheng, G. M. Rubin, M. D. Adams, and J. C. Venter. 2000. A wholegenome assembly of Drosophila. Science 287:2196-2204. Nicolas, F. J., R. M. Ruiz-Vazquez, and F. J. Murillo. 1994. A genetic link between light response and multicellular development in the bacterium Myxococcus xanthus. Genes Dev. 8~2375-2387. Nierman, W. C., A. Pain, M. J. Anderson, J. R. Wortman, H. S. Kim, J. Arroyo, M. Berriman, K. Abe, D. B. Archer, C. Bermejo, J. Bennett, P. Bowyer, D. Chen, M. Collins, R. Coulsen, R. Davies, P. S. Dyer, M. Farman, N. Fedorova, N. Fedorova, T. V. Feldblyum, R. Fischer, N. Fosker, A. Fraser, J. L. Garcia, M. J. Garcia, A. Goble, G. H. Goldman, K. Gomi, S. Griffith-Jones, R. Gwilliam, B. Haas, H. Haas, D. Harris, H. Horiuchi, J. Huang, S. Humphray, J. Jimenez, N. Keller, H. Khouri, K. Kitamoto, T. Kobayashi, S. Konzack, R. Kulkarni, T. Kumagai, A. Lafton, J. I?. Latge, W. Li, A. Lord, C. Lu, W. H. Majoros, G. S. May, B. L. Miller, Y. Mohamoud, M. Molina, M. Monod, I. Mouyna, S. Mulligan, L. Murphy, S. O’Neil, I. Paulsen, M. A. Penalva, M. Pertea, C. Price, B. L. Pritchard, M. A. Quail, E. Rabbinowitsch, N. Rawlins, M. A. Rajandream, U. Reichard, H. Renauld, G. D. Robson, S. Rodriguez de Cordoba, J. M. RodriguezPena, C. M. Ronning, S. Rutter, S. L. Salzberg, M. Sanchez, J. C. Sanchez-Ferrero, D. Saunders, K. Seeger, R. Squares, S. Squares, M. Takeuchi, F. Tekaia, G. Turner, C. R. Vazquez de Aldana, J. Weidman, 0. White, J. Woodward, J. H. Yu,
16. THEGENOMES OF 211.
XANTHUS AND
S. AURANTIACA
C. Fraser, J. E. Galagan, K. Asai, M. Machida, N. Hall, B. Barrell, and D. W. Denning. 2005. Genomic sequence of the pathogenic and allergenic filamentous fungus Aspergillus fumigatus. Nature 438:1151-1156. O’Connor, K. A., M. J. McBride, M. West, H. Yu, L. Trinh, K. Yuan, T. Lee, and D. R. Zusman. 1996. Photolyase of Myxococcus xanthus, a Gram-negative eubacterium, is more similar to photolyases found in Archaea and “higher” eukaryotes than to photolyases of other eubacteria. J . Biol. Chem. 271:6252-6259. Offner, S., A. Hofacker, G. Wanner, and F. Pfeifer. 2000. Eight of fourteen gvp genes are sufficient for formation of gas vesicles in halophilic archaea. J. Bacteriol. 182:4328-4336. Paitan, Y., E. Orr, E. Z. Ron, and E. Rosenberg. 1999. Genetic and functional analysis of genes required for the post-modification of the polyketide antibiotic TA of Myxococcus xanthus. Microbiology 145(Pt. 11):3059-3067. Plaga, W., I. Stamm, and H. U. Schairer. 1998. Intercellular signaling in Stigmatella aurantiaca: purification and characterization of stigmolone, a myxobacterial pheromone. Proc. Natl. Acad. Sci. USA 9 5 1 1263-1 1267. Plaga, W., and S. H. Ulrich. 1999. Intercellular signalling in Stigmatella aurantiaca. Cur%Opin. Microbiol. 2593-597. Pospiech, A., B. Cluzel, J. Bietenhader, and T. Schupp. 1995. A new Myxococcus xanthus gene cluster for the biosynthesis of the antibiotic saframycin M x l encoding a peptide synthetase. Microbiology 141(Pt. 8):1793-1803. Qualls, G. T., K. Stephens, and D. White. 1978. Light-stimulated morphogenesis in the fruiting myxobacterium Stigmatella aurantiaca. Science 201:444-445. Rau, W. 1988. Functions of carotenoids other than in photosynthesis, p. 231-255. In T. Goodwin (ed.), Plant Pigments. Academic Press, London, United Kingdom. Riley, M. 1993. Functions of the gene products of Escherichia coli. Microbiol Rev. 575362-952. Salzberg, S. L., A. L. Delcher, S. Kasif, and 0. White. 1998. Microbial gene identification using interpolated Markov models. Nucleic Acids Res. 26544-548. Sancar, G. B. 1990. DNA photolyases: physical properties, action mechanism, and roles in dark repair. Mutat. Res. 236:147-160. Shimkets, L., and C. R. Woese. 1992. A phylogenetic analysis of the myxobacteria: basis for their classification. Proc. Natl. Acad. Sci. USA 89:9459-9463. Shimkets, L.J. 1990. Social and developmental biology of the Myxobacteria. Microbiol. Rev. 54:473-501. Silakowski, B., H. Ehret, and H. U. Schairer. 1998. fbfB, a gene encoding a putative galactose oxidase, is involved in Stigmatella aurantiaca fruiting body formation. J. Bacteriol. 180:1241-1247. Silakowski, B., B. Kunze, and R. Muller. 2000a. Stigmatella aurantiaca Sg a15 carries genes encoding type I and type I1 3-deoxy-D-arabino-heptulosonate-7-phosphate synthases: involvement of a type I1 synthase in aurachin biosynthesis. Arch. Microbiol. 173:403-411. Silakowski, B., B. Kunze, and R. Muller. 2001a. Multiple hybrid polyketide synthasehon-ribosomal peptide synthetase gene clusters in the myxobacterium Stigmatella aurantiaca. Gene 275:233-240.
297 Silakowski, B., B. Kunze, G. Nordsiek, H. Blocker, G. Hofle, and R. Muller. 2000b. The myxochelin iron transport regulon of the myxobacterium Stigmatella aurantiaca Sg a l 5 . Eur. J. Biochem. 267:6476-6485. Silakowski, B., G. Nordsiek, B. Kunze, H. Blocker, and R. Muller. 2001b. Novel features in a combined polyketide synthasehon-ribosomal peptide synthetase: the myxalamid biosynthetic gene cluster of the myxobacterium Stigmatella aurantiaca Sga1.5. Chem. Biol. 859-69. Silakowski, B., A. Pospiech, B. Neumann, and H. U. Schairer. 1996. Stigmatella aurantiaca fruiting body formation is dependent on the fbfA gene encoding a polypeptide homologous to chitin synthases. J. Bacteriol. 178:6706-6713. Silakowski, B., H. U. Schairer, H. Ehret, B. Kunze, S. Weinig, G. Nordsiek, P. Brandt, H. Blocker, G. Hofle, S. Beyer, and R. Muller. 1999. New lessons for combinatorial biosynthesis from myxobacteria. The myxothiazol biosynthetic gene cluster of Stigmatella aurantiaca DW4/3-1. J. Biol. Chem. 2 7 4 ~739 3 1-37399. Skladny, H., M. Heidelbach, and H. U. Schairer. 1992. Cloning and DNA sequence of sigB gene of Stigmatella aurantiaca. Nucleic Acids Res. 20:6416. Skladny, H., M. Heidelbach, and H. U. Schairer. 1994. Cloning and characterization of the gene encoding the major sigma factor of Stigmatella aurantiaca. Gene 143:123-127. Spormann, A. M. 1999. Gliding motility in bacteria: insights from studies of Myxococcus xanthus. Microbiol. Mol. Biol. Rev. 63:621-641. Sproer, C., H. Reichenbach, and E. Stackebrandt. 1999. The correlation between morphological and phylogenetic classification of myxobacteria. Int. J. Syst. Bacteriol. 49(Pt. 3 ) : 1255-1262. Stamm, I., F. Lottspeich, and W. Plaga. 2005. The pyruvate kinase of Stigmatella aurantiaca is an indole binding protein and essential for development. Mol. Microbiol. 56:13861395. Thony-Meyer, L., and D. Kaiser. 1993. devRS, an autoregulated and essential genetic locus for fruiting body development in Myxococcus xanthus. J. Bacteriol. 175:74507462. Toal, D. R., S. W. Clifton, B. A. Roe, and J. Downard. 1995. The esg locus of Myxococcus xanthus encodes the E l alpha and E l beta subunits of a branched-chain keto acid dehydrogenase. Mol. Microbiol. 16:177-189. Tojo, N., S. Inouye, and T. Komano. 1993. The lonD gene is homologous to the lon gene encoding an ATP-dependent protease and is essential for the development of Myxococcus xanthus. J. Bacteriol. 175:4545-4549. Trudeau, K. G., M. J. Ward, and D. R. Zusman. 1996. Identification and characterization of FrzZ, a novel response regulator necessary for swarming and fruiting-body formation in Myxococcus xanthus. Mol. Microbiol. 20:645-655. Ueki, T., and S. Inouye. 2003. Identification of an activator protein required for the induction of fruA, a gene essential for fruiting body development in Myxococcus xanthus. Proc. Natl. Acad. Sci. U S A 100:8782-8787. van Keulen, G., D. A. Hopwood, L. Dijkhuizen, and R. G. Sawers. 2005. Gas vesicles in actinomycetes: old buoys in novel habitats? Trends Microbiol. 13:350-354.
298 Vlamakis, H. C., J. R. Kirby, and D. R. Zusman. 2004. The Che4 pathway of Myxococcus xantbus regulates type IV pilus-mediated motility. Mol. Microbiol. 52:1799-1811. Wall, D., P. E. Kolenbrander, and D. Kaiser. 1999. The Myxococcus xantbus pilQ (sglA) gene encodes a secretin homolog required for type IV pilus biogenesis, social motility, and development. J. Bacteriol. 181:24-33. Ward, M. J., H. Lew, and D. R. Zusman. 2000. Social motility in Myxococcus xantbus requires FrzS, a protein with an extensive coiled-coil domain. Mol. Microbiol. 371357-1371. Waterman, M. S. 1988. Computer analysis of nucleic acid sequences. Methods Enzymol. 164:765-793. Wenzel, S. C., B. Kunze, G. Hofle, B. Silakowski, M. Scharfe, H. Blocker, and R. Muller. 2005. Structure and biosynthesis of myxochromides S1-3 in Stigmatella aurantiaca: evidence for an iterative bacterial type I polyketide synthase and for module skipping in nonribosomal peptide biosynthesis. Chembiochem 6:375-3 85.
MYXOBACTERIAL GENOMICS AND POSTGENOMICS White, D., W. Shropshire, Jr., and K. Stephens. 1980. Photocontrol of development by Stigmatella aurantiaca. J. Bacterial. 142:1023-1024. Wu, S. S., and D. Kaiser. 1995. Genetic and functional evidence that Type IV pili are required for social gliding motility in Myxococcus xantbus. Mol. Microbiol. 18547558. Wu, S. S., J. Wu, Y. L. Cheng, and D. Kaiser. 1998. The pilH gene encodes an ABC transporter homologue required for type IV pilus biogenesis and social gliding motility in Myxococcus xantbus. Mol. Microbiol. 29:1249-1261. Wu, S. S., J. Wu, and D. Kaiser. 1997. The Myxococcus xantbus pilT locus is required for social gliding motility although pili are still produced. Mol. Microbiol. 23:109-121. Youderian, P., N. Burke, D. J. White, and P. L. Hartzell. 2003. Identification of genes required for adventurous gliding motility in Myxococcus xanthus with the transposable element mariner. Mol. Microbiol. 49255-570.
Myxobacteria: Multicellularity and Differentiation Edited by David E. Whitworth 0 2008 ASM Press, Washington, D.C.
Garret Suen Barry S. Goldman Roy D. Welch
A Postgenomic Overview
17
of the Myxobacteria
The first genome sequence of a myxobacterium appeared in 2001, when the Monsanto Company released a partial genome sequence of Myxococcus xanthus strain DK1622. This initial sequence, known as the “Mlgenome,” represented approximately 98% of the total sequence at 4.5X coverage and consisted of 1,273 nonoverlapping contiguous DNA sequences (Jakobsen et al., 2004). The impact of the Mlgenome on almost every active area of M. xanthus research was immediate and profound. Since this beginning, The Institute for Genomic Research (TIGR) finished and assembled the genome, and now, 6 years and several updates later, this genome is considered an essential research tool. As a direct demonstration of this fact, more than 75 research articles on M. xanthus have been published that utilize data from the genome. This is due in large part to the genome’s inherent value as the complete data set of every DNA base pair required to create and maintain an M . xanthus bacterium. From a practical perspective, the real utility of the M. xanthus genome largely depends on the specifics of any given research project. At a minimum, the genome
represents a searchable database for all of the DNA sequence in M. xanthus and, as such, it has been used in conjunction with screens or libraries to simplify the identification of genes. The genome has been used as a starting point for mutagenesis and for providing information on sets of paralogous genes that can be studied together as targets for disruption or deletion. The completed sequence has also allowed for the application of high-throughput technologies, such as a gene expression microarray, and the development of functional genomictechniques to study the relationship between genes and genetic networks. Currently, M. xanthus is the only species of myxobacteria to mature into the postgenomic phase, and therefore, this chapter focuses on this organism. Given the current speed of genomic development, however, M. xanthus will not be alone for long. We predict that this will be the only edition of the Myxobacteria series to contain a postgenomics chapter; by the next edition, application of the genome will be so pervasive that it will be considered an integral part of other topics, rather than a topic unto itself. In many ways this has already occurred for M. xanthus.
Garret Suen and Roy D. Welch, Department of Biology, Syracuse University, Syracuse, NY 13244. Barry S. Goldman, Monsanto Company, St. Louis, M O 63167.
299
MYXOBACTERIAL GENOMICS AND POSTGENOMICS
300
APPLYING GENOMICS TO EXPERIMENTAL BIOLOGY After the initial release of 98% of the M . xanthus Mlgenome (Jakobsen et al., 2004), many experimental researchers shifted their focus to characterizing large numbers of genes. This is reflected in recent publications, where emphasis has shifted from single genes to the characterization of multiple genes involved in a particular function. In this section, we highlight some of the research occurring on the genomic scale using experimental biology. We begin our discussion with genomic screens using transposable elements, continue with the paralogous analysis of gene families, and conclude with whole-genome yeast two-hybrid (Y2H) assays.
Genomic Screens Using Transposable Elements Transposable insertion elements are pieces of genetic material that can insert themselves into new locations in the genome and possibly disrupt coding sequences (Mahillon and Chandler, 1998). Random mutagenesis through transposable insertion elements has traditionally been used to create strains of M. xanthus that can be screened for specific phenotypes such as motility. Previous studies using the transposon Tn5 identified mutants defective for adventurous (A) motility and only a handful of these loci were mapped onto the genome (MacNeil et al., 1994; Wu and Kaiser, 1995). With the completion of the M. xanthus genome, the ability to map large numbers of transposon insertions became feasible. This technique involves screening a library of transposon insertions for mutants exhibiting a desired phenotype. The transposon insertion can be used as a primer to sequence the disrupted region, and an alignment program such as BLAST (Altschul et al., 1997) can be used to map the sequence to a specific locus on the genome. In this way, rather than focusing on one or a few open reading frames (ORFs),all of the ORFs can be identified and associated with a specific phenotype. Youderian and coworkers (2003) first demonstrated the utility of this approach by screening a magellan-4 random mutagenesis library for M. xanthus strains deficient in A-motility. They sequenced regions of the M . xanthus Mlgenome (Jakobsen et al., 2004) adjacent A
B
C
D
E
F
G
H
I
J
K
L
to magellan-4 insertions and mapped 115 A-motility mutants to 33 putative ORFs. Three of these ORFs were previously characterized as A-motility genes, whereas the remaining 30 were novel. The putative annotations of these ORFs support a current model of A-motility: the extrusion of polysaccharide “slime” through pores located at the polar ends of each cell (Wolgemuth et al., 2002). For example, many of the ORFs identified in this screen are thought to be involved in the production and transport of polysaccharides, and six ORFs encode homologues of the To1 transport system, suggesting that M. xanthus may extrude exopolysaccharide (EPS) by using this system. The Tn5 transposable element has also been used to create random mutagenesis libraries. Using a method similar to that of Youderian and coworkers, Lu and coworkers (2005) screened a Tn5 mutagenesis library for mutants deficient for social (S) motility and EPS production. S-motility, a motility system whereby groups of M. xanthus cells move cooperatively, requires the production of EPS, presumably for cell-cell adhesion and as a substrate for coordinated movement via pilus retraction (Li et al., 2003). Screening of over 5,000 colonies resulted in the identification of 68 mutants, mapped to two specific regions on the M. xanthus chromosome. The first region, a 37.2-kb region termed the eps (EPSsynthesis) region, contains 26 ORFs (Fig. l),the majority of which had homologs to genes involved in EPS biosynthesis in other organisms. The second region, a 1.7-kb region termed the eas (EPS-associated) region, contains two ORFs annotated as hypothetical proteins. Further analyses of these two ORFs did not reveal any function of these genes in EPS production. More recently, Youderian and coworkers (2006) revisited their random mutagenesis screen by screening a magellan-4 insertion library for mutants deficient in S-motility. They isolated 132 mutants which were then mapped to 68 ORFs on the completed M . xanthus genome (Goldman et al., 2006). Many of the discovered ORFs were classified into four categories known to be required for S-motility. The first category, type IV pili (TFP) biogenesis, contains 15 ORFs that map to a 27-kb region on the M. xanthus genome, as shown in Fig. 2A. Disruption M
N
O P Q R S T U
V
W
I
Figure 1 Schematic of the EPS (eps)region in M. xanthus. A total of 26 ORFs were identified by Lu and coworkers (2005), of which 12 contained Tn5 insertions found through a random mutagenesis screen (indicated by black dots). Reprinted and modified with permission from Lu and coworkers (2005).
X
Y
Z
(A) Type IV pilin biogenesis
dMI0 5766
5767 5758
5770 S771
5772 5773 5774 5?75 5776
5777
5778 5779 5760 578? 3782 5783 5784
5786 5167
5785
5788
(B) Exopolysaccharide biosynthesis
x 7479
epsZepsE
x 7416 7417 74f8 7420 7421 7422 7423 7425 7430 epsY epsx epsW epsV epsU 74.?4 7426 7431
7433 eps0
7435
epsN
-----------------*-c--*-c+--------c
-
7439 74$0 7441 epsJ e~Unla24epsfi
7442 7443
7444
epsG
epsF
7445 7647 7448 7449 7460 746; ewD ep& epSe epsA/cpsF
epsE
+=----A ___I__,
(C) LPS Biosynthesis and known S-motility genes
1 %
//-
8
3;
11_r-/I
E71ihii 2920
2921 sgd
-+-
2922 sgmK
f8
f
4f49
UPrS
h
4750 4757
sgmd
------+-
/1
'
s;
dhm 4673 4614 sgmP 46'5
+-- *-+ -
L
I
4816 sgmQ
4617
461a
4619 wb%B
4626 sgmR
4627 rfbC
4622
rfbs
4623 rfba
Figure 2 Location of magellan-4 insertions in specific regions of the M. xanthus genome. Disruption of putative ORFs resulted in deficiencies in S-motility. Four groups were identified including genes involved in TFP biogenesis (A), EPS biosynthesis (B), and LPS biosynthesis and known S-motility genes (C). Inverted triangles represent magellan-4 insertion sites. Reprinted and modified with permission from Youderian and coworkers (2006).
EAhB -
4638
4639
wmH
sgmS
4640 sgmT
302 of these ORFs likely caused deficiencies in M. xanthus’s ability to produce TFP. The second category, EPS biosynthesis, contained 10 ORFs mapped to the eps cluster previously discovered by Lu and coworkers (2005), as shown in Fig. 2B. A third category, lipopolysaccharide synthesis, contained a cluster of five ORFs predicted to be required for S-motility (Bowden and Kaplan, 1998). The fourth category contained additional genes known to be required for S-motility (Fig. 2C), including tgl, a gene encoding an outer membrane lipoprotein (Nudleman et al., 2005), and frzS, a gene that encodes a protein required for directed motility that is localized to the cell pole (Mignot et al., 2005). In addition to recovering genes known to be involved in S-motility, they also found 31 ORFs related to a variety of other processes including transcriptional regulation, primary metabolism, and polysaccharide metabolism.
A Paralog Analysis of Gene Families One of the first types of analyses that became possible with the completed genome is the paralogous analysis of gene families. Paralog analysis of the M . xanthus genome has focused on the signaling pathways utilized to respond to the changing environment characteristic of its soil habitat. The general technique for this approach is to identify members of specific gene families by aligning all predicted ORFs in the genome against the sequence of a known gene within the gene family. Characterization of each predicted member through genetic disruption and phenotypic analysis identifies a hypothetical role for each gene in the gene family. In M. xanthus, two gene families have been extensively characterized by use of this method: the NtrC-like transcriptional activators (TA) and the serinehhreonine protein kinases (STPIC). The NtrC-like TAs are enhancer binding proteins that recruit os4-bound RNA polymerases to transcribe genes and are often found as response regulators in two-component signaling pathways (Morett and Segovia, 1993). A survey of the M. xanthus genome reveals a total of 52 NtrC-like TAs (Jelsbak et al., ZOOS), a number that is large, relative to other bacteria (Goldman et al., 2006). The NtrC-like TAs are known to participate in multicomponent signaling pathways, and it has been suggested that these complex signaling networks are the evolutionary beginnings of multicellularity (Goldman et al., 2006). The characterization of these TAs has thus been the subject of recent work by some laboratories in the M . xanthus research community. Caberoy and coworkers presented the first global analysis of the NtrC-like TAs in 2003. Through a paralog search of the M . xanthus Mlgenome (Jakobsen et al., 2004), they identified 37 putative NtrC-like TAs, 28 of
MYXOBACTERIAL GENOMICS AND POSTGENOMICS which were uncharacterized. Using homologous recombination, they made disruptions in these 28 ORFs and tested the resulting mutants for developmental defects. Eight of these mutants were defective for development and became the focus of their study. They tested these mutants for defects in A- and S-motility and found that three mutants, nla2 (NtrC-like activator), nlal9, and nlu23 were defective for S-motility, while nlu24 was defective for both motility systems. The three mutants defective for S-motility were slightly delayed for aggregation, consistent with previous findings that an intact S-motility system is essential for normal aggregation and fruiting body production. Interestingly, the other four mutants (rtlu4, d a b , n h l 8 , and nla28) were not defective for motility but failed to aggregate or form spores. These findings suggest that the NtrC-like TAs play an important role in the M. xanthus starvation response by regulating genes required for development. Further insight into the role that NtrC-like TAs play in specific signaling pathways was provided by Jelsbak and coworkers (ZOOS). Using Pfam (Finn et al., 2006) analysis, they identified 52 ORFs annotated as NtrC-like TAs. Further analysis revealed that 12 of these contained forkhead-associated (FHA)domains, as shown in Fig. 3 . Proteins containing FHA domains are believed to interact with STPKs. Characterization of one of these, mx488.5, revealed abnormal and delayed aggregation and a severe reduction in sporulation. Analysis of this protein’s FHA domain showed that it contains a phosphothreonine recognition site. Since many NtrC-like TAs are found adjacent to STPKs in the genome, they suggested that these pairs of genes might possibly interact. The second major set of characterized paralogs in M. xanthus are the STPKs. These proteins are known to play critical roles in sensing environmental signals and initiating signal transduction pathways in eukaryotes (Inouye et al., 2000; Kaiser, 1986; Ueki and Inouye, 2006). Interestingly, few prokaryotes are known to use these signaling molecules, and M. xanthus is a rare exception, with an estimated 97 STPKs in its genome (Goldman et al., 2006); this number is similar to that found in lower eukaryotes such as Saccharomyces cerevisiae. The characterization of STPKs has been the pioneering work of Sumiko Inouye’s group, and the paralogous identification and systematic inactivation of 94 of these STPKs showed that at least 20 are essential for development (Inouye et al., 2000; Munoz-Dorado et al., 1991).STPKs appear to be involved in fruiting body formation by controlling the timing of this event. For example, the STPKs Pknl, Pkn2, and Pkn5 are negative regulators of fruiting body formation, as inactivation of these genes results in faster fruiting body formation (Munoz-Dorado et al., 1991;
17. POSTGENOMIC OVERVIEW OF THE MYXOBACTERIA
Domain organization Sensory domain
303
ORF
Sigma54 interaction ATPase domain
-
previOus'yl
identified
DNA binding domain Generic EBP
..-..._.....*._-..--..*--**-..-..--....---*..---.-..---..-..-.-.-.--"--. Mx2469 (449 aa) Mxa249 Mx5079 f446 aa) Mxa264 Mx0888 (476 aa) Nla18 Mxl288 (459 aa) Mx4901 (472 aa) Nla14 Mx2176 (454 aa) Nla27 Mx4885 (459 aa) Mxl502 (546 aa) Nla5 Mxl757 (476 aa) Mxal91 Mx3725 (566 aa) Mxl598 (630 aa) Mxa213 Mx4562 (581 aa) Nla12
100 aa
Sigma54 Interaction
I
Figure 3 Domain organization of the 12 FHA-associated NtrC-like activators in M. xanthus. The domain organization of a generic NtrC-like activator is shown for reference. Reprinted with permission from Jelsbak et al., 2005.
Udo et al., 1996; Zhang et al., 1996). In contrast, inactivation of Pkn6 and Pkn9 results in slower fruiting body formation (Hanlon et al., 1997; Zhang et al., 1996). Recently, the Inouye group has proposed a model suggesting that both STPKs and histidine kinases can regulate the activity of the same TA (Nariya and Inouye, 2005), as shown in Fig. 4. This model includes an STPK signaling cascade, the first confirmed cascade of its type in any prokaryote (Lux and Shi, 2005). The STPK Pkn8 is a transmembrane protein capable of receiving an external signal and phosphorylates the STPK Pknl4, which in turn phosphorylates the cyclic AMP receptor protein, MrpC. The mrpC gene is part of the three-gene mrpABC operon (Sun and Shi, 2001a, 2001b) and modulates the expression of genes required for development, such as
fruA (Ueki and Inouye, 2003). The mrpA gene encodes a histidine kinase, which phosphorylates the NtrC-like TA mrpB, which then activates the transcription of mrpC. Using eight putative binding regions found upstream of mrpC, they searched the 111. xanthus genome for other potential target binding regions for MrpC. They found binding sites upstream of a number of genes, including the development gene fmA, a gene involved in the production of antibiotic TA, a multidrug efflux pump gene, and a peptidase gene. Availability of the 211. xanthus genome sequence makes the paralogous characterization of gene families possible. The characterization of gene families such as the NtrClike TAs and the STPKs has increased our understanding of signal transduction pathways and, more importantly,
MYXOBACTERIAL GENOMICS AND POSTGENOMICS
304
Vegetative growth
~e~elopIiien~a1 gene expression
=I[
Protein SeriThr kinase cascade Protein Ser/Ttir kinase Histicline kinase Sigma 54-dependent response regulato CRP famiiy t r ~ n s c ~ p ~ i oactivator nal Response regulator (=> Lori family swine protease
~ W o - ~ o ~ ~signal ~ I itrarisduction e n ~ S e r ~ ~ - ~ h o s p hgroup or~l High energy phiosphoryl group ~Iiviro?iment~l signal Phosp~o~lation Re~la~io~Acti~ation Re~laEioIi/Ii~l~bi tion
Figure 4 Schematic representation of the MrpC pathway. MrpC is found to be regulated using both a two-component signal transduction HPK and an STPK cascade system. Details of this pathway are outlined in the text. Reprinted with permission from Nariya and Inouye (2006).
allows researchers to construct genetic networks that describe the genetic underpinnings of M . xanthus’s complex life cycle. The findings of the Inouye group confirm the hypothesis that M . xanthus can co-opt multiple sensory input components such as the STPKs to regulate specific signaling pathways. As researchers continue to characterize other paralogous groups in the genome, our understanding of how these families contribute to the genetic pathways utilized by M. xanthus will eventually be made clear.
Genome-Wide Protein Interactions Using the Y2H System The Y2H method, first introduced by Fields and Song (1989), is a powerful technique capable of detecting pairs of protein-protein interactions. This method has been utilized by the M. xanthus field to explore the protein interactions that occur during its life cycle. For example, the previously discussed STPK signal cascade pathway was confirmed through the use of Y2H assays. In addition, Y2H assays have been used to characterize
17. POSTGENOMIC OVERVIEW OF THE MYXOBACTERIA a number of interactions between proteins, including the two-component signal transduction pathway encoded by the red operon (Higgs et al., ZOOS), the light-induced carotenogenesis operon car (Browning et al., 2003; Whitworth and Hodgson, 2001), the interaction between the operons mas and mgl involved in A-motility (Thomasson et al., 2002), and the interaction between the chemotaxis operon frz and an operon encoding for ATP-binding cassette proteins (Ward et al., 1998). With the availability of the genome sequence, researchers are now conducting Y2H screens in a manner similar to that of random mutagenesis screens. The general technique involves constructing a whole-genome library using chromosomal DNA, conducting the Y2H screen using a desired bait protein, and identifying the yeast colonies that contain positive interactions. Sequencing of the plasmid in positive colonies reveals the DNA region encoding the positive interaction. Comparison of this sequence against the M. xanthus genome identifies the ORF discovered through the screen. In M. xanthus, a small number of large-scale studies using this method have been reported. Yang and coworkers began the first Y2H screen using the M. xanthus genome in 2004 (Yang et al., 2004). They utilized this approach to investigate specific proteins that interact with the cytoplasmic GTPase encoded by mglA, a gene required for both A- and S-motility systems. From their screen, they isolated aglZ, which encodes a protein related to type-2 myosin. Characterization of aglZ through inactivation showed that it is involved in A-motility and to some extent in S-motility. Interestingly, the interaction between a GTPase and a myosinlike protein has never been reported in prokaryotes; in eukaryotes this interaction is essential for vesicular transport. Based on these findings and the current model of A-motility (Wolgemuth et al., 2002), they suggest that this interaction may be involved in the transport of substances extruded out of the cell. An elegant demonstration of the Y2H screen was reported by Lancero and coworkers in 2005 (Lancero et al., 2005). Utilizing the genes within the dif operon, which encode six proteins essential for fruiting body formation, fibril production, and S-motility, they conducted a Y2H screen against the whole M. xanthus proteome. Interestingly, they found that the Dif proteins were highly specific and tend to interact with other Dif proteins. From their results they constructed a model of the Dif signaling pathway, which mirrors many of the interactions found within the chemotaxis system of other bacteria (Szurmant and Ordal, 2004). DifA, a methyl-accepting chemotaxis protein, forms a ternary complex with DifC and DiE, homologs of Chew and
305
CheA, respectively. DifE is found to interact with DifD, a homolog of CheY. DifF, a homolog of CheC, interacts with DifE, suggesting that DifF may function by dephosphorylating DifE in a manner similar to that observed in Bacillus subtilis (Szurmant and Ordal, 2004). DifB, which has no known homolog, was not found to interact with any of the other Dif proteins. The interactions in this pathway have since been independently confirmed by Yang and Li (2005), also using the Y2H system. Pham and coworkers (2005) used a full-proteome Y2H screen to find proteins that interact with SdeK, a histidine kinase essential for development. They recovered six interaction partners including a hybrid response regulator (MXAN-3879), an NtrC-like TA (MXAN5048), an ABC transporter permease (MXAN-3773), a chorismate mutase/prephenate dehydratase (MXAN3221), a DnaK homolog (MXAN-5323), and a Baf-like protein (MXAN-4151). Inactivation of these ORFs showed that only one, MXAN-4151 (termed brgE for Baf-like regulatory effector), had a phenotype similar to that of sdeK mutants for development. Further analysis of this interaction showed that both participate in the same pathway during development. The use of Y2H screens to identify protein interactions in M. xanthus is greatly facilitated by a genome sequence. Any positive colonies retrieved from the screen can be sequenced, and the corresponding ORF can be identified. So far, studies utilizing Y2H screens have primarily focused on the interactions of motility and chemosensory proteins. Y2H screens are a valuable technique that can be used to quickly identify new components of pathways. Combined with other techniques, such as random mutagenesis screens and paralog analyses, we expect our understanding of M. xanthus genetic network topology to rapidly increase in the very near future.
NEW EXPERIMENTAL GENOMICS TECHNOLOGIES In the past decade, a plethora of high-throughput genomics technologies have been developed, made possible by genome sequencing. Starting from the sequence, experimental techniques can measure the different features of all of the genes in the genome, spawning the so-called “omics” revolution. These include the whole suite of array technologies such as DNA microarrays (transcriptomics), protein microarrays (proteomics), and phenotype arrays (phenomics).Each technique provides a wealth of data that requires the application of automated data management and analysis through bioinformatics techniques. Currently, the only high-throughput
306 technology available to the M. xanthus community is a DNA microarray.
The M. xanthus Microarray The first-generation M . xanthus microarray was constructed in 2001, based on the Mlgenome sequence. A total of 7,242 ORFs were predicted from this sequence, and amplicons corresponding to these ORFs were spotted onto anarray (Jakobsenet a1.,2004). Asecond-generation M . xanthus microarray has since been constructed by TIGR, based on the completed genome sequence, and is currently under preliminary evaluation by several members of the M. xanthus community (M. Singer, personal communication). The first use of microarray technology for M. xanthus was reported by Jakobsen and coworkers (Jakobsen et al., 2004). They identified 53 putative NtrC-like TAs from the Mlgenome and constructed a microarray with amplicons generated from the sequence of these ORFs. In addition to these, 224 other ORFs annotated as putative DNA binding proteins involved in the regulation of transcription were also included. As a control, 94 previously characterized M. xanthus genes were also spotted. All of these 371 ORFs were spotted in duplicate and assayed for transcriptional levels of developing cells at 12 h. The set of mRNA transcripts from vegetatively growing wildtype strains was used as a reference sample. Based on their results, they found that six of the NtrClike TAs were expressed at higher levels during development than during vegetative growth. Three of these genes, sasR, spdR, and pilR, were previously characterized and implicated in development. Disruption of the other three ORFs showed that only one of these genes, Mx-3320, was defective for development, specifically with respect to aggregation and sporulation. In addition, a number of genes known to be involved in motility and cell-cell signaling were found to be up-regulated. For example, four S-motility genes from the difand pil operons were found to be up-regulated, consistent with the observation that S-motility is required for normal development. Finally, a number of one-component transcriptional regulators were also found to be up-regulated during development, including those belonging to the AraC, ArsR, TetR, MarR, and LysR families. The first full-scale analysis of the M . xanthus genome using the microarray was recently reported by Diodati and coworkers (2006). They compared a mutant strain inactivated for the gene nlaZ8 under vegetative growing against wild-type cells. Previous investigations of nlaZ8, a gene coding for an NtrC-like TA (Caberoy et al., 2003), showed that disruption of this TA caused defects in both aggregation and sporulation. Microarray
MYXOBACTERIAL GENOMICS AND POSTGENOMICS experiments were conducted using mRNA transcripts isolated from nla18 and wild-type cells growing under vegetative conditions. More than 700 ORFs were found to be significantly affected by the mutation. Classification of these ORFs showed that the largest group contains those encoding putative membrane and membraneassociated proteins. Based on these results, they hypothesized that nlal8 might be involved in the maintenance and synthesis of membrane-associated proteins. To test this, they generated membrane protein profiles for nlaZ8 and wildtype M. xanthus cells and found that these profiles were remarkably different; some proteins were either missing or underrepresented in the nla18 mutant. Based on these observations, they concluded that nlal8 is an important regulator of membrane proteins. In addition to the membrane-associated proteins, many other classes of genes were also found to be up- or down-regulated in the nlal8 mutant, including genes involved in metabolism, translation, and transcriptional regulation. Both of the findings presented in this section are significant because they represent how the application of a genome sequence can aid in the identification of genes involved in specific processes such as development and vegetative growth. The ability to rapidly characterize the expression levels of a large number of genes allows researchers to investigate the genetic networks responsible for the complex shifts in phenotype observed during the life cycle of M . xanthus.
FUNCTIONAL GENOMICS: MAKING PREDICTIONS All of the postgenomics methods presented in this chapter so far focus on using the genome sequence to facilitate the characterization of genes using experimental techniques. Functional genomics seeks to use the genomic data generated by these experimental techniques to make predictions about novel interactions (Ge et al., 2003; Marcotte and Date, 2001; Vidal, 2001). Genomescale data sets, such as sequence and microarray data, can be processed by integration algorithms to make predictions about the putative interaction partners of genes within the genome. These methods have been successfully used to recapitulate known pathways such as the galactose utilization pathway in S. cerevisiae (Hwang et al., 2005a, 2005b). It is thought that the most accurate predictions are made by integrating large numbers of genome-scale data sets. For example, the recapitulation of the galactose utilization pathway in S . cerevisiae was predicted by integrating 18 different genome scale data sets.
17. POSTGENOMIC OVERVIEW OF THE MYXOBACTERIA Genome scale data sets can be divided into two categories: those based on sequence data and those based on high-throughput experiments (Suen et al., 2007).The sequence-based genome scale data sets include those based on gene fusions as “Rosetta Stones” (Marcotte et al., 1999), the tracking of correlated mutations (Gertz et al., 2003; Pazos and Valencia, 2002), the enumeration of conserved operons (Overbeek et al., 1999), and the calculation of phylogenetic profiles (Pellegrini et al., 1999). The experiment-based genome scale data sets include those based on DNA microarray experiments (Lockhart and Winzeler, 2000), protein microarray experiments (LaBaer and Ramachandran, 2005), metabolomics (Fiehn, 2002), and ChIP-on-chip (Buck and Lieb, 2004). M. xanthus currently has two welldeveloped genome scale data sets: sequence-based phylogenetic profiling and experiment-based microarray data. Both of these data sets have been used to make functional predictions about interactions in the genome. In this section, we highlight the advances that have been made using these two methods.
Functional Predictions Using Phylogenomic Mapping The availability of a large number of bacterial genomes presents an attractive data set that can be exploited to explore a hypothesis of bacterial evolution: coinheritance. Coinheritance posits that bacterial genomes retain groups of genes that participate in a specific function and dispense with genes that do not (Pellegrini et al., 1999). Based on this hypothesis, the genes of a given bacterial species can be compared against the genes of other bacterial species to construct a phylogenetic profile for each gene within the bacterial species; this is done by assessing the presence or absence of a particular gene’s homolog in every other bacterial species. A clustering algorithm can be applied to this set of phylogenetic profiles and thereby group genes that have similar profiles. In this way, genes that participate in the same function can be predicted. This methodology, termed phylogenomic mapping, has been applied to the genome of M . xanthus by Srinivasan and coworkers (2005).The predicted proteome of M . xanthus was aligned against a local database containing the protein sequences of 205 sequenced bacteria by using BLASTP (Altschul et al., 1997), as shown in Color Plate 3a. A matrix of raw bit scores was constructed with each row corresponding to a predicted M. xanthus protein and each column representing a different sequenced bacterial strain (Color Plate 3b). Spearman’s rank correlation was then used to produce a similarity matrix containing correlation scores (Color Plate 3c). The top
307
50 positive correlates for each protein in the M. xanthus genome were retained, and a combination of multidimensional scaling and force-directed placement was applied to assign each protein an ( x , y) coordinate in a plane. The resulting graph was visualized as a topographical map in three dimensions using the computer program VxInsight (Davidson et al., 2001), as shown in Color Plate 3d. Each mountain on this map represents clusters of proteins that share similar evolutionary histories, and presumably similar functions. To experimentally validate this approach, selected proteins known to be associated with motility were identified in five mountains on the map. A total of 15 uncharacterized proteins were selected from these mountains (Color Plate 4), their associated ORFs were inactivated, and each resulting strain was assayed for defects in motility. A total of 12 ORFs (or 80%) were found to be defective for either A- or S-motility. Analyses of the putative annotations of these proteins reveal that many are involved in cell division, cell wall biosynthesis, and transport. Interestingly, a number of ORFs are homologous to proteins involved in the To1 transport system, consistent with the findings of Youderian and coworkers (2003) for proteins associated with A-motility. This method presents an exciting approach for the detection of functionally related proteins and is currently being utilized by members of the M. xanthus community to make further functional predictions.
Functional Predictions Using Gene Expression Mapping Microarray experiments are typically used to investigate the gene expression patterns of a given gene under specific environmental conditions. As discussed previously, the M . xanthus microarray has been used to monitor the expression levels of genes during vegetative growth (Diodati et al., 2006). When analyzing microarray data, only those genes that exhibit up- and down-regulation, relative to the reference sample, are typically considered. However, there is a wealth of additional data that accompanies any microarray experiment: gene expression levels are recorded for each gene on the microarray. These data can be harnessed and used to make functional predictions through the process of gene expression mapping, a technique similar to that of phylogenomic mapping. The central idea behind gene expression mapping is to identify those genes that share similar expression patterns regardless of the experimental conditions. Presumably, sets of genes that are turned on or off at the same time in response to changing environmental conditions represent genes that are under tightly linked transcriptional control and therefore may be functionally linked.
308 This idea, first introduced by Kim and coworkers (2OO1),was applied to the nematode Caenorhabditiseleguns. They combined over 500 sets of microarray experiments and constructed a topographical map, which was then visualized using VxInsight. Analysis of this map revealed that many genes known to be involved in the same function were found clustered into specific mountains, such as those genes involved in spermatogenesis. Recently Suen and coworkers (2006)produced a gene expression map for M . xanthus based on over 200 microarray experiments. Construction of this map followed the same process outlined for phylogenomics. Briefly, a log (cy3/cy5) matrix of intensity ratios was constructed with each row corresponding to each ORF represented on the microarray and each column representing a specific microarray experiment. This matrix was then clustered using Spearman’s rank correlation to generate a similarity matrix. The top 50 positive correlates for each ORF were retained and a combination of multidimensional scaling and force-directed placement was used to assign each ORF an ( x , y) coordinate on a plane. This resulting graph was then visualized using VxInsight as a topographical map where each mountain represents a clustered set of ORFs. Preliminary analysis of this map shows that genes involved in the same process such as transcription and translation were found to cluster in a single mountain. This map has since been distributed to various labs in the M . xanthus community, who are using it to make functional predictions.
MYXOBACTERIAL DATABASE RESOURCES Most postgenomic applications focus on utilizing the genome to solve problems, such as identifying genes involved in S-motility. One of the ultimate goals of myxobacterial genomics is to determine the complete set of genetic networks that contribute to the complex life cycle of the myxobacteria. The next era of postgenomics will be accompanied by an influx of massive amounts of data which must be managed and processed. Making sense of these data will require the concerted effort of the whole community of those involved in myxobacterial research to pool resources and share data. Several model organism research communities have facilitated this endeavor by constructing databases, and the maturity of any model organism’s genomic development often coincides with the establishment of a community database made available on the World Wide Web. Publicly accessible databases currently exist for a number of model organisms including Dictyostelium discoideum (Chisholm et al., 2006), C. eleguns (Schwarz et al., 2006), S . cerevisiae (Dwight et al., 2004), Mus musculus
MYXOBACTERIAL GENOMICS AND POSTGENOMICS (Eppig et al., 2005), Drosophila melanogaster (Grumbling and Strelets, 2006), and Escherichia coli (Keseler et al., 2005). A similar community-driven database has been developed for M . xanthus.
XanthusBase: A Community-Driven Online Database’ At the 32nd International Conference on the Biology of the Myxobacteria in Harrison Hot Springs, British Columbia, Canada, the community of myxobacterial researchers decided to support a project aimed at establishing an online database for M . xanthus. This project has spawned the creation of XanthusBase (available at http://xanthusbase.org) (Arshinoff et al., 2007), a publicly accessible online database that currently houses the majority of genomic resources related to M. xanthus. This database contains many of the elements found in other model organism databases: an online browser for viewing the genome, profiles for each gene within the genome, a database of publications relating to M . xanthus, and an M . xanthus genome BLAST server. From an extensive analysis of the currently available database platforms, the model currently employed by the D. discoideum community, http://dictybase.org (Chisholm et al., 2006), was selected as the model for the implementation of XanthusBase. This model is implemented using the collection of database software packages collectively known as The Generic Model Organism Database Project (Stein et al., 2002), a set of implementation tools that is quickly becoming standard in the model organism database community. In addition to the basic services offered by XanthusBase, M. xanthus has included a community-driven annotation scheme which departs from the traditional database model of manual curation by a single laboratory (Stein, 2001). While technical maintenance of the physical database is handled by a single laboratory, it was decided that opening up the complete database to the community would be a practical model for the curation of M . xanthus’s genome for a number of reasons. With the proliferation of postgenomic activity, genes within the genome are being characterized at a rapid pace, with respect to both function and annotation, and it is expected that some type of annotation will be available for every gene in the genome in the very near future. It is also accepted that each laboratory within the M . xanthus community is an authority on different aspects of M . xanthus biology. Based on these factors, XanthusBase has incorporated a recent community-driven model of curation which has been widely successful: the Wikipedia model (available at http://Wikipedia.org). The Wikipedia model is centered on an open-source philosophy, where any expert
17. POSTGENOMIC OVERVIEW OF THE MYXOBACTERIA can annotate and post information pertaining to any topic. The database simply houses each Wikipedia entry as it is created o r updated over time. A recent article in Nature explored the Wikipedia phenomenon and found that content on Wikipedia “comes close to Britannica in terms of the accuracy of its science entries” (Giles, 2006). While this model appears successful in terms of its utility, it has yet to find its way into the biological database community. For XanthusBase, the Wikipedia model of curation allows any member of the community to update and make changes to the annotations of genes within the genome. Entries associated with every change are instantaneously posted online while seamlessly integrated into the database. Given the expertise of the community, it is expected that a Wikipedia-style model would facilitate the dissemination of M. xanthus research to the community as a whole.
CONCLUSIONS AND THE FUTURE OF MYXOBACTERIAL POSTGENOMICS In this chapter, we have presented an overview of postgenomics in the myxobacteria with a particular focus on M. xanthus, which has the most well-developed set of postgenomics tools. The application of random mutagenesis and Y2H screens coupled with paralogous analysis of gene families have greatly increased o u r understanding of the genetic networks that result in the complex phenotypes exhibited by M. xanthus. The underlying theme of these studies centers on the complexity of M. xanthus’s signaling pathways. Many signals appear to be co-opted by M. xanthus to control multiple genetic networks, and this topography may suggest how multicellularity evolved. Much of the experimental biology is greatly facilitated by the availability of a genome sequence, and we expect the rapid characterization of a large number of genetic networks in the very near future. In addition to these techniques, high-throughput methods for exploring the M. xanthus genome are also available. Exploratory work using microarrays is currently under way in an effort to identify the temporal expression of genes under a host of different environmental conditions. This work has thus far yielded insight into the number of genes associated with TAs under vegetative conditions. Microarray technology is currently being utilized by a number of researchers in the M. xanthus community, and we expect further insight into the classes of genes that participate in M. xanthus’s complex life cycle in the next few years. To further harness the power of genome scale experiments, functional genomics tools such as phylogenomic and gene expression mapping have also been developed
309
to predict putative interactions within the genome. These mapping tools have been used to successfully predict genes associated with motility, and serve as a useful complement to the experimental tools discussed in this chapter. As postgenomics in M. xanthus continues to mature, we expect that a host of other experimental and functional genomics techniques will also be developed, allowing for the rapid identification of the genetic interactions that exist within the organism. One of the major challenges of the postgenomics era will be to effectively manage the large amount of data that will soon be made available. This is currently being accomplished through the development of the community-driven database XanthusBase. While this database is modeled after existing model organism databases, XanthusBase incorporates a Wikipedia-style annotation approach, allowing the whole community to participate in the management of this data. M. xanthus research is now poised to make major strides in understanding the genomics of this complex organism. As new techniques are developed, we will come ever closer to uncovering the vast network of interacting genes that allow M. xanthus to maintain its social lifestyle in response to its changing environment.
References Altschul, S. F., T. L. Madden, A. A. Schaffer, J. Zhang, Z . Zhang, W. Miller, and D. J. Lipman. 1997. Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res. 25:3389-3402. Arshinoff, B. I., G. Suen, E. M. Just, S. M. Merchant, W. A. Kibbe, R. L. Chisholm, and R. D. Welch. 2007. Xanthusbase: adapting wikipedia principles to a model organism database. Nucleic Acids Res. 35:D422-D426. Bowden, M. G., and H. B. Kaplan. 1998. The Myxococcus xanthus lipopolysaccharide 0-antigen is required for social motility and multicellular development. Mol. Microbiol. 301275-284. Browning, D. F., D. E. Whitworth, and D. A. Hodgson. 2003. Light-induced carotenogenesis in Myxococcus xanthus: functional characterization of the ECF sigma factor CarQ and antisigma factor CarR. Mol. Microbiol. 48:237-251. Buck, M. J., and J. D. Lieb. 2004. ChIP-chip: considerations for the design, analysis, and application of genome-wide chromatin immunoprecipitation experiments. Genomics 83~349-360. Caberoy, N. B., R. D. Welch, J. S. Jakobsen, S. C. Slater, and A. G. Garza. 2003. Global mutational analysis of NtrC-like activators in Myxococcus xanthus: identifying activator mutants defective for motility and fruiting body development. J. Bacteriol. 185:6083-6094. Chisholm, R. L., P. Gaudet, E. M. Just, I<. E. Pilcher, l? Fey, S. N. Merchant, and W. A. Kibbe. 2006. dictyBase, the model organism database for Dictyostelium discoideum. Nucleic Acids Res. 34:D423-D427.
320 Davidson, G., B. Wylie, and K. Boyack. 2001. Cluster stability and the use of noise in interpretation of clustering. p. 23-30. In Proceedings of the IEEE Symposium on Information Visualization (INFOVIS’Ol), October 22-23, 2001. IEEE Computer Society, Washington, DC. Diodati, M. E., F. Ossa, N. B. Caberoy, I. R. Jose, W. Hiraiwa, M. M. Igo, M. Singer, and A. G. Garza. 2006. Nla18, a key regulatory protein required for normal growth and development of Myxococcus xanthus. J. Bacteriol. 188:1733-1743. Dwight, S. S., R. Balakrishnan, K. R. Christie, M. C. Costanzo, K. Dolinski, S. R. Engel,B.Feierbach,D. G.Fisk, J. Hirschman, E. L. Hong, L. Issel-Tarver, R. S. Nash, A. Sethuraman, B. Starr, C. L. Theesfeld, R. Andrada, G. Binkley, Q. Dong, C. Lane, M. Schroeder, S. Weng, D. Botstein, and J. M. Cherry. 2004. Saccharomyces genome database: underlying principles and organisation. Brief. Bioinform. 5:9-22. Eppig, J. T., C. J. Bult, J. A. Kadin, J. E. Richardson, J. A. Blake, A. Anagnostopoulos, R. M. Baldarelli, M. Baya, J. S. Beal, S. M. Bello, W. J. Boddy, D. W. Bradt, D. L. Burkart, N. E. Butler, J. Campbell, M. A. Cassell, L. E. Corbani, S. L. Cousins, D. J. Dahmen, H. Dene, A. D. Diehl, H. J. Drabkin, K. S. Frazer, P. Frost, L. H. Glass, C. W. Goldsmith, P. L. Grant, M. Lemon-Pierce, J. Lewis, I. Lu, L. J. Maltais, M. McAndrews-Hill, L. McClellan, D. B. Miers, L. A. Miller, L. Ni, J. E. Ormsby, D. Qi, T. B. Reddy, D. J. Reed, B. Richards-Smith, D. R. Shaw, R. Sinclair, C. L. Smith, P. Szauter, M. B. Walker, D. 0. Walton, L. L. Washburn, I. T. Witham, and Y. Zhu. 2005. The Mouse Genome Database (MGD):from genes to mice-a community resource for mouse biology. Nucleic Acids Res. 33:D471-D475. Fiehn, 0. 2002. Metabolomics-the link between genotypes and phenotypes. Plant Mol. Biol. 48:155-171. Fields, S., and 0.Song. 1989. A novel genetic system to detect protein-protein interactions. Nature 340:245-246. Finn, R. D., J. Mistry, B. Schuster-Bockler, S. Griffiths-Jones, V. Hollich, T. Lassmann, S. Moxon, M. Marshall, A. Khanna, R. Durbin, S. R. Eddy, E. L. Sonnhammer, and A. Bateman. 2006. Pfam: clans, web tools and services. Nucleic Acids Res. 34:D247-D251. Ge, H., A. J. Walhout, and M. Vidal. 2003. Integrating ‘omic’ information: a bridge between genomics and systems biology. Trends Genet. 19551-560. Gertz, J., G. Elfond, A. Shustrova, M. Weisinger, M. Pellegrini, S. Cokus, and B. Rothschild. 2003. Inferring protein interactions from phylogenetic distance matrices. Bioinformatics 19~2039-2045. Giles, J. 2006. Internet encyclopaedias go head to head. Nature 438~900-901. Goldman, B. S., W. C. Nierman, D. Kaiser, S. C. Slater, A. S. Durkin, J. A. Eisen, C. M. Ronning, W. B. Barbazuk, M. Blanchard, C. Field, C. Halling, G. Hinkle, 0. Iartchuk, H. S. Kim, C. Mackenzie, R. Madupu, N. Miller, A. Shvartsbeyn, S. A. Sullivan, M. Vaudin, R. Wiegand, and H. B. Kaplan. 2006. Evolution of sensory complexity recorded in a myxobacterial genome. Proc. Natl. Acad. Sci. USA 103:15200-15205. Grumbling, G., and V. Strelets. 2006. FlyBase: anatomical data, images and queries. Nucleic Acids Res. 34:D484-D488. Hanlon, W. A., M. Inouye, and S. Inouye. 1997. Pkn9, a Serl Thr protein kinase involved in the development of Myxococcus xanthus. Mol. Microbiol. 23:459-471.
MYXOBACTERIAL GENOMICS AND POSTGENOMICS Higgs, P. I., K. Cho, D. E. Whitworth, L. S. Evans, and D. R. Zusman. 2005. Four unusual two-component signal transduction homologs, RedC to RedF, are necessary for timely development in Myxococcus xanthus. J. Bacteriol. 187: 8191-8195. Hwang, D., A. G. Rust, S. Ramsey, J. J. Smith, D. M. Leslie, A. D. Weston, P. de Atauri, J. D. Aitchison, L. Hood, A. F. Siegel, and H. Bolouri. 2005a. A data integration methodology for systems biology. Proc. Natl. Acad. Sci. U S A 102~17296-17301. Hwang, D., J. J. Smith, D. M. Leslie, A. D. Weston, A. G. Rust, S. Ramsey, P. de Atauri, A. F. Siegel, H. Bolouri, J. D. Aitchison, and L. Hood. 2005b. A data integration methodology for systems biology: experimental verification. Proc. Natl. Acad. Sci. USA 102:17302-17307. Inouye, S., R. Jain, T. Ueki, H. Nariya, C. Y. Xu., M. Y. Hsu, B. A. Fernandez-Luque, J. Munoz-Dorado, E. Farez-Vidal, and M. Inouye. 2000. A large family of eukaryotic-like protein SeriThr kinases of Myxococcus xanthus, a developmental bacterium. Microb. Comp. Genomics 5:103-120. Jakobsen, J. S., L. Jelsbak, L. Jelsbak, R. D. Welch, C. Cummings, B. Goldman, E. Stark, S. Slater, and D. Kaiser. 2004. Sigma54 enhancer binding proteins and Myxococcus xanthus fruiting body development. J. Bacteriol. 186:4361-4368. Jelsbak, L., M. Givskov, and D. Kaiser. 2005. Enhancer-binding proteins with a forkhead-associated domain and the sigma54 regulon in Myxococcus xanthus fruiting body development. Proc. Natl. Acad. Sci. U S A 102:3010-3015. Kaiser, D. 1986. Control of multicellular development: Dictyostelium and Myxococcus. Annu. Rev. Genet. 20539-566. Keseler, I. M., J. Collado-Vides, S. Gama-Castro, J. Ingraham, S. Paley, I. T. Paulsen, M. Peralta-Gil, and P. D. Karp. 2005. EcoCyc: a comprehensive database resource for Escherichia coli. Nucleic Acids Res. 33:D334-D337. Kim, S. K., J. Lund, M. Kiraly, K. Duke, M. Jiang, et al. 2001. A gene expression map for Caenorhabditis elegans. Science 293:2087-2092. LaBaer, J., and N. Ramachandran. 2005. Protein microarrays as tools for functional proteomics. Curr. Opin. Chem. Biol. 9: 14-19. Lancero, H. L., S. Castaneda, N. B. Caberoy, X. Ma, A. G. Garza, and W. Shi. 2005. Analysing protein-protein interactions of the Myxococcusxanthus Dif signallingpathway using the yeast two-hybrid system. Microbiology 151:1535-1541. Li, Y., H. Sun, X. Ma, A. Lu, R. Lux, D. Zusman, and W. Shi. 2003. Extracellular polysaccharides mediate pilus retraction during social motility of Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 1005443-5448. Lockhart, D. J., and E. A. Winzeler. 2000. Genomics, gene expression and DNA arrays. Nature 4055327-836. Lu, A., K. Cho, W. P. Black, X. Y. Duan, R. Lux, Z. Yang, H. B. Kaplan, D. R. Zusman, and W. Shi. 2005. Exopolysaccharide biosynthesis genes required for social motility in Myxococcus xanthus. Mol. Microbiol. 55:206-220. Lux, R., and W. Shi. 2005. A novel bacterial signalling system with a combination of a Ser/Thr kinase cascade and a His/ Asp two-component system. Mol. Microbiol. 58:345-348. MacNeil, S. D., F. Calara, and P. L. Hartzell. 1994. New clusters of genes required for gliding motility in Myxococcus xanthus. Mol. Microbiol. 14:61-71.
17. POSTGENOMIC OVERVIEW OF THE MYXOBACTERIA Mahillon, J., and M. Chandler. 1998. Insertion sequences. Microbiol. Mol. Biol. Rev. 62:725-774. Marcotte, E., and S. Date. 2001. Exploiting big biology: integrating large-scale biological data for function inference. Brief. Bioinform. 2:363-374. Marcotte, E. M., M. Pellegrini, H. L. Ng, D. W. Rice, T. 0. Yeates, and D. Eisenberg. 1999. Detecting protein function and protein-protein interactions from genome sequences. Science 285:751-753. Mignot, T., J. P. Merlie, Jr., and D. R. Zusman. 2005. Regulated pole-to-pole oscillations of a bacterial gliding motility protein. Science 310:855-857. Morett, E., and L. Segovia. 1993. The sigma 54 bacterial enhancer-binding protein family: mechanism of action and phylogenetic relationship of their functional domains. J. Bacteriol. 175:6067-6074. Munoz-Dorado, J., S. Inouye, and M. Inouye. 1991. A gene encoding a protein serinekhreonine kinase is required for normal development of M. xanthus, a gram-negative bacterium. Cell 67:995-1006. Nariya, H., and S. Inouye. 2005. Modulating factors for the Pkn4 kinase cascade in regulating 6-phosphofructokinase in Myxococcus xanthus. Mol. Microbiol. 56:13 14-1 328. Nudleman, E., D. Wall, and D. Kaiser. 2005. Cell-to-cell transfer of bacterial outer membrane lipoproteins. Science 309~125-127. Overbeek, R., M. Fonstein, M. DSouza, G. D. Pusch, and N. Maltsev. 1999. The use of gene clusters to infer functional coupling. Proc. Natl. Acad. Sci. USA 96:2896-2901. Pazos, F., and A. Valencia. 2002. In silico two-hybrid system for the selection of physically interacting protein pairs. Proteins 47:219-227. Pellegrini, M., E. M. Marcotte, M. J. Thompson, D. Eisenberg, and T. 0. Yeates. 1999. Assigning protein functions by comparative genome analysis: protein phylogenetic profiles. Proc. Natl. Acad. Sci. USA 96:4285-4288. Pham, V. D., C. W. Shebelut, E. J. Zumstein, and M. Singer. 2005. BrgE is a regulator of Myxococcus xanthus development. Mol. Microbiol. 57:762-773. Schwarz, E. M., I. Antoshechkin, C. Bastiani, T. Bieri, D. Blasiar, P. Canaran, J. Chan, N. Chen, W. J. Chen, P. Davis, T. J. Fiedler, L. Girard, T. W. Harris, E. E. Kenny, R. Kishore, D. Lawson, R. Lee, H. M. Muller, C. Nakamura, P. Ozersky, A. Petcherski, A. Rogers, W. Spooner, M. A. Tuli, K. Van Auken, D. Wang, R. Durbin, J. Spieth, L. D. Stein, and P. W. Sternberg. 2006. WormBase: better software, richer content. Nucleic Acids Res. 34:D475-D478. Srinivasan, B. S., N. B. Caberoy, G. Suen, R. G. Taylor, R. Shah, F. Tengra, B. S. Goldman, A. G. Garza, and R. D. Welch. 2005. Functional genome annotation through phylogenomic mapping. Nut. Biotechnol. 23:691-698. Stein, L. 2001. Genome annotation: from sequence to biology. Nut. Rev. Genet. 2:493-503. Stein, L. D., C. Mungall, S. Shu, M. Caudy, M. Mangone, A. Day, E. Nickerson, J. E. Stajich, T. W. Harris, A. Arva, and S. Lewis. 2002. The generic genome browser: a building block for a model organism system database. Genome Res. 12~1599-1610. Suen, G., J. S. Jakobsen, B. S. Goldman, M. Singer, A. G. Garza, and R. D. Welch. 2006. Bacterial postgenomics: the
311
promise and peril of systems biology. J. Bacteriol. 188:79998004. Suen, G., B. I. Arshinoff, R. G. Taylor, and R. D. Welch. 2007. Practical applications of bacterial functional genomics. Biotechnol. Genet. Eng. Rev. 24:213-242. Sun, H., and W. Shi. 2001a. Genetic studies of mrp, a locus essential for cellular aggregation and sporulation of Myxococcus xanthus. J. Bucteriol. 183:4786-4795. Sun, H., and W. Shi. 2001b. Analyses of mrp genes during Myxococcus xanthus development. J. Bacteriol. 183:67336739. Szurmant, H., and G. W. Ordal. 2004. Diversity in chemotaxis mechanisms among the bacteria and archaea. Microbiol. Mol. Biol. Rev. 68:301-319. Thomasson, B., J. Link, A. G. Stassinopoulos, N. Burke, L. Plamann, and P. L. Hartzell. 2002. MglA, a small GTPase, interacts with a tyrosine kinase to control type IV pilimediated motility and development of Myxococcus xanthus. Mol. Microbiol. 46:1399-1413. Udo, H., M. Inouye, and S. Inouye. 1996. Effects of overexpression of Pkn2, a transmembrane protein serinehhreonine kinase, on development of Myxococcus xanthus. J. Bacteriol. 178~6647-6649. Ueki, T., and S. Inouye. 2003. Identification of an activator protein required for the induction of fruA, a gene essential for fruiting body development in Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 10053782-8787. Ueki, T., and S. Inouye. 2006. A novel regulation on developmental gene expression of fruiting body formation in Myxobacteria. Appl. Microbiol. Biotechnol. 72:21-29. Vidal, M. 2001. A biological atlas of functional maps. Cell 104:333-339. Ward, M. J., K. C. Mok, D. P. Astling, H. Lew, and D. R. Zusman. 1998. An ABC transporter plays a developmental aggregation role in Myxococcus xanthus. J. Bacteriol. 1805697-5 703. Whitworth, D. E., and D. A. Hodgson. 2001. Light-induced carotenogenesis in Myxococcus xanthus: evidence that Cars acts as an anti-repressor of CarA. Mol. Microbiol. 42:809-819. Wolgemuth, C., E. Hoiczyk, D. Kaiser, and G. Oster. 2002. How myxobacteria glide. Curr. Biol. 12:369-377. Wu, S. S., and D. Kaiser. 1995. Genetic and functional evidence that Type IV pili are required for social gliding motility in Myxococcus xanthus. Mol. Microbiol. 18547-558. Yang, R., S. Bartle, R. Otto, A. Stassinopoulos, M. Rogers, L. Plamann, and P. Hartzell. 2004. AglZ is a filament-forming coiled-coil protein required for adventurous gliding motility of Myxococcus xunthus. J. Bacteriol. 186:6168-6178. Yang, Z., and Z. Li. 2005. Demonstration of interactions among Myxococcus xanthus Dif chemotaxis-like proteins by the yeast two-hybrid system. Arch. Microbiol. 183:243252. Youderian, P., N. Burke, D. J. White, and P. L. Hartzell. 2003. Identification of genes required for adventurous gliding motility in Myxococcus xanthus with the transposable element mariner. Mol. Microbiol. 49:555-570. Zhang, W., M. Inouye, and S. Inouye. 1996. Reciprocal regulation of the differentiation of Myxococcus xanthus by Pkn5 and Pkn6, eukaryotic-like Ser/Thr protein kinases. Mol. Microbiol. 20:435-447.
Stigmatella and Sorangium
Myxobacteria: Multicellularity and Differentiation Edited by David E. Whitworth 0 2008 ASM Press, Washington, D.C.
Wulf Plaga
The Challenge of Structural Complexity: Stigmatella awantiaca as an Alternative
18
Myxobacterial Model Roland Thaxter was the first who recognized Stigmatella aurantiaca (Chondromyces aurantiaca) as a myxobacterium in 1892 (Color Plate 5) (Thaxter, 1892; Reichenbach and Dworkin, 1969). The genus Stigmatella is distributed in soil worldwide. S. aurantiaca preferably lives on bark and rotting wood (Dawid, 1979,2000)and shows the typical myxobacterial multicellular development which is induced by starvation. Hundred of thousands of vegetative cells of a swarm, which are about 7 pm by 0.7 pm, join in an aggregation center. From each center a highly structured fruiting body develops (Color Plate 6). The fruiting body consists of a stalk and several sporangioles (ca. 50 pm by 35 pm) attached by pedicels at the top (Reichenbach and Dworkin, 1969; Gerth and Reichenbach, 1978). The stalk is most likely a cellular structure consisting of cells and slime, but during maturation of the fruiting body more and more of the cells lyse, resulting in empty tubules (Vasquez et al., 1985; Voelz and Reichenbach, 1969; Grilione and Pangborn, 1975; Stephens and White, 1980b). The sporangioles each contain several thousands of dormant myxospores (Liinsdorf et al., 1995); the size of the spores is about 3 pm by 1 pm (Voelz and Reichenbach, 1969), and one fruiting body contains lo4 to lo5 spores (Neumann
et al., 1993). A high number of cells neither become part of the fruiting body structure nor differentiate to spores but (auto)lyseto support the developmental process under starvation conditions. When the conditions improve again, spores from at least one sporangiole germinate to establish a new swarm. The sporangiole can therefore be considered to be a dissemination unit. At least during the early phase of development the Stigmatella pheromone stigmolone is required (Plaga et al., 1998). Cellular development (spore formation) can be separated from cooperative morphogenesis of the fruiting body. Many substances such as 3-methylindole, indole, n-butanol, or 2-phenylethanol induce the differentiation of vegetative cells into spores without fruiting body formation (Gerth et al., 1993). This shortcut has also been observed with Myxococcus xanthus (Dworkin and Gibson, 1964), but much more detailed knowledge has been accumulated from experiments with S. aurantiaca. A stimulating effect of light and guanine derivatives on fruiting body formation has been reported for S. aurantiaca, especially when low cell densities were used in the experiments (Qualls et al., 1978a; Stephens and White, 1980a; White et al., 1980; Inouye et al., 1989). Some results could be strain specific: when 10 different
Wulf Plaga, Zentrum fur Molekulare Biologie der Universitat Heidelberg (ZMBH), University of Heidelberg, 69120 Heidelberg, Germany.
315
316
S. aurantiaca isolates were analyzed, five unique genomic fingerprints were determined (Neumann et al., 1992). At present most experiments use S. aurantiaca strain DW4/3-1, whose genome sequence has been determined recently (The Institute for Genomic Research). DW4/ 3-1 is a streptomycin-resistant derivative of DW4 which is a dispersed-growing strain derived from strain CCf (Qualls et al., 197813). The species S. aurantiaca has also been extensively studied by electron microscopy of cells, spores, and fruiting bodies (for examples, see Grilione and Pangborn, 1975; Vasquez et al., 1985; Voelz and Reichenbach, 1969; and Stephens and White, 1980b). Seven years ago, high-resolution scanning electron microscopic images revealed helical surface features which were interpreted as gliding-associated surface patterns (Liinsdorf and Schairer, 2001). Similar structures have been suggested to support adventurous gliding (A-motility) of 211. xanthus (Freese et al., 1997; Mignot et al., 2007). However, a second equivalent suggestion is that A-motility relies on nozzle-like structures for slime secretion (Kaiser, 2003).
GENOME AND TRANSCRIPTIONAL AND TRANSLATIONAL MACHINERIES Genome and Retroelements S. aurantiaca has a genome size of 10.27 Mbp (GenBank accession no. AAMD00000000), which is intermediate when compared to M. xanthus (9.14 Mbp; GenBank accession no. CPOOOll3) and Sorangium cellulosum (12.2 Mbp [Pradella et al., 20021). However, among these species Stigmatella clearly forms the most complex fruiting body, and therefore morphological complexity is not correlated directly with genome size. It might well be that diversity of secondary metabolism has the dominant impact on genome size. Retroelements are found in S. aurantiaca and in many other (myxo)bacteria (Dhundale et al., 1985; Rice and Lampson, 1996).An S. aurantiaca cell contains approximately 500 copies of a short single-stranded linear DNA (multicopy single-stranded DNA [msDNA]); its size is 162 or 163 bases. At its 5' end the msDNA is attached to an RNA by a 2',5' linkage. The msDNA is encoded by a retroelement which also contains the gene for a reverse transcriptase responsible for the synthesis of the msDNA (Furuichi et al., 1987a, 1987b;Hsuet al., 1992).Deletion of the retrons has no phenotype in M. xanthus (Inouye et al., 1990), and thus, the function of the elements in myxobacteria remains unknown. For Escherichia coli it was observed that cells producing large amounts of msDNA showed a higher rate of mutation, but the
STIGMATELLA AND SORANGIUM molecular mechanism for this elevated mutation rate is still unknown (Jeong and Lim, 2004).
RNA Polymerase and Sigma Factors The DNA-dependent RNA polymerase with the main IT factor bound has been purified from vegetative cells to homogeneity as analyzed by sodium dodecyl sulfatepolyacrylamide gel electrophoresis with the aim to characterize cr factors and development-specific promoters in vitro. The apparent molecular masses have been determined for the a, p, and p' subunits as well as for the main cr factor to be 40,000, 146,000, 146,000, and 105,000, respectively (Heidelbach et al., 1992). The sigA gene (encoding the major cr factor SigA) and the genes for the alternative cr factors SigB and SigC were cloned and sequenced (Skladny et al., 1992, 1994; Silakowski et al., 2001) (accession numbers M94370, 214970, and SAU27311). SigA shares an identity of about 90% with the major cr factor crSoof M. xanthus and is detected under all physiological conditions tested. SigB and SigC are members of the cr70 protein family. sigB and sigC expression was analyzed by Western blotting and reverse transcription-PCR. Both cr factors were found to be expressed during fruiting body formation and indole-induced sporulation but not after heat shock. The expressions of sigB and sigC appeared to be independent from each other (Coudart, 1998). Independent analyses by Western blotting performed with another serum revealed that the SigB protein was not present in vegetative cells but was present in indole-induced spores, in fruiting bodies, and in cells after heat shock. The antiserum raised against SigB in this study cross-reacted with at least one further alternative cr factor whose expression pattern is similar to that of SigB. SigB expression was specifically determined by analysis of a merodiploid sigB mutant harboring a sigB-lacZ fusion gene, and transcription of sigB was analyzed by reverse transcriptionPCR. The results obtained by these different methods of analysis corroborated the Western blot data including the observation that SigB occurs in cells after heat shock (Silakowski et al., 2001). The inactivation of the sigB gene did not give rise to a phenotype during fruiting body formation or indoleinduced sporulation (Coudart, 1998; Silakowski et al., 2001), and possible SigB targets such as DnaK, GroEL, or HspA were not affected in the mutant. The loss of SigB in the mutant seems to be compensated by other cr factors, which are most likely encoded by the genome: in Southern blot analyses using oligonucleotides encoding conserved regions of cr factors, up to eight hybridizing DNA restriction fragments were detected (Silakowski et al., 2001).
18. S. AURANTIACA AS ALTERNATIVE MYXOBACTERIAL MODEL Transcription and Translation Factors (CarD, IF2, EF-Tu) CarD of M. xanthus is a transcription factor which is essential for fruiting body formation as well as for the expression of the light-inducible carQRS operon in M. xanthus. This operon is involved in the regulation of carotenogenesis. CarD is a prokaryotic analog of eukaryotic high-mobility group A proteins. A carD gene was proven to also exist in S. aurantiaca by Southern hybridization using carD of M. xanthus as a probe. Furthermore, the CarD protein could be detected in Western blot analyses using an antiserum against CarD of M. xanthus. The deduced amino acid sequences of M. xanthus and S. aurantiaca carD show 73% identical residues. Interestingly, most differences in the primary structures localize to the high-mobility group-type domains. Nevertheless the carD gene of S. aurantiaca could complement the carD mutant of M . xanthus concerning fruiting body formation and carotenogenesis (Cayuela et al., 2003). Translation initiation factor 2 (IF2) of S. aurantiaca is encoded by the in@ gene. The C-terminal region and the G domain are homologous to the respective IF2 regions of many bacteria. Remarkably, the N-terminal region is extended and does not show any significant homology to IF2 N termini of E. coli, Bacillus subtilis, Bacillus stearothermophilus, or Streptococcus faecium but does to that of Synechocystis. IF2 of Stigmatella as well as an N-terminally truncated version complemented an E. coli in@ mutant. Possibly the N-terminal part of the IF2 of Stigmatella plays a role in development (Bremaud et al., 1997).IF2 of M. xanthus showed the same characteristics as IF2 of S. aurantiaca (Tiennault-Desbordeset al., 2001). Elongation factor EF-Tu is encoded by the tufB gene of S. aurantiaca, which is cotranscribed with four tRNA
317
genes upstream of tu@. The predicted molecular mass of EF-Tu is 43.4 kDa, and it displays extensive homologies with the EF-Tu of E. coli or Thermus thermophilus (Bremaud e t al., 1995).
ESTABLISHED DEVELOPMENTAL ELEMENTS Stigmolone, a Myxobacterial Pheromone In 1982 a pheromone activity was demonstrated to be important during fruiting body formation of S. aurantiaca (Stephenset al., 1982). Later the requirement of diffusible signal factor(s) was proven by a dialysis experiment. Starving cells were unable to aggregate on a dialysis membrane as long as secreted molecule(s)were dialyzed away (Fig. 1) (Plaga et al., 1998). Vice versa the dialysate mediated an acceleration of fruiting body formation, which was used as a biotest during the purification of the pheromone. Material secreted from 8 X l O I 3 cells was purified to yield finally about 2 pmol of pure pheromone for structure determination. Using mass spectrometry, infrared spectroscopy, lH-nuclear magnetic resonance (NMR) and I3C-NMR, the structure was determined to be 2,5,8-trimethyl-8hydroxy-nonan-4-one (Fig. 2), and the compound was named stigmolone. Chemical synthesis of stigmolone unequivocally demonstrated that stigmolone serves the purpose of a true pheromone by representing an extracellular signal substance required for successful intercellular communication of S. aurantiaca. The dose-response curve showed that stigmolone is biologically active at about 1 nM (Fig. 2). It therefore represents a highly bioactive prokaryotic pheromone with a unique structure. Stigmolone does not accelerate fruiting body formation in bioassays with M . xanthus (Plaga et al., 1998; Hull et al., 1998).
Figure 1 Prevention of fruiting body formation by dialysis. Stigmatella cells on a dialysis membrane were unable to form aggregates during dialysis (A) but aggregated normally after dialysis had been stopped (B). Bar, 5 mm. Reprinted from Plaga et al., 1998.
STIGMATELLA AND SORANGIUM
318
I
I
I I I I
Oll
I
I
I I I I
1!0 stigmolone [nM]
I I I I
1'0
'
Figure 2 Dose-response curve of S. aurantiaca to stigmolone. The inset shows the structural formula of stigmolone. Reprinted from Plaga et al., 1998.
The stigmolone structure shows one asymmetric center, so it would be of interest to investigate whether the racemate is synthesized by Stigmatella or only one of the enantiomers and whether the biological activity resides in only one of the enantiomers or in both. To solve this question, the enantiomers (S)- and (R)-stigmolone were synthesized and tested for biological activity (Morikawa et al., 1998; Enders and Ridder, 2000). The activity of the racemate was equal to that of each of the enantiomers in biotests. Since a racemization under the conditions of the biotests is possible and its extent was not determined, it is not demonstrated yet whether both enantiomers are synthesized in vivo and whether they have the same biological activity.
The fbf Genes, Important for Fruiting Body Formation Genes essential for fruiting body formation (fbf)were searched by screening of transposon insertion mutants defective in development. Four fbf genes (A through D ) were found in a gene cluster where fbfC and fbfD are organized in an operon (Fig. 3 ) . fbfmutants (Table 1) were generated which could form only clumps instead of fruiting bodies. Some of them were partially phenotypically complemented in development by mixing with another developmental mutant, AP191. This mutant is not characterized in detail but is unable to produce aggregates even during starvation (Table 1) (Pospiech et al., 1993; Silakowski et al., 1996, 1998; Muller, 2002).
CsgA of Stigmatella CsgA of M. xanthus is required for rippling, aggregation, and sporulation (Bonner and Shimltets, 2001; see also chapter 4). To investigate the role of this protein in Stigmatella, a csgA gene of S. aurantiaca was cloned using the csgA gene of M . xanthus as a hybridization probe (Butterfag, 1992). The deduced amino acid sequence showed 56% identity to CsgA of M. xanthus. Under starvation conditions a csgA disruption mutant was able to make fruiting bodies like the wild type, and the myxospores formed were able to germinate. However, rippling was abolished completely and the migration and aggregation patterns were altered. When compared to M. xanthus the csgA phenotype of S. aurantiaca is less prominent. Interestingly, it was additionally observed that the csgA mutant did not respond to stigmolone in the standard bioassay (Milosevic, 2003). Since CsgA of Stzgmatella has sequence similarities to short-chain alcohol dehydrogenases and CsgA of M. xanthus is a short-chain alcohol dehydrogenase with unknown substrate specificity (Lee et al., 1995), it could be speculated that stigmolone is a substrate for CsgA in Stigmatella.
'
fbf5
fbfA
'
fbfC
'
fbfD
"
500bp
-
Figure 3 Physical map of the fbfgene cluster of S. auruntiucu.
18.
s. AURANTIACA AS ALTERNATIVE MYXOBACTERIAL MODEL
319
Table 1 The f6fgeiies'
f6f gene A B
C D
Homologous gene product NodC (N-acetylglucosamine transferase of Rhizobza) GaoA (galactose oxidase of Dactylium dendroides)
ORF2 (putative protein of M. xnnchus not involved in fruiting body formation [Harris and Singer, 19981)
Duration (h) of starvation required for expression
Phenotypical complementation by mixing with strain AP193
8"
Partial
14"
Partial
4c; 8" 4c
None None
'>Thedata are taken from Silakowski et al., 1996 (fbfA),Silakowski et al., 1998 (fbfB),and Miiller, 2002 (fbfC and fbfo). "Detection based on P-galactosidase activity of Atrp-lucZ reporter constructs. 'Transcripts detected using reverse transcription-PCR
(Artificial) Induction of Sporulation Cellular and cooperative morphogenesis can artificially be uncoupled. For Stigmatella many inducers belonging to different substance classes are known, and the ultrastructure of the artificially induced myxospores has been analyzed. In this respect more detailed data have been gathered for Stigmatella than for M . xunthus (Reichenbach et al., 1969; Reichenbach and Dworkin, 1970; Gerth and Reichenbach, 1978,1994; Gerth et al., 1993). Fruiting body spores and artificially induced spores are very similar, since merely quantitative differences concerning the capsule, wall folding, and polyphosphate granules were found. They could be explained by the different timing of the two processes: fruiting body formation takes place over many hours, whereas the change of cell shape during artificial spore formation needs only about 15 min (Reichenbach et al., 1969; Voelz and Reichenbach, 1969). Indole and some indole derivatives are very potent examples of the known inducers. The optimum concentrations for induction are 0.1 mM for indole and 0.07 mM for 3-methylindole. On the other hand, structurally related compounds are inhibitors of chemically induced spore formation. For instance, in the presence of 0.3 mM oxindole the induction by 0.1 mM indole was suppressed (Gerth et al., 1993). Gerth et al. (1993)were able to classify the inducers into four groups and postulated three inducer-specific, independent receptors, two of which should interact with indole derivatives. They also speculated that the natural inducer could be a compound of the indole family, since several indole derivatives have been isolated from myxobacteria as secondary metabolites (Bohlendorf et al., 1996). Recently, the pyruvate kinase and an aldehyde dehydrogenase were isolated from S. aurantiaca by exploiting
their capacity to bind indole. The activity of the pyruvate kinase was stimulated in the presence of indole. Sporulation induced by indole was strongly delayed in a p y k A mutant, and the mutant strikingly revealed that pyruvate kinase is essential for multicellular development: the fruiting body formation of the mutant was abolished, and rippling during starvation was never observed (Fig. 4) (Stamm et al., 2005). It could be that the aldehyde dehydrogenase, the second putative indole receptor, is used as a bypass in artificially induced sporulation and therefore it is not abolished in the p y k mutant but delayed only. The characterization of aldehyde dehydrogenase of S. aurantiaca is in progress. Interestingly, homologous pyruvate kinases and aldehyde dehydrogenases have been described in higher organisms, in which they also bind a small hydrophobic molecule, the thyroid hormone, and seem to be involved in developmental processes (Ashizawa and Cheng, 1992; Yamauchi et al., 1999; Yamauchi and Tata, 2001).
PROTEINS AND PROCESSES WHICH COULD CONTRIBUTE TO DEVELOPMENT The Low-Molecular-Weight Heat Shock Protein HspA (SP21) The stress protein HspA (formerly SP21) is synthesized during fruiting body formation, artificially induced sporulation, heat shock, and anoxia. It is a member of the wcrystallin family of low-molecular-weight heat shock proteins (Heidelbach et al., 1993a, 199313). HspA was localized by immunoelectron microscopy using protein A-gold conjugates. In fruiting-body-derived spores the protein was found mainly at the cell wall, in indoleinduced spores it was either at the cell periphery or within the cytoplasm, and in heat-shocked cells it was
STIGMATELLA AND SORANGIUM
320
Figure 4 Developmental phenotype of p y k A mutant (A) in comparison to the wild type (B). The fruiting bodies of the wild-type strain are visible in the left part of panel B. Bars, 1 mm. Reprinted, with permission, from Stamm et al., 2005. found at the cell periphery. HspA was also observed to be associated with cellular remnants in the stalk of fruiting bodies (Liinsdorf et al., 1995). An hspA deletion mutant behaved like the wild type during vegetative growth, fruiting body formation, sporulation, and spore germination; the thermotolerance was also not affected in the mutant. The extent of oligomerization of recombinant HspA (HspA,,,) was determined by size exclusion chromatography, since other small heat shock proteins are known to oligomerize. The predominant species found corresponded to a molecular mass of 560 kDa, which reflects a complex of about 25 HspA molecules. This oligomeric HspA,,, was able to interact with unfolded citrate synthase and prevented its precipitation, but the enzyme activity was not recovered. An interaction of HspA with the unfolded B chain of insulin was not observed. Nevertheless the data could hint at a chaperone function of HspA (Shen and Schairer, 1999; Shen, 1999) as known for other members of the lowmolecular-weight heat shock proteins (see Bukau, 1999).
Adenylyl Cyclases The concentration of cyclic AMP (CAMP)was measured intra- as well as extracellularly during starvation in liquid medium. During the first hour the intracellular concentration increased by a factor of two. Then, during the next 4 h, the intracellular concentration declined to one-half of the initial concentration by excretion. The extracellular cAMP is degraded. Synthesis of cAMP is accomplished by adenylyl cyclases, and two adenylyl cyclases, AC1 and AC2 of Stigmatella, encoded by the genes cyaA and cyaB, have been cloned (Coudart-Cavalli et al., 1997). Complementation of a cya mutant of E. coli was exploited to clone both genes. Because of sequence similarities AC1
and AC2 belong to class I11 adenylyl cyclases (Danchin, 1993). The enzyme activities of AC1 and AC2 are inhibited by adenosine, which is also an inhibitor known for other adenylyl cyclases. Adenosine is a cell density signal in M. xanthus (Shimkets and Dworkin, 1981) and could also play an important role in Stigmatella development, possibly via its impact on adenylyl cyclase activity (Coudart-Cavalli et al., 1997).
Glucosaminidase An endo-N-acetyl-P-D-glucosaminidasewas purified from culture medium of S. aurantiaca to homogeneity, to investigate a possible connection between glycoprotein metabolism and development. This enzyme does not act as a murein hydrolase but has glycoproteins and glycoasparagines as substrates. In vivo it might be used to eliminate oligosaccharide moieties from polypeptides contained in the food to facilitate the degradation by proteases, or it could be responsible for the release of developmental signals from secreted N-glycosylproteins (Bourgerie et al., 1994).In M. xanthus this enzyme activity was found to be secreted during vegetative growth as well as during development. Importantly, the secreted activity was higher during development, especially during spore development, which may reflect an implication in the maturation of the spore coat (Barreaud et al., 1995).
Inositide Degradation and Inositol Phospholipid Synthesis Stimulated by the role of inositol phosphates in eukaryotic signal transduction, inositol phospholipid synthesis and degradation in StigmatelEa were analyzed. Inositol phospholipid synthesis and degradation were stimulated during starvation in liquid medium containing Ca2+, conditions which promote clumping of Stigmatella cells.
18. S. AURANTIACA AS ALTERNATIVE MYXOBACTERIAL MODEL Furthermore inositol phosphate and inositol bisphosphate were formed, which could have a role as second messengers. In addition a phospholipase C activity increased, stimulated by GTPyS. This prompted the speculation that an interaction with a G protein could be involved (Benaissa et al., 1994).
A Membrane-Associated GTP-Binding Protein G proteins are important transducers in eukaryotic cells, and therefore, it was investigated whether S . aurantiaca harbors G proteins. In a photoaffinity labeling experiment membrane preparations were treated with radioactively labeled GTP. One 54-kDa polypeptide was labeled by this procedure. The GTP binding was specific since an excess of ATP, CTP, or UTP did not interfere; moreover, GDP competed with GTP. These data were interpreted as evidence for the existence of an 01 subunit of a G protein in S. aurantiaca, comparable to the G, proteins in eukaryotes (Dtrijard et al., 1989). Since G proteins can be connected to phosphoinositide metabolism or other signal transduction pathways, a G protein in Stigmatella could participate in signal transduction during development. It should be noted, however, that specific binding of GTP is not absolutely indicative of a true G protein.
SECONDARY METABOLITES AND CHEMICAL COMPOSITION Secondary Metabolites Stigmatella is a rich source of secondary metabolites, which is true for myxobacteria in general (Fig. 5) (Reichenbach, 2001). Only a few data of the Stigmatella
321
system are discussed here since a comprehensive overview is given in chapter 15. Volatiles which are released by S. aurantiaca were analyzed by gas chromatographymass spectrometry. The substances found represent such different compound classes as ketones, esters, lactones, terpenes, and sulfur and nitrogen compounds (Dickschat et al., 2 0 0 5 ~ )Secondary . metabolites which exert some biological effect affect mainly electron transport. Examples of such compounds are aurafuron, aurachin, myxalamid, myxochromide, myxothiazol, and stigmatellin. Structurally most of these compounds are polyketides, peptides, or terpenoids. Eukaryotes are more susceptible to the various substances than gram-positive prokaryotes; gram-negative prokaryotes are hardly affected. Besides electron transport the cytoskeleton and polymerases are well-known targets for biologically active myxobacterial secondary metabolites (Kunze et al., 1984, 1987, 2005; Beyer et al., 1999; Wenzel et al., 2005; Silakowski et al., 1999). Many of the compounds synthesized by Stigmatella are also produced by other organisms. For instance, the well-known earthy smelling compound geosmin is produced by S. aurantiaca as well as by Nannocystis exedens, Streptomyces grz'seus, fungi, plants, etc. (Trowitzsch et al., 1981; Dickschat et al., 2005a). Gene loci associated with the synthesis of secondary metabolites can be interesting targets for genetic experiments. The mtaB gene, part of the gene cluster responsible for myxothiazol synthesis, could be used as a locus for ectopic complementations of, e.g., developmentally relevant genes, because it is not involved in development. Remarkably, the mta cluster is located only about
CONHp
Myxothiazol
Aurachin
I
OCH3 OCH3
0HQCO
0
OH
Myxalamid
Stigmatellin
Geosrnin
Figure 5
Examples of secondary metabolites of S. uuruntzucu.
STIGMATELLAAND SORANGIUM
322 800 bp downstream of the developmentally important
fbfB locus (Silakowski et al., 1998, 1999; Stamm et al., 2005).
Lipids and Pigments The main phospholipids of vegetative S. aurantiaca cells are phosphatidylethanolamine (50%), phosphatidylinositol (20%), lysophosphatidylethanolamine (17%), and phosphatidylglycerol (12%). Alkyl ether linkages in phospholipid species of Stigmatella were found (Caillon et al., 1983; Reichenbach and Dworkin, 1992). Only about 40% of the fatty acid content of a Stigmatella cell is bound in phospholipids (Schroder and Reichenbach, 1970).The majority of fatty acids found are oddnumbered, isobranched, and unsaturated. Only small amounts of even-numbered and unbranched fatty acids are present; 2-hydroxy and 3-hydroxy fatty acids are also found (Fautz et al., 1979; Dickschat et al., 2005b; Reichenbach and Dworkin, 1992). The fatty acid composition does not change significantly during myxospore formation (Schroder and Reichenbach, 1970). The synthesis of pigments is stimulated by light in Stigmatella. Carotenoids are among the main pigments of most myxobacteria. For Stigmatella 1',2'-dihydro-1'hydroxy-3,4-dehydro-toruleneglucoside (myxobactin) and 1',2'-dihydro- 1'-hydroxy-4-keto-torulene glucoside (myxobacton) are the two main carotenoids which occur as monoesters of various fatty acids and account for at least 80% of the total carotenoids (Kleinig and Reichenbach, 1970). Nine minor carotenoids were additionally identified, which account for about 10% of the total carotenoids (Kleinig and Reichenbach, 1969). Melanoid pigments seem also to be produced by Stigmatella because liquid cultures turn black soon after reaching the stationary phase (Reichenbach and Dworkin, 1969,1992).
METHODS S. aurantiaca DW4/3-1 cells can be grown either dispersed in liquid tryptone medium or on solid media (Plaga et al., 1998). The generation time in liquid medium at 32°C is 6 to 7 h. Vegetative cells from a liquid culture reproducibly form fruiting bodies if transferred to a solid starvation medium as follows: cells from the liquid culture (about 1.5 X lo8 cells/ml) are harvested by centrifugation, washed with buffer at pH 7.2 (100 mM HEPES/NaOH, 10 mM CaCl,), and suspended in this buffer to a final density of 4 X 1O1O cells/ ml. Volumes of 5 or 10 pl are spotted onto starvation agar plates (1.5% agar, 6.8 m M CaCl,), or on filter paper squares (Whatman 3MM Chr) sitting on this agar.
Excess liquid is dried away in a hood, and the plates are then incubated at 32°C with illumination. Fruiting bodies appear after 20 to 25 h (Plaga et al., 1998; Stamm et al., 1999). The process of fruiting body formation can be well approached by molecular genetic methods since many of them are well established for Stigmatella. It is nowadays certainly true that S. aurantiaca and M . xanthus are similarly amenable to genetic experiments. However, a problem that still persists for both organisms is the lack of replicating plasmids.
Mutagenesis When molecular genetic work with Stigmatella was in its beginnings, transfer of DNA was performed by conjugation with E. coli using IncP plasmids or derivatives of the conjugative plasmid pSUP102 (Glomp et al., 1988; Pospiech et al., 1993). This method for DNA transfer was replaced by electroporation when an appropriate protocol became available (Stamm et al., 1999). Random mutagenization of S. aurantiaca, to isolate developmental mutants and identify developmentally regulated genes, was achieved using a Tn5-based transposon (Pospiech et al., 1993). This screening identified the fbf gene cluster and generated the mutant AP191 (see above). Once the electroporation method for DNA transfer had been established, random plasmid insertions into the genome were easily feasible. A genomic plasmid library comprising 400- to 2,000-bp fragments could be successfully used for insertions by homologous recombination (Stamm and Plaga, 2000). Furthermore, this strategy also allowed defined gene disruptions even when only 126 bp of homologous DNA mediated the recombination event (Stamm et al., 2005). Markerless in-frame deletions in any nonessential gene can be made using sacB of B. subtilis for counterselection as introduced for M. xanthus (Wu and Kaiser, 1996; Weinig et al., 2003).
Integration of Plasmids into the attB Site The phage attachment site attB of M. xanthus was used for ectopic gene expression by integration of plasmids containing the Mx8 intP-attP gene (Li and Shimkets, 1988; Fisseha et al., 1996). Therefore, the attB site of S . aurantiaca was characterized in detail (Miiller et al., 2006). The attB site of S. aurantiaca is located in the trnD gene (encoding tRNAAsp)which is found in an operon with the trnV gene (encoding tRNAV"')(Fig. 6). Integration of plasmids harboring the Mx8 intP-attP gene resulted in a strong developmental defect. Possibly, the decreased expression of trnVD and the block in the processing of the trnD transcript caused by the integration event hamper protein synthesis during
18. S. AURANTIACA AS ALTERNATIVE MYXOBACTERIAL MODEL attB
trnD
323
several consecutive rounds of sorting, alternately with or without stigmolone addition.
trn V
Figure 6 Physical map of the attB locus of S. uuruntzucu. The uttB site (gray square) is located in the trnD gene. Reprinted, with permission, from Muller e t al., 2006.
development (Muller et al., 2006). In contrast, no developmental phenotype was reported for M . xanthus strains which had integrated such plasmids at the attB site. However, the activities of several promoters were reduced when expressed from this site in M. xanthus (Fisseha et al., 1996). Taken together, integration of plasmids at the attB site is of limited use in Stigmatella and unsuitable for most developmental studies.
Use of the cspA Promoter A cold-shock-like protein of S. aurantiaca, CspA, was characterized, and during the study it was found that the cspA promoter is rather strong, effecting transcription at high levels during vegetative growth as well as development (Stamm et al., 1999). To test the suitability of this promoter for heterologous gene expression in Stigmatella, a gfp gene was expressed under its control. The gfp-expressing cells were easily detected by their prominent green fluorescence, and they were not affected in development (Stamm and Plaga, 2000). Therefore, the cspA promoter seems to be useful to express heterologous genes in Stigmatella. For instance, it could be used to introduce new resistance genes which are not expressed from their own promoters in Stigmatella. This could facilitate the generation of strains with several genes inactivated simultaneously.
Use of a Promoter Trap Vector for Differential Fluorescence Induction Stigmatella cells harboring one gfp copy under the control of the cspA promoter in their genome can be distinguished from wild-type cells by using a fluorescenceactivated cell sorter. A promoter trap vector pTRAPl was constructed which allows the creation of gene fusions to a promoterless gfp gene. This vector was randomly integrated into the genome as a promoter probe (Stamm and Plaga, 2000). The resulting Stigrnatella strains were sorted using the fluorescence-activated cell sorting technique (Hauer and Eipel, 1997) with the final goal to isolate strains which have the promoter probe integrated downstream of a stigmolone-regulated promoter. At least an enrichment of such strains was achieved after
2D Analysis of the Proteome The proteome of S. aurantiaca at different developmental stages and after artificially induced sporulation was analyzed by two-dimensional (2D) electrophoresis. As expected it could be shown that the protein patterns change remarkably during development and sporulation (Hofmann, 2004). 2D electrophoretic analysis of such changes will be a very powerful method to analyze development in general as well as the effects of defined mutations on the protein pattern of the mutant cell. This is especially true since the recent availability of the genome sequence greatly facilitates the annotation of the various protein spots.
CONCLUDING REMARKS The most prominent feature of S. aurantiaca is the formation of a complex and highly structured fruiting body. This complexity represents the specific challenge of the Stigmatella system and distinguishes Stigmatella from Myxococcus and Sorangium. Many intermediate steps during fruiting body formation can be defined, and the influence of mutations on each of them can be studied. Research on the molecular biology of Stigmatella started later than that of Myxococcus, but much information has been gathered during the last decade. At present genetic tools are similarly available for the two organisms. The pheromone stigmolone represents a unique feature of the Stigmatella system. Its biosynthesis needs to be elucidated, as does the downstream processing of the stigmolone signal. This will certainly establish a new and specific signal transduction chain and also reveal how stigmolone signaling is used for cell communication. The sporulation process per se may be studied further using artificial spore induction. This cellular differentiation has been figured out in more detail for Stigmatella than for Myxococcus. Results obtained with the model organism Stigmatella may stimulate the work on M . xanthus in the future as did work on M. xanthus for research on Stigmatella in the past. Since both genomes have been sequenced, comparisons of the two systems are greatly facilitated now and many new results will certainly be obtained from interspecies genetic complementation experiments. Along these lines it might even become apparent which genetic elements shape the fascinating fruiting body of Stigmatella.
324
References Ashizawa, K., and S.-Y. Cheng. 1992. Regulation of thyroid hormone receptor-mediated transcription by a cytosol protein. Proc. Natl. Acad. Sci. USA 89:9277-9281. Barreaud, J.-P., S. Bourgerie, R. Julien, J. F. Guespin-Michel, and Y. Karamanos. 1995. An endo-N-acetyl-P-D-glucosaminidase, acting on the di-N-acetylchitobiosyl part of Nlinked glycans, is secreted during sporulation of Myxococcus xanthus. J. Bacteriol. 177:916-920. BenaYssa, M., J. Vieyres-Lubochinsky, R. Odtide, and B. Lubochinsky. 1994. Stimulation of inositide degradation in clumping Stigmatella aurantiaca. J. Bacteriol. 176:13901393. Beyer, S., B. Kunze, B. Silakowski, and R. Miiller. 1999. Metabolic diversity in myxobacteria: identification of the myxalamid and the stigmatellin biosynthetic gene cluster of Stigmatella aurantiaca Sg a15 and a combined polyketide(po1y)peptide gene cluster from the epothilone producing strain Sorangium cellulosum So ce90. Biochim. Biophys. Acta 1445:185-195. Bohlendorf, B., E. Forche, N. Bedorf, K. Gerth, H. Irschik, R. Jansen, B. Kunze, W. Trowitzsch-Kienast, H. Reichenbach, and G. Hofle. 1996. Indole and quinoline derivatives as metabolites of tryptophan in myxobacteria. Liebigs Ann. 1996~49-53. Bonner, P. J., and L. J. Shimkets. 2001. Piecing together a puzzling pathway: new insights into C-signaling. Trends Microbiol. 9:462-464. Bourgerie, S., Y. Karamanos, T. Grard, and R. Julien. 1994. Purification and characterization of an endo-N-acetyl-p-Dglucosaminidase from the culture medium of Stigmatella aurantiaca DW4. J. Bacteriol. 176:6170-6174. Bremaud, L., C. Fremaux, S. Laalami, and Y. Cenatiempo. 1995. Genetic and molecular analysis of the t-RNA-tufB operon of the myxobacterium Stigmatella aurantiaca. Nucleic Acids Res. 23:1737-1743. Bremaud, L., S. Laalami, B. Derijard, and Y. Cenatiempo. 1997. Translation initiation factor IF2 of the myxobacterium Stigmatella aurantiaca: presence of a single species with an unusual N-terminal sequence. J. Bacteriol. 179:2348-2355. Bukau, B. (ed.). 1999. Molecular Chaperones and Folding Catalysts: Regulation, Cellular Function and Mechanisms. Harwood Academic Publishers, Amsterdam, The Netherlands. Butterfag, H.-J. 1992. Isolierung und Charakterisierung des csgA-Genes aus Stigmatella aurantiaca. Diploma thesis. Ruprecht-Karls-Universitat, Heidelberg, Germany. Caillon, E., B. Lubochinsky, and D. Rigomier. 1983. Occurrence of dialkyl ether phospholipids in Stigmatella aurantiaca DW4. J. Bacteriol. 153:1348-1351. Cayuela, M. L., M. Elias-Amanz, M. Peiialver-Mellado, S. Padmanabhan, and F. J. Murillo. 2003. The Stigmatella aurantiaca homolog of Myxococcus xanthus high-mobility-group A-type transcription factor CarD: insights into the functional modules of CarD and their distribution in bacteria. J. Bacterial. 185:3527-3537. Coudart, M.-P. 1998. Independent patterns of expression of two alternative sigma factors, SigB and SigC, of the myxobacterium Stigmatella aurantiaca during development. Mol. Biol. Rep. 25:183-188.
STIGMATELLA AND SORANGIUM Coudart-Cavalli, M.-P., 0. Sismeiro, and A. Danchin. 1997. Bifunctional structure of two adenylyl cyclases from the myxobacterium Stigmatella aurantiaca. Biochimie 79:757767. Danchin, A. 1993. Phylogeny of adenylyl cyclases. Adv. Second Messenger Phosphoprotein Res. 27: 109-1 62. Dawid, W. 1979. Vorkornmen und Verbreitung Fruchtkorperbildender Myxobakterien im Siebengebirge. Z . Allg. Mikrobiol. 19:705-719. Dawid, W. 2000. Biology and global distribution of myxobacteria in soil. FEMS Microbiol. Rev. 24:403-427. DCrijard, B., M. Ben A h a , B. Lubochinsky, and Y. Cenatiempo. 1989. Evidence for a membrane-associated GTP-binding protein in Stigmatella aurantiaca, a prokaryotic cell. Biochem. Biophys. Res. Commun. 158562-568. Dhundale, A. R., T. Furuichi, S. Inouye, and M. Inouye. 1985. Distribution of multicopy single-stranded DNA among myxobacteria and related species. J. Bacteriol. 164:914-917. Dickschat, J. S., H. B. Bode, T. Mahmud, R. Miiller, and S. Schulz. 2005a. A novel type of geosmin biosynthesis in myxobacteria. J. Org. Chem. 705174-5182. Dickschat, J. S., H. B. Bode, R. M. Kroppenstedt, R. Miiller, and S. Schulz. 2005b. Biosynthesis of iso-fatty acids in myxobacteria. Org. Biomol. Chem. 3:2824-2831. Dickschat, J. S., H. B. Bode, S. C. Wenzel, R. Miiller, and S. Schulz. 2005c. Biosynthesis and identification of volatiles released by the myxobacterium Stigmatella aurantiaca. ChemBioChem 6:2023-2033. Dworkin, M., and S. M. Gibson. 1964. A system for studying microbial morphogenesis: rapid formation of microcysts in Myxococcus xanthus. Science 146:243-244. Enders, D., and A. Ridder. 2000. First asymmetric synthesis of stigmolone: the fruiting body inducing pheromone of the myxobacterium Stigmatella aurantiaca. Synthesis 2000:1848-1851. Fautz, E., G. Rosenfelder, and L. Grotjahn. 1979. Iso-branched 2- and 3-hydroxy fatty acids as characteristic lipid constituents of some gliding bacteria. J. Bacteriol. 140:852-858. Fisseha, M., M. Gloudemans, R. Gill, and L. Kroos. 1996. Characterization of the regulatory region of a cell interactiondependent gene in Myxococcus xanthus. ]. Bacteriol. 178: 2539-2550. Freese, A., H. Reichenbach, and H. Liinsdorf. 1997. Further characterization and in situ localization of chain-like aggregates of the gliding bacteria Myxococcus fulvus and Myxococcus xanthus. J. Bacteriol. 179:1246-1252. Furuichi, T., A. Dhundale, M. Inouye, and S. Inouye. 1987a. Branched RNA covalently linked to the 5’ end of a singlestranded DNA in Stigmatella aurantiaca: structure of msDNA. Cell 48:47-53. Furuichi, T., S. Inouye, and M. Inouye. 1987b. Biosynthesis and structure of stable branched RNA covalently linked to the 5’ end of rnulticopy single-stranded DNA of Stigmatella aurantiaca. Cell 4855-62. Gerth, K., and H. Reichenbach. 1978. Induction of myxospore formation in Stigmatella aurantiaca (myxobacterales). I. General characterization of the system. Arch. Microbiol. 117~173-182.
18.
s. AURANTIACA AS ALTERNATIVE MYXOBACTERIAL MODEL
Gerth, K., R. Metzger, and H. Reichenbach. 1993. Induction of myxospores in Stigmatella aurantiaca (myxobacteria): inducers and inhibitors of myxospore formation, and mutants with a changed sporulation behaviour. J. Gen. Microbiol. 139:865-871. Gerth, K., and H. Reichenbach. 1994. Induction of myxospores in Stigmatella aurantiaca (myxobacteria):analysis of inducer-inducer and inducer-inhibitor interactions by doseresponse curves. Microbiology 140:3241-3247. Glomp, I., P. Saulnier, J. Guespin-Michel, and H. U. Schairer. 1988. Transfer of IncP plasmids into Stigmatella aurantiaca leading to insertional mutants affected in spore development. Mol. Gen. Genet. 214:213-217. Grilione, P. L., and J. Pangborn. 1975. Scanning electron microscopy of fruiting body formation by myxobacteria. J . Bacteriol. 124:1558-1565. Harris, B. Z., and M. Singer. 1998. Identification and characterization of the Myxococcus xanthus argE gene. J. Bacteriol. 180:64 12-6414. Hauer, B., and H. Eipel. 1997. Flow cytometry: useful tool for analyzing bacterial populations cell by cell, p. 273-291. In J. A. Shapiro and M. Dworkin (ed.), Bacteria as Multicellular Organisms. Oxford University Press, Oxford, United Kingdom. Heidelbach, M., H. Skladny, and H. U. Schairer. 1992. Purification of the DNA-dependent RNA polymerase from the myxobacterium Stigmatella aurantiaca. J. Bacteriol. 174:2733-2735. Heidelbach, M., H. Skladny, and H. U. Schairer. 1993a. Purification and characterization of SP21, a developmentspecific protein of the myxobacterium Stigmatella aurantiaca. J. Bacteriol. 175:905-908. Heidelbach, M., H. Skladny, and H. U. Schairer. 1993b. Heat shock and development induce synthesis of a low-molecularweight stress-responsive protein in the myxobacterium Stigmatella aurantiaca. J. Bacteriol. 175:7479-7482. Hofmann, D. 2004. Proteinanalyse zur Charakterisierung der Lebensformen von Stigmatella aurantiaca. Ph.D. thesis. Ruprecht-ICarls-Universitat, Heidelberg, Germany. Hsu, M.-Y., C. Xu, M. Inouye, and S. Inouye. 1992. Similarity between the Myxococcus xanthus and Stigmatella aurantiaca reverse transcriptase genes associated with multicopy, single-stranded DNA. J. Bacteriol. 174:2384-2387. Hull, W. E., A. Berkessel, and W. Plaga. 1998. Structure elucidation and chemical synthesis of stigmolone, a novel type of prokaryotic pheromone. Proc. Natl. Acad. Sci. U S A 95: 11268-1 1273. Inouye, S., D. White, and M. Inouye. 1989. Development of Stigmatella aurantiaca: effects of light and gene expression. J. Bacteriol. 141:1360-1365. Inouye, S., P. J. Herzer, and M. Inouye. 1990. Two independent retrons with highly diverse transcriptases in Myxococcus xanthus. Proc. Natl. Acad. Sci. U S A 87:942-945. Jeong, M.-A., and D. Lim. 2004. A proteomic approach to study msDNA function in Escherichia coli. J. Microbiol. 42:200-204. Kaiser, D. 2003. Coupling cell movement to multicellular development in myxobacteria. Nut. Rev. Microbiol. 1:45-54.
325
Kleinig, H., and H. Reichenbach. 1969. Carotenoid pigments of Stigmatella aurantiaca (myxobacterales). I. The minor carotenoids. Arch. Mikrobiol. 68:210-217. Kleinig, H., and H. Reichenbach. 1970. Carotenoid pigments of Stigmatella aurantiaca (myxobacterales). 11. Acylated carotenoid glucosides. Arch. Mikrobiol. 74:223-234. Kunze, B., T. Kemmer, G. Hofle, and H. Reichenbach. 1984. Stigmatellin, a new antibiotic from Stigmatella aurantiaca (myxobacterales). I. Production, physico-chemical and biological properties. J. Antibiot. 37:454-461. Kunze, B., G. Hofle and H. Reichenbach. 1987. The aurachins, new quinoline antibiotics from myxobacteria: production, physico-chemical and biological properties. J. Antibiot. 40: 258-265. Kunze, B., H. Reichenbach, R. Miiller, and G . Hofle. 2005. Aurafuron A and B, new bioactive polyketides from Stigmatella aurantiaca and Archangium gephyra (myxobacteria): fermentation, isolation, physico-chemical properties, structure and biological activity. J. Antibiot. 5 8~244-251. Lee, B. U., K. Lee, J. Mendez, and L. J. Shimkets. 1995. A tactile sensory system of Myxococcus xanthus involves an extracellular NAD(P)+-containingprotein. Genes Dev. 9: 2964-2973. Li, S., and L. Shimkets. 1988. Site-specific integration and expression of a developmental promoter in Myxococcus xanthus. J. Bacteriol. 1705552-5556. Lunsdorf, H., H. U. Schairer, and M. Heidelbach. 1995. Localization of the stress protein SP21 in indole-induced spores, fruiting bodies, and heat-shocked cells of Stigmatella aurantiaca. J. Bacteriol. 1777092-7099. Liinsdorf, H., and H. U. Schairer. 2001. Frozen motion of gliding bacteria outlines inherent features of the motility apparatus. Microbiology 147:939-947. Mignot, T., J. W. Shaevitz, P. L. Hartzell, and D. R. Zusman. 2007. Evidence that focal adhesion complexes power bacterial gliding motility. Science 315:853-856. Milosevic, A. 2003. CsgA, a putative signal molecule of the myxobacterium Stigmatella aurantiaca involved in fruiting: characterization of the csgA gene and influence of csgA inactivation on development. Ph.D. thesis. Ruprecht-KarlsUniversitat, Heidelberg, Germany. Morikawa, Y., S. Takayama, R. Fudo, S. Yamanaka, K. Mori, and A. Isogai. 1998. Absolute chemical structure of the myxobacterial pheromone of Stigmatella aurantiaca that induces the formation of its fruiting body. FEMS Microbiol. Lett. 165:29-34. Muller, S. 2002. Morphogenese bei Stigmatella aurantiaca: Studien zur Fruchtkorperbildung. Ph.D. thesis. RuprechtKarls-Universitat, Heidelberg, Germany. Miiller, S., H. Shen, D. Hofmann, H. U. Schairer, and J. R. Kirby. 2006. Integration into the phage attachment site, attB, impairs multicellular differentiation in Stigmatella aurantiaca. J. Bacteriol. 188:1701-1709. Neumann, B., A. Pospiech, and H. U. Schairer. 1992. Size and stability of the genomes of the myxobacteria Stigmatella aurantiaca and Stigmatella erecta. J. Bacteriol. 174:63076310.
326 Neumann, B., A. Pospiech, and H. U. Schairer. 1993. A physical and genetic map of Stigmatella aurantiaca DW4/3.1 chromosome. Mol. Microbiol. 10:1087-1099. Plaga, W., I. Stamm, and H. U. Schairer. 1998. Intercellular signaling in Stigmatella aurantiaca: purification and characterization of stigmolone, a myxobacterial pheromone. Proc. Natl. Acad. Sci. USA 95:11263-11267. Pospiech, A., B. Neumann, B. Silakowski, and H. U. Schairer. 1993. Detection of developmentally regulated genes of the myxobacterium Stigmatella aurantiaca with the transposon TnSlacZ. Arch. Microbiol. 159:201-206. Pradella, S., A. Hans, C. Sproer, H. Reichenbach, K. Gerth, and S. Beyer. 2002. Characterisation, genome size and genetic manipulation of the myxobacterium Sorangium cellulosum So ce56. Arch. Microbiol. 178:484-492. Qualls, G. T., K. Stephens, and D. White. 1978a. Light-stimulated morphogenesis in the fruiting myxobacterium Stigmatella aurantiaca. Science 201:444-445. Qualls, G. T., K. Stephens, and D. White. 1978b. Morphogenetic movements and multicellular development in the fruiting myxobacterium, Stigmatella aurantiaca. Dev. Biol. 66~270-274. Reichenbach, H., and M. Dworkin. 1969. Studies on Stigmatella aurantiaca (myxobacterales).J. Gen. Microbiol. 58:3-14. Reichenbach, H., H. Voelz, and M. Dworkin. 1969. Structural changes in Stigmatella aurantiaca during myxospore induction.]. Bacteriol. 97:905-911. Reichenbach, H., and M. Dworkin. 1970. Induction of myxospore formation in Stigmatella aurantiaca (Myxobacterales) by monovalent cations. J. Bateriol. 101:325-326. Reichenbach, H., and M. Dworkin. 1992. The Myxobacteria, p. 3416-3487. In A. Balows, H. G. Triiper, M. Dworkin, W. Harder, and K.-H. Schleifer (ed.), The Prokaryotes, 2nd ed. Springer-Verlag, Heidelberg, Germany. 3rd ed., http://link. springer-ny.com/link/service/books/lO 1251. Reichenbach, H. 2001. Myxobacteria, producers of novel bioactive substances.]. Ind. Microbiol. Biotechnol. 27:149156. Rice, S. A., and B. C. Lampson. 1996. Bacterial reverse transcriptase and msDNA. Virus Genes 11:95-104. Schroder, J., and H. Reichenbach. 1970. The fatty acid composition of vegetative cells and myxospores of Stigmatella aurantiaca (myxobacterales). Arch. Mikrobiol. 71:384390. Shen, H., and H. U. Schairer. 1999. Transcriptional regulation of hspA of Stigmatella aurantiaca and function investigation of the genve product. Chin. Med. J. 11253.57. Shen, H. 1999. Transcriptional regulation of hspA gene in Stigmatella aurantiaca and function analysis of HspA protein. Ph.D. thesis. Ruprecht-Karls-Universitat, Heidelberg, Germany. Shimkets, L. J., and M. Dworkin. 1981. Excreted adenosine is a cell density signal for the initiation of fruiting body formation in Myxococcus xanthus. Dev. Biol. 8451-60. Silakowski, B., A. Pospiech, B. Neumann, and H. U. Schairer. 1996. Stigmatella aurantiaca fruiting body formation is dependent on the f 6 f A gene encoding a polypeptide homologous to chitin synthases. J. Bacteriol. 178:6706-6713.
STIGMATELLAAND SORANGIUM Silakowski, B., H. Ehret, and H. U. Schairer. 1998. FbfB, a gene encoding a putative galactose oxidase, is involved in Stigmatella aurantiaca fruiting body formation. J. Bacteriol. 180:1241-1247. Silakowski, B., H. U. Schairer, H. Ehret, B. Kunze, S. Weinig, G. Nordsiek, P. Brandt, H. Blocker, G. Hofle, S. Beyer, and R. Muller. 1999. New lessons for combinatorial biosynthesis from myxobacteria. J. Biol. Chem. 274:37391-37399. Silakowski, B., S. Muller, H. Skladny, H. Ehret, and H. U. Schairer. 2001. SigB, an alternative sigma factor of the myxobacterium Stigmatella aurantiaca, is synthesized during development and heat shock. Microbiology 1422265-2273. Skladny, H., M. Heidelbach, and H. U. Schairer. 1992. Cloning and DNA sequence of sigB gene of Stigmatella aurantiaca. Nucleic Acids Res. 20:6416. Skladny, H., M. Heidelbach, and H. U. Schairer. 1994. Cloning and characterization of the gene encoding the major sigma factor of Stigmatella aurantiaca. Gene 143:123127. Stamm, I., A. Leclerque, and W. Plaga. 1999. Purification of cold-shock-like proteins from Stigmatella aurantiacamolecular cloning and characterization of the cspA gene. Arch. Microbiol. 172:175-181. Stamm, I., and W. Plaga. 2000. Fluorescence-based analysis of gene expression during the developmental cycle of the myxobacterium Stigmatella aurantiaca (abstract). Biospektrum (Sonderheft 2000) 6:78. Stamm, I., F. Lottspeich, and W. Plaga. 2005. The pyruvate kinase of Stigmatella aurantiaca is an indole binding protein and essential for development. Mol. Microbiol. 56:13861395. Stephens, K., and D. White. 1980a. Morphogenetic effects of light and guanine derivatives on the fruiting myxobacterium Stigmatella aurantiaca. J. Bacteriol. 144:322326. Stephens, K., and D. White. 1980b. Scanning electron micrographs of fruiting bodies of the myxobacterium Stigmatella aurantiaca lacking a coat and revealing a cellular stalk. FEMS Microbiol. Lett. 9:189-192. Stephens, K., G. D. Hegeman, and D. White. 1982. Pheromone produced by the myxobacterium Stigmatella aurantiaca. J. Bacteriol. 149:739-747. Thaxter, R. 1892. Contributions from the Cryptogamic Laboratory of Harvard University. XVIII. On the Myxobacteriaceae, a new order of Schizomycetes. Bot. Gaz. 23:395411. Tiennault-Desbordes, E., Y. Cenatiempo, and S. Laalami. 2001. Initiation factor 2 of Myxococcus xanthus, a large version of prokaryotic translation initiation factor 2. J. Bacteriol. 183:207-2 13. Trowitzsch, W., L. Witte, and H. Reichenbach. 1981. Geosmin from earthy smelling cultures of Nannocystis exedens (myxobacterales). FEMS Microbiol. Lett. 12:257-260. Vasquez, G. M., F. Qualls, and D. White. 1985. Morphogenesis of Stigmatella aurantiaca fruiting bodies. J. Bacteriol. 163515-521. Voelz, H., and H. Reichenbach. 1969. Fine structure of fruiting bodies of Stigmatella aurantiaca (Myxobacterales).J. Bacterial. 99:856-866.
18. S. AURANTIACA AS ALTERNATIVE MYXOBACTERIAL MODEL Weinig, S., T. Mahmud, and R. Miiller. 2003. Markerless mutations in the myxothiazol biosynthetic gene cluster: a delicate megasynthase with a superfluous nonribosomal peptide synthetase domain. Chem. Biol. 10:953-960. Wenzel, S. C., B. Kunze, G . Hofle, B. Silakowski, M. Scharfe, H. Blocker, and R. Miiller. 2005. Structure and biosynthesis of myxochromides S1-3 in Stigmatella aurantiaca: evidence for an iterative bacterial type I polyketide synthase and for module skipping in nonribosomal peptide biosynthesis. ChemBioChem 6:375-385. White, D., W. Shropshire, and K. Stephens. 1980. Photocontrol of development by Stigmatella aurantiaca. 1. Bacteriol. 142:1023-1 024.
32 7
Wu, S. S., and D. Kaiser. 1996. Markerless deletions of pi1 genes in Myxococcus xanthus generated by counterselection with the Bacillus subtilis sacB gene. J. Bacteriol. 17858175821. Yamauchi, K., J.-I. Nakajima, H. Hayashi, R. Horiuchi, and J. R. Tata. 1999. Xenopus cytosolic thyroid hormone-binding protein (xCTBP) is aldehyde dehydrogenase catalyzing the formation of retinoic acid. J. Biol. Chem. 274:84608469. Yamauchi, K., and J. R. Tata. 2001. Characterization of Xenopus cytosolic thyroid-hormone-binding protein (xCTBP) with aldehyde dehydrogenase activity. Chem. Biol. Interact. 130-1 3 2 ~09-32 3 1.
Myxobacteria: Multicellularity and Differentiation Edited by David E. Whitworth 0 2008 ASM Press, Washington, D.C.
Klaus Gerth Olena Perlova Rolf Muller
Sorangium cellulosurn
HISTORY The story of cellulose-degrading myxobacteria and their systematics and morphology was first reviewed by Imshenetski in 1959 (Imshenetski, 1959). On enrichment plates for cellulose-degrading bacteria from soil samples taken at the Volga riverbanks, he observed orangecolored regions on the wetted filter paper. Later, brownish elevations appeared in the center of the colored zone, which were identified as microcysts from myxobacteria by microscopic observation. The bacterium was identified as Polyangium cellulosum by the use of the 3rd edition of Bergey’s Manual of Determinative Bacteriology, and the results were published in 1936 (Imshenetski and Solntseva, 1936). Later this isolate was renamed according to the new Bergey’s nomenclature by Jahn into Sorangium compositum, a myxobacterium which was first described by Thaxter in 1904, who, however, did not recognize its capability to degrade cellulose (Krzemieniewska and Krzemieniewski, 1937a). In 1937 Imshenetski and Solntseva isolated a new species of cellulose-degrading myxobacteria, which they called Sorangium cellulosum. All further isolates were grouped into a Polyangium cellulosum group, which was characterized by large dark-brown fruiting bodies with
19
polygonal cysts and large vegetative cells (3.5 to 8.5 pm long with a diameter of 0.8 to 1.2 pm), and a Sorangium cellulosum group with reddish-brown but not very characteristic fruiting bodies and smaller cells (2.2 to 4.5 pm long with a diameter of 0.4 pm). In the following years several new cellulolytic species of Sorangium like S. nigrescens, S. niger, and S. spumosum were described (Krzemieniewska and Krzemieniewski, 1937b; Mishustin, 1938; Imshenetski, 1959), but with the exception of P. cellulosum no culturable reference material is available today for any of them (Sproer et al., 1999).
TAXONOMY AND SYSTEMATICS A classification based almost exclusively on fruiting body characteristics like size or color does not take into account the tremendous variations in fruiting body morphology and the effect of culture conditions like media. McCurdy discussed a “Polyangium-Sorangiumcellulolytic complex” because he was not able to unambiguously assign any of his own isolates of cellulolytic myxobacteria to either of these two groups (McCurdy, 1970).J. E. Peterson described S. cellulosum and P. cellulosum and perhaps most other cellulolytic
Klaus Gerth, Helmholtz-Zentrum fur Infektionsforschung GmbH, Inhoffenstrage 7 , 3 8 124 Braunschweig, Germany. Olena Perlova and Rolf Miiller, Institut fur Pharmazeutische Biotechnologie, Universitat des Saarlandes, Postfach 15 11 50,66041 Saarbriicken, Germany.
329
330 myxobacteria as identical or, at best, variants (Peterson, 1969b). 16s rRNA gene sequencing of nine isolates of Sorangium and their comparison with P. cellulosum as the reference strain proved a close phylogenetic relationship (evolutionary distance, less than 3 % on the nucleotide level). According to these results they all belong to a single genus (Yan et al., 2003). Analysis of two more cellulose-degrading isolates of “S. cellulosum,” i.e., So ce90 (producer of epothilone) and So ce56 (genome sequenced), confirm this classification of Sorangium. Therefore, the proposed use of the genus name Sorangium for cellulose degraders of the Polyangium cellulosum complex (Sproer et al., 1999) seems to be reasonable, but the assignment has to be further validated. Future sequencing of representative isolates from our collection of more than 2,000 Sorangium strains at the Gesellschaft fur Biotechnologische Forschung (GBF) in Braunschweig may perhaps reveal new species of cellulose-degrading myxobacteria. From the phylogenetic point of view, based on 16s rRNA analysis, myxobacteria belong to the Proteobacteria (Sproer et al., 1999). They represent a deep trifurcated but phylogenetically coherent group, the order Myxococcales. One lineage is defined by the genera Cystobacter, Angiococcus, Archangium, Melittgangium, M ~ X O C O C Cand U S ,Stigmatella. The second lineage contains the genus Chondromyces and the species PolyangiumlSorangium cellulosum, while the third lineage is comprised of Nannocystis and a strain identified as Polyangium vitellinum (Sproer et al., 1999). The new taxonomy of myxobacteria (Brenner et al., 2005) takes these results into account: the Myxococcales are now divided into three suborders, the “ Cystobacterineae,” the “Sorangiineae,” and the “Nannocystineae.” In the literature there is some confusion with respect to the classification of certain gliding bacteria. Erroneously one Lysobacter isolate was classified as Sorangium by E. A. Peterson (Peterson et al., 1966). By light microscopic observation the cells of lysobacteria can resemble the vegetative cells of Sorangium isolates, and also the GC content of 65 to 70% comes close to that of Sorangium (70 to 72%) (Reichenbach, 1992). Nevertheless, they can be distinguished by their growth velocity and the color of the colonies (Sorangium and Lysobacter colonies are orange to red and pale white to yellow, respectively). Growth of Sorangium is slow with generation times usually between 8 and 16 h, while Lysobacter grows very fast (2- to 3-h generation time). Some Lysobacter isolates were called Myxobacter in the past, an obsolete myxobacterial genus, which can be found in literature even today.
STIGMATELLA AND SORANGIUM
ISOLATION AND MORPHOLOGY Distribution Like other myxobacteria the genus Sorangium is ubiquitous and a prominent component of the soil microflora. It was isolated from the Kola Peninsula in northern Russia, where it is fairly common in virgin and cultivated podzol and peat marsh soils (Peterson, 1969b),in all Scandinavian countries (Peterson and NorCn, 1967), in tremendous profusion in the Sonoran desert soils of Mexico (90 to 95% of all soil piles were positive E. Peterson, personal communication]), in soils of the central United States (Lampky, 1971), in India (Singh and Singh, 1971), in China (Yan et al., 2003), and in Israel (Gaspari et al., 2004). In studies on the myxobacterial biodiversity in an established oak-hickory forest, S. cellulosum was found to be a predominant bacterial species (Neil et al., 2005). At the GBF Sorangium was isolated from soil samples taken at many places from four continents. Interestingly, populations of this organism seem to be very much associated with human cultivation. Extensive studies in Missouri soils and patterns of occurrence of Sorangium in Sweden revealed a strong correlation between the profusion of Sorangium and the degree of manuring and ploughing under of plant debris (Peterson, 1965). Statistically Sorangium can be isolated from about 15 to 30% (Peterson, personal communication) or 5% (Dawid, 2000) of all soil samples. The sample origin, i.e., from cultivated areas or undisturbed soils, can explain the observed differences.
u.
Enrichment and Isolation One outstanding characteristic of Sorangium is its ability to decompose cellulose. In order to take advantage of this potential for the isolation of cellulolytic myxobacteria, Solntseva has recommended the use of a complete mineral salt agar with filter paper as a substrate (Solntseva, 1939). The filter paper method is best for the isolation of S. cellulosum strains even today. According to the classical methods the enrichment plates are incubated at room temperature or at 30°C. Within 2 to 3 weeks yellow-orange spots can be detected adjacent to the soil, a first indication of growth of Sorangium. Later the degradation of cellulose and first sporangioles or fruiting bodies can be monitored under the dissection microscope (Color Plate 7a). Lysed areas are contaminated with a complex mixture of other cellulolytic or mostly noncellulolytic bacteria and fungi. If fresh soil samples are used, the plate is often crowded with amoebae and nematodes. Slimes which cover the fruiting bodies harbor contaminants and shelter them. Addition of 100 Fg/ml of cycloheximide (Actidion) (Peterson, 1969b) is useful to reduce
19. SORANGIUMCELLULOSUM
33 1
growth of fungal contaminants, while the propagation of nematodes and amoebae is almost completely inhibited by the addition of 100 p,g/ml of levamisole (Gerth and Miiller, 2005). Both compounds can be added to the agar from the very beginning without any effect on the growth of myxobacteria. The main problems during the isolation procedure are therefore bacterial contaminants. The use of single antibiotics or better combinations of several antibiotics that act on different targets may be helpful, because Sorangium species usually turn out to be multiresistant (Gerth et al., 2003). Useful combinations are cephalosporin, which acts on cell wall synthesis, and/or kanamycin, an inhibitor of protein synthesis, or polymyxine (active on membranes) and/or trimethoprim as an inhibitor of folic acid synthesis. Often, however, a reduced number of different resistant bacteria will remain. According to our experience Sorangium is usually sensitive against rifampicin, chloramphenicol, streptomycin, and fusidic acid. If the strains begin to swarm outside the filter paper (Color Plate 7b), uncontaminated cells can sometimes be scraped from the swarm edges and transferred onto fresh filter paper. The tendency of Sorangium to penetrate the agar can also be used for purification. Inverting the agar medium makes possible to pick material from the deepest portion of the penetrating swarm, which often is no longer contaminated (McCurdy, 1969). If these procedures are repeated several times, pure cultures can be obtained and tested by transfer onto complex medium like medium M (Muller and Gerth, 2006).
Cell Morphology Cells of the suborder “Cystobacterineae” on the one hand and “Sorangiineae” and “Nannocystineae” on the other hand can easily be distinguished because they differ in cell morphology, as can be detected using phase-contrast microscopy. While members of the former family are small and slender with tapered ends, Sorangium cells are robust rigid cylindrical rods with blunt rounded ends, 2 to 10 km long and 0.4 to 1.2 Frn in diameter. In older cultures, refractive inclusion bodies redolent of endospores from bacilli can often be seen. These may be accumulations from poly-P-hydroxybutyric acid, which was detected in cell extracts (R. Jansen, unpublished results). The conversion of vegetative Sorangium cells to resting cells is not accompanied by a typical cellular morphogenesis as we know it from M~xOcOccus species. Myxos??ores are only slightly shorter ( 2to 4 Fm by 0.8 to 1.2 pm) than the vegetative cells and are not phase dense (Fig. 1).
Colonies and Swarms On poor media like vy/2 agar plates (Reichenbach and Dworkin, 1981), the vegetative cells move by gliding
Figure 1 Raster electron microscopic pictures of fruiting sorangium. (a) Vegetative Swarm colonY. (b) SPorangioles on the agar surface, some of which are broken. (c) Myxospores of Sorangium. The surface Structure is the result of drying. Pictures by K. Gerth and H. Lfinsdorf.
and typical swarm colonies appear; in contrast, compact nonswarming colonies arise on rich peptone maltose media.
332 The consistency of the colonies varies: sometimes the isolates form soft-slimy colonies and one can easily scrape off cells with an inoculation loop. The distribution on fresh agar plates or inoculation of liquid cultures is difficult with rigid gristly colonies because the cell masses stick together. Swarms of Sorangium can be very different in shape, but as a rule, regularly arranged rippling patterns as we know them from other myxobacteria like Myxococcus or Corallococcus can never be seen. On cellulose filter paper one can often recognize different zones (Color Plate 7a), i.e., a yellow-orange outer zone where Sorangium is actively growing, a brownish region where the filter paper becomes increasingly degraded, and a center with dark red-brown masses of fruiting bodies. Fruiting Bodies Sporangioles which lie directly on the substrate are packed together in small or large, loose or dense parcels, “primary cysts” (Icrzemieniewska and Krzemieniewski, 1937b), or “sporangia” (Peterson, 1969a) with up to 100 sporangioles. They are usually covered by a slime layer (Color Plate 7d). Because of the pressure inside these parcels, the spherical sporangioles are often of a polyhedral shape. The sum of parcels, the fruiting bodies, can be produced in enormous quantities on digested filter paper (Color Plate 7a). Sometimes sporangioles can be detected not in the form of parcels but grouped in long rows which correlate in their direction with the orientation of previous cellulose fibers (K. Gerth, unpublished data). The size of sporangioles varies from 10 to 60 pm in diameter (Color Plate 7c). A subdivision into two species of Sorangium, those with small spherical sporangioles (10 to 30 pm) and those with large, polyhedrical sporangioles (30 to 60 Fm), was not supported by 16s rRNA gene sequence analysis (Yan et al., 2003). The color of fruiting bodies varies from yellow, orange, red-brown, and brown to black. When the cultures are propagated over a longer period, the capability of fruiting body formation is usually lost. We believe that a selection of mutants which can grow in homogenous cell suspension is often accompanied by a loss in slime secretion, one prerequisite of fruiting body formation.
PHYSIOLOGY Aerobic cellulose degradation is not very common in bacteria. Strains of Byssophaga cruenta, characterized by their intense blood-red color, and the very common S. cellulosum strains are the only myxobacteria which degrade crystalline cellulose and can use it as the sole
STIGMATELLA AND SORANGIUM carbon source (Brenner et al., 2005). Hence, Sorangium is much more versatile with respect to carbon source than, for example, Myxococcus; besides cellulose, xylans, starch, and chitin are also degraded. Amazingly, this very common soil bacterium is almost overlooked in the literature related to cellulose degradation and soil ecology. The first physiological investigations with cellulosedegrading myxobacteria were performed by H. and S. Krzemieniewski. These authors proved the aerobic degradation of cellulose which is used as the sole and best carbon source in the presence of nitrate or ammonium salts and investigated the pH and temperature dependence of the process (Icrzemieniewska and Krzemieniewski, 1937a). A cellulolytic activity was detected in culture filtrates, which hydrolyzed insoluble cellulose to cellobiose and glucose (Coucke and Voets, 1968). In addition to these soluble cellulases, cell-bound cellulases can be produced (Sarao et al., 1985). First annotation results of the S. celZulosum So ce56 genome clearly prove the existence of an exoglucanase as well as numerous endoglucanases. Xylan-degrading enzymes, which are responsible for the lysis of hemicelluloses, the main components of plant fibers, are also present. In addition to cellulose and starch, their degradation products (the disaccharides cellobiose and maltose) are excellent carbon sources used in synthetic media (Miiller and Gerth, 2006). In contrast to glucose and fructose, which are alternative inexpensive substrates that allow growth, mannose is a unique substrate which enables fast growth up to a high cell density. Sucrose and ribose cannot be metabolized by Sorangium. In our experience, xylose can be used only by some strains, e.g., So ce12 (Hoischen, 1986), but does not support growth of So ce56 (Miiller and Gerth, 2006). The same is true for organic acids like acetate or lactate as the sole carbon source (Miiller and Gerth, 2006). Key enzymes for carbohydrate metabolism, glycolysis, pentose shunt pathway, tricarboxylic acid cycle, and glyxoylate shunt have been described (Sarao et al., 1985).Ammonium sulfate and asparagine are favored nitrogen sources for the growth of most Sorangium isolates; others seem to prefer nitrate. Glutamate and aspartate cannot replace one of the nitrogen sources mentioned above. Magnesium sulfate, calcium chloride, potassium phosphate, and ferrous sulfate are essential minerals which are required for growth (Coucke and Voets, 1967). Favored substrates for growth of Sorangium strains cultivated so far in our laboratories are complex media based on soy peptones and maltose as in medium M (Miiller and Gerth, 2006). Growth on peptone as the sole carbon source as described by Sarao could not be confirmed by us (Sarao et al., 1985).
19. SORANGIUMCELLULOSUM
333
Myxobacteriologists have always considered their bacteria to be mesophiles with an optimum growth temperature between 30 and 34°C (Dawid, 2000). This is generally also true for Sorangium, but some strains were described to tolerate even 38 to 40”C, accompanied, however, with a declined growth velocity. It came as a surprise when we first isolated moderately thermophilic Sorangium strains with a temperature optimum increased by approximately 10°C (Fig. 2). The origin of the soil samples for isolation of such myxobacteria is a key factor for the detection of these strains; semiarid warm climates, as found in Mediterranean countries, seem to favor moderately thermophilic myxobacteria in their natural environment. The growth of these Sorangium strains at 30°C is slow, which is why these strains have been overlooked in the past (Gerth and Miiller, 2005). To date, true thermophilic Sorangium strains, however, have not been detected. Mesothermophilic as well as moderately thermophilic Sorangium isolates share a natural multiple resistance against antibiotics. Most isolates are inherently resistant to numerous amino glycosides like kanamycin, neomycin, ribostamycin, or tobramycin and against beta-lactams, i.e., ampicillin or cephalosporin. Resistance against the membrane active polymyxin or trimethoprim-an inhibitor of folic acid synthesis-is typical for many strains. Fusidic acid, an antibiotic
r
16
So ce26
16 Y
.= E
14
C
.-0 4-l
2
12
a,
c
8
lo
8
6 28
30
32
34
36
38
40
42
4-1
46
Incubationtemperature [“C] Figure 2 Dependence of the generation time of Sorangium strains on incubation temperature. So ce26 is a mesothermophilic isolate. An increase of the temperature from 30 to 40°C results in an increase of the generation time from 11 to 19 h. GT-46 and GT-41 are moderately thermophilic Sorangium strains. The generation time decreases with an increase in temperature. At 42°C the temperature optimum is reached with a generation time of 6.5 h.
which acts preferentially against gram-positive bacteria, inhibits growth of most Sorangium isolates (Gerth and Miiller, 2005). Nevertheless, there are always exceptions to this rule. Recently we isolated a moderately thermophilic s. cellulosum strain, which is apparently sensitive to most of the antibiotics mentioned above, kanamycin included (Gerth, unpublished).
SECONDARY METABOLISM Antibiotic activities from myxobacteria were described as early as in 1950 (Finck, 1950). In 1966 the first detailed description of “a wide-spectrum activity produced by a species of Sorangium” was published (Peterson et al., 1966). The structure of the corresponding metabolite, myxin, was elucidated a year later (Weigele and Leimgruber, 1967), but the producer strain obviously was a Lysobacter and not a myxobacterium (Reichenbach, 1992).The producer of the first real myxobacterial secondary metabolite, the antifungal ambruticin (Ringel et al., 1977),was an S. ceElulosum isolated by J. E. Peterson. His pioneering work on Sorangium opened the door to others.
The Potential of the Genus Sorangium At the GBF an isolation and screening program was initiated in the late 1970s to evaluate the potential of the different genera of myxobacteria as producers of secondary metabolites. A close cooperation between microbiologists, analytical chemists, and fermentation engineers enabled this work. Today we know that strains of S. cellulosum are by far the most potent myxobacteria with respect to secondary metabolite production (Gerth et al., 2003). Almost 50% of the more than 100 novel metabolites and some of the most interesting ones-sorangicins (Irschik et al., 1987), soraphens (Gerth et al., 1994), and epothilones (Gerth et al., 1996a)-are produced by members of this group (Fig. 3). Surprisingly, only one Sorangium metabolite, the antibiotic pyrrolnitrin, had been isolated before from a member of another suborder of myxobacteria as well as from a nonmyxobacterial isolate. In general, a family of closely related compounds is produced rather than a single secondary metabolite. From the culture broth of strain So ce26,28 structurally related soraphens were isolated, and strain So ce90 was shown to excrete more than 30 different epothilones (Gerth et al., 2003). Almost 90% of the more than 2,000 isolated Sorangium strains of the GBF collection are actually producers of some natural products. There are strains which synthesize only one or few compounds. But there are also strains that are multiproducers of many unrelated
STIGMATEL LA AND SORANGI UM
334 Sorangium 47,6%
-
‘xococcus 12,60/
Hyalocystis 1,0% Angiococcus 1,9% Corallococcus 1,9940 Byssophaga 2,9% Cystobacter 2,9%
Polyangium 3,9%
Stigmtatella
I
Chondromyces 6,8%
I
Nannocystis 4,9% Archangium 5,8%
Figure 3 Myxobacterial producers of novel secondary metabolites. With 47% of total production, Sorangium strains are the most outstanding producers of novel metabolites.
compounds like So cel525, which excretes chivosazols, disorazols, sorangicins, soraphens, sulfangolide, sorangiolide, chlorotonils, and some up-to-now-unknown compounds simultaneously (Gerth et al., 2003). All these compounds are polyketides, nonribosomally made peptides, or hybrids thereof, and they are different with respect to their chemical structure, their biosynthesis, and their targets. Sometimes a consistent combination of metabolites is produced (50% of the epothilone producers also make spirangiens), while epothilone is never found in combination with sorangicin (which itself is usually accompanied by disorazols [according to Irschik et al., 1995b, 50% of the sorangicin producers]) or chivosazols ([27% of the sorangicin producers]). The frequency with which a compound is detected varies (Fig. 4). Some compounds like icumazol or spirangien (Niggemann et al., 2005) are widespread, and others like epothilone or chivosazol (Irschik et al., 1995a) are frequently found in screening, while some compounds are actually rare, like etnangien or jerangolid (Gerth et al., 199610). The distribution of producers of a certain compound seems to be independent from the origin of the soil sample they were isolated from. During our screening program, many soraphen- and epothiloneproducing strains of S. cellulosum were isolated from soil samples collected worldwide. Between 1.1and 3.6% of the isolates from Europe, Asia, Africa, and the United States produced soraphen, whereas 1 to 2.5% of these were shown to produce epothilone (Gerth et al., 2003). There is no preferred occurrence in one continent. The
production rate and the nutritional requirements are, however, strain specific and vary over a broad range, as is exemplified for different producer strains of soraphen and epothilone. While soraphen production is almost completely inhibited by the addition of 0.2% of peptone in strain So ce26, these conditions are almost optimal for strain So ce539 (Gerth et al., 1994). Similar differences in physiology were detected with the epothilone producer strains So ce90 and So ce1198. The production of So ce90 is stimulated by increasing concentrations of glucose; the production of strain So cell98 is, however, inhibited by the same compound (Gerth et al., 2003). The initial concentrations produced by the different producer strains vary between 0.3 to 25 mg/liter for epothilones and 0.1 to 70 mg/liter for soraphens. The compounds so far elucidated from Sorangium strains are mostly macrocyclic lactone rings, linear polyketides, and cyclic peptides. Hybrid compounds synthesized by nonribosomal peptide synthetases and polyketide synthases (e.g., disorazol or epothilone) are very common and characteristic. Although the metabolism of polysaccharides is unique for this genus of myxobacteria, compounds with attached sugar moieties are rare, e.g., sorangicin, chivosazol, or icumazol. The production of secondary metabolites from myxobacteria and from Sorangium has been reviewed recently (Hofle and Reichenbach, 1995; Reichenbach and Hofle, 1999), which is why only some novel^' compounds are presented here (Fig. 5), which until now were published only in the “Annual Scientific Reports of the GBF”
19. SORANGIUM CELLULOSUM
335 Spirangien
9% lcumazole
Diisorazol A-
Sorapkien
I
Am bruticin
431
Chivosazol
\
Disorazol427 Epothilone
Figure 4 Frequency of some selected metabolites derived from S. cellulosum strains. The data are given as numbers of producer strains from 1,700 screened isolates. From Gerth et al. (2003)with the permission of Elsevier, B.V.
(Hofle, 1996-2002). Some of these metabolites are inactive in our test systems and were detected by a highperformance liquid chromatography (HPLC) screening because of their unknown UV spectra. These drug-like compounds are interesting candidates for future screenings on novel targets. The majority of secondary metabolites from Sorangium act against eukaryotes, i.e., fungi, which might reflect the pressure of competition of these cellulose degraders with wood-destroying fungi in their natural biotope.
Improvement of Yields Under nonoptimized conditions the natural products are produced in a concentration range between 0.2 to 20 mg/ liter, very rarely up to 100 mg/liter. The production of industrially relevant secondary metabolites thus requires a simultaneous improvement of the producer strains, the culture media, and the fermentation processes. This is possible, as was convincingly demonstrated with the industrially important antifungal metabolite soraphen for the first time. The productivity was increased from 3 mg/liter in 1986 to 1.5 g/liter in 1990 without the help of molecular biology, which was not yet established for Sorangium at that time. Fermentation processes run with partners from the industry have been scaled up to 60m3 scales. Another example is epothilone, which is being produced by fermentation for the pharmaceutical market as an anticancer compound. Both examples contradict
statements that the biotechnological production of natural products using S. cellufosum is “economically impractical” (Tang et al., 2000).
MOLECULAR BIOLOGY OF THE GENUS SORANGIUM
S. cellulosum and the Largest Known Prokaryotic Genome To increase our understanding of the physiology of myxobacteria and to take advantage of the biosynthetic potential of the genus Sorangium, several molecular biological and genetic approaches were developed and are being further improved in addition to the classical methods of strain isolation and production yield enhancement. Understanding the molecular basics of the processes involved in secondary metabolism increases the chances to isolate new natural products and to use the metabolic capacity of these fascinating microorganisms more efficiently. At present, it is possible to use molecular similarities to deduce relationships of genes and organisms. Sequencing and analysis of 16s rRNA allows the phylogenetic classification of microorganisms. Hybridization approaches lead to the identification of the regions of interest on the DNA molecule of the target organisms or in genomic or metagenomic libraries, which are used for the identification of biosynthetic genes based on their similarity in different organisms. By now,
STIGMATELLA AND SORANG I U M
336
0
Kulkenon 0
Eliamid
0
OH
Carolacton
OH
OHH
OH
~
l
t
e
p
o
l
i
d
A
Soracumen 0
Socein
Tuscolid
HO
HO
Pellasoren
Thuggacin Maracen A
OH
Etnangien OH
OH
Figure 5 A survey of “novel” metabolites from Sorungium. Typical linear and macrocyclic polyketides are presented. Some of them are likely to be biosynthesized by combinations of peptide synthetases and polyketide synthetases, e.g., eliamid. Socein is one of the rare polypeptides active against fungi and yeasts.
the sequencing of whole genomes has become a routine procedure. Genomics, transcriptomics, and proteomics represent large-scale analyses of the organism, providing a plethora of information which allows insights into the physiology as well as explanations of various regulatory processes of an organism. To date, hundreds of microbial genomes have been sequenced, and even more prokaryotic genome sequencing projects (914!)are being pursued
at the moment (http://www.genomesonline.org/). Many of the target organisms are interesting for industrial applications (e.g., as biocatalysts or producers of antibiotics). Only a few of all sequenced bacterial species represent the delta group of proteobacteria; among them are the myxobacteria Myxococcus xanthus DK1622, S. cellulosum So ce56, and Anaeromyxobacter dehalogenans 2CP-C. Anaeromyxobacter spp. exhibit anaerobic growth and
19. S O R A N G ~ UCELLULOSUM M
33 7
other and form “biosynthetic gene clusters” often spanlack the ability to form fruiting bodies characteristic for ning genomic regions larger than 50 kbp. It has been myxobacteria (Sanford et al., 2002). M. xanthus represhown that the genomes of many bacterial producers of sents the best-studied myxobacterium, and its genome is secondary metabolites contain more biosynthetic gene discussed in chapter 16 of this book. S. cellulosum So clusters responsible for the production of the secondary ce56 was chosen as an additional model strain because of metabolites than could be expected from the number some advantageous and reproducible features: this bacof known compounds isolated from the corresponding terium as well as other myxobacteria is able to glide in bacterial strain (Silakowski et al., 2001; Bentley et al., swarms over solid surfaces, and it shows a complex life2002; Ikeda et al., 2003; Bode and Muller, 2005). This style including cooperative and cellular morphogenesis that is not lost when growing in liquid cultures. S. celfinding is also true for S. cellulosum So ce56. In addilulosum So ce56 grows in homogeneous submerged cultion to the biosynthetic gene clusters involved in the tures with a relatively short generation time of about 7 h, biosynthesis of chivosazol, etnangien, and myxochelin, some other chromosomal loci carry genes encoding prowhich makes this strain particularly suitable for molecuteins similar to polyketide synthases (PIG) or nonribosomal lar biological and genetic applications, especially since the peptide synthetases (NRPS), which are known to be genetic tools for this microorganism have already been responsible for the formation of many microbial secondestablished (see below). This strain is also known as a proary metabolites (see chapter 15).These PKS and NRPS are ducer of at least three secondary metabolites-chivosazol, S. probably involved in the biosynthesis of natural products etnangien, and myxochelin. All these features describe that have not been detected under laboratory conditions cellulosum So ce56 as an outstanding model organism so far. Genetic approaches are useful to activate such sofor a functional genome project. The genome sequenccalled “silent” genes and express them either in the same ing is being performed within the Bielefeld GenoMik netorganism by modifying the regulation processes or in a work of the German Ministry of Education and Research (http://~~~.genetik.uni-bielefeld.de/GenoMik/cluster6.suitable heterologous host. The information obtained from the genome data will be helpful to investigate natuhtml) and provides insight into the biology of the Sorangium group of proficient secondary metabolite producral product biosynthesis and explore the complex regulatory networks involved in morphological differentiation ers. After establishing the genome sequence starting from as well as primary and secondary metabolism. a whole-genome shotgun sequencing approach (Kaiser et al., 2003), the project has recently been finished with Uncommon Plant-Like Genes in S. cellulosum the functional annotation process. With a genome size of approximately 13 Mbp, S. cellulosum So ce56 harbors In the “pregenomic” era Sorangium strains had been extensively analyzed for secondary metabolite producthe largest prokaryotic genome known to date (the size tion, and more recently, genes involved in the biosynof 12.3 Mbp has been determined by macrorestriction thetic processes have been described. Unexpectedly, analyses [Pradella et al., 20021). It is expected that the molecular biological studies revealed the presence of genome encodes approximately 10,000 genes, which represents significantly more genes than were found in some unusual enzymes and genes which were formerly numerous eukaryotes, e.g., the yeast Schizosaccharobelieved to occur exclusively in plants. myces pombe (4,824 genes) (Wood et al., 2002) or SacA ddc gene product, which was previously found only charornyces cerevisiae (5,885 genes) (Goffeau et al., in eukaryotes, has also been identified in Sorangium strains. The gene product from S. cellulosum So ce90, 1996).Other well-established and completely sequenced Ddc (L-dopa decarboxylase), converts L-dihydroxy phebacterial producers of secondary metabolites also nylalanine (L-dopa) to dopamine (Muller et al., 2000). belong to the group of prokaryotes with exceptionally The function of the gene in S. cellulosum So ce90 is large bacterial genomes and a complex lifestyle (http:// unknown, but the activity of the enzyme has been experiwww.genomesonline.org; http://www.tigr.org/tdb/md b/ mentally proven after heterologous expression in Eschemdbinprogress.htm1) (e.g., Streptomyces coelicolor richia coli. This enzyme is more related to plant enzymes (8.67 Mbp), S. avermitilis (9.03 Mbp), 211. xanthus than to animal or bacterial amino acid decarboxylases (9.45 Mbp), and Nostoc punctiforme (9.2 Mbp) (Bode and Miiller, 2005). Interestingly, there seems to be (AADs).In plants AADs typically catalyze the decarboxylation of tyrosine, L-dopa, or tryptophan and represent some positive correlation between the genome size and a branching point from primary into secondary metabothe number of genes involved in secondary metabolism (Konstantinidis and Tiedje, 2004). lism, as the reaction products tyramine and dopamine provide the organisms with precursors for the formation The biosynthetic genes directing the formation of secof alkaloids (Facchini et al., 2000; Facchini, 2001). The ondary metabolites are mostly located adjacent to each
338 Ddc-encoding gene was also identified in S. cellulosum So ce56, and this Ddc is phylogenetically related to plant and animal AADs as well (Bode and Muller, 2003; 0. Perlova and R. Miiller, unpublished data). However, with the increasing number of sequenced genomes the number of putative ddc genes also grows, although these genes do not represent common attributes of the genomes. Genes with high similarity to ddc from S . cellulosum So ce90 were found, for example, in Solibacter usitatus (44% identity on amino acid level), Pseudomonas putida (42% identity), Mezorhizobium loti (41%), Yersinia pestis (40%), and other organisms with lower similarity. The corresponding gene products have not been characterized functionally, and these findings are based on the similarity of amino acid composition only. Since hybridization experiments under low-stringency conditions showed that the ddc gene is not widespread among other myxobacteria, its presence in some Sorangium strains might be a result of horizontal gene transfer (Muller et al., 2000). Phenylalanine ammonia lyase (PAL)-encoding genes represent another example of plant-like genes found in Sorangium. Like ddc, PAL-encoding genes are rarely found in prokaryotes (Xiang and Moore, 2005). In higher plants PAL catalyzes the nonoxidative deamination of phenylalanine to cinnamic acid, which is involved in the biosynthesis of phenylpropanoids. As early as 1970 it was shown that PAL is involved in the biosynthesis of cinnamate in Streptomyces verticillatus. It was thought that cinnamate serves as a biosynthetic precursor for benzoyl-coenzyme A (CoA),a starter molecule in secondary metabolite formation. Recently, the first bacterial PAL-encoding gene, encP, was found in the marine bacterium “Streptomyces maritimus” and EncP was characterized as specific for L-phenylalanine. The protein shares many biochemical properties with eukaryotic PALS (Xiangand Moore, 200.9, although the gene is more similar to prokaryotic histidine ammonia lyases (HALs). In “S. maritimus” EncP is involved in the biosynthesis of wailupemycin and enterocin (Hertweck and Moore, 2000). Whereas HAL enzymes are common in bacteria, the enzymes with PAL and tyrosine-ammonia lyase (TAL) activity are very rare in bacteria. The first bacterial TAL is characterized by a higher affinity for tyrosine than for phenylalanine and was isolated from Rhodobacter capsulatus (Kyndt et al., 2002). Recently, a second TAL was biochemically characterized from the actinomycete Saccarothrix espanaensis, although its sequence was highly similar to that of prokaryotic HALs (Berner et al., 2006). In myxobacteria phenylalanine degradation via PAL might be a widespread feature as PAL activity has been shown indirectly in the soraphen-producing
STIGMATELLA AND SORANGIUM S. cellulosum strain So ce26, and genes similar to encP were identified in Stigmatella aurantiaca (Silakowski et al., 2001) and in the the model strain S. cellulosum So ce56. In S. cellulosum So ce26 PAL seems to be involved in the formation of benzoyl-CoA, which is incorporated into the soraphen molecule (Fig. 6) (Bode and Muller, 2003; Gerth et al., 2003). In addition to soraphen, other secondary metabolites-crocacin (Jansen et al., 1999), phenalamid (Trowitzsch-Kienast et al., 1992), and thiangazol (Kunze et al., 1993; Jansen et al., 1993)-with a benzoic acid moiety that might also require a PAL activity for the biosynthesis were isolated from the myxobacteria belonging to genera Chondromyces, Myxococcus, and Polyangium (Fig. 6). In addition to L-dopa decarboxylase or PAL genes there are more genes typical for plants that can be found in Sorangium strains. Type I11 PKS similar to the superfamily of plant stilbene synthases and chalcone synthases were also found in S. cellulosum So ce56 (Gross et al., 2006a). In plants, these enzymes are implicated in the biosynthesis of phytoalexins and flavonoids for which a wide range of biological activities have been elucidated. In bacteria, the type I11 PKS RppA is used for the biosynthesis of flaviolin via 1,3,6,8-tetrahydroxynaphthalene and various secondary metabolites containing a naphtoquinone ring (Funa et al., 1999). The DpgA type I11 PKS catalyzes the formation of the nonproteinogenic amino acid DPG [(S)-3,5-dihydroxyphenylglycine](Li et al., 2001; Pfeifer et al., 2001). Although very few type I11 PKS have been characterized biochemically, the number of described genes from bacteria increases rapidly with the growing number of sequenced bacterial genomes. In S. cellulosum So ce56 two type I11 PKS genes have been identified. One of them is similar to bacterial RppA genes (61% identity), and the second gene shows 32% identity to plant chalcone synthases. These genes are thought to be “silent” under the tested laboratory conditions; their function in vivo is not clear, as no corresponding products could be isolated from S. cellulosum. Using molecular biological tools it was possible to activate the rppA-like gene in the heterologous host P. putida and to determine the product of the corresponding protein (see below) (Gross et al., 2006a). The function of the second type I11 PKS gene from S. cellulosum So ce56 remains to be determined.
Molecular Biological Tools for S . cellulosum To investigate the function of certain proteins in an organism, the inactivation of the corresponding genes is often required. Although S. cellulosum strains are known from extensive investigations as producers of many interesting secondary metabolites, a detailed study of biosynthetic
339
19. SORANGIUMCELLULOSUM
Crocacin A
a OMe
Phenalamid
Q
b
oO % - H cH = \c a/
7% H,N+-CH I COOH phenylalanine
E5 0
acid
E4
SCoA
''
o >H2 cH2 - c ~
SCoA
benzoyl-CoA
Figure 6 (a) Structures of myxobacterial secondary metabolites with a benzoic acid moiety. (b) Benzoyl-CoA biosynthesis in S. cellulosum So ce26 (soraphen producer). Mutants E4 and E5 are nonproducer mutants. Mutant E4 excretes traces of cinnamic acid and high concentrations of phenyl propionic acid into the culture supernantant. Mutant E5 recovers the ability of soraphen production in the presence of these compounds. From Gerth et al. (2003) with the permission of Elsevier, B.V.
mechanisms and regulation is difficult because the genetic manipulation in most Sorungium strains is not a routine procedure yet (Kopp et al., 2004). It is therefore necessary to develop and to adapt novel molecular biological techniques including DNA transfer and mutagenesis
systems. In 1992, the transfer of foreign DNA into S. cellulosum by conjugation using mobilizable E. coli plasmids was described (Jaoua et al., 1992).The transfer efficiency into Sorungium strains is low, which triggered the establishment of a variety of protocols since that time.
340 Pradella et al. described an improved triparental mating protocol for S. cellulosum So ce56 in 2002 (Pradella et al., 2002). Later it was reported that the same procedure of DNA transfer is not applicable for other Sorungium strains and the protocol needs to be optimized for each strain (Kopp et al., 2004). Since no plasmid replicating in Sorungium strains could be found, foreign DNA has to be integrated into the chromosome for all molecular engineering purposes. Although this makes Sorangium difficult to handle, some success was achieved in developing genetic manipulation systems: (i) transposon mutagenesis was established; (ii)conjugational transfer using biparental mating in S. cellulosum is possible; (iii) homologous recombination via single crossover leads to gene disruption; and (iv)an electroporation protocol has been established.
Transposon Mutagenesis Transposon mutagenesis systems are very helpful for identifying the gene clusters involved in natural products biosynthesis and for understanding the regulation of this process; they are also particularly important for the functional genome project of S. cellulosum. Two different research groups developed transposons that are useful in S. cellulosum (Julien and Fehd, 2003; Kopp et al., 2004). Both transposons are based on the eukaryotic muriner family of transposons. These transposons that require only the dinucleotide TA recognition site for integration into the chromosome not only are already broadly used in eukaryotic cells but also function in archaea and eubacteria, among them the myxobacterium 211. xanthus (Zhang et al., 1998; Rubin et al., 1999; Golden et al., 2000; Zhang et al., 2000; Youderian et al., 2003). One of the first transposons used in Sorangium-the conjugative plasmid pKOS183-3harbors the mariner tnp gene under the Lac1 repressible T7A1 promoter plus inverted repeats flanking bleomycin and kanamycin resistance genes (Julien and Fehd, 2003). This construct has been shown to enable transposition in the epothilone producer S. cellulosum So ce90 with an efficiency greater than per cell. In strain So ce12 the efficiency was significantly lower, indicating again that every protocol has to be optimized for each Sorangium strain. Another muriner-based transposon carries the hygromycin resistance gene (which is useful as a selection marker in Sorungium), the transposase (under the control of the mycobacterial T6 promoter), the oriT region (which allows conjugational transfer into Sorungium strains), and in addition, a conditional E. coli origin of replication (hpir dependent). This origin allows the recovery of the transposon DNA that was inserted into
STIGMATELLA AND SORANGIUM the chromosome together with the adjacent parts of host DNA from the mutants after successful transposition (“vector recovery”) (Fig. 7) (Kopp et al., 2004). The obtained mutants can be screened for different phenotypes in comparison to the wild type, e.g., via bioassays or HPLC for secondary metabolite production. With this method four disorazol-negative mutants were identified in S. cellulosum So ce12 by screening of 1,100 transposon mutants. Recovery of the transposon allowed the isolation of the biosynthetic and regulatory genes for disorazol from the bacterial artificial chromosome library, determination of the whole sequence, and consequently, the postulation of the hypothetical pathway to disorazol (Kopp et al., 2004).
Gene Inactivation by Insertion Resulting from Homologous Recombination Another way to inactivate a gene of interest in Sorangium is the integration of a selection marker into the chromosomal region encoding the corresponding gene by homologous recombination (so far only single-crossover experiments were reported from Sorungium). For this purpose, a DNA fragment approximately 1,000 bp in size is required. If the whole-genome sequence is not available, it is also possible to identify the biosynthetic genes by using an inactivation fragment, which can be obtained based on sequence similarity to known PKS- or NRPS-encoding genes. The homologous fragment can be amplified using degenerate PCR primers. This fragment is then cloned into a suitable vector and introduced into the bacterial strain for inactivation. The application of this method in S. cellulosum has led to the identification of the chivosazol biosynthetic gene cluster in the model strain So ce56 (Fig. 8). Two different IG fragments were obtained by PCR starting from chromosomal DNA prior to genome sequencing and used for inactivation. Subsequent analysis of phenotypes with HPLC and bioassays showed a lack in the production of chivosazol from which it could be concluded that both mutants belong to the same biosynthetic gene cluster. Further sequence analysis enabled the identification of the complete nucleotide sequence and proposal of a biosynthetic hypothesis (Perlova et al., 2006). (Fig. 8). Homologous recombination and consequential gene disruption has been described in S. cellulosum So ce26 for genes involved in the biosynthesis of soraphen (Schupp et al., 1995) and in swarming motility (Zirkle et al., 2004b), in So ce90 for the disruption of the PKS encoding genes involved in epothilone biosynthesis (Molnar et al., 2000), and in So ce690 for the identification of sorangicin biosynthetic genes (M. Kopp and R. Muller, unpublished data).
19. SORANGIUM CELLULOSUM
341
I
Transposase
transposition Primer I
Primer 2
A''...\
chromosomal DNA isolation digestion, e.g. with Mlul religation Primer I
analysis of mutants, e.g. bioassay Southern hybridization
Primer 7
d
C romosomal mutant DNA
1
sequencing and identification of target gene
Figure 7 Transposon mutagenesis in S. cellulosum. (a) mariner-based transposon. IR, inverted repeats, PaphII,promoter of the aphII gene; SZ, transcription terminator of the hygromycin resistance gene (HygR);oriRGKy, conditional origin of replication. (b) Transposon region when integrated into the chromosome. (c) Transposon recovery, consisting of ligation of chromosomal DNA from mutants after restriction with an enzyme which does not cut inside the transposed element (e.g., MluI). Using Primer 1 and Primer 2 the flanking chromosomal regions can be sequenced from the recovered plasmid. (d) Analysis of mutants, using a bioassay (e.g., comparison of nonproducers of chivosazol obtained by transposon mutagenesis with the wild type, which shows an inhibition zone on the s. cerevisiae indicator plate) (Kopp et al., 2004).
The phenotypic analysis of the mutants is expected to result in further insights into the physiology of Sorangium and to shed light onto the molecular mechanisms of natural product biosynthesis and regulation. It was already shown by combining different molecular biological techniques such as gene inactivation, regulatory protein fishing, and transcriptional analysis of chivosazol biosynthetic gene cluster expression by quantitative PCR that the ChiR-like protein acts as a direct or indirect activator of chivosazol gene expression and
binds to the promoter region of the biosynthetic gene cluster. Moreover, this protein is somehow involved in the regulation of the morphological differentiation in S. cellulosum, as the mutants lacking chiR are not able to differentiate and form fruiting bodies (S. Rachid and R. Muller, unpublished data; see chapter 15). Such studies together with the sequence analysis of the largest bacterial genome will result in deciphering complex regulatory networks, which seem to occupy a large coding capacity of the genome.
STIGMATELLAAND SORANGIUM
342
a
Conjugation single crossover
x
chromosome Mutant 2
Mutant 1
b off2 chiB
O r f l O orfl
chiA
C
Otf3
Orf4
chic
chiD
Mutant 1
I
I
,
I
I
0
2
4
6
8
I
10
orf6 chiF
O f l
chiG
d
Chivosazoles
1
Orf5 Chi€
>
12
,
14
I
16
Mutant 2
Wild-type
*
18
Time [min]
Figurer 8 Identification of the chivosazol biosynthetic gene cluster by inactivation of PKS genes using plasmid integration by homologous recombination. (a) Inactivation plasmid containing the selection marker HygRand a homologous region obtained by PCR using degenerate PKS primers. (b)Biosynthetic gene cluster; localization of mutations is marked. ( c )HPLC chromatogram of culture extracts of S. cellulosum So ce56 wild type (wt) and chivosazolnegative mutants (Mutant1 and Mutant2). (d) Bioassay for chivosazol production using Hansenula anomala.
Recent Improvements in the Molecular Biology of Sorungium The development of genetic manipulation systems for Sorangium (transposon mutagenesis and insertion inactivation) was further improved by an electroporation protocol for foreign DNA transfer, which was successfully applied for So ce12 (Kopp et al., 2005). In contrast to other myxobacteria (for example, M. xanthus), a longer incubation time for the phenotypical expression of the selection marker is needed. About 20 h (more than two doubling times) was used to enable the transposition into the S . cellulosum So ce12 genome and the expression of the hygromycin resistance. Electroporation is a more convenient and efficient method than conjugation, and the development of this technique for Sorangium
strains is a significant improvement in the development of the genetic manipulation system. Moreover, protocols for quantitative reverse transcriptase PCR and proteome analysis in Sorangium have been established (Kegler et al., 2006; Y. Elnakady, A. Alici, R. Miiller, and K. Niehaus, unpublished data). Interestingly, it has been shown for the first time by realtime PCR analysis that the presumed large secondary metabolic transcripts (exemplified by the chivosazol and etnangien transcripts from So ce56, which are assumed to be approximately 90 kbp in size) seem to be unusually stable, with a half-life time of 30 min in contrast to control transcripts (e.g., phosphopantetheinyl transferase from So ce56), which were shown to be degraded completely after 10 to 15 min (Kegler et al., 2006).
19. SORANGIUMCELLULOSUM Identified Secondary Metabolite Biosynthetic Gene Clusters from Sorungium Strains Recently investigations concerning the molecular basis of the biosynthesis of secondary metabolites in strains of the genus Sorungium have become more extensive (Schupp et al., 1995; Julien et al., 2000; Tang et al., 2000; Ligon et al., 2002; Pradella et al., 2002; Gerth et al., 2003; Kopp et al., 2004, 2005; Perlova et al., 2006). Chivosazol, disorazol, soraphen, and epothilone genes represent examples for published biosynthetic gene clusters. Disorazol and chivosazol genes were analyzed recently (Carvalho et al., 2005; Kopp et al., 2005; Perlova et al., 2006), and the genetic basis of other secondary metabolites biosyntheses is currently being investigated (leupyrrin, sorangicin, etnangien, and spirangiene; 0. Perlova, M. Kopp, B. Frank, and R. Muller, unpublished data). The disorazol and chivosazol biosynthetic gene clusters belong to a group of lately described truns-acyltransferase (trans-AT)type I PKS (Cheng et al., 2003; Piel et al., 2004) with a discrete AT domain located on a separate protein (Kopp et al., 2005; Perlova et al., 2006). In addition, both gene clusters are characterized by highly unusual properties such as “split-modules” located on separate proteins and hybrid proteins containing NRPS and PKS modules on the same polypeptide (Wenzel and Miiller, 2005). Both biosynthetic gene clusters showed additional exceptions from the textbook rules, which are based on work with better-investigated secondary metabolite biosynthetic systems (Carvalho et al., 2005; Kopp et al., 2005; Perlova et al., 2006). More extensive biochemical studies are required to explain the details of the biosynthesis in each case.
Heterologous Expression of the Sorungium Biosynthetic Genes Although S. cellulosum strains possess great biotechnological importance, they are slow-growing bacteria and it is therefore desirable to produce the natural products of these microorganisms (e.g., epothilone as a potential anticancer agent, see above) in an alternative host with a better fermentation potential and thus enable the overproduction of the substance and/or generate altered products. Different heterologous expression systems are currently being developed (seeJulien and Shah, 2002; Zirkle et al., 2004a; and Wenzel et al., 2005). But the heterologous expression remains a challenging task, and only a relatively small number of successfully expressed genes in foreign strains are known to date. Several examples in the literature describe not only the expression of the biosynthetic gene clusters in phylogenetically related strains (e.g., expression of bacitracin from Bacillus licheniformis
343 in B. subtilis [Eppelmann et al., 20011 and expression of griseorhodin A from an environmental Streptomyces isolate in S. Zividuns [Li and Piel, 20021) but also the successful expression in less related organisms. Some efforts have been made for the heterologous expression of the biosynthetic gene clusters from Sorungium. The biosynthetic genes for the production of epothilone, soraphen, and flaviolin from Sorungium strains were expressed in different heterologous hosts (Tang et al., 2000; Julien and Shah, 2002; Zirkle et al., 2004a; Gross et al., 2006a). In 2000, genes for epothilone biosynthesis were successfully expressed in the nonrelated streptomycete S. coelicolor (Tang et al., 2000). In this experiment, the expression of the biosynthetic genes cloned into the plasmids was driven by the act1 promoter. The transformants produced epothilones A and B and, after the deletion of the epoK gene, the epothilones C and D. The initial yield of epothilones in S. coelicolor was 50 to 100 pg/liter, which is probably due to the bacteriostatic effect of this compound in the host strain. Next, the epothilone genes were introduced stepwise from different cosmids into the chromosome of M. xunthus, assuming that this microorganism possesses all required characteristics for the heterologous biosynthesis of epothilone (Julien and Shah, 2002), as this strain is a secondary metabolite producer itself (Wenzel et al., 2006; Simunovic et al., 2006). Despite some advantages which the related myxobacterial strain offers for the biosynthesis, the yields of heterologously produced epothilones are still low in comparison to yield improvements that have been achieved by classical strain mutagenesis of S. cellulosum. One of the disadvantages in using M. xunthus is that the genes had to be integrated stepwise into the chromosome from different cosmids because of a lack of plasmids in myxobacteria. However, the fermentation process in M. xunthus could be optimized by incorporating an adsorber resin, the identification of a suitable carbon source, the supplement of trace metals, and the implementation of a fed-batch culture. The yield of the produced epothilones increased from 0.16 to 23 mg/liter (Lau et al., 2002), which is comparable to the natural producer but still far from the titer of the optimized S . cellulosum strain C18-1. Recently, the heterologous production of epothilone in E. coli has also been achieved by the researchers from Kosan Biosciences (Mutka et al., 2006). However, the synthetic epo genes had to be designed for this purpose and introduced into the engineered E. coli strain, which had to undergo several modifications to allow the expression of each single protein of the gene cluster. Moreover, the expression of the largest protein-EpoD (765 kDa)-required separation into two polypeptides. Successful epothilone
STIGMATELLAAND SORANGIUM
344 production was possible only in combination with a lower temperature, the coexpression of chaperones, and alternative promoters. Another example which sheds light onto the molecular level of the biosynthetic machinery is the heterologous production of the antifungal polyketide soraphen A of S. cellulosum So ce26 by introducing the biosynthetic genes into S. lividuns (Zirkle et al., 2004a). These genes were cloned on two integrative plasmids and on an autonomously replicating expression plasmid. In order to facilitate soraphen biosynthesis, feedings with different precursors of soraphen were required. Although the yields of heterologously produced soraphen are still low (less than 0.3 mg/liter) and the host strain needs further improvement, this system allows us to draw conclusions about the genetics of the biosynthesis and the formation of the soraphen “glycolate” polyketide extender unit of unclear biosynthetic origin (Zirkle et al., 2004a). Recent studies have shown that also pseudomonadswhich are known as producers of secondary metabolites, have a codon usage and a GC content similar to that of myxobacteria, possess an intrinsic broad-substrate Ppant transferase (Gross et al., 2005), and grow very fast in culture-represent suitable hosts for the heterologous expression of myxobacterial gene clusters (Wenzel et al., 2005). This was demonstrated recently for the expression of a type I11 PIC5 protein from s. cellulosum So ce56 (Gross et al., 2006a). As has been described above (see the section on plantlike genes above), s. cellulosum So ce56 possesses two uncommon plant-like genes which seem to encode gene products similar to chalcone synthases. Extensive analysis of the secondary metabolites from this strain did not show any products which could be correlated to these genes, as inactivation mutants did not show phenotypical properties different from those of the wild type. Under the tested laboratory conditions both genes seem to be silent. The heterologous expression of one of these genes (rppA-like gene similar to other bacterial type I11 PKS, see above), however, helped to find a novel myxobacterial metabolite, the formation of which is catalyzed by the gene product (Gross et al., 2006a). HPLC analysis as well as mass spectrometry and nuclear magnetic resonancebased structure elucidation led to the conclusion that the biosynthetic product is flaviolin, which is also responsible for the pigmentation of the liquid culture of the heterologous host P. putida (Color Plate 8). Although it has been shown that P. putidu is a suitable heterologous host for some myxobacterial biosynthetic genes (Wenzel et al., 2005; Gross et al., 2005, 2006a), the system has to be further engineered for the expression of other gene
clusters. This was shown recently for the expression of the myxobacterial secondary metabolite myxothiazol, which required the introduction of the genes for the formation of methylmalonyl-CoA as biosynthesis precursor in P. putida (Gross et al., 2006b). Alternatively, the newly described group of fastgrowing thermophilic myxobacteria, among them also Sorungium strains, might replace the slow-growing isolates in favor of a cost-saving production of myxobacterial secondary metabolites; this group of myxobacteria can probably also be used as heterologous hosts for the expression of gene clusters from slow-growing strains (Gerth and Miiller, 2005). As discussed in this chapter, the fascinating microorganisms of the genus Sorangium attract more and more attention, because they undergo a complex life cycle, possess the largest bacterial genomes known to date, and show a high potential as producers of biotechnologically important natural products. Therefore, many new techniques are being developed to make work with these microorganisms more effective. When applied to Sorungium, these new methods provide the opportunity to obtain results which were not imaginable some years ago and to access and manipulate the enormous diversity of the natural products derived from these myxobacteria. Dedicated to Prof. J. E. Peterson for his pioneering work on Sorangium. K.G. and O.P. contributed equally to this chapter.
References Bentley, S. D., I<. F. Chater, A. M. Cerdeno-Tarraga, G. L. Challis, N. R. Thomson, K. D. James, D. E. Harris, M. A. Quail, H. Kieser, D. Harper, A. Bateman, S. Brown, G. Chandra, C. W. Chen, M. Collins, A. Cronin, A. Fraser, A. Goble, J. Hidalgo, T. Hornsby, S. Howarth, C. H. Huang, T. Kieser, L. Larke, L. Murphy, K. Oliver, S. O’Neil, E. Rabbinowitsch, M. A. Rajandream, K. Rutherford, S. Rutter, K. Seeger, D. Saunders, S. Sharp, R. Squares, S. Squares, K. Taylor, T. Warren, A. Wietzorrek, J. Woodward, B. G. Barrell, J. Parkhill, and D. A. Hopwood. 2002. Complete genome sequence of the model actinomycete Streptomyces coelicolor A3(2).Nature 417:141-147. Berner, M., D. Krug, C. Bihlmaier, A. Vente, R. Miiller, and A. Bechthold. 2006. Genes and enzymes involved in caffeic acid biosynthesis in the actinomycete Saccharothrix espanaensis. J. Bacteriol. 188:2666-2673. Bode, H. B., and R. Miiller. 2003. Possibility of bacterial recruitment of plant genes associated with the biosynthesis of secondary metabolites. Plant Physiol. 132:1153-1161. Bode, H. B., and R. Miiller. 2005. The impact of bacterial genomics on natural product research. Angew. Chern. Int. Ed. Engl. 44:6828-6846. Brenner, D. J., N. R. Krieg, and J. T. Staley. 2005. The alpha-, beta-, delta-, and epsilonproteobacteria. In G. M. Garrity (ed.), Bergey’s Manual of Systematic Bacteriology, vol. 2. The Proteobacteria. Springer, New York, NY.
19. SORANGIUMCELLULOSUM Carvalho, R., R. Reid, N. Viswanathan, H. Gramajo, and B. Julien. 2005. The biosynthetic genes for disorazoles, potent cytotoxic compounds that disrupt microtubule formation. Gene 359:91-98. Cheng, Y. Q., G. L. Tang, and B. Shen. 2003. Type I polyketide synthase requiring a discrete acyltransferase for polyketide biosynthesis. Proc. Natl. Acad. Sci. USA 100:3 149-3154. Coucke, P., and J. P. Voets. 1967. The mineral requirements of Polyangium cellulosum. 2.Allg. Microbiol. 7:175-182. Coucke, P., and J. P. Voets. 1968. Etude de la cellulolyse enzymatique par Sorangium compositum. Ann. Inst. Pasteur 116549-5 60. Dawid, W. 2000. Biology and global distribution of myxobacteria in soils. FEMS Microbiol. Rev. 24:403-427. Eppelmann, K., S. Doekel, and M. A. Marahiel. 2001. Engineered biosynthesis of the peptide antibiotic bacitracin in the surrogate host Bacillus subtilis. J . Biol. Chem. 276:3482434831. Facchini, P. J., K. L. Huber-Allanach, and L. W. Tari. 2000. Plant aromatic-amino acid decarboxylases: evolution, biochemistry, regulation, and metabolic engineering applications. Phytochemistry 54:121-138. Facchini, P. J. 2001. Alkaloid biosynthesis in plants: biochemistry, cell biology, molecular regulation, and metabolic engineering applications. Annu. Rev. Plant Physiol. Plant Mol. Biol. 52:29-66. Finck, G. 1950. Biologische und stoffwechselphysiologische Studien an Myxococcaceen. Arch. Mikrobiol. 15:358-388. Funa, N., Y. Ohnishi, I. Fujii, M. Shibuya, Y. Ebizuka, and S. Horinouchi. 1999. A new pathway for polyketide synthesis in microorganisms. Nature 400:897-899. Gaspari, F., Y. Paitan, M. Mainini, D. Losi, E. Z. Ron, and F. Marinelli. 2004. Myxobacteria isolated in Israel as potential source of new anti-infectives.J. Appl. Microbiol. 98:429-439. Gerth, K., N. Bedorf, H. Irschik, G. Hofle, and H. Reichenbach. 1994. The soraphens: a family of novel antifungal compounds from Sorangium cellulosum (Myxobacteria). I. Soraphen A l : fermentation, isolation, biological properties. J. Antibiot. 47:23-31. Gerth, K., B. Bedorf, G. Hofle, H. Irschik, and H. Reichenbach. 1996a. Epothilons A and B: antifungal and cytotoxic compounds from Sorangium cellulosum (Myxobacteria) production, physico-chemical and biological properties. 1. Antibiot. 49560-563. Gerth, K., P. Washausen, G. Hofle, H. Irschik, and H. Reichenbach. 199610. The jerangolids: a family of new antifungal compounds from Sorangium cellulosum (Myxobacteria). Production, physico-chemical and biological properties of jerangolid A.J. Antibiot. 49:71-75. Gerth, K., S. Pradella, 0. Perlova, S. Beyer, and R. Miiller. 2003. Myxobacteria: proficient producers of novel natural products with various biological activities-past and future biotechnological aspects with the focus on the genus Sorangium.]. Biotechnol. 106:233-253. Gerth, K., and R. Miiller. 2005. Moderately thermophilic myxobacteria: novel potential for production of natural products. Environ. Microbiol. 7:874-880. Goffeau, A., B. G. Barrell, H. Bussey, R. W. Davis, B. Dujon, H. Feldmann, F. Galibert, J. D. Hoheisel, C. Jacq,
345 M. Johnston, E. J. Louis, H. W. Mewes, Y. Murakami, P. Philippsen, H. Tettelin, and S. G. Oliver. 1996. Life with 6000 genes. Science 274546,563-567. Golden, N. J., A. Camilli, and D. W. Acheson. 2000. Random transposon mutagenesis of Campylobacter jejuni. Infect. Immun. 685450-5453. Gross, F., D. Gottschalk, and R. Miiller. 2005. Posttranslational modification of myxobacterial carrier protein domains in Pseudomonas sp. by an intrinsic phosphopantetheinyl transferase. Appl. Microbiol. Biotechnol. 68:66-74. Gross, F., N. Luniak, 0. Perlova, N. Gaitatzis, H. JenkeKodama, K. Gerth, D. Gottschalk, E. Dittmann, and R. Miiller. 2006a. Bacterial type I11 polyketide synthases: phylogenetic analysis and potential for the production of novel secondary metabolites by heterologous expession in pseudomonads. Arch. Microbiol. 185:28-38. Gross, F., M. W. Ring, 0. Perlova, J. Fu, S. Schneider, K. Gerth, S. Kuhlmann, F. Stewart, Y. Zhang, and R. Miiller. 2006b. Red/ET-subcloning and heterologous expression of methylmalonyl-CoA biosynthesis genes of Sorangium cellulosum So ce56 in Pseudomonas putida KT2440. Chem. Biol. 13:1253-1264. Hertweck, C., and B. S. Moore. 2000. A plant-like biosynthesis of benzoyl-CoA in the marine bacterium ‘Streptomyces maritimus’. Tetrahedron 56:9115-9120. Hofle, G., and H. Reichenbach. 1995. The biosynthetic potential of the Myxobacteria, p. 61-78. In W. Kuhn and H. P. Fiedler (ed.), Sekundarmetabolismus bei Mikroorganismen. Attempo Verlag, Tubingen, Germany. Hofle, G. 1996-2002. Isolation, structure elucidation and chemistry. In Scientific Annual Report. Gesellschaft fur Biotechnologische Forschung, Braunschweig, Germany. Hoischen, C. 1986. Untersuchungen zum Zuckerstoffwechsel von Sorangium cellulosum, p. 1-97. Fakultat fur Biologie der Universitat, Tubingen, Germany. Ikeda, H., J. Ishikawa, A. Hanamoto, M. Shinose, H. Kikuchi, T. Shiba, Y. Sakaki, M. Hattori, and S. Omura. 2003. Complete genome sequence and comparative analysis of the industrial microorganism Streptomyces avermitilis. Nat. Biotechnol. 21526-531. Imshenetski, A. A., and L. Solntseva. 1936. On aerobic cellulose-decomposing bacteria. Izv. Akad. Nuuk SSSR Cl. Sci. Math. Natl. Ser. Biol. 1936:1115-1172. (In Russian with English summary.) Imshenetski, A. A. 1959. Mikrobiologie der Cellulose. Akademie Verlag, Berlin, Germany. Irschik, H., R. Jansen, K. Gerth, G. Hofle, and H. Reichenbach. 1987. The sorangicins, novel and powerful inhibitors of eubacterial RNA polymerase isolated from myxobacteria. 1. Antibiot. 40:7-13. Irschik, H., R. Jansen, K. Gerth, G. Hofle, and H. Reichenbach. 1995a. Chivosazol A, a new inhibitor of eukaryotic organisms isolated from myxobacteria. J. Antibiot. 48:962-966. Irschik, H., R. Jansen, K. Gerth, G. Hofle, and H. Reichenbach. 199513.Disorazol A, an efficient inhibitor of eukaryotic organisms isolated from myxobacteria. J. Antibiot. 48:3 1-35. Jansen, R., D. Schomburg, and G. Hofle. 1993. Thiangazole, a new Tris(thiazo1ine) derivative from Polyangium spec.: absolute configuration. Liebigs Ann. 1993:701-704.
346 Jansen, R., P. Washausen, B. Kunze, H. Reichenbach, and G. Hofle. 1999. Antibiotics from gliding bacteria, LXXXIIIThe crocacins, novel antifungal and cytotoxic antibiotics from Chondromyces crocatus and Chondromyces pediculatus (Myxobacteria):isolation and structure elucidation. Eur. J. Org. Chem. 1999:1085-1089. Jaoua, S., S. Neff, and T. Schupp. 1992. Transfer of mobilizable plasmids to Sorangium cellulosum and evidence for their integration into the chromosome. Plasmid 28:157-165. Julien, B., S. Shah, R. Ziermann, R. Goldman, L. Katz, and C. Khosla. 2000. Isolation and characterization of the epothilone biosynthetic gene cluster from Sorangium cellulosum. Gene 249:153-160. Julien, B., and S. Shah. 2002. Heterologous expression of epothilone biosynthetic genes in Myxococcus xanthus. Antimicrob. Agents Chemother. 46:2772-2778. Julien, B., and R. Fehd. 2003. Development of a mariner-based transposon for use in Sorangium cellulosum. Appl. Environ. Microbiol. 69:6299-6301. Kaiser, O., D. Bartels, T. Bekel, A. Goesmann, S. Kespohl, A. Piihler, and F. Meyer. 2003. Whole genome shotgun sequencing guided by bioinformatics pipelines-an optimized approach for an established technique. J. Biotechnol. 106:121-133. Kegler, C., K. Gerth, and R. Miiller. 2006. Establishment of a real-time PCR protocol for expression studies of secondary metabolite biosynthetic gene clusters in the G/C-rich myxobacterium Sorangium cellulosum So ce56. 1. Biotechnol. 121:201-212. Konstantinidis, K. T., and J. M. Tiedje. 2004. Trends between gene content and genome size in prokaryotic species with larger genomes. Proc. Natl. Acad. Sci. USA 101:3160-3165. Kopp, M., H. Irschik, F. Gross, 0. Perlova, A. Sandmann, K. Gerth, and R. Miiller. 2004. Critical variations of conjugational DNA transfer into secondary metabolite multiproducing Sorangium cellulosum strains So ce12 and So ce56: development of a mariner-based transposon mutagenesis system. J. Biotechnol. 107:29-40. Kopp, M., H. Irschik, S. Pradella, and R. Miiller. 2005. Production of the tubulin destabilizer disorazol in Sorangium cellulosum: biosynthetic machinery and regulatory genes. ChemBioChem 6:1277-1286. Krzemieniewska, H., and S. Krzemieniewski. 1937a. Uber die Zersetzung der Zellulose durch Myxobakterien. Bull. Acad. Pol. Sci. Lett. C1. Sci. Math. Nut. B I:33-59. Krzemieniewska, H., and S. Krzemieniewski. 1937b. Die zellulosezersetzenden Myxobakterien. Bull. Acad. Pol. Sci. Lett. Cl. Sci. Math. Nut. B I:11-31. Kunze, B., R. Jansen, L. Pridzun, E. Jurkiewicz, G. Hunsmann, G. Hofle, and H. Reichenbach. 1993. Thiangazole, a new thiazoline antibiotic from Polyangium sp. (myxobacteria): production, antimicrobial activity and mechanism of action. J. Antibiot. 46:1752-1755. Kyndt, J. A., T. E. Meyer, M. A. Cusanovich, and J. J. Van Beeumen. 2002. Characterization of a bacterial tyrosine ammonia lyase, a biosynthetic enzyme for the photoactive yellow protein. FEBS Lett. 512:240-244. Lampky, J. R. 1971. Distribution of Sorangium cellulosum. Appl. Microbiol. Biotechnol. 22:93 7-93 8.
STIGMATELLAAND SORANGIUM Lau, J., S. Frykman, R. Regentin, S. Ou, H. Tsuruta, and P. Licari. 2002. Optimizing the heterologous production of epothilone D in Myxococcus xanthus. Biotechnol. Bioeng. 78:280-288. Li, A., and J. Piel. 2002. A gene cluster from a marine Streptomyces encoding the biosynthesis of the aromatic spiroketal polyketide griseorhodin A. Chem. Biol. 9:1017-1026. Li, T., 0. Choroba, H. Hong, D. Williams, and J. Spencer. 2001. Biosynthesis of the vancomycin group of antibiotics: characterisation of a type I11 polyketide synthase in the pathway to (S)-3,5-dihydroxyphenylglycine.Chem. Commun. 20~2156-2157. Ligon, J., S. Hill, J. Beck, R. Zirkle, I. Molnar, J. Zawodny, S. Money, and T. Schupp. 2002. Characterization of the biosynthetic gene cluster for the antifungal polyketide soraphen A from Sorangium cellulosum So ce26. Gene 285:257-267. McCurdy, H. D. 1969. Studies on the taxonomy of the Myxobacterales. I. Record of Canadian isolates and survey of methods. Can. J. Microbiol. 15:1453-1461. McCurdy, H. D., J . 1970. Studies on the taxonomy of the Myxobacterales 11. Polyangium and the demise of the Sorangiaceae. lnt. J. Syst. Bacteriol. 20:283-296. Mishustin, E. N. 1938. Cellulose-decomposing myxobacteria. Microbiologiya 7:427-444. Molnar, I., T. Schupp, M. Ono, R. Zirkle, M. Milnamow, B. Nowak-Thompson, N. Engel, C. Toupet, A. Stratmann, D. D. Cyr, J. Gorlach, J. M. Mayo, A. Hu, S. Goff, J. Schmid, and J. M. Ligon. 2000. The biosynthetic gene cluster for the microtubule-stabilizing agents epothilones A and B from Sorangium cellulosum So ce90. Chem. Biol. 7:97-109. Miiller, R., K. Gerth, P. Brandt, H. Blocker, and S. Beyer. 2000. Identification of an L-dopa decarboxylase gene from Sorangium cellulosum So ce90. Arch. Microbiol. 173:303-306. Miiller, R., and K. Gerth. 2006. Development of simple media which allow investigations into the global regulation of chivosazole biosynthesis with Sorangium celllulosum So ce56. J. Biotechnol. 121:192-200. Mutka, S. C., J. R. Carney, Y. Liu, and J. Kennedy. 2006. Heterologous production of epothilone C and D in Escherichia coli. Biochemistry 45:1321-1330. Neil, R. B., D. Hite, M. I. Kelrick, M. L. Lockhart, and K. Lee. 2005. Myxobacterial biodiversity in an established oakhickory forest and a savanna restoration site. Curr. Microbiol. 50638-95. Niggemann, J., N. Bedorf, U. Florke, H. Steinmetz, K. Gerth, H. Reichenbach, and G. Hofle. 2005. Spirangien A and B, highly cytotoxic and antifungal spiroketals from the Myxobacterium Sorangium cellulosum: isolation, structure elucidation and chemical modifications. Eur. J. Org. Chem. 235013-5018. Perlova, O., K. Gerth, A. Hans, 0.Kaiser, and R. Miiller. 2006. Identification and analysis of the chivosazol biosynthetic gene cluster from the myxobacterial model strain Sorangium cellulosum So ce56.J. Biotechnol. 121:174-191. Peterson, E. A., D. C. Gillespie, and F. D. Cook. 1966. A widespectrum antibiotic produced by a species of Sorangium. Can. J. Microbiol. 12:221-230. Peterson, J. E. 1965. Group of strongly cellulolytic Myxobacteria previously unreported in North American soils. Am. J. Bot. 52636.
19. SORANGIUM CELLULOSUM Peterson, J. E., and B. Norin. 1967. The occurrence of the cellulose-decomposing myxobacterium, Sorangium cellulosum, in Scandinavian soils. Am.]. Bot. 54:648. Peterson, J. E. 1969a. The fruiting Myxobacteria: their properties, distribution and isolation. ]. Appl. Bacteriol. 325-12. Peterson, J. E. 1969b. Isolation, cultivation and maintenance of the myxobacteria, p. 185-210. In J. R. Norris and D. W. Ribbons (ed.), Methods in Microbiology, vol. 3B. Academic Press, New York, NY. Pfeifer, V., G. J. Nicholson, J. Ries, J. Recktenwald, A. B. Schefer, R. M. Shawky, J. Schroder, W. Wohlleben, and S. Pelzer. 2001. A polyketide synthase in glycopeptide biosynthesisthe biosynthesis of the non-proteinogenic amino acid (S)-3,5dihydroxyphenylglycine. J. Biol. Chem. 276:38370-38377. Piel, J., D. Hui, N. Fusetani, and S. Matsunaga. 2004. Targeting modular polyketide synthases with iteratively acting acyltransferases from metagenomes of uncultured bacterial consortia. Environ. Microbiol. 6:921-927. Pradella, S., A. Hans, C. Sproer, H. Reichenbach, K. Gerth, and S. Beyer. 2002. Characterisation, genome size and genetic manipulation of the myxobacterium Sorangium cellulosum So ce56. Arch. Microbiol. 178:484-492. Reichenbach, H. 1992. The Genus Lysobacter, p. 3256-3275. In A. Balows, H. G. Truper, M. Dworkin, W. Harder, and K. H. Schleifer (ed.), The Prokaryotes. Springer-Verlag, New York, NY. Reichenbach, H., and G. Hofle. 1999. Myxobacteria as producers of secondary metabolites, p. 149-179. In s. Grabley and R. Thieriecke (ed.), Drug Discovery from Nature. Springer Verlag, Berlin, Germany. Reichenbach, R., and M. Dworkin. 1981. The order Myxobacterales, p. 328-355. In H. Stolp, M. P. Starr, H. G. Truber, A. Balows, and H. G. Schlegel (ed.), The Prokaryotes: A Handbook on Habitats, Isolation, and Identification of Bacteria, vol. I. Springer-Verlag KG, Berlin, Germany. Ringel, S. M., R. C. Greenough, S. Roemer, D. Connor, A. L. Gutt, B. Blair, G. Kanter, and M. von Strandtmann. 1977. Ambruticin" (W7783), a new antifungal antibiotic. J. Antibiot. 30:371-375. Rubin, E., B. Akerley, V. Novik, D. Lampe, R. Husson, and J. Mekalanos. 1999. In vivo transposition of mariner-based elements in enteric bacteria and mycobacteria. Proc. Natl. Acad. Sci. USA 96:1645-1650. Sanford, R. A., J. R. Cole, and J. M. Tiedje. 2002. Characterization and description of Anaeromyxobacter dehalogenans gen. nov., sp. nov., an aryl-halorespiring facultative anaerobic myxobacterium. Appl. Environ. Microbiol. 685393-900. Sarao, R., H. D. McCurdy, and L. Passador. 1985. Enzymes of the intermediary carbohydrate metabolism of Polyangium cellulosum. Can. J. Microbiol. 31:1142-1146. Schupp, T., C. Toupet, B. Cluzel, S. Neff, S. Hill, J. J. Beck, and J. M. Ligon. 1995. A Sorangium cellulosum (myxobacterium) gene cluster for the biosynthesis of the macrolide antibiotic soraphen A: cloning, characterization, and homology to polyketide synthase genes from actinomycetes. 1.Bacteriol. 177:3673-3679. Silakowski, B., B. Kunze, and R. Miiller. 2001. Multiple hybrid polyketide synthasehon-ribosomal peptide synthetase gene clusters in the myxobacterium Stigmatella aurantiaca. Gene 27.5~233-240.
347 Simunovic, V., J. Zapp, S. Rachid, D. Krug, P. Meiser, and R. Miiller. 2006. Myxovirescin biosynthesis is directed by an intriguing megasynthetase consisting of hybrid polyketide synthaseshonribosomal peptide synthetase, 3-hydroxy-3methylglutaryl CoA synthases and trans-acting acyltransferases. ChemBioChem 21206-1220. Singh, B. N., and H. R. Singh. 1971. Distribution of fruiting myxobacteria in Indian soils, bark of trees and dung of herbivorous animals. Indian ]. Microbiol. 11:47-92. Solntseva, L. 1939. Methoden zum Kultivieren der Myxobakterien. Microbiologiya 8:959-963. Sproer, C., H. Reichenbach, and E. Stackebrandt. 1999. The correlation between morphological and phylogenetic classification of myxobacteria. Int. J. Syst. Bacteriol. 49:12551262. Tang, L., S. Shah, L. Chung, J. Carney, L. Katz, C. Khosla, and B. Julien. 2000. Cloning and heterologous expression of the epothilone gene cluster. Science 287:640-642. Trowitzsch-Kienast, W., E. Forche, V. Wray, H. Reichenbach, E. Jurkiewicz, G. Hunsmann, and G. Hofle. 1992. Phenalamide, neue HIV-Inhibitoren aus Myxococcus stipitatus Mx s40. Liebigs Ann. Chem. 16:659-664. Weigele, M., and W. Leimgruber. 1967. The structure of myxin. Tetrahedron Lett. 1967:715-718. Wenzel, S. C., F. Gross, Y. Zhang, J. Fu, F. A. Stewart, and R. Miiller. 2005. Heterologous expression of a myxobacterial natural products assembly line in pseudomonads via redlET recombineering. Chem. Biol. 12:349-356. Wenzel, S. C. and R.Miiller. 2005. Formation of novel secondary metabolites by bacterial multimodular assembly lines: deviations from text book biosynthetic logic. Curr. Opin. Chem. Biol. 9:447-458. Wenzel, S. C., I? Meiser, T. Binz, T. Mahmud, and R. Miiller. 2006. Nonribosomal peptide biosynthesis: point mutations and module skipping lead to chemical diversity. Angew. Chem. Int. Ed. Engl. 45:2296-2301. Wood, V., R. Gwilliam, M. A. Rajandream, M. Lyne, R. Lyne, A. Stewart, J. Sgouros, N. Peat, J. Hayles, S. Baker, D. Basham, S. Bowman, K. Brooks, D. Brown, S. Brown, T. Chillingworth, C. Churcher, M. Collins, R. Connor, A. Cronin, P. Davis, T. Feltwell, A. Fraser, S. Gentles, A. Goble, N. Hamlin, D. Harris, J. Hidalgo, G. Hodgson, S. Holroyd, T. Hornsby, S. Howarth, E. J. Huckle, S. Hunt, K. Jagels, K. James, L. Jones, M. Jones, S. Leather, S. McDonald, J. McLean, P. Mooney, S. Moule, K. Mungall, L. Murphy, D. Niblett, C. Odell, K. Oliver, S. O'Neil, D. Pearson, M. A. Quail, E. Rabbinowitsch, I<. Rutherford, S. Rutter, D. Saunders, K. Seeger, S. Sharp, J. Skelton, M. Simmonds, R. Squares, S. Squares, K. Stevens, K. Taylor, R. G. Taylor, A. Tivey, S. Walsh, T. Warren, S. Whitehead, J. Woodward, G. Volckaert, R. Aert, J. Robben, B. Grymonprez, I. Weltjens, E. Vanstreels, M. Rieger, M. Schafer, S. Muller-Auer, C. Gabel, M. Fuchs, A. Dusterhoft, C. Fritzc, E. Holzer, D. Moestl, H. Hilbert, K. Borzym, I. Langer, A. Beck, H. Lehrach, R. Reinhardt, T. M. Pohl, P. Eger, W. Zimmermann, H. Wedler, R. Wambutt, B. Purnelle, A. Goffeau, E. Cadieu, S. Dreano, S. Gloux, V. Lelaure, S. Mottier, F. Galibert, S. J. Aves, Z. Xiang, C. Hunt, K. Moore, S. M. Hurst, M. Lucas, M. Rochet, C . Gaillardin, V. A. Tallada, A. Garzon, G. Thode, R. R. Daga, L. Cruzado, J. Jimenez, M. Sanchez, F. del Rey, J. Benito,
348 A. Dominguez, J. L. Revuelta, S. Moreno, J. Armstrong, S. L. Forsburg, L. Cerutti, T. Lowe, W. R. McCombie, I. Paulsen, J. Potashkin, G. V. Shpakovski, D. Ussery, B. G. Barrell, and P. Nurse. 2002. The genome sequence of Schizosaccharomyces pombe. Nature 415:871-880. Xiang, L., and B. S. Moore. 2005. Biochemical characterization of a prokaryotic phenylalanine ammonia lyase. J. Bacteriol. 187:4286-4289. Yan, Z. C., B. Wang, Y. Z. Li, X. Gong, H. Q. Zhang, and P. J. Gao. 2003. Morphologies and phylogenetic classification of cellulolytic Myxobacteria. Syst. Appl. Microbiol. 26:104-109. Youderian, P., N. Burke, D. J. White, and P. L. Hartzell. 2003. Identification of genes required for adventurous gliding motility in Myxococcus xanthus with the transposable element mariner. Mol. Microbiol. 49555-570. Zhang, J. K., M. A. Pritchett, D. J. Lampe, H. M. Robertson, and W. W. Metcalf. 2000. In vivo transposon mutagenesis
STIGMATELLAAND SORANGIUM of the methanogenic archaeon Methanosarcina acetivorans C2A using a modified version of the insect mariner-family transposable element Himarl. Proc. Natl. Acad. Sci. USA 97:9665-9670. Zhang, L., U. Sankar, D. J. Lampe, H. M. Robertson, and F. L. Graham. 1998. The Himarl mariner transposase cloned in a recombinant adenovirus vector is functional in mammalian cells. Nucleic Acids Res. 26:3687-3693. Zirkle, R., J. M. Ligon, and I. Molnar. 2004a. Heterologous production of the antifungal polyketide antibiotic soraphen A of Sorangium cellulosum So ce26 in Streptomyces lividans. Microbiology 150:2761-2774. Zirkle, R., J. M. Ligon, and M. Molnar. 2004b. Cloning, sequence analysis and disruption of the mglA gene involved in swarming motility of Sorangium cellulosum So ce26, a producer of the antifungal polyketide antibiotic soraphen A. 1.Biosci. Bioeng. 97:267-274.
Analogous Systems ~
Myxobacteria: Multicellularity and Differentiation Edited by David E. Whitworth 0 2008 ASM Press, Washington, D.C.
K. J. Evans, L. Hobley, C. Lambert, R. E. Sockett
BdeZZovibrio: Lone Hunter
2
“Cousin” of the “Pack Hunting” Myxobacteria
LIFESTYLE, PHYLOGENY, AND ECOLOGY OF BDELLOVIBRIO Bdellovibrio organisms are predatory members of the Deltaproteobacteria, a group consisting of bacteria with a variety of lifestyles and metabolic processes, including the sulfate-reducing Desulfovibrio, iron [Fe(III)]reducing Geobacter, and the myxobacteria, which are the main subject of this volume. As this chapter describes, Bdellovibrio and myxobacteria can be isolated from similar soil environments, and both employ an arsenal of hydrolytic enzymes to kill and digest other bacteria to provide for their own growth and division. Although they are both members of the Deltaproteobacteria, there are significant genomic differences between them. Also, while both express type IV pili, the myxobacteria use these for social motility on surfaces, while the Bdellovibrio are flagellate and fast swimming, and thus, the water content of the soil microenvironments which they inhabit may sometimes be quite different. As research with both bacteria has only recently benefited from genomic data, the molecular details of their related predatory lifestyles are still being revealed. Here we concentrate on the ecology and
motility systems of Bdellovibrio, followed by an overview of the hydrolytic enzymes used in prey digestion.
Predatory Life Cycle Bdellovibrio strains have a predatory lifestyle, but unlike the myxobacteria, which exude enzymes to digest an external “buffet” of diverse prey, BdelEovibrio isolates use flagellar motility and taxis to locate prey-rich environments (Lambert et al., 2003), and then each bdellovibrio penetrates a single, gram-negative prey cell, sealing itself within the prey’s periplasm to “dine alone” on its cytoplasmic contents and to elongate and then replicate (Fig. 1 and 2); at high predator-to-prey ratios, however, multiple infection may occur (Nunez et al., 2003). Only when the prey contents are exhausted and the bdellovibrios have septated into progeny cells do they resume external life by escaping the prey outer membrane (Fig. 1) (reviewed in Sockett and Lambert, 2004). There is not a strict relationship between Bdellovibrio and a single prey bacterium; predation by different Bdellovibrio organisms occurs over a range of gram-negative bacteria. How this range is specified is not yet understood; however, the ecology of Bdellovibrio is at least in part influenced by
K.J. Evans, L. Hobley, C. Lambert, and R. E. Sockett, Institute of Genetics, School of Biology, University of Nottingham Medical School, Nottingham NG7 2UH, United Kingdom.
351
ANALOGOUS SYSTEMS
352 Infrequent septation to give free-swimming Bdellovibrio
(ii)Bdellovibrio
on rich media
Figure 1 The predatory life cycle and host-independent growth phases of Bdellovibrio.
the diversity of heterotrophic and/or autotrophic prey bacteria in different niches (Jurkevitch et al., 2000). A fraction of any predatory population of Bdellovibrio can be cultured in a host-independent (HI) manner, axenically on rich growth media; such HI cells are not well understood, but they seem to mimic the filamentous phase of growth that predatory bdellovibrios undergo within the bdelloplast (Fig. 2B). Whether HI growth of Bdellovibrio occurs commonly outside laboratory environments has not yet been elucidated.
Phylogeny Bdellovibrio and related bacteria with a similar invasive predatory lifestyle, Bdellovibrio-like organisms (BLO) (Snyder et al., 2002; Davidov and Jurkevitch, 2004), are contained within the order Bdellovibrionales within one of two separate families: Bdellovibrionaceae and Bacteriovoracaceae (Fig. 3 ) .The only current known exception to this is the alphaproteobacterial Micavibrio which has been recently added to the BLO lineage, but not within the order Bdellovibrionales (Davidov et al., 2006).
Figure 2 Transmission electron micrographs of B. bacteriovorus attaching to E. coli DFB225 prey (A) and B. bacteriovorus forming a bdelloplast and preying upon E. coli DFB225 (B). Bars, 1 km. Both stained with 1%phosphotungstic acid (pH7).
20.
PREDATORY
LIFESTYLE O F BDELLOVIBRIO
I
353
1 Lx=l e Delta-Proteobacteria
I
Myxococcales
Cystobacterineae
Myxococcaceae
Bdellovibrio
Peredibacter
Bacteriovorax
bacteriovorus DKI 622T Cow dung, Iowa G+C = 69% Genome 9.2 Mb
HDIOOT Soil, California G+C = 50.7% Genome 3.8 Mb
A3.12T Soil, California G+C = 43.5%
Uki2T Soil, Kentucky G+C = 41.8%
SJT Coastal waters, US Virgin Islands G+C = 37.7 38.3% Genome 3.4Mb
-
JS!jT Crab gill, Maryland G+C = 37.8%
Ecological, genomic, and phylogenetic relationships between the type strains of Bdellovibrio and the myxobacteria. Figure 3
The Bdellovibrionaceae currently consist of one described genus, Bdellovibrio, which in turn is characterized by one species, Bdellovibrio bacteriovorus, with type strain HDIOOT (Stolp and Starr, 1963), for which a whole-genome sequence is published (Rendulic et al., 2004). In contrast, the Bacteriovoracaceae are divided into two genera, Bacteriovorax and Peredibacter. Peredibacter is currently described as containing one species, Peredibacter starrii (type strain A3 .12T) (Stolp and Starr, 1963; Baer et al., ZOOO), whereas the genus Bacteriovorax is characterized by three known species: Bacteriovorax marinus (typestrain SJT)(Schoeffieldet al., 1991; Baer et al., 2004), for which a genome sequence is being completed by the Sanger Centre; B. litoralis (JSST) (Schoeffieldet al., 1991; Baer et al., 2004); and B. stolpii (Uki2T)(Seidler et al., 1972; Baer et al., 2000). There has been one report of a gammaproteobacterial BLO isolate found at James Island in the United States (Snyder et al., 2002), but no further detail as to whether this is a true predatory bacterium has yet been published. Recent phylogenetic examination of 16s rRNA sequences of the Bdellovibrionales has shown that the two families do not form a monophyletic group (Davidov and Jurkevitch, 2004). The Bacteriovoracaceae have been shown to consistently cluster within the Deltaproteobacteria, whereas the Bdellovibrionaceae seem
to form an early branching group, which Davidov and Jurkevitch suggest may indicate the existence of a common predatory ancestor to the Deltaproteobacteria, which gave rise to both the Bdellovibrionales and the myxobacteria. The division between the Bdellovibrionales (Fig. 3 ) is reinforced by the GC content of each of the type strains. B. bacteriovorus HD1OOT,with a 3.85Mb genome, has a GC content of 50.7 mol% (Rendulic et al., 2004), whereas the GC content of the Bacteriovoracaceae ranges from between 37 to 44 mol% (Baeret al., 2004; Seidler et al., 1972). In contrast the Myxobacteria show a highly GC-biased genome content of 69 mol% and a genome size of 9.2 M b for type strain DK1622, with others ranging between 5 and 13.1 Mb. Insofar as GC bias and genome sizes give an indication of significant adaptive evolution, it is clear that the present-day bdellovibrios and the myxobacteria are very different from each other and that their grouping within the Deltaproteobacteria in some ways masks these differences. As we see later in this chapter, they can be isolated from similar terrestrial environments and they employ some similar biochemical processes in their predatory lifestyles. There are other predatory bacteria reported in the literature, apart from the Bdellovibrionales and the myxobacteria. One of these is the recently characterized BLO species Micavibrio, which phylogenetically belongs to
354 the Alphaproteobacteria (Davidov et al., 2006), others include bacteria belonging to the genus Ensifer within the order Rhizobiales, again Alphaproteobacteria, and a third set are gram-positive bacteria belonging to the genus Saprospira (Shi et al., 2006), which grows epibiotically using cyanobacteria as prey. A further two are the bacteria Daptobacter (Guerrero et al., 1986), which has been shown to grow within the cytoplasm of its prey (instead of the periplasm as in the case of Bdellovibrio), and Vampirococcus (Guerrero et al., 1986), an anaerobe which consumes its prey by attaching itself to the surface of its prey and epibiotically consuming the cytoplasmic contents of its prey. Phylogenetic analysis of Varnpirococcus and Daptobacter has yet to be undertaken (as they have not yet been grown in culture); thus, whether they share a common ancestor with the Bdellovibrionales is as yet unknown. However, the possibility of a common predatory ancestor of modern predatory Deltaproteobacteria is lent some weight by the recent characterization of Bdellovibrio strains JSS and KL8. Isolated on Caulobacter crescentus (Shemesh et al., 2003), JSS is phylogenetically placed within the family Bdellovibrio (Davidov and Jurkevitch, 2004), yet both isolates exhibited extremely unusual predatory behavior for Bdellovibrio. They appear to grow epibiotically, in a manner similar to that of Vampirococcus, suggesting that Vampirococcus may in fact be related to the Bdellovibrionales. Interestingly, Daptobacter is reported to be facultatively-and Vampirococcus obligately-anaerobic; yet anaerobic Bdellovibrio organisms have yet to be isolated, although they do survive microaerophilic conditions. As is the case for myxobacteria, among which Anaeromyxobacter has only been recently isolated and studied, it may be that a greater understanding of Bdellovibrio ecology will lead to the isolation of anaerobes (Sanford et al., 2002). There have also been reports of predatory bacteria growing within bacteria and mitochondria within eukaryotic cells. An as-yet-unidentified predatory bacterium has been seen within both the periplasm and the cytoplasm of Thiothrix prey, a gammaproteobacterial symbiont of the mayfly larva (Larkin et al., 1990). The alphaproteobacterium IricES1 is a predatory symbiont of Ixodes ticks, invading and growing within the mitochondria of oocytes (Sacchi et al., 2004), seemingly as Bdellovibrio grows within its bacterial prey. The IricES1 bacteria appear to be passed on from one tick generation to the next, suggesting a long-standing symbiosis/ parasitism within such degenerate gram-negative “bacteria” as mitochondria (Gray et al., 2001). Further isolation and purification of such diverse predatory bacteria for prey-free 16s rRNA gene sequencing
ANALOGOUS SYSTEMS will support or disprove the idea that the ability of Bdellovibrio to grow within another bacterial host is not unique to them and may be found in other Proteobacteria. It is clear that the nonobligately symbiotic or parasitic, predatory bacteria like Bdellovibrio and Myxobacteria have large genomes akin to those of heterotrophs; thus, determining whether transfer of predatory gene islands is responsible for apparently quite diverse bacteria adapting to fit certain predatory niches is not trivial and can ultimately be answered only by full comparative analysis of multiple predatory genomes, although there is little evidence of recent horizontal gene transfer in the B. bacteriovorus HDlOO genome (Gophna et al., 2006).
Ecology of the Bdellovibrionales B. bacteriovorus HD1OOT and Peredibacter starrii A3. 12T were among the first set of isolates of the Bdellovibrionales recorded, both of which were isolated from soil in 1963 by Stolp and Starr (Stolp and Starr, 1963). As Fig. 3 shows, Bdellovibrionales have been isolated from numerous terrestrial and aquatic habitats around the world and in addition 16s rRNA sequences that fall within the two clades have been found in various metagenomic studies. As Bdellovibrio isolates, unlike myxobacteria, must replicate within prey, the true ecological description of isolated Bdellovibrio organisms must include a consideration of prey strains. B. bacteriovorus HDIOOT was first isolated on Erwinia amylovora, a gammaproteobacterium; Peredibacter starrii A3.12T was first isolated on Pseudomonas fluorescens, again a gammaproteobacterium; and both B . marinus SJ’ and B. litoralis JSST were originally isolated on Vibrio parahaemolyticus, another gammaproteobacterium. The host used for the isolation of strain Uk was Escherichia coli Blr; this strain was then turned HI, giving rise to B. stolpii Uki2T. However, Bdellovibrionales are not restricted to a host range consisting of just Gammaproteobacteria: they have been isolated on, and shown to prey on, a variety of gram-negative hosts; for example, strain JSS was isolated on Caulobacter crescentus (Shemesh et al., 2003), an alphaproteobacterium that exhibits another unusual life cycle (Quardokus and Brun, 2003). B. bacteriovorus strain W was isolated from sewage in Germany on Rhodospirillum rubrum, a purple nonsulfur alphaproteobacterium, inside which it has the unusual ability to form resting structures known as bdellocysts (Burger et al., 1968). A recent study has shown that Bdellovibrionales can be isolated from the gut flora of humans, horses, and chickens by using the enterobacterium Proteus mirabilis as prey, and can be isolated from sewage by using Citrobacter freundii as prey (Schwudke et al., 2001). The five
20. PREDATORYLIFESTYLEOF BDELLOVIBRIO isolates from gut flora all belong to the Bdellovibrionaceae, whereas the sewage isolate belongs to the genus Peredibacter. As myxobacteria have been isolated from soil and cow dung, it is clear that they may encounter Bdellovibrio in their natural environment. This is further supported by studies (Ravenschlag et al., 1999) in which researchers made 16s rRNA clone libraries from marine sediments and were able to find both Bdellovibrio- and Myxococcus-related sequences therein. Observations from our laboratory (R. Till and R. E. Sockett, unpublished data) show that for the type strains at least, B. bacteriovorus HDlOO isolates are unable to prey upon Myxococcus xanthus DK1622, either on overlay plaque assay plates or in liquid lysates.
MOTILITY SYSTEMS IN BDELLOUBRIO COMPARED TO MYXOCOCCUS The recent sequencing of both the B. bacteriovorus HDlOO (Rendulic et al., 2004) and M. xanthus DK1622 genomes by The Institute for Genomic Research (TIGR) has enabled proper comparisons of the motility systems in these Deltaproteobacteria for the first time. The main difference between the two bacteria is in the use of flagellar motility: Myxococcus is a nonflagellate topsoil bacterium (Henrichson, 1972), whereas Bdellovibrio uses a single polar flagellum for motility through liquid environments (Lambert et al., 2006). Both species, howevel; share conserved type IV pilus (TFP) systems which are used for S-motility in Myxococcus and which are involved in the prey cell entry characteristic of the predatory Bdellovibrio (Rendulic et al., 2004; Evans et al., 2007). As mentioned earlier (Fig. 1 and 2) Bdellovibrio enters its gram-negative prey after attachment and generation of a temporary “pore” in the prey cell outer membrane. Electron microscopic studies show the predator squeezing through a pore smaller than the size of the cell (Burnham et al., 1968), which would need considerable force against the pressure of the prey periplasm. Once in the periplasm, the Bdellovibrio establishes itself and begins modifying and degrading the prey to replicate itself (Fig. 2). Bdellovibrio attachment to prey cells when entering is extremely stable and cannot be disrupted by vortexing or brief sonication (Burnham et al., 1968). TFP have been shown in other species to generate large forces through PilT-mediated retraction of pili, up to 100 pN in Neisseria gonorrhoeae (Maier et al., 2002), which indicates that the use of TFP in prey entry could generate such force as needed for Bdellovibrio entry in prey. Pili have been shown in other species, such as Myxococcus, Neisseria, and Pseudomonas, to have diverse roles, from S- and twitching motility to biofilm formation and
355 pathogenesis (reviewed in Mattick, 2002), so the suggested use of TFP for prey entry in Bdellovibrio is reasonable. Pilus-like fibers were first observed on the nonflagellate pole of Bdellovibrio by Shilo (1969)and Abram and Davis (1970) using transmission electron microscopy. Pili are difficult to visualize in Bdellovibrio, with some 70% of attack phase cells being nonpiliated (Evans et al., 2007) and those with pili having only three or four pili extending outside the cell. Bdellovibrio HDlOO strains swim at extremely high velocity, recorded at up to 160 pm s-l (Lambert et al., 2006). For an organelle such as a TFP, which is strong under tension rather than not, damage would very likely occur if the pili were outside the cell during periods of such high-speed flagellum-mediated motility. We postulate that Bdellovibrio would extrude and use pili only when having come to rest and being in contact with prey after using high-speed flagellar motility to arrive. This is in contrast to Myxococcus strains which use pili for low-speed (up to 20 pm min-’ on low concentration agar) S-motility constitutively (Shi and Zusman, 1993), although Bdellovibrio may use pili in an analogous way when preying on biofilms. Homology searches using reciprocal NCBI BLASTS (Altschul et al., 1997) and ClustalW protein alignments (Higgins et al., 1994) show that Bdellovibrio has conserved homologues of Myxococcus genes encoding TFP (Table 1).Additional pi1 genes found in other piliated species such as Pseudomonas were not found in Bdellovibrio. Both deltaproteobacteria have the same pi1 gene order as is seen within the Myxococcus operon (Fig. 4), but Bdellovibrio seems to have undergone genomic rearrangements which have scattered parts of the operon to different areas of the genome. Despite this genome fragmentation, the Bdellovibrio pi1genes are expressed (Evans et al., 2007). Gliding motility genes are implicated in myxococcal motility, but these systems seem to be not present in Bdellovibrio, or they were made redundant so long ago that the genes involved are no longer recognizable as such. Homology searches found no significant homologues of Myxococcus genes involved in gliding (Youdeian et al., 2003). The most obvious example of such a gene is the mglA gene, the protein product of which in Myxococcus is absolutely required for both A- and S-motility (Hartzell and Kaiser, 1991a) and is present in Bdellovibrio (Bd3734), but its counterparts in Myxococcus are required only for gliding, such as mglB, cglB, and aglZ (Hartzell and Kaiser, 1991b; Rodriguez and Spormann, 1999; Yang et al., 2004), the homologues of which are not found within the Bdellovibrio genome.
ANALOGOUS SYSTEMS
356
Table 1 Pilus genes found in Myxococcus compared to the Bdellovibrio genome, using reciprocal homology searches and protein level alignment to determine appropriate homologues Gene
Function of gene product
pilB pilT
ATPase, pilus extrusion ATPase, retraction
pilC
Polytopic membrane protein required for pilus biogenesis Negative regulator pilA expression, 2-component system with pilR Regulates pilA expression, 2-component system with Pils Pilus fiber protein Part of ABC transporter required for pilus biogenesis with PilHI Part of ABC transporter required for pilus biogenesis with PilGI Part of ABC transporter required for pilus biogenesis with PilGH Prepilin pedptidase Homologues PilSR in Myxococcus Required for pilus biogenesis, FtsA/MreB homology Involved in pilus biogenesis
pilS pilR pilA pilG pilH pilI pilD pilS2/R2 pilM pilN pi10 pilP pilQ
td
Involved in pilus biogenesis Lipoprotein needed to stabilize PilQ multimer formation Outer membrane multimer complex needed for PilA fiber extrusion Lipoprotein with TPR protein interaction domains that is required for pilQ multimer formation
Bdellovibrio homologue
Bdellovibrio gene no.
Reference(s)
pilB pilT ( 2 are annotated, this best homologue) pilC
Bdl509 Bd3852
Turner et al., 1993 Wu et al., 1997
Bd1511
pilS
Bd1512
Nunn et al., 1990; Mattick, 2002 Wu and Kaiser, 1997
pilR
Bdl513
Wu and Kaiser, 1997
pilA Bd1291
Bd1290 Bd1291
Wu and Kaiser, 1995 Wu et al., 1998
Annotated phnC, upstream of pill pilI
Bd0860
Wu et al., 1998
Bd0861
Wu et al., 1998
pilD No definite homologues pilM (best homologue); pilM pilN
Bd0862 NIA Bd0863; Bd1585
Nunn et al., 1990 Jelsbak and Kaiser, 2005 Martin et al., 1995
Bd0864
pi10 pilP
Bd0865 Bd0866
pilQ (most significant homologue of 3 pilQs annotated) Most likely candidate is annotated pilF in genome
Bd0867
Martin et al., 1995; Mattick 2002 Mattick, 2002 Drake et al., 1997; Nudleman et al., 2006 Wall et al., 1999
Neither does Bdellovibrio appear to have the frz gene system that is used in Myxococcus for switching the pole at which pili are extruded during social motility (reviewed in Ward and Zusman, 1999) (Fig. 5 ) . The pole-switching Myxococcus frzS gene (Mignot et al., 2005) has no significant homologues in the Bdellovibvio genome, and the other frz genes show only partial homology to general monocyte chemoattractant protein or 2-component sensor-regulators, likely required for flagellar motility or unrelated processes. Biologically, this makes sense because Myxococcus uses the pilus pole switching for reversal of the direction of cell motility as a nonflagellate soil surface bacterium (Fig. 5B). Bdellovibrio, on the other hand, uses flagellummediated motility through liquid environments and has a single pole dedicated to housing the flagellum (Fig. 5A). It follows, therefore, that there would never be a requirement for FrzS-mediated switching of the
Bd3829
Rodriguez-Soto and Kaiser, 1997a, 1997b
piliated pole in Bdellovibrio as in this bacterium TFP are at the nonflagellate pole where prey cell entry occurs (Fig. 5A). Taken together, the conclusions are that there is a conserved cell movement pathway between the two bacteria in the form of TFP, where the genes may be ancestral to both. Recent work has determined that such TFPmediated cell movement in Bdellovibrio is at the prey penetration step. After the speciation and divergence of these predators, Myxococcus retained gliding genes and developed slow surface motility behaviors, feeding socially and cooperatively on prey, in rich soil/dung environments, by external enzymic hydrolysis. Bdellovibrio retained and/or acquired the genes necessary for flagellar motility and forsook slow glidingkwitching motility for rapid, lone, and adventurous seeking of distant prey across more dilute areas of water within soil, sediment, and aquatic environments.
20. PREDATORYLIFESTYLEOF BDELLOVIBRIO
35 7
A
B
A
pilD
pilM
I
pilN
I
pi10
I
pilP
I
pilQ
Figure 4 Comparison of the operon structure of pi1 genes in Myxococcus and Bdellovibrio. (A) Organization of Myxococcus pi1 genes, taken from Wall and Kaiser, 1999. (B) pi1 genes are scattered around the Bdellovibrio genome, but a Myxococcus-like, possibly ancestral, organization can be seen with the gene order being conserved. The starred annotated pilT within the operon is a good homologue, but Bd3852 on the left is a better homologue, suggesting a duplication event. Bdellovibrio does not have significant homologues of pilR2/S2.
HYDROLYTIC ENZYMES OF BDELLOVIBNO AND MYXOCOCCUS As mentioned at the beginning of the chapter, Bdellovibrio and Myxococcus are microorganisms which prey upon and lyse other microorganisms, but the specificity and mechanisms by which they do this differ. Bdellovibrio preys upon only gram-negative bacteria, albeit a wide range of these (Martin, 2002), as it penetrates the outer layers and enters the periplasmic space of its prey. This lifestyle requires an initially directed and tempered hydrolytic assault as first only a small pore is formed in the prey outer membrane and cell wall and resealed after prey entry, but during the following few hours the prey is rapidly degraded, and the contents of the cytoplasm and inner membrane are then broken down in a controlled manner, followed by outer layer components as the prey “ghost” is lysed to release progeny Bdellovibrio (Engelking and Seidler, 1974; Shilo and Bruff, 1965; Tudor et al., 1990). In contrast, Myxococcus can prey upon a much wider range of microorganisms including gram-negative and gram-positive bacteria and even fungi (Rosenberg and Varon, 1984). They do this by surrounding the prey cells in a swarm and releasing hydrolytic enzymes to kill and digest them, starting with the outer layers and then digesting the cell contents after prey cell lysis. These different prey ranges and killing strategies require many similar hydrolytic enzymes, but also many different ones. The ongoing study of how these hydrolytic enzymes allow the bacterial predators to kill and degrade their prey is not only intrinsically interesting but also potentially useful as they may ultimately be developed as novel antibiotics.
The first striking similarity between the two bacteria’s predatory arsenals is the very large number of genes predicted to be coding for hydrolytic enzymes, with the largest group being proteases. Indeed, Bdellovibrio has the second-highest density of genes encoding these hydrolytic enzymes found in all bacterial genomes sequenced so far, after the minimalist, specialist genome of the symbiont Buchnera aphidicola. The recently completed Myxococcus genome (TIGR) seems to have almost as high a proportion as Bdellovibrio, despite its much larger genome size.
Modes of Delivery of Hydrolytic Enzymes Toxic proteins are often delivered across bacterial membranes and to their targets by Type I11 or Type IV secretion, which involves the building of protein complexes specifically for this task. However, neither Bdellovibrio nor Myxococcus has a complete set of genes for these transport systems, except those dedicated to assembling flagella in Bdellovibrio and TFP pili in both. Both bacteria, as expected, have full sets of genes for the general secretion pathway (Sec-dependent),and many hydrolytic enzymes of all types from both organisms are predicted to have signal sequences for secretion across the inner membrane by this pathway. Both also have a full set of tat genes (for twin-arginine transport) and a subset of gene products from both genomes have the characteristic signal sequences with the twin-arginine motif, suggesting that they are transported across the predator inner membrane by this system. Interestingly, however, while Myxococcus has some 16 proteases with the TAT signal peptide (as predicted by the
ANALOGOUS SYSTEMS
358
A
Figure 5 Comparison (not to scale) of motility systems in Bdellovibrio (A) and myxobacterial (B) cells. The cytoplasm is shown in white and the periplasm in gray in each. EPS, extracellular polysaccharide.
tatP program [http://www.cbs.dtu.dk/services/TatP-1.O/]), Bdello~ibriohas no hydrolytic enzymes with the motif. A large number of Bdellovibrio hydrolytic enzymes must be either transported across the inner membrane by the Sec pathway or possibly by as-yet-uncharacterized methods. There is less indication of how, or if, they cross the outer membrane, as it is not yet established whether Bdellovibrio retains an intact outer membrane throughout the predatory cycle. In addition these enzymes may also have to cross an intact prey cytoplasmic membrane. A number of hydrolytic enzymes may be passenger domains of autotransporters and thus use Type V secretion (Henderson et al., 2004). It is possible that completely novel transport systems exist in either organism for the specific task of delivering hydrolytic enzymes across the outer layers to their targets. As it is not yet known how many hydrolytic enzymes are exported out of the cell, it is likely that many of those predicted to be periplasmic (Table 2) (Odelson et al., 1982) or of unknown location may actually be extracellular. A clearer picture of the organization and methods of export awaits further transcript profiling and proteomic analysis.
Proteases By far the largest group of hydrolases in both bacteria is proteases; this probably reflects the importance to predatory bacteria in breaking down prey proteins for uptake and consumption by the predator. Proteins not only provide a source of carbon, hydrogen, and oxygen but also provide nitrogen, which is essential t o the synthesis of nucleic acids and nucleotide cofactors as well as proteins. In the case of Bdellovibrio, as explained in “Nucleases” below, there is a significant need to synthesize nucleotides from prey raw materials during replication. Table 2 shows the types of proteases and their predicted location by using the PSORTb program (http://www.psort.org/psortb/). The two genomes appear to have a similar complement of proteases in that the majority are serine- and metal-catalytic types. (Rao et al., 1998). Not discussed are the large array of L-amino-peptidases, which must have significant roles in prey degradation. Bdellovibrio and myxobacteria have a full complement of proteases involved in both housekeeping and shock responses, including genes such as lon and clp
20. PREDATORY LIFESTYLE OF BDELLOVIBRIO Table 2
359
Predicted location and classes of Mvxococcus and Bdellovibrio Droteasesa
~
External
Outer membrane
Periplasm
Myxo metal dependent Bdello metal dependent
12 1
1 0
2 2
Myxo serine catalytic type Bdello serine catalytic type
8 11
1 0
Myxo cysteine catalytic type Bdello cysteine catalytic type
0 0
Myxo unclassified catalytic type Bdello unclassified catalytic type
1 0
Class
Inner membrane
Cytoplasm
Unknown
8 7
9 5
38 38
9 5
0 7
0 4
18 47
0 0
0 1
1 2
0 1
2 4
0 0
2 2
1 1 (plus 1 aspartic type)
11 0
20 11
aValues represent the numbers of proteases
(Gottesman, 2003). These may also have been exploited, amplified, and diversified for predatory roles.
Peptidoglycan Hydrolysis There are a number of putative carboxypeptidases which are predicted to disrupt bonds between D-amino acids, and some of these in both bacteria will be involved in remodeling of the bacterium’s own peptidoglycan during growth and development. However, some of these may be involved in the de-cross-linking and breakdown of prey peptidoglycan and hence prey lysis. Here there is a difference between the requirements of the two predators. Bdellovibrio initially forms a small pore in the prey peptidoglycan which it then reseals, modifying the prey cell wall further to form a rounded bdelloplast structure; only later does it degrade the remainder from within. MYXOCOCCUS, on the other hand, attacks the prey peptidoglycan from without, having a wider variety of substrate peptidoglycans to act upon as it also attacks gram-positive cells. Myxococcus also undergoes autolysis as a developmental step during fruiting body development (Wireman and Dworkin, 1977), also requiring specific peptidoglycan hydrolytic enzymes. These differences may be reflected in peptidase diversity, although few obvious trends of difference can be distinguished by sequence prediction alone. One group of hydrolases that may be involved in these differing manipulations of peptidoglycan are the murein lytic transglycosylases (MLTs). These are involved in breaking the glycan bonds to make holes in peptidoglycan layers and as such may well function in the predatory life described above. In other bacteria they function to form local pores in which to insert machinery such as flagella and Type 111 or Type IV secretion apparatuses (Zahrl et al., 2005), and some could possibly be used to construct novel secretion apparatuses or junctions between predator and prey membranes in Bdellovibrio. There are
eight predicted MLTs and three other glycanases in the Bdellovibrio genome and five predicted MLTs in Myxococcus. This is more than in most bacteria (e.g., E. coli has one), strongly suggesting that these are used in the predatory lifestyle. The fact that Bdellovibrio has more than Myxococcus could also be because of the many different staged modifications of the peptidoglycan that Bdellovibrio requires as it establishes and grows within the prey periplasm, breaking the cell wall completely only later to lyse the prey. BLAST analysis shows that the Bdellovibrio MLTs tend to have homology to MLTs from other gram-negative bacteria, which is to be expected, as they are to act upon the peptidoglycan of gram-negative organisms. Those of Myxococcus, however, have homology to MLTs from eukaryotes and a range of diverse bacteria, and this probably reflects their varied host range as they could act upon a range of cell walls.
Nucleases Biochemical studies have shown that Bdellovibrio breaks down prey DNA and RNA in a controlled manner (Hespel1 et al., 1975), and the genome reveals a number of genes with homology to endonucleases which are predicted to be exported. Bdellovibrio typically produces three to six progeny from one E. coli prey cell, and this presents it with the problem of needing to produce three to six genomes, each of which is around the same size as that of its prey. Hence, it is probably very important for Bdellovibrio to salvage as many nucleotides as possible from its prey and then to synthesize the remainder required. Little is known of the dynamics of Myxococcus breaking down prey nucleotides, but a similar situation may exist, as it has one of the largest bacterial genomes and so may also require significant amounts of synthesis. However, there are very few genes with homology to known nucleases and fewer still that are predicted to be
ANALOGOUS SYSTEMS
360 exported. There are a number of genes annotated as having some similarity to nucleases/hydrolases, the export of which cannot be accurately predicted by sequence alone, and it is possible that these are involved in the breakdown of prey nucleotides. In addition to these nucleases that are probably involved in the predatory lifestyle of both bacteria, they also have a set of nucleases for the recycling of the cell’s own nucleotides and for defense against phages.
Other Hydrolases Myxococcus has four genes predicted to be chitinases, with class I and class I1 chitinases represented, and these are presumably involved in the breakdown of fungal cell walls, as these are also prey for Myxococcus. Bdellovibrio lacks any chitinases presumably because it does not prey upon fungi. Both organisms have approximately 15 genes predicted to encode lipases of various types. In addition to the remodeling of their own lipid layers and recycling of cellular lipids, some of these are likely to be used to degrade prey lipids. A possible alternative uptake of lipids may involve partial fusion of lipid membranes between prey and predator, particularly with Bdellovibrio because there is contact between membranes. Both genomes also have a large number of other hydrolases including those which may degrade polysaccharides and others of unknown specificity. Both Bdellovibrio and Myxococcus have life cycle stages in which they are not in contact with prey cells; thus, they must transcriptionally coordinate a hydrolytic assault upon detection of their prey. As described above, there are various stages of prey hydrolysis by Bdellovibrio and we are working to determine the expression patterns of hydrolytic genes at each stage.
CONCLUSION Bdellovibrio and Myxococcus are both bacteria with a very large complement of hydrolytic enzymes which they use to kill and digest other bacteria. Due to the internal predatory phase of Bdellovibrio, it has remained somewhat of a “Cinderella” in bacterial research, while mutant studies have illuminated much of the regulatory controls of the growth and development of myxobacteria, as a book of this magnitude implies. However, with the benefit of genome sequences and reverse genetic, transcriptional, and proteomic approaches, Bdellovibrio too is starting to be understood. It is clear that, like the myxobacteria, it uses TFP and it may employ novel secretion systems and also use the TAT, autotransporter, and Sec systems in predation to help export its arsenal
of hydrolytic enzymes. Studying how these bacteria coordinate their hydrolytic attack on prey is fascinating deltaproteobacterial biology in its own right, and it might also highlight candidates for use as antibiotics as many prey organisms are pathogens of humans, animals, and plants. Research in our laboratory is funded by grants received from The Wellcome Trust, The British Society for Antimicrobial Chemotherapy, and The Human Frontier Science Program and by studentships from N E R C and BBSRC. We thank Rob Till, Michael Capeness, Colin Breakwell, and Marilyn Whitworth for technical assistance.
References Abram, D., and B. K. Davis. 1970. Structural properties and features of parasitic Bdellovibrio bacteriovorus. J. Bacteriol. 104~948-965. Altschul, S. F., T. L. Madden, A. A. Schaffer, J. Zhang, Z. Zhang, W. Miller, and D. J. Lipman. 1997. Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res. 25:3389-3402. Baer, M. L., J. Ravel, S. A. Pineiro, D. Guether-Borg, and H. N. Williams. 2004. Reclassification of salt-water Bdellovibrio sp. as Bacteriovorax marinus sp. nov. and Bacteriovorax litoralis sp. nov. Int. J. Syst. Evol. Microbiol. 54: 1011-1016. Baer, M. L., J. Ravel, J. Chun, R. T. Hill, and H. N. Williams. 2000. A proposal for the reclassification of Bdellovibrio stolpii and Bdellovibrio starrii into a new genus, Bacteriovorax gen. nov., as Bacteriovorax stolpii comb. nov. and Bacteriovorax starrii comb. nov., respectively. lnt. J. Syst. Evol. Microbiol. 50:219-224. Burger, A., G. Drews, and R. Ladwig. 1968. Host range and infection cycle of a newly isolated strain of Bdellovibrio bacteriovorus. Arch. Mikrobiol. 61:261-279. (In German.) Burnham, J. C., T. Hashimoto, and S. F. Conti. 1968. Electron microscopic observations on the penetration of Bdellovibrio bacteriovorus into gram-negative bacterial hosts. J. Bacteriol. 96:1366-138 1. Davidov, Y., and E. Jurkevitch. 2004. Diversity and evolution of Bdellovibrio-and-like organisms (BALOs), reclassification of Bacteriovorax starrii as Peredibacter starrii gen. nov., comb. nov., and description of the Bacteriovorax-Peredibacter clade as Bacteriovoracaceae fam. nov. lnt. J. Syst. Evol. Microbiol. 54:1439-1452. Davidov, Y., D. Huchon, S. F. Koval, and E. Jurkevitch. 2006. A new alpha-proteobacterial clade of Bdellovibrio-like predators: implications for the mitochondria1 endosymbiotic theory. Environ. Microbiol. 8:2179-2188. Drake, S. L., S. A. Sandstedt, and M. Koomey. 1997. PilP, a pilus biogenesis lipoprotein in Neisseria gonorrhoeae, affects expression of PilQ as a high-molecular-mass mulitmer. Mol. Microbiol. 23657. Engelking, H. M., and R. J. Seidler. 1974. The involvement of extracellular enzymes in the metabolism of Bdellovibrio. Arch. Mikrobiol. 95:293-304.
20. PREDATORY LIFESTYLEOF BDELLOVIBRIO Evans, K. J., C. Lambert, and R. E. Sockett. 2007. Predation by Bdellovibrio bacteriovorus HDlOO requires type IV pili. J. Bacteriol. 189:4850-4859. Gophna, U., R. L. Charlebois, and W. F. Doolittle. 2006. Ancient lateral gene transfer in the evolution of Bdellovibrio bacteriovorus. Trends Microbiol. 14:64-69. Gottesman, S. 2003. Proteolysis in bacterial regulatory circuits. Annu. Rev. Cell Dev. Biol. 19565-587. Gray, M. W., G. Burger, and B. F. Lang. 2001. The origin and early evolution of mitochondria. Genome Biol. 2: reviews101 8.1-101 8.5 Guerrero, R., C. Pedros-Alio, I. Esteve, J. Mas, D. Chase, and L. Margulis. 1986. Predatory prokaryotes: predation and primary consumption evolved in bacteria. Proc. Natl. Acad. Sci. USA 83:2138-2142. Hartzell, P., and D. Kaiser. 1991a. Function of MglA, a 22kilodalton protein essential for gliding in Myxococcus xanthus. J. Bacteriol. 173:7615-7624. Hartzell, I?, and D. Kaiser. 1991b. Upstream gene of the mgl operon controls the level of the MglA protein in Myxococcus xanthus. J. Bacteriol. 173:7625-7635. Henderson, I. R., F. Navarro-Garcia, M. Desvaux, R. C. Fernandez, and D. Ala'Aldeen. 2004. Type V protein secretion pathway: the autotransporter story. Microbiol. Mol. Biol. Rev. 68:692-744. Henrichson, J. 1972. Bacterial surface translocation: a survey and a classification. Bacteriol. Rev. 36:478-503. Hespell, R. B., G. F. Miozzari, and S. C. Rittenberg. 1975. Ribonucleic acid destruction and synthesis during intraperiplasmic growth of Bdellovibrio bacteriovorus. J. Bacteriol. 123:48 1-491. Higgins, D., J. Thompson, T. Gibson, J. D. Thompson, D. G. Higgins, and T. J. Gibson. 1994. CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Res. 22:46734680. Jelsbak, L., and D. Kaiser. 2005. Regulating pilin expression reveals a threshold for S motility in Myxococcus xanthus. J. Bacteriol. 1872105-2112. Jurkevitch, E., D. Minz, B. Ramati, and G. Barel. 2000. Prey range characterization, ribotyping, and diversity of soil and rhizosphere Bdellovibrio spp. isolated on phytopathogenic bacteria. Appl. Environ. Microbiol. 66:2365-2371. Lambert, C., K. J. Evans, R. Till, L. Hobley, M. J. Capeness, S. Rendulic, S. C. Schuster, S.-I. Aizawa, and R. E. Sockett. 2006. Characterising the flagella filament and the role of motility in bacterial prey-penetration by Bdellovibrio bacteriovorus. Mol. Microbiol. 60:274-286. Lambert, C., M. C. M. Smith, and R. E. Sockett. 2003. A novel assay to monitor predator-prey interactions for Bdellovibrio bacteriovorus l09J reveals a role for methyl-accepting chemotaxis proteins in predation. Environ. Microbiol. 5:127132. Larkin, J. M., M. C. Henk, and S. D. Burton. 1990. Occurrence of a Thiothrix sp. attached to mayfly larvae and presence of parasitic bacteria in the Thiothrix sp. Appl. Environ. Microbiol. 56:357-361.
361 Maier, B., L. Potter, M. So, H. S. Seifert, and M. P. Sheetz. 2002. Single pilus motor forces exceed 100pN. Proc. Natl. Acad. Sci. USA 99:16012-16017. Martin, M. 0. 2002. Predatory prokaryotes: an emerging research opportunity. J. Mol. Microbiol. Biotechnol. 4:467477. Martin, P. R., A. A. Watson, T. F. McCaul, and J. S. Mattick. 1995. Characterisation of a five-gene cluster required for the biogenesis of type 4 fimbriae in Pseudomonas aeruginosa. Mol. Microbiol. 16:497-508. Mattick, J. S. 2002. Type IV pili and twitching motility. Annu. Rev. Microbiol. 56:289-314. Mignot, T., J. P. Merlie, and D. R. Zusman. 2005. Regulated pole-to-pole oscillations of a bacterial gliding motility protein. Science 310:855-857. Nudleman, E., D. Wall, and D. Kaiser. 2006. Polar assembly of the type IV pilus secretin in Myxococcus xanthus. Mol. Microbiol. 60:16-29. Nunez, M. E., M. 0. Martin, L. K. Duong, E. Ly, and E. M. Spain. 2003. Investigations into the life cycle of the bacterial predator Bdellovibrio bacteriovorus 109J at an interface by atomic force microscopy. Biophys. J. 84:3379-3388. Nunn, D., S. Bergman, and S. Lory. 1990. Products of three accessory genes, pilB, pilC, and pilD, are required for biogenesis of Pseudomonas aeruginosa pili. J. Bacteriol. 172:2911-2919. Odelson, D. A., M. A. Patterson, and R. B. Hespell. 1982. Periplasmic enzymes in Bdellovibrio bacteriovorus and Bdellovibrio stolpii. J. Bacteriol. 151:756-763. Quardokus, E. M., and Y. V. Brun. 2003. Cell cycle timing and developmental checkpoints in Caulobacter crescentus. Curr. Opin. Microbiol. 6541-549. Rao, M. B., A. M. Tanksale, M. S. Ghatge, and V. V. Deshpande. 1998. Molecular and biotechnological aspects of microbial proteases. Microbiol. Mol. Biol. Rev. 62597-635. Ravenschlag, K., K. Sahm, J. Pernthaler, and R. Amann. 1999. High bacterial diversity in permanently cold marine sediments. Appl. Environ. Microbiol. 65:3982-3989. Rendulic, S., P. Jagtap, A. Rosinus, M. Eppinger, C. Baar, C. Lanz, H. Keller, C. Lambert, K. J. Evans, A. Goesmann, F. Meyer, R. E. Sockett, and S. C. Schuster. 2004. A predator unmasked: life cycle of Bdellovibrio bacteriovorus from a genomic perspective. Science 303:689-692. Rodriguez, A. M., and A. M. Spormann. 1999. Genetic and molecular analysis of cglB, a gene essential for single cell gliding in Myxococcus xanthus. J. Bacteriol. 181:4381-4390. Rodriguez-Soto, J. P., and D. Kaiser. 1997a. The tgl gene: social motility and stimulation in Myxococcus xanthus. J. Bacterial. 179:4361-4371. Rodriguez-Soto, J. P., and D. Kaiser. 199713. Identification and localization of the Tgl protein, which is required for Myxococcus xanthus social motility. J. Bacteriol. 179: 4372-4381. Rosenberg, E., and M. Varon. 1984. Antibiotics and lytic enzymes, p. 109-125. In E. Rosenberg (ed.), Myxobacteria: Development and Cell Interactions. Springer, New York, NY. Sacchi, L., E. Bigliardi, S. Corona, T. Beninati, N. Lo, and A. Franceschi. 2004. A symbiont of the tick Ixodes ricinus invades and consumes mitochondria in a mode similar to
362 that of the parasitic bacterium Bdellovibrio bacteriovorus. Tissue Cell 36:43-53. Sanford, R. A., J. R. Cole, and J. M. Tiedje. 2002. Characterization and description of Anaeromyxobacter dehalogenans gen. nov., sp. nov., an aryl-halorespiring facultative anaerobic myxobacterium. Appl. Environ. Microbiol. 68:893-900. Schoeffield, A. J., W. A. Falkler, D. Desai, and H. N. Williams. 1991. Serogrouping of halophilic bdellovibrios from Chesapeake Bay and environs by immunodiffusion and immunoelectrophoresis. Appl. Environ. Microbiol. 57:3470-3475. Schwudke, D., E. Strauch, M. Krueger, and B. Appel. 2001. Taxonomic studies of predatory bdellovibrios based on 16s rRNA analysis, ribotyping and the hit locus and characterization of isolates from the gut of animals. Syst. Appl. Microbiol. 24:385-394. Seidler, R. J., M. Mandel, and J. N. Baptist. 1972. Molecular heterogeneity of the bdellovibrios: evidence of two new species. J. Bacteriol. 109:209-217. Shemesh, Y., Y. Davidov, S. Koval, and E. Jurkevitch. 2003. Small eats big: ecology and diversity of Bdellovibrio and like organisms, and their dynamics in predator-prey interactions. Agronomie 23:433-439. Shi, M., L. Zou, X. Liu, Y. Gao, Z. Zhang, W. Wu, D. Wen, Z. Chen, and C. An. 2006. A novel bacterium Saprospira sp. strain PdY3 forms bundles and lyses cyanobacteria. Front. Biosci. 11:1916-1 923. Shi, W., and D. R. Zusman. 1993. The two motility systems of Myxococcus xanthus show different selective advantages on various surfaces. Proc. Natl. Acad. Sci. USA 90:3378-3382. Shilo, M. 1969. Morphological and physical aspects of the interaction of Bdellovibrio with host bacteria. Curr. Top. Microbiol. and Immunol. 50:174-204. Shilo, M., and B. Bruff. 1965. Lysis of gram-negative bacteria by host-independent ectoparasitic Bdellovibrio bacteriovorus isolates. J. Gen. Microbiol. 40:317-328. Snyder, A. R., H. N. Williams, M. L. Baer, K. E. Walker, and 0. C. Stine. 2002. 16s rDNA sequence analysis of environmental Bdellovibrio-and-like organisms (BALO) reveals extensive diversity. Int. J . Syst. Evol. Microbiol. 52:20892094. Sockett, R. E., and C. Lambert. 2004. Bdellovibrio as therapeutic agents: a predatory renaissance? Nut. Rev. Microbiol. 2~669-675. Stolp, H., and M. P. Starr. 1963. Bdellovibrio bacteriovorus gen. et sp. n., a predatory, ectoparasitic, and bacteriolytic microorganism. Antonie Leeuwenhoek 29:217-248.
ANALOGOUS SYSTEMS Tudor, J. J., M. P. McCann, and I. A. Acrich. 1990. A new model for the penetration of prey cells by bdellovibrios. J. Bacteriol. 172:2421-2426. Turner, L. R., J. C. Lara, D. N. Nunn, and S. Lory. 1993. Mutations in the consensus ATP-binding sites of XcpR and PilB eliminate extracellular protein secretion and pilus biogenesis in Pseudomonas aeruginosa. J. Bacteriol. 175: 4962-4969. Wall, D., P. E. Kolenbrander, and D. Kaiser. 1999. The Myxococcus xanthus pilQ (sglA)gene encodes a secretin homologue required for Type IV pilus biogenesis, social motility, and development. J. Bacteriol. 181:24-33. Wall, D., and D. Kaiser. 1999. Type IV pili and cell motility. Mol. Microbiol. 32:l-10. Ward, M., and D. Zusman. 1999. Motility in M . xanthus and its role in developmental aggregation. Cum Opin. Microbiol. 2:624-629. Wireman, J. W., and M. Dworkin. 1977. Developmentally induced autolysis during fruiting body formation by Myxococcus xanthus. J. Bacteriol. 129:798-802. Wu, S. S., J. Wu, and D. Kaiser. 1997. The Myxococcus xanthus pilT locus is required for social gliding motility although pili are still produced. Mol. Microbiol. 23:109-121. Wu, S. S., and D. Kaiser. 1995. Genetic and functional evidence that Type IV pili are required for social gliding motility in Myxococcus xanthus. Mol. Microbiol. 18547. Wu, S. S., and D. Kaiser. 1997. Regulation of expression of the pilA gene in Myxococcus xanthus. J. Bacteriol. 179:77487758. Wu, S. S., J. Wu, Y. L. Cheng, and D. Kaiser. 1998. The pilH gene encodes an ABC transporter homologue required for type IV pilus biogenesis and social gliding motility in Myxococcus xanthus. Mol. Microbiol. 29:1249. Yang, R., S. Bartle, R. Otto, A. Stassinopoulos, M. Rogers, L. Plamann, and P. Hartzell. 2004. AglZ is a filament-forming coiled-coil protein required for adventurous gliding motility of Myxococcus xanthus. J. Bacteriol. 186:6168-6178. Youderian, P., N. Burke, D. J. White, and P. L. Hartzell. 2003. Identification of genes required for adventurous gliding motility in Myxococcus xanthus using the transposable element mariner. Mol. Microbiol. 49:555-570. Zahrl, D., M. Wagner, K. Bischof, M. Bayer, B. Zavecz, A. Beranek, C. Ruckenstuhl, G. E. Zarfel, and G . Koraimann. 2005. Peptidoglycan degradation by specialized lytic transglycosylases associated with type I11 and type IV secretion systems. Microbiology 151:3455-3467.
Myxobacteria: Multicellularity and Differentiation Edited by David E. Whitworth 0 2008 ASM Press, Washington, D.C.
Lee Kroos Patrick J. Piggot Charles P. Moran, Jr.
21
Bacillus su btilis Sporulation and Other Multicellular Behaviors
In some ways, Bacillus subtilis is very different from the myxobacteria. It is classified in a different phylum, Firmicutes, because it stains positively in the Gram test. This reflects its thick cell wall. In contrast, gram-negative bacteria like the myxobacteria have a thinner cell wall and an extra cell envelope layer, the outer membrane. B. subtilis has a much smaller genome than Myxococcus xanthus (4.2 versus 9.1 Mb) with a much lower G+C content (43.5 versus 68.9%). Another major difference is that B. subtilis can swim, powered by peritrichous flagella, while myxobacteria lack flagella and are limited to gliding on solid surfaces. Metabolically, B. subtilis is equipped to transport and utilize a wide variety of carbohydrates, whereas M. xanthus lacks this ability and is suited to an Atkins-like diet of proteins and lipids. Perhaps these differences allow B. subtilis and M . xanthus to exploit distinct niches in their common soil environment, though at times they likely are competitors. B. subtilis is the best-studied gram-positive bacterium. Two books have described various aspects of its biology, one in the pregenome era (Sonenshein et al., 1993) and one postgenomically (Sonenshein et al., 2002). Here,
we focus on B. subtilis multicellularity, emphasizing the two-cell differentiation process of endospore formation and attempting to note similarities to M. xanthus.
MULTICELLULAR BEHAVIORS Populations of B. subtilis can self-organize into macrofibers, complex colonies, and bioconvection patterns (reviewed in Mendelson, 1999).Macrofibers form when cells divide but fail to separate, forming chains that twist together. Complex colony morphologies ranging from concentric rings to branched patterns can be produced by placing the same strain on solid surfaces differing in agar and nutrient concentration. The patterns produced result from the interplay between cell growth, motility, chemotaxis, nutrient availability, and moisture content of the agar surface. B. subtilis in static liquid culture swims up the oxygen concentration gradient toward the fluidair interface and then sinks in plumes of a bioconvection pattern set in motion by gravity, which carries oxygen to cells below. While cell growth, division, motility, and chemotaxis clearly play roles in forming bioconvection patterns, complex colonies, and macrofibers, these ~~
~~
~
Lee Kroos, Department of Biochemistry and Molecular Biology, Michigan State University, East Lansing, MI 48824. Patrick J. Piggot, Department of Microbiology and Immunology, Temple University School of Medicine, Philadelphia, PA 19140. Charles P. Moran, Jr., Department of Microbiology and Immunology, Emory University School of Medicine, Atlanta, Georgia 30322.
363
364 multicellular phenomena have not yet been subjected to systematic genetic analysis. In contrast, recently discovered multicellular behaviors of biofilm formation (Branda et al., 2001; Hamon and Lazazzera, 2001) and swarming motility (Dixitet al., 2002; Kearns and Losick, 2003) are rapidly being elucidated by genetic and genomic approaches (Branda et al., 2004, 2006; Calvio et al., 2005; Chapeau and Saier, 2004; Chu et al., 2006; Connelly et al., 2004; Hamon et al., 2004; Kearns et al., 2004, 2005; Senesi et al., 2004; Stanley et al., 2003). Natural isolates of B. subtilis form biofilms with aerial structures that are preferential sites for sporulation (Branda et al., 2001), in some ways resembling M. xanthus fruiting bodies. Under other conditions, natural isolates exhibit swarming motility (Kearns and Losick, 2003). Cells become hyperflagellated and move cooperatively in groups, analogous to M. xanthus social motility, though powered differently. Surfactant production plays an important role in B. subtilis swarming motility and biofilm formation (Branda et al., 2001; Kearns and Losick, 2003). While there is evidence that M . xanthus produces surfactant (Dworkin et al., 1983), its role in motility and fruiting body formation is unclear. On the other hand, extracellular matrix material clearly plays a critical role in M . xanthus swarming and development (see chapter 6), it is implicated in B. subtilis biofilm formation (Branda et al., 2001,2004; Kearns et al., 2005), and its role in B. subtilis swarming motility remains to be explored. The most studied and best understood multicellular behaviors of B. subtilis are the development of genetic competence (the ability to take up exogenous DNA) and sporulation. Both of these behaviors are regulated by quorum sensing mediated by peptide signals (reviewedin Lazazzera and Grossman, 1998; Perego and Brannigan, 2001; Piggot and Hilbert, 2004; and Tortosa and Dubnau, 1999). The ComX pheromone is a 10-amino-acid peptide that is predicted to be farnesylated on a tryptophan residue in strain 168, and in strain RO-E-2, ComX has been shown to be a 6-amino-acid peptide with a geranylated tryptophan residue (Okada et al., 2005). It signals competence development by interacting with ComP, a membrane-bound histidine protein kinase. This has parallels with M. xanthus A-signaling, which measures cell density early in development by monitoring the extracellular concentration of amino acids and peptides via the Sass histidine protein kinase (see chapter 3). Other peptides are secreted by B. subtilis and signal cell density by being taken up through oligopeptide permease and then interacting with proteins that regulate competence and sporulation pathways (reviewed in Lazazzera and Grossman, 1998; Perego and Brannigan, 2001;
ANALOGOUS SYSTEMS Piggot and Hilbert, 2004; and Tortosa and Dubnau, 1999). Whether M. xanthus uses similar mechanisms remains to be seen. Recently, a putative antibiotic-like peptide and a small (63-amino-acid)protein have been described, that appear to be secreted from B. subtilis having entered the sporulation pathway, and kill nonsporulating cells in the population (Gonzalez-Pastor et al., 2003). This cannibalistic behavior is proposed to delay commitment to sporulation as the secreting cells feed on nutrients released from siblings. Such a delay would seem advantageous under conditions of transient nutrient deprivation. Cells producing the putative antibiotic-like peptide appear to resist killing by also making an export pump. They resist the small protein toxin by synthesizing an immunity protein (Ellermeier et al., 2006). A role for autolysis in M. xanthus development has been proposed (Wireman and Dworkin, 1975), but more recent experiments challenge the idea (O’Connor and Zusman, 1988). Whether a given B. subtilis cell in a population is a cannibal or a victim appears to be determined by its level of phosphorylated SpoOA (Gonzalez-Pastor et al., 2003). Likewise, initiation of sporulation appears to be governed by a phosphorelay leading to formation of SpoOA-P (Burbulys et al., 1991), as well as by additional regulatory mechanisms (see below) that constitute a bistable switch, so that some cells activate SpoOA and others do not (Chung et al., 1994; Veening et al., 2005). The master regulator of competence, ComK, also exhibits bistability, which requires positive autoregulation (Maamar and Dubnau, 2005; Smits et al., 2005). It would not be surprising if a bistable switch governs which M. xanthus cells enter fruiting bodies and which remain outside as peripheral rods (O’Connor and Zusman, 1991).
INITIATION OF SPORULATION In addition to extracellular peptide signaling of cell density, mentioned above, intracellular signals including nutrient limitation, the metabolic state of the cell with respect to glucose utilization and tricarboxylic acid cycle activity, and the status of the chromosome with respect to damage, replication, and segregation all feed into a regulatory network that controls the transitions from growth to stationary phase to cannibalism and, as a last resort, to sporulation (reviewed in Burkholder and Grossman, 2000; Piggot and Hilbert, 2004; and Sonenshein, 2000). Recent work on key components of the network, such as the transcription factors Cody (reviewed in Sonenshein, 2005) and AbrB (reviewed in Phillips and Strauch, 2002), the kinases and
366 (Fujita et al., 2005). M. xanthus genes that depend on C-signaling for expression have been shown to respond to different threshold levels of C-signal (Kim and Kaiser, 1991) and are regulated spatially (Julien et al., 2000; Sager and Kaiser, 1993), so it is tempting to speculate that FruA, which mediates responses to C-signaling (see chapters 4 and 9), uses mechanisms similar to that of B. subtilis SpoOA to directly regulate transcription of many genes expressed at different times and in different domains during M . xanthus fruiting body development.
OVERVIEW OF SUBSEQUENT MORPHOLOGICAL CHANGES AND CELL-SPECIFIC (T FACTORS Shortly after completion of asymmetric division, up becomes active in the forespore, generating a signal that leads to the formation of active aE in the mother cell (Fig. 1C). Differential transcription in the two cell types results in synthesis of proteins necessary for engulfment, a phagocytic-like process in which the mother cell membrane migrates around the forespore, fusing near the pole to pinch off the forespore (now surrounded by a second membrane often referred to as the outer forespore membrane [OFM]) and release it into the mother cell cytoplasm (Fig. 1D). Completion of engulfment seems to trigger activation of aGin the forespore, and this generates a signal that leads to formation of active aK in the mother cell (Fig. 1E). A loosely cross-linked peptidoglycan layer called the cortex is synthesized between the inner and outer forespore membranes (Fig. 1E). Proteins made in the mother cell assemble on the forespore surface to produce the spore coat (Fig. 1F). The spore matures within the mother cell and is eventually released by programmed autolysis of the mother cell. For more-detailed accounts of the morphological changes during B. subtilis endospore formation, and especially the regulation of compartmentalized gene expression, including comprehensive citation of earlier work, the reader is referred to Piggot and Losick, 2002, and Hilbert and Piggot, 2004. Also, a shorter review highlighting recent findings has appeared (Piggot and Hilbert, 2004). The general picture to emerge is that completion of two morphological events (polar division and engulfment) triggers activation of two foresporespecific a factors (aFand a") that initiate signal transduction pathways leading to formation of two mother cell-specific a factors (aEand aK).Hence, gene expression in each cell type is governed by a different a factor cascade, but the two cascades are coupled to morphogenesis and coordinated by intercellular signaling. The extent to which M. xanthus development relies on a factor
ANALOGOUS SYSTEMS cascades remains to be determined. C-signaling appears to couple developmental gene expression to morphogenesis (Kruse et al., 2001), apparently by monitoring contact between cells (see chapter 4). This is quite different from the septation and membrane fusion events that seem to be monitored during B. subtilis sporulation. Also, while signals between the forespore and mother cell must be transmitted across the two membranes that separate the cytoplasms of the two cell types, they need not overcome outer membrane and peptidoglycan barriers, as is the case for signaling between M. xanthus cells. Therefore, some of the mechanisms of intercellular signaling and morphological coupling are likely to be quite different in the two organisms. So, too, are the products of the two differentiation processes quite different, with B. subtilis endospores being considerably more resistant to environmental insults (reviewed in Driks, 2002, and Nicholson et al., 2000) than myxospores. Below, we summarize our understanding of how morphogenesis and intercellular signaling control the activity of cell-specific a factors, focusing on recent progress and attempting to identify questions that remain. We also review the results of genomic approaches to characterize the regulon of each cell-specific a factor and the functions of some of the gene products.
REGULATION OF uFACTIVITY uFis the first sporulation-specific sigma factor to become active during formation of spores by B. subtilis. Appearance of the activities of the later sporulation-specific sigma factors depends on aFactivity. aFis formed in the predivisional cell after the start of spore formation but is held inactive until completion of the sporulation division septum. It becomes active exclusively in the forespore (Gholamhoseinian and Piggot, 1989; Margolis et al., 1991; Piggot and Losick, 2002). It becomes active very soon after the asymmetric division and before the chromosome has completely partitioned into the forespore (Frandsen et al., 1999; Wu and Errington, 1994). Genetic and biochemical studies have identified three proteins that are involved in the regulation of aFactivity: SpoIIAB, SpoIIAA, and SpoIIE. Yudkin and Clarkson (Yudkin and Clarkson, 2005) have recently written an excellent review emphasizing the critical nature of the biochemical reactions controlling aFactivation, while Barak and Wilkinson (Barak and Wilkinson, 2005) provide an excellent review of compartmentalized aF activation that nicely complements that of Clarkson and Yudkin in its approach. Hilbert and Piggot (Hilbert and Piggot, 2004) provide a historical perspective on aF studies.
21. B. sumxrs SPORULATION When first formed, uF is kept inactive in a complex with the anti-sigma factor SpoIIAB. SpoIIAA acts as an anti-anti-sigma factor to disrupt the complex (Duncan et al., 1996; Schmidt et al., 1990). However, early in spore formation SpoIIAA is present in a phosphorylated form, which is inactive (Carniol et al., 2004; Magnin et al., 1997). It is SpoIIAB that functions as the kinase to phosphorylate SpoIIAA (Min et al., 1993). SpoIIE is the phosphatase that dephosphorylates SpoIIAA-PO, (Duncan et al., 1995), thus activating it and enabling it to interact with the SpoIIAB-uFcomplex, bind SpoIIAB, and hence release and thus activate oF.We now have considerable insight into these controls, but there remain questions about how uF is activated so quickly after septum formation and why its activity is completely compartmentalized. The dramatic activation that occurs exclusively in the forespore after septation requires an equally dramatic regulation. The structural gene for oF,spoIIAC, is the third gene of the spoIIA operon. SpoIIAB is the product of the second and SpoIIAA of the first gene in the operon. Transcription of the spoUA operon is strongly induced after the start of spore formation and before the sporulation division. Induction requires the master regulator SpoOA, which is activated by the phosphorelay (reviewed in Hilbert and Piggot, 2004). uF is a 255-residue protein of the u70 class. SpoIIAB is a 146-residue protein that in the presence of ATP (or ADP) binds to and inhibits uF(Alper et al., 1994; Magnin et al., 1997).The inhibitory complex contains two SpoIIAB molecules to one uF (Campbell and Darst, 2000). SpoIIAB can also form a complex with SpoIIAA in the presence of ATP or ADP (Alper et al., 1994; Magnin et al., 1997). In the presence of ATP it can act as a kinase to phosphorylate SpoIIAA on residue Ser.58. SpoIIAA is a 117-residue protein, and in its phosphorylated form it is unable to bind SpoIIAB (Diederich et al., 1994; Min et al., 1993). Prior to spore septum formation, SpoIIAA is predominantly in its phosphorylated form (Carniol et al., 2004; Magnin et al., 1997), with uF held inactive as the SpoIIAB2: oF complex. The spoIIE gene is monocistronic. Its induction after the start of spore formation also depends on SpoOA (York et al., 1992). SpoIIE is an 827-residue protein with 10 putative membrane-spanning regions in its N-terminal domain (Arigoni et al., 1999). Its C-terminal domain is a phosphatase that dephosphorylates SpoIIAA-PO, (Duncan et al., 1995).Its central domain interacts with FtsZ (Lucet et al., 2000), and it colocalizes with FtsZ at the site of the asymmetrically located sporulation division (Levin et al., 1997). It is required for efficient movement of FtsZ to the site of the sporulation division
367 and for efficient division (Ben-Yehudaand Losick, 2002; Khvorova et al., 1998). SpoIIE is present at the site of septation before the spore septum is completed (Arigoniet al., 1995).Presumably it has the potential to dephosphorylate SpoIIAAPO,. However, not much dephosphorylated SpoIIAA is detected and uFis not active (Carniol et al., 2004; Magnin et al., 1997).Something about completion of the septum either activates SpoIIE or in some other way changes the balance in the uFregulatory system, or both. In support of this catchall statement, hyperactive mutants of SpoIIE cause the activation of uFwithout the need for septation (Carniol et al., 2004; Feucht et al., 2002; Hilbert and Piggot, 2003). The interactions among SpoIIAB, SpoIIAA, SpoIIE, and uFare complex (Fig.2A). Three factors that are presently thought to contribute to uF activation and to its continued activity in the forespore are considered here: the concentration in the forespore of SpoIIE, the stability of the complex that sequesters SpoIIAB and SpoIIAA, and transient genetic asymmetry. The most striking feature of the division during sporulation of B. subtilis is its asymmetric location. The septum is formed so close to the pole of the dividing cell that it is difficult to measure its position accurately by fluorescence microscopy, but a four- to eightfold difference in volume between mother cell and forespore seems plausible (Carniol et al., 2004; Iber et al., 2006). The published evidence suggests that SpoIIE is distributed about equally between the two faces of the septum when that is first formed (Kinget al., 1999). Because its phosphatase activity is low, most SpoIIE may be associated with its substrate, SpoIIAA-PO,, while the rest of the reactants (Fig. 2B) are cytoplasmic and plausibly have a uniform concentration in the forespore and the mother cell. If correct, then the Spo1IE:SpoIIAA-PO, concentration is four to eight times higher in the forespore than in the mother cell (Fig. 2B) (Iber et al., 2006). Importantly, the concentration of SpoIIE is not linearly related to activation of uFin vitro; in the right conditions, a 10-fold increase in the concentration of SpoIIE largely dephosphorylates SpoIIAA-PO, and results in uF activation (Clarkson et al., 2004). Mathematical modeling, based on published in vitro experiments of the reactions known to be involved in uF activation, suggests that a sudden increase in Spo1IE:SpoIIAA-PO, concentration in the forespore upon completion of septation would be sufficient to dephosphorylate SpoIIAA and activate crF (Iber et al., 2006). However, experiments with a strain that synthesized only the soluble, C-terminal domain of SpoIIE suggest that, even if the above is correct, there must be other partially redundant mechanisms contributing to the forespore-specific activation of uF.This
ANALOGOUS SYSTEMS
368 A
I
4
1 ,)r
IIAA
+ IIABA,p-oF+
IIABATp-IIAA
+
oF
1
IIAA-P
+ IIAB,,
I B
of spoZZAB
only a mild effect on compartmentalization, and hence spore formation. Any explanation of why uFis activated on completion of septation must also explain why its activity is maintained. In other words, why is uF activity not rapidly damped down with the various players returning to their previous equilibrium? A major determinant is likely to be a long-lived intermediate that sequesters SpoIIAB and SpoIIAA: immediately after the sudden accumulation of dephosphorylated SpoIIAA in the forespore, formation of this complex has the effect of impeding the ability of SpoIIAB to (i) bind to and inhibit uFand (ii) phosphorylate and inactivate SpoIIAA. So long as any SpoIIAA-PO, continues to be actively dephosphorylated, the longlived complex favors continued uF activity (Yudkin and Clarkson, 2005). On the other hand, before septation the same long-lived complex effectively sequesters any small amount of dephosphorylated SpoIIAA present, so that the system is buffered against activating uF when there is limited dephosphorylation of SpoIIAA (Carniol et al., 2004). The results suggest that a threshold concentration of unphosphorylated SpoIIAA needs to be reached in order to trigger the activation of uF(Barak and Wilkinson, 2005; Carniol et al., 2004; Yudkin and Clarkson, 2005). Another consideration is the transient genetic asymmetry that characterizes the forespore (and mother cell) when first formed. Upon closure of the sporulation septum, only the origin-proximal 30% of the chromosome destined for the forespore is actually present in the forespore (Wu and Errington, 1998).It requires the action of the DNA translocase SpoIIIE to transfer the rest of the chromosome into the forespore (Bath et al., 2000; Wu et al., 1995), a process that has been estimated to take about 15 min (Khvorova et al., 2000; Pogliano et al., 1999). Frandsen et al. (Frandsen et al., 1999) proposed that this transient genetic asymmetry could be important for the establishment of compartmentalized uF activity. Notably, the spollA locus is located near the terminus and so is not present in the forespore when it is first formed. Frandsen et al. (Frandsen et al., 1999) showed that moving spolIAC, encoding uF,near the origin conferred some ability to form spores in a strain with spoIIAB remaining near the terminus, even in the absence of SpoIIAA and SpoIIE, which are normally essential for spore formation. Related experiments have reinforced the importance of transient genetic asymmetry with respect to spoIIA and point towards lack of replenishment of SpoIIAB in the newly formed forespore as being critical (Dworkin and Losick, 2001; Dworkin, 2003; Pan et al., 2001). In support of this model, SpoIIAB is unstable when it is not bound to either SpoIIAA or uF, and a strain encoding a stable derivative sporulates poorly (Pan et al., 2001).
i SpoIIAB protein degraded in forespore
Figure 2 Regulation of uFactivity. (A) Illustration of the reactions that result in uFbeing activated in the forespore. Critical determinants are thought to be the concentrations of SpoIIE and SpoIIAA-PO,; a long-lived SpoIIAA-SpoIIAB-ADP complex (a SpoIIAAlSpoIIAB “sink”);and instability of free SpoIIAB combined with transient genetic asymmetry so that both copies of the spoIIAB gene are in the mother cell. (B) Schematic illustration of the effects of the sporulation division on the regulators of uF.The SpoIIAA-PO, protein (open pentagons) is presumed to be evenly distributed throughout the cytoplasm so that most is present in the mother cell. The SpoIIE protein (filled diamonds) is associated with the septum, and most of it may be associated with SpoIIAA-PO,, such that the SpoIIE: SpoIIAA-PO, complex is distributed equally between mother cell and forespore. Only the origin-proximal 30% of a chromosome is present in the forespore when first formed. As a consequence both copies of the spoIIAB gene are present in the mother cell (MC),and degradation disproportionally reduces the SpoIIAB concentration in the forespore.
strain formed spores with moderate efficiency, about 30% of the parental strain, and uF activity was forespore specific in about one-half of the bacteria displaying activity (Arigoni et al., 1999). These results indicate that equal partitioning of SpoIIE between the forespore and mother cell faces of the septum may be important for the compartmentalization of uFactivation but is not essential for it. Rather there appears to be more than one mechanism ensuring compartmentalization so that loss of just one, such as equimolar partitioning of SpoIIE, has
21. B. SUBTILIS
369
SPORULATION
However, the half-life of wild-type SpoIIAB is about 30 min, and this seems too long to be critical in the first few minutes after septation when uFbecomes active in the forespore; it seems more suggestive of a mechanism that enforces and prolongs uF activity after it has first been established. Such an enforcer role, which can be partly compensated by proper localization of SpoIIE to the septum (Dworkin and Losick, 2001), may be important for efficient spore formation. The origin proximal location of spoIIE appears to be less important for spore formation than the origin distal location of spoIIA (McBride et al., 2005). It is appropriate to introduce a related species of endospore former, Sporosarcina ureae. S. ureae is a coccus, and there is little or no volume asymmetry between mother cell and forespore when first formed, yet it sporulates and displays compartmentalized activity of uF and uE (Zhang et al., 199713; V. Chary and P. J. Piggot, unpublished data). The ratios of septally located to soluble proteins should not differ greatly between mother cell and forespore as a result of division and so should not trigger compartmentalized gene expression. However, spore formation requires the DNA translocase SpoIIIE, suggesting that transient genetic asymmetry may be important, possibly even more important than in B. subtilis, for establishing compartmentalized uFactivity (Chary et al., 2000).
THE uFREGULON The uF regulon has recently been carefully analyzed by microarray by two groups (Steil et al., 2005; Wang et al., 2006). The results are in general agreement. They indicate 48 to 55 genes, many encoding proteins of unknown function, whose transcription is directed by uF. The analyses confirmed the previous impression (Piggot and Losick, 2002) that uF had the smallest of the four regulons directed by the sporulation-specific sigma factors. The analysis of Steil et al. (Steil et al., 2005) greatly extended previous results indicating two temporal classes of uF-directed genes. There is overlap in promoter specificity between uF and uc (Helmann and Moran, 2002; Wang et al., 2006); those promoters that are recognized by uGare in the second temporal class (Steil et al., 2005). Chromosome position (transient genetic asymmetry) may be important for some oF-directed genes, because moving the spoIIR locus from its normal origin-proxima1 location to the terminus severely impairs expression and spore formation (Khvorova et al., 2000; Zupancic et al., 2001). Key genes in the regulon include genes required for the expression of all the later sigma factors: spoIR, which
is required for activation of uE (Karow et al., 1995; Londono-Vallejo and Stragier, 1995); spoIIIG, which is the structural gene for uG(Sun et al., 1991); spoIIQ, which appears to have several roles including regulation of uc activity and of spoIIIG transcription (LondonoVallejo et al., 1997; Sun et al., 2000); and spoIVB and bofC, which encode regulators of uK(Gomez and Cutting, 1996, 1997). As their spo name implies, most of these regulators are essential for spore formation. Transcription of spoIIIG, which falls in the second temporal class, is unusual among uF-directed genes in that it requires a uE-directed signal from the mother cell (Partridge and Errington, 1993). The uF regulon includes two regulators of uF, lonB and rsfA, although neither is critical to spore formation (Serrano et al., 2001; Wu and Errington, 2000). Also of note is an inhibitor of cell division (Hilbert et al., 2004), which has recently been identified as spoIIl’ and has been associated with the phenomenon of commitment to form spores (Dworkin and Losick, 2005); spollP is unusual in that its transcription is directed by uE in the mother cell (Frandsen and Stragier, 1995) as well as by uFin the forespore from the upstream gpr promoter (Dworkin and Losick, 2005).
REGULATION OF mE ACTIVITY Like the regulation of uFactivity, that of uEappears to be ensured by multiple factors. These include preferential expression of the uEprecursor, pro-uE,in the mother cell (Arabolaza et al., 2003; Fujita and Losick, 2003) and degradation of pro-uE in the forespore (Fujita and Losick, 2002; Ju et al., 1997, 1998; Pogliano et al., 1997), ensuring that uEbecomes active only in the mother cell. Timing of pro-uEactivation, which involves proteolytic removal of its N-terminal 27-amino-acid prosequence (LaBell et al., 1987), is tightly linked to uF activation in the forespore. uF RNAP transcribes the spoIIR gene which encodes a signal protein (Hofmeister et al., 1995; Karow et al., 1995; Londono-Vallejo and Stragier, 1995) that leads to activation of SpoIIGA (Fig. 3 ) , the putative protease believed to cleave pro-oE (Jonas et al., 1988; Masuda et al., 1990; Stragier et al., 1988). This must all happen rapidly so that genes under uERNAP control are expressed in the mother cell and the products prevent formation of a second polar septum (Eichenberger et al., 2001) and commit the mother cell to sporulation (Dworkin and Losick, 2005). Transcriptional regulation of the bicistronic spoIIG operon, which encodes the putative protease SpoIIGA and its apparent substrate pro-uE(also called SpoIIGB), ensures that these proteins are produced predominantly in the mother cell. Transcription of the spoIIG promoter
ANALOGOUS SYSTEMS
3 70
forespore
moth.er cell
[@<]
'..
._
Figure 3 Signaling pathway governing pro-uE processing. The upper part shows the sporangium after the asymmetric septum forms. The lower part shows components of the signaling pathway. SpoIIR made in the forespore is translocated across the forespore membrane of the septum and activates SpoIIGA to cleave p r o d .
by crARNAP is activated by phosphorylated SpoOA prior to asymmetric septum formation but after septation persists mainly in the mother cell (Fujita and Losick, 2003). SpoIIAA (specifically, its nonphosphorylated form) appears to inhibit phosphorylation of SpoOA in the forespore and therefore reduce transcription of SpoOAP04-dependent genes and operons like spoIIG (Arabolaza et al., 2003). As noted above, SpoIIAA likely reaches a higher level in the smaller forespore compartment than in the mother cell due to SpoIIE phosphatase activity. Transient genetic asymmetry aids the differential expression, as the spoIIG operon is origin-distal, and therefore, two copies are present in the mother cell at the time of septation and prior to chromosome translocation into the forespore (McBride et al., 2005). Pro-aE made in the forespore is not processed but is instead degraded (Fujita and Losick, 2002; Ju et al., 1997). However, the normal confinement of pro-oE processing and oEaccumulation to the mother cell breaks down in a spoIIIE null mutant unable to translocate one copy of the chromosome into the forespore (Pogliano et al., 1997), especially if the spoIIG operon is experimentally fused to a oF-dependent promoter and placed in the origin-proximal region of the chromosome (which is captured in the forespore at the time of septation) (Ju et al., 1998). These results have been interpreted as evidence that an origin-distal protease( s) normally degrades pro-aE, SpoIIGA, and oEmade in the forespore (Ju et al., 1998). Consistent with this idea, expression of the spoIIG operon from a constitutive oA-dependent promoter did not impair sporulation, and fluorescence
from pro-aE/oEtagged with the green fluorescent protein (GFP) disappeared selectively from the forespore after septation (Fujita and Losick, 2002). The putative protease(s) responsible for preventing aEactivity in the forespore has not been identified. It is important to do so, because while it appears that the protease(s) allows sporulation in the absence of normal spoIIG transcriptional regulation, it would be interesting to test whether transcriptional regulation alone, in the absence of the protease(s), would permit normal sporulation. It appears that processing of pro-oEin the mother cell begins within minutes after septation and activation of uF in the forespore, during the approximately 15-min period of transient genetic asymmetry before a copy of the chromosome is fully translocated into the forespore (Khvorova et al., 2000; Zupancic et al., 2001). This conclusion is inferred from studies in which the spoIIR gene, normally origin proximal and therefore present in the forespore at the time of septation, was moved to origindistal chromosomal locations, delaying and reducing both oF-dependent transcription of spoIIR in the forespore and processing of pro-aE in the mother cell, and decreasing sporulation efficiency due to the formation of a second polar septum in the mother cell. Normally, the products of &dependent genes prevent the second septum from forming (see below). Hence, synthesizing SpoIIR too late is detrimental. On the other hand, making SpoIIR too early, prior to septation, is of relatively little consequence (Pogliano et al., 1997; Zhang et al., 1996), as observed for pro-oE and SpoIIGA (Fujita and Losick, 2002), apparently because the mechanisms discussed above ensure that active aEaccumulates only in the mother cell. SpoIIR is believed to be secreted from the forespore into the septa1 space and has been proposed to interact with SpoIIGA, activating its putative protease domain to cleave pro-oE and release active aEinto the mother cell (Hofmeister et al., 1995; Karow et al., 1995; Londono-Vallejo and Stragier, 1995). SpoIIGA and pro-aE are membrane associated and localize to the polar septum (Fawcett et al., 1998; Hofmeister, 1998; Ju et al., 1997; Peters and Haldenwang, 1991), but other aspects of the model remain to be tested. The putative protease domain of SpoIIGA exhibits features of an aspartyl (Stragier et al., 1988) or serine protease (Masuda et al., 1990). All of the elements necessary for pro-aEprocessing reside in its N-terminal 27-amino-acid prosequence (Carlson et al., 1996), and a Glu residue at position 25 has been shown to be important (Peters et al., 1992). Changing Glu at position 25 to Lys makes processing inefficient, but this mutation can be suppressed by Pro-to-Leu substitution at position 259 in the putative
21. B. SUBTILIS SPORULATION protease domain of SpoIIGA (Peters and Haldenwang, 1994). This argues that SpoIIGA interacts directly with pro-uE, but the suppression was not allele specific, so other interpretations are possible. The N-terminal 55 amino acids of pro-aEis sufficient to target GFP to the polar septum (Ju et al., 1997), and this is necessary for processing (Ju and Haldenwang, 2003), consistent with localization of SpoIIGA-GFP to the septum (Fawcett et al., 1998) and direct interaction of the two proteins. Additional evidence for a direct interaction between prooEand SpoIIGA comes from the unexpected finding that both proteins are necessary for aG to accumulate in the mother cell of mutants defective in producing aFand its anti-sigma factor SpoIIAB (Chary et al., 2005). It is hypothesized that pro-aEprevents SpoIIGA from stimulating proteolysis of aG,possibly by the LonA protease. Of course, under normal circumstances aGaccumulates in the forespore, not in the mother cell (see below).
THE uEREGULON Four studies have used microarray analysis to define genes transcribed by aE RNAP (Eichenberger et al., 2003, 2004; Feucht et al., 2003; Steil et al., 2005). Though the studies differ in their estimate of the size of the aEregulon, they agree that it is the largest of the cell-type regulons, comprising 154 to 262 genes. Null mutations were created in many of the newly identified genes and operons, and 15 of these exhibited defects in sporulation (Eichenberger et al., 2003; Feucht et al., 2003). Together with previous mutant analysis, about 20% of the genes in the oEregulon are required for sporulation (Eichenberger et al., 2003). This is a high percentage compared to a study of M. xanthus in which only 3 Tn.5 lac insertions caused a developmental defect among a set of 29 insertions in distinct developmentally regulated transcription units (Kroos et al., 1986). Nevertheless, in both cases, many developmental genes appear to have redundant or nonessential functions, at least under laboratory conditions and within the limitations of phenotypic observation employed. Of the 45 crE-controlledgenes required for B. subtilis sporulation, many (about 70%) are present in the genomes of other endospore-forming gram-positive bacteria, but not in the genome of Listeria monocytogenes, a close relative that does not form spores (Eichenberger et al., 2003). Conversely, of the 46 aE-controlled genes conserved among endospore formers, about 30% are not required for B. subtilis sporulation. Why these genes are conserved and why other genes are required for sporulation of B. subtilis but not other endospore formers are important evolutionary questions.
3 71
Some of the key functions of oE-dependentgene products have been mentioned above. SpoIIP, made under uE control in the mother cell and under aF control (from the promoter of the upstream gpr gene) in the forespore, blocks further growth (with SpoIIQ also involved in the forespore), committing each cell type to its fate (Dworkin and Losick, 2005). SpoIIP, together with oE-controlled SpoIID and SpoIIM, is involved in septa1 peptidoglycan hydrolysis, and these three proteins help prevent formation of a second polar septum in the mother cell (Eichenberger et al., 2001) and are crucial for engulfment (Abanes-De Mello et al., 2002; Pogliano et al., 1999). Other key oE-controlled gene products include enzymes involved in synthesis of the spore cortex, structural proteins of the spore coat (mainly the inner coat layer, as well as several important for coat assembly), proteins that support metabolic activity in the mother cell (enzymes involved in lipid metabolism and protein turnover and transporters to scavenge nutrients from the environment), proteins required for activation of aG in the forespore upon completion of engulfment (products of the spoIIIA operon, see below), pro-aKand proteins that govern its activity (see below), and the transcription factors SpoIIID and GerR (Eichenberger et al., 2003,2004; Feucht et al., 2003; Steil et al., 2005). GerR appears to be a negative regulator of 14 genes in the oE regulon, while SpoIIID acts more broadly as a negative regulator (112 genes) but also acts positively (10 genes) (Eichenberger et al., 2004), in many cases acting directly as a repressor or activator of transcription (Halberg and Kroos, 1994; Ichikawa and Kroos, 2000; Kroos et al., 1989; Zhang et al., 1997a). Since SpoIIID is itself under aEcontrol, genes requiring SpoIIID for transcription are induced slightly later than genes requiring only oERNAP (Steilet al., 2005). Genes negatively regulated by SpoIIID and GerR are expressed in a pulse (Eichenberger et al., 2004). Hence, GerR and especially SpoIIID diversify the patterns of gene expression in the aEregulon.
REGULATION OF uGACTIVITY Regulation of uGsynthesis and activity is not completely understood, but it is clear that its activity must be tightly regulated because upon activation aGdirects transcription of its own structural gene, spolllG or sigG, creating a positive-feedback loop that drives additional synthesis of active aG(Sun et al., 1991). Transcription of sigG is initiated from two promoters (PspollGand PsigG)(Masuda et al., 1988; Sun et al., 1991) (Fig. 4). The spoIIG operon is located immediately upstream from sigG, and some transcripts from the spoIlG promoter run through the end of the spollG operon into sigG (Masuda et al.,
ANALOGOUS SYSTEMS
3 72
I
I
ll"'" spollGA
sigE
I
1 sigG
Figure 4 Organization and transcription of the spoIIG and sigG operons. Indicated are the two genes of the spoIIG operon, spoIIGA, the protease believed to process pro-uE,and sigE, the structural gene for pro-oE. Also shown is sigG, the gene encoding uG.An open reading frame located downstream from sigG, ylmA (not shown), is cotranscribed with sigG. Transcription from PspoIIGrequires a*-RNAP and phosphorylated SpoOA. Some of the PspoIIG transcript reads through the sigG operon. P,, is used weakly by uFRNAP and more strongly by uG RNAP.
1988; Sun et al., 1991).The spoIIG promoter is active in the mother cell after septation (Fujita and Losick, 2003), raising the possibility that uG could be produced in the mother cell. However, at least three factors act to prevent accumulation or activity of uG in the mother cell. First, comparison of transcriptional and translational lac2 fusions in sigG indicates that very little uGis translated from the read-through transcripts (Sun et al., 1991).Two additional factors prevent activity and accumulation of uGif synthesized from the read-through transcript in the mother cell. The anti-sigma factor SpoIIAB can inhibit uGactivity in vivo when coexpressed with the sigma factor, and in vitro SpoIIAB binds uG (Kellner et al., 1996; Kirchman et al., 1993; Serrano et al., 2004). However, because SpoIIAB also regulates uFactivity and uFdirects transcription of sigG, a simple genetic test to examine the effect of a null mutation in spolIAB on uGactivity is problematic. An alternative test involved examination of two different single amino acid substitutions in uGthat prevented binding by SpoIIAB without affecting uGfunction (Kellner et al., 1996; Serrano et al., 2004). These amino acid substitutions resulted in increased uGactivity in vivo; however, this activity occurred primarily in the mother cell compartment. Therefore, SpoIIAB appears to play a role in preventing uGactivity in the mother cell. This regulation of uG by SpoIIAB in the mother cell may be important because a low level of active uG,produced by readthrough transcription from the upstream spoIIG operon, would lead to runaway expression of uG in the mother cell if not prevented by SpoIIAB. LonA protease also appears to play a similar role by reducing the level of uGaccumulation in both the mother cell and the predivisional cell (Serrano et al., 2001). The sigG promoter (Psjgc) is located between the spollG operon and the beginning of sigG (Masuda et al., 1988; Sun et al., 1991) (Fig. 4). uF-directedtranscription
from this promoter in the forespore produces a transcript that is efficiently translated and results in accumulation of uG.However, the uGdoes not become active until the completion of forespore engulfment (Errington et al., 1992; Illing and Errington, 1991). It is unknown how uGactivity is regulated in response to engulfment. The N-terminal sequence of uGis the same as that predicted from the gene sequence (Sun et al., 1989), and the uG that accumulates before engulfment is able to function in vitro (C. P. Moran, unpublished data); therefore, uG evidently does not require covalent modification or proteolytic processing for its activation. Although it seems likely that there is a direct negative regulator of uGin the forespore, such a regulator remains to be discovered. SpoIIAB is likely not the sole negative regulator of uGin the preengulfment forespore since the previously described experiments with the mutant forms of uGthat are not bound by SpoIIAB result in uGincreased activity mainly in the mother cell and not in the forespore (Serran0 et al., 2004). Mutations in two operons (spolll] and spoIIIA) prevent activation of uG,although uGaccumulates and engulfment of the forespore by the mother cell appears to be normal in these mutants (Errington et al., 1992; Illing and Errington, 1991).The spoIIIJ operon encodes two genes, only one of which, spolll], is required for uGactivation and sporulation (Errington et al., 1992). SpoIIIJ is a member of the YidC/Oxal/Alb3 family, which includes proteins that are required for the proper insertion or folding of proteins into membranes (Serrano et al., 2003; Tjalsma et al., 2003; Yi and Dalbey, 2005). The B. subtilis genome also encodes a SpoIIIJ paralog called YqjG. Both genes are expressed during growth, and at least one, SpoIIIJ or YqjG, is required for viability, indicating some redundancy of function. However, only SpoIIIJ is required for sporulation (Serrano et al.,
21. B. s u m m SPORULATION 2003). Evidently, both SpoIIIJ and YqjG can mediate the insertion of membrane proteins required for growth, but proper insertion into the membrane of at least one protein that is necessary for the activation of uc requires SpoIIIJ. One such sporulation-essential membrane protein may be SpoIIIAE, which is encoded by the spollIA operon. Evidence supporting a model in which SpoIIIAE is a substrate of SpoIIIJ includes a synthetic lethal phenotype that is created by expression of SpoIIIAE in vegetative B. subtilis cells of a spoIIl] mutant, and the observation that SpoIIIJ and SpoIIIAE can be shown to interact in cross-linking experiments when coexpressed in Escherichia coli (M. Serrano, E Vieira, C. P. Moran, and A. 0. Henriques, unpublished data). Thus, dependence on SpoIIIJ for uGactivation appears to be due to its role in insertion of SpoIIIAE into the membrane where it functions. SpoIIIAE is encoded within the spolIlA operon, which encodes eight proteins. Seven of these are predicted to be integral membrane proteins, and the eighth, SpoIIIAA, is possibly a peripheral membrane ATPase. Each of the eight spoIIIA genes (spoIIIAA through spoIIIAH) is required for uGactivation (Kellner et al., 1996; Moran, unpublished). The spoIlA operon is transcribed by uERNAP in the mother cell (Eichenberger et al., 2004; Illing and Errington, 1991; Steil et al., 2005).Although produced in the mother cell, SpoIIIAH becomes localized to the membrane surrounding the forespore (Blaylock et al., 2004). SpoIIIAH spans the mother cell membrane with its C terminus protruding into the space between the mother cell and forespore. Initially, SpoIIIAH is inserted randomly around the mother cell membrane. However, after septa1 thinning, diffusion of SpoIIIAH leads to its capture at the septum between the two cells through direct binding to the C terminus of SpoIIQ, which protrudes from the forespore membrane (Blaylock et al., 2004) (Fig. 5).The two proteins, SpoIIQ, which is synthesized in the forespore, and SpoIIIAH, which is made in the mother cell, zipper one another around the forespore as engulfment is completed (Blaylock et al., 2004). It seems likely, and some preliminary data suggest (Moran, unpublished), that the other spoIllA products are also localized around the forespore membrane, where they form a complex. The function of the proteins is unknown. However, given the size of this complex, its location around the forespore, and the potential ATPase encoded by SpoIIIAA, it is tempting to speculate that the complex formed by these proteins could serve to translocate a substrate from the mother cell into the intermembrane space between the mother cell and forespore membranes, or into or out of the forespore. In the former scenario, completion of engulfment would isolate the intermembrane space
3 73
A
IFM
OFM
B CtpB C
pro-oK
C
Figure 5 Model for the regulation of pro-oKprocessing. (A) The upper part shows the sporangium after engulfment of the forespore within the mother cell. Black ovals represent a protein complex that bridges the two membranes surrounding the forespore. The lower part shows an expanded view of the protein complex that bridges from the inner forespore membrane (IFM) to the outer forespore membrane (OFM). SpoIVB is a serine protease made initially under uFcontrol in the forespore and believed to be translocated across the IFM. A low level of SpoIVB is sufficient to cause proteolysis of the SpoIIQ extracellular domain. uG RNAP boosts the level of SpoIVB and it cleaves the C-terminal extracellular domain of SpoIVFA. An unknown protein (Protein X) is proposed to localize the pro-uKprocessing machinery to the SpoIIQ-SpoIIIAH complex. (B) Loss of SpoIVFA renders BofA susceptible to cleavage by CtpB, a serine protease made under uEcontrol in the mother cell and under uGcontrol in the forespore. CtpB is believed to be translocated into the space between the two membranes, where it targets a short C-terminal extracellular domain of BofA. (C) Loss of BofA allows SpoIVFB to cleave pro-uK, releasing active uKinto the mother cell. See the text for references.
3 74 from the external milieu; therefore, the action of the SpoIIIA pump would result in a rapid shift in substrate concentration within the intermembrane space, possibly signaling the completion of engulfment. It is less obvious how the completion of engulfment would be sensed in the latter model, where the pump moves a substrate into or out of the forespore. Nevertheless, this model must also be considered because recent results show that SpoIIIAE may be able to function when expressed in the forespore (Serrano et al., unpublished), suggesting that in wild-type cells SpoIIIAE may span both mother cell and forespore membranes. Therefore, we are left with two fundamental questions regarding uGactivation: how is the completion of engulfment sensed, and what is the direct regulator of uGactivity?
THE uGREGULON uGdirects transcription of at least 95 genes distributed throughout the chromosome. Two studies using microarrays (Steil et al., 2005; Wang et al., 2006) have found that the uGregulon includes a few regulatory genes but that most of the genes encode proteins that are devoted to building the spore and preparing it for dormancy and subsequent germination. Very few of the genes, about half a dozen, encode metabolic enzymes. These function in glucose uptake, glycolysis, or glycine metabolism. However, it appears that most of the energy and building blocks required at this stage of development are supplied by the mother cell. Many of the genes in the uGregulon encode proteins that prepare the spore for dormancy. For example, the spo VA operon encodes proteins that import dipicolinic acid from the mother cell (Tovar-Rojo et al., 2002). Dipicolinic acid makes up about 10% of the dry weight of spores and is essential to the spore’s resistance to wet heat and maintenance of dormancy. The regulon also includes over a dozen genes encoding small acid soluble proteins (SASPs), which are abundant, small proteins that bind DNA to protect the chromosome from damage (reviewed in Nicholson et al., 2000). Other uGregulon members, including the spl and ykoVU operons, encode proteins involved in DNA repair. ykoV and ykoU encode DNA break repair proteins that mend DNA breaks during germination, probably by a nonhomologous endjoining mechanism (Wang et al., 2006). Like ykoVU, other uc-dependent genes encode products that function during germination. sleB encodes a cortex lytic enzyme that degrades the peptidoglycan cortex during germination (Wang et al., 2006). The receptors that signal germination are encoded in four tricistronic operons (gerA,gerB, gerK, and probably the
ANALOGOUS SYSTEMS ynd operon) all of which are transcribed by uG RNAP (Steil et al., 2005; Wang et al., 2006). The 6-dependent gerD gene also encodes a protein required for signaling germination, but its mechanism is not known. During germination the SASPs are degraded by a protease that is encoded by the &dependent gene gpr (Sanchez-Salas et al., 1992; Setlow, 2003). The regulatory genes expressed as part of the uc regulon include spolllG, or sigG, which encodes ac. As described above, activation of uGproduces a positivefeedback loop in which uG directs its own synthesis. uG also directs expression of spoVT, which encodes a DNA-binding protein that affects expression of 47 uGdependent genes (Bagyan et al., 1996; Shcheptov et al., 1997; Wang et al., 2006). SpoVT acts with uc in both coherent and incoherent feedforward loops, activating or stimulating expression of 20 genes and repressing 27 genes respectively (Wang et al., 2006). SpoVT appears to contribute to the switch from the final stage of forespore gene expression to the final stage of mother cell gene expression by repressing transcription of sigG, and by stimulating transcription of sgolVB, which signals uK activation in the mother cell (Wang et al., 2006).
REGULATION OF uKACTIVITY The regulation of uK activity shares some similarities with the regulation of uE,but there are clear mechanistic differences. Like uE, aKis made as an inactive precursor, pro-uK,that associates with membranes via its prosequence (Kroos et al., 1989; Prince et al., 2005; Stragier et al., 1989; Zhang et al., 1998). Also, the prosequence is cleaved by a membrane-embedded protease in response to a signal from the forespore (Cutting et al., 1990,1991a; Lu et al., 1990, 1995; Rudner et al., 1999; Yu and Kroos, 2000; Zhou and Kroos, 2004). Until recently, the only known signal was SpoIVB, a serine protease synthesized under dual control of uFand crc in the forespore and secreted into the space between the two membranes surrounding the forespore after engulfment (Cutting et al., 1991a; Gomez and Cutting, 1996; Hoa et al., 2002; Wakeley et al., 2000) (Fig. 5A). Recent results suggest that the pro-aKprocessing machinery is recruited to a protein complex that bridges the space between the two membranes during engulfment (Blaylock et al., 2004; Doan et al., 2005; Jiang et al., 2005; Rudner and Losick, 2002; Rudner et al., 2002), and it has been proposed that this synapse-like structure directly couples engulfment to aKactivation (Jiang et al., 2005). SpoIVFA is a key localization and assembly factor for the pro-aKprocessing machinery (Doan et al., 2005; Rudner and Losick, 2002), which includes the putative
21. B. SUBTILIS SPORULATION metalloprotease, SpoIVFB, believed to cleave pro-aK(Lu et al., 1995; Rudner et al., 1999; Yu and Kroos, 2000; Zhou and Kroos, 2004), and BofA, the primary inhibitor of SpoIVFB (Zhou and Kroos, 2004). It appears that SpoIVB cleaves SpoIVFA (Campo and Rudner, 2006; Dong and Cutting, 2003; Zhou and Kroos, 2005), and then BofA is cleaved by a second serine protease in the cascade, CtpB (Zhou and Kroos, 2005) (Fig. SB), relieving inhibition of SpoIVFB so that it can cleave pro-aK (Fig. SC). aKreleased from the OFM into the mother cell directs transcription of genes whose products help form the spore coat and eventually lyse the mother cell to release the spore (Eichenberger et al., 2004; Steil et al., 2005). Coupling between forespore events and mother cell processing of pro-aKwas discovered long ago (Cutting et al., 1990; Lu et al., 1990), but only recently has evidence emerged that pro-oKprocessing is both directly and indirectly coupled to engulfment (Jiang et al., 2005). The indirect mechanism is better understood. As described above, aGRNAP becomes active in the forespore upon completion of engulfment. This boosts transcription of spoIVB (Cutting et al., 1991b), which encodes a serine protease (Hoa et al., 2002; Wakeley et al., 2000) that appears to cleave SpoIVFA (Campo and Rudner, 2006; Dong and Cutting, 2003; Zhou and Kroos, 2005), triggering a proteolytic cascade that leads to aKactivity (Zhou and Kroos, 2005) (Fig. 5 ) . A direct mechanism coupling engulfment to pro-oK processing is inferred from the finding that a bofA null mutation, which eliminates the primary inhibitor of the apparent processing enzyme, SpoIVFB, allows processing in the absence of uGand SpoIVB (i.e., bypasses the indirect mechanism) but does not allow processing in the absence of engulfment proteins (SpoIID, SpoIIM, and SpoIIP) or proteins that interact in the space between the two membranes during engulfment (SpoIIQ and SpoIIIAH) (Jiang et al., 2005). These proteins are required to properly localize GFP-SpoIVFA (Doan et al., 2005) and SpoIVFB-GFP (Jiang et al., 2005) during engulfment. Moreover, BofA is required for proper localization of GFP-SpoIVFA (Doan et al., 2005; Rudner and Losick, 2002) and both BofA and SpoIVFA are needed for GFP-SpoIIQ to properly track the engulfing mother cell membrane (Jiang et al., 2005). Yet there is no evidence of direct interaction between engulfment proteins or the SpoIIQ-SpoIIIAH complex and the SpoIVFA-BofASpoIVFB complex, and careful analysis of the localization results suggests that an additional component(s) is involved (Doan et al., 2005; Jiang et al., 2005) (Protein X in Fig. 5A). The direct mechanism coupling pro-oKprocessing (as well as perhaps aGactivation) to completion
3 75 of engulfment has been proposed to involve the assembly of a synapse-like protein complex bridging the space between the two membranes (Jiang et al., 2005). The protein complex minimally would include the proteins shown in the OFM of Fig. 5A and SpoIIQ in the inner forespore membrane (IFM).SpoIID, SpoIIM, and SpoIIP would be required for proper localization of the complex during engulfment. Upon completion of engulfment, the low level of SpoIVB synthesized under aFcontrol is sufficient to cause proteolysis of the SpoIIQ domain in the space between the forespore membranes (Jiang et al., 2005). This domain is similar to the C-terminal domain of SpoIVFA (Rudner and Losick, 2002), which is cleaved by SpoIVB (Campo and Rudner, 2006; Dong and Cutting, 2003; Zhou and Kroos, 2005). Recently, it was shown that SporVB cleaves SpoIIQ, although this was not required for activation of ac or aK,or for spore formation (Chiba et al., 2007) (Fig. 5A). However, completion of engulfment did release SpoIIQ from an immobile complex, so it was proposed that membrane fusion-dependent reorganization of the complex might be the mechanism that directly couples activation of ac and aKto completion of engulfment. To simplify studies of the pro-aKprocessing machinery and its regulation, components have been expressed in various combinations in E. coli under the control of T7 RNAP. Coexpression of SpoIVFB and pro-aKallowed accurate and abundant processing (Zhou and Kroos, 2004), confirming the results of an earlier study (Lu et al., 1995) and supporting the idea, based on mutational analyses of spoIVFB and sequence comparisons (Rudner et al., 1999; Yu and Kroos, 2000), that SpoIVFB is a metalloprotease that carries out regulated intramembrane proteolysis (RIP) of pro-aK. RIP involves cleavage of a protein within a membrane or near the membrane surface. SpoIVFB provided the first bacterial example of RIP, which has emerged as an important and widely conserved mechanism that controls signaling pathways in both prokaryotes and eukaryotes (reviewed in Brown et al., 2000, and Wolfe and Kopan, 2004). Coexpressing BofA with SpoIVFB and pro-aKin E. coli markedly inhibited RIP of pro-aK, identifying BofA as the primary inhibitor of SpoIVFB (Zhou and Kroos, 2004). Adding SpoIVFA to the system completely inhibited RIP, and adding a fifth component, the SpoIVB serine protease normally made in the forespore, partially restored RIP in E. coli (Zhou and Kroos, 2005). SpoIVB has a signal sequence capable of translocating alkaline phosphatase to the E. coli periplasm (Wakeley et al., ZOOO), so presumably SpoIVB acts from the periplasm. SpoIVB appears to target SpoIVFA in E. coli and in sporulating
3 76 B. subtilis (Campo and Rudner, 2006; Zhou and Kroos, ZOOS), in agreement with in vitro studies (Campo and Rudner, 2006; Dong and Cutting, 2003) (Fig. 5A). The finding that RIP of pro-uKis delayed by about 1h in a ctpB mutant identified another regulator (Pan et al., 2003). CtpB is a serine protease made under uEcontrol in the mother cell (Pan et al., 2003) and under uGcontrol in the forespore (Steil et al., 2005; Wang et al., 2006) (Fig. 5B). Experiments in sporulating B. subtilis, in E. coli, and in vitro provide evidence that CtpB cleaves BofA near its C terminus, after SpoIVB has cleaved SpoIVFA (Zhou and Kroos, 2005). CtpB can also cleave SpoIVFA (Campo and Rudner, 2006). However, neither cleavage by CtpB is essential for RIP of pro-aK,since appearance of aKis delayed but not abolished in a ctpB mutant (Pan et al., 2003). CtpB cleavage of SpoIVFA is partly, if not fully, redundant with SpoIVB cleavage of SpoIVFA (Campo and Rudner, 2006). A backup mechanism to cleave BofA also appears to exist since loss of BofA correlated with the onset of pro-oKRIP in wild-type or ctpB mutant cells (Zhou and Kroos, 2005). How does SpoIVFB cleave pro-oKwithin a membrane or near its surface? The exact mechanism by which any intramembrane-cleaving protease (I-CLIP) recognizes its substrate and bends or unfolds it (transmembrane domains are typically or-helical, making peptide bonds inaccessible for cleavage) for peptide bond hydrolysis is unknown, but is of great interest since I-CLiPs govern pathways involved in human disease (Brown et al., 2000; Lichtenthaler and Steiner, 2007; Wolfe and Kopan, 2004). Pro-uKis cleaved after amino acid 20 in sporulating B. subtilis (Kroos et al., 1989) and in E. coli coexpressing SpoIVFB (Zhou and Kroos, 2004). The prosequence is necessary for membrane association in sporulating B. subtilis (Zhang et al., 1998) and amino acids 1 through 27 are sufficient for membrane localization of a GFP chimera; however, a distal region between amino acids 109 and 117is required for RIP (Prince et al., 2005). SpoIVFB does not measure the distance from the N terminus of pro-uKto the cleavage site, since addition or deletion of five amino acids near the N terminus did not alter the site of cleavage, but a lysine at position 1 3 is important for RIP (Prince et al., 2005). RIP of pro-uK by SpoIVFB provides an excellent model to understand I-CLIP function.
THE uKREGULON Once active aKis released from the OFM, the pattern of transcription in the mother cell changes rapidly. A clever method was devised to test whether transcription of a uE-dependentgene and a aK-dependentgene occurs simul-
ANALOGOUS SYSTEMS taneously in the same mother cell, and little or no overlap was found (Li and Piggot, 2001). uKRNAP activity negatively regulates transcription of the sigE gene (Zhang and Kroos, 1997; Zhang et al., 1999), and uKdisplaces uE from core RNAP (Ju et al., 1999),presumably hastening the switch from transcription of the oEregulon to transcription of the uKregulon. On the other hand, about 30 genes are members of both regulons (Eichenberger et al., 2004; Steil et al., 2005), either being transcribed from separate aE-and uK-dependent promoters or from the same promoter recognized by both holoenzymes. Based on microarray analyses, the uK regulon is smaller than the uE regulon, with estimates in good agreement, ranging from 132 to 144 genes (Eichenberger et al., 2004; Steil et al., 2005). Genes in the oKregulon are less highly conserved among endospore-forming bacteria than genes in the uE regulon, perhaps because more genes in the uKregulon encode components of the outer surface of the spore and these undergo greater evolutionary adaptation to the organism’s environment (Eichenberger et al., 2004). In addition to encoding many proteins of the outer coat layer (Kim et al., 2006), genes in the uKregulon encode enzymes that synthesize polysaccharides found in the coat, cell wall hydrolases that lyse the mother cell, and proteins necessary for the spore to germinate (Eichenberger et al., 2004; Steil et al., 2005). Of the germination proteins, GerE is a transcription factor that negatively regulates 55 genes in the oK regulon and positively regulates 48 genes (36 of these depend strongly upon GerE) (Eichenberger et al., 2004). GerE temporally diversifies the pattern of gene expression in the uKregulon (Eichenberger et al., 2004; Steil et al., 2005), playing a role akin to that of SpoIIID in the oEregulon. GerE can directly activate or repress transcription (Ichikawa et al., 1999; Ichikawa and Kroos, 2000; Zhang et al., 1994; Zheng et al., 1992), and its structure (Ducros et al., 2001) and two regions required for transcriptional activation (Crater and Moran, 2002) are known.
CONCLUSIONS AND FUTURE DIRECTIONS B. subtilis exhibits a number of multicellular behaviors. Among these, biofilm formation and swarming motility share similarities with M. xanthus fruiting body development and social motility, although there are clear differences as well. It will be interesting to compare these behaviors as molecular insights continue to emerge. The best-characterized multicellular behaviors of B. subtilis, sporulation and the development of competence to take up exogenous DNA, are regulated by extracellular
21. B. SUBTILIS SPORULATION peptide signaling, analogous to M . xanthus A-signaling (see chapter 3). A multitude of intracellular signals also influence the initiation of sporulation, feeding into a network of regulators in which SpoOA, uH, AbrB, Cody, and the Soj-SpoOJ system are key nodes. In M . xanthus, (p)ppGppis an early intracellular signal of starvation (see chapter 3 ) and uD RNAP, d4RNAP, and as4-activator proteins are key early regulators (see chapter 9), and undoubtedly, important signals and regulators remain to be discovered. As B. subtilis sporulation progresses, SpoOA continues to regulate genes, temporally and spatially. FruA, responding to C-signaling, seems to play a similar role as M . xanthus development proceeds (see chapters 4 and 9). The strategy for regulating genes in the forespore and mother cell after polar septation of B. subtilis is clear. Polar division triggers activation of aFin the forespore, and this rapidly signals activation of aEin the mother cell. Engulfment ensues, and its completion leads to activation of acin the forespore, generating signals that permit activation of uKin the mother cell. In addition to these interconnected u cascades, at least five auxiliary transcription factors (RsfA and SpoVT in the forespore and SpoIIID, GerR, and GerE in the mother cell) diversify the patterns of gene expression by up- or down-regulating particular genes. An analogous global picture of the M . xanthus gene regulatory network is just beginning to emerge (see chapter 9). Clearly, contact-dependent Csignaling is an important morphogenetic input (see chapter 4). The mechanism of this input is likely somewhat different from the input mechanisms of polar septation and engulfment completion that couple B. subtilis morphogenesis to developmental gene expression. Further elucidating these mechanisms is an important goal of future research. Although the detailed mechanisms of signaling and gene regulation surely differ in some respects between organisms, concepts learned from studies of cell-typespecific u factor activation in B. subtilis provide paradigms for myxobacteria and their neighbors. Regulation of uF activity involves multiple mechanisms that are partially redundant, including localization of the SpoIIE phosphatase to the polar septum, formation of a stable complex that sequesters the SpoIIAB anti-u, and transient genetic asymmetry together with susceptibility of SpoIIAB to proteolysis. At least one gene in the uF regulon, spoIIR, must be present in the origin-proximal region of the chromosome so its product is made quickly. SpoIIR appears to signal a membrane-embedded protease to cleave pro-aE, but the mechanism of signaling is unknown. Preferential expression of pro-uE in the mother cell and degradation of pro-uE/uEin the forespore
3 77
ensure compartmentalized activation. Again, multiple mechanisms that are partially redundant seem to be in place. Products of the uEregulon prevent a second polar septum from forming in the mother cell, commit it to sporulation even if nutrients become available, and drive engulfment, all by mechanisms not fully understood. Neither is it clear how completion of engulfment leads to activation of uGin the forespore and uKin the mother cell, although engulfment appears to properly localize a protein complex that includes SpoIIQ and SpoIIIAH bridging the space between the IFM and OFM, and the inhibited SpoIVFA-BofA-SpoIVFB complex in the OFM, where it is poised to process pro-aK. A cascade of proteolytic cleavages involving serine proteases SpoIVB and CtpB in the space between the membranes surrounding the forespore culminates in RIP of pro-uKby SpoIVFB. How I-CLiPs like SpoIVFB recognize their substrates and how they release them from membranes are fundamental questions relevant to human health, since I-CLiPs are involved in crucial signaling and gene regulatory pathways. Proteolysis is already known to play roles in A-, B-, and C-signaling during M. xanthus development (see chapters 3 and 4), and it seems likely that many more roles will be uncovered, based on studies of B. subtilis. Just as these studies have provided a host of paradigms, so too will continued investigation of the myxobacteria and their neighbors continue to yield novel insights of medical, economic, and environmental benefit. We are grateful to R. Zhou, P.Himes, L. Wang, and D. Imamura for helpful comments on the manuscript and to ]. Clarkson and M. Yudkin for sharing results prior to publication. Research on B. subtilis in the lab of L.K. was supported by N I H grant GM43585, and L.K. was supported in part by the Michigan Agricultural Experiment Station. Research in the lab of P.J.P. was supported by NIH grant GM43577. Research in the lab of C.P.M. was supported by N I H grant GM54395.
References Abanes-De Mello, A., Y. L. Sun, S. Aung, and K. Pogliano. 2002. A cytoskeleton-like role for the bacterial cell wall during engulfment of the Bacillus subtilis forespore. Genes Dev. 16:3253-3 264. Alper, S., L. Duncan, and R. Losick. 1994. An adenosine nucleotide switch controlling the activity of a cell type-specific transcription factor in B. subtilis. Cell 77195-205. Arabolaza, A. L., A. Nakamura, M. E. Pedrido, L. Martelotto, L. Orsaria, and R. R. Grau. 2003. Characterization of a novel inhibitory feedback of the anti-anti-sigma SpoIIAA on SpoOA activation during development in Bacillus subtilis. Mol. Microbiol. 471251-1263. Arigoni, F., K. Pogliano, C. D. Webb, P. Stragier, and R. Losick. 1995. Localization of protein implicated in establishment of cell type to sites of asymmetric division. Science 270:637640.
3 78 Arigoni, F., A. M. Guerout-Fleury, I. Barak, and P. Stragier. 1999. The SpoIIE phosphatase, the sporulation septum and the establishment of forespore-specifictranscription in Bacillus subtilis: a reassessment. Mol. Microbiol. 31:1407-1415. Bagyan, I., J. Hobot, and S. Cutting. 1996. A compartmentalized regulator of developmental gene expression in Bacillus subtilis. J. Bacteriol. 178:45004507. Barak, I., and A. J. Wilkinson. 2005. Where asymmetry in gene expression originates. Mol. Microbiol. 5 7 6 11-620. Bath, J., L. J. Wu, J. Errington, and J. C. Wang. 2000. Role of Bacillus subtilis SpoIIIE in DNA transport across the mother cell-prespore division septum. Science 290:995-997. Ben-Yehuda, S., and R. Losick. 2002. Asymmetric cell division in B. subtilis involves a spiral-like intermediate of the cytokinetic protein FtsZ. Cell 109:257-266. Ben-Yehuda, S., D. Z. Rudner, and R. Losick. 2003. Assembly of the SpoIIIE DNA translocase depends on chromosome trapping in Bacillus subtilis. Curr. Biol. 13:2196-2200. Ben-Yehuda, S., M. Fujita, X. S. Liu, B. Gorbatyuk, D. Skoko, J. Yan, J. F. Marko, J. S. Liu, P. Eichenberger, D. Z. Rudner, and R. Losick. 2005. Defining a centromere-like element in Bacillus subtilis by identifying the binding sites for the chromosome-anchoring protein RacA. Mol. Cell 17:773-782. Blaylock, B., X. Jiang, A. Rubio, C. P. Moran, Jr., and K. Pogliano. 2004.Zipper-like interaction between proteins in adjacent daughter cells mediates protein localization. Genes Dev. 18:2916-2928. Branda, S. S., J. E. Gonzalez-Pastor, S. Ben-Yehuda, R. Losick, and R. Kolter. 2001. Fruiting body formation by Bacillus subtilis. Proc. Natl. Acad. Sci. USA 98:11621-11626. Branda, S. S., J. E. Gonzalez-Pastor, E. Dervyn, S. D. Ehrlich, R. Losick, and R. Kolter. 2004. Genes involved in formation of structured multicellular communities by Bacillus subtilis. J. Bacteriol. 186:39 70-3979. Branda, S. S., F. Chu, D. B. Kearns, R. Losick, and R. Kolter. 2006. A major protein component of the Bacillus subtilis biofilm matrix. Mol. Microbiol. 59:1229-1238. Britton, R. A., l?Eichenberger,J. E. Gonzalez-Pastor,P. Fawcett, R. Monson, R. Losick, and A. D. Grossman. 2002. Genomewide analysis of the stationary-phase sigma factor (sigma-H) regulon of Bacillus subtilis. J. Bacteriol. 184:488 1 4 8 9 0 . Brown, M. S., J. Ye, R. B. Rawson, and J. L. Goldstein. 2000. Regulated intramembrane proteolysis: a control mechanism conserved from bacteria to humans. Cell 100:391-398. Burbulys, D., K. A. Trach, and J. A. Hoch. 1991. Initiation of sporulation in B. subtilis is controlled by a multicomponent phosphorelay. Cell 64545-552. Burkholder, W. F., and A. D. Grossman. 2000. Regulation of the initiation of endospore formation in Bacillus subtilis, p. 151-166. In Y. V. Brun and L. J. Shimkets (ed.), Prokaryotic Development. ASM Press, Washington, DC. Burkholder, W. F., I. Kurtser, and A. D. Grossman. 2001. Replication initiation proteins regulate a developmental checkpoint in Bacillus subtilis. Cell 104:269-279. Calvio, C., F. Celandroni, E. Ghelardi, G. Amati, S. Salvetti, F. Ceciliani, A. Galizzi, and S. Senesi. 2005. Swarming differentiation and swimming motility in Bacillus subtilis are controlled by swrA, a newly identified dicistronic operon. J. Bacteriol. 1875356-5366.
ANALOGOUS SYSTEMS Campbell, E. A., and S. A. Darst. 2000. The anti-sigma factor SpoIIAB forms a 2:l complex with oF,contacting multiple conserved regions of the sigma factor.]. Mol. Biol. 300:1728. Campo, N., and D. Z. Rudner. 2006. A branched pathway governing the activation of a developmental transcription factor by regulated intramembrane proteolysis. Mol. Cell 23:25-35. Carlson, H. C., S. Lu, L. Kroos, and W. G. Haldenwang. 1996. Exchange of precursor-specificelements between Pro-& and Pro-oKof Bacillus subtilis. 1. Bacteriol. 178546-549. Carniol, K., P. Eichenberger, and R. Losick. 2004. A threshold mechanism governing activation of the developmental regulatory protein uF in Bacillus subtilis. J. Biol. Chem. 279:14860-14870. Chagneau, C., and M. H. Saier, Jr. 2004. Biofilm-defective mutants of Bacillus subtilis. J. Mol. Microbiol. Biotechnol. 8:177-1 88. Chary, V. K., D. W. Hilbert, M. L. Higgins, and P. J. Piggot. 2000. The putative DNA translocase SpoIIIE is required for sporulation of the symmetricallydividing coccal species Sporosarcina ureae. Mol. Microbiol. 35:612-622. Chary, V. K., M. Meloni, D. W. Hilbert, and P. J. Piggot. 2005. Control of the expression and compartmentalization of uG activity during sporulation of Bacillus subtilis by regulators of uFand uE.J. Bacteriol. 187:6832-6840. Chiba, S., K. Coleman, and K. Pogliano. 2007. Impact of membrane fusion and proteolysis on SpoIIQ dynamics and interaction with SpoIIIAH.J. Biol. Chem. 282:2576-2586. Chu, F., D. B. Kearns, S. S. Branda, R. Kolter, and R. Losick. 2006. Targets of the master regulator of biofilm formation in Bacillus subtilis. Mol. Micro biol. 5 9: 1216-1 22 8. Chung, J. D., G. Stephanopoulos, K. Ireton, and A. D. Grossman. 1994. Gene expression in single cells of Bacillus subtilis: evidence that a threshold mechanism controls the initiation of sporulation. J. Bacteriol. 176:1977-1984. Clarkson, J., I. D. Campbell, and M. D. Yudkin. 2004. Efficient regulation of crF, the first sporulation-specific sigma factor in B. subtilis. J. Mol. Biol. 342:1187-1195. Connelly, M. B., G. M. Young, and A. Sloma. 2004. Extracellular proteolytic activity plays a central role in swarming motility in Bacillus subtilis. J. Bacteriol. 186:4159-4167. Crater, D. L., and C. P. Moran, Jr. 2002. Two regions of GerE required for promoter activation in Bacillus subtilis. J. Bacteriol. 184:241-249. Cutting, S., V. Oke, A. Driks, R. Losick, S. Lu, and L. Kroos. 1990. A forespore checkpoint for mother-cell gene expression during development in Bacillus subtilis. Cell 62:239-250. Cutting, S., A. Driks, R. Schmidt, B. Kunkel, and R. Losick. 1991a. Forespore-specifictranscription of a gene in the signal transduction pathway that governs pro-oKprocessing in Bacillus subtilis. Genes Dev.5:456466. Cutting, S., S. Roels, and R. Losick. 1991b. Sporulation operon sPoIVF and the characterization of mutations that uncouple mother-cell from forespore gene expression in Bacillus subtilis. J. Mol. Biol. 221:1237-1256. Diederich, B., J. F. Wilkinson, T. Magnin, S. M. A. Najafi, J. Errington, and M. D. Yudkin. 1994. Role of interactions between SpoIIAA and SpoIIAB in regulating cell-specific
21. B. SUBTILIS SPORULATION transcription factor uFof Bacillus subtilis. Genes Dev. 8: 2653-2663. Dixit, M., C. S. Murudkar, and K. K. Rao. 2002. epr is transcribed from a uDpromoter and is involved in swarming of Bacillus subtilis. J. Bacteriol. 184596-599. Doan, T., K. A. Marquis, and D. Z. Rudner. 2005. Subcellular localization of a sporulation membrane protein is achieved through a network of interactions along and across the septum. Mol. Microbiol. 55:1767-1781. Dong, T. C., and S. M. Cutting. 2003. SpoIVB-mediated cleavage of SpoIVFA could provide the intercellular signal to activate processing of Pro-uKin Bacillus subtilis. Mol. Microbiol. 49:1425-1434. Driks, A. 2002. Maximum shields: the assembly and function of the bacterial spore coat. Trends Microbiol. 10:251-254. Ducros, V. M., R. J. Lewis, C. S. Verma, E. J. Dodson, G. Leonard., J. P. Turkenburg, G. N. Murshudov, A. J. Wilkinson, and J. A. Brannigan. 2001. Crystal structure of GerE, the ultimate transcriptional regulator of spore formation in Bacillus subtilis. J. Mol. Biol. 306:759-771. Duncan, L., S. Alper, F. Arigoni, R. Losick, and P. Stragier. 1995. Activation of cell-specific transcription by a serine phosphatase at the site of asymmetric division. Science 270:641-644. Duncan, L., S. Alper, and R. Losick. 1996. SpoIIAA governs the release of the cell-type specific transcription factor cFfrom its anti-sigma factor SpoIIAB. J. Mol. Biol. 260:147-164. Dworkin, J., and R. Losick. 2001. Differential gene expression governed by chromosomal spatial asymmetry. Cell 107:339346. Dworkin, J. 2003. Transient genetic asymmetry and cell fate in a bacterium. Trends Genet. 19:107-112. Dworkin, J., and R. Losick. 2005. Developmental commitment in a bacterium. Cell 121:401-409. Dworkin, M., K. H. Keller, and D. Weisberg. 1983. Experimental observations consistent with a surface tension model of gliding motility of Myxococcus xanthus. J. Bacteriol. 155:1367-1371. Eichenberger, P., P. Fawcett, and R. Losick. 2001. A threeprotein inhibitor of polar septation during sporulation in Bacillus subtilis. Mol. Microbiol. 42:1147-1162. Eichenberger, P., S. T. Jensen, E. M. Conlon, C. van Ooij, J. Silvaggi, J. E. Gonzalez-Pastor, M. Fujita, S. Ben-Yehuda, P. Stragier, J. S. Liu, and R. Losick. 2003. The uEregulon and the identification of additional sporulation genes in Bacillus subtilis. J. Mol. Biol. 322945-972. Eichenberger, P., M. Fujita, S. T. Jensen, E. M. Conlon, D. Z. Rudner, S. T. Wang, C. Ferguson, K. Haga, T. Sato, J. S. Liu, and R. Losick. 2004. The program of gene transcription for a single differentiating cell type during sporulation in Bacillus subtilis. PLoS Biol. 2:1664-1683. Ellermeier, C. D., E. C. Hobbs, J. E. Gonzalez-Pastor, and R. Losick. 2006. A three-protein signaling pathway governing immunity to a bacterial cannibalism toxin. Cell 124549559. Errington, J., L. Appleby, R. A. Daniel, H. Goodfellow, S. R. Partridge, and M. D. Yudkin. 1992. Structure and function of the spoIIIJ gene of Bacillus subtilis: a vegetatively expressed gene that is essential for uc activity at an intermediate stage of sporulation. J. Gen. Microbiol. 138:2609-2618.
3 79 Errington, J. 2003. Regulation of endospore formation in Bacillus subtilis. Nut. Rev. Microbiol. 1:117-126. Fawcett, P., A. Melnikov, and P. Youngman. 1998. The Baciflus SpoIIGA protein is targeted to sites of spore septum formation in a SpoIIE-independent manner. Mol. Microbiol. 28:931-943. Feucht, A., L. Abbotts, and J. Errington. 2002. The cell differentiation protein SpoIIE contains a regulatory site that controls its phosphatase activity in response to asymmetric septation. Mol. Microbiol. 45:1119-1130. Feucht, A., L. Evans, and J. Errington. 2003. Identification of sporulation genes by genome-wide analysis of the uEregulon of Bacillus subtilis. Microbiology 149:3023-3034. Frandsen, N., and P. Stragier. 1995. Identification and characterization of the Bacillus subtilis spoIIP locus. J. Bacteriol. 172716-722. Frandsen, N., I. Barak, C. Karmazyn-Campelli, and P. Stragier. 1999. Transient gene asymmetry during sporulation and establishment of cell specificity in Bacillus subtilis. Genes Dev. 13:394-399. Fujita, M., and R. Losick. 2002. An investigation into the compartmentalization of the sporulation transcription factor uE in Bacillus subtilis. Mol. Microbiol. 43:27-38. Fujita, M., and R. Losick. 2003. The master regulator for entry into sporulation in Bacillus subtilis becomes a cell-specific transcription factor after asymmetric division. Genes Dev. 17:1166-1 174. Fujita, M., J. E. Gonzalez-Pastor, and R. Losick. 2005. Highand low-threshold genes in the SpoOA regulon of Bacillus subtilis. J. Bacteriol. 1821357-1368. Gholamhoseinian, A., and P. J. Piggot. 1989. Timing of spoII gene expression relative to septum formation during sporulation of Bacillus subtilis. J. Bacteriol. 1715747-5749. Gomez, M., and S. M. Cutting. 1996. Expression of the Bacillus subtilis spoIVB gene is under dual oF/uCcontrol. Microbiology 142:3453-3457. Gomez, M., and S. M. Cutting. 1997. 6ofC encodes a putative forespore regulator of the Bacillus subtilis uK checkpoint. Microbiology 143:157-170. Gonzalez-Pastor, J. E., E. C. Hobbs, and R. Losick. 2003. Cannibalism by sporulating bacteria. Science 301510-513. Halberg, R., and L. Kroos. 1994. Sporulation regulatory protein SpoIIID from Bacillus subtilis activates and represses transcription by both mother-cell-specific forms of RNA polymerase. J. Mol. Biol. 243:425-436. Hamon, M. A., and B. A. Lazazzera. 2001. The sporulation transcription factor SpoOA is required for biofilm development in Bacillus subtilis. Mol. Microbiol. 42:1199-1209. Hamon, M. A., N. R. Stanley, R. A. Britton, A. D. Grossman, and B. A. Lazazzera. 2004. Identification of AbrB-regulated genes involved in biofilm formation by Bacillus subtilis. Mol. Microbiol. 52:847-860. Helmann, J. D., and C. P. Moran, Jr. 2002. RNA polymerase and sigma factors, p. 289-312. In A. L. Sonenshein, J. A. Hoch, and R. Losick (ed.), Bacillus subtilis and Its Closest Relatives: from Genes to Cells. ASM Press, Washington, DC. Hilbert, D. W., and P. J. Piggot. 2003. Novel spoIIE mutation that causes uncompartmentalized uFactivation in Bacillus subtilis. J. Bacteriol. 185: 1590-1 59 8.
380 Hilbert, D. W., V. K. Chary, and P. J. Piggot. 2004. Contrasting effects of uEon compartmentalization of uF activity during sporulation of Bacillus subtilis. J. Bacteriol. 186:19831990. Hilbert, D. W., and P. J. Piggot. 2004. Compartmentalization of gene expression during Bacillus subtilis spore formation. Microbiol. Mol. Biol. Rev. 68:234-262. Hoa, N. T., J. A. Brannigan, and S. M. Cutting. 2002. The Bacillus subtilis signaling protein SpoIVB defines a new family of serine peptidases. J. Bacteriol. 184:191-199. Hofmeister, A. 1998. Activation of the proprotein transcription factor Pro-uEis associated with three changes in its subcellular localization during sporulation in Bacillus subtilis. J. Bacteriol. 180:2426-2433. Hofmeister, A. E. M., A. Londono-Vallejo, E. Harry, P. Stragier, and R. Losick. 1995. Extracellular signal protein triggering the proteolytic activation of a developmental transcription factor in B. subtilis. Cell 83:219-226. Iber, D., J. Clarkson, M. D. Yudkin, and I. D. Campbell. 2006. The mechanism of cell differentiation in Bacillus subtilis. Nature 441:371-374. Ichikawa, H., R. Halberg, and L. Kroos. 1999. Negative regulation by the Bacillus subtilis GerE protein. J. Biol. Chem. 274:8322-8327. Ichikawa, H., and L. Kroos. 2000. Combined action of two transcription factors regulates genes encoding spore coat proteins of Bacillus subtilis. ]. Biol. Chem. 275:13849-13855. Illing, N., and J. Errington. 1991. The spoIIIA operon of Bacillus subtilis defines a new temporal class of mother-cellspecific sporulation genes under the control of the uEform of RNA polymerase. Mol. Microbiol. 5:1927-1940. Jiang, X., A. Rubio, S. Chiba, and K. Pogliano. 2005. Engulfmentregulated proteolysis of SpoIIQ: evidence that dual checkpoints control sigma activity. Mol. Microbiol. 58:102-115. Jonas, R. M., E. A. Weaver, T. J. Kenney, C. P. Moran, Jr., and W. G. Haldenwang. 1988. The Bacillus subtilis spoIIG operon encodes both uEand a gene necessary for uE activation. J. Bacteriol. 170507-511. Ju, J., T. Luo, and W. Haldenwang. 1997. Bacillus subtilis prouEfusion protein localizes to the forespore septum and fails to be processed when synthesized in the forespore. J . Bacterial. 179:48 8 8-4893. Ju, J., T. Luo, and W. Haldenwang. 1998. Forespore expression and processing of the SigE transcription factor in wild-type and mutant Bacillus subtilis. J. Bacteriol. 180:1673-1681. Ju, J., T. Mitchell, H. K. Peters, and W. G. Holdenwang. 1999. Sigma factor displacement from RNA polymerase during Bacillus subtilis sporulation. J. Bacteriol. 18k4969-4977. Ju, J., and W. G. Haldenwang. 2003. Tethering of the Bacillus subtilis uEproprotein to the cell membrane is necessary for its processing but insufficient for its stabilization. J. Bacterial. 1855897-5900. Julien, B., A. D. Kaiser, and A. Garza. 2000. Spatial control of cell differentiation in Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 979098-9103. Karow, M. L., P. Glaser, and P. J. Piggot. 1995. Identification of a gene, spoIIR, that links the activation of uEto the transcriptional activity of uFduring sporulation in Bacillus subtilis. Proc. Natl. Acad. Sci. USA 92:2012-2016.
ANALOGOUS SYSTEMS Kearns, D. B., and R. Losick. 2003. Swarming motility in undomesticated Bacillus subtilis. Mol. Microbiol. 4 9 5 8 1-590. Kearns, D. B., F. Chu, R. Rudner, and R. Losick. 2004. Genes governing swarming in Bacillus subtilis and evidence for a phase variation mechanism controlling surface motility. Mol. Microbiol. 52:357-369. Kearns, D. B., F. Chu, S. S. Branda, R. Kolter, and R. Losick. 2005. A master regulator for biofilm formation by Bacillus subtilis. Mol. Microbiol. 55:739-749. Kellner, E. M., A. Decatur, and C. P. Moran, J . 1996. Two-stage regulation of an anti-sigma factor determines developmental fate during bacterial endospore formation. Mol. Microbiol. 21:913-924. Khvorova, A., L. Zhang, M. L. Higgins, and P. J. Piggot. 1998. The spoIIE locus is involved in the SpoOA-dependent switch in the location of FtsZ rings in Bacillus subtilis. J. Bacteriol. 180~1256-1260. Khvorova, A., V. K. Chary, D. W. Hilbert, and P. J. Piggot. 2000. The chromosomal location of the Bacillus subtilis sporulation gene spoIIR is important for its function. J. Bacteriol. 182:4425-4429. Kim, H., M. Hahn, P. Grabowski, D. C. McPherson, M. M. Otte, R. Wang, C. C. Ferguson, P. Eichenberger, and A. Driks. 2006. The Bacillus subtilis spore coat protein interaction network. Mol. Microbiol. 59:487-502. Kim, S. K., and D. Kaiser. 1991. C-factor has distinct aggregation and sporulation thresholds during Myxococcus development. 1. Bacteriol. 173:1722-1728. King, N., 0. Dreesen, P. Stragier, K. Pogliano, and R. Losick. 1999. Septation, dephosphorylation, and the activation of uF during sporulation in Bacillus subtilis. Genes Dev. 13:11561167. Kirchman, P. A., H. DeGrazia, E. M. Kellner, and C. 11. Moran, Jr. 1993. Forespore-specific disappearance of the sigma-factor antagonist SpoIIAB: implications for its role in determination of cell fate in Bacillus subtilis. Mol. Microbiol. 8:663-671. Kroos, L., A. Kuspa, and D. Kaiser. 1986. A global analysis of developmentally regulated genes in Myxococcus xanthus. Dev. Biol. 117252-266. Kroos, L., B. Kunkel, and R. Losick. 1989. Switch protein alters specificity of RNA polymerase containing a compartment-specific sigma factor. Science 243526-529. Kroos, L., and J. R.Maddock. 2003. Prokaryotic development: emerging insights. J. Bacteriol. 185:1128-1146. Kruse, T., S. Lobedanz, N. M. Berthelsen, and L. SsgaardAndersen. 2001. C-signal: a cell surface-associated morphogen that induces and co-ordinates multicellular fruiting body morphogenesis and sporulation in Myxococcus xanthus. Mol. Microbiol. 40: 156-1 68. LaBell, T. L., J. E. Trempy, and W. G. Haldenwang. 1987. Sporulation-specific sigma factor, u29,of Bacillus subtilis is synthesized from a precursor protein, P3*.Proc. Natl. Acad. Sci. USA 84:1784-1788. Lazazzera, B. A., and A. D. Grossman. 1998. The ins and outs of peptide signaling. Trends Microbiol. 6:288-294. Levin, P. A., and R. Losick. 1996. Transcription factor SpoOA switches the localization of the cell division protein FtsZ from a medial to a bipolar pattern in Bacillus subtilis. Genes Dev. 10:478-488.
21. B. SUBTILIS SPORULATION Levin, P. A., R. Losick, P. Stragier, and F. Arigoni. 1997. Localization of the sporulation protein SpoIIE in Bacillus subtilis is dependent upon the cell division protein FtsZ. Mol. Microbiol. 25:839-846. Li, Z., and P. J. Piggot. 2001. Development of a two-part transcription probe to determine the completeness of temporal and spatial compartmentalization of gene expression during bacterial development. Proc. Natl. Acad. Sci. USA 98~12538-12543. Lichtenthaler, S . F., and H. Steiner. 2007. Sheddases and intramembrane-cleaving proteases: RIPpers of the membrane. Symposium on Regulated Intramembrane Proteolysis. EMBO Rep. 8537-541. Liu, J., and P. Zuber. 2000. The ClpX protein of Bacillus subtilis indirectly influences RNA polymerase holoenzyme composition and directly stimulates aH-dependenttranscription. Mol. Microbiol. 37:885-897. Londono-Vallejo, J. A., and P. Stragier. 1995. Cell-cell signaling pathway activating a developmental transcription factor in Bacillus subtilis. Genes Dev. 9503-508. Londono-Vallejo, J. A., C. Frehel, and P. Stragier. 1997. SpoIIQ, a forespore-expressed gene required for engulfment in Bacillus subtilis. Mol. Microbiol. 24:29-39. Lu, S., R. Halberg, and L. Kroos. 1990. Processing of the mother-cell a factor, aK,may depend on events occurring in the forespore during Bacillus subtilis development. Proc. Natl. Acad. Sci. USA 87:9722-9726. Lu, S., S. Cutting, and L. Kroos. 1995. Sporulation protein SpoIVFB from Bacillus subtilis enhances processing of the sigma factor precursor pro-aKin the absence of other sporulation gene products. J. Bacteriol. 177:1082-1085. Lucet, I., A. Feucht, M. D. Yudkin, and J. Errington. 2000. Direct interaction between the cell division protein FtsZ and the cell differentiation protein SpoIIE. EMBO J. 19:1467-1475. Maamar, H., and D. Dubnau. 2005. Bistability in the Bacillus subtilis K-state (competence) system requires a positive feedback loop. Mol. Microbiol. 56:615-624. Magnin, T., M. Lord, and M. D. Yudkin. 1997. Contribution of partner switching and SpoIIAA cycling to regulation of crF activity in sporulating Bacillus subtilis. J. Bacteriol. 179:3922-3927. Margolis, P., A. Driks, and R. Losick. 1991. Establishment of cell type by compartmentalized activation of a transcription factor. Science 2 5 4 5 6 2 4 6 5 . Masuda, E. S., H. Anaguchi, K. Yamada, and Y. Kobayashi. 1988. Two developmental genes encoding sigma factor homologs are arranged in tandem in Bacillus subtilis. Proc. Natl. Acad. Sci. USA 85:7637-7641. Masuda, E. S., H. Anaguchi, T. Sato, M. Takeuchi, and Y. Kobayashi. 1990. Nucleotide sequence of the sporulation gene spoIIGA from Bacillus subtilis. Nucleic Acids Res. 18:657. McBride, S. M., A. Rubio, L. Wang, and W. G. Haldenwang. 2005. Contributions of protein structure and gene position to the compartmentalization of the regulatory proteins aE and SpoIIE in sporulating Bacillus subtilis. Mol. Microbiol. 57~434-451. Mendelson, N. H. 1999. Bacillus subtilis macrofibres, colonies and bioconvection patterns use different strategies to achieve multicellular organization. Environ. Microbiol. 1:471-477.
381 Min, K., C. M. Hilditch, B. Diederich, J. Errington, and M. D. Yudkin. 1993. aF,the first compartment-specific transcription factor of Bacillus subtilis, is regulated by an anti-a factor that is also a protein kinase. Cell 74:735-742. Molle, V., M. Fujita, S. T.Jensen, P. Eichenberger, J. E. GonzalezPastor, J. S. Liu, and R. Losick. 2003. The SpoOA regulon of Bacillus subtilis. Mol. Microbiol. 50:1683-1701. Nicholson, W. L., N. Munakata, G. Horneck, H. J. Melosh, and P. Setlow. 2000. Resistance of Bacillus endospores to extreme terrestrial and extraterrestrial environments. Microbiol. Mol. Biol. Rev. 64548-572. O’Connor, K. A., and D. R. Zusman. 1988. Reexamination of the role of autolysis in the development of Myxococcus xanthus. J. Bacteriol. 170:4103-4112. O’Connor, K. A., and D. R. Zusman. 1991. Development in Myxococcus xanthus involves differentiation into two cell types, peripheral rods and spores. J. Bacteriol. 173:33183333. Okada, M., I. Sato, S. J. Cho, H. Iwata, T. Nishio, D. Dubnau, and Y. Sakagami. 2005. Structure of the Bacillus subtilis quorum-sensing peptide pheromone ComX. Nut. Chem. Biol. 1:23-24. Pan, Q., D. A. Garsin, and R. Losick. 2001. Self-reinforcing activation of a cell-specific transcription factor by proteolysis of an anti-sigma factor in B. subtilis. Mol. Cell 8:873883. Pan, Q., R. Losick, and D. Z. Rudner. 2003. A second PDZcontaining serine protease contributes to activation of the sporulation transcription factor aK in Bacillus subtilis. J. Bacteriol. 185:6051-6056. Partridge, S. R., and J. Errington. 1993. Importance of morphological events and intercellular interactions in the regulation of prespore-specific gene expression during sporulation in Bacillus subtilis. Mol. Microbiol. 8:945-955. Perego, M., and J. A. Brannigan. 2001. Pentapeptide regulation of aspartyl-phosphate phosphatases. Peptides 22:15411547. Perego, M., and J. A. Hoch. 2002. Two-component systems, phosphorelays, and regulation of their activities by phosphatases, p. 473-482. In A. L. Sonenshein, J. A. Hoch, and R. Losick (ed.), Bacillus subtilis and Its Closest Relatives: from Genes to Cells. ASM Press, Washington, DC. Peters, H. K., and W. G. Haldenwang. 1991. Synthesis and fractionation properties of SpoIIGA, a protein essential for Pro-oEprocessing in Bacillus subtilis. J. Bacteriol. 173:782 17827. Peters, H. K., H. C. Carlson, and W. G. Haldenwang. 1992. Mutational analysis of the precursor-specific region of Bacillus subtilis aE.J. Bacteriol. 174:4629-4637. Peters, H. K., and W. G. Haldenwang. 1994. Isolation of a Bacillus subtilis spoIIGA allele that suppresses processingnegative mutations in the Pro-aE gene (sigE).J. Bacteriol. 176~7763-7766. Phillips, Z. E., and M. A. Strauch. 2002. Bacillus subtilis sporulation and stationary phase gene expression. Cell. Mol. Life Sci. 59:392-402. Piggot, P. J., and R. Losick. 2002. Sporulation genes and intercompartmental regulation, p. 483-518. In A. L. Sonenshein, J. A. Hoch, and R. Losick (ed.), Bacillus subtilis
382 and Its Closest Relatives: from Genes to Cells. ASM Press, Washington, DC. Piggot, P. J., and D. W. Hilbert. 2004. Sporulation of Bacillus subtilis. Curr. Opin. Microbiol. 7579-586. Pogliano, J., N. Osborne, M. D. Sharp, A. Abanes-De Mello, A. Perez, Y. L. Sun, and K. Pogliano. 1999. A vital stain for studying membrane dynamics in bacteria: a novel mechanism controlling septation during Bacillus subtilis sporulation. Mol. Microbiol. 31:1149-1159. Pogliano, K., A. Hofmeister, and R. Losick. 1997. Disappearance of the uEtranscription factor from the forespore and the SpoIIE phosphatase from the mother cell contributes to establishment of cell-specific gene expression during sporulation in Bacillus subtilis. J. Bacteriol. 179:33313341. Prince, H., R. Zhou, and L. Kroos. 2005. Substrate requirements for regulated intramembrane proteolysis of Bacillus subtilis pro-uK.J. Bacteriol. 187:961-971. Rowland, S. L., W. F. Burkholder, K. A. Cunningham, M. W. Maciejewski, A. D. Grossman, and G . F. King. 2004. Structure and mechanism of action of Sda, an inhibitor of the histidine kinases that regulate initiation of sporulation in Bacillus subtilis. Mol. Cell 13:689-701. Rudner, D., P. Fawcett, and R. Losick. 1999. A family of membrane-embedded metalloproteases involved in regulated proteolysis of membrane-associated transcription factors. Proc. Natl. Acad. Sci. USA 96:14765-14770. Rudner, D. Z., and R. Losick. 2002. A sporulation membrane protein tethers the pro-uKprocessing enzyme to its inhibitor and dictates its subcellular localization. Genes Dev. 16~1007-1018. Rudner, D. Z., Q. Pan, and R. M. Losick. 2002. Evidence that subcellular localization of a bacterial membrane protein is achieved by diffusion and capture. Proc. Natl. Acad. Sci. USA 99:8701-8706. Sager, B., and D. Kaiser. 1993. Spatial restriction of cellular differentiation. Genes Dev. 7:1645-1653. Sanchez-Salas, J. L., M. L. Santiago-Lara, B. Setlow, M. D. Sussman, and P. Setlow. 1992. Properties of Bacillus megaterium and Bacillus subtilis mutants which lack the protease that degrades small, acid-soluble proteins during spore germination. J. Bacteriol. 174:807-814. Schmidt, R., P. Margolis, L. Duncan, R. Coppolecchia, Jr., C. P. Moran, and R. Losick. 1990. Control of developmental transcription factor uF by sporulation regulatory proteins SpoIIAA and SpoIIAB in Bacillus subtilis. Proc. Natl. Acad. Sci. USA 87:9221-9225. Senesi, S., E. Ghelardi, F. Celandroni, S. Salvetti, E. Parisio, and A. Galizzi. 2004. Surface-associatedflagellum formation and swarming differentiation in Bacillus subtilis are controlled by the ifm locus. J. Bacteriol. 186:1158-1164. Serrano, M., S. Hovel, C. P. Moran, Jr., A. 0. Henriques, and U. Volker. 2001. Forespore-specific transcription of the lonB gene during sporulation in Bacillus subtilis. J. Bacteriol. 183~2995-3003. Serrano, M., L. Corte, J. Opdyke, C. I?. Moran, Jr., and A. 0. Henriques. 2003. Expression of spoIIIJ in the prespore is sufficient for activation of uG and for sporulation in Bacillus subtilis. J. Bacteriol. 185:3905-3917.
ANALOGOUS SYSTEMS Serrano, M., A. Neves, C. M. Soares, C. P. Moran, Jr., and A. 0. Henriques. 2004. Role of the anti-sigma factor SpoIIAB in regulation of uG during Bacillus subtilis sporulation. J. Bacteriol. 186:4000-4013. Setlow, P. 2003. Spore germination. Curr. Opin. Microbiol. 6:550-556. Shcheptov, M., G. Chyu, I. Bagyan, and S. Cutting. 1997. Characterization of csgA, a new member of the forespore-expressed regulon from Bacillus subtilis. Gene 184:133-140. Smits, W. K., C. C. Eschevins, K. A. Susanna, S. Bron, 0. P. Kuipers, and L. W. Hamoen. 2005. Stripping Bacillus: ComK auto-stimulation is responsible for the bistable response in competence development. Mol. Microbiol. 56:604-614. Sonenshein, A. L., J. A. Hoch, and R. Losick. 1993. Bacillus subtilis and Other Gram-Positive Bacteria: Biochemistry, Physiology, and Molecular Genetics. ASM Press, Washington, DC. Sonenshein, A. L. 2000. Control of sporulation initiation in Bacillus subtilis. Curr. Opin. Microbiol. 3561-566. Sonenshein, A. L., J. A. Hoch, and R. Losick. 2002. Bacillus subtilis and Its Closest Relatives: from Genes to Cells. ASM Press, Washington, DC. Sonenshein, A. L. 2005. Cody, a global regulator of stationary phase and virulence in Gram-positive bacteria. Curr. Opin. Microbiol. 8:203-207. Stanley, N. R., R. A. Britton, A. D. Grossman, and B. A. Lazazzera. 2003. Identification of catabolite repression as a physiological regulator of biofilm formation by Bacillus subtilis by use of DNA microarrays.]. Bacteriol. 185:19511957. Steil, L., M. Serrano, A. 0. Henriques, and U. Volker. 2005. Genome-wideanalysis of temporally regulated and compartment-specific gene expression in sporulating cells of Bacillus subtilis. Microbiology 151:399-420. Stragier, P., C. Bonamy, and C. Karmazyn-Campelli. 1988. Processing of a sporulation sigma factor in Bacillus subtilis: how morphological structure could control gene expression. Cell 52:697-704. Stragier, P., B. Kunkel, L. Kroos, and R. Losick. 1989. Chromosomal rearrangement generating a composite gene for a developmental transcription factor. Science 243 507-5 12. Sun, D., P. Stragier, and I? Setlow. 1989. Identification of a new a-factor involved in compartmentalized gene expression during sporulation of Bacillus subtilis. Genes Dev. 3: 141-149. Sun, D., R. M. Cabrera-Martinez, and P. Setlow. 1991. Control of transcription of the Bacillus subtilis spoIIIG gene, which codes for the forespore-specific transcription factor a'. J. Bacteriol. 173:2977-2984. Sun, Y. L., M. D. Sharp, and K. Pogliano. 2000. A dispensable role for forespore-specificgene expression in engulfment of the forespore during sporulation of Bacillus subtilis. J. Bacteriol. 182:2919-2927. Tjalsma, H., S. Bron, and J. M. van Dijl. 2003. Complementary impact of paralogous Oxal-like proteins of Bacillus subtilis on post-translocational stages in protein secretion. J. Biol. Chem. 278:15622-15632. Tortosa, P., and D. Dubnau. 1999. Competence for transformation: a matter of taste. Curr. Opin. Microbiol. 2588-592.
21. B. SUBTlLIS
SPORULATION
Tovar-Rojo, F., M. Chander, B. Setlow, and P. Setlow. 2002. The products of the spoVA operon are involved in dipicolinic acid uptake into developing spores of Bacillus subtilis. J. Bacteriol. 184584-587. Veening, J. W., L. W. Hamoen, and 0. P. Kuipers. 2005. Phosphatases modulate the bistable sporulation gene expression pattern in Bacillus subtilis. Mol. Microbiol. 56:1481-1494. Wakeley, P. R., R. Dorazi, N. T. Hoa, J. R. Bowyer, and S. M. Cutting. 2000. Proteolysis of SpoIVB is a critical determinant in signalling of Pro-uKprocessing in Bacillus subtilis. Mol. Microbiol. 36:1336-1348. Wang, S. T., B. Setlow, E. M. Conlon, J. L. Lyon, D. Imamura, T. Sato, P. Setlow, R. Losick, and P. Eichenberger. 2006. The forespore line of gene expression in Bacillus subtilis. J . Mol. Biol. 358:16-37. Wireman, J. W., and M. Dworkin. 1975. Morphogenesis and developmental interactions in myxobacteria. Science 189516-523. Wolfe, M. S., and R. Kopan. 2004. Intramembrane proteolysis: theme and variations. Science 305:1119-1123. Wu, L. J., and J. Errington. 1994. Bacillus subtilis SpoIIIE protein required for DNA segregation during asymetric cell division. Science 264572-575. Wu, L. J., P. J. Lewis, R. Allmansberger, P. M. Hauser, and J. Errington. 1995. A conjugation-like mechanism for prespore chromosome partitioning during sporulation in Bacillus subtilis. Genes Dev. 9:1316-1326. Wu, L. J., and J. Errington. 1998. Use of asymmetric cell division and spolIIE mutants to probe chromosome orientation and organization in Bacillus subtilis. Mol. Microbiol. 27~777-786. Wu, L. J., and J. Errington. 2000. Identification and characterization of a new prespore-specific regulatory gene, rsfA, of Bacillus subtilis. J. Bacteriol. 182:418-424. Yi, L., and R. E. Dalbey. 2005. OxallAlb3NidC system for insertion of membrane proteins in mitochondria, chloroplasts and bacteria (review). Mol. Membr. Biol. 22:lOl111. York, K., T. J. Kenny, S. Satola, and C. P. Moran, Jr. 1992. SpoOA controls the &dependent activation of Bacillus subtilis sporulation-specific transcription unit spollE. J. Bacterial. 174:2648-2658. Yu, Y.-T. N., and L. Kroos. 2000. Evidence that SpoIVFB is a novel type of membrane metalloprotease governing intercompartmental communication during Bacillus subtilis sporulation. J . Bacteriol. 182:3305-3309.
383 Yudkin, M. D., and J. Clarkson. 2005. Differential gene expression in genetically identical sister cells: the initiation of sporulation in Bacillus subtilis. Mol. Microbiol. 56578589. Zhang, B., R. Daniel, J. Errington, and L. Kroos. 1997a. Bacillus subtilis SpoIIID protein binds to two sites in the spoVD promoter and represses transcription by uERNA polymerase. J . Bacteriol. 179:972-975. Zhang, B., and L. Kroos. 1997. A feedback loop regulates the switch from one sigma factor to the next in the cascade controlling Bacillus subtilis mother cell gene expression. J. Bacteriol. 179:613 8-6144. Zhang, B., A. Hofmeister, and L. ICroos. 1998. The prosequence of pro-uKpromotes membrane association and inhibits RNA polymerase core binding. J. Bacteriol. 180:2434-2441. Zhang, B., P. Struffi, and L. Kroos. 1999. uKcan negatively regulate sigE expression by two different mechanisms during sporulation of Bacillus subtilis. J. Bacteriol. 181:40814088. Zhang, J., H. Ichikawa, R. Halberg, L. Kroos, and A. I. Aronson. 1994. Regulation of the transcription of a cluster of Bacillus subtilis spore coat genes.]. Mol. Biol. 240:405-415. Zhang, L., M. L. Higgins, P. J. Piggot, and M. L. Karow. 1996. Analysis of the role of prespore gene expression in the compartmentalization of mother-cell gene expression during sporulation of Bacillus subtilis. J. Bacteriol. 178:28132817. Zhang, L., M. L. Higgins, and P. J. Piggot. 199710. The division during bacterial sporulation is symmetrically located in Sporosarcina ureae. Mol. Microbiol. 25:1091-1098. Zheng, L., R. Halberg, S. Roels, H. Ichikawa, L. ICroos, and R. Losick. 1992. Sporulation regulatory protein GerE from Bacillus subtilis binds to and can activate or repress transcription from promoters for mother-cell-specific genes. J. Mol. Biol. 226:1037-1050. Zhou, R., and L. Kroos. 2004. BofA protein inhibits intramembrane proteolysis of pro-uK in an intercompartmental signaling pathway during Bacillus subtilis sporulation. Proc. Natl. Acad. Sci. U S A 101:6385-6390. Zhou, R., and L. Kroos. 2005. Serine proteases from two cell types target different components of a complex that governs regulated intramembrane proteolysis of pro-uKduring Bacillus subtilis development. Mol. Microbiol. 58:835-846. Zupancic, M. L., H. Tran, and A. E. Hofmeister. 2001. Chromosomal organization governs the timing of cell type-specific gene expression required for spore formation in Bacillus subtilis. Mol. Microbiol. 39:1471-1481.
Myxobacteria: Multicellularity and Differentiation Edited by David E. Whitworth 0 2008 ASM Press, Washington, D.C.
Deanne L. Pierce Yves V. Brun
Developmental Control in Caulobacter crescentus; Strategies for Survival in Oligotrophic Environments Bacteria must coordinate a multitude of cellular processes in order to thrive in their environment. In many wellstudied bacterial systems, such as Myxococcus xanthus and Bacillus subtilis, development is environmentally signaled. M. xanthus cells grow and divide vegetatively when nutrients are sufficient, but when nutrients become less available and cells are at a high density on a solid surface, M. xanthus shifts into a developmental mode (Dworkin, 2000). M . xanthus forms a fruiting body that is filled with metabolically inactive and somewhat resistant resting cells called myxospores, until conditions become favorable for vegetative growth again (Dworkin, 2000). Similarly, when faced with nutrient limitation, bacteria like those of the genus Bacillus form a dormant spore, which is highly resistant to heat and desiccation (Jiang et al., 2000; Stephenson and Hoch, 2002). In contrast, in stalked alphaproteobacteria such as Caulobacter crescentus, Asticcacaulis biprosthecum, and Hyphomicrobium neptunium, development is not triggered by environmental changes but is a consequence of normal progression through the cell cycle (Brun and Janakiraman, 2000). In these aquatic gramnegative organisms, cell division produces two different
22
cell types, a sessile cell with one or more extensions of the cell envelope called the stalk (or prosthecae), and a motile swarmer cell with a single polar flagellum. After an obligatory period as a nonreplicating motile cell, the swarmer cell differentiates into a stalked cell and initiates DNA replication. The location of the stalks and other appendages on the cells of these different species varies, but their overall developmental cycle is similar (Brun and Janakiraman, 2000). C. crescentus has a single polar stalk with an adhesive holdfast at the tip and a flagellum and pili at the opposite pole. A. biprosthecum has two lateral stalks, one on either side of the cell, with a holdfast and a flagellum at opposite poles of the cell. H. neptunium has a polar stalk from which the new swarmer cell buds, with the chromosome traversing the length of the stalk to enter the budding daughter cell (Brun and Janakiraman, 2000). C. crescentus is by far the best-studied prosthecate bacterium and has emerged as an outstanding model organism in which to study cellular organization, signal transduction, and the cell cycle. C. crescentus stalked cells produce a motile, swarmer daughter cell during every cell cycle (Fig. 1)(Martin and Brun, 2000). The swarmer
Deanne L. Pierce and Yves V. Brun, Department of Biology, Indiana University, Bloomington, IN 47405.
385
ANALOGOUS SYSTEMS
386
t I
I 1
Figure 1 The cell cycle of C. crescentus. Each round of cell division produces a stalked cell and a motile swarrner cell. The swarmer cell must differentiate into a stalked cell prior to undergoing cell division. cell has a polar flagellum and pili at the same pole and is incapable of initiating DNA replication until after differentiation into a stalked cell. During differentiation, the flagellum is shed, the pili are retracted, and a stalk with an adhesive holdfast at its tip is extended from the same pole that the flagellum previously occupied. The holdfast allows the cell to remain strongly attached to biotic and abiotic surfaces. The stalked cell is replication competent and divides to produce a new swarmer cell at the opposite pole. This intricate developmental pathway is likely to play many different roles in C. crescentus to confer survival advantages in its nutrient-limited environment. In contrast to M. xanthus and B. subtilis, which alternate between a feast or famine existence and produce dormant cells when nutrients become scarce, the oligotrophic C. crescentus has adopted a different strategy to cope with almost constant famine. First, the production of a chemotactically competent swarmer cell allows dispersal towards more nutritionally rich environments. Since the swarmer cell cannot start a new cell cycle for a fixed
portion of its life cycle, the motile period lasts longer when nutrients are less plentiful because the growth rate is slower. The longer motile period improves the chances that a better environment will be found. Furthermore, the production of a swarmer cell by a stalked cell attached to a surface should reduce the likelihood of competition between the stalked mother cell and its daughter swarmer cell since it increases the chances that the swarmer cell will disperse farther away from the stalked cell in nutrientpoor environments. Second, the successive synthesis of a flagellum, pili, and holdfast at the same pole of the cell optimizes attachment of swarmer cells to surfaces, thus improving the cell's access to nutrients absorbed to surfaces. Finally, once attached tightly to a surface by its holdfast, the cell synthesizes a stalk, which dramatically improves nutrient uptake in the diffusion-limited environment where C. crescentus typically lives. This chapter focuses on the mechanisms of surface adhesion and stalk function, as well as how the overall developmental cycle is controlled.
22. DEVELOPMENTAL CONTROL IN C.
387
CRESCENTUS
SURFACE ATTACHMENT AND HOLDFAST SYNTHESIS Differentiation is an essential process for adhesion of C. crescentus to surfaces (Bodenmiller et al., 2004). While the holdfast of C. crescentus generates a very strong, permanent attachment to surfaces, the holdfast alone is not sufficient for optimal attachment (Bodenmiller et al., 2004; Levi and Jenal, 2006; Tsang et al., 2006). A working flagellum is necessary for optimal attachment, presumably because it helps C. crescentus to overcome the repulsive barrier at a surface. Once the cell has made initial contact with a surface, the polar pili retract to bring the cell pole in closer contact with the surface. Pilus retraction positions the proper pole of the cell at the surface for holdfast secretion, allowing permanent attachment of the cell (Fig. 2). This model explains why growth is required for adhesion in C. crescentus; growth is necessary for the transition from the swarmer cell to the stalked cell (Bodenmilleret al., 2004). While polar development enhances the adhesion of swarmer cells to surfaces, adhesion is not required for swarmer cell differentiation. The strong adhesion of cells to a surface is mediated by the polar holdfast, found at the tip of C. crescentus stalks. Experiments in which individual cells were pulled off a glass surface gave an average adhesion force of 0.59 (20.62) p.N for single cells, with a range from 0.11 to 2.26 p.N (Tsang et al., 2006). This is by far the strongest force of adhesion measured for single microbial cells ( Abu-Lail and Camesano, 2003). With an average force per unit area of at least 80 N/mm2, equivalent to a force that could support a weight of 700 kg/cm2, the adhesion of the holdfast is the strongest ever measured for biological adhesives. The holdfast is an elastic material (Li et al., 2005) composed of polysaccharides and probably proteins and is absolutely required for strong adhesion to surfaces (Merker and Smit, 1988; Ong et al., 1990). The fluorescently labeled lectin fluorescein isothiocyanate-wheat germ agglutinin (FITC-WGA)binds to N-acetylglucosamine (GlcNac or NAG) and was shown
to bind specifically to the holdfast, indicating that the holdfast contains NAG (Merker and Smit, 1988). All mutants that cannot be stained with FITC-WGA are completely deficient in surface adhesion, indicating that the NAG polysaccharide is a critical component of the holdfast. The integrity of NAG polymers is critical for strong adhesion (Tsang et al., 2006). The holdfast can be partially degraded by lysozyme, which cleaves NAG polymers (Merker and Smit, 1988),and this partial degradation drastically reduces the force of cell adhesion (Tsang et al., 2006) and increases the elasticity of the holdfast (Li et al., 2005). In order to identify genes involved in holdfast biosynthesis, C. crescentus transposon mutants were screened for reduced binding to surfaces (Mitchell and Smit, 1990; Kurtz and Smit, 1992, 1994; Smith et al., 2003). These screens identified three classes of mutants: developmental mutants (pod] and p l e c ) , holdfast synthesis mutants (hfs),and holdfast attachment mutants (hfu).hfs mutants are unable to synthesize or export the holdfast, whereas hfu mutants are able to synthesizeand export the holdfast but are unable to keep it attached to the tip of the stalk. All of the hfs mutants mapped to a putative polysaccharide export gene cluster (hfsDABC),and all of the hfu mutants mapped to a different gene cluster (hfuABD). Sequence analysis, outlined below, suggests that these two gene clusters contain genes necessary for the export and attachment of the holdfast polysaccharide. Mutants with in-frame deletions of hfsA, hfsB, and hfsD have no detectable holdfasts and are unable to attach to surfaces or to form rosettes, groups of cells attached to each other by their holdfasts (Smith et al., 2003). HfsA and HfsB have sequence similarity to gram-negative exopolysaccharide export proteins (Smith et al., 2003). HfsD resembles Wza proteins, which function as outer membrane channels for export of capsular polysaccharide (Paulsen et al., 1997; Drummelsmith and Whitfield, 2000). HfsA has sequence similarity to the inner membrane periplasmic auxiliary (MPA-1 or Wzc) family of
Figure 2 Diagram of the stages of C. crescentus adhesion. From left to right, a swarmer cell approaches a substrate so that the pili and/or the flagellum binds the surface; weak attachment persists transiently, during which time the pili retract until tight attachment forms, mediated by the holdfast.
388 polysaccharide export proteins (Paulsen et al., 1997). Gram-negativeWzc proteins have a cytoplasmic C-terminal ATP-binding domain, but HfsA lacks this domain. In fact, HfsA is more similar to the gram-positiveMPA-1+C proteins, in which the transmembrane component and the ATP-bindingcomponents are encoded separately (Paulsen et al., 1997).HfsB is similar to the ATP-bindingcomponent (C component) of the gram-positive MPA-1+ C proteins such as CapB of Staphylococcus aureus. These proteins are tyrosine autokinases involved in polysaccharide chain length/export control (Whitfieldand Paiment, 2003). Wza and Wzc form a polysaccharide export channel spanning the periplasm and outer membrane (Dong et al., 2006; Collins et al., 2007). Therefore, HfsA, HfsB, and HfsD appear to be involved in holdfast polysaccharide export. Recently, a group of genes involved in the synthesis of the holdfast has been identified adjacent to the hfsABD genes (E. Toh and Y. V. Brun, unpublished data). The hfaA, hfaB, and hfaD genes are required for attachment of the holdfast to the tip of the stalk. Mutations in these genes result in shedding of the holdfast into the medium and cell detachment from holdfasts that are attached to a surface, suggesting that their protein products serve to anchor the holdfast in the outer membrane of the stalk (Kurtz and Smit, 1992; Cole et al., 2003; Smith et al., 2003). HfaB, a lipoprotein, and HfaD are membrane-associated proteins (Cole et al., 2003). Both HfaB and HfaD are enriched in the stalk compared to the cell body, consistent with their predicted function (Cole et al., 2003).It has been suggestedthat the HfaA and HfaD proteins are involved in the formation of a fimbria-like structure on the cell surface at the tip of the stalk. These fimbria-like structures would interact with the holdfast polysaccharide and anchor the holdfast to the cell. Part of this model is based on the similarity of Hfa proteins to curlin proteins. Curli are masses of thin, irregular, and highly aggregated surface fibers that are involved in adhesion events in Escherichia coli and Salmonella enterica serovar Enteritidis (Loferer et al., 1997). HfaB has sequence similarity to the curli secretin CsgG (Loferer et al., 1997; Robinson et al., 2006). HfaA is a small protein and has some structural similarity to the E. coli curlin, CsgA. The holdfast polysaccharide may be associated with the HfaA fimbriae, much like the curli-like thin aggregative fimbriae of S. enterica serovar Enteritidis are associated with extracellular polysaccharides (White et al., 2003).
SYNTHESIS AND FUNCTION OF THE STALK The stalk is an extension of the cell wall and membranes devoid of ribosomes, DNA (Poindexter and Bazire, 1964), and cytoplasmic proteins (Ireland et al., 2002;
ANALOGOUS SYSTEMS Wagner et al., 2006). The stalk is transected perpendicularly by crossbands synthesized during each round of cell division, which may serve to compartmentalize the stalk from the cell body (Poindexter and Staley, 1996). The compartmentalization of the stalk and cell body extends to inner membrane proteins, most of which appear to be absent from the stalk (Ireland et al., 2002; Wagner et al., 2006). In contrast, periplasmic proteins are present in the stalk (Wagner et al., 2006). There is growing evidence that suggests the stalk of Caulobacter is involved in nutrient uptake (Fig. 3 ) (Wagner and Brun, 2007). In rich media, Caulobacter stalks are usually only a couple of micrometers long, but in environments with low phosphate concentration the stalks dramatically elongate to tens of micrometers (Wagner and Brun, 2007). Some other prosthecate bacteria respond to carbon limitation by elongating their prosthecae (Whittenbury and Dow, 1977). When stalks are sheared from A. biprosthecum, they can deplete glucose and amino acids from growth medium (Larson and Pate, 1976; Tam and Pate, 1985). Similarly, purified C. crescentus stalks can take up and hydrolyze organic phosphate (Wagner et al., 2006). Since the stalk contains outer membrane and periplasmic proteins but no inner membrane proteins, nutrients probably bind to periplasmic nutrient binding proteins after being taken up through the outer membrane. The periplasmic binding protein-nutrient complexes would then diffuse to the cell body, where the nutrients can be taken up through the inner membrane using ABC transporters. C. crescentus is generally found in environments where nutrient uptake is diffusion limited, i.e., in environments where there is no flow past the cell. In such environments, the rate of nutrient uptake is proportional to the length of the cell rather than its surface area (Berg and Purcell, 1977; Berg, 1993; Wagner et al., 2006). Mathematical modeling indicates that the long, thin shape of the stalk is highly optimized from a biophysical standpoint for contacting diffusive nutrients while minimizing increases of both cell volume and surface area (Wagner et al., 2006). For example, a l-pm-long cell with a 2.4-pm stalk has a maximum rate of nutrient uptake 1.8 times greater than that of a 2-pm-long stalkless cell with the same surface area. The stalk is still useful under flow conditions (as opposed to diffusion) since a stalked cell and a stalkless cell of identical surface areas also have identical rates of nutrient uptake. However, since the stalk contributes less to volume than an elongated cell body shape, stalk synthesis is still highly economical from a bioenergetics standpoint. Stalk synthesis is a specialized type of cell elongation. Stalk synthesis requires the proteins PBP2 (Seitz and
22. DEVELOPMENTAL CONTROL IN C.
389
CRESCENTUS
/
i Cell body ~ u t r i ~ ~n t
~
~
a
k
~
Figure 3 Nutrient uptake model of stalk function in C. crescentus. Two nonexclusive models are proposed to explain nutrient uptake by the stalk. The periplasmic diffusion model illustrates nutrients binding to periplasmic receptors and being transported to the cell body through the periplasm, where they can then be taken up by the cell. The stalk core diffusion model shows the possibility of nutrients binding to periplasmic receptors and then being taken into the core of the stalk by transport proteins. The nutrients would then diffuse into the cell body. The available experimental data support the periplasmic diffusion model. Reprinted from Molecular Microbiology (Wagner and Brun, 2007) with permission of the publisher.
Brun, 1998), RodA, and MreB (Wagner et al., 2005), all of which are also involved in peptidoglycan synthesis and cell shape determination. Inhibition of PBP2 affects stalk elongation and morphogenesis (Seitz and Brun, 1998), as does depletion of RodA and MreB (Wagner et al., 2005). These proteins also define polar regions of the cell (Wagner et al., 2005), which would be expected to dictate the location of stalk synthesis. Stalk synthesis in C. crescentus is regulated by two different phosphorelay pathways. RpoN ( d4), an alternative sigma factor, and the response regulator TacA control the expression of a transcription factor, StaR, involved in stalk biogenesis (Biondi et al., 2006b). TacA is a member of the family of response regulators required for the activation of open complex formation by oS4-RNApolymerase holoenzyme. The phosphorelay that
culminates in activation of TacA includes ShkA, a hybrid histidine kinase, and ShpA, a histidine phosphotransfer (HPt) protein (Biondi et al., 2006b). ShkA autophosphorylates, after which ShpA transfers the phosphoryl group from ShkA to TacA, and then TacA and RpoN activate StaR, which in turn likely regulates expression of other genes involved in stalk synthesis (Biondi et al., 2006b). tucA and rpoN expression are positively regulated by CtrA when it is expressed after DNA replication initiation in the stalked cell. This regulation by CtrA ties their expression to the cell cycle, as discussed below. Stalk elongation is regulated by the Pho regulon. The PhoB response regulator is required for cells to elongate stalks in response to phosphate starvation (Gonin et al., 2000). PhoB controls expression of the pstSCAB genes, which encode high-affinity phosphate transport
ANALOGOUS SYSTEMS
390 proteins. Mutations in the p s t genes lead to a constitutive long-stalk phenotype, even in high-phosphate medium (Gonin et al., 2000), presumably due to induction of the Pho regulon. In a pst mutant, the Pho regulon is induced because the histidine kinase PhoR is no longer inactivated by the Pst proteins, resulting in activation of PhoB by phosphorylation and a long-stalk phenotype.
COORDINATING CELL DIVISION AND POLAR DEVELOPMENT IN C. CRESCENTUS The succession of external structures synthesized at the poles of C. crescentus is highly ordered and occurs at specific times during progression through the cell cycle. A complex signal transduction pathway ensures that polar development, DNA replication, and cell division are coordinated (Fig. 4).Much of this regulation
““‘y
controls the activity of the response regulator CtrA and the cyclic-di-GMP(c-di-GMP)synthetase PleD, enabling development to occur at the proper time in the cell cycle. CtrA inhibits the initiation of DNA replication by binding to the origin of replication (Marczynski and Shapiro, 2002) and directly regulates the transcription of 95 genes in the cell, many of which are involved in cell division and polar morphogenesis (Laub et al., 2002). CtrA expression is autoregulated (Domian et al., 1999), and the timing of CtrA activity is controlled by its own phosphorylation and proteolysis (Domian et al., 1997; Jenal and Fuchs, 1998). CtrA is present in swarmer cells, is degraded during swarmer cell differentiation, and is synthesized in predivisional cells (Domian et al., 1997). The response regulator PleD is required for the transition from the motile state to the nonmotile, adhered state during Cuulobucter development (Aldridge et al.,
PleD
x i v J ; x )C’*”’i
PleC
PleC PleD-P
Development
DivK-P
1_
k
n
c-di-GMP
x
CckA-P
I I+
’
GcrA
-
ChpT
‘7
T3-s:
CpdR-P
GcrA
c
P1
ChpT-P
CtrA-P
P2
+ CPdR + ClpXP+ RcdA= Proteolysis
I
T 4
1 Origin of Replication
c
Cell Division and Development
Figure 4 Regulatory pathway controlling cell division and polar development in C. crescentus. Sensor kinases and response regulators, along with other factors, coordinate polar development with cell cycle progression. Figure adapted from Biondi et al. (Biondi et al., 2006a).
22. DEVELOPMENTAL CONTROL IN C.
CRESCENTUS
391
2003). PleD was first identified as a bypass suppressor of a pleC mutant lacking motility (Sommer and Newton, 1989). pleD mutants have a supermotile phenotype, in which cells do not deactivate flagellar rotation and do not eject the flagellum or form stalks (Hecht and Newton, 1995). PleD plays a role in the degradation of the FliF flagellar anchor protein and in efficient ejection of the flagellum during the swarmer-to-stalked cell transition (Aldridge and Jenal, 1999). PleD is unique in that it has two N-terminal receiver domains and a C-terminal GGDEF output domain, which exhibits c-di-GMP synthetase (also called diguanylate cyclase) activity pending phosphorylation of the D1 receiver domain (Aldridge et al., 2003; Paul et al., 2004). c-di-GMP is a widespread second messenger in bacteria, with levels controlled by the opposing effects of diguanylate cyclases and phosphodiesterases (Jenal and Malone, 2006). Two histidine kinases, PleC and DivJ, which localize at opposite poles of predivisional cells, impact the activity of PleD directly and CtrA indirectly. Mutations in pleC result in cells that do not form stalks or pili and have a paralyzed flagellum (Wang et al., 1993).PleC localizes at the flagellar pole in swarmer and predivisional cells, while DivJ localizes at the stalked pole (Wheeler and Shapiro, 1999). div] was identified in a screen for mutations that suppressed the motility defect of a temperature-sensitive pleC mutant at high temperature (37°C) but simultaneously conferred a cell division defect at cold temperature (24°C) (Sommer and Newton, 1991).A div] null mutant is filamentous and often forms long, misplaced stalks. DivJ and PleC both regulate the phosphorylation state of the response regulator DivK, which couples development to cell division (Wheeler and Shapiro, 1999). DivJ serves as the kinase for DivK, while PleC acts primarily as a phosphatase for phosphorylated DivK (DivK-P) (Matroule et al., 2004). The phosphorylation state of DivK controls its localization, with DivK colocalizing with DivJ at the stalked pole of the predivisional cell (Color Plate 9). DivK is phosphorylated by DivJ, and then DivK-P subsequently localizes to the opposite pole with PleC, where PleC dephosphorylates DivK-P. Once cell compartmentalization is complete, DivK in the swarmer compartment is no longer phosphorylated since it cannot diffuse to DivJ, and DivK-P in the stalked cell cannot be dephosphorylated because PleC is not present. This restriction of DivK phosphorylation and dephosphorylation to exclusive compartments signals compartmentalization to the cell, allowing swarmer-progeny-specific development in the new swarmer cell and allowing the cell cycle to continue in the stalked cell. In addition to its role in signaling cell compartmentalization, DivK-P also plays a role in the degradation of CtrA, discussed
below. DivJ activates DivK, and activated DivK signals CtrA proteolysis, which is likely the reason for the filamentous nature of div] mutant cells. DivJ and PleC also directly regulate the phosphorylation of PleD (Paul et al., 2004), with DivJ acting as a kinase primarily and PleC likely acting as a phosphatase or in another manner to negatively affect PleD-P levels (Aldridge et al., 2003), much as they act on the response regulator DivK (Wheeler and Shapiro, 1999; Matroule et al., 2004). PleD is localized to the stalked pole in stalked and predivisional cells and randomly dispersed in swarmer cells (Paul et al., 2004). DivJ phosphorylates PleD at the swarmer-to-stalked cell transition, allowing PleD to act in the transition from a motile to a nonmotile state, and PleC dephosphorylates PleD-P in the swarmer compartment of the predivisional cell, inactivating it so that flagellar rotation is turned on (Aldridge et al., 2003). CtrA activation plays an important role in cell cycle regulation. The hybrid kinase CckA is required for phosphorylation of CtrA in vivo (Jacobs et al., 1999), and the tyrosine kinase DivL phosphorylates CtrA in vitro (Wu et al., 1999). CckA and CtrA are phosphorylated at the same time during the cell cycle and regulate the same genes (Jacobs et al., 1999,2003). The newly identified ChpT phosphotransferase transfers the phosphoryl group from the CckA receiver domain to CtrA (Biondi et al., 2006a). CtrA has two promoters which are active at different times in the predivisional cell: P1 is repressed by CtrA, whereas P2 is activated (Domian et al., 1999). After CtrA is degraded during swarmer cell differentiation, ctrA transcription is initiated by the weak P1 promoter in early predivisional cells, producing a small amount of CtrA. However, once a threshold amount of CtrA is produced and phosphorylated, CtrA-P represses transcription from the P1 promoter and activates high levels of transcription from the P2 promoter (Fig. 4) (Domian et al., 1999). This continues until proteolysis of CtrA breaks the feedback loop for P2. The tyrosine kinase DivL is essential for cell division and viability. This kinase is unusual in that it has the structure and conserved sequence of a bacterial histidine kinase, but it contains a tyrosine residue instead of the conserved histidine residue (Wu et al., 1999).Phosphorylated DivL can transfer phosphate to the global response regulator CtrA in vitro, and suppressor mutations in CtrA suppress the cell division defect seen in conditional DivL mutants, suggesting that DivL functions, at least in part, through CtrA (Wu et al., 1999). Additionally, mutations in DivL and CckA suppress a div] deletion mutant by reducing the levels of active CtrA in the cell, again indicating that DivL and CckA act through CtrA (Pierce et al., 2006). Interestingly, in addition to its role
392 in potentially activating CtrA, DivL has been shown to interact with the single-domain response regulator DivK in a yeast two-hybrid screen (Ohta and Newton, 2003). In addition to CtrA, the C. crescentus cell cycle is also regulated by GcrA, a regulatory protein that influences transcription of 125 genes, including ctrA (Holtzendorff et al., 2004). GcrA antagonizes CtrA activity through an unknown mechanism. CtrA and GcrA levels oscillate in the cell, with GcrA initiating ctrA expression, either by directly binding the CtrA P1 promoter or by binding another protein which binds the promoter (Holtzendorff et al., 2004). The subsequent accumulation of CtrA represses expression of the gcrA promoter. Upon controlled proteolysis of CtrA in the stalked compartment of the predivisional cell and expression of the replication initiation factor DnaA (Hottes et al., 2005), gcrA expression is no longer repressed and GcrA can accumulate to begin the cycle again (Holtzendorff et al., 2004). Many of the 125 genes directly or indirectly regulated by GcrA are involved in DNA replication, cell motility, and polar development (including the gene for the polar development protein PodJ, described below). There are 25 regulatory genes affected by GcrA, which include genes encoding the signal transduction proteins PleC, DivK, and of course, CtrA (Holtzendorff et al., 2004). CtrA activity is cell cycle regulated by proteolysis, in addition to phosphorylation (Domian et al., 1997). CtrA is degraded by the ClpXP protease in the stalkedcell compartment of the predivisional cell and at the swarmer-to-stalked cell transition (Domian et al., 1997; Jenal and Fuchs, 1998). ClpXP requires an unknown signal present in the first 56 amino acids of the receiver domain and the 15 amino acids at the C terminus of CtrA in order to degrade it efficiently (Ryan et al., 2002). CtrA accumulates at the cell poles of predivisional cells and swarmer cells, just prior to proteolysis; the phosphorylation state of CtrA does not have an effect on its proteolysis or localization (Ryan et al., 2002,2004). The receiver domain signal within amino acids 1to 56 is required for polar localization of CtrA; however, the C-terminal 15 amino acids of CtrA required for degradation are not required for localization (Ryan et al., 2004). Transcription of the single-domain response regulator DivK is regulated by CtrA (Hung and Shapiro, 2002), and DivK in turn plays a role in the proteolysis of CtrA. When DivK is conditionally inactivated, degradation of CtrA is blocked (Hung and Shapiro, 2002). Inactivation of DivK also affects the polar localization of CtrA, resulting in a continuous low level of polar foci, instead of the higher numbers of foci seen in a wild-type strain (Ryan et al., 2004). However, CtrA localization at the cell poles does not depend on the colocalization of DivK, just the activity
ANALOGOUS SYSTEMS of DivK (Ryan et al., 2004). Taken together, these results suggest that DivK promotes the proper localization of CtrA during the cell cycle, allowing it to be degraded in the correct cell types at the correct times during the cell cycle. Additionally, it has recently been shown that cells overproducing DivK have delocalized CckA and a significant decrease in CtrA-P, both of which depend on the presence of DivJ, the kinase for DivK (Biondi et al., 2006a). This suggests that the role of DivK-P in the inactivation and degradation of CtrA is mediated by the effects of DivK-P on CckA localization and activity (Biondi et al., 2006a). In addition to the ClpXP protease, the degradation of CtrA requires the concerted action of a polar localization factor, RcdA, and a response regulator, CpdR. The RcdA protein was found by mining genomic data for CtrA-regulatory proteins (McGrath et al., 2006). RcdA localizes to the stalked pole a t the same time as CtrA, is required for the localization and degradation of CtrA, and requires the ClpX subunit of ClpXP for its localization (McGrath et al., 2006). ClpX, CtrA, and RcdA interact to form a complex at the cell pole (McGrath et al., 2006). The CpdR response regulator shares the localization pattern of CtrA, RcdA, and ClpXP and is required for CtrA proteolysis (Iniesta et al., 2006). CpdR is required for ClpX localization at the cell pole, where the two proteins interact (Iniesta et al., 2006). Localization of CpdR (and therefore ClpXP) is regulated by the phosphorylation state of CpdR. When CpdR is phosphorylated, it is not localized, which prevents proteolysis of CtrA (Iniesta et al., 2006). CpdR is phosphorylated by the hybrid kinase CckA, via ChpT (Biondi et al., 2006a), which also phosphorylates CtrA (Biondi et al., 2006a; Iniesta et al., 2006). Thus, the order of localization is as follows: CpdR, ClpX, RcdA, and finally CtrA, leading to the degradation of CtrA. This complex process likely ensures that CtrA is degraded only at the proper time and place in the cell, particularly since CpdR is required for the localization of the protease and CtrA and is prevented from localizing by being phosphorylated by the same proteins, CckA and ChpT, that activate CtrA.
ADDITIONAL FACTORS IN POLAR DEVELOPMENT AND REGULATION As mentioned above, the expression of the developmental factor PodJ is regulated by GcrA (Holtzendorff et al., 2004) and negatively by CtrA (Crymes et al., 1999). PodJ is a polar organelle development protein that is required for the synthesis of pili and holdfast, as well as normal motility in low-concentration agar and flagellar release, all of which also require the histidine
22. DEVELOPMENTAL CONTROL IN C.
CRESCENTUS
393
kinase PleC (Viollier et al., 2002; Hinz et al., 2003). Interestingly, PodJ is required for proper polar localization of PleC, although it is not known if PleC localization is a factor in the regulation of its activity (Viollier et al., 2002; Hinz et al., 2003). PodJ is a modular protein with different domains required for different functions. The periplasmic portion of PodJ is required for pilus synthesis, while the cytoplasmic portion is necessary for swarming motility and holdfast synthesis (Viollier et al., 2002; Lawler et al., 2006).PodJ is synthesized in its fulllength form (PodJ,) in stalked cells, where it localizes to the pole opposite the stalk. PodJ, is then proteolytically cleaved into a shorter form (PodJ,) at the time of cell division (Viollier et al., 2002; Hinz et al., 2003) in a reaction that requires the PerP periplasmic protease (Chen et al., 2006). PodJ, remains at the swarmer pole until it is degraded upon differentiation of the swarmer cell to a stalked cell by the MmpA protease (Chen et al., 2005). PodJ, ensures that PleC is properly localized to the swarmer pole, where PleC contributes to the signal that cell compartmentalization has taken place by reducing the level of DivK phosphorylation. Reduction of DivK-P in the swarmer compartment results in CtrA activation, which in turn activates PerP expression, directly or indirectly, resulting in a regulatory loop (Chen et al., 2006). Clearly, cell polarity is a major factor in the development of C. crescentus. The new poles of the daughter cells formed at each round of cell division are marked by the protein TipN (Huitema et al., 2006; Lam et al., 2006). The new pole remains marked until late in cell division when TipN moves from the pole to the site of division. As such, upon division, both daughter cells have TipN at the new pole. The release of TipN from the poles of the cell depends on the size of the predivisional cell, while the relocalization of TipN is dependent on cell division (Huitema et al., 2006; Lam et al., 2006). In cells lacking TipN, the polarity of the cells is disturbed, which resulted in misplacement of the flagella at the old pole approximately 75 to 85% of the time (Huitema et al., 2006; Lam et al., 2006). This is due at least in part to the mislocalization of TipF, a flagellum assembly factor with c-di-GMP-specific phosphodiesterase activity that requires TipN for proper localization (Huitema et al., 2006).This c-di-GMP degradation activity of TipF may tie in with the role of the PleD diguanylate cyclase (Huitema et al., 2006).
the successive synthesis of polar structures, C. crescentus can bring itself in closer contact with adsorbed nutrients. At the same time, by producing a motile progeny cell, the stalked parent cell ensures that the progeny will be capable of swimming away to avoid local competition for nutrients. The stalk of C. crescentus is able to take up nutrients and can be elongated when phosphate and possibly other nutrients are limited, allowing a cell to scavenge for nutrients even when it is sessile. The same regulatory factors that control PleD activity at the swarmer-to-stalked cell transition also control activation of DivK and therefore CtrA activation and degradation. PleD is required for the transition from the motile to sessile adhered state in C. crescentus, while DivK and CtrA are involved in the control of the cell cycle, as well as other aspects of polar development. It is not surprising that such a complex signal transduction pathway is required to coordinate the multiple stages of polar development with the DNA replication and cell division cycle. Development and cell cycle regulation have been studied mostly under conditions where nutrients are plentiful. In the future, it will be interesting to investigate how the regulatory pathways that have been uncovered might be modulated by cell physiology.
CONCLUSION The developmental life cycle of C. crescentus is well suited to the nutrient-limiting aquatic environments where it lives. By attaching to surfaces, a process optimized by
References Abu-Lail, N. I., and T. A. Camesano. 2003. Polysaccharide properties probed with atomic force microscopy. J. Microsc. 17-23 8. 2 12~2 Aldridge, P., and U. Jenal. 1999. Cell cycle-dependent degradation of a flagellar motor component requires a novel-type response regulator. Mol. Microbiol. 32:379-391. Aldridge, P., R. Paul, P. Goymer, P. Rainey, and U. Jenal. 2003. Role of the GGDEF regulator PleD in polar development of Caulobacter crescentus. Mol. Microbiol. 47:1695-1 70 8. Berg, H. C. 1993. Random Walks in Biology. Princeton University Press, Princeton, NJ. Berg, H. C., and E. M. Purcell. 1977. Physics of chemoreception. Biophys. 1.20~193-219. Biondi, E. G., S. J. Reisinger, J. M. Skerker, M. Arif, B. S. Perchuk, K. R. Ryan, and M. T. Laub. 2006a. Regulation of the bacterial cell cycle by an integrated genetic circuit. Nature 444:899-904. Biondi, E. G., J. M. Skerker, M. Arif, M. S. Prasol, B. S. Perchuk, and M. T. Laub. 2006b. A phosphorelay system controls stalk biogenesis during cell cycle progression in Caulobacter crescentus. Mol. Microbiol. 59:386-401. Bodenmiller, D., E. Toh, and Y. V. Brun. 2004. Development of surface adhesion in Caulobacter crescentus. J. Bacteriol. 186~1438-1447. Brun, Y. V., and R. Janakiraman. 2000. The dimorphic life cycle of Caulobacter and stalked bacteria, p. 297-317. In Y. V. Brun and L. J. Shimkets (ed.), Prokaryotic Development. ASM Press, Washington, DC.
394 Chen, J. C., A. K. Hottes, H. H. McAdams, P. T. McGrath, P. H. Viollier, and L. Shapiro. 2006. Cytokinesis signals truncation of the PodJ polarity factor by a cell cycle-regulated protease. EMBO J. 25:377-386. Chen, J. C., P. H. Viollier, and L. Shapiro. 2005. A membrane metalloprotease participates in the sequential degradation of a Caulobacter polarity determinant. Mol. Microbiol. 55:1085-1103. Cole, J., G. G. Hardy, D. Bodenmiller, E. Toh, A. Hinz, and Y. V. Brun. 2003. The HfaB and HfaD adhesion proteins of Caulobacter crescentus are localized in the stalk. Mol. Microbiol. 49: 1671-1 683. Collins, R. F., K. Beis, C. Dong, C. H. Botting, C. McDonnell, R. C. Ford, B. R. Clarke, C. Whitfield, and J. H. Naismith. 2007. The 3D structure of a periplasm-spanning platform required for assembly of group 1 capsular polysaccharides in Escherichia coli. Proc. Natl. Acad. Sci. USA 104: 2390-23 95. Crymes, W. B., Jr., D. Zhang, and B. Ely. 1999. Regulation of podJ expression during the Caulobacter crescentus cell cycle. J. Bacteriol. 181:3967-3973. Domian, I. J., K. C. Quon, and L. Shapiro. 1997. Cell typespecific phosphorylation and proteolysis of a transcriptional regulator controls the G1-to-S transition in a bacterial cell cycle. Cell 90:415-424. Domian, I. J., A. Reisenauer, and L. Shapiro. 1999. Feedback control of a master bacterial cell-cycle regulator. Proc. Natl. Acad. Sci. USA 96:6648-6653. Dong, C., K. Beis, J. Nesper, A. L. Brunkan-Lamontagne, B. R. Clarke, C. Whitfield, and J. H. Naismith. 2006. Wza, the translocon for E. coli capsular polysaccharides, defines a new class of membrane protein. Nature 444:226-229. Drummelsmith, J., and C. Whitfield. 2000. Translocation of group 1capsular polysaccharide to the surface of Escherichia coli requires a multimeric complex in the outer membrane. E M B O J. 1957-66. Dworkin, M. 2000. Introduction to the Myxobacteria, p. 221-242. In Y. V. Brun and L. J. Shimkets (ed.),Prokaryotic Development. ASM Press, Washington, DC. Gonin, M., E. M. Quardokus, D. O’Donnol, J. Maddock, and Y. V. Brun. 2000. Regulation of stalk elongation by phosphate in Caulobacter crescentus. J. Bacteriol. 182: 337-347. Hecht, G. B., and A. Newton. 1995. Identification of a novel response regulator required for the swarmer-to-stalked-cell transition in Caulobacter crescentus. J. Bacteriol. 177:62236229. Hinz, A. J., D. E. Larson, C. S. Smith, and Y. V. Brun. 2003. The Caulobacter crescentus polar organelle development protein PodJ is differentially localized and is required for polar targeting of the PleC development regulator. Mol. Microbiol. 47929-941. Holtzendorff, J., D. Hung, P. Brende, A. Reisenauer, P. H. Viollier, H. H. McAdams, and L. Shapiro. 2004. Oscillating global regulators control the genetic circuit driving a bacterial cell cycle. Science 304:983-987. Hottes, A. K., L. Shapiro, and H. H. McAdams. 2005. DnaA coordinates replication initiation and cell cycle transcription in Caulobacter crescentus. Mol. Microbiol. 58:1340-1353.
ANALOGOUS SYSTEMS Huitema, E., S. Pritchard, D. Matteson, S. K. Radhakrishnan, and P. H. Viollier. 2006. Bacterial birth scar proteins mark future flagellum assembly site. Cell 124:1025-1037. Hung, D. Y., and L. Shapiro. 2002. A signal transduction protein cues proteolytic events critical to Caulobacter cell cycle progression. Proc. Natl. Acad. Sci. USA 99:13160-13165. Iniesta, A. A., P. T. McGrath, A. Reisenauer, H. H. McAdams, and L. Shapiro. 2006. A phospho-signaling pathway controls the localization and activity of a protease complex critical for bacterial cell cycle progression. Proc. Natl. Acad. Sci. U S A 103:10935-10940. Ireland, M. M., J. A. Karty, E. M. Quardokus, J. P. Reilly, and Y. V. Brun. 2002. Proteomic analysis of the Caulobacter crescentus stalk indicates competence for nutrient uptake. Mol. Microbiol. 45:1029-1041. Jacobs, C., N. Ausmees, S. J. Cordwell, L. Shapiro, and M. Laub. 2003. Functions of the CckA histidine kinase in Caulobacter cell cycle control. Mol. Microbiol. 47:1279-1290. Jacobs, C., I. J. Domian, J. R. Maddock, and L. Shapiro. 1999. Cell cycle-dependentpolar localization of an essential bacterial histidine kinase that controls DNA replication and cell division. Cell 97:lll-120. Jenal, U., and T. Fuchs. 1998. An essential protease involved in bacterial cell-cycle control. E M B O J. 175658-5669. Jenal, U., and J. Malone. 2006. Mechanisms of cyclic-di-GMP signaling in bacteria. Annu. Rev. Genet. 40:385-407. Jiang, M., W. Shao, M. Perego, and J. A. Hoch. 2000. Multiple histidine kinases regulate entry into stationary phase and sporulation in Bacillus subtilis. Mol. Microbiol. 38535-542. Kurtz, H. D., Jr., and J. Smit. 1992. Analysis of a Caulobacter crescentus gene cluster involved in attachment of the holdfast to the cell. J. Bacteriol. 174:687-694. Kurtz, H. D., Jr., and J. Smit. 1994. The Caulobacter crescentus holdfast: identification of holdfast attachment complex genes. FEMS Microbiol. Lett. 116:175-182. Lam, H., W. B. Schofield, and C. Jacobs-Wagner. 2006. A landmark protein essential for establishing and perpetuating the polarity of a bacterial cell. Cell 124:1011-1023. Larson, R. J., and J. L. Pate. 1976. Glucose transport in isolated prosthecae of Asticcacaulis biprosthecum. J. Bacteriol. 126:282-293. Laub, M. T., S. L. Chen, L. Shapiro, and H. H. McAdams. 2002. Genes directly controlled by CtrA, a master regulator of the Caulobacter cell cycle. Proc. Natl. Acad. Sci. U S A 99~4632-4637. Lawler, M. L., D. E. Larson, A. J. Hinz, D. Klein, and Y. V. Brun. 2006. Dissection of functional domains of the polar localization factor PodJ in Caulobacter crescentus. Mol. Microbiol. 59:301-3 16. Levi, A., and U. Jenal. 2006. Holdfast formation in motile swarmer cells optimizes surface attachment during Caulobacter crescentus development. J. Bacteriol. 18853155318. Li, G., C. S. Smith, Y. V. Brun, and J. X. Tang. 2005. The elastic properties of the Caulobacter crescentus adhesive holdfast are dependent on oligomers of N-acetylglucosamine./. Bacteriol. 182257-265. Loferer, H., M. Hammar, and S. Normak. 1997. Availability of the fibre subunit CsgA and the nucleator protein CsgB
CONTROL IN C. 22. DEVELOPMENTAL
CRESCENTUS
395
during assembly of fibronectin-binding curli is limited by the intracellular concentration of the novel lipoprotein CsgG. Mol. Microbiol. 26:ll-23. Marczynski, G. T., and L. Shapiro. 2002. Control of chromosome replication in Caulobacter crescentus. Annu. Rev. Microbiol. 56:625-656. Martin, M. E., and Y. V. Brun. 2000. Coordinating development with the cell cycle in Caulobacter. Curr. Opin. Microbiol. 3589-595. Matroule, J. Y., H. Lam, D. T. Burnette, and C. Jacobs-Wagner. 2004. Cytokinesis monitoring during development: rapid pole-to-pole shuttling of a signaling protein by localized kinase and phosphatase in Caulobacter. Cell 118579-590. McGrath, P. T., A. A. Iniesta, K. R. Ryan, L. Shapiro, and H. H. McAdams. 2006. A dynamically localized protease complex and a polar specificity factor control a cell cycle master regulator. Cell 124535-547. Merker, R. I., and J. Smit. 1988. Characterization of the adhesive holdfast of marine and freshwater caulobacters. Appl. Environ. Microbiol. 54:2078-2085. Mitchell, D., and J. Smit. 1990. Identification of genes affecting production of the adhesion organelle of Caulobacter crescentus CB2. J . Bacteriol. 1725425-5431. Ohta, N., and A. Newton. 2003. The core dimerization domains of histidine kinases contain recognition specificity for the cognate response regulator. J. Bacteriol. 185:4424-4431. Ong, C. J., M. L. Y. Wong, and J. Smit. 1990. Attachment of the adhesive holdfast organelle to the cellular stalk of Caulobacter crescentus. J. Bacteriol. 172:1448-1456. Paul, R., S. Weiser, N. C. Amiot, C. Chan, T. Schirmer, B. Giese, and U. Jenal. 2004. Cell cycle-dependent dynamic localization of a bacterial response regulator with a novel di-guanylate cyclase output domain. Genes Dev. 18:715-727. Paulsen, I. T., A. M. Beness, and M. H. Saier, Jr. 1997. Computer-based analyses of the protein constituents of transport systems catalysing export of complex carbohydrates in bacteria. Microbiology 143:2685-2699. Pierce, D. L., D. S. O’Donnol, R. C. Allen, J. W. Javens, E. M. Quardokus, and Y. V. Brun. 2006. Mutations in DivL and CckA rescue a div] null mutant of Caulobacter crescentus by reducing the activity of CtrA. J. Bacteriol. 188:2473-2482. Poindexter, J. L. S., and G. C. Bazire. 1964. The fine structure of stalked bacteria belonging to the family Caulobacteraceae. J. Cell Biol. 23587-607. Poindexter, J. S., and J. T. Staley. 1996. Caulobacter and Asticcacaulis stalk bands as indicators of stalk age. 1. Bacteriol. 178:3939-3948. Robinson, L. S., E. M. Ashman, S. J. Hultgren, and M. R. Chapman. 2006. Secretion of curli fibre subunits is mediated by the outer membrane-localized CsgG protein. Mol. Microbiol. 595370-88 1. Ryan, K. R., S. Huntwork, and L. Shapiro. 2004. Recruitment of a cytoplasmic response regulator to the cell pole is linked to its cell cycle-regulated proteolysis. Proc. Natl. Acad. Sci. USA 101:7415-7420. Ryan, I(. R., E. M. Judd, and L. Shapiro. 2002. The CtrAresponse regulator essential for Caulobacter crescentus cell-cycle progression requires a bipartite degradation signal for temporally controlled proteolysis. J. Mol. Biol. 324:443455.
Seitz, L. C., and Y. V. Brun. 1998. Genetic analysis of mecillinam-resistant mutants of Caulobacter crescentus deficient in stalk biosynthesis. J. Bacteriol. 1805235-5239. Smith, C. S., A. Him, D. Bodenmiller, D. E. Larson, and Y. V. Brun. 2003. Identification of genes required for synthesis of the adhesive holdfast in Caulobacter crescentus.]. Bacteriol. 185~1432-1442. Sommer, J. M., and A. Newton. 1991. Pseudoreversion analysis indicates a direct role of cell division genes in polar morphogenesis and differentiation in Caulobacter crescentus. Genetics 129:623-630. Sommer, J. M., and A. Newton. 1989. Turning off flagellum rotation requires the pleiotropic gene pleD: pleA, pleC, and pleD define two morphogenic pathways in Caulobacter crescentus.]. Bacteriol. 171:392-401. Stephenson, K., and J. A. Hoch. 2002. Evolution of signalling in the sporulation phosphorelay. Mol. Microbiol. 46:297-304. Tam, E., and J. L. Pate. 1985. Amino acid transport by prosthecae of Asticcacaulis biprosthecum: evidence for a broadrange transport system. J. Gen. Microbiol. 131:2687-2699. Tsang, P. H., G. Li, Y. V. Brun, L. B. Freund, and J. X. Tang. 2006. Adhesion of single bacterial cells in the micronewton range. Proc. Natl. Acad. Sci. USA 1035764-5768. Viollier, P. H., N. Sternheim, and L. Shapiro. 2002. Identification of a localization factor for the polar positioning of bacterial structural and regulatory proteins. Proc. Natl. Acad. Sci. USA 99:13831-13836. Wagner, J. K., and Y. V. Brun. 2007. Out on a limb: how the Caulobacter stalk can boost the study of bacterial cell shape. Mol. Microbiol. 64:28-33. Wagner, J. I<., C. D. Galvani, and Y. V. Brun. 2005. Caulobacter crescentus requires RodA and MreB for stalk synthesis and prevention of ectopic pole formation. J. Bacteriol. 187544-553. Wagner, J. K., S. Setayeshgar, L. A. Sharon, J. P. Redly, and Y. V. Brun. 2006. A nutrient uptake role for bacterial cell envelope extensions. Proc. Natl. Acad. Sci. USA 103: 11772-1 1777. Wang, S. P., P. L. Sharma, P. V. Schoenlein, and B. Ely. 1993. A histidine protein kinase is involved in polar organelle development in Caulobacter crescentus. Proc. Natl. Acad. Sci. USA 90:630-634. Wheeler, R. T., and L. Shapiro. 1999. Differential localization of two histidine kinases controlling bacterial cell differentiation. Mol. Cell 4:683-694. White, A. P., D. L. Gibson, S. K. Collinson, P. A. Banser, and W. W. Kay. 2003. Extracellular polysaccharides associated with thin aggregative fimbriae of Salmonella enterica serovar Enteritidis. J. Bacteriol. 1855398-5407. Whitfield, C., and A. Paiment. 2003. Biosynthesis and assembly of Group 1 capsular polysaccharides in Escherichia coli and related extracellular polysaccharides in other bacteria. Carbohydr. Res. 338:2491-2502. Whittenbury, R., and C. S. Dow. 1977. Morphogenesis and differentiation in Rhodomicrobium vannielii and other budding and prosthecate bacteria. Bacteriol. Rev. 413754-808. Wu, J., N. Ohta, J. L. Zhao, and A. Newton. 1999. A novel bacterial tyrosine kinase essential for cell division and differentiation. Proc. Natl. Acad. Sci. USA 96:13068-13073.
Myxobacteria: Mtrlticelhlarity and Differentiation Edited by David E. Whitworth 0 2008 ASM Press, Washington, D.C.
Jindong Zhao C. Peter Wolk
Developmental Biology of Heterocysts, 2006
HETEROCYST STRUCTURE AND FUNCTIONS Over 2 billion years ago, with the growth of large numbers of O,-producing cyanobacteria, our planet emerged from anaerobiosis. The cyanobacteria needed only light, water, and inorganic nutrients, with CO, as a carbon source, to replicate. The 0, that they produced was a very-broad-spectrum biocide. As a result, cyanobacteria were likely the dominant life form on earth. Despite an atmosphere composed mostly of N,, available nitrogen limited the growth of cyanobacteria because the N2-fixing enzyme, nitrogenase, was inactivated by 0,. Certain filamentous cyanobacteria overcame that limitation by inventing true multicellular division of labor. That is, some of the cells in a filament traded the potentiality of replication for service, by becoming N,-fixing heterocysts, cells able to fix N, in an 0,-containing environment and to provide the fixed nitrogen to the other cells in the filament. For information beyond the scope of this chapter, the reader is referred to recent reviews on heterocysts (Adams and Duggan, 1999; Golden and Yoon, 2003; Meeks and Elhai, 2002; Thiel, 2004; Wolk, 2000; Wolk et al., 1994; Zhang et al., 2006).
23
Depending on the organism, heterocysts usually form either at semiregular intervals along the cyanobacterial filaments or only at one or both ends of such a filament. In the former situation, new heterocysts normally arise approximately midway between two preexisting heterocysts as the interval separating those two approximately doubles. When heterocysts are found only at the termini of filaments, either a filament breaks into two filaments, with a heterocyst forming at each newly created end, or a pair of heterocysts forms near the middle of such a filament and then splits apart, leaving two filaments, each ending in a new heterocyst. Five years ago, one of us concluded that “Two questions concerning the developmental biology of heterocysts appear finally to be yielding to experimentation” (Wolk, 2000), namely, how is it decided which cells will, and which will not, differentiate; and how is the subsequent progression of the differentiation process regulated? To date, the first question has yielded more than the second. To provide a place of refuge for the 0,-labile enzyme, nitrogenase, in an 0,-containing milieu, heterocysts differ from vegetative cells in various ways. They stop producing 0,. To reduce the entry of O,, they deposit a circumferential barrier comprising laminae of glycolipids.
Jindong Zhao, State Key Laboratory of Protein and Plant Genetic Engineering, College of Life Sciences, Peking University, Beijing 100871, China. C . Peter Wolk, MSU-DOE Plant Research Laboratory and Department of Plant Biology, Michigan State University, E. Lansing, MI 48824.
397
ANALOGOUS SYSTEMS
398 Presumably because that barrier is easily damaged, they first deposit an outer, protective layer of polysaccharide. The chemical structures of these materials are known (Wolk et al., 1994; Wolk, 2000). Because N, must penetrate to be assimilated and because N, and 0, are physically similar, some 0, also enters. To remove permeant 0,, heterocysts greatly increase their rate of respiration. The reductant that is used to fix N, and to reduce 0, comes from photolysis of water, a process that takes place only in vegetative cells. Therefore, the vegetative cells provide the reductant in exchange for fixed nitrogen. Numerous genes have been identified that are required specifically for synthesis and deposition of heterocyst envelope glycolipids and polysaccharide, respiratory processes in heterocysts, and cessation of division in those cells that will become heterocysts. Genes have also been identified whose products are required for the regulation of those processes, as have been yet other genes that are required for heterocyst maturation but whose roles or mechanisms have not yet been discerned. More generally, much remains to be clarified about how the progression of the differentiation process is regulated. Stage:
Initiation
Some filamentous cyanobacteria also form akinetes (a spore form), which may have been the evolutionary precursor of heterocysts. Akinetes differentiate either specifically adjacent to heterocysts (or sometimes near but not contiguous), distant from heterocysts, or independent of heterocysts. The formation of patterns of spaced heterocysts has been the subject of extensive research. The formation of patterns of different, juxtaposed cells has not yet received a similar degree of investigation, in part because facile genetic manipulation has not been found for any organism that forms such patterns.
SIGNALS FOR HETEROCYST DEVELOPMENT AND THEIR TRANSDUCERS Nitrogen Deprivation, 2-Oxoglutarate, GlnB, NtcA, and HetR Signals for heterocyst development and their transducers are depicted in Fig. 1. The initiation of heterocyst development is suppressed completely by ammonium or very extensively by nitrate in most heterocyst-forming cyanoCommitment
Proheterocyst
Maturity
NtcA
.
1 NtcA
1
nif/hup rearrangements
0 2-OG?
Vegetative Cell
RbcLXS
cell division
Figure 1 Stages of heterocyst development. An increase in intracellular 2-OG (2-oxoglutarate) signals nitrogen deprivation, which may be transduced by NtcA, a protein upon which numerous developmental processes depend. Itself autoregulatory, NtcA participates in autoregulation of HetR, the central regulator of heterocyst formation. Beyond commitment, heterocysts do not revert to vegetative growth. The ability to fix N, and to reassimilate H,, both of which require internally micro-oxic conditions, represents maturity. Semicolons demarcate known or presumptive regulatory proteins from presumptive enzymatic proteins. Short vertical arrows up represent increases, arrows down next to N and 0, represent decreases, horizontal arrows represent the temporal direction from precedent to subsequent and often connote a lack of knowledge of regulatory mechanisms, and blunt-ended arrows indicate negative influences. Diagonal and curved lines, and other arrows down, indicate positive influences. Short-dashed curves indicate proteolysis.
23. DEVELOPMENTAL BIOLOGYOF HETEROCYSTS, 2006 bacteria. However, despite the strong effect of ammonium, its suppression of heterocyst differentiation is believed to be indirect. In cyanobacteria, the glutamine synthetase (GS)-glutamate synthase (glutamine oxo-glutarate aminotransferase [GOGAT]) cycle plays a central role in nitrogen metabolism (Herrero et al., 2004). In this cycle, 2-oxoglutarate (2-OG; a-ketoglutarate) and glutamine are used to synthesize two molecules of glutamate, part of which is then used to assimilate ammonium to reform glutamine. Therefore, an increase of glutamine or a depletion of 2-OG could signal that there is a sufficient supply of nitrogen. When assimilation of ammonium by Anabaena cylindrica is inhibited by L-methionineDL-sulfoximine (MSX), an inhibitor of GS, heterocysts form in the presence of ammonium (Rowel1 and Stewart, 1975).Therefore, ammonium must be assimilated to suppress heterocyst differentiation. Furthermore, depletion of glutamine or an increase of 2-OG could indicate an insufficiency of combined nitrogen and could function as a signal for heterocyst differentiation. Glutamine can serve as nitrogen source for growth of Anabaena variabilis while not blocking heterocyst differentiation, suggesting that glutamine “may not be the molecule directly responsible for repression of the differentiation of heterocysts” (Thiel and Leone, 1986). In cyanobacteria, 2-OG is produced by isocitrate dehydrogenase, the product of the gene icd. Cyanobacteria lack 2-OG dehydrogenase, a key enzyme of the citric acid cycle, and use 2-OG mainly for incorporation of ammonium. As first shown in a unicellular cyanobacterium (Muro-Pastor et al., 2001), cellular 2-OG accumulates in various cyanobacteria under nitrogen-limiting conditions. It is now known that 2-OG plays a critical role as a signal for nitrogen status in cyanobacteria. Although microarray results (Ehira and Ohmori, 2006a) showed no significant up-regulation of icd (open reading frame [ORF] alrl827) upon nitrogen deprivation, Muro-Pastor et al. (1996)observed a moderately greater abundance of Icd and of transcripts of icd when Anabaena sp. was grown on N, than when it was grown on ammonium or nitrate. Icd and icd transcripts were especially high in isolated heterocysts. This effect (if found also in cells en route to differentiation), combined with diminished availability of fixed nitrogen, could lead to an increased intracellular concentration of 2-OG. The role of 2-OG in a heterocyst-forming cyanobacterium has recently been elucidated. Li et al. (2003) expressed a gene encoding 2-OG permease in Anabaena sp. strain PCC 7120 (hereinafter denoted Anabaena sp.) and found that the frequency of heterocysts increased in the strain in response to exogenous 2-OG. Laurent et al. (2005) determined that the concentration of cellular 2-OG in Anabaena sp. increased rapidly in response
399
to nitrogen deprivation, to a maximum of approximately 0.1 mM within 1 h after nitrogen step-down. 2,2Difluoropentanedioic acid (DFPA)is a nonmetabolizable analogue of 2-OG. In a strain expressing 2-OG permease, DFPA was found to induce heterocyst formation even in the presence of ammonium. These results constitute strong evidence that as in unicellular cyanobacteria, 2-OG signals nitrogen deprivation in heterocyst-forming cyanobacteria. In nondifferentiating cyanobacteria, 2-OG signals the nitrogen status through both the “NtcA” and the “PII” regulatory circuits (Merida et al., 1991, 1992; Muro-Pastor et al., 2001; Tapia et al., 1996). Much has been learned about the regulatory role of NtcA (Frias et al., 1994; Wei et al., 1994), a transcription factor whose encoding gene is up-regulated in response to nitrogen deprivation. There is a rich literature on NtcA in Anabaena sp.; the reader is referred to a fine review by Herrero et al. (2004) and a commendable alternative view (Muro-Pastor and Florencio, 2003). Su et al. (2005) analyzed the regulatory network of NtcA in nine cyanobacteria whose genomes have been sequenced. A member of the CRP (sometimes known as CAP) superfamily of transcription factors (Herrero et al., 2001; Vega-Palas et al., 1992), Anabaena sp. NtcA responds to nitrogen deprivation by activating the nir-nar operon for uptake and reduction of nitrate and nitrite, as well as genes required for assimilation of urea and ammonium and for heterocyst differentiation (Frias et al., 1994; Herrero et al., 2001, 2004; Luque et al., 1994; Wei et al., 1994).As in the unicellular cyanobacterium Synechococcus elongatus, in which NtcA binds to its own promoter (Tanigawa et al., 2002), Anabaena sp. ntcA is autoregulatory (Muro-Pastor et al., 2002). The autoregulatory increase in transcription of hetR, which is required for heterocyst differentiation, is dependent on NtcA in Anabaena sp. (Muro-Pastor et al., 2002), as is expression of hetC (Muro-Pastor et al., 1999), which is also required early in heterocyst formation (Khudyakov and Wolk, 1997). In Anabaena sp., as in S. elongatus (Tanigawa et al., 2002), the site-specific DNA-binding activity of NtcA (consensus binding site, GTAN,TAC) is enhanced by 2-OG, and in Anabaena sp. also by the nonmetabolizable analogue DFPA of 2-OG (Laurent et al., 2005). Accordingly, even though NtcA is not restricted to heterocyst formation, it is currently the most promising candidate for transduction of the 2-OG signal in heterocyst differentiation. NtcA also affects later stages in the developmental process (see Fig. 1 and below). In proteobacteria, nitrogen deprivation is sensed by the glnB product, PI, (Merrick and Edwards, 1995).PI, is the only component of the proteobacterial PI, regulatory pathway with a cyanobacterial ortholog, suggesting that
400 PI, regulation in cyanobacteria (reviewed by Forchhammer, 2004) differs from that in proteobacteria. Unlike in proteobacteria, PI, in cyanobacteria is modified not by uridylylation but by phosphorylation (Forchhammer and Tandeau de Marsac, 1994; Irmier et al., 1997; Liotenberg et al., 1996). In studying PI, of Anabaena sp., Laurent et al. (2004)showed that although a mutantglnB encoding PIIS49Acannot be phosphorylated, heterocyst formation occurred normally, indicating that heterocyst differentiation does not require phosphorylation of PI,. It was suggested that PI, was probably not involved in heterocyst formation (Laurent et al., 2004). hetR is a master gene that controls heterocyst differentiation (Buikema and Haselkorn, 1991).In its absence, no heterocysts form. When it is overexpressed, clusters of heterocysts form and heterocysts are formed under conditions of nitrogen abundance that would normally suppress heterocyst formation. Positive autoregulation of hetR, which is critical to the process of heterocyst differentiation (Black et al., 1993) because it enforces the decision to differentiate once the process is initiated, is dependent upon ntcA (Muro-Pastor et al., 2002). However, (i) the promoter region of hetR and, in particular, those of its transcription start sites that are ntcA dependent do not contain a canonical NtcA-binding site, nor does NtcA bind, in vitro, upstream from those start sites (Muro-Pastor et al., 2002); and (ii) overexpression of hetR with a copper-inducible petE promoter (Buikema and Haselkorn, 2001) results in heterocyst differentiation even in the presence of ammonium, which reduces expression of ntcA (Muro-Pastor et al., 2002). On the basis of observations (i) and (ii), up-regulation of hetR by NtcA is believed to be indirect. Recent studies suggest that a transcription factor, NrrA (A114312),provides a link between ntcA and hetR. nrrA is rapidly and greatly up-regulated upon nitrogen deprivation (Ehira and Ohmori, 2006a) in an NtcA-dependent fashion (Muro-Pastor et al., 2006).NrrA is also found to recognize a sequence in the promoter region of hetR, and supernumerary copies of nrrA enhance expression of hetR (Ehira and Ohmori, 2006b), whereas up-regulation of hetR is delayed in the absence of nrrA (Ehira and Ohmori, 2006a). The heterocysts that form when hetR is overexpressed in the presence of ammonium are nonfunctional, presumably because processes of heterocyst maturation (see “Heterocyst Maturation” below) depend on NtcA (Herrero et al., 2004). Overexpression of ntcA also produces nonfunctional heterocysts (Olmedo-Verd et al., ZOOS).”
Calcium Ca2+is a universal messenger of signal transduction in eukaryotic cells, where its roles in cellular processes are
ANALOGOUS SYSTEMS well established (Campbell, 1983). With exceptions, e.g., in sporulation of Bacillus (O’Hara and Hageman, 1990) and chemotaxis of Escherichia coli (Tisa et al., 1993), the role of Ca2+in bacterial cellular processes is in general less clear than that in eukaryotes but is receiving increasing attention (Dominguez, 2004; Michiels et al., 2002; Youatt, 1993).An early systematic investigation of a role of Ca2+in cyanobacteria came from study of its role in photosynthetic electron transfer (Becker and Brand, 1985; Brand and Becker, 1988). A structural role of calcium in photosystem I1 was later shown by determination of the three-dimensional structure of a cyanobacterial photosystem I1 (Zouni et al., 2001). Torrecilla et al. (2000) constructed a [Ca2+],reporter system with recombinant aequorin and showed that the [Ca”], in Anabaena sp. was maintained at approximately 100 nM, a concentration similar to that in eukaryotes, suggesting that a mechanism of calcium homeostasis in cyanobacteria exists. Two unicellular cyanobacteria have genes that may encode P-type Ca2+pumps (Berkelman et al., 1994; Geisler et al., 1993), multiple orthologs of which are found in all cyanobacterial genomic sequences thus far reported, including those of the heterocystforming strains Anabaena sp., A. variabilis, and Nostoc punctiforme. A requirement for Ca2+in heterocyst differentiation was first suggested by Wood and Haselkorn (1979) in their study of calcium-dependent proteases required for protein turnover early in heterocyst differentiation. The gene encoding one calcium-dependent protease was later cloned and sequenced (Maldener et al., 1991) and was found to be inessential for heterocyst formation. Smith et al. (1987) observed that the exogenous Ca2+ concentration influenced the frequency of heterocysts in Nostoc sp. strain PCC 6720, suggesting that Ca2+is a critical factor in heterocyst differentiation; a similar study was reported by Zhao et al. (1991). Torrecilla et al. (2004)found that a specific Ca2+signalwas observed upon nitrogen deprivation of Anabaena sp. Their use of various inhibitors of calcium-binding proteins demonstrated that Ca2+is critical to cellular differentiation in Anabaena sp. HetR is a serine-type protease that binds Ca2+(Zhou et al., 1998b); its protease activity depends on Ca2+(Y. Shi and J. Zhao, unpublished data). Early reports of calmodulin-like Ca2+-bindingproteins in heterocyst-forming cyanobacteria based on indirect assays and immunology (Onek and Smith, 1992; Pettersson and Bergman, 1989) may have been attributable to other Ca2+-bindingproteins. However, Onek et al. (1994) isolated and characterized a 21-kDa calmodulinlike protein from Nostoc sp. strain PCC 6720 that could activate pea NAD kinase, as does eukaryotic calmodulin,
23. DEVELOPMENTAL BIOLOGYOF HETEROCYSTS, 2006 and cross-reacted with antibodies raised against spinach calmodulin. Their study provided the first biochemical evidence of calmodulin activity of a calcium-binding protein from a heterocyst-forming cyanobacterium. The gene encoding the protein has not yet been reported. Anabaena sp. strain ATCC 29151 (PCC 7119) has an adenylate cyclase activity that is activated by Ca2+-calmodulin and has acalmodulin-like activity, and so it is a prokaryote “in which an adenylate cyclase is regulated by an endogenous calmodulin-like activity” (Bianchini et al., 1990). Recently, a Ca2+-bindingprotein, CcbP, was isolated and characterized in Anabaena sp. (Zhao et al., 2005). CcbP is present in other heterocyst-forming cyanobacteria and absent from unicellular cyanobacteria. CcbP has 126 amino acid residues and is acidic, with a PI of 4.2. It contains no EF hand; its calcium-binding ability is probably due to its surface charge. When cc6P was deleted, filaments of Anabaena sp. deprived of nitrogen formed multiple contiguous heterocysts (the Mch phenotype). When cc6P was overexpressed, hetR up-regulation was suppressed and heterocyst formation was not observed. Overexpression in Anabaena sp. and A. variabilis of the gene encoding rice calmodulin suppressed heterocyst formation in both cyanobacteria, suggesting strongly that CcbP regulates heterocyst differentiation by sequestering Caz+.By using recombinant obelin as a reporter of Ca2+,Zhao et al. (2005) demonstrated that the overall Ca2+concentration in filaments increased starting at about 4 h after nitrogen step-down. Thereafter, but still before proheterocysts could be observed, individual cells with an increased [Ca2+Iicould also be observed, and a pattern of cells with higher [Ca2+Iiwas established along the filaments of Anabaena sp. (Color Plate 10). These results showed that the increase of [Ca2+Iiis an early signal in heterocyst differentiation. Because heterocysts were likely present 2 billion years ago (Tomitani et al., 2006), the involvement of calcium in heterocyst differentiation suggests that calcium played an important role in cellular differentiation, probably prior to the evolution of eukaryotes.
Other Signals, Including in Symbiosis Some nitrogen-fixing cyanobacteria establish symbiotic relationships with different plants and fungi. These cyanobacteria, mostly in the genus Nostoc, are able to form symbioses with bryophytes, ferns, gymnosperms, and angiosperms and provide fixed nitrogen to the plants (for a review, see Meeks and Elhai, 2002). The symbiotic association between cyanobacteria and plants is usually not as specific as in root nodules of legumes: a cyanobacterial strain isolated from one symbiotic association can often establish a symbiosis with another plant. How-
401
ever, one common consequence of the establishment of a symbiotic relationship of plants with cyanobacteria is the induction of heterocyst formation. The regulation of heterocyst differentiation of cyanobacteria in symbiosis seems to be more complex than that in free-living cyanobacteria. Heterocyst frequency in symbiosis is often much higher than that in free-living cyanobacteria, and much of the nitrogen fixed by heterocysts is excreted. Unlike in free-living cyanobacteria, combined nitrogen often does not directly repress heterocyst formation in cyanobacteria in symbiosis. Meeks and Elhai (2002)suggested that there is a specific plant signal for induction of heterocyst formation and that this signal enters the normal heterocyst control circuit at the stage of ntcA or earlier. No such putative plant signal required for heterocyst formation in symbiosis has yet been identified. The concentration of phosphorylated guanosine (ppGpp) increases within 30 min of nitrogen deprivation (Akinyanju and Smith, 1979, 1987). In some bacteria, ppGpp is important for cellular responses to environmental stress. For example, cells of Bacillus sp. are not able to differentiate normally when the ppGppdependent stringent response is impaired (Ochi et al., 1981). However, the significance of the increase of ppGpp in heterocyst-forming cyanobacteria is unclear and it has not been reported that ppGpp production is required for heterocyst development. Cyclic AMP is a key molecule in signal transduction in both bacteria and eukaryotes, and the level of cyclic AMP is responsive to nitrogen deprivation (reviewed by Wolk et al., 1994). Cyclic AMP was also observed to influence the heterocyst pattern of A. variabilis (Smith and Ownby, 1981), but its role in signaling heterocyst differentiation is unclear. Anabaena sp. contains several potential adenylate cyclase genes and approximately a dozen genes that could be involved in cyclic dinucleotide synthesis (Kaneko et al., 2001). Because cyclic di-GMP is an important messenger in various cellular processes in bacteria (D’Argenio and Miller, 2004), and orthologs of the corresponding GGDEF domain (http://www.ncbi.nlm.nih.gov/Structure/cdd/ cddsrv.cgi?uid =cdO 1949&version =v2.06) are present in 14 ORFs of Anabaena sp., it will be interesting to ascertain whether any of these ORFs function in heterocyst formation.
HETEROCYST PATTERN FORMATION Initiation of Differentiation The pattern of spaced heterocysts is a strikingly simple example of a multicellular pattern. Heterocysts, which lack active photosystem I1 and Rubisco, derive fixed
402 carbon (Elhai and Wolk, 1990; Wolk, 1968; Wolk et al., 1994) and certain other nutrients from vegetative cells; and when there is a paucity of fixed nitrogen in the environment, vegetative cells depend on nitrogen fixed by heterocysts. The semiregularity of the pattern presumably facilitates the exchange of materials between heterocysts and vegetative cells. Which cells differentiate and which do not requires regulatory processes that are being extensively investigated. Fritsch (1951)wrote: As a working hypothesis it may be suggested that heterocysts during the vegetative period secrete substances that stimulate growth and cell-division and, when the concentration of these substances falls off as the cells come to lie further and further away from heterocysts, production of other heterocysts is induced. When the reproductive period sets in, the nature of the secretion from the heterocysts changes, and something is formed that stimulates akinete formation; or perhaps there is merely an alteration in the reaction of the contents of the cells involved. The idea, implicitly proposed by Fritsch, that heterocysts inhibit nearby cells from becoming heterocysts was later established experimentally (Wilcox et al., 1973; Wolk, 1967). The simplest mechanistic model that has been proposed (Fay et al., 1968) to account for the perpetuation of the pattern during growth is that a product of N, fixation acts, as does exogenously supplied NH,+ or a derivative of it, as an inhibitor of heterocyst formation. A possible alternative (or simultaneous) model is that vegetative cells produce a readily diffusible stimulator of heterocyst formation. One could imagine that stimulator being 2-OG, with heterocysts acting as localized sinks for that stimulator perhaps by producing glutamine that reacts in vegetative cells with 2-OG to produce glutamate (see above). Thomas et al. (1977) concluded that heterocysts of A. cylindrica transfer glutamine from heterocysts to vegetative cells in exchange for glutamate coming from vegetative cells. Their conclusion was strengthened by the results of enzymatic assays, Western blots, and Northern blots that showed that ferredoxin-dependent glutamate synthase is absent from, but glutamine synthetase and isocitrate dehydrogenase are present in, heterocysts of Anabaena sp. (Martin-Figueroa et al., 2000). Reaction with NH,+ would act as a metabolic sink for the same stimulator, be it 2-OG or another substance. Neither of these models is appropriate for the initial establishment of the pattern, prior to the inception of fixation of dinitrogen, unless (see Fleming and Haselkorn, 1974) release of nitrogen from some polymerized form such as protein is characteristic of a very early stage of differentiation. It remains controversial whether the
ANALOGOUS SYSTEMS establishment of the pattern and its perpetuation are based on the same mechanism (Thiel and Pratte, 2001; Yoon and Golden, 2001). Progress has been made in understanding both heterocyst differentiation and its inhibition.
HetR, Pats, and Their Interactions As noted above, hetR is central to the control of heterocyst differentiation (Buikema and Haselkorn, 1991). However, orthologs of hetR are present also in some cyanobacteria in which heterocysts do not differentiate (Buikema and Haselkorn, 1991; Janson et al., 1998; Lundgren et al., 2005; Mes and Stal, 2005; Zhou et al., 1998a). Its function in those organisms is unknown. In heterocyst-forming cyanobacteria, hetR mutants can grow with combined nitrogen at a rate similar to that of the wild type and show no sign of differentiation when transferred to a medium free of combined nitrogen. When filaments of Anabaena sp. are deprived of fixed nitrogen, up-regulation of hetR can be observed within 1h and the concentration of hetR mRNA is maximal within 3 h (Buikema and Haselkorn, 1991; Huang et al., 2004). Among the genes that function early in the differentiation process, hetR is not the only gene that shows positive feedback in regulation of gene expression. The expression of ntcA is dependent on the presence of an intact ntcA (Ramasubramanian et al., 1996), and the expression of hetC may also be positively autoregulated (Khudyakov and Wolk, 1997). However, only for hetR is it known that its positive autoregulation cannot be bypassed. HetR, the product of hetR, is a serine-type protease (Zhou et al., 1998b) whose active serine residue in Anabaena sp. is Ser152, as determined by labeling with dansyl fluoride (Dong et al., 2000) and the observation that HetR-S152A mutants do not self-digest. Autodegradation was also absent in a HetR-S179N mutant (Zhou et al., 1998b), suggesting that Ser179 is also involved in proteaseactivity. BeyondHetRitself (Zhouetal., 1998b), the only identified substrate of HetR proteolysis is CcbP; its degradation releases calcium, enhancing HetR activity (Shi et al., 2006). Although HetR has no predicted DNAbinding motif, recent studies have shown that HetR, like Lon (Bretzet al., 2002), is a DNA-binding protein (Huang et al., 2004). The DNA-binding activity of HetR requires dimerization of HetR through its sole cysteine residue (Cys48 in Anabaena sp.). Replacing Cys48 with other residues abolishes dimerization of HetR and its DNAbinding activity. In Anabaena sp., both monomer and dimer can be observed in vivo, suggesting that the two forms of HetR are in equilibrium. In strain C48 of Anabaena sp., which carries a hetR gene with a C48A mutation, filaments could not initiate differentiation
23. DEVELOPMENTAL BIOLOGYOF HETEROCYSTS, 2006 when deprived of fixed nitrogen (Huang et al., 2004). Northern blotting showed only limited expression of hetR and only a single transcript, in contrast to three major transcripts in the wild type. Real-time PCR showed that up-regulation of hetR was limited to the first several hours of nitrogen deprivation; the level of its total mRNA then declined to the level before induction. Certain findings concerning Serl52 and Cys48 have been challenged (Risser and Callahan, 2007); why discrepant results were obtained remains unclear. HetR binds DNA fragments from the promoter regions of hetR, hepA, and pats. Because the expression of these genes requires active HetR, HetR may function as a transcriptional activator or may degrade a repressor(s)that also binds to the promoter regions of these genes. Isoelectrofocusing gel electrophoresis showed that there are several isoforms of HetR with different PI values (Zhou et al., 1998b), suggesting that HetR may be modified in vivo. The nature of the modification remains to be determined. One candidate for modifying HetR is PatA (see “HetN, HetF, and PatA” below). Wilcox et al. (1973) noticed that the initiation of heterocyst differentiation in Anabaena catenula and A. cylindrica seemed to start with contiguous proheterocysts. One cell in a string of such cells continues to differentiate while the others reinitiate vegetative growth. It was speculated that the cell that continues to differentiate also influences neighboring cells to revert to a vegetative condition. How series of one or more cells are chosen to initiate differentiation is unknown. Mitchison et al. (1976) observed that in A. catenula, heterocyst differentiation occurred in cells only 6 to 8 h after cell division, implying that the cell cycle may influence the choice of cells to start differentiating in response to nitrogen deprivation. Recent study of ftsZ of Anabaena sp. also suggested that the cell cycle may be a factor in regulation of differentiation (Kuhn et al., 2000). SulA of E. coli is a protein that inhibits the GTPase activity of FtsZ and prevents FtsZ-ring formation. By expressing the E. coli sulA gene in Anabaena sp., Sakr et al. (2006) showed that cell division is required for heterocyst formation, emphasizing the possibility that the cell cycle may play a regulatory role in heterocyst development. One factor produced by heterocysts and differentiating cells that negatively influences the differentiation of neighboring cells is Pats, the product of pats. The pats gene, first reported by Yoon and Golden (1998), from Anabaena sp. is located on one of two large fragments that suppress heterocyst formation when present in a multicopy plasmid (Bauer et al., 1997). The gene encodes a 17- or 13-amino-acid peptide, depending upon which Met codon is used by the translation machinery. Pats may be further trimmed to a smaller peptide (Wu et al., 2004). Overexpression of pats
403
results in complete suppression of heterocyst formation, whereas deletion of pats leads to an Mch phenotype upon nitrogen deprivation. When any one of four of the C-terminal five residues (RGSGR) was mutated, the resulting clone could no longer suppress heterocyst formation. Addition of synthetic pentapeptide RGSGR to the medium of Anabaena sp. at a concentration of 0.1 p M fully suppressed heterocyst formation under conditions of nitrogen deprivation. The expression of pats is restricted to those cells that have at least initiated differentiation to mature heterocysts (Yoon and Golden, 1998) and is dependent upon an active hetR (Huang et al., 2004). The up-regulation of pats started approximately 6 h after nitrogen deprivation, several hours after hetR (Huang et al., 2004; Yoon and Golden, 2001). It has been speculated, but not yet shown, that part or all of Pats moves from its cell of origin to neighboring cells (Yoon and Golden, 1998). The pats product may suppress heterocyst formation by inhibiting the DNA-binding activity of dimeric HetR: in vitro, synthetic RGSGR inhibits gel shifting by HetR dimers (Huang et al., 2004). Because DNA-binding by HetR dimers is required for heterocyst formation, inhibition of DNA-binding by HetR dimers in vivo may lead to suppression of heterocyst differentiation. The pats product is capable of inhibiting the function of HetR downstream as well as upstream of hetR transcription, because overexpression of pats in Anabaena sp. is epistatic to overexpression of hetR (Orozcoet al., 2006). Additional evidence that pats suppresses heterocyst formation through HetR comes from the report by Khudyakov and Golden (2004). In searching for mutants of Anabaena sp. whose heterocyst differentiation is not suppressed by Pats, they isolated a strain carrying a mutant hetR gene encoding HetR with an R223W mutation. The strain showed no sign of suppression of heterocyst formation by pats and hetN (see “HetN, HetF, and PatA” below) and had an Mch phenotype. Heterocysts were randomly distributed, suggesting that hetR is also critically involved in control of heterocyst pattern formation. Therefore, the HetR-Pats interaction plays a key role in establishing heterocyst pattern and, as is discussed next, this interaction is one of the best examples of regulation of biological pattern formation.
Turing’s Model of Pattern Formation More than a half century ago, Turing (1952)provided a mathematical model to elucidate biological pattern formation. He showed that the interaction of two substances with different diffusion rates can generate a pattern of concentrations of the substances even when the initial distribution is nearly uniform. This important discovery was little noticed at that time in the field of developmental
ANALOGOUS SYSTEMS
404 biology, not to mention in biology in general. Subsequently, Gierer and Meinhardt (1972)proposed a model system for biological pattern formation based on reaction-diffusion dynamics. Their model, which was similar to the model proposed by Turing (1952),emphasized the activating and inhibitory effects of the substances. A key contribution of their model was the recognition of the great importance of autocatalysis and lateral inhibition in biological pattern formation. Pattern formation in many organisms may be explained or predicted by their model (Meinhardt and Gierer, 2000). Many early researchers, eg., Wigglesworth (1940), recognized the importance of lateral inhibition by an inhibitory substance to explain biological spacing patterns. However, lateral inhibition alone could not solve the problem of pattern formation because it could not explain why the inhibitory substance does not prevent differentiation of the cells or tissues that produce it. For example, why does Pats not inhibit the differentiation of the cells that produce PatS? That question can be answered by the models proposed by Turing (1952) and Gierer and Meinhardt (1972), in which they covered several important points. First, two substances are produced, a self-enhancing substance (the “activator”) that has a short or restricted transfer range, and a second substance (the “inhibitor”) that inhibits production of the activator and is laterally distributed with a relatively long transfer range. Second, a pattern can be generated from a nearly uniform distribution (or fluctuations from a uniform distribution) and is not dependent upon a preexisting pattern. At the final stage of pattern formation, the activator will have a high local concentration surrounded by the inhibitory substance. Third, because degradation of the activator increases as the concentration of the activator increases, a nonlinear factor should govern production of the activator. The local concentration of the activator could then increase even in the presence of the inhibitory substance. The nonlinear factor might, for example, result from dimerization of the activator or amplification of gene expression through an expression cascade. The simplest case for pattern formation would consist of an autocatalytic activator and an inhibitory substance whose production is dependent on the activator. Among many examples of biological pattern formation, the interaction of HetR and Pats is the simplest, and one of the best studied, systems that can be described by the models of Turing (1952) and Gierer and Meinhardt (1972). First, hetR is positively autoregulatory, its up-regulation depending on the presence of a functional hetR (Black et al., 1993). Second, HetR is a protease with autodegrading activity (Zhou et al., 1998b). The instability allows HetR to disappear once the
up-regulation of hetR is not sustainable or is inhibited by a repressor. Third, the up-regulation of hetR depends on the formation of HetR homodimer. As mentioned above, dimerization is a nonlinear factor that allows positive feedback. Fourth, pats up-regulation depends upon a functional hetR and lags several hours behind that of hetR (Huang et al., 2004). Finally, Pats inhibits hetR by preventing the DNA-binding activity of HetR. What has not yet been shown is that Pats has the lateral mobility predicted for the inhibitor, so that it would not steadily increase in concentration but would be dispersed to contiguous cells where it would inhibit transcription of hetR. Nonetheless, the above features approach the predicted explanation of why Pats does not inhibit the function of HetR in the cells that produce PatS. That said, the reality of biological decision making and stabilization appears to be, as discussed in the next section, considerably more complex than the model. Meeks and Elhai (2002) proposed a two-stage model for heterocyst formation, in which the first stage is initiation of groups of cells, possibly siblings at a similar phase of the cell cycle, and the second stage is competitive resolution. Except that Turing’s model does not invoke co-initiation of grouped cells, this model is in general agreement with Turing’s. It is even possible that all cells may initiate differentiation in response to nitrogen step-down and that resolution halts the differentiation of many cells, sometimes leaving the strings of cells observed by Wilcox et al. (1973) initiating differentiation along the filaments. Khudyakov and Golden (2004) and Borthakur et al. (2005) observed, respectively, that overexpression of a hetR-R223 W gene in a hetRR223 W background and simultaneous mutation of pats and hetN led to differentiation of nearly all cells. Importantly, however, neither group observed simultaneous differentiation. (There are significant differences in the distributions of numbers of contiguous vegetative cells in the mutants studied by these two groups, especially after protracted periods of nitrogen deprivation. Moreover, the first group presented evidence that differentiation was random, whereas the second group concluded that differentiation was not random but, rather, biased toward clustering of heterocysts.) It is possible that the genetic manipulations somehow disturbed regulation of the cell cycle, leading to altered initiation or resolution during heterocyst formation.
HetN, HetF, and PatA HetN and HetF are thought to be involved in the autoregulation of hetR. Despite data suggestive of interconversion of wild-type and mutant alleles of hetN, Black and Wolk (1994)concluded:
23. DEVELOPMENTAL BIOLOGYOF HETEROCYSTS, 2006 Because most insertions within hetN result in the production of multiple contiguous heterocysts, while supernumerary copies of hetN suppress heterocyst formation, a substance produced or modified by the product of hetN may inhibit heterocyst differentiation. Such a substance may even mediate intercellular interactions that regulate development. In an elegant piece of work, Callahan and Buikema (2001) expressed hetN from a Cu2+-regulatable petE promoter, so that a mutant could be isolated with hetN active and the mutant phenotype then observed upon inactivation of the promoter by lowering the concentration of Cu2+.They observed that overexpression of hetN blocked normal expression of hetR and prevented heterocysts from forming, but in the absence of expression of hetN, a pattern of multiple contiguous heterocysts was observed 48 h after the onset of nitrogen deprivation. Having also observed that in wild-type filaments hetN is expressed principally in heterocysts, these authors concluded that the normal function of hetN is to prevent the formation of contiguous heterocysts after initial pattern formation has taken place but that overexpressed HetN can prevent normal autoregulation of hetR. Like hetR, hetF, heretofore studied virtually only in N. punctiforme, is required for heterocysts to differentiate in response to nitrogen deprivation, and supernumerary copies of hetF lead to the formation of clustered heterocysts. In a hetF mutant, nitrogen deprivation leads to expression of Phctn::gfpin all cells, although hetR is expressed more slowly than in the wild-type strain. In the presence of abundant nitrogen, a plasmid-borne hetR::gfp fusion that is expressed heterocyst-specifically in a nitrogen-deprived, wild-type strain is also expressed in all cells. It appears, therefore, that HetF facilitates autoregulation of hetR and is required for accumulation of the HetR protein in differentiating heterocysts (Wong and Meeks, 2001). A PatA mutant has the interesting phenotype that heterocysts form nearly exclusively from the terminal cells of filaments (Liang et al., 1992), implying that PatA is required for the formation of intercalary heterocysts. Even when hetR is overexpressed from a petE promoter, which in a wild-type background elicits heterocyst formation in the presence of NH,+, a patA mutant does not form intercalary heterocysts (Buikema and Haselkorn, 2001). More general tests of epistasis were interpreted as suggesting that PatA antagonizes the negative effects of HetN and Pats on differentiation and also stimulates differentiation independent of its antagonistic effect on the activities of HetN and Pats (Orozco et al., 2006). PatA resembles two-component response regulators that lack
405
a DNA-binding motif and is therefore thought to interact with some other molecule in the cell. What that molecule may be in the case of PatA is unknown, but Buikema and Haselkorn (2001) suggest that PatA may lead, perhaps indirectly, to posttranslational modification, possibly phosphorylation, of HetR. A mutant allele of hetR that, in certain genetic backgrounds, results in a patA-like phenotype increases the likelihood that PatA may, directly or indirectly, modify or interact with HetR (unpublished results cited by Orozco et al., 2006). Whether overexpression of patA would result in a phenotype different from that of the wild-type strain, perhaps with more closely spaced heterocysts, has not yet been determined.
Gradients of Fixed Nitrogen Although Pats mediates spacing initially upon nitrogen step-down, its influence decreases over time, an effect attributed by Yoon and Golden (2001) to products of nitrogen fixation controlling the pattern. Others argue, however (see above), that the decrease in influence is due to an effect on the hetR-(positive-)autoregulatoryloop by HetN produced in heterocysts (Callahan and Buikema, 2001; Li et al., 2002). Also possibly in conflict with the interpretation of Yoon and Golden are results of Thiel and Pratte (2001).A. variabilis has two molybdenum-containing nitrogenases, whose polypeptides are encoded by .if2 and nif2, respectively. The .if2 genes, like the nif genes of Anabaena sp., are expressed specifically in heterocysts, whereas the nif2 genes are expressed in all cells, but only under anoxic conditions (Thiel et al., 1995). Wild-type A. variabilis, a .if2 mutant, and a mutant incapable of forming heterocysts were grown with ammonium and shifted under anoxic conditions to a medium that lacked combined nitrogen but contained fructose and N,. All three grew exponentially, producing what, for the wild-type strain and the nifl mutant, appeared to be a normal pattern and number of heterocysts (Thiel and Pratte, 2001). These authors considered the simplest interpretation of their results to be that (i) no product of nitrogen fixation controls heterocyst pattern formation, (ii) pattern formation does not depend on a gradient of fixed nitrogen that diffuses along the filament, and (iii) starvation for fixed nitrogen is not a prerequisite for heterocyst differentiation. Perhaps the following speculation is worth consideration. Glutamine may be produced principally by heterocysts (see “Initiation of Differentiation” above) even in a .if2 mutant, while the fructose that was fed may have increased the concentration of heterocyst-eliciting (Laurent et al., 2005) 2-OG throughout a filament. If
Next Page
406 the glutamine is metabolized together with 2-OG by glutamate synthase within vegetative cells adjacent to heterocysts, the concentration of 2-OG near heterocysts could be less than in cells farther away in those filaments (Thiel et al., 1995), resulting in a normal pattern of heterocyst formation.
HETEROCYST MATURATION Genes That Encode Catalytically Active Proteins Heterocyst maturation is depicted in Fig. 1. Ehira et al. (2003), in a ground-breaking paper, scanned along the genome of Anabaena sp., looking for ca. 3-kb regions, derived from M13mp18 clones that contain part or all of up to eight chromosomal genes, whose expression showed in excess of a fivefold increase in transcription as of 1, 3, 8, or 24 h of nitrogen deprivation. They probed their arrays with cDNAs from intact filaments that had been deprived of fixed nitrogen for 0,1,3,8, and 24 h, and with probes generated from RNA extracted from isolated, mature heterocysts. Despite the low frequency of developing heterocysts as a fraction of the total population of cells, and despite the disadvantage that highly significant change of expression of one gene on an element could be obscured by strong, and perhaps oppositely directed, expression of others on the same element, gene clusters were observed to be up-regulated, and others down-regulated, in response to nitrogen deprivation (Ehira et al., 2003). By this analysis, they identified several clusters of highly expressed genes. Subsequently, when gene-by-gene microarrays became available for their studies, such experiments were repeated within the context of analysis of a newly discovered nitrogen-regulatory response-regulator, NrrA (Ehira and Ohmori, 2006a), and the results were presented in supplementary tables on the Web. As of 1 h following nitrogen step-down, Ehira et al. (2003) found only a single genetic cluster highly expressed, namely, the region of nitrogen uptake and reduction genes nirA(alr0607)nrtABCD- narB(alr0612)that had earlier been identified (Cai and Wolk, 1997) as rapidly activated, in response to nitrogen deprivation, by use of transposons bearing ZuxAB as a reporter of transcription. The 37 ORFs that showed about a fivefold or greater average increase of expression at 3 h and the 59 ORFs that showed about an eightfold or greater average increase at 8 h were less strongly expressed at 24 h. These ORFs included the following. (i) By 3 h, clustered ORFs a112004-alZ2006, which putatively encode a cation-transporting ATPase and unknown proteins, and most ORFs in a more protracted region, ah3057 through alr3073, many of which putatively encode glycosyl transferases (Ehira and Ohmori,
ANALOGOUS SYSTEMS 2006a). (The ORFs of the Anabaena sp. chromosome are numbered consecutively from an AvrII site [Kaneko et al., 20011; the letters a, 1 [or s], and r [or I] preceding the number of the ORF refer to the genus, the length of the ORF, and its orientation.) It is notable that in a near-saturating transposon mutagenesis of Anabaena sp. directed at identifying genes required specifically for heterocyst differentiation and function (Q. Fan and C. P. Wolk, unpublished data), only a single transposon was found within the latter region, well within the frequency of false positives found in that screen. In contrast, 63 transposon insertions were found within a region required for synthesis of heterocyst envelope polysaccharide. As suggested also by the rapid expression of the region aZr3057-aZr3073 in response to nitrogen deprivation, its ORFs are probably not required specifically for aerobic fixation of dinitrogen. Intriguingly, this same series of ORFs is extensively inactivated in response to several regulatory mutations that block heterocyst maturation (Lechno-Yossef et al., 2006). It is unclear what polysaccharide may be synthesized by the proteins that these ORFs presumably encode. (ii)Two cytochrome oxidase respiratory genes, coxA2 (aZr2515) and coxC2(aZr2516). The cox11 and cox111 respiratory genes may be induced specifically in heterocysts (and differentiating heterocysts) upon nitrogen step-down. Only if both of these sets of genes are inactivated does Anabaena lose its ability to grow diazotrophically (Valladares et al., 2003). PatB, encoded by the nearby ORF a11251 2, has a helix-turn-helix motif and, like the product of the adjacent ORF, asr2513, has characteristics of a ferredoxin. Liang et al. (1993)suggested that PatB “may function as a transcriptional regulator in response to the redox state or other redox factors in the cell.” Subsequently, patB was shown (Jones et al., 2003) to be expressed primarily, perhaps exclusively, in heterocysts. Jones and coauthors determined that whereas point mutations in patB resulted in very slow growth on N,, a patB deletion mutant almost completely stopped growing on N, within 24 h and formed multiple contiguous heterocysts. However, whereas a frameshift mutation of patB strikingly reduced transcription of the nearby cox11 locus, alr251 4-aZr2516, deletion mutants of PatB had very little effect on transcription of that locus. (iii) ah2825 to alr2841, and alr3699. ah3699 is contiguous with hepB (alr3698) and is itself a Fox gene (Wang et al., 2007), i.e., a gene required specifically for nitrogen fixation in the presence of oxygen. Many of these ORFs are required for synthesis of heterocyst envelope polysaccharide (Huang et al., 2005). (iv)alr3712, encoding devA, one of a cluster of ABC transporter genes whose products have been proposed to
Previous Page
23. DEVELOPMENTAL BIOLOGYOF HETEROCYSTS, 2006 export either heterocyst envelope glycolipid or possibly a scaffold for the deposition of those glycolipids (Fiedler et al., 1998; Maldener et al., 1994). Notably, the genes for synthesis and deposition of those glycolipids are expressed more strongly at 24 h than at 8 h (see below). (v) a110688, the gene hupS, which, together with hupL, encodes an uptake hydrogenase that captures and recycles the reducing equivalents of hydrogen gas wastefully produced by nitrogenase. (vi) n i p , n i p , and nifK of nitrogenase (aka all1455, all1454, and all1440 [Ehira and Ohmori, 2006a; Lammers and Haselkorn, 1983; Mazur and Chui, 1982; Mevarech et al., 1980]),plus, according to the paper by Ehira et al. (2003),chromosomal segments whose products are involved in synthesis of the iron-molybdenum cofactor of nitrogenase and other niffunctions (Buikema and Haselkorn, 1993); cotranscribed genes nifB and fdxN (Mulligan and Haselkorn, 1989),aka alll 51 7 and alll 516, respectively; and xisA (aka alr1442), whose product removes a DNA element from within n i p (Golden, 1998). (vii)alr2732, one of the coxIII series of genes involved in respiration (Jones and Haselkorn, 2002; Valladares et al., 2003). (viii) a115342-alr5358, which include genes whose products synthesize the aglycones of the heterocyst envelope glycolipids (Hgls); other genes that are involved in deposition of those glycolipids (Fan et al., 2005); and hetN, which plays a role in preventing the formation of clusters of heterocysts (Borthakur et al., 2005; Callahan and Buikema, 2001). Genes identified as up-regulated in mature, isolated heterocysts (Ehira et al., 2003) suggested extension of the hgl region in both directions, to a115341-ah53 60, thus including a115341. The predicted product of a115341 is a glycosyl transferase, and inactivation of that ORF blocks glycosylation of the aglycone portions of the heterocyst envelope glycolipids (Awai and Wolk, 2007). Putative start sites for transcription of a115347 (hgdB) and alr.5351 (hglE,) have been identified in N2-grown, wild-type PCC 7120 and found to be up-regulated by 0.2 M NaCl, but not in a cyaC mutant (CyaC is an adenylate cyclase that contains two-component hybrid sensor and regulator domains: Imashimizu et al., 2005). The functions of numerous ORFs activated at 3,8, and 24 h remain obscure. For example, of the 80 ORFs whose expression was observed (Ehira and Ohmori, 2006a) to have increased, on average, fourfold or more at 3 h of nitrogen-deprivation, 42 are annotated as “unknown” or “hypothetical”; of 139 ORFs that were first activated, on average, fourfold or more at 8 h, but not as of 3 h, 64 are similarly annotated; and of 97 ORFs that were first
407
activated an average of fourfold or more at 24 h, but not as of 8 h, 51 had those same annotations (grand total: 157 of 316 ORFs). (It should be recognized that visualization of a fourfold increase by use of microarrays is apt to be an underestimate due to background. For example, Northern blotting enabled Jones et al. [2003] to identify a 35-fold increase in coxll transcript over the time course of differentiation [normalized to 23s rRNA], whereas microarray data brought to light an average of 12.5- and 8.8-fold increases for coxA2 and coxC2, respectively.) Evidently, much remains to be learned about functions that are exercised during heterocyst maturation. Genetic segments down-regulated in heterocysts visA-vis whole filaments (Ehira et al., 2003) were shown to include genes petE, aroK, psbV, psbAI, and rbcLXS. Their down-regulation presumably reflects, in part, the loss of evolution of 0, by photosystem I1 (encoded by psb genes) and of activity of ribulose bisphosphate carboxylase (encoded by rbc genes) in mature heterocysts (see paragraph 1 of “Heterocyst Pattern Formation” above). Biliproteins that comprise photosynthetic antennae and are among the most abundant proteins in vegetative cells are degraded, in part to serve as a nitrogen source (Wood and Haselkorn, 1980), in response to nitrogen deprivation. Their genes are soon conspicuously inactivated in response to nitrogen deprivation (Ehira and Ohmori, 2006a) but are later reactivated once nitrogen fixation has begun.
HetC and HetP Because transcriptional events have great importance for the establishment of the pattern of spaced heterocysts, and for the progression of the process of heterocyst differentiation, we now consider the regulatory pathways that control changes in transcription. HetC is an unlikely candidate for a regulatory molecule, because its product is similar to ATP-binding cassette transporters of proteins, peptides, and polysaccharides. Nonetheless, HetC acts at an intermediate point of differentiation because a hetC mutant appears by light microscopy to be devoid of differentiation, but when viewed by fluorescence microscopy, it shows a pattern of spaced cells that exhibit little or no biliprotein-derived fluorescence (hereafter we refer to these cells as having low autofluorescence) but are foci of expression of a hetR::gfp fusion (Xu and Wolk, 2001). Contrariwise, a hetC mutation blocks expression of fusions of gfp to heterocyst differentiation genes hglD and hglE, which are involved in biosynthesis of heterocyst envelope glycolipid, and to patB, nifB, and xisA (Wang and Xu, 2005; see above). A hetC mutation also results in very low expression of patA::gfp (Wang and Xu, 2005). According to the model of Orozco
408
et al. (2006),very low expression of patA would enhance the differentiation-inhibitory activity of pats and hetN. Thus, a hetC mutant achieves a nearly mature state of pattern formation, but a very immature state of cellular differentiation. One very important way in which its differentiation is immature is that the cells with low autofluorescence, unlike mature heterocysts, remain capable of cell division (Wang and Xu, 2005; Xu and Wolk, 2001), producing pairs or longer groupings of cells that are diminished in size (Xu and Wolk, 2001). In this way, they are reminiscent of cells of the motile filaments known as hormogonia (Meeks and Elhai, 2002; Tandeau de Marsac, 1994). Whereas heterocysts lack the principal cell division protein FtsZ (Kuhn et al., ZOOO), the cells with low autofluorescence show enhanced fluorescence of an ftsZ::gfp fusion (Wang and Xu, 2005). Of much interest are the observations that transcriptional activation of hetC is under the control of ntcA (Muro-Pastor et al., 1999), but not of hetR (Muro-Pastor et al., 2002). hetP is the first gene 3' from hetC in the genome. As of 8 h after nitrogen step-down, the level of hetP message increased an average of 89-fold (Ehira and Ohmori, 2006a). A hetP mutation, like a hetC mutation, results in a Het- phenotype by light microscopy, suggesting that HetC and HetP act in concert, but shows a less clear pattern of little-fluorescent cells. A complemented hetP mutant shows, relative to a wild-type strain, an increased propensity for the formation of short series of contiguous heterocysts (Fernindez-Piiias et al., 1994).
Later-Acting Regulatory Proteins Later-acting regulatory proteins are listed in Table 1. The essence of being a heterocyst is to provide a nitrogenase-friendly environment, i.e., (i) 0, production by photosystem I1 is inactivated (Wolk et al., 1994), (ii) respiration is enhanced (Wolk et al., 1994),and (iii)an envelope is deposited that comprises a Hep layer and a Hgl layer. Mutations that lead to loss of one or the other of the two kinds of envelope layer are referred to as having a Hep- or Hgl- phenotype, respectively. The Hgl layer, which is deposited within-and therefore locally after-the Hep layer, largely blocks penetration of 0, (Walsby, 1985). Ultrastructural examination of mutants mutated in genes that presumptively encode biosynthetic and regulatory proteins required for biosynthesis of Hep has consistently shown damaged Hgl layers, leading to the interpretation that the Hep layer protects the structure of the Hgl layer (Wolk, 2000; Zhou and Wolk, 2003; Fan et al., 2006). Because limitation of penetration of 0, implies limitation of penetration of the structurally similar molecule N,, there has to be a compromise on blockage of penetration. The evidence seems fairly good that
ANALOGOUS SYSTEMS how much glycolipid is synthesized depends, if only indirectly, on how great is the extracellular concentration of 0, (Kangatharalingam et al., 1992; Rippka and Stanier, 1978), but the presumably 0,-responsive mechanism of regulation of the Hgl layer remains unknown. As has been abundantly shown by numerous authors and recapitulated and extended by the results of Ehira and Ohmori (2006a), major landmarks in heterocyst differentiation are regulated transcriptionally. Therefore, analysis of transcriptional regulatory pathways is essential for understanding the progression of differentiation. Binding sites for NtcA (see above) are found in the promoter region of a number of the genes that are (i) activated in differentiating cells; (ii) active in both vegetative cells and heterocysts but transcribed differentially from different transcriptional start sites depending on whether combined nitrogen, or only N,, is available; or (iii) inactivated in differentiating cells. A DNAbinding protein, DevH, that closely resembles NtcA was shown also to be required for Hgl, but not Hep, formation (Ramirez et al., 2005). Gel retardation experiments with sequences upstream from a Hep gene, purification of the DNA-binding proteins, and comparison of the sequences of tryptic fragments from these proteins with proteomic predictions identified potential DNA-binding proteins, two of which (A111939 and Alr3608) were shown by insertional mutagenesis of the corresponding genes to be required for Hgl formation but not Hep formation (Koksharova and Wolk, 2002b). (Because these proteins bound upstream from a Hep gene and resemble, respectively, a processing protease and an S-layer endoglucanase, these results were unexpected.) Two-component regulatory systems often control bacterial responses to environmental stimuli (Hoch and Silhavy, 1995).Such systems consist of a self-phosphorylating protein-histidine kinase that acts as a sensor and a response-regulator protein, to which its phosphate is transferred. The latter protein acts as a metabolic effector, often one that site-specifically binds DNA. One or more phosphate acceptor-donor proteins may intercede between the sensor and the regulator (e.g., Burbulys et al., 1991).HepK (A114496)and DevR, (Alr0442), the Anabaena sp. ortholog of N. punctiforme protein DevR (Campbell et al., 1996), interact as a His kinase and a response regulator, respectively, of an Anabaena sp. twocomponent regulatory system (Zhou and Wolk, 2003). hepK and devR, mutants share a Hep- Hgl' phenotype. Ning and Xu (2004) reported, and we have confirmed, that an alrOll7 (aka hepN [Table 11)mutant lacks the polysaccharide layer of the heterocyst envelope and, consistent with their observation that alr5351 (hglE,, one of two Anabaena sp. homologs of N. punctiforme gene
23. DEVELOPMENTAL BIOLOGYOF HETEROCYSTS, 2006
409
Table 1 Genes whose products regulate heterocyst formation and maturation ORF
Annotation
Protein
Mutant phenotype
ah0117
His kinase
HepN
Hep- Hgl+
a110187
Transcriptional regulator
ConR
Hep+ Hgl+
alr0442
Response regulator
DevR,
Hep- Hgl'
a110521
Response regulator
PatA
alrl086
Response regulator
HenR
Heterocysts terminal in filaments Hep- Hgl-
all1939
DNA-binding protein
Abp2
Hep+ Hgl-
as12301
Oligopeptide
Pats
Mch
ah2339
DNA-binding protein, protease
HetR
Het-
a112512
Ferredoxin-like transcriptional regulator Ser/Thr kinase
PatB
HepS
Heterocysts sometimes paired Hep- Hgl+
ATP-binding cassette transporter
HetC
Pattern' Het-
HetP HetF,
Pattern+'- HetHet-
a112760 ah281 7
alr2818 ah3546
ah3608
DNA-binding protein
Abp3
Hep+ Hgl-
alr3952
Transcriptional regulator
DevH
Hep' Hgl-
a114312
Transcriptional regulator
NrrA
alr4392
Nitrogen-regulatory transcriptional regulator His kinase
NtcA
Het-
HepK
Hep- Hgl'
HetN
Different mutant phenotypes Mch
a114496
ah5348 alr5358
Possible transcriptional regulator Ketoacyl reductase
Com m ent
Heterocyst envelopes often open at one or both ends Interacts with HepK Down-regulates Pats and HetN Heterocysts have only fibrous enveloping layer Similar to processing protease Involved in initial spacing of heterocysts Central regulator of heterocyst formation
Required for cessation of division during differentiation
Similar to S-layer endoglucanase
Mediates effect of NtcA on hetR
Reference(s) Ning and Xu, 2004; Fan et al., 2006; Lechno-Yossef et al., 2006 Fan et al., 2006
Campbell et al., 1996; Zhou and Wolk, 2003 Liang et al., 1992; Orozco et al., 2006 Fan et al., 2006; Lechno-Yossef et al., 2006 Koksharova and Wok, 2002b Borthakur et al., 2005; Orozco et al., 2006; Wu et al., 2004; Yoon and Golden, 1998 Buikema and Haselkorn, 1991; Huang et al., 2004; Zhou et al., 19981, Jones et al., 2003; Liang et al., 1993 Fan et al., 2006; Lechno-Yossef et al., 2006 Khudyakov and Wolk, 1997; Wang and Xu, 2005; Xu and Wolk, 2001 Fernindez-Pifias et al., 1994 Curtis and Hebbar, 2001; Wong and Meeks, 2001; Wolk et al., 2007 Koksharova and Wolk, 2002b Hebbar and Curtis, 2000; Ramirez et al., 2005 Ehira and Ohmori, 2006a, 2006b; Muro-Pastor et al., 2006 Frias et al., 1994; Herrero et al., 2004; Wei et al., 1994
Interacts with DevR,
Zhou and Wolk, 2003; Zhu et al., 1998; Lechno-Yossef et al., 2006 Fan et al., unpublished
Delayed regulation of spacing of heterocysts
Bauer et al., 1997; Black and Wolk, 1994; Borthakur et al., 2005; Callahan and Buikema, 2001; Orozco et al., 2006
420 hglE) is transcribed, we find that a glycolipid envelope layer is present in heterocysts of a hepN mutant (Fan et al., 2006). In addition, we find that transposon mutations in the genes that encode the presumptive Ser/Thr kinase A112760 (designated HepS), like mutations in hepK, devR,, and ah01 17, result in a Hep- Hgl+ phenotype, as does also one kind of alrS348 mutant (Fan et al., 2005; Fan et al. and Ehira et al., unpublished observations). Whether all of these genes are part of the same regulatory cascade and possibly act via the SpoOJlike domain (which is, however, only 73.3% aligned!) of Alr5348 is unknown. Other bacteria that differentiate express sets of genes in response to an ordered appearance of sigma factors (Apelian and Inouye, 1990; Brun and Shapiro, 1992; Chater, 1989; Losick and Stragier, 1992). SigA, the major sigma factor of Anabaena (Brahamsha and Haselkorn, 1991), is supplemented during heterocyst differentiation by SigB and SigC, but mutants in which sigB, sigC, or both are disrupted differentiate and grow normally on N, (Brahamsha and Haselkorn, 1992). sigD and sigE mutants also show very little phenotype (Khudyakov and Golden, 2001). Even so, one need not conclude that Anabaena differentiation lacks such a regulatory mechanism: its genome is annotated (http://www.kazusa.or.jp/cyano/ Anabaena/kwd.html) as having four other sigma factors, three anti-sigma factors, ca. 12 anti-sigma factor antagonists, and other sigma-factor interactive proteins. ORF ah1086 (designated he&) presumptively encodes a response regulator that, like PatA and DevR,, lacks a known DNA-binding motif. This ORF has a mutant phenotype in which heterocysts are Hep- Hglbut have a fibrous enveloping layer (Fan et al., 2006). The product of Ah1086 also has a protein phosphatase 2C (PP2C) domain that is characteristic of one of the four major classes of mammalian Ser/Thr-specific protein phosphatases. The 3’ terminus of alr2086 overlaps that of ~111087,whose product is similar to RsbV, an anti-sigma factor antagonist from B. subtilis. Alr1086 most closely resembles a B. subtilis response regulator, RsbU, which interacts with RsbV. Yet unknown is whether Alr1086 functions via A111087; via known mechanisms (see above) that regulate, independently, production of Hep or Hgl; or independently of all of them. However, because of the similarities mentioned, analysis of the mechanism of action of Alr1086 may implicate an anti-sigma factor antagonist, and thereby, perhaps, one or more sigma factors, in the regulation of heterocyst maturation. Cell wall metabolism evidently plays an important role in heterocyst formation, as was first shown by the observation that genes rf6P and rf6Z participate in the normal formation of vegetative cell lipopolysaccharide, and yet are Fox genes, required only for aerobic
ANALOGOUS SYSTEMS growth on N, (Xu et al., 1997). Similarly, alr5101 is a Fox gene whose sequence predicts a penicillin-binding protein and whose mutation affects vegetative cell walls (Lhzaro et al., 2001; LeganCs et al., 2005). Mutations in conR (a110287 [Fan et al., 2006]), which presumptively encodes a transcriptional regulatory protein, and a112981 (Leganis et al., 2005) both affect the topology of the heterocyst envelope, whose polysaccharide layer often fails to be normally constricted at one or both ends. It is unknown whether A110187 may act via A112981. In contrast to a112981 and alrS201, ah4579 predicts a penicillin-binding protein but lacks, specifically, the polysaccharide layer of the heterocyst envelope (Leganis et al., 2005). Finally, hcwA is a Fox gene that encodes N-acetylmuramoyl-L-alanine amidase (Zhu et al., 2001). A possible interpretation of the effects of the ah4579 and hcwA mutations is that precursors of the Hep layer cannot penetrate the peptidoglycan layer of the vegetative cell wall. These findings suggest, collectively, that the wall of the differentiating vegetative cell may have to be refashioned if the heterocyst envelope is to be deposited correctly (Legants et al., 2005).
OTHER DIFFERENTIATION PROCESSES IN CYANOBACTERIA Much effort has been devoted to the study of heterocyst differentiation, due in part to the role of heterocysts in N, fixation and in part to fascination with the problem of how cells “count to 10” (Haselkorn, 1998).However, there remains another equally intriguing set of problems concerning a second differentiation process in many of the same cyanobacteria. That is, in most cyanobacteria in which cells differentiate into heterocysts, other cells differentiate into akinetes, a spore form (Adams and Duggan, 1999). Akinetes can be about the same size as heterocysts, or much larger. They are also of ecological importance, because they serve as a means for the organism forming them to survive cold, desiccation, or phosphate deprivation. There are two kinds of juxtaposition patterns that involve akinetes. In one such pattern, seen in Cylindrospermum spp. and in A. cylindrica, akinetes form specifically adjacent to heterocysts. The small amount of relevant experimental evidence indicates strongly that in those instances, heterocysts in some way induce adjacent cells to become akinetes (Hirosawa and Wolk, 1980; Wolk, 1965, 1966). A second such pattern very likely involves only akinetes and is seen in many strains of Nostoc and of other genera in which akinetes form first at a distance from heterocysts. Often, there is then progressive, serial differentiation of vegetative cells into akinetes centrifugally away from the initial position of
23. DEVELOPMENTAL BIOLOGYOF HETEROCYSTS, 2006 differentiation, as though akinetes themselves induce adjacent vegetative cells to become akinetes. A reader may wonder, when “akinetes form first at a distance from heterocysts,” how does that differ from the position in which a heterocyst would be expected to differentiate? The straightforward answer is, it does not appear to differ. A great, classical cyanobacteriologist (though he would never have called himself that), Lothar Geitler, wrote (1925, p. 199; we translate), “Metabolic processes taking place in filaments call forth, at particular loci, changes that are equally favorable for the formation of spores and heterocysts, and the decision as to which is to be formed comes late, or at least later.” In short, one may pose an additional, interesting question: What is the relationship, if any, between heterocyst differentiation and akinete differentiation? One of us suggested that heterocysts may have evolved from akinetes (Wolk et al., 1994). This suggestion was based, in part, on the following observations: (i) in two species of Anabaena, the polysaccharide that forms a principal constituent of the envelope of heterocysts is experimentally identical to a principal constituent of the envelope of akinetes (Cardemil and Wolk, 1979, 1981); (ii) a hepA mutation that prevents heterocyst envelope polysaccharide from forming also prevents akinete envelope polysaccharide from forming (Legants, 1994); and (iii) in one species of Nostoc, hetR is required for the formation of akinetes as well as heterocysts and is strongly expressed in akinetes (Legants et al., 1994). Thus persuaded, you may be ready to take up your Eppendorf pipette and initiate study of akinetes, but there is a problem: with which strain should you work? Just as people present pictures of Chondromyces but work on M ~ X O C O C CA. U Scylindrica , looks wonderful but A. variabilis and N. punctiforme are much more easily manipulated experimentally. It is sad to write, but there is currently no genetically proficient strain in which to study the juxtaposition of heterocysts and akinetes. A. variabilis and N. punctiforme, both of which have fully sequenced genomes and a substantial literature, are contenders for studying the other questions. An akinete marker gene identified for A. variabilis (Zhou and Wolk, 2002) is expressed also in vegetative cells, albeit very weakly; it serves as an akinete marker also for N. punctiforme (Argueta et al., 2004; Argueta and Summers, 2005). Two other types of differentiation that are amenable to investigation are the formation of hormogonia, heterocyst-free filaments that serve for dispersal and for the initiation of symbiosis (Meeks and Elhai, 2002; Tandeau de Marsac, 1994), and the phenomenon of true branching (Gugger and Hoffmann, 2004). Concerning the latter phenomenon, we have found no study of
411
whether there is regulation of the frequency with which cells of a true-branching filamentous cyanobacterium (order Stigonematales in the older literature; Subsection V of the taxonomic system of the Pasteur Culture Collection) divide in a direction perpendicular to their normal axis of division, or whether that frequency is subject to environmental manipulation.
APPLIED BIOLOGY OF DIFFERENTIATING CYANOBACTERIA It is possible to calculate very roughly that one-quarter to one-half billion humans-subsistence rice farmers and their family members who are too poor to afford the purchase of commercial fertilizer and who therefore continue to use traditional, low-yielding varieties of ricederive a significant proportion of their nutrition from the N, fixed, largely contributed by heterocyst-forming cyanobacteria, in their rice paddies. Some cyanobacteria, including heterocyst-forming strains, are eaten; some, engineered with endotoxin genes, have been used for biocontrol of mosquitoes; others have been used as a source of recombinant pigments (Koksharova and Wolk, 2002a). However, the potential of using heterocysts to protect hydrogen-producing enzymes as a significant source of hydrogen gas (H,) in an energy economy based on H, awaits development (Tamagnini et al., 2002). Tsygankov et al. (2002) determined that in London, United Kingdom, A. variabilis produced 0.25 liter of H, (liter photobioreactor)-’day-l with a photobioreactor whose area was <0.25 m2, while simultaneously converting 7- to 21-fold more solar energy to biomass. That rate of H, production was ca. 40-fold too low to be commercially practical. The authors cited other work in which a different cyanobacterium, Arthrospira platensis, was found to convert about eightfold more solar energy to biomass in other kinds of outdoor photobioreactors. In short, genetic engineering of A. variabilis to convert a far higher fraction of solar energy to H, while making much less biomass, together with use of different photobioreactors, may permit far higher conversion of solar energy to H,. Cyanobacteria, once a dominant life form on earth and still the most numerous photoautotrophs in the ocean, might once again come into their own, as a servant of humankind, transducing solar radiation and water into energy currency.
SUMMARY The last 6 years have seen dramatic increases in our understanding of mechanisms that underlie formation of patterns of spaced heterocysts. The critically important central regulator of heterocyst formation, HetR, is
ANALOGOUS SYSTEMS
412 now recognized to be a self-degrading protease that is also, when dimerized, a transcription factor. Two principal inhibitors of its expression, namely Pats and HetN, have been identified; when neither can be synthesized, essentially all cells differentiate (Fig. 2). HetR not only
is autoregulatory but also stimulates transcription of PatS. Pats or a fragment of it and a presumptive product of HetN may mediate intercellular inhibition of differentiation at early and late stages of differentiation, respectively.
D
0
24
48
72
96 120 144 168 192 Time (h)
Figure 2 In the absence of fixed nitrogen, Pats and HetN are principally responsible for preventing extensive differentiation. Because hetN mutants are highly susceptible to suppressor mutations, a pats mutant of Anabaena sp. was modified to express hetN from a promoter, PpetE,that requires provision of Cu2+for expression, forming strain UHM100. (A) As of 2 days with neither fixed nitrogen nor Cu2+,UHMlOO shows many instances of multiple contiguous heterocysts (the Mch phenotype). (B) As of 8 days, long series of heterocysts can be seen. (C) On agar after 30 days, heterocyst formation is nearly confluent. (D) Time course of heterocyst accumulation, as a percentage of total cells, after transfer to medium lacking both fixed nitrogen and Cu2+of wild-type Anabaena sp. (+), a pats deletion mutant (A), a P,,,,-hetN mutant (*),and UHMlOO (H). Reproduced from Fig. 4 of Borthakur et al., 2005, with the kind permission of Blackwell Scientific Publications and the corresponding author.
23. DEVELOPMENTAL BIOLOGYOF HETEROCYSTS, 2006 HetR and NtcA, a classical transcription factor that is involved in diverse aspects of nitrogen metabolism, are mutually reinforcing. In addition, NtcA plays a role in transcription of numerous genes that are required for heterocyst maturation. The observation that HetR dimer binds upstream from hepA suggests that like NtcA, HetR may play a role in the progression of differentiation as well as in its inception. Use of green fluorescent protein has allowed a number of laboratories to image the loci of expression of numerous genes, while 2,2-difluoropentanedioic acid, recently introduced, provides a direct trigger of differentiation. Significant progress is also being made in analysis of mechanisms that lead to heterocyst maturation. Numerous genes, including regulatory genes, have been identified that are required specifically for maturation, and microarray experiments have identified many “unknown” and “hypothetical” genes that are activated strongly at different times after nitrogen deprivation as well as clusters of genes for which considerable functional information has been obtained. However, no network of interacting proteins has yet been identified that may guide the way to heterocyst maturation. We thank Anneliese Ernst for championing the theoretical work of Turing and Meinhardt. Our work is supported by a grant (30230040) from the National Science Foundation of China (J.Z.) and by U.S. Department of Energy grant DOE-
FG02-91ER20021 (C.P.W).
References Adams, D. G., and P. S. Duggan. 1999. Heterocyst and akinete differentiation in cyanobacteria. New Phytol. 144:3-33. Akinyanju, J., and R. J. Smith. 1979. Accumulation of ppGpp and pppGpp during nitrogen deprivation of the cyanophyte Anabaena cylindrica. FEBS Lett. 107:173-176. Akinyanju, J. A., and R. J. Smith. 1987. The accumulation of phosphorylated guanosine nucleotides in Anabaena cylindrica. New Phytol. 105:117-122. Apelian, D., and S. Inouye. 1990. Development-specific (T factor essential for late stage differentiation of Myxococcus xanthus. Genes Dev. 41396-1403. Argueta, C., K. Yuksek., and M. Summers. 2004. Construction and use of GFP reporter vectors for analysis of celltype-specific gene expression in Nostoc punctiforme. 1.Microbiol. Methods 59:181-188. Argueta, C., and M. L. Summers. 2005. Characterization of a model system for the study of Nostoc punctiforme akinetes. Arch. Microbiol. 183:338-346. Awai, K., and C. P. Wolk. 2006. Identification of the glycosyl transferase required for synthesis of the principal glycolipid characteristic of heterocysts of Anabaena sp. strain PCC 7120. FEMS Microbiol. Lett. 266:98-102. Bauer, C. C., K. S. Ramaswamy, S. Endley, L. A. Scappino, J. W. Golden, and R. Haselkorn. 1997. Suppression of heterocyst
413
differentiation in Anabaena PCC 7120 by a cosmid carrying wild-type genes encoding enzymes for fatty acid synthesis. FEMS Microbiol. Lett. 151:23-30. Becker, D. W., and J. J. Brand. 1985. Anacystis nidulans demonstrates a photosystem I1 cation requirement satisfied only by Ca2+and Na+. Plant Physiol. 79552-558. Berkelman, T., P. Garret-Engele, and N. E. Hoffman. 1994. The pacL gene of Synechococcus sp. strain PCC 7942 encodes a Ca2+-transportingATPase.]. Bacteriol. 176:44304436. Bianchini, G. M., A. C. Pastini, J. P. Muschietti, M. T. TtllezIiion, H. E. Martinetto, H. N. Torres, and M. M. Flawia. 1990. Adenylate cyclase activity in cyanobacteria: activation by Ca2+-calmodulinand a calmodulin-like activity. Biochim. Biophys. Acta 1055:75-81. Black, T. A., and C. P. Wolk. 1994. Analysis of a Het- mutation in Anabaena sp. strain PCC 7120 implicates a secondary metabolite in the regulation of heterocyst spacing.]. Bacteriol. 176:2282-2292. Black, T. A., Y. Cai, and C. P. Wolk. 1993. Spatial expression and autoregulation of hetR, a gene involved in the control of heterocyst development in Anabaena. Mol. Microbiol. 9:77-84. Borthakur, P. B., C. C. Orozco, S. S. Young-Robbins, R. Haselkorn, and S. M. Callahan. 2005. Inactivation of pats and hetN causes lethal levels of heterocyst differentiation in the filamentous cyanobacterium Anabaena sp. PCC 7120. Mol. Microbiol. 57:111-123. Brahamsha, B., and R. Haselkorn. 1991. Isolation and characterization of the gene encoding the principal sigma factor of the vegetative cell RNA polymerase from the cyanobacterium Anabaena sp. strain PCC 7120.1. Bacteriol. 173:24422450. Brahamsha, B., and R. Haselkorn. 1992. Identification of multiple RNA polymerase sigma factor homologs in the cyanobacterium Anabaena sp. strain PCC 7120: cloning, expression, and inactivation of the sigB and sigC genes. J. Bacteriol. 174:7273-7282. Brand, J. J., and D. W. Becker. 1988. In vivo effect of calcium on photosystem 11. Methods Enzymol. 167:280-285. Bretz, J., L. Losada, K. Lisboa, and S. W. Hutcheson. 2002. Lon protease functions as a negative regulator of type I11 protein secretion in Pseudomonas syringae. Mol. Microbiol. 45:397-409. Brun, Y., and L. Shapiro. 1992. A temporally controlled afactor is required for polar morphogenesis and normal cell division in Caulobacter. Genes Dev. 6:2395-2408. Buikema, W. J., and R. Haselkorn. 1991. Characterization of a gene controlling heterocyst differentiation in the cyanobacterium Anabaena 7120. Genes Dev. 5:321-330. Buikema, W. J., and R. Haselkorn. 1993. Molecular genetics of cyanobacterial development. Annu. Rev. Plant Physiol. Plant Mol. Biol. 4433-52. Buikema, W. J., and R. Haselkorn. 2001. Expression of the Anabaena hetR gene from a copper-regulated promoter leads to heterocyst differentiation under repressing conditions. Proc. Natl. Acad. Sci. USA 98:2729-2734. Burbulys, D., K. A. Trach, and J. A. Hoch. 1991. Initiation of sporulation in B. subtilis is controlled by a multicomponent phosphorelay. Cell 64545-552.
414 Cai, Y., and C. P. Wolk. 1997. Nitrogen deprivation of Anabaena sp. strain PCC 7120 elicits rapid activation of a gene cluster that is essential for uptake and utilization of nitrate. J. Bacteriol. 179:258-266. Callahan, S. M., and W. J. Buikema. 2001. The role of HetN in maintenance of the heterocyst pattern in Anabaena sp. PCC 7120. Mol. Microbiol. 40:941-950. Campbell, A. K. 1983. Intracellular Calcium: Its Universal Role as Regulator. Wiley, New York, NY. Campbell, E. L., K. D. Hagen, M. F. Cohen, M. L. Summers, and J. C. Meeks. 1996. The devR gene product is characteristic of receivers of two-component regulatory systems and is essential for heterocyst development in the filamentous cyanobacterium Nostoc sp. strain ATCC 29133. J. Bacteriol. 178:2037-2043. Cardemil, L., and C. P. Wolk. 1979. The polysaccharides from heterocyst and spore envelopes of a blue-green alga. Structure of the basic repeating unit.]. Biol. Chem. 254:736-741. Cardemil, L., and C. P. Wolk. 1981. Polysaccharides from the envelopes of heterocysts and spores of the blue-green algae Anabaena variabilis and Cylindrospermum licheniforme. J. Phycol. 17~234-240. Chater, K. F. 1989. Sporulation in Streptomyces, p. 277-299. In I. Smith, R. A. Slepecky, and P. Setlow (ed.),Regulation of Procaryotic Development. American Society for Microbiology, Washington, DC. Curtis, S. E., and P. B. Hebbar. 2001. A screen for sequences up-regulated during heterocyst development in Anabaena sp. strain PCC 7120. Arch. Microbiol. 175:313-322. D’Argenio, D. A., and S. I. Miller. 2004. Cyclic di-GMP as a bacterial second messenger. Microbiology 150:2497-2502. Dominguez, D. C. 2004. Calcium signalling in bacteria. Mol. Microbiol. 54:291-297. Dong, U., X. Huang, X.-Y. Wu, and J. Zhao. 2000. Identification of the active site of HetR protease and its requirement for heterocyst differentiation in the cyanobacterium Anabaena sp. strain PCC 7120. J. Bacteriol. 182:1575-1579. Ehira, S., M. Ohmori, and N. Sato. 2003. Genome-wide expression analysis of the responses to nitrogen deprivation in the heterocyst-forming cyanobacterium Anabaena sp. strain PCC 7120. D N A Res. 10:97-113. Ehira, S., and M. Ohmori. 2006a. NrrA, a novel nitrogenregulating response regulator regulates heterocyst development in the cyanobacterium Anabaena sp. strain PCC 7120. Mol. Microbiol. 59:1692-1703. Ehira, S., and M. Ohmori. 2006b. NrrA directly regulates expression of hetR during heterocyst differentiation in the cyanobacterium Anabaena sp. strain PCC 7120.1. Bacteriol. 188:8520-8525. Elhai, J., and C. P. Wolk. 1990. Developmental regulation and spatial pattern of expression of the structural genes for nitrogenase in the cyanobacterium Anabaena. EMBO ]. 9:33793388. Fan, Q., G. Huang, S. Lechno-Yossef, C. P. Wolk, T. Kaneko, and S. Tabata. 2005. Clustered genes required for synthesis and deposition of envelope glycolipids in Anabaena sp. strain PCC 7120. Mol. Microbiol. 58:227-243. Fan, Q., S. Lechno-Yossef, S. Ehira, T. Kaneko, M. Ohmori, N. Sato, S. Tabata, and C. P. Wolk. 2006. Signal transduction
ANALOGOUS SYSTEMS genes required for heterocyst maturation in Anabaena sp. strain PCC 7120.1. Bacteriol. 188:6688-6693. Fay, P., W. D. P. Stewart, A. E. Walsby, and G. E. Fogg. 1968. Is the heterocyst the site of nitrogen fixation in blue-green algae? Nature 220:810-812. Fernandez-Piiias, F. Legants, and C. P. Wolk. 1994. A third genetic locus required for the formation of heterocysts in Anabaena sp. strain PCC 7120. J. Bacteriol. 1765277-5283. Fiedler, G., M. Arnold, S. Hannus, and I. Maldener. 1998. The DevBCA exporter is essential for envelope formation in heterocysts of the cyanobacterium Anabaena sp. strain PCC 7120. Mol. Microbiol. 271193-1202. Fleming, H., and R. Haselkorn. 1974. The program of protein synthesis during heterocyst differentiation in nitrogen-fixing blue-green algae. Cell 3:159-170. Forchhammer, K. 2004. Global carbodnitrogen control by PI, signal transduction in cyanobacteria: from signals to targets. FEMS Microbiol. Rev. 28:319-333. Forchhammer, K., and N. Tandeau de Marsac. 1994. The PI, protein in the cyanobacterium Synechococcus sp. strain PCC 7942 is modified by serine phosphorylation and signals the cellular N-status. J. Bacteriol. 176:84-91. Frias, J. E., E. Flores, and A. Herrero. 1994. Requirement of the regulatory protein NtcA for the expression of nitrogen assimilation and heterocyst development genes in the cyanobacterium Anabaena sp. PCC 7120. Mol. Microbiol. 145323432. Fritsch, F. E. 1951. The heterocyst: a botanical enigma. Proc. Linn. SOC. London 162:194-211. Geisler, M., J. Richter, and J. Schumann. 1993. Molecular cloning of a P-type ATPase gene from the cyanobacteriuin Synechocystis sp. PCC 6803. Homology to eukaryotic Ca2+ATPases. J . Mol. Biol. 234:1284-1289. Geitler, L. 1925. Synoptische Darstellung der Cyanophyceen in morphologischer und systematischer Hinsicht. Beih. Botan. Centralbl. 41:163-294. Gierer, A., and H. Meinhardt. 1972. A theory of biological pattern formation. Kybernetik 12:30-39. Golden, J. 1998. Programmed DNA rearrangements in cyanobacteria, p. 162-173. In F. J. de Bruijn, J. R. Lupski, and G. M. Weinstock (ed.), Bacterial Genomes: Physical Structure and Analysis. Chapman & Hall, New York, NY. Golden, J. W., and H. S. Yoon. 2003. Heterocyst development in Anabaena. Curr. Opin. Microbiol. 6557-563. Gugger, M. F., and L. Hoffmann. 2004. Polyphyly of true branching cyanobacteria (Stigonematales). Int. J. Syst. Evol. Microbiol. 54:349-357. Haselkorn, R. 1998. How cyanobacteria count to 10. Science 282~891-892. Hebbar, P. B., and S. E. Curtis. 2000. Characterization of devH, a gene encoding a putative DNA binding protein required for heterocyst function in Anabaena sp. strain PCC 7120. J. Bacteriol. 182:3572-358 1. Herrero, A., A. M. Muro-Pastor, and E. Flores. 2001. Nitrogen control in cyanobacteria. J. Bacteriol. 183:411-425. Herrero, A., A. M. Muro-Pastor, A. Valladares, and E. Flores. 2004. Cellular differentiation and the NtcA transcription factor in filamentous cyanobacteria. FEMS Microbiol. Rev. 28:469-487.
23. DEVELOPMENTAL BIOLOGYOF HETEROCYSTS, 2006 Hirosawa, T., and C. P. Wolk. 1980. Factors controlling the formation of akinetes adjacent to heterocysts in the cyanobacterium Cylindrospermum licheniforme TZiitz. J. Gen. Microbiol. 114:423-432. Hoch, J. A., and T. J. Silhavy (ed.). 1995. Two-Component Signal Transduction. ASM Press, Washington, DC. Huang, G., Q. Fan, S. Lechno-Yossef, E. Wojciuch, C. P. Wolk, T. Kaneko, and S. Tabata. 2005. Clustered genes required for the synthesis of heterocyst envelope polysaccharide in Anabaena sp. strain PCC 7120. J. Bacteriol. 187:1114-1123. Huang, X., Y. Dong, and J. Zhao. 2004. HetR homodimer is a DNA-binding protein required for heterocyst differentiation, and the DNA-binding activity is inhibited by Pats. Proc. Natl. Acad. Sci. USA 101:4848-4853. Imashimizu, M., H. Yoshimura, H. Katoh, S. Ehira, and M. Ohmori. 2005. NaCl enhances cellular CAMP and upregulates genes related to heterocyst development in the cyanobacterium, Anabaena sp. strain PCC 7120. FEMS Microbiol. Lett. 252:97-103. Irmier, A., S. Sanner, H. Dierks, and K. Forchhammer. 1997. Dephosphorylation of the phosphoprotein PI, in Synechococcus PCC 7942: identification of an ATP and 2-oxoglutarate phosphatase activity. Mol. Microbiol. 26:81-90. Janson, S., A. Matveyev, and B. Bergman. 1998. The presence and expression of hetR in the non-heterocystous cyanobacteriuum Symploca PCC 8002. FEMS Microbiol. Lett. 168~173-179. Jones, K. M., and R. Haselkorn. 2002. Newly identified cytochrome c oxidase operon in the nitrogen-fixing cyanobacterium Anabaena sp. strain PCC 7120 specifically induced in heterocysts. J. Bacteriol. 184:2491-2499. Jones, K. M., W. J. Buikema, and R. Haselkorn. 2003. Heterocyst-specific expression of patB, a gene required for nitrogen fixation in Anabaena sp. strain PCC 7120. J . Bacteriol. 185:2306-2314. Kaneko, T., Y. Nakamura, C. P. Wolk, T. Kuritz, S. Sasamoto, A. Watanabe, M. Iriguchi, A. Ishikawa, K. Kawashima, T. Kimura, Y. Kishida, M. Kohara, M. Matsumoto, A. Matsuno, A. Muraki, N. Nakazaki, s. Shimpo, M. Sugimoto, M. Takazawa, M. Yamada, M. Yasuda, and S. Tabata. 2001. Complete genomic sequence of the filamentous nitrogenfixing cyanobacterium Anabaena sp. strain PCC 7120. D N A Res. 8:205-213. Kangatharalingam, N., J. C. Priscu, and H. W. Paerl. 1992. Heterocyst envelope thickness, heterocyst frequency and nitrogenase activity in Anabaena flos-aquae: influence of exogenous oxygen tension. J. Gen. Microbiol. 138:26732678. Khudyakov, I., and C. P. Wolk. 1997. hetC, a gene coding for a protein similar to bacterial ABC protein exporters, is involved in early regulation of heterocyst differentiation in Anabaena sp. strain PCC 7120. J. Bacteriol. 179:6971-6978. Khudyakov, I. Y., and J. W. Golden. 2001. Identification and inactivation of three group 2 sigma factor genes in Anabaena sp. strain PCC 7120. J. Bacteriol. 183:6667-6675. Khudyakov, I. Y., and J. W. Golden. 2004. Different functions of HetR, a master regulator of heterocyst differentiation in Anabaena sp. PCC 7120, can be separated by mutation. Proc. Natl. Acad. Sci. USA 101:16040-16045.
415
Koksharova, 0. A., and C. P. Wolk. 2002a. Genetic tools for cyanobacteria. Appl. Microbiol. Biotechnol. 58:123-137. Koksharova, 0.A., and C. P. Wolk. 2002b. Novel DNA-binding proteins in the cyanobacterium Anabaena sp. strain PCC 7120. J. Bacteriol. 184:3931-3940. Kuhn, I., L. Peng, S. Bedu, and C.-C. Zhang. 2000. Developmental regulation of the cell division protein FtsZ in Anabaena sp. strain PCC 7120, a cyanobacterium capable of terminal differentiation. J. Bacteriol. 182:4640-4643. Lammers, P. J., and R. Haselkorn. 1983. Sequence of the n i p gene coding for the a subunit of dinitrogenase from the cyanobacterium Anabaena. Proc. Natl. Acad. Sci. USA 80~4723-4727. Laurent, S., K. Forchhammer, L. Gonzalez, T. Heulin, C.-C. Zhang, and S. BCdu. 2004. Cell-type specific modification of PI, is involved in the regulation of nitrogen metabolism in the cyanobacterium Anabaena PCC 7120. FEBS Lett. 576:261-265. Laurent, S., H. Chen, S. Bidu, F. Ziarelli, L. Peng, and C.-C. Zhang. 2005. Nonmetabolizable analogue of 2-oxoglutarate elicits heterocyst differentiation under repressive conditions in Anabaena sp. PCC 7120. Proc. Natl. Acad. Sci. USA 102:9907-9912. Lazaro, S., F. Fernandez-Piiias, E. Fernandez-Valiente, A. BlancoRivero, and F. LeganCs. 2001. pbpB, a gene coding for a putative penicillin-binding protein, is required for aerobic nitrogen fixation in the cyanobacterium Anabaena sp. strain PCC7120. J. Bacteriol. 183:628-636. Lechno-Yossef, S., Q. Fan, S. Ehira, N. Sato, and C. P. Wolk. 2006. Mutations in four regulatory genes have interrelated effects on heterocyst maturation in Anabaena sp. strain PCC 7120. J. Bacteriol. 188:7387-7395. Legants, F. 1994. Genetic evidence that hepA gene is involved in the normal deposition of the envelope of both heterocysts and akinetes in Anabaena variabilis ATCC 29413. FEMS Microbiol. Lett. 123:63-67. Leganis, F., F. Fernandez-Piiias, and C. P. Wolk. 1994. Two mutations that block heterocyst differentiation have different effects on akinete differentiation in Nostoc ellipsosporum. Mol. Microbiol. 12:679-684. Legants, F., A. Blanco-Rivero, F. Fernhndez-Picas, M. Redondo, E. Fernandez-Valiente, Q. Fan, S. Lechno-Yossef, and C. P. Wolk. 2005. Wide variation in the cyanobacterial complement of presumptive penicillin-binding proteins. Arch. Microbiol. 184:234-248. Li, B., X. Huang, and J. Zhao. 2002. Expression of hetN during heterocyst differentiation and its inhibition of hetR upregulation in the cyanobacterium Anabaena sp. PCC 7120. FEBS Lett. 517537-91. Li, J.-H., S. Laurent, V. Konde, S. BCdu, and C.-C. Zhang. 2003. An increase in the level of 2-oxoglutarate promotes heterocyst development in the cyanobacterium Anabaena sp. strain PCC 7120. Microbiology 149:3257-3263. Liang, J., L. Scappino, and R. Haselkorn. 1992. The patA gene product, which contains a region similar to CheY of Escherichia coli, controls heterocyst pattern formation in the cyanobacterium Anabaena 7120. Proc. Natl. Acad. Sci. USA 895655-5659.
416 Liang, J., L. Scappino, and R. Haselkorn. 1993. The patB gene product, required for growth of the cyanobacterium Anabaena sp. strain PCC 7120 under nitrogen-limiting conditions, contains ferredoxin and helix-turn-helix domains. J. Bacteriol. 175:1697-1704. Liotenberg, S., D. Campbell, A.-M. Castets, J. Houmard, and N. Tandeau de Marsac. 1996. Modification of the PI, protein in response to carbon and nitrogen availability in filamentous heterocystous cyanobacteria. FEMS Microbiol. Lett. 144~185-190. Losick, R., and P. Stragier. 1992. Crisscross regulation of celltype-specific gene expression during development in B. subtilis. Nature 355:601-604. Lundgren, P., S. Janson, S. Jonasson, A. Singer, and B. Bergman. 2005. Unveiling of novel radiations within Trichodesmium cluster by hetR gene sequence analysis. Appl. Environ. Microbiol. 71:190-196. Luque, I., E. Flores, and A. Herrero. 1994. Molecular mechanism for the operation of nitrogen control in cyanobacteria. EMBO J. 13~2862-2869. Maldener, I., W. Lockau, Y. Cai, and C. P. Wolk. 1991. Calcium-dependent protease of the cyanobacterium Anabaena: molecular cloning and expression of the gene in Escherichia coli, sequencing and site-directed mutagenesis. Mol. Gen. Genet. 225:113-120. Maldener, I., G . Fiedler, A. Ernst, F. Fernandez-Piiias, and C. P. Wolk. 1994. Characterization of devA, a gene required for the maturation of proheterocysts in the cyanobacterium Anabaena sp. strain PCC 7120. J. Bacteriol. 176:7543-7549. Martin-Figueroa, E., F. Navarro, and F. J. Florencio. 2000. The GS-GOGAT pathway is not operative in the heterocysts. Cloning and expression of glsF gene from the cyanobacterium Anabaena sp. PCC 7120. FEBS Lett. 476:282-286. Mazur, B. J., and C. F. Chui. 1982. Sequence of the gene coding for the P-subunit of dinitrogenase from the blue-green alga Anabaena. Proc. Natl. Acad. Sci. USA 79:6782-6786. Meeks, J. C., and J. Elhai. 2002. Regulation of cellular differentiation in filamentous cyanobacteria in free-living and plant-associated symbiotic growth states. Microbiol. Mol. Biol. Rev. 66:94-121. Meinhardt, H., and A. Gierer. 2000. Pattern formation by local self-activation and lateral inhibition. Bioessays 22:753760. Merida, A., P. Candau, and F. J. Florencio. 1991. Regulation of glutamine synthetase activity in the unicellular cyanobacterium Synechocystis sp. strain PCC 6803 by the nitrogen source: effect of ammonium. J. Bacteriol. 173: 4095-4100. Merida, A., E. Flores, and F. J. Florencio. 1992. Regulation of Anabaena sp. strain PCC 7120 glutamine synthetase activity in a Synechocystis sp. strain PCC 6803 derivative strain bearing the Anabaena glnA gene and a mutated host glnA gene. J. Bacteriol. 174:650-654. Merrick, M. J., and R. A. Edwards. 1995. Nitrogen control in bacteria. Microbiol. Rev. 59:604-622. Mes, T. H. M., and L. J. Stal. 2005. Variable selection pressures across lineages in Trichodesmium and related cyanobacteria based on the heterocyst differentiation protein gene hetR. Gene 346:163-171.
ANALOGOUS SYSTEMS Mevarech, M., D. Rice, and R. Haselkorn. 1980. Nucleotide sequence of a cyanobacterial n i p gene coding for nitrogenase reductase. Proc. Natl. Acad. Sci. USA 77:6476-6480. Michiels, J., C. Xi, J. Verhaert, and J. Venderleyden. 2002. The function of CaZ+in bacteria: a role for EF-hand proteins? Trends Microbiol. 1097-93. Mitchison, G. J., M. Wilcox, and R.J. Smith. 1976. Measurement of an inhibitory zone. Science 191:866-868. Mulligan, M. E., and R. Haselkorn. 1989. Nitrogen fixation (nif) genes of the cyanobacterium Anabaena species strain PCC 7120. The nip-fdxN-nifS-nifU operon. J. Biol. Chem. 264:19200-19207. Muro-Pastor, A. M., A. Valladares, E. Flores, and A. Herrero. 1999. The hetC gene is a direct target of the NtcA transcriptional regulator in cyanobacterial heterocyst development. J. Bacteriol. 18 1:66 64-66 6 9. Muro-Pastor, A. M., A. Valladares, E. Flores, and A. Herrero. 2002. Mutual dependence of the expression of the cell differentiation regulatory protein HetR and the global nitrogen regulator NtcA during heterocyst development. Mol. Microbiol. 44:1377-1385. Muro-Pastor, A. M., E. Olmedo-Verd, and E. Flores. 2006. A1143 12, an NtcA-regulated two-component response regulator in Anabaena sp. strain PCC 7120. FEMS Microbiol. Lett. 256:171-177. Muro-Pastor, M. I., and F. J. Florencio. 2003. Regulation of ammonium assimilation in cyanobacteria. Plant Physiol. Biochem. 41595-603. Muro-Pastor, M. I., J. C. Reyes, and F. J. Florencio. 1996. The NADP+-isocitrate dehydrogenase gene (icd)is nitrogen regulated in cyanobacteria. J. Bacteriol. 178:4070-4076. Muro-Pastor, M. I., J. C. Reyes, and F. J. Florencio. 2001. Cyanobacteria perceive nitrogen status by sensing intracellular 2-oxoglutarate levels. J. Biol. Chem. 276:38320-38328. Ning, D., and X. Xu. 2004. alrO117, a two-component histidine kinase gene, is involved in heterocyst development in Anabaena sp. PCC 7120. Microbiology 150:447-453. Ochi, K., J. C. Kandala, and E. Freese. 1981. Initiation of Bacillus subtilis sporulation by the stringent response to partial amino acid deprivation. J. Biol. Chem. 256:6866-6875. O’Hara, M. B., and J. H. Hageman. 1990. Energy and calcium ion dependence of proteolysis during sporulation of Bacillus subtilis cells. J. Bacteriol. 172:4161-4170. Olmedo-Verd, E., E. Flores, A. Herrero, and A. M. Muro-Pastor. 2005. HetR-dependent and -independent expression of heterocyst-related genes in an Anabaena strain overproducing the NtcA transcription factor. J. Bacteriol. 18719851991. Onek, L. A., and R. J. Smith. 1992. Calmodulin and calcium mediated regulation in prokaryotes. J. Gen. Microbiol. 138:1039-1049. Onek, L. A., P. J. Lea, and R. J. Smith. 1994. Isolation and characterization of a calmodulin-like protein from the cyanobacterium Nostoc sp. PCC 6720. Arch. Microbiol. 161:352-3 5 8. Orozco, C. C., D. D. Risser, and S. M. Callahan. 2006. Epistasis analysis of four genes from Anabaena sp. strain PCC 7120 suggests a connection between PatA and Pats in heterocyst pattern formation. J. Bacteriol. 188:1808-1816.
23. DEVELOPMENTAL BIOLOGYOF HETEROCYSTS, 2006 Pettersson, A., and B. Bergman. 1989. Calmodulin in heterocystous cyanobacteria: biochemical and immunological evidence. FEMS Microbiol. Lett. 60:95-100. Ramasubramanian, T. S., T. F. Wei, A. K. Oldham, and J. W. Golden. 1996. Transcription of the Anabaena sp. strain PCC 7120 ntcA gene: multiple transcripts and NtcA binding. J. Bacteriol. 178:922-926. Ramirez, M. E., P. B. Hebbar, R. Zhou, C. P. Wolk, and S. E. Curtis. 2005. Anabaena sp. strain PCC 7120 gene devH is required for synthesis of the heterocyst glycolipid layer. J. Bacteriol. 187:2326-2331. Rippka, R., and R. Y. Stanier. 1978. The effects of anaerobiosis on nitrogenase synthesis and heterocyst development by nostocacean cyanobacteria. J. Gen. Microbiol. 105533-94. Risser, D. D., and S. M. Callahan. 2007. Mutagenesis of hetR reveals amino acids necessary for HetR function in the heterocystous cyanobacterium Anabaena sp. strain PCC 7120. J. Bacteriol. 189:2460-2467. Rowell, R., and W. D. P. Stewart. 1975. Effects of L-methionineDL-SUlfOXimine on the assimilation of newly fixed NH,, acetylene reduction and heterocyst production in Anabaena cylindrica. Biochem. Biophys. Res. Commun. 65:846-856. Sakr, S., R. Jeanjean, C.-C. Zhang, and T. Arcondeguy. 2006. Inhibition of cell division suppresses heterocyst development in Anabaena sp. strain PCC 7120. J. Bacteriol. 188:13961404. Shi, Y., W. Zhao, W. Zhang., Z. Ye, and J. Zhao. 2006. Regulation of intracellular free calcium concentration during heterocyst differentiation by HetR and NtcA in Anabaena sp. PCC 7120. Proc. Natl. Acad. Sci. USA 103:11334-11339. Smith, G., and J. D. Ownby. 1981. Cyclic AMP interferes with pattern formation in the cyanobacterium Anabaena variabilis. FEMS Microbiol. Lett. 11:175-180. Smith, R. J., S. Hobson., and I. R. Ellis. 1987. Evidence for calcium-mediated regulation of heterocyst frequency and nitrogenase activity in Nostoc 6720. New Phytol. 105531-541. Su, Z., V. Olman., F. Mao, and Y. Xu. 2005. Comparative genomics analysis of NtcA regulons in cyanobacteria: regulation of nitrogen assimilation and its coupling to photosynthesis. Nucleic Acids Res. 3351.56-5171. Tamagnini, P., R. Axelsson, P. Lindberg, F. Oxelfelt, R. Wunschiers, and P. Lindblad. 2002. Hydrogenases and hydrogen metabolism of cyanobacteria. Microbiol. Mol. Biol. Rev. 66:l-20. Tandeau de Marsac, N. 1994. Differentiation of hormogonia and relationships with other biological processes, p. 825842. In D. A. Bryant (ed.), The Molecular Biology of Cyanobacteria. Kluwer Academic Publishers, Dordrecht, The Netherlands. Tanigawa, R., M. Shirokane, S. I. Maeda, T. Omata, K. Tanaka, and H. Takahashi. 2002. Transcriptional activation of NtcA-dependent promoters of Synechococcus sp. PCC 7942 by 2-oxoglutarate in vitro. Proc. Natl. Acad. Sci. USA 99:42S 1-4255. Tapia, M. I., J. A. G. Ochoa de Alda, M. J. Llama, and J. L. Serra. 1996. Changes in intracellular amino acids and organic acids induced by nitrogen starvation and nitrate or ammonium resupply in the cyanobacterium Phormidium laminosum. Planta 198526-531.
41 7
Thiel, T. 2004. Nitrogen fixation in heterocyst-forming cyanobacteria. In W. Klipp, B. Masepohl, J. R. Gallon, and W. E. Newton (ed.), Genetics and Regulation of Nitrogen Fixing Bacteria. Kluwer Academic Publishers, Dordrecht, The Netherlands. Thiel, T., and M. Leone. 1986. Effect of glutamine on growth and heterocyst differentiation in the cyanobacterium Anabaena variabilis. J. Bacteriol. 168:769-774. Thiel, T., and B. Pratte. 2001. Effect on heterocyst differentiation of nitrogen fixation in vegetative cells of the cyanobacterium Anabaena variabilis ATCC 29413. J. Bacteriol. 183:280-286. Thiel, T., E. M. Lyons, J. C. Erker, and A. Ernst. 1995. A second nitrogenase in vegetative cells of a heterocystforming cyanobacterium. Proc. Natl. Acad. Sci. USA 92: 9358-9362. Thomas, J., J. C. Meeks, C. P. Wolk, P. W. Shaffer, and S. M. Austin. 1977. Formation of glutamine from [13N]ammonia, [I3N]dinitrogen, and [14C]glutamate by heterocysts isolated from Anabaena cylindrica. J. Bacteriol. 129:1545-1555. Tisa, L.S., B. M. Olivera, and J. Adler. 1993. Inhibition of Escherichia coli chemotaxis by wconotoxin, a calcium ion channel blocker. 1.Bacteriol. 175:1235-1238. Tomitani, A., A. H. Knoll, C. M. Cavanaugh, and T. Ohno. 2006. The evolutionary diversification of cyanobacteria: molecular-phylogenetic and paleontological perspectives. Proc. Natl. Acad. Sci. USA 1035442-5447. Torrecilla, I., F. LeganCs, I. Bonilla, and F. Fernandez-Piiias. 2000. Use of recombinant aequorin to study calcium homeostasis and monitor calcium transients in response to heat and cold shock in cyanobacteria. Plant Physiol. 123:161176. Torrecilla, I., F. LeganCs, I. Bonilla, and F. Fernandez-Pifias. 2004. A calcium signal is involved in heterocyst differentiation in the cyanobacterium Anabaena sp. PCC 7120. Microbiology 150:373 1-3739. Tsygankov, A. A., A. S. Fedorov, S. N. Kosourov, and K. K. Rao. 2002. Hydrogen production by cyanobacteria in an automated outdoor photobioreactor under aerobic conditions. Biotechnol. Bioeng. 80:777-783. Turing, A.M. 1952. The chemical basis of morphogenesis. Phi10s. Trans. Roy. Soc. Lond. B 237:37-72. Valladares, A., A. Herrero, D. Pils, G. Schmetterer, and E. Flores. 2003. Cytochrome c oxidase genes required for nitrogenase activity and diazotrophic growth in Anabaena sp. PCC 7120. Mol. Microbiol. 47:1239-1249. Vega-Palas, M.A., E. Flores, and A. Herrero. 1992. NtcA, a global nitrogen regulator from the cyanobacterium Synechococcus that belongs to the Crp family of bacterial regulators. Mol. Microbiol. 6:1853-1859. Walsby, A. E. 1985. The permeability of heterocysts to the gases nitrogen and oxygen. Proc. R. SOL. Lond. B 226:345-366. Wang, Y., and X. Xu. 2005. Regulation by hetC of genes required for heterocyst differentiation and cell division in Anabaena sp. strain PCC 7120. J. Bacteriol. 187:84898493. Wang, Y., S. Lechno-Yossef,Y. Gong, Q. Fan, C. P. Wolk, and X. Xu. 2007. Predicted glycosyl transferase genes located outside the HEP Island are required for formation of heterocyst
418 envelope polysaccharide in Anabaena sp. strain PCC 7120. J. Bacteriol189:5372-5378. Wei, T.-F., T. S. Ramasubramanian, and J. W. Golden. 1994. Anabaena sp. strain PCC 7120 ntcA gene required for growth on nitrate and heterocyst development. J. Bacteriol. 176:4473-4482. Wigglesworth, V. B. 1940. Local and general factors in the development of “pattern” in Rhodnius prolixus. J. Exp. Biol. 17:lSO-200. Wilcox, M., G. J. Mitchison, and R. J. Smith. 1973. Pattern formation in the blue-green alga, Anabaena. I. Basic mechanisms. J. Cell Sci. 12:707-723. Wolk, C. P. 1965. Control of sporulation in a blue-green alga. Dev. Biol. 12:15-35. Wolk, C. P. 1966. Evidence of a role of heterocysts in the sporulation of a blue-green alga. Am.]. Bot. 53:260-262. Wolk, C. P. 1967. Physiological basis of the pattern of vegetative growth of a blue-green alga. Proc. Natl. Acad. Sci. USA 57~1246-1251. Wolk, C. P. 1968. Movement of carbon from vegetative cells to heterocysts in Anabaena cylindrica. J. Bacteriol. 96:21382143. Wolk, C. P. 2000. Heterocyst formation in Anabaena, p. 83-104. In Y. V. Brun and L. J. Shimkets (ed.), Prokaryotic Development. ASM Press, Washington, DC. Wolk, C. P., A. Ernst, and J. Elhai. 1994. Heterocyst metabolism and development, p. 769-823. In D. A. Bryant (ed.), The Molecular Biology of Cyanobacteria. Kluwer Academic Publishers, Dordrecht, The Netherlands. Wolk, C. P., Q. Fan, R. Zhou, G. Huang, S. Lechno-Yossef, T. Kuritz, and E. Wojciuch. 2007. Paired cloning vectors for complementation of mutations in the cyanobacterium Anabaena sp. strain PCC 7120. Arch. Microbiol. Jul. 17: epub ahead of print. Wong, F. C., and J. C . Meeks. 2001. The hetF gene product is essential to heterocyst differentiation and affects HetR function in the cyanobacterium Nostoc punctiforme.]. Bacteriol. 183~2654-2661. Wood,N. B., andR. Haselkorn. 1979. Proteinase activity during heterocyst differentiation in nitrogen-fixing cyanobacteria, p. 159-166. In G. H. Cohen and H. Holzer (ed.), Limited Proteolysis in Microorganisms. U.S. DHEW Publication No. (NIH) 79-1591, NIH, Bethesda, MD. Wood, N. B., and R. Haselkorn. 1980. Control of phycobiliprotein proteolysis and heterocyst differentiation in Anabaena. J. Bacteriol. 141:1375-1 3 85. Wu, X., D. Liu, M. H. Lee, and J. W. Golden. 2004. pats minigenes inhibit heterocyst development of Anabaena sp. strain PCC 7120.1. Bacteriol. 186:6422-6429.
ANALOGOUS SYSTEMS Xu, X., and C. P. Wolk. 2001. Role for hetC in the transition to a nondividing state during heterocyst differentiation in Anabaena sp.]. Bacteriol. 183:393-396. Xu, X., I. Khudyakov, and C. P. Wolk. 1997. Lipopolysaccharide dependence of cyanophage sensitivity and aerobic nitrogen fixation in Anabaena sp. strain PCC 7120. J. Bacteriol. 1792884-2891. Yoon, H.-S., and J. W. Golden. 1998. Heterocyst pattern formation controlled by a diffusible peptide. Science 282:935-938. Yoon, H.-S., and J. W. Golden. 2001. Pats and products of nitrogen fixation control heterocyst pattern. J. Bacteriol. 183:2605-2613. Youatt, J. 1993. Calcium and microorganisms. Crit. Rev. Microbiol. 19:83-97. Zhang, C.-C., S. Laurent, S. Sakr, L. Peng, and S. Bkdu. 2006. Heterocyst differentiation and pattern formation in cyanobacteria: a chorus of signals. Mol. Microbiol. 59:367-375. Zhao, J., J. W. LaClaire 11, and J. J. Brand. 1991. Calcium and heterocyst development in Anabaena 7120, p. 105. In Abstr VII. Int. S y m p . Photosynthetic Prokaryotes. Zhao, Y., Y. Shi, W. Zhao, X. Huang, D. Wang, N. Brown, J. Brand, and J. Zhao. 2005. CcbP, a calcium-binding protein from Anabaena sp. PCC 7120, provides evidence that calcium ions regulate heterocyst differentiation. Proc. Natl. Acad. Sci. USA 1025744-5748. Zhou, R., and C. P. Wolk. 2002. Identification of an akinete marker gene in Anabaena variabilis. J. Bacteriol. 184:25292532. Zhou, R., and C. P. Wolk. 2003. A two-component system mediates developmental regulation of biosynthesis of a heterocyst polysaccharide. J. Biol. Chem. 278:19939-19946. Zhou, R., Z. Cao, and J. Zhao. 1998a. Characterization of HetR protein turnover in Anabaena sp. PCC 7120. Arch. Microbiol. 169:4 1 7 4 2 3 . Zhou, R., X. Wei, N. Jiang, H. Li, Y. Dong, K. L. His, and J. Zhao. 1998b. Evidence that HetR protein is an unusual serine-type protease. Proc. Natl. Acad. Sci. USA 95:49594963. Zhu, J., R. Kong, and C. P. Wolk. 1998. Regulation of hepA of Anabaena sp. strain PCC 7120 by elements 5‘ from the gene and by he9K.J. Bacteriol. 180:4233-4242. Zhu, J., K. Jager, T. Black, K. Zarka, 0. Koksharova, and C. P. Wolk. 2001. HcwA, an autolysin, is required for heterocyst maturation in Anabaena sp. strain PCC 7120. J. Bacteriol. 183:6841-6851. Zouni, A., H.-T. Witt, J. Kern, P. Fromme, N. Krauss, W. Saenger, and P. Orth. 2001. Crystal structure of photosystem I1 from Synechococcus elongatus at 3.8 A resolution. Nature 409:739-743.
Myxobacteria: Multicellularity and Differentiation Edited by David E. Whitworth 02008 ASM Press, Washington, D.C.
Marie A. Elliot Mark J. Buttner Justin R. Nodwell
Multicellular Development in Streptomyces
The colony structure (Fig. 1)and life cycle of the grampositive, soil-dwelling bacterium Streptomyces coelicolor provide a fascinating exception to the view of bacteria as simple unicellular microorganisms. Growth commences when spores encounter a suitable source of nutrients and germinate. The emerging vegetative cells, referred to as “substrate hyphae,” are filamentous rather than unicellular and grow by apical tip extension and branching, forming a tangled network of filamentous cells, referred to as a “substrate mycelium.” Septation is a relatively rare event in the substrate hyphae, with individual cells consisting of long compartments containing multiple chromosomes. Consequently, and in contrast to all other organisms investigated to date, cell division is dispensable for viability in streptomycetes, although it is necessary for sporulation later in their life cycle (McCormick et al., 1994; McCormick and Losick, 1996). As the vegetative colonies age, they produce a second filamentous cell type that grows into the air away from the substrate hyphae (typically occurring between 24 and 48 h under laboratory conditions). These “aerial hyphae” do not branch; rather, each is fated to undergo a single, synchronous round of septation that subdivides
it into 40 to 60 compartments of equal size. These “prespore” compartments, each containing a single chromosome, go on to metamorphose into spores (Fig. 1). Spores undergo a number of maturation steps, culminating in the deposition of a gray polyketide pigment on the spore surface that turns the aerial mycelium from white to gray (Davis and Chater, 1990; Kelemen et al., 1998). This aerial morphogenesis and formation of reproductive spores provide streptomycetes with a mechanism of dispersal to new environments, as they are otherwise nonmotile and exhibit no obvious chemotactic behavior. The initiation of aerial hypha formation coincides with the production of a diverse arsenal of secondary metabolites, many of which have significant medical application. Included among these are the majority of the medically important antibiotics, several antifungal drugs, and a variety of other important chemotherapeutic agents, such as anticancer drugs and immunosuppressants. S. coelicolor has been used extensively as a model for secondary metabolite production, in part because two of the antibiotics that it makes, actinorhodin and undecylprodigeosin, are pigmented blue and
Marie A. Elliot, Department of Biology, McMaster University, 1280 Main St. West, Hamilton, ON, L8S 4K1, Canada. Mark J. Buttner, Department of Molecular Microbiology, John Innes Centre, Colney Lane, Norwich, NR4 7UH, United Kingdom. Justin R. Nodwell, Department of Biochemistry and Biomedical Sciences, Health Sciences Centre, McMaster University, Hamilton, ON, L8N 3 2 5 , Canada.
419
ANALOGOUS SYSTEMS
420
Figure 1 Scanning electron micrograph of a side-on view of a mature S. coelicolor colony showing long chains of spores in the air supported by a layer of substrate mycelium (photo courtesy of Kim Findlay and Mark Buttner, John Innes Centre).
red, respectively, facilitating analysis. Intriguingly, in S. coelicolor the production of antibiotics not only is temporally correlated with the formation of aerial hyphae but also shares regulatory elements. Mutations in genes involved in morphogenesis alter colony appearance but do not usually compromise viability (nonsporulating mutants are easily propagated vegetatively). A number of genes required for the formation of aerial hyphae have been identified (Table l), many of which have been given a bld gene designation (for “bald”), reflecting the lack of fuzzy aerial hyphae that results from mutation of these genes. Of these bld genes, mutations in bldA, bldB, bldC, bldD, bldG, bldH, and bld] also impair antibiotic production, consistent with the existence of regulatory links between morphogenesis and secondary metabolism. The majority of genes identified as being important for aerial hypha formation encode regulatory proteins (Table 1);however, recent work has resulted in the characterization of two classes of structural molecules that are necessary for aerial development: the SapB surfactant peptide (specified by the ram gene cluster) and eight chaplin proteins (ChpA through H).
Cessation of aerial hyphal extension precedes the subdivision of the aerial filaments into single-genome prespore compartments, which subsequently round off and mature into spores (Flardh et al., 1999). A second class of developmental mutations, many of which have been given a whi gene designation (for “white”), allow the formation of aerial hyphae but prevent the differentiation of these reproductive structures into mature graypigmented spores, resulting in colonies which remain white, even on prolonged incubation. These genes include whiA, whiB, whiD, whiE, whiG, whiH, whil, whi], whiL, whiM, whiO, sigF, and the recently identified ssg family of genes. Of these, all but whiL, whiM, and whiO have been identified and studied at the molecular level (Table 2) (Ryding et al., 1999). While the first 20 years of research into the morphogenetic processes of S . coelicolor was mostly concerned with the identification and cloning of the genes listed above, recent progress, including in particular the use of fluorescence microscopy, has opened basic cellular processes such as cell division and chromosome segregation to scrutiny. We are finding that while Streptomyces has many of the conventional genes that are necessary
Table 1 Genes required for formation of aerial hyphae Gene name
SCO no.
SAV no.
Alternative name
bldA bldB
SC05723
SAV2529
bldC
SC04091
SAV4130
bldD
SC01489
SAV6861
bldG
sc03549
SAV46 14
bldH
SC02792
SAV5261
bldJ bldK bldL
Not cloned SCO5112-16 Not cloned
SAV3172-76
bldM
SC04768
SAV4998
whiK
bldN ramR
SC03323 SC06685
SAV4735 SAV7499
whiN amfR
ramCSAB
SCO6681-84
SAV7500-03
amfTSAB
chpA-H
SAV1230 (B); 6478 (E); 6636-37(C&H)
citA acoA cYa clpP1
SCO1674-75; 1800; 2699; 2705; 2716-17; 7257 SC02736 SC05999 SC04928 SC02619
SAV5330 SAV2258 SAV3329 SAV5447
catB dasR
SC00666 SC05231
SAV348 SAV3023
brgA
Not cloned
adpA
sacA
Product
Close homologuc distribution
Actinomycctcs
Lawlor ct al., 1987; Lcskiw et al., 1991 Pope et al., 1998; Ecclcston ct al., 2002, 2006 Hunt et al., 2005
Actinomycetcs
Elliot ct al., 1998, 1999,2001
Bacteria
Bigncll et al., 2000
Bacteria
Takano ct al., 2003
Bacteria
Willcy ct al., 1993 Nodwell et al., 1999 Nodwcll ct al., 1999
Bacteria
Molle and Buttner, 2000
Bacteria Bacteria
Bibb ct al., 2000 Ma and Kendall, 1994; Kcijscr ct al., 2002; Nguycn ct al., 2002; O’Connor ct al., 2002 Ma and Kcndall, 1994; O’Connor ct al., 2002; Kodani ct al., 2004
tRNA Small protein; may be DNA-binding Putative McrR-like DNA-binding protein DNA-binding transcription factor Putative anti-anti-sigma factor DNA-binding transcription factor
All organisms Actinomycctes
Oligopcptidc pcrmease Unknown; bldK-like characteristics Two-component response regulator ECF sigma factor Two-component response regulator
C: lantibiotic biosynthctic Streptomyccs protein; S: lantibioticlike molecule; A and B: permeases Sporulating Hydrophobic cell wall-associatcd proteins actinomycctcs Citrate synthasc Aconitasc Adcnylatc cyclasc Clp protcasc protcolytic subunit Catalasc GntR-like transcription factor
Reference(s)
Clacsscn et al., 2003; Elliot ct al., 2003
All organisms All organisms All organisms Bacteria
Viollicr ct al., 2001b Viollier et al., 2001a Derouaux et al., 2004 dc Cr6cy-Lagard ct al., 1999
Bacteria Bacteria
Cho ct al., 2000 Rigali et al., 2006 Shima et al., 1996
ANALOGOUS SYSTEMS
422 Table 2
Genes required for sporulation
Gene name
SCO no.
whiA whiB
SC01950 SC03034
whiD
SC04767
whiE
SC05314SC05321 SC05621
whiG
SAV no.
Unknown Putative transcription factor SAV4997 (wblB) Putative transcription factor SAV2835-2842 Polyketide; gray spore pigment SAV2630 Sigma factor
whir
SC06029
SAV2230
whi]
SCP4543
SAV3425
sigF
SC04035 SC02082
SAV4185 SAV6124
ssgR
SC03926 SC01541 SC0728 9; 6722; 3158; 7175; 2924 SC03925
SAV4267 SAV6810 SAV6722 (D); 3158 (E); 580 (F) SAV4268
samR
SC02935
SAV5141
ssgA ssgB ssgC-F
Gram-positive bacteria Actinomycetes
SAV6294 SAV5042
whiH
ftSZ
Close homologue distribution
Product
Actinomycetes
Reference(s) Ainsa et al., 2000 Davis and Chater, 1992; Soliveri et al., 2000 Molle et al., 2000; Jakimowicz et al., 2005b Davis and Chater, 1990
GntR-like transcription factor Two-component response regulator Putative transcription factor Sigma factor Tubulin-like cell division protein Possible regulator? Possible regulator? Possible regulator?
Bacteria
MCndez and Chater, 1987; Chater et al., 1989 Ryding et al., 1998
Bacteria
Ainsa et al., 1999
Actinomycetes Bacteria Bacteria
Gehring et al., 2000; Chater and Chandra, 2006 PotkkovA et al., 1995 McCormick et al., 1994
Sporulating actinomycetes Sporulating actinomycetes Sporulating actinomycetes
van Wezel et al., 2000 Keijser et al., 2003 Noens et al., 2005
IclR-like transcription factor IclR-like transcription factor
Bacteria
Traag et al., 2004
Bacteria
Tan et al., 2002
for these processes to occur, Streptomyces cells are organized very differently from other bacteria and these differences are highly relevant to colony development and spore formation.
GERMINATION AND APICAL GROWTH In simple rod-shaped bacteria such as Escherichia coli, cell growth occurs through the isotropic insertion of new cell wall material throughout the lateral walls. This is interrupted at intervals by cytokinesis (cell division) at the cell midpoint to create two new cells and hence two new poles. Further, replication of the E. coli chromosome and cytokinesis are tightly linked. This is in complete contrast to the mycelial growth habit of Streptomyces, in which growth arises solely through the deposition of new cell wall material at hyphal tips and chromosome replication occurs without associated cell division. The cues that trigger spore germination in Bacillus subtilis are relatively well understood (see Setlow, 2003, for a review); however, this is not the case for
Bacteria
Streptomyces, for which very little is currently known. Cyclic AMP has been implicated in the germination process, as mutations in both adenylate cyclase (Susstrunk et al., 1998) and a cyclic AMP-receptor-like protein-encoding gene (Derouaux et al., 2004) result in severe germination defects and other developmental abnormalities. Several groups have proposed that there is a stable mRNA population in dormant spores that is actively translated at the outset of germination (Quir6s et al., 1986; Mikulik et al., 2002); however, the resulting proteins have not yet been identified. A likely candidate for early expression in germinating spores is the classical cell-division-controlling protein, DivIVA. In B. subtilis, DivIVA is a key determinant of accurate division at the midpoint of the cell (Edwards and Errington, 1997). The protein is localized to the cell poles and is responsible for recruiting MinCD to this location, thereby preventing polar cell division during vegetative growth (Marston et al., 1998). In Streptomyces, however, DivIVA is an essential protein that does not seem to be associated with cell division but rather is
24. MULTICELLULAR DEVELOPMENT IN STREPTOMYCES crucial for coordinating cell wall growth (Flardh, 2003a, 2003b). As S. coelicolor spores germinate, DivIVA forms discrete foci that localize to, and remain associated with, the new cell tip (Color Plate l l a ) . All new tips that form as a result of branching similarly have DivIVA associated with their apices. Altering the abundance of DivIVA has dramatic consequences for cell growth (Flardh, 2003a, 2003 b). Overexpression of DivIVA during the initiation of germination results in cells that are incapable of germ tube emergence and instead become swollen and rounded, as if cell wall growth were delocalized. In contrast, overexpression of DivIVA in an established substrate mycelium results in the formation of large numbers of new branches, consistent with the activation of multiple new sites of cell wall synthesis. Overall, these results suggest that DivIVA establishes hyphal tip growth and that it is responsible, directly or indirectly, for the recruitment of the cell wall biosynthetic machinery to hyphal tips and new branch points. Tip-associated cell wall biosynthesis can be visualized in germlings stained with fluorescently labeled vancomycin (Color Plate l l b ) (Daniel and Errington, 2003; Flardh, 2003a, 2003b). Vancomycin interacts with the un-cross-linked peptide cross-bridges associated with nascent, immature peptidoglycan. Vancomycin also labels nascent peptidoglycan associated with new vegetative cross walls in Streptomyces. Similar observations using fluorescently labeled wheat germ agglutinin (Schwedock et al., 1997) and radioactively labeled peptidoglycan precursors (Brafia et al., 1982; Gray et al., 1990) support this conclusion. These data suggest that the majority of cell wall biosynthesis occurs at the growing tip of the hyphal cell. It is not known whether membrane or teichoic acid biosynthesis is similarly localized. This mode of cell wall biosynthesis and therefore cell growth differ significantly from those of all other bacteria studied to date. In E. coli and B. subtilis, penicillin-binding proteins (theenzymes responsible for the synthesis of new peptidoglycan) have specialized roles, with some found primarily at sites of septation and others occurring with an “irregular distribution” in the lateral wall (Scheffers et al., 2004). While these data are consistent with the production of new peptidoglycan at distinct foci, the foci themselves are more or less evenly dispersed across the cell wall and the deposition of peptidoglycan is therefore relatively even throughout the cell envelope (reviewed in Archibald et al., 1993). Other gram-positive organisms synthesize cell wall material in a more localized manner: in coccoid cells such as Staphylococcus aureus, all new peptidoglycan is associated with septum formation, starting in a band around the cell’s midpoint, and as the daughter cell grows, one half of the wall is newly
423
synthesized while the other half consists entirely of old cell wall material (reviewed in Giesbrecht et al., 1998). A biochemical picture of teichoic acid biosynthesis is emerging (reviewed in Neuhaus and Baddiley, 2003), but the issue of localized or delocalized biosynthesis has not been addressed. Despite these obvious differences, the S. coelicolor vegetative cell wall is indistinguishable by electron microscopy from that of other gram-positive bacteria, and they appear to have a similar battery of penicillin-binding proteins and teichoic acid biosynthetic enzymes. Questions of major importance are the manner in which the cell wall machinery is localized to the growing tip of Streptomyces hyphae and the nature of the interactions between these proteins and others sharing the same subcellular “address.”
METABOLIC TRIGGERS The switch from substrate hyphal growth to aerial growth is thought to coincide with the sensing of environmental stresses and nutrient deficiencies. Nitrate depletion, for instance, is known to coincide with the onset of aerial hypha formation (Karandikar et al., 1997), whiie the presence of glucose can inhibit aerial hypha formation (Redshaw et al., 1976). Similarly, high concentrations of N-acetylglucosamine arrest development during vegetative growth; this arrest is mediated by the regulator DasR, which controls the expression of several genes involved in sugar transport (ptsH, cry, and ptsl) (Rigali et al., 2006). Interestingly, many bld mutants are defective in their ability to sense glucose and consequently may be unable to sense/signal starvation (Pope et al., 1996).Most bld mutants are also unable to neutralize the organic acids that are secreted during vegetative growth, resulting in a decrease in environmental pH that is associated with inhibition of aerial hypha formation (Viollier et al., 2001b). It seems likely that a functional Krebs cycle is necessary for the initiation of aerial mycelium formation since S. coelicolor mutants lacking citrate synthase or aconitase have a bald phenotype (Viollier et al., 2001a, 2001b), and this may be associated with the ability to sense glucose. Similarly, a functional Krebs cycle is also required for the initiation of sporulation in Bacillus (Ireton et al., 1995). Aconitase is known to be a multifunctional enzyme, not only acting in the tricarboxylic acid cycle but also having the capacity to respond to oxidative stress through its posttranslational regulatory ability to bind to iron regulatory elements within mRNAs to maintain iron homeostasis (reviewed in Beinert and Kennedy, 1993). Oxidative stress has also been implicated in Streptomyces morphogenesis because deletion of a developmentally regulated catalase
424
gene (catB) blocks aerial mycelium formation (Cho et al., 2000). Intriguingly, deletion of the anti-sigma factor gene rsrA (which controls the activity of the disulfide stress response sigma factor, a”)does not affect the formation of aerial hyphae but blocks their differentiation into spore chains (Paget et al., 2001), while deletion of the anti-sigma factor gene rsuA (which is predicted to control the activity of the stress response sigma factor, uU)prevents the erection of aerial hyphae (Gehring et al., 2001). Furthermore, deletion of crB (the osmotic/ oxidative stress response sigma factor) prevents the formation of aerial hyphae (Lee et al., 2005), while deletion of the related stress response sigma factors uL and uM adversely affects sporulation (Lee et al., 2005). Recent transposon mutagenesis experiments have demonstrated developmental defects resulting from mutations in putative metabolic genes such as metH (involved in vitamin BIZ-dependentmethionine synthesis [Gehring et al., 2004]), bkdR (encoding a regulator of genes involved in the catabolism of branched-chain amino acids [Sprusansky et al., 2005]), and SC06938 (encoding a gene similar to those involved in fatty acid/ branched-chain amino acid degradation [Gehring et al., 20041). More-targeted investigations have revealed a cycle of glycogen storage and utilization correlating with the morphogenetic pathway (Bruton et al., 1995; Ye0 and Chater, 2005). At present it is not possible to form an integrated hypothesis based on these incongruent observations; however, they clearly suggest that nutrient availability, primary metabolism, and environmental stresses all have a significant impact on the developmental program.
SIGNALING AND REGULATION How environmental conditions are sensed and how extracellular signals are transmitted are currently unknown; however, there is much evidence to suggest that multiple signals contribute to the onset of morphological differentiation. A possible route for the transduction of these signals is through the action of permeases. One of the characterized bld loci, bldK, encodes an oligopeptide permease that is likely to import a signaling molecule necessary for the initiation of aerial hypha formation (Nodwell et al., 1996). The bldK mutant phenotype is “leaky,” such that sparse aerial hyphae are formed upon extended incubation; however, there are a number of oligopeptide permease gene clusters present in the S. coelicolor genome, one of which is highly similar to the bldK locus, and so it is conceivable that there is some degree of functional redundancy. Much work has gone into trying to identify
ANALOGOUS SYSTEMS the substrates of these transporters, as well as other signals involved in morphological differentiation (Nodwell and Losick, 1998); however, they have so far eluded identification. One class of signaling molecules that play very important roles in the biology of streptomycetes are the ybutyrolactones (Takano, 2006), which are distinct from the homoserine lactone signaling molecules involved in quorum sensing in gram-negative bacteria. In Streptomyces griseus, a y-butyrolactone called A-factor acts to induce the transcription of a pleiotropic regulator known as AdpA (for A-factor-dependent protein A), which controls both antibiotic production and aerial hypha formation (reviewed by Ohnishi et al., 2005). An AdpA homologue (AdpA,,) has also been implicated in aerial development in S. coelicolor (Takano et al., 2003; Nguyen et al., 2003), but production of this protein does not appear to depend upon a y-butyrolactone. In fact, there is currently no evidence to suggest that y-butyrolactones have any significant effect on development and morphogenesis in S. coelicolor, instead appearing to be required exclusively for the control of secondary metabolite production (Takano et al., 2000; Takano, 2006). The expression of AdpA,, does, however, require bldA (Takano et al., 2003; Nguyen et al., 2003), which encodes the only tRNA capable of efficiently translating the rare UUA leucine codon (Streptomyces DNA is typically -70% G+C, and there appear to be no essential genes containing a UUA codon in S. coelicolor [Lawlor et al., 1987; Leskiw et al., 19911). Recent work has shown that the inability of bldA mutants to make aerial mycelium is due exclusively to the presence of a TTA codon in adpAsc(which is also known as bldH) (Takano et al., 2003; Nguyen et al., 2003). Several AdpA,,-dependent genes have been identified, including the melC operon (Zhu et al., 2005), which specifies the melanin pigment, and a gene encoding a serine protease inhibitor (Kim et al., 2005); however, none of these appears to have an essential role in morphological differentiation. While the extracellular signals that contribute to morphological differentiation have not been identified, the control of development inside the cell is better understood. The majority of developmental genes that have been characterized in S. coelicolor appear to encode regulatory proteins, including sigma factors encoded by bldN (Bibb et al., ZOOO), whiG (Chater et al., 1989), and sigF (PotGckov6 et al., 1995); “orphaned” response regulators encoded by bldM (Molle and Buttner, 2000), whiI ( A h a et al., 1999), and ramR (Ma and Kendell, 1994); and GntR-like regulators, encoded by whiH (Ryding et al., 1998),devA (Hoskisson et al., 2006), and dasR (Rigali et al., 2007). The three characterized sigma
24. MULTICELLULAR DEVELOPMENT IN STREPTOMYCES factors each act at different stages of development: uBldN is required for aerial hypha formation but does not influis one of the earliestence antibiotic production; uwhiG acting sporulation factors, with mutations in whiG being epistatic to mutations in any other whi gene, while uFis required late in the sporulation process, during the spore maturation phase. Direct targets have been identified for two of these sigma factors: aBldN directs the transcription of the regulatory gene bZdM (Bibb et al., 2000), which has a very similar mutant phenotype, and oWhiC directs the transcription of the regulatory genes whiH (Ryding et al., 1998) and whiI (Ainsa et al., 1999).Unfortunately, no direct targets of BldM, WhiH, or WhiI have yet been identified; however, it is notable that whiH and certain ftsZ mutant alleles have similar sporulation phenotypes (Flardh et al., 2000). These sigma factor cascades are at least partially connected through the activity of BldD, which negatively regulates the expression of both bldN and whiG during vegetative growth (Elliot et al., 2001), as well as those of the stress-response sigma factor sigH (Kelemen et al., 2001) and an as-yet-uncharacterized regulator termed bdtA (Elliot et al., 2001). In S. griseus, BldD also controls the expression of the ram (or amf) gene cluster (Ueda et al., 2005); however, this connection is not conserved in S. coelicolor. The ram (for rapid aerial mycelium formation [Ma and Kendall, 19941) cluster consists of the response regulator gene rumR, which is positioned convergently to the rumCSAB operon. The expression of ramCSAB is completely dependent on direct activation by RamR, and ramCSAB is one of two gene clusters under its control (O’Connor et al., 2002, 2005; San Paolo et al., 2006). Expression of the entire ram cluster depends on the activity of all the bld genes examined (apart from bldM and bldN), although the nature of this dependence has yet to be elucidated. There are three classes of developmental genes that appear to be actinomycete specific (Chater and Chandra, 2006). bldB encodes a 98-amino-acidYdimeric protein which has an extremely low PI and is required for efficient antibiotic production and formation of aerial hyphae (Pope et al., 1998; Eccleston et al., 2002). The genes abaA and whiJ encode similar proteins that have been implicated in antibiotic production and spore formation, respectively (Fernhndez-Moreno et al., 1992; Gehring et al., 2000). The biochemical roles of these proteins are not well understood; however, the S . coelicolor genome encodes >20 members of this family (Gehring et al., 2000; Eccleston et al., 2002). whiB encodes the founding member of the “Wbl” (for WhiB-like) class of proteins (Soliveri et al., ZOOO), of which WhiD is also a member (Molle et al., 2000). These small proteins
425
(-100 amino acids) have four absolutely conserved Cys residues that are individually essential for protein function (Jakimowicz et al., 2005b). In WhiD and WhiB, these residues have been shown to coordinate the binding of an oxygen-sensitive (4Fe-4s) cluster, in a manner typical of many redox-regulated proteins (Jakimowicz et al., 2005b). The biochemical function of Wbl proteins has not been clearly resolved. Circumstantial evidence suggests that Wbl proteins might function as transcription factors (Steyn et al., 2002; Bentley et al., 2004; Molle et al., 2000; Jakimowicz et al., 2005b). However, recently Alam et al. (2007) presented evidence that Wbl proteins can function in vitro as protein disulfide reductases and that loss of the oxygen-sensitive [4Fe-4S] cluster is required to release enzymatic activity. Regardless of their biochemical function, the presence of an oxygen-sensitive [4Fe-4S] cluster in WhiB and WhiD means it is likely that they respond to redox changes or contribute to redox signaling during the course of development, influencing the expression of different subsets of genes accordingly (WhiB acts before WhiD in the sporulation process). A third, more specialized family of proteins, found only in the sporulating actinomycetes,is the ssg family, which was originally identified in S. griseus (Kawamoto et al., 1997). In S. griseus, ssgA expression depends on the activity of A-factor and AdpA (Yamazaki et al., 2003); whether such a connection between ssgA and AdpA,, exists in S. coelicolor has not been investigated.There are seven ssgAlike genes in S. coelicolor (ssgA-G [Keijser et al., 20031) and six in S. avermitilis. Like the wbls, these genes appear to play different roles at different stages of development (van Wezel et al., 2000; Keijser et al., 2003; Noens et al., 2005) and, as discussed below, may be involved in the control of cell division and growth cessation. Expression of ssgA is independent of all whi genes tested and depends completely on the upstream gene ssgR, which encodes an IclR-like regulator (Traag et al., 2004). A number of other predicted regulatory proteins have been identified and subjected to preliminary characterization: bldC encodes an unusual MerR-like regulator that lacks the effector domain and possesses only the DNA-binding domain (Hunt et al., 2005); and bldG encodes a protein bearing homology to anti-anti-sigma factors, although it has not been shown to interact with any known anti-sigma factors (Bignellet al., 2000). Currently there is nothing known about the targets of either of these proteins. It is readily apparent that while many genes have been implicated in development and many regulators have been identified, there is still much work that needs to be done to understand how these regulators coordinate their activity and amalgamate the signals needed for
ANALOGOUS SYSTEMS
426 differentiation to proceed. The advent of genomics technologies, like microarrays and proteomic techniques, will undoubtedly help to advance our understanding of these regulators and their targets.
MODIFICATIONS OF THE AERIAL CELL WALL While we are continuing to identify regulators and their targets involved in morphological differentiation, we are also now starting to make inroads into understanding the biochemical and structural components necessary for aerial hypha formation and sporulation. In order to break surface tension and grow into the air, streptomycetes must coat their aerial structures with a hydrophobic surface layer and, on rich media, they also have to synthesize a surfactant peptide called SapB. The hydrophobic surface layer (which in S. coelicolor is apparent as a basketwork of paired rodlets on the surface of spores and aerial hyphae [Fig. 21) was first recognized through electron microscopy in the early 1960s (Hopwood and Glauert, 1961; Wildermuth, 1970; Wildermuth et al.,
1971; Smucker and Pfister, 1978), and SapB was first identified in the early 1990s (Willey et al., 1991, 1993), yet both eluded genetic and biochemical characterization, even after the completion of the S. coelicolor genome sequence. The recent discovery of the protein constituents of the hydrophobic surface layer, the chaplins, and the solution of the structure and biochemical mode of production of SapB are therefore major breakthroughs. SapB production is dependent on all bld genes, except for bldM and bldN, and this production is limited to growth on rich media. SapB has recently been shown to be a “lantibiotic-like” molecule (Fig. 3A); lantibiotics are ribosomally synthesized oligopeptide antibiotics produced by gram-positive bacteria that are translated as inactive prepeptides and undergo extensive posttranslational modification before being cleaved to yield the mature peptide (Kodani et al., 2004). The characterization of SapB also explains the role of the ram gene cluster in Streptomyces development (Fig. 3B). SapB is the posttranslationally processed product of the rams gene (Kodani et al., 2004; Willey et al., 2006). rams, which is part of a four-gene operon (ramCSAB)under the control
Figure 2 Spore rodlet ultrastructure. Freeze-etch preparation of a spore showing the basketwork of paired rodlets characteristic of the hydrophobic surface layer of S. coelicolor aerial hyphae and spores. The sample was prepared by freeze-etching followed by the creation of a replica, which was examined by transmission electron microscopy. The spore envelopes are partly fractured away, revealing part of the spore wall and an expanse of the plasma membrane. Image kindly provided by Hansrudi Wildermuth and David Hopwood, John Innes Centre.
24. MULTICELLULAR DEVELOPMENT IN STREPTOMYCES
42 7
A
B SapB
Removal of leader
Rams Figure 3 (A) Covalen ructure of SapB (Kodani et al., 2004). Note that following dehydration of serine to Dha and lanthionine bridge formation by a nearby cysteine residue, both the Dha residue and Cys residues involved in thioether formation are, by convention, designated as alanine residues. (B) Model for SapB maturation and export complex (Willey et al., 2006). The unmodified rumS gene product (Rams) is modified by dehydration and thioether formation, presumably catalyzed by RamC, which is hypothesized to have LanM-like bifunctional enzyme activity. RamC is known to function as a dimer (Hudson and Nodwell, 2004) and is associated with the membrane (Hudson et al., 2002). The modified product, PreSapB, is exported, and the leader sequence is cleaved to yield mature SapB. Currently there are no good candidates for the leader peptidase, so it is unclear if processing occurs before, during, or following export.
of RamR, encodes a 42-amino-acid prepeptide that is the likely substrate of RamC, a possible lantibiotic synthetase. Modification of the C-terminal half of Rams by RamC results in the dehydration of four serine residues to dehydroalanine, two of which react with cysteine thi01s to create intramolecular lanthionine rings. Following
these modifications, the N-terminal 1eaL;r peptide is removed by an unknown protease, to yield the mature 21-amino-acid lantibiotic-like SapB surfactant molecule (Fig. 3A). Mutations resulting in the loss of SapB (deletion of ramR, rumC, or ramCSAB) give a bld mutant phenotype on rich media (Keijser et al., 2002; Nguyen
ANALOGOUS SYSTEMS
428
Inner membrane
I I
Figure 4 Localization of the chaplins. The long chaplins are covalently anchored to the cell wall a t their C termini. A peptidoglycan-spanning domain is shown as a thick black line, and the two chaplin domains are shown as hatched diamonds. The short chaplins consist of a single chaplin domain and are thought to be anchored to the cell wall by the long chaplins.
et al., 2002, O'Connor et al., 2002), but these mutant strains can produce aerial mycelium and spores on some minimal media, showing that there is a SapB-independent pathway for aerial hypha formation. The most likely candidates for mediating SapB-independent production of aerial hyphae are the chaplins (Fig. 4; Table 3 ) . The chaplin family consists of eight secreted proteins, ChpA through H (ChpA-H), that share a highly conserved hydrophobic "chaplin domain" (Claessen et al., 2003; Elliot et al., 2003). The three long chaplins (ChpA-C) have two chaplin domains and are covalently attached to the cell wall peptidoglycan of Table 3
aerial hyphae by a sortase enzyme, while the five short chaplins (ChpD-H) have a single chaplin domain and are likely anchored to the cell surface through the heteropolymerization of the short and long chaplins into hydrophobic chaplin fibrils. Unlike SapB, the chaplins are expressed on both rich and minimal media. Deletion of the chaplin genes results in a strain that is severely deficient in the production of aerial hyphae, particularly on minimal media. A model for the role of SapB and the chaplins in aerial hypha formation is shown in Fig. 5. SapB and the chaplins are similar in that they are highly hydrophobic
Cell wall decoration
Gene name
SCO no.
Protein product Hydrophobic cell wall-associated proteins
rdlAIB sapB
SC02716; 7257; 1674; 2717; 1800; 2699; 2705; 1675 SC02718; 2719 SC06682
Secreted cell wall-associated proteins Lantibiotic-like peptide surfactant
sapA sap C sapD sapE
SC00409 SCP1.297 SCP1.304 SCP1.303
Spore-associated protein Spore-associated protein Putative acetyltransferase Putative acyl dehydratase
chpA-H
Reference(s) Claessen et al., 2003; Elliot et al., 2003 Claessen et al., 2002 Willey et al., 1991; 1993; Kodani et al., 2004 Guijarro et al., 1998 Bentley et al., 2004 Bentley et al., 2004 Bentley et al., 2004
24. MULTICELLULAR DEVELOPMENT IN STREPTOMYCES
Figure 5
Model of morphogenetic protein activity in the formation of aerial hyphae in S. coelicolor. During vegetative growth (A and B), SapB is secreted (black circles) and assembles to form an amphiphilic sheet at the air-water interface (straight black line). This reduces surface tension and allows the emergence of aerial hyphae. Chaplin synthesis and secretion (hatched diamonds) begin during late vegetative growth (B) and continue throughout aerial hyphal growth ( C and D). The chaplins polymerize to form a hydrophobic sheath surrounding the aerial filament, which further facilitates the growth into the air.
429
ANALOGOUS SYSTEMS
430 and have surfactant activity believed to reduce surface tension at an air-water interface, a necessary function during the emergence of aerial hyphae from the moist vegetative environment. Interestingly, loss of both SapB and the chaplins results in a strain that is predominantly bald on all media, suggesting that they are the major surfactants needed for the physical emergence of hyphae into the air (Capstick et al., 2007). It is not currently known whether SapB and the chaplins interact. There is, however, evidence that suggests the chaplins interact with a third class of proteins termed the rodlins (Claessen et al., 2004) (Table 3). The rodlins were initially isolated in a search for fungal hydrophobin-like molecules in S. coelicolor and were identified as being important for the formation of a characteristic “rodlet ultrastructure” seen on the surface of aerial hyphae and spores of S. coelicolor (Fig. 2 ) (Claessen et al., 2002). Unlike the chaplins and SapB, the rodlins are not found universally in the sporulating streptomycetes, as S. auermitilis does not possess rodlin genes. The loss of the rodlin proteins and the concomitant loss of the rodlet ultrastructure do not affect aerial morphogenesis. It has been proposed that the rodlins interact with the chaplin proteins to form the paired rodlet ultrastructure and that the chaplin fibrils are disorganized inrodlin mutants (Claessenet al., 2004). Consistent with this, the surface ultrastructure of S. auermitilis aerial hyphae and spores is smooth and resembles that of S. coelicolor rodlin mutants (Claessen et al., 2004). Several other cell surface proteins have been identified in S. coelicolor, including the chromosomally encoded SapA (Guijarro et al., 1988), and the plasmid (SCP1)encoded SapC, D, and E (Bentley et al., 2004) (Table 3 ) . No function has been assigned to any of these proteins. Table 4
fts Q
fts w ftSX
ftSZ divIC divIVA ssgA-F
ssgR
CELL DIVISION AND CHROMOSOME PARTITIONING The genomes of S. coelicolor and S. auermitilis encode similar arrays of putative cell division proteins including probable or demonstrated orthologues of FtsE, FtsI, FtsK, FtsL, FtsQ, FtsW, FtsX, FtsZ, DivIC, and DivIVA (Table 4). The basic mechanism of Z-ring formation appears to be shared between S. coelicolor and other prokaryotes; however, there are important differences in how Streptomyces employs and executes cell division. Strikingly, a number of cell division genes that are important or essential for proper division in other bacteria are absent in Streptomyces, including FtsA, MinC, and MinD (important in E. coli and B. subtilis), FtsB, FtsN, and ZipA (E. coli), and ZapA and EzrA ( B . subtilis). Understanding how S. coelicolor can live without these proteins is an important priority. In addition to missing some proteins, S. coelicolor makes alternative uses of others. As described in “Germination and Apical Growth” above, the S. coelicolor DivIVA homologue drives and organizes cell wall biosynthesis at the hyphal tips (Flardh, 2003a, 2003b), while in B. subtilis, DivIVA
Cell division genes in S. coelicolor and S. auermitilis
Gene name
ftsE fts I fts K fts L
It is interesting that the chaplins, rodlins, and SapA have conventional signal peptides and appear to be secreted by the Sec pathway and SapC has a Seclike signal peptide but is not processed, while SapD and E have no obvious signal sequences and no readily apparent means of being secreted (SapB is likely secreted by the ABC transporter encoded by ramAB [S. H. Au-Young, T. J. O’Connor, and J. R. Nodwell, unpublished data]).
SCO no.
SAV no.
Protein product
SC02969 SC02090 SC05750 SC02091 SC02083 and SC03853 SC02085 SC02968 SC02082
SAV6104 SAV6116 SAV4542 SAV6115 SAV6123 andSAV4332 SAV61 21 SAV6105 SAV6124
ABC transporter Penicillin binding protein Membrane protein Small membrane protein Membrane protein
sc03095 SC02077 SC03926; 1541; 7289; 6722; 3158; 7175; 2924 sco3925
SAV3532 SAV6129 SAV4267; 6810; 580; 570; 1687; 3605
Integral membrane protein ABC transporter Tubulin-like cell division protein Small membrane protein Tip-associated protein Small cytoplasmic proteins
SAV4268
IclR-like transcription factor
Reference(s)
Wang et al., 2007 McCormick and Losick, 1996
McCormick et al., 1994
Flardh, 2003 van Wezel et al., 2000; Keijser et al., 2003; Noens et al., 2005 Traag et al., 2004
24. MULTICELLULAR DEVELOPMENT IN STREFTOMYCES
43 1
B
Figure 6 Transmission electron micrographs of S. coelicolor. (A) Substrate hyphae showing a branch (B) and a vegetative cross wall (VC). (B) Aerial hypha showing synchronous sporulation septation. Note that septation and chromosome segregation are taking place simultaneously.
acts to ensure accurate septum placement (Edwards and Errington, 1997). In Streptomyces, cell division is relatively rare in the substrate hyphae, and as a result each vegetative cell consists of a long chamber containing multiple chromosomes (Fig. 6A). In contrast, each aerial hypha undergoes a synchronous round of cell division that serves to divide it into 40 to 60 identical prespore compartments, each of which contains a single chromosome (Fig. 6B; Color Plate 12). Cell division is therefore more important for spore formation than for vegetative growth. Perhaps not unexpectedly, but in contrast to all other prokaryotes, cell division is dispensable for viability in Streptomyces (McCormick et al., 1994). Both ftsQ and ftsZ null mutants grow less vigorously than wild-type strains but can establish a substrate mycelium and raise a substantial aerial mycelium (McCormick et al., 1994; McCormick and Losick, 1996). They are unable to form spore
chains, however, as they cannot produce sporulation septa. Intriguingly, the rod-shape-determining protein MreB, an essential protein in B. subtilis, is also not essential in Streptomyces, with an mreB null mutant being unaffected in vegetative growth but defective in spore formation (Mazza et al., 2006). In B. subtilis, the initiation of sporulation is characterized by an asymmetric cell division event that divides a vegetative cell into a small forespore compartment and a larger mother cell compartment. This requires a redeployment of FtsZ ring formation from the middle of the cell to one side through the formation of a transient helical filament that appears to migrate longitudinally in a manner reminiscent of a corkscrew (Ben-Yehuda et al., 2002). Careful analysis of the localization of an FtsZeGFP fusion protein as a function of time in S. coelicolor aerial hyphae undergoing sporulation has led to a picture of sporulation septation that similarly involves several
432 classes of helical intermediates (Grantcharova et al., 2005). It is possible that an initial vegetative septation event at the base of a nascent aerial filament may serve a function analogous to that of the asymmetrically positioned septum in B. subtilis. The cytoplasmic concentration of FtsZ then increases in the aerial tip compartment and begins to segregate into short, separate helices that are relatively evenly spaced. These helices then resolve into FtsZ rings that are similar in appearance to those of other bacteria. Following this, sporulation septa are laid down to create the cylindrical unigenomic prespore compartments, which subsequently mature into a long chain of rounded spores. Vegetative and sporulation septa are structurally, functionally, and genetically distinct from each other: the vegetative septa are very similar to those of other grampositive bacteria but remain as fixed, peptidoglycancontaining cross walls separating adjacent compartments that do not progress to membrane constriction and cell separation. Sporulation septa are generally thicker and, unlike their vegetative counterparts, do support cytokinesis. It is likely that the differences between vegetative and sporulation septation are reflected in the proteins involved in septation and cytokinesis. Indeed, Grantcharova, Flardh, and coworkers employed an ingenious genetic selection to identify an ftsZ allele that permitted vegetative but not sporulation septation (Grantcharova et al., 2003). Mutants bearing only this allele grew in a manner typical of a wild-type strain and had nearly wildtype levels of vegetative septa; however, sporulation septation was almost completely abolished. Closer analysis revealed that while the mutant FtsZ was produced at elevated levels in aerial hyphae as is usual, it failed to assemble into the ladder-like array of rings characteristic of sporulation-associated septation. The mutation, an A249T substitution, was modeled onto the Methanococcus jannaschii FtsZ structure (Lowe and Amos, 1998) and revealed the Ala 249 to pack into the hydrophobic core of the protein. The threonine substitution must therefore cause a local rearrangement that disrupts the formation of helical filaments necessary for sporulation septation but not for the less-well-understood process that leads to vegetative septation. An important question is how S. coelicolor brings about different patterns of septum formation and cell division in the substrate and aerial hyphae. Part of the answer to this question appears to involve a simple regulatory mechanism. The ftsZ gene has three promoters in S. coelicolor (pl, p2, and p3), with evidence for transcriptional read-through from the upstream ftsQ gene as well (Flardh et al., 2000). Transcripts from the p l and p3 promoters and the read-through transcript from ftsQ were
ANALOGOUS SYSTEMS detectable at low levels throughout development and appear to be spatially confined to the substrate hyphae. In contrast, transcription from p2 is upregulated upon initiation of aerial hypha formation and appears to be restricted to the aerial filaments. The greater level of ftsZ expression in the aerial hyphae might therefore support a larger number of cell division events than is possible in the substrate hyphae. As might be expected based on this model, inactivation of the p2 promoter gives rise to a sporulation-defective phenotype (Flardh et al., 2000). The upregulation of p2 is compromised in whiA, whiB, whiG, whiH, whil, and whi] mutants (Flardh et al., 2000), all of which are defective in their ability to form sporulation septa. The direct regulator(s) of p2 has not been identified. One critical question is how aerial filaments ensure that septation takes place at evenly spaced intervals and how the resulting spores each acquire a single intact chromosome. Two mechanisms are known to control septum placement in other bacteria: the min system and nucleoid occlusion. Streptomyces has no homologues of the B. subtilis or E. coli MinC or MinD proteins and uses its DivIVA for functions apparently unrelated to cell division. If there is a related mechanism in Streptomyces, it must utilize genes that have very little or no sequence similarity to their counterparts in other bacteria. The phenotypes of two of the sporulation mutants, whiH and whir), are noteworthy in this regard. In wild-type S. coelicolor, nucleoid segregation is not seen until sporulation septa have started forming, and in whiA, whiB, and whiG mutants, which do not initiate septation, the DNA appears to be continuous throughout the aerial hyphae. In contrast, whiH mutants divide their aerial hyphae into compartments of highly variable size through the deposition of rare sporulation septa and within these aerial hyphae the nucleoids are partitioned into condensed bodies of different sizes and irregular distribution (Flardh et al., 1999).Thus, after terminating aerial growth, whiH mutants appear to initiate later sporulation processes but execute them incorrectly (Flardh et al., 1999).The whiD mutation impairs spore formation (75% fewer spores than the wild type) but also adversely affects cell division itself, as a number of mini compartment^" (much smaller than normal spore compartments) are produced (Molle et al., 2000). These minicompartments appear to lack DNA, implying that there was therefore no “guillotine” effect on chromosomes by the mispositioned septa (Molle et al., 2000). Nucleoid occlusion is less well understood than the min system: to date only two genes have been identified that are important for this process: noc in B. subtilis (Wu and Errington, 2004) and slmA in E. coli (Bernhardt and de Boer, 2005). S. coelicolor
24. MULTICELLULAR DEVELOPMENT I N STREPTOMYCES and S. avermitilis encode paralogues of both, but none of them has been investigated. In B. subtilis the SpoIIIE protein is important for transporting a chromosome into the forespore before the asymmetrically positioned septum comes down on it (Bath et al., 2000). S . coelicolor encodes several SpoIIIE-like proteins, and it is possible that one or more of these helps ensure that chromosomes have been segregated to the prespores prior to cytokinesis. Recent work by Wang et al. (2007) has shown that one SpoIIIE/FtsK homologue (SC05750)localizes to the sporulation septa and its deletion results in a chromosome segregation defect during sporulation, leading to a substantial number of spores containing chromosomes with deletions at one or both ends. A second SpoIIIE/ FtsK homologue (SC01416) has also been shown to localize to the sporulation septa; however, it appears to be involved in other or additional processes important for cell division and sporulation (Ausmees et al., in press). Perhaps the functions of some of the “missing” cell division genes are fulfilled by putative cell division genes that are specific to the streptomycetes. Recent work has identified a class of proteins referred to as the SALPs (for SsgA-like proteins). Null mutations in ssgA and ssgB cause defects in sporulation septation. Mutations in ssgC, ssgD, ssgF, and ssgG cause more-complex defects that include alterations in placement of sporulation septa, misshapen spores, or prespore compartments of altered size. A mutation in ssgE causes rapid autolysis at the sporulation septa such that free spores are released from the aerial hyphae prematurely (Noens et al., 2005). Other work has revealed an actinomycete-specificintegral membrane protein, CrgA, whose overexpression inhibits the formation of productive Z-rings, suggesting an important role in coordinating cytokinesis (Del Sol et al., 2006). Like the myxococci, the streptomycetes have a large chromosome: the S. coelicolor and S. avermitilis chromosomes have been sequenced and shown to have lengths of -8.7 and -9.0 Mb and to have 7,822 and 7,577 protein-coding genes, respectively (Bentley et al., 2002; Ikeda
433 et al., 2003). Unlike most other prokaryotes, however, the Streptomyces chromosomes are linear (Lin et al., 1993; Hopwood, 2006) and have evolved novel mechanisms for end replication. Chromosome replication initiates at a centrally located origin of replication and proceeds divergently to the ends of the chromosome, where it leaves 3’ leading strand overhangs (and correspondingly, 5 ’ recessed ends [Chang and Cohen, 19941).These telomerelike ends are then extended through the concerted activity of a telomere-binding complex. This complex consists of DNA polymerase, a topoisomerase, and two other streptomycete-specific proteins (Bao and Cohen, 2004). One of them is a telomere-protecting protein (Tpg),which is covalently attached to the 5’ end and may act as a primer for lagging strand DNA synthesis (Yang et al., 2002). The second is a telomere-associated protein that interacts with both Tpg and the 3’ overhang (Bao and Cohen, 2003). Relatively little is known about chromosome organization in Streptomyces, but it appears that the chromosome ends interact to form a circular structure, despite the fact that the S. coelicolor chromosome is linear (Yang and Losick, 2001). Whether this is mediated through interaction between the telomere-specific proteins (Tpg or telomere-associated protein) is currently unknown. Chromosome partitioning proceeds, at least in part, through the binding of ParB complexes to multiple pars sites surrounding the origin of replication (Jakimowicz et al., 2002,2005a) (Table 5 ) . As has been seen for f t s z , ParB accumulation is controlled in part by the whi genedependent upregulation of parB transcription during aerial hypha formation (Jakimowicz et al., 2006). It is not yet clear what, if any, other proteins are involved in chromosome segregation. Another big question that remains to be answered is what directs or marks the sites to which chromosomes will segregate in the aerial filaments: in Bacillus, RacA, a DNA-binding protein, associates with sequences surrounding the origin and anchors the chromosomes to the cell poles; however, such a protein is not found in Streptomyces.
Table 5 Partitioning and cytoskeletal genes Gene name
parA parB smc ftSZ
mreBlmb1
SCO no. SC038 86 SC038 87 9205577 SC02082
SAV no. SAV4309 SAV430 8 SAV2658 SAV6124
Protein product ATPase DNA binding protein Chromosome-associated ATPase Tubulin-like cell division protein
SC02611 SAV5455 Actin-like ATPase and SC02451 and SAV5720
Reference(s) Kim et al., 2000 Jakimowicz et al., 2002,20OSa, 2006 McCormick et al., 1994; Schwedock et al., 1997; Flardh et al., 2000; Grantcharova et al., 2005 Burger et al., 2000; Mazza et al., 2006
434
FUTURE DIRECTIONS It is becoming increasingly apparent that streptomycetes are unusual amongst prokaryotes, not only for their filamentous life cycle but also in how they execute fundamental processes such as spore germination, hyphal growth, and cell division. Given the complexity of development and metabolism in the streptomycetes, a major challenge will be to integrate our existing knowledge into a cohesive picture, where we understand how signaling and regulation are coordinated, what metabolic and environmental cues trigger a switch to aerial development and secondary metabolism, and how cell division and chromosome segregation are synchronized, both spatially and temporally. Addressing these questions will be facilitated not only by advances in postgenomic technologies but also by the development of fundamental genetic and cell biology tools, such as a robust transcriptional reporter system (Craney et al., 2007) and an expanding repertoire of fluorescent probes available for use in Streptomyces (Ausmees et al., in press).
References A h a , J. A., N. J. Ryding, N. Hartley, K. C. Findlay, C. J. Bruton, and K. F. Chater. 2000. WhiA, a protein of unknown function conserved among gram-positive bacteria, is essential for sporulation in Streptomyces coelicolor A3(2).J. Bacteriol. 1 8 2 5 4 7 0 4 4 7 8 . A k a , J. A., H. D. Parry, and K. F. Chater. 1999. A response regulator-like protein that functions at an intermediate stage of sporulation in Streptomyces coelicolor A3(2).Mol. Microbiol. 34:607-619. Alam, M. S., S. K. Garg, and P. Agrawal. 2007. Molecular function of WhiB4lRv3681c of Mycobacterium tuberculosis H37Rv: a [4Fe-4S] cluster co-ordinating protein disulphide reductase. Mol. Microbiol. 63:1414-1431. Archibald, A. R., I. C . Hancock, and C. R. Harwood. 1993. Cell wall structure, synthesis and turnover. In A. L. Sonenshein, J. A. Hoch, and R. Losick (ed.), Bacillus subtilis and Other Gram-Positive Bacteria: Biochemistry, Physiology, and Molecular Genetics. ASM Press, Washington, DC. Ausmees, N., H. Wahlstedt, S. Bagchi, M. Elliot, M. Buttner, and K. Flardh. SmeA, a small membrane protein with multiple functions in Streptomyces sporulation including targeting of a SpoIIIE/FtsK-domain protein to cell division septa. Mol. Microbiol., in press. Bao, K., and S. N. Cohen. 2003. Recruitment of terminal protein to the ends of Streptomyces linear plasmids and chromosomes by a novel telomere-binding protein essential for linear DNA replication. Genes Dev. 17:774-785. Bao, K., and S . N. Cohen. 2004. Reverse transcriptase activity innate to DNA polymerase I and DNA topoisomerase I proteins of Streptomyces telomere complex. Proc. Natl. Acad. Sci. USA 101:14361-14366. Bath, J., L. J. Wu, J. Errington, and J. C. Wang. 2000. Role of Bacillus subtilis SpoIIIE in DNA transport across the mother cell-prespore division septum. Science 290:995-997.
ANALOGOUS SYSTEMS Beinert, H., and M. C. Kennedy. 1993. Aconitase, a twofaced protein: enzyme and iron regulatory factor. FASEB J. 71442-1449. Bentley, S. D., S. Brown, L. D. Murphy, D. E. Harris, M. A. Quail, J. Parkhill, B. G. Barrell, J. R. McCormick, R. I. Santamaria, R. Losick, M. Yamasaki, H. Kinashi, C. W. Chen, G. Chandra, D. Jakimowicz, H. M. Kieser, T. Kieser, and K. F. Chater. 2004. SCP1, a 356,023 bp linear plasmid adapted to the ecology and developmental biology of its host, Streptomyces coelicolor A3(2).Mol. Microbiol. 5 1:1615-1628. Bentley, S. D., K. F. Chater, A. M. Cerdeno-Tarraga, G. L. Challis, N. R. Thomson, K. D. James, D. E. Harris, M. A. Quail, H. Kieser, D. Harper, A. Bateman, S. Brown, G. Chandra, C. W. Chen, M. Collins, A. Cronin, A. Fraser, A. Goble, J. Hidalgo, T. Hornsby, S. Howarth, C. H. Huang, T. Kieser, L. Larke, L. Murphy, K. Oliver, S. O’Neil, E. Rabbinowitsch, M. A. Rajandream, K. Rutherford, S. Rutter, K. Seeger, D. Saunders, s. Sharp, R. Squares, s. Squares, K. Taylor, T. Warren, A. Wietzorrek, J. Woodward, B. G. Barrell, J. Parkhill, and D. A. Hopwood. 2002. Complete genome sequence of the model actinomycete Streptomyces coelicolor A3(2). Nature. 417141-147. Ben-Yehuda, S., and R. Losick. 2002. Asymmetric cell division in B. subtilis involves a spiral-like intermediate of the cytokinetic protein FtsZ. Cell 109:257-266. Bernhardt, T. G., and l? A. de Boer. 2005. SlmA, a nucleoidassociated, FtsZ binding protein required for blocking septal ring assembly over chromosomes in E. coli. Mol. Cell 18~555-564. Bibb, M. J., V. Molle, and M. J. Buttner. 2000. uBldN, an extracytoplasmic function RNA polymerase sigma factor required for aerial mycelium formation in Streptomyces coelicolor A3(2).J. Bacteriol. 182:4606-4616. Bignell, D. R., J. L. Warawa, J. L. Strap, K. F. Chater, and B. K. Leskiw. 2000. Study of the bldG locus suggests that an antianti-sigma factor and an anti-sigma factor may be involved in Streptomyces coelicolor antibiotic production and sporulation. Microbiology 146:2161-2173. Braiia, A. F., M.-B. Manzanal, and C. Hardisson. 1982. Mode of cell wall growth of Streptomyces antibioticus. FEMS Microbiol. Lett. 13:231-235. Bruton, C. J., K. A. Plaskitt, and K. F. Chater. 1995. Tissuespecific glycogen branching isoenzymes in a multicellular prokaryote, Streptomyces coelicolor A3(2). Mol. Microbiol. 18539-99. Burger, A., K. Sichler, G. Kelemen, M. Buttner, and W. Wohlleben. 2000. Identification and characterization of the mre gene region of Streptomyces coelicolor A3(2). Mol. Gen. Genet. 263:1053-1060. Capstick, D. S., J. M. Willey, M. J. Buttner, and M. A. Elliot. 2007. SapB and the chaplins: connections between morphogenetic proteins in Streptomyces coelicolor. Mol. Microbiol. 64:602-613. Chang, P. C., and S. N. Cohen. 1994. Bidirectional replication from an internal origin in a linear streptomyces plasmid. Science 265:952-954. Chater, K. F., C. J. Bruton, K. A. Plaskitt, M. J. Buttner, C. MCndez, and J. D. Helmann. 1989. The developmental fate of S. coelicolor hyphae depends upon a gene product homologous with the motility sigma factor of B. subtilis. Cell 59:133-143.
24. MULTICELLULAR DEVELOPMENT IN STREPTOMYCES Chater, K. F., and G. Chandra. 2006. The evolution of development in Streptomyces analysed by genome comparisons. FEMS Microbiol. Rev. 30:651-672. Cho, Y. H., E. J. Lee, and J. H. Roe. 2000. A developmentally regulated catalase required for proper differentiation and osmoprotection of Streptomyces coelicolor. Mol. Microbiol. 35~150-160. Claessen, D., R. Rink, W. de Jong, J. Siebring, P. De Vreugd, F. G. Boersma, L. Dijkhuizen, and H. A. Wosten. 2003. A novel class of secreted hydrophobic proteins is involved in aerial hyphae formation in Streptomyces coelicolor by forming amyloid-like fibrils. Genes Dev. 17:1714-1726. Claessen, D., I. Stokroos, H. J. Deelstra, N. A. Penninga, C. Bormann, J. A. Salas, L. Dijkhuizen, and H. A. Wosten. 2004. The formation of the rodlet layer of streptomycetes is the result of the interplay between rodlins and chaplins. Mol. Microbiol. 53:433-443. Claessen, D., H. A. Wosten, G. van Keulen, 0. G. Faber, A. M. Alves, W. G. Meijer, and L. Dijkhuizen. 2002. Two novel homologous proteins of Streptomyces coelicolor and Streptomyces lividans are involved in the formation of the rodlet layer and mediate attachment to a hydrophobic surface. Mol. Microbiol. 44:1483-1492. Craney., A., T. Hohenauer, Y. Xu,N. K. Navani, Y. Li, and J. Nodwell. 1 March 2007. A synthetic luxCDABE gene cluster optimized for expression in high-GC bacteria. Nucleic Acids Res. 35:e46 [Epub ahead of print.] Daniel, R. A., and J. Errington. 2003. Control of cell morphogenesis in bacteria: two distinct ways to make a rod-shaped cell. Cell 113:767-776. Davis, N. K., and K. F. Chater. 1992. The Streptomyces coelicolor whiB gene encodes a small transcription factor-like protein dispensable for growth but essential for sporulation. Mol. Gen. Genet. 232:351-358. Davis, N. K., and K. F. Chater. 1990. Spore colour in Streptomyces coelicolor A3 (2) involves the developmentally regulated synthesis of a compound biosynthetically related to polyketide antibiotics. Mol. Microbiol. 4:1679-1691. de CrCcy-Lagard, V., P. Servant-Moisson, J. Viala, C. Grandvalet, and P. Mazodier. 1999. Alteration of the synthesis of the Clp ATP-dependent protease affects morphological and physiological differentiation in Streptomyces. Mol. Microbiol. 32505-517. Del Sol, R., J. G. Mullins, N. Grantcharova, K. Flardh, and P. Dyson. 2006. Influence of CrgA on assembly of the cell division protein FtsZ during development of Streptomyces coelicolor. J. Bacteriol. 188:1540-1550. Derouaux, A., S. Halici, H. Nothaft, T. Neutelings, G. Moutzourelis, J. Dusart, F. Titgemeyer, and S. Rigali. 2004. Deletion of a cyclic AMP receptor protein homologue diminishes germination and affects morphological development of Streptomyces coelicolor.J. Bacteriol. 186:1893-1897. Eccleston, M., R. A. Ali, R. Seyler, J. Westpheling, and J. Nodwell. 2002. Structural and genetic analysis of the BldB protein of Streptomyces coelicolor. J. Bacteriol. 184:42704276. Eccleston, M., A. Willems, A. Beveridge, and J. R. Nodwell. 2006. Critical residues and novel effects of over-expression of the Streptomyces coelicolor developmental protein BldB: evidence for a critical interacting partner. J Bacteriol. 188~8189-8195.
435
Edwards, D. H., and J. Errington. 1997. The Bacillus subtilis DivIVA protein targets to the division septum and controls the site specificity of cell division. Mol. Microbiol. 24:905915. Elliot, M., F. Damji, R. Passantino, K. Chater, and B. Leskiw. 1998. The 6ldD gene of Streptomyces coelicolor A3(2): a regulatory gene involved in morphogenesis and antibiotic production. J. Bacteriol. 180:1549-1555. Elliot, M. A., M. J. Bibb, M. J. Buttner, and B. K. Leskiw. 2001. BldD is a direct regulator of key developmental genes in Streptomyces coelicolor A3(2).Mol. Microbiol. 40:257-269. Elliot, M. A., N. Karoonuthaisiri, J. Huang, M. J. Bibb, S. N. Cohen, C. M. Kao, and M. J. Buttner. 2003. The chaplins: a family of hydrophobic cell-surface proteins involved in aerial mycelium formation in Streptomyces coelicolor. Genes Dev. 17~1727-1740. Elliot, M. A., and B. I(. Leskiw. 1999. The BldD protein from Streptomyces coelicolor is a DNA-binding protein. J. Bacteriol. 181:6832-6835. Fernandez-Moreno, M. A., A. J. Martin-Triana, E. Martinez, J. Niemi, H. M. Kieser, D. A. Hopwood, and F. Malpartida. 1992. abaA, a new pleiotropic regulatory locus for antibiotic production in Streptomyces coelicolor. ]. Bacteriol. 174:2958-2967. Flardh, K. 2003a. Essential role of DivIVA in polar growth and morphogenesis in Streptomyces coelicolor. Mol. Microbiol. 49~1523-1536. Flardh, K. 2003b. Growth polarity and cell division in Streptomyces. Curr. Opin. Microbiol. 6564-571. Flardh, K., K. C. Findlay, and K. F. Chater. 1999. Association of early sporulation genes with suggested developmental decision points in Streptomyces coelicolor A3 (2).Microbiology 145~2229-2243. Flkdh, K., E. Leibovitz, M. J. Buttner, and K. F. Chater. 2000. Generation of a non-sporulating strain of Streptomyces coelicolor A3(2) by the manipulation of a developmentally controlled ftsZ promoter. Mol. Microbiol. 38:737-749. Gehring, A. M., J. R. Nodwell, S. M. Beverley, and R. Losick. 2000. Genomewide insertional mutagenesis in Streptomyces coelicolor reveals additional genes involved in morphological differentiation. Proc. Natl. Acad. Sci. USA 97:9642-9647. Gehring, A. M., S. T. Wang, D. B. Kearns, N. Y. Storer, and R. Losick. 2004. Novel genes that influence development in Streptomyces coelicolor.J. Bacteriol. 186~3570-3577. Gehring, A. M., N. J. Yoo, and R. Losick. 2001. RNA polymerase sigma factor that blocks morphological differentiation by Streptomyces coelicolor. ]. Bacteriol. 183: 5991-599 6. Giesbrecht, P., T. Kersten, H. Maidhof, and J. Wecke. 1998. Staphylococcal cell wall: morphogenesis and fatal variations in the presence of penicillin. Microbiol. Mol. Biol. Rev. 62~1371-1414. Grantcharova, N., U. Lustig, and K. Flardh. 2005. Dynamics of FtsZ assembly during sporulation in Streptomyces coelicolor A3(2).J. Bacteriol. 187:3227-3237. Grantcharova, N., W. Ubhayasekera, S. L. Mowbray, J. R. McCormick, and K. Flardh. 2003. A missense mutation in ftsZ differentially affects vegetative and developmentally controlled cell division in Streptomyces coelicolor A3(2). Mol. Microbiol. 47645-656.
436 Gray, D. I., G. W. Gooday, and J. I. Prosser. 1990. Apical hyphal extension in Streptomyces coelicolor A3(2).J. Gen. Microbiol. 136:1077-1084. Guijarro, J., R. Santamaria, A. Shauer, and R. Losick. 1988. Promoter determining the timing and spatial localization of transcription of a cloned Streptomyces coelicolor gene encoding a spore-associated polypeptide. J. Bacteriol. 170:1895-1 901. Hopwood, D. A. 2006. Soil to genomics: the Streptomyces chromosome. Annu. Rev. Genet. 4O:l-23. Hopwood, D. A., and A. M. Glauert. 1961. Electron microscope observations on the surface structures of Streptomyces violaceoruber. 1. Gen. Microbiol. 26:325-330. Hoskisson, P. A., S . Rigali, K. Fowler, K. C. Findlay, and M. J. Buttner. 2006. DevA, a GntR-like transcriptional regulator required for development in Streptomyces coelicolor. J. Bacteriol. 188:50 14-5023. Hudson, M. E., D. Zhang, and J. R. Nodwell. 2002. Membrane association and kinase-like motifs of the RamC protein of Streptomyces coelicolor. J. Bacteriol. 184:4920-4924. Hudson, M. E., and J. R. Nodwell. 2004. Dimerization of the RamC morphogenetic protein of Streptomyces coelicolor. J. Bacteriol. 186:1330-1336. Hunt, A. C., L. Servin-Gonzklez, G. H. Kelemen, and M. J. Buttner. 2005. The bldC developmental locus of Streptomyces coelicolor encodes a member of a family of small DNAbinding proteins related to the DNA-binding domains of the MerR family.]. Bacteriol. 187:716-728. Ikeda, H., J. Ishikawa, A. Hanamoto, M. Shinose, H. Kikuchi, T, Shiba, Y. Sakaki, M. Hattori, and S. Omura. 2003. Complete genome sequence and comparative analysis of the industrial microorganism Streptomyces avermitilis. Nut. Biotechnol. 21526-53 1. Ireton, K., S. Jin, A. D. Grossman, and A. L. Sonenshein. 1995. Krebs cycle function is required for activation of the SpoOA transcription factor in Bacillus subtilis. Proc. Natl. Acad. Sci. USA 92:2845-2849. Jakimowicz, D., K. Chater, and J. Zakrzewska-Czerwinska. 2002. The ParB protein of Streptomyces coelicolor A3(2) recognizes a cluster of pars sequences within the originproximal region of the linear chromosome. Mol. Microbiol. 45:1365-1377. Jakimowicz, D., B. Gust, J. Zakrzewska-Czenvinska, and K. F. Chater. 2005a. Developmental-stage-specificassembly of ParB complexes in Streptomyces coelicolor hyphae. J. Bacteriol. 187:3572-3 5 80. akimowicz, D., S . MOW,J. Zakrzewska-Czerwinska, and K. F. Chater. 2006. Developmental control of a parAB promoter leads to formation of sporulation-associated ParB complexes in Streptomyces coelicolor. J. Bacteriol. 188: 1710-1 720. akimowicz, P., M. R. Cheesman, W. R. Bishai, K. F. Chater, A. J. Thomson, and M. J. Buttner. 2005b. Evidence that the Streptomyces developmental protein WhiD, a member of the WhiB family, binds a [4Fe-4S] cluster. J. Biol. Chem. 280:8309-8315. Karandikar, A., G. P. Sharples, and G. Hobbs. 1997. Differentiation of Streptomyces coelicolor A3(2) under nitratelimited conditions. Microbiology 143:358 1-3590.
ANALOGOUS SYSTEMS Kawamoto, S., H. Watanabe, A. Hesketh, J. C. Ensign, and K. Ochi. 1997. Expression analysis of the ssgA gene product, associated with sporulation and cell division in Streptomyces griseus. Microbiology 143:1077-1086. Keijser, B. J., E. E. Noens, B. Kraal, H. K. Koerten, and G. P. van Wezel. 2003. The Streptomyces coelicolor ssgB gene is required for early stages of sporulation. FEMS Microbiol. Lett. 22559-67. Keijser, B. J., G. P. van Wezel, G. W. Canters, and E. Vijgenboom. 2002. Developmental regulation of the Streptomyces lividans ram genes: involvement of RamR in regulation of the ramCSAB operon. J. Bacteriol. 184:4420-4429. Kelemen, G. H., P. Brian, K. Fliirdh, L. Chamberlin, K. F. Chater, and M. J. Buttner. 1998. Developmental regulation of transcription of whiE, a locus specifying the polyketide spore pigment in Streptomyces coelicolor A3(2).J. Bacteriol. 180~2515-2521. Kelemen, G. H., P. H. Viollier, J. Tenor, L. Marri, M. J. Buttner, and C. J. Thompson. 2001. A connection between stress and development in the multicellular prokaryote Streptomyces coelicolor A3(2).Mol. Microbiol. 40:804-814. Kim, D. W.,K. Chater, K. J. Lee, and A. Hesketh. 2005. Changes in the extracellular proteome caused by the absence of the bldA gene product, a developmentally significant tRNA, reveal a new target for the pleiotropic regulator AdpA in Streptomyces coelicolor. J. Bacteriol. 1872957-2966. Kim, H. J., M. J. Calcutt, F. J. Schmidt, and K. F. Chater. 2000. Partitioning of the linear chromosome during sporulation of Streptomyces coelicolor A3(2) involves an oriC-linked parAB locus. J. Bacteriol. 182:1313-1320. Kodani, S., M. E. Hudson, M. C. Durrant, M. J. Buttner, J. R. Nodwell, and J. M. Willey. 2004. The SapB morphogen is a lantibiotic-like peptide derived from the product of the developmental gene rams in Streptomyces coelicolor. Proc. Natl. Acad. Sci. USA 101:11448-11453. Lawlor, E. J., H. A. Baylis, and K. F. Chater. 1987. Pleiotropic morphological and antibiotic deficiencies result from mutations in a gene encoding a tRNA-like product in Streptomyces coelicolor A3(2). Genes Dev. 1:1305-1310. Lee, E. J., N. Karoonuthaisiri, H. S. Kim, J. H. Park, C. J. Cha, C. M. Kao, and J. H. Roe. 2005. A master regulator crB governs osmotic and oxidative response as well as differentiation via a network of sigma factors in Streptomyces coelicolor. Mol. Microbiol. 571252-1264. Leskiw, B. K., M. J. Bibb, and K. F. Chater. 1991. The use of a rare codon specificallyduring development? Mol. Microbiol. 5~2861-2867. Lin, Y. S., H. M. Kieser, D. A. Hopwood, and C. W. Chen. 1993. The chromosomal DNA of Streptomyces lividans 66 is linear. Mol. Microbiol. 10:923-933. Lowe, J., and L. A. Amos. 1998. Crystal structure of the bacterial cell division protein FtsZ. Nature 391:203-206. Ma, H., and K. Kendall. 1994. Cloning and analysis of a gene cluster from Streptomyces coelicolor that causes accelerated aerial mycelium formation in Streptomyces lividans. J. Bacteriol. 176:3800-3811. Marston, A. L., H. B. Thomaides, D. H. Edwards, M. E. Sharpe, and J. Errington. 1998. Polar localization of the MinD protein of Bacillus subtilis and its role in selection of the mid-cell division site. Genes Dev. 21:3419-3430.
24. MULTICELLULAR DEVELOPMENT IN STREPTOMYCES Mazza, P., E. E. Noens, K. Schirner, N. Grantcharova, A. M. Mommaas, H. K. Koerten, G. Muth, K. Fliirdh, G. P. Van Wezel, and W. Wohlleben. 2006. MreB of Streptomyces coelicolor is not essential for vegetative growth but is required for the integrity of aerial hyphae and spores. Mol. Microbiol. 60: 838-852. McCormick, J. R., and R. Losick. 1996. Cell division gene ftsQ is required for efficient sporulation but not growth and viability in Streptomyces coelicolor A3(2). J. Bacteriol. 178:5295-5301. McCormick, J. R.,E. P. Su, A. Driks, and R. Losick. 1994. Growth and viability of Streptomyces coelicolor mutant for the cell division gene ftsZ. Mol. Microbiol. 14:243-254. MCndez, C., and K. F. Chater. 1987. Cloning of whiG, a gene critical for sporulation of Streptomyces coelicolor A3(2). J. Bacteriol. 1695715-5720. Mikulik, K., J. Bobek, S. BezouSkova, 0. Benada, and 0. Kofrokva. 2002. Expression of proteins and protein kinase activity during germination of aerial spores of Streptomyces granaticolor. Biochem. Biophys. Res. Commun. 299:335342. Molle, V., and M. J. Buttner. 2000. Different alleles of the response regulator gene 6ldM arrest Streptomyces coelicolor development at distinct stages. Mol. Microbiol. 36:12651278. Molle, V., W. J. Palframan, K. C. Findlay, and M. J. Buttner. 2000. WhiD and WhiB, homologous proteins required for different stages of sporulation in Streptomyces coelicolor A3(2).J. Bacteriol. 182:1286-1295. Neuhaus, F, C., and J. Baddiley. 2003. A continuum of anionic charge: structures and functions of D-alanyl-teichoic acids in gram-positive bacteria. Microbiol. Mol. Biol. Rev. 67:686723. Nguyen, K. T., J. M Willey, L. D. Nguyen, L. T. Nguyen, P. H. Viollier, and C. J. Thompson. 2002. A central regulator of morphological differentiation in the multicellular bacterium Streptomyces coelicolor. Mol. Microbiol. 46:1223-123 8. Nguyen, K. T.,J. Tenor, H. Stettles, L. T. Nguyen, L. D. Nguyen, and C. J. Thompson. 2003. Colonial differentiation in Streptomyces coelicolor depends on translation of a specific codon within the adpA gene.]. Bacteriol. 185:7291-7296. Nodwell, J. R., and R. Losick. 1998. Purification of an extracellular signaling molecule involved in production of aerial mycelium by Streptomyces coelicolor. J. Bacteriol. 180~1334-1337. Nodwell, J. R., I<. McGovern, and R. Losick. 1996. An oligopeptide permease responsible for the import of an extracellular signal governing aerial mycelium formation in Streptomyces coelicolor. Mol. Microbiol. 22:8 81-893. Nodwell, J. R., M. Yang, D. Kuo, and R. Losick. 1999. Extracellular complementation and the identification of additional genes involved in aerial mycelium formation in Streptomyces coelicolor. Genetics 151569-5 84. Noens, E. E., V. Mersinias, B. A. Traag, C. P. Smith, H. K. Koerten, and G. P. van Wezel. 2005. SsgA-like proteins determine the fate of peptidoglycan during sporulation of Streptomyces coelicolor. Mol. Microbiol. 58:929-944. O’Connor, T. J., P. Kanellis, and J. R. Nodwell. 2002. The ramC gene is required for morphogenesis in Streptomyces
43 7
coelicolor and expressed in a cell type-specific manner under the direct control of RamR. Mol. Microbiol. 45: 45-57. O’Connor, T. J., and J. R. Nodwell. 2005. Pivotal roles for the receiver domain in the mechanism of action of the response regulator RamR of Streptomyces coelicolor. J. Mol. Biol. 351:1030-1047. Ohnishi, Y., H. Yamazaki, J. Y. Kato, A. Tomono, and S. Horinouchi. 2005. AdpA, a central transcriptional regulator in the A-factor regulatory cascade that leads to morphological development and secondary metabolism in Streptomyces griseus. Biosci. Biotechnol. Biochem. 69:431-439. Paget, M. S., J. B. Bae, M. Y. Hahn, W. Li, C. Kleanthous, J. H. Roe, and M. J. Buttner. 2001. Mutational analysis of RsrA, a zinc-binding anti-sigma factor with a thiol-disulphide redox switch. Mol. Microbiol. 39:1036-1047. Pope, M. K., B. D. Green, and J. Westpheling. 1996. The bld mutants of Streptomyces coelicolor are defective in the regulation of carbon utilization, morphogenesis and cell-cell signalling. Mol. Microbiol. 19:747-756. Pope, M. K., B. Green, and J. Westpheling. 1998. The bldB gene encodes a small protein required for morphogenesis, antibiotic production, and catabolite control in Streptomyces coelicolor. J. Bacteriol. 180:1556-1562. Potiickova, L., G. H. Kelemen, K. C. Findlay, M. A. Lonetto, M. J. Buttner, and J. Kormanec. 1995. A new RNA polymerase sigma factor, sigma F, is required for the late stages of morphological differentiation in Streptomyces spp. Mol. Microbiol. 1237-48. Quiros, L. M., C. Hardisson, and J. A. Salas. 1986. Isolation and properties of Streptornyces spore membranes. 1.Bacterial. 165:923-928. Redshaw, P. A., P. A. McCann, L. Sankaran, and B. M. Pogell. 1976. Control of differentiation in streptomycetes: involvement of extrachromosomal deoxyribonucleic acid and glucose repression in aerial mycelia development. J. Bacteriol. 125:698-705. Rigali, S., H. Nothaft, E. E. Noens, M. Schlicht, S. Colson, M. Miiller, B. Joris, H. K. Koerten, D. A. Hopwood, F. Titgemeyer, and G. P. van Wezel. 2006. The sugar phosphotransferase system of Streptomyces coelicolor is regulated by the GntR-family regulator DasR and links N-acetylglucosamine metabolism to the control of development. Mol. Microbiol. 61~1237-1251. Ryding, N. J., M. J. Bibb, V. Molle, I(. C. Findlay, K. F. Chater, and M. J. Buttner. 1999. New sporulation loci in Streptomyces coelicolor A3(2).]. Bacteriol. 1815419-5425. Ryding, N. J., G. H. Kelemen, C. A. Whatling, K. Flardh, M. J. Buttner, and K. F. Chater. 1998. A developmentally regulated gene encoding a repressor-like protein is essential for sporulation in Streptomyces coelicolor A3(2). Mol. Microbiol. 29:343-357. San Paolo, S., J. Huang, S. N. Cohen, and C. J. Thompson. 2006. rag genes: novel components of the RamR regulon that trigger morphological differentiation in Streptomyces coelicolor. Mol. Microbiol. 61:1167-1186. Scheffers, D. J., L. J. Jones, and J. Errington. 2004. Several distinct localization patterns for penicillin-binding proteins in Bacillus subtilis. Mol. Microbiol. 51:749-764.
438 Schwedock, J., J. R. McCormick, E. R. Angert, J. R. Nodwell, and R. Losick. 1997. Assembly of the cell division protein FtsZ into ladder-like structures in the aerial hyphae of Streptomyces coelicolor. Mol. Microbiol. 25:847-858. Setlow, P. 2003. Spore germination. Curr. Opin. Microbiol. 6550-556. Shima, J., A. Hesketh, S. Okamoto, S. Kawamoto, and K. Ochi. 1996. Induction of actinorhodin production by rpsL (encoding ribosomal protein S12) mutations that confer streptomycin resistance in Streptomyces lividans and Streptomyces coelicolor A3(2).J. Bacteriol. 178:7276-7284. Smucker, R. A., and R. M. Pfister. 1978. Characteristics of Streptomyces coelicolor A3(2) aerial spore rodlet mosaic. Can. J. Microbiol. 24:397-408. Soliveri, J. A., J. Gomez, W. R. Bishai, and K. F. Chater. 2000. Multiple paralogous genes related to the Streptornyces coelicolor developmental regulatory gene whiB are present in Streptomyces and other actinomycetes. Microbiology 146:33 3-343. Sprusansky, O., K. Stirrett, D. Skinner, C. Denoya, and J. Westpheling. 2005. The bkdR gene of Streptomyces coelicolor is required for morphogenesis and antibiotic production and encodes a transcriptional regulator of a branched-chain amino acid dehydrogenase complex. J. Bucteriol. 187664-671. Steyn, A. J., D. M. Collins, M. K. Hondalus, W. R. Jacobs, Jr., R. P. Kawakami, and B. R. Bloom. 2002. Mycobacterium tuberculosis WhiB3 interacts with RpoV to affect host survival but is dispensable for in vivo growth. Proc. Natl. Acad. Sci. USA 99:3147-3152. Susstrunk, U., J. Pidoux, S. Taubert, A. Ullmann, and C. J. Thompson. 1998. Pleiotropic effects of CAMP on germination, antibiotic biosynthesis and morphological development in Streptomyces coelicolor. Mol. Microbiol. 30:33-46. Takano, E., T. Nihira, Y. Hara, J. J. Jones, C. J. Gershater, Y. Yamada, and M. Bibb. 2000. Purification and structural determination of SCB1, a gamma-butyrolactone that elicits antibiotic production in Streptomyces coelicolor A3(2). J. Biol. Chem. 275:11010-11016. Takano, E., M. Tao, F. Long, M. J. Bibb, L. Wang, W. Li, M. J. Buttner, M. J. Bibb, Z. X. Deng, and K. F. Chater. 2003. A rare leucine codon in adpA is implicated in the morphological defect of bldA mutants of Streptomyces coelicolor. Mol. Micro biol. 5 0:475-4 86. Takano, E. 2006. y-butyrolactones: Streptomyces signaling molecules regulating antibiotic production and differentiation. Curr. Opin. Microbiol. 9:287-294. Tan, H., Y. Tian, H. Yang, G. Liu, and L. Nie. 2002. A novel Streptomyces gene, sumR, with different effects on differentiation of Streptomyces ansochromogenes and Streptomyces coelicolor. Arch. Microbiol. 177274-278. Traag, B. A., G. H. Kelemen, and G . P. van Wezel. 2004. Transcription of the sporulation gene ssgA is activated by the IclR-typeregulator SsgR in a whi-independent manner in Streptomyces coelicolor A3(2).Mol. Microbiol. 53:985-1000. Ueda, K., H. Takano, M. Nishimoto, H. Inaba, and T. Beppu. 2005. Dual transcriptional control of amfTSBA, which regulates the onset of cellular differentiation in Streptomyces griseus. J. Bacteriol. 187135-142.
ANALOGOU s SYSTEMS van Wezel, G. P., J. van der Meulen, S. Kawamoto, R. G. Luiten, H. K. Korten, and B. Kraal. 2000. ssgA is essential for sporulation of Streptomyces coelicolor A3(2) and affects hyphal development by stimulating septum formation. J. Bacteriol. 1825653-5662. Viollier, P. H., K. T. Nguyen, W. Minas, M. Folcher, G. E. Dale, and C. J. Thompson. 2001a. Roles of aconitase in growth, metabolism, and morphological differentiation of Streptomyces coelicolor.J. Bacteriol. 183:3193-3203. Viollier, l? H., W. Minas, G. E. Dale, M. Folcher, and C. J. Thompson. 2001b. Role of acid metabolism in Streptomyces coelicolor morphological differentiation and antibiotic biosynthesis. J. Bacteriol. 183:3184-3192. Wang, L., Y. Yu, X. He, X. Zhou, Z. Deng, K. F. Chater, and M. Tao. 2007. Role of an FtsK-like protein in genetic stability in Streptomyces coelicolor A3(2).J. Bacteriol. 189:2310-2318. Wildermuth, H. 1970. Surface structure of streptomycete spores as revealed by negative staining and freeze-etching. J. Bucteriol. 101:318-322. Wildermuth, H., E. Wehrli, and R. W. Horne. 1971. The surface structure of spores and aerial mycelium in Streptomyces coelicolor. J. Ultrustruct. Res. 35:168-180. Willey, J., R. Santamaria, J. Guijarro, M. Geistlich, and R. Losick. 1991. Extracellular complementation of a developmental mutation implicates a small sporulation protein in aerial mycelium formation by S. coelicolor. Cell 65:641-650. Willey, J., J. Schwedock, and R. Losick. 1993. Multiple extracellular signals govern the production of a morphogenetic protein involved in aerial mycelium formation by Streptomyces coelicolor. Genes Dev. 7:895-903. Willey, J. M., A. Willems, S. Kodani, and J. R. Nodwell. 2006. Morphogenetic surfactants and their role in the formation of aerial hyphae in Streptomyces coelicolor. Mol. Microbiol. 59:73 1-742. Wu, L. J., and J. Errington. 2004. Coordination of cell division and chromosome segregation by a nucleoid occlusion protein in Bacillus subtilis. Cell 117:915-925. Yamazaki, H., Y. Ohnishi, and S. Horinouchi. 2003. Transcriptional switch on of ssgA by A-factor, which is essential for spore septum formation in Streptomyces griseus. J. Bacteriol. 185:1273-1 2 83. Yang, C. C., C. H. Huang, C. Y. Li, Y. G. Tsay, S. C. Lee, and C. W. Chen. 2002. The terminal proteins of linear Streptomyces chromosomes and plasmids: a novel class of replication priming proteins. Mol. Microbiol. 43:297-305. Yang, M. C., and R. Losick. 2001. Cytological evidence for association of the ends of the linear chromosome in Streptomyces coelicolor. J. Bacteriol. 183:5180-5186. Yeo, M., and K. Chater. 2005. The interplay of glycogen metabolism and differentiation provides an insight into the developmental biology of Streptomyces coelicolor. Microbiology 151:855-861. Zhu, D., X. He, X. Zhou, and Z. Deng. 2005. Expression of the melC operon in several Streptomyces strains is positively regulated by AdpA, an AraC family transcriptional regulator involved in morphological development in Streptomyces coelicolor. J. Bacteriol. 187:3180-3 187.
Myxobacteria: Multicellulurity and Differentiation Edited by David E. Whinvorth 0 2008 ASM Press, Washington, D.C.
Derrick Brazill Richard H. Gomer
A Eukaryotic Neighbor:
25
Dictyostelium discoideum
A SIMPLE DEVELOPMENTAL CYCLE MAKES DTCTYOSTELTUM USEFUL FOR THE STUDY OF BASIC MECHANISMS The first study of Dictyostelium was done by Brefeld (Brefeld, 1869), who found the organism on horse dung and, having observed the aggregation streams and fruiting bodies, combined the Greek words dikty (net-like) and stili (column) to name this remarkable organism. Phylogenetic analysis suggests that Dictyostelium separated from the animal/fungus lineage after the intitial divergence of plants, but before fungi diverged from animals (Eichinger et al., 2005). Dictyostelium appears to be ubiquitous in soils throughout the world, where it preys upon a wide variety of bacteria and possibly yeasts (Cavender and Raper, 1968). The relative simplicity of Dictyostelium discoideum lends itself to the study of many fundamental questions (Kessin, 2001). This eukaryote normally exists as vegetative amoebae that eat bacteria on soil and decaying leaves. The amoebae, which are haploid, increase in number by fission. When the amoebae are starved for bacteria, they cease dividing and begin secreting an 80-kDa glycoprotein called conditioned medium factor (CMF). When there is a high density of starving cells, as indicated by a
high concentration of CMF (Jain et al., 1992; Yuen et al., 1995), the cells aggregate using relayed pulses of cyclic AMP (CAMP)as the chemoattractant (Aubry and Firtel, 1999) (Fig. 1).Aggregation occurs between 5 and 10 h after starvation. The aggregating cells form large streams that break up into groups of -20,000 cells. Each group forms a wormlike slug that crawls towards light. When the slug finds itself in a brightly lit, dry, open area (a location favorable for spore dispersal), it develops into a fruiting body consisting of a mass of spore cells supported on a -2-mm-high column of stalk cells. Approximately 10 to 20% of the cells vacuolate and become stalk cells; the rest are found in the spore mass. There are many basic mechanisms that can be studied using Dictyostelium as a model system. At the single-cell level, these include nutrient and starvation sensing, as well as the impact of these pathways on cell proliferation, directed cell motility, symmetry breaking, and cell type differentiation. At the multicellular level, one can study how the composition of a tissue or group of cells is sensed and maintained, as the starving population of Dictyostelium cells has a mechanism to sense the percentage of starving cells. In addition, cells in the slug appear to be able to sense the percentage of stalk and spore
Derrick Brazill, Department of Biology, Hunter College, 695 Park Ave., New York, NY 10021. Richard H. Gomer, Howard Hughes Medical Institute and Department of Biochemistry and Cell Biology, MS-140, Rice University, 6100 S. Main St., Houston, TX 77005.1892.
439
ANALOGOUS SYSTEMS
440
Spores
1111,
f
1
Aggregation
Figure 1 The D. discoideum life cycle. Single-celled amoebae feed on soil bacteria. When the food source is exhausted and the cells begin to starve, they initiate CAMP signaling, leading to aggregation and mound formation. Precursors of the two main cell types begin to differentiate in the mound, forming prestalk cells (black) and prespore cells (spotted). The mound follows a predefined morphogenic program, producing a multicellular slug which migrates in response to light and heat. The program continues leading to culmination of the slug into a mature fruiting body consisting of a stalk (black) and a spore mass (spotted). Spores are released and when food is present, germinate into vegetative amoebae.
precursors and correct this ratio if necessary. Dictyostelium also has a simple mechanism that regulates the size of the fruiting body. The basic physics of multicellular morphogenesis is still very poorly understood, and if one thinks of the fruiting body as a sphere on a cylinder, this is a very simple system to investigate the morphogenesis of such structures. Several mathematical models of development, especially the aggregation of starving cells, are helping to guide experiments and verify hypotheses (Dallon and Othmer, 1997; Dormann et al., 1998; Hofer and Maini, 1997; Maree and Hogeweg, 2001; Martiel and Goldbeter, 1985; Pate and Odell, 1981; Pate and Othmer, 1986; Roisin-Bouffay et al., 2000). A very new and interesting field of biology involves the molecular
basis of altruism. The Dictyostelium cells that become stalk cells sacrifice themselves and die to help the spore cells get dispersed, and studies of how different strains either cooperate or cheat to become spore cells is helping to open this field. Finally, there are several human diseases or therapies for diseases that can be usefully studied using the simplicity of Dictyostelium to uncover molecular mechanisms.
DICTYOSTELIUM IS EASY TO WORK WITH The simplicity of Dictyostelium development is complemented by the ease with which one can work with the system (Sussman, 1987). Some strains of Dictyostelium
25.
A EUKARYOTIC NEIGHBOR: D. DISCOIDEUM
do not require bacteria as a food source. These strains, referred to as axenic, can be grown at room temperature in inexpensive serum-free broth similar to media used for growing bacteria. The cell division time is approximately 8 h, and cells can be grown to 1010 cells/liter (approximately 7 g [wet weight] of cells/liter). The cells normally grow and develop at room temperature, which makes growth and developmental studies accessible to high schools or researchers with limited budgets. In addition, the growth at room temperature makes time-lapse videomicroscopy studies on live cells quite easy. Strains can be preserved by lyophilization of spores, desiccation of spores on silica, or freezing vegetative cells in media containing glycerol or dimethyl sulfoxide (Sussman, 1987). Performing biochemical experiments with Dictyostelium is relatively straightforward. Cells can be grown in shaker flasks or fermentors in large quantities easily and inexpensively, thereby enabling biochemical studies. The cells are -10 p,m in diameter and can be lysed by passage through a 5-pm-pore-size syringe filter with only a minimal amount of organelle disruption. This rapid lysis facilitates experiments which examine processes with fast kinetics, such as signal transduction biochemistry, where one can add a ligand to the cells and watch the kinetics of a cytosolic protein translocating to the plasma membrane and then returning to the cytosol (Lilly and Devreotes, 1995). A wide variety of genetic tools are available for studying Dictyostelium. Genetic approaches, facilitated by the fact that Dictyostelium colonies can be grown from single cells or spores, have been used for a large number of studies, and there exists a wide variety of mutants (Kuspa et al., 1995; Loomis, 1987). Both extrachromosoma1 and integrating transformation vectors have been developed for Dictyostelium (Knecht et al., 1990; Nellen et al., 1984). These have been used for studies on cisacting regulatory sequences, as well as the expression of foreign proteins, site-directed mutations of DictyosteZium proteins, or green fluorescent protein fusions in cells (Cohen et al., 1986; Datta et al., 1986; Esch et al., 1992; Fields et al., 2002; Pears and Williams, 1987; Reymond et al., 1986). Transformation of cells with antisense or homologous recombination constructs can be used to abolish the expression of selected genes or families of genes (Crowley et al., 1985; De Lozanne and Spudich, 1987; Gomer, 1998). Mutagenesis screens can be done with either random insertion of a linearized plasmid containing a selectable marker (restriction enzyme-mediated integration [REMI]) or shotgun antisense (Kuspa and Loomis, 1992; Spann et al., 1996). Genetic networks can be elucidated by the use of REMI to isolate secondsite suppressors (Shaulsky et al., 1996). A stock center
441 provides a valuable repository of strains, transformants, and transformation vectors. Once a gene is identified by PCR of the cDNA in the shotgun antisense construct or by inverse PCR to sequence some of the DNA flanking a REMI-tagged gene, additional sequence can then be obtained from a Japanese project in which -9,000 different cDNAs have been sequenced (there are -12,500 genes in Dictyostelium) or from the recently completed project to sequence, assemble, and annotate the -34 Mb genome (Eichinger et al., 2005). An online bioinformatics resource is available at http://dictybase.org/. As with other model systems, cDNA microarrays can be used for genome-wide expression analysis (Iranfar et al., 2003; Van Driessche et al., 2002). Taken together, these characteristics make Dictyostelium an attractive model system.
INITIATING THE DEVELOPMENTAL PROGRAM: SENSING STARVATION Vegetative Dictyostelium cells consume bacteria and divide by binary fission. However, when the cells sense they are starving, they stop dividing. Leaving the vegetative stage of their life cycle and entering into development is an important decision for Dictyostelium. Early cessation of growth would give other cells extra time to divide and could lead to being outcompeted. In addition, initiating development long after starvation had begun could make it difficult for the cells to produce the required proteins under the limited food and energy supplies. Therefore, Dictyostelium cells must be able to sense the approach of starvation and initiate development at the appropriate time. The cells accomplish this by secreting a glycoprotein called prestarvation factor (PSF). Exponentially growing cells continuously synthesize and secrete PSF, which accumulates outside the cells in proportion to cell density (Burdine and Clarke, 1995; Clarke and Gomer, 1995; Rathi et al., 1991). PSF regulates the expression of genes involved in aggregation, such as discoidin-1, CAMP receptor cAR1, cell adhesion protein gp24, and lysosomal protein a-mannosidase. PSF does not reach high enough concentrations to activate these genes until cells are four generations from exiting exponential growth phase. Thus, PSF is able to regulate transcription based upon cell density. In addition, PSF activity is inhibited by the presence of bacteria, allowing its activity to be modulated not only by cell density but also by available food supplies. Therefore, PSF ensures that the cell is physiologically prepared to switch from vegetative growth to development by monitoring the ratio of cell density to food density in growing cells.
442 Once PSF levels determine that the conditions are appropriate to initiate development, the cell cycle must be arrested. While the cell cycle of Dictyostelium is controlled by many of the same proteins that regulate other eukaryotic cell cycles, much remains to be understood about how starvation causes the cell cycle to stop. One of the proteins involved in starvation-induced cell cycle arrest is the protein kinase YakA (Souza et al., 1998; Taminato et al., 2002). YakA belongs to a family of protein kinases found in yeast, plants, mammals, worms, and mice. In all cases, these proteins are involved in controlling the cell cycle. Dictyostelium cells overexpressing YakA arrest prematurely, while cells lacking YakA cycle faster than normal. In addition to arresting the cell cycle, YakA also plays a crucial role in shifting gene transcription from a vegetative program to a developmental one. To enter into development, cells must halt production of proteins required for vegetative growth and initiate production of proteins required for aggregation and development. PSF levels are monitored by YakA, with high levels of PSF leading to activated YakA. Active YakA is necessary to down regulate growth-specific genes and up regulate developmental genes. YakA accomplishes this by increasing protein kinase A (PKA) activity (Taminato et al., 2002). As will be seen later, PKA plays a central role in several developmental processes in Dictyostelium. Thus, the approach of starvation is sensed by secreted PSF, while the disruption of the cell cycle is controlled by YakA, which itself is responsive to PSF levels.
COORDINATING DEVELOPMENT: CELL DENSITY (QUORUM)SENSING There can be different microenvironments in the 1- to 10-mm-diameter zone of cells that generates an initial aggregate of Dictyostelium cells. One can envision cells in one microenvironment starving earlier than cells in an adjacent microenvironment. Without coordination, small cohorts of cells that happened to starve at the same time would form fruiting bodies that were too small and thus too close to the ground for efficient spore dispersal. To have as many cells as possible aggregate and form a tall fruiting body, the amoebae need to coordinate their entry into the developmental pathway. This coordination appears to be mediated by a mechanism that senses the density of starved cells and allows aggregation to occur only when there is a sufficiently high density of starved cells to form a properly sized fruiting body. Insight into how such a quorum-sensing mechanism functions was gained when it was observed that D. discoideum cells starved at low cell densities differentiate only in buffer previously conditioned by a high density of starved
ANALOGOUS SYSTEMS cells (Grabel and Loomis, 1978; Kay and Jermyn, 1983; Mehdy and Firtel, 1985). Thus, D. discoideum cells are able to sense the number of starving cells present and are able to make a developmental decision based on this information. Interestingly, this ability is not dependent on cell-cell contact but appears to involve a secreted, soluble factor (Mehdy and Firtel, 1985). This factor, CMF, was purified, and the gene coding it was cloned (Brazill et al., 1998; Deery et al., 2002; Jain et al., 1992; Van Haastert et al., 1996; Yuen et al., 1995). Cells lacking CMF are unable to aggregate and develop unless exogenous CMF is added. CMF is produced in growing cells but is not secreted until the cells begin to starve. Once a cell begins to starve, it secretes CMF at a constant rate, making CMF an ideal molecule for density/quorum sensing. As the number of starving cells in a given area increases, the level of CMF in that area rises. At a certain time, there are enough starving cells to form a functional fruiting body, and at this time the CMF levels are high enough to allow development to proceed. CMF is able to exert this control by regulating the initial step of development, aggregation. CMF accomplishes this by controlling cAMP signal transduction (Jain et al., 1992; Yuen et al., 1995) and therefore chemotaxis towards the aggregation center. CMF regulates cAMP signal transduction by activating its own signal transduction pathway involving a seven-transmembrane receptor which activities a G protein, phospholipase C, and phospholipase D (Brazill et al., 1998; Chen et al., 2005; Deery et al., 2002; Deery and Gomer, 1999).Thus, until there is a high density of starving cells, as indicated by a high concentration of CMF, the cells will not respond to pulses of CAMP.
DIFFERENTIATION AND SYMMETRYBREAKING: A MUSICAL CHAIRS MECHANISM TELLS CELLS WHETHER TO BECOME PRESPORE OR PRESTALK When development is first initiated, all of the cells are undifferentiated. Yet by the slug stage the precursors to the stalk and spore cells, called prestalk and prespore cells, respectively, can be found. Thus, symmetry has been broken and differentiation has begun. However, initial choice of cell type is determined by the phase of the cell cycle that a cell happens to be in at the time of starvation (Fig. 2). Like some fungi, the Dictyostelium cell cycle has no observable GI phase; cells enter S phase immediately after mitosis; they undergo cytokinesis during S phase and then spend most of the cell cycle in G, (Weijer et al., 1984). In a normal population, cells are randomly distributed throughout the cell cycle. Prestalk cells (identified using
25. A EUKARYOTIC NEIGHBOR: D. DISCOIDEUM
443
Figure 2 Initial cell type choice is dependent on the cell cycle phase at starvation. Prestalk cells, cells with a predisposition to become stalk cells, are derived from cells that were in S phase and early G, phase at the time of starvation. Prespore cells, cells with a predisposition to become spore cells, are derived from cells that were in late G, or M phase at the time of starvation. Like some fungi, Dictyostelium has a very small GI phase.
antibodies against an antigen called CP2) are derived from cells starved in S and early G,, and prespore cells are derived from cells starved in late G, (Araki et al., 1994; Azhar et al., 2001; Gomer and Firtel, 1987a; McDonald and Durston, 1984; Weijer et al., 1984). Cells of a third type, null cells, are derived from the sister cells of the prespore and prestalk cells (Clay et al., 1995; Gomer and Firtel, 198713; Wood et al., 1996). This mechanism regulates only initial differentiation. The eventual fate of the cell is still plastic, and a variety of factors such as adenosine, ammonia, a chlorinated hydrocarbon called DIF, and oxygen can change the final fate (Brookman et al., 1987; Gross et al., 1983; Kay et al., 1989; Kwong and Weeks, 1989; Schaap and Wang, 1986; Sternfeld, 1988; Williams et al., 1987; Xie et al., 1991).Thus, initial predisposition towards a specific cell fate is decided at the initiation of starvation on an individual, cell-by-cell basis.
INITIATING MULTICELLULARITY AGGREGATION The transition to multicellularity begins when individual starving cells aggregate. Upon starvation, individual cells use relayed pulses of cAMP as a chemoattractant to aggregate. Aggregation occurs between 5 and 10 h after
starvation. It is still unclear exactly how the pulses begin and how an aggregation center is chosen. Starved cells respond to a pulse of cAMP in three ways. First, the cells move towards the source of CAMP. This is the driving force behind aggregation. Second, the cells release a burst of cAMP themselves. This relays the aggregation signal to cells that are further from the aggregation center. Third, the cells activate or deactivate expression of specific classes of genes (Mann and Firtel, 1987). The incoming cAMP pulse is detected by cell surface cAMP receptors (Fig. 3 ) . D. discoideum cells express several cAMP receptor genes, all of which are developmentally regulated (Saxeet al., 1991). The cAMP receptor that mediates chemotaxis, cAR1, contains seven transmembrane domains, typical of G-protein-coupled receptors (Gilman, 1987; Firtel, 1996). The binding of cAMP to the cAMP receptors causes the receptors to activate G proteins (Theibert and Devreotes, 1986; Van Haastert, 1984; Van Haastert et al., 1987). At least eight different G protein a subunit genes are expressed during development (Wu and Devreotes, 1991), and several more can be found in the genome. However, Ga2 is the subunit that mediates many of the downstream responses associated with the activation of cAR1, including chemotaxis and gene expression (Kumagai et al., 1991; Kumagai et al., 1989).
ANALOGOUS SYSTEMS
444
c~~~ Relay Signal to Other Cells
cAMP Receptor
3
I
t Guanylyl I Phosphorylation
Chemotaxis Figure 3 cAMP signaling during chemotaxis. cAMP secreted from starving cells binds to and activates the cAMP receptor. The receptor in turn activates its associated G protein, Ga2py, by catalyzing the exchange of GTP for GDP. The G protein separates, and the Py subunit, along with CRAC (cytosolic regulator of adenylyl cyclase) activates adenylyl cyclase. This leads to the secretion of cAMP and thus the relay of the aggregation signal to other cells. The activation of the G protein also leads to the initiation of chemotaxis towards the source of cAMP and thus movement towards the center of aggregation.
Activation of CAR1 and its associated G protein leads to transient activation of guanylyl cyclase and adenylyl cyclase. In addition, there is a transient uptake of extracellular Ca2+(Kuwayama and van Haastert, 1998; Milne and Devreotes, 1993). Guanylyl cyclase is activated by cAMP through the CAR1 receptor and its associated G protein a subunit Ga2. This causes a transient elevation in intracellular cGMP (Newell, 1995; Van Haastert and Kuwayama, 1997). Activation of guanylyl cyclase by the cAMP receptor appears to play a major role in general chemotaxis in Dictyostelium cells (Kuwayama et al., 1993). The activation of adenylyl cyclase occurs over several minutes and is also associated with CAR1 (Devreotes, 1989). The Py heterodimer moiety of an activated G protein activates adenylyl cyclase (Devreotes, 1989). In addition, adenylyl cyclase activation requires an 88-kDa cytosolic protein (Theibert and Devreotes, 1986),which has been designated cytoplasmic regulator of adenylyl cyclase (CRAC) (Insall et al., 1994).
CHEMOTACTIC SIGNALS REGULATE A N ACTIN/MYOSIN-BASED CYTOSKELETON T O REGULATE MOTILITY Motility in D. discoideum is mediated by actin-driven protrusion of the leading edge and cell body translocation driven at least in part by myosin II-mediated contraction of the actin cortex in areas away from the leading edge (Chung et al., 2001; Elson et al., 1999; Manstein, 1993). Cells which are morphologically polarized (for instance, due to exposure to a cAMP gradient) have a clear cytoskeletal polarity, with actinfilled protrusions at the front and myosin I1 accumulation at the rear (Condeelis, 1992; Dharmawardhane et al., 1989; McRobbie and Newell, 1983; Noegel and Luna, 1995). Myosin I1 heavy-chain phosphorylation and assembly are regulated by myosin I1 heavy-chain kinases (de la Roche and Cote, 2001; Liang et al., 2002). Myosin heavy-chain kinase is recruited to these new protrusions and phosphorylates myosin I1 in order
25. A EUKARYOTIC NEIGHBOR: D. DISCOIDEUM to prevent minifilament assembly (Berlot et al., 1985; Luck-Vielmetter et al., 1990; Sabry et al., 1997; Steimle et al., 2001). A kinase called PAKa regulates myosin I1 assembly, cell polarity, and cell motility via the myosin I1 heavy-chain kinases (Chung and Firtel, 1999; Chung et al., 2001). PAKa is in turn phosphorylated and regulated by Akt/PKB (Chung et al., 2001). Akt/PKB binds to phosphatidylinositol3,4,5-trisphosphate(PIP,) via a pleckstrin homology (PH)domain (Tanaka et al., 1999). The pulses of cAMP that mediate chemotaxis activate PI3 kinases that cause a transient appearance of PIP, and phosphatidylinositol( 3,4)P, at the leading edge of cells, which causes Akt/PKB to in turn transiently bind to the leading edge of the cell and become activated (Funamoto et al., 2001; Huang et al., 2003; Meili et al., 1999). Thus, the presence of cAMP causes the activation of a localized signaling pathway, culminating in the phosphorylation and deactivation of myosin at the leading edge of the cell. Another protein that binds to PIP, at the leading edge of cells is PhdA (Funamoto et al., 2001). PhdA regulates the assembly of actin at the leading edge of cells (Funamoto et al., 2001), suggesting that activation of PI3 kinase at the leading edge of cells causes both the disassembly of myosin and the appearance of an actin pseudopod at the leading edge. Therefore, aggregation occurs when the CAR1 cAMP receptor is activated by the pulses of CAMP.This activates the heterotrimeric G protein whose 01 subunit is Go12 (Meili et al., 1999). This in turn causes a transient activation of PI3 kinase (Huang et al., 2003), leading to the formation of a pseudopod via actin reorganization and myosin disassembly at the leading edge of the cell.
445 smlAas (Spann et al., 1996). Disruption of the smlA gene by homologous recombination produced mutant cells lacking smlA, which had the same phenotype as smlAas cells. The exudate from starving cells lacking
A
B
C
A SECRETED MULTIPOLYPEPTIDE FACTOR REGULATES DICTYOSTELIUM GROUP SIZE Aggregating cells in an aggregation field form radial streams flowing toward a common center. Regulation of the size of this field is important since there is a limit to the strength of the stalk, and too large a field will result in the formation of a fruiting body that will collapse under its own weight. Therefore, Dictyostelium has evolved a mechanism that senses the number of cells in a stream and causes the stream to break up into groups if there are too many cells in it (Shaffer, 1957) (Fig. 4), thus allowing the formation of a functional fruiting body. To elucidate the mechanism causing the stream to break up into groups, shotgun antisense was used to mutagenize Dictyostelium cells in a screen for abnormally sized fruiting bodies. A transformant that formed large numbers of very small fruiting bodies due to excessive stream breakup was identified and designated
t high adhesion
Figure 4 Fruiting body size regulation. Panels A and B show a field of aggregating cells at approximately 7 h (A) and 8 h (B) after starvation. The cells were at a high density, and as a consequence the streams broke into groups. Panel C shows a computer simulation of the breakup. If the cells in a stream have a high cell-cell adhesion (and/or a low random cell motility), the stream will tighten (downward arrow). If the cells in a stream have a low cell-cell adhesion (and/or a high random cell motility), the stream will dissipate and breaks will form in the stream (right arrow). The cells in the break regions chemotax towards the cells in the nearby groups, further accentuating the breaks. If cell-cell adhesion then increases (and/or random cell motility decreases), the groups coalesce. Dictyostelium uses a secreted factor to modulate adhesion and motility and thus the extent of stream breakup and group size. Bar (panel B), 1 mm.
446
smZA causes wild-type cells to form small fruiting bodies (Brock et al., 1996). The factor oversecreted by the cells lacking smlA (and secreted by wild-type cells, albeit at a lower level) was partially purified and found to be an -450-kDa complex of polypeptides now called counting factor (CF) (Brock and Gomer, 1999). Disruption of countin, a gene encoding one of the components of CF, results in streams not breaking up. These streams then coalesce into one huge group that forms a huge fruiting body, which then collapses, spilling the spores onto the ground. The same phenotype is seen in colonies of cells lacking CF45-1 or CF50, other components of CF (Brock et al., 2002,2003). The effect of CF on group size suggests that CF is part of a negative-feedback loop, with a high concentration of CF inducing stream breakup. Thus, aggregation field size is monitored and regulated by secreted CF. As more cells enter the field, CF levels rise until enough cells are present to form a properly sized fruiting body. At this point, CF causes the streams of cells to break up, ensuring that each aggregation field does not contain too many cells. To predict possible parameters that CF could control, which could regulate the morphogenesis of a stream of cells into a series of groups of cells, a computer simulation was used to model stream behavior (RoisinBouffay et al., 2000). The simulations predicted that if the cell-cell adhesion was low, the stream would start to dissipate. If the adhesion then increased, the dissipated stream would coalesce into groups (Fig. 4). In accordance with the computer predictions, overexpressing an adhesion protein during Dictyostelium development causes the formation of unbroken streams and large aggregates, while blocking cell surface adhesion proteins with monoclonal antibodies causes the formation of broken streams and many small aggregates (Kamboj et a]., 1990; Roisin-Bouffay et al., 2000; Siu and Kamboj, 1990). As predicted by these observations and the simulations, CF inhibits cell-cell adhesion (Roisin-Bouffay et al., 2000) and thus decreases group size. In addition to decreased adhesion, the simulations also predicted that increasing cell motility would increase stream dissipation and hence stream breakup. In agreement with the computer models, decreasing motility increases group size (Tang et al., 2002), and CF, which decreases group size, increases motility (Tang et al., 2002). CF appears to regulate adhesion and motility and thus group size in part through a signal transduction pathway that involves glucose or a glucose metabolite (Gao et al., 2004; Jang et al., 2002; Jang and Gomer, 2005). This supports and explains previous observations that growing cells in the presence of glucose causes the formation of large groups (Garrod and Ashworth, 1972). Therefore, CF controls
ANALOGOUS SYSTEMS group size, and consequently fruiting body size, by modulating motility and cell-cell adhesion.
MORPHOGENESIS OF THE GROUPS OF CELLS Once chemotaxis has begun and group size is determined, a mound is formed. The cell-autonomous musical chairs cell-type choice mechanism causes prespore and prestalk cells to appear at random places within the mound. From this random distribution of prestalk and prespore cells emerges a tipped mound where prestalk cells are found at the apical tip of the mound and prespore cells make up the base. This happens through a process of cell sorting. Cells in the mound are highly motile and are constantly moving in a circular pattern within the mound (Clow et al., 2000; Kellerman and McNally, 1999; Weijer, 1999). As the cells move, differential adhesion and motility rates lead to cell sorting with the prestalk cells localizing to the apical tip of the mound. The tip of the mound elongates, forming the first finger. Then, under certain conditions, the first finger will fall over and become a migrating slug or pseudoplasmodium. These slugs migrate towards light and heat, presumably to reach the surface of the soil, where spore dispersal will be more efficient. By the time the slug has formed, the cells have differentiated even further. The prestalk cells have subdivided into several groups (Jermyn et al., 1989). PstA cells are found at the very anterior tip of the slug. Inside the PstA group of cells is a small core of PstAB cells. Slightly posterior to the PstA cells are the PstO cells. These three cell types inhabit the anterior fourth of the slug. The remaining three-fourths of the slug consists of a preponderance of prespore cells with a smattering of anterior-like cells (ALC), followed by rearguard cells. Both ALC and rearguard cells are of prestalk origin (Blaschke et al., 1986; Casademunt et al., 2002; Devine and Loomis, 1985; Sternfeld and David, 1982). As the slug migrates, the rearguard cells are lost. To maintain the proper ration of prestalk to prespore cells, prespore cells convert to ALC, which convert to PstO and then PstA cells. This ability to interconvert is not relegated to this gradual process but can be quite rapid. If the tip of a slug is removed, leaving only the prespore region, this process will be used to create a whole new tip (Bonner, 1952). A fully functional, although smaller, slug will be re-formed. Thus, there is constant monitoring and adjustment of cell types to actively maintain proper patterning. One of the major morphogens involved in this process is DIF-1, a small chlorinated hexaphenone (Fukuzawa et al., 2003; Kay and Jermyn, 1983). DIF-1 induces the expression of prestalk genes
25. A EUKARYOTIC NEIGHBOR: D. DISCOIDEUM and represses the expression of prespore genes. DIF-1 is found mainly in the posterior of the slug. It still remains unclear as to the molecular mechanisms by which patterning in the slug is actively maintained, but chemotaxis to cAMP and differential adhesion, the same processes seen in the mound, appear to be involved. Culmination of the slug into a fruiting body is controlled by a variety of environmental factors including light, temperature, and ammonia concentration (Harwood et al., 1992; Jermyn and Williams, 1991). During culmination, the slug stops moving and begins to stand up on its tail. The rearguard cells and ALCs form a base that will eventually become the basal disk. The stalk tube begins to form around the PstAB cells. The stalk tube then elongates down from the anterior tip of the culminant to the base as PstA cells move into the tube. Eventually the stalk tube reaches the basal disk. The cells that make up the stalk develop a cellulose-based cell wall, become highly vacuolated and die as stalk cells. While this is occurring, the spore mass is pulled up off the ground. As the spores move up the stalk, they are encapsulated. Finally, the upper and lower cups of the sorus are formed by PstAB cells. The entire process of final differentiation into spore and stalk cells is highly coordinated and requires cAMP and the activation of PKA. Two secreted spore differentiation factors (SDF-1 and SDF-2) are also involved (Anjard et al., 1997; Wang et al., 1999). Little is known about the signaling done by SDF-1; however, much insight has been gained about SDF-2. SDF-2 activates a two-component system. SDF-2 activates the histidine kinase receptor DhkA. DhkA phosphorylates and inactivates RegA, a response regulator. RegA is a phosphodiesterase, which destroys CAMP.Its inactivation by DhkA allows cAMP levels to rise and thus causes the activation of PKA. Specifics about the components of terminal differentiation downstream of PKA remain unclear. However, a transcription factor with homology to mammalian Serum Response Factor, SrfA, is required for terminal spore differentiation. While it is still unknown how this system regulates stalk development, it is interesting that SDF-2 sets up a positive-feedback loop in stalk cells that causes more SDF-2 to be released.
DICTYOSTELIUM IS A N EXCELLENT SYSTEM TO STUDY SOME HUMAN DISEASES In addition to being an excellent organism to study chemotaxis, development, and differentiation, Dictyostelium is also used to understand the molecular basis of disease. One of the diseases under investigation is
44 7
Legionnaires’ disease, which is caused by an infection by Legionella pneumophila bacteria. In humans, inhaled L. pneurnophila is phagocytosed by macrophages present in the lungs. This process destroys most bacteria; however, instead of being digested, L. pneumophila replicates within the macrophages. Ingested L. pneumophila organisms traffic to and grow within an unusual phagosomal compartment. Transport to this compartment in phagocytic cells is required for bacterial growth, as mutants defective in this process are not virulent. However, it is unclear what specific proteins in the macrophage interact with the bacteria to create this replicative compartment, thus preventing digestion of the bacteria and allowing for its growth. As with macrophages, L. pneumophila is phagocytosed by Dictyostelium and is able to replicate in phagosomal compartments (Otto et al., 2004; Solomon et al. 2000). Studies with Dictyostelium have identified a number of cytoskeletal and vesicle fusion proteins that affect growth of L. pneumophila. Success with L. pneumophila has led to the use of Dictyostelium to study infection by Pseudomonas aeruginosa, Cryptococcus neoformans, and Mycobacterium spp. Dictyostelium is also being used to understand the genetic basis of tumor resistance to the chemotherapeutic drug, cisplatin (Alexander et al., 2003). Cisplatin is used to treat numerous cancers including small-cell and non-small-cell lung cancer, non-Hodgkin’s lymphoma, testicular cancer, ovarian cancer, head and neck cancer, esophageal cancer, and bladder cancer. Unfortunately, acquired resistance to the drug reduces the efficacy of therapy in these cancers. In addition innate resistance prevents the use of the drug with other cancers. A genetic screen for cisplatin resistance in Dictyostelium has led to the identification of a number of genes believed to be involved in cisplatin action, none of which had been previously associated with cisplatin resistance. For instance, two genes that code for sphingosine kinases, sgkA and sgkB, mediate cisplatin resistance in Dictyostelium. Overexpression of the genes makes cells resistant to the drug, while disruption of the genes leads to hypersensitivity to the drug. Interestingly, sgkA and sgkB have homologues in humans, suggesting that regulation of their human cognates may increase the effectiveness of cisplatin treatment of tumors. Lithium and valproic acid are used to treat bipolar disorder, but their mechanisms of action are poorly understood. Use of Dictyostelium has led to the understanding that these drugs may work though a common pathway, inositol metabolism (Williams et al., 2002). Development in Dictyostelium is sensitive to both drugs. A genetic screen for mutants resistant to lithium identified a gene coding for a prolyl oligopeptidase. Loss of the
448
gene causes an increase in inositol triphosphate levels and confers resistance to both lithium and valproic acid. This led to the finding that in Dictyostelium lithium and valproic acid both cause a decrease in inositol triphosphate levels. In accordance with the findings in Dictyostelium, the plasma concentrations of prolyl oligopeptidase are elevated in patients suffering from mania and decreased in patients with depression. The studies in Dictyostelium have helped to clarify the molecular basis for bipolar disorder as well as delineate the method of action of drugs used to treat it. In addition, they have identified an exciting new direction for developing diagnostics and treatments for certain mental disorders.
MOLECULAR SOCIOLOGY The fact that 20% of the cells that form the fruiting body kill themselves to form stalk cells demonstrates that Dictyostelium cells exhibit altruism. These cells are sacrificing themselves so that the remaining 80% can create the next generation. Normally, such altruism is seen in metazoans such as social wasps and ants, where only a subpopulation of individuals is reproductively active. The genetic tools available for Dictyostelium research can be used to elucidate molecular mechanisms underlying altruism. In a mixed population of cells, there is an evolutionary advantage to becoming a spore cell. Strains that preferentially become spores in a mixture of different strains will have a greater chance to pass their genes on to the next generation. These strains are called “cheaters” because they unfairly become spores at a higher rate (Dao et al., 2000). In the wild, genetically different strains of Dictyosteliurn live in close enough proximity to each other to form mixed fruiting bodies. In these cases, it is quite possible for cheaters to exist and thrive. Indeed, in the wild, several cheater strains have been identified. Genetic dissection of these cheaters is currently being pursued. Thus, Dictyostelium lends itself to be conducive to studying not only developmental mechanisms but also social behavior.
COMPARISONS TO MYXOBACTERIA D. discoideum and myxobacteria share a number of traits. They both exist as individual cells. Upon starvation, both aggregate to form a multicellular structure, the fruiting body. During formation of the fruiting body, cells differentiate to become heat- and desiccationresistant spores. This differentiation is dependent on cellular motion within the forming aggregate, cell density, and cell-cell signaling (Julien at al., 2000). When food becomes available, these spores germinate, releasing
ANALOGOUS SYSTEMS a new generation of individuals. On the surface, their lifestyles seem identical, yet upon closer inspection some differences become apparent. Dictyostelium cells are eukaryotes and as such display a number of structural and physiological differences from myxobacteria. Dictyostelium uses cAMP as a chemoattractant while Myxococcus xanthus, for example, uses phosphatidyl ethanolamine. Some of the signal transduction mechanisms are also different. Dictyostelium uses both G-protein-coupled receptors and two-component receptors, while myxobacteria use two-component systems. More importantly, the stalk of a Dictyostelium fruiting body is made of terminally differentiated, dead cells, while the stalk of a myxobacterial fruiting body is made of secreted compounds. Therefore, Dictyostelium cells display a form of altruism in the creation of a fruiting body whereas no cells are sacrificed to make a myxobacterial fruiting body. While it has been postulated that a large portion of the developing myxobacteria undergo autolysis in order to aid in the development of the remaining cells, no evidence has been presented that demonstrates an obligatory role for such a putative sacrifice (Wireman and Dworkin, 1975; Rosenbluh et al., 1989). Thus, while both organisms evolved the same basic response to nutrient deprivation, they have unique methods for eliciting that response. A comparison of their biochemical pathways can lend insight into how different organisms can use different mechanisms to achieve the same goals. W e thank Yitai Tang for the photograph used in Fig. 1. This work was supported by grant number C-1555 from the Robert A. Welch Foundation.
References Alexander, H., A. N. Vomund, and S. Alexander. 2003. Viability assay for Dictyostelium for use in drug studies. BioTechniques 35:464-470. Anjard, C., M. van Bemmelen, M. Veron, and C . D. Reymond. 1997. A new spore differentiation factor (SDF) secreted by Dictyostelium cells is phosphorylated by the cAMP dependent protein kinase. Differentiation 62:4349. Araki, T., H. Nakao, I. Takeuchi, and Y. Maeda. 1994. Cellcycle-dependent sorting in the development of Dictyostelium cells. Dev. Biol. 162:221-228. Aubry, L., and R. A. Firtel. 1999. Integration of signaling networks that regulate Dictyostelium differentiation. Annu. Rev. Cell Dev. Biol. 15:469-517. Azhar, M., P. Kennady, G. Pande, M. Espiritu, W. Holloman, D. Brazill, R. Gomer, and V. Nanjundiah. 2001. Cell cycle phase, cellular Ca2+and development in Dictyostelium discoideum. Int. J. Dev. Biol. 45:405-414. Berlot, C . H., J. A. Spudich, and P. N. Devreotes. 1985. Chemoattractant-elicited increases in myosin phosphorylation in Dictyostelium. Cell 43:307-314.
25. A EUKARYOTIC NEIGHBOR: D. DISCOIDEUM Blaschke, A., C. Weijer, and H. MacWilliams. 1986. Dictyostelium discoideum: cell-type proportioning, cell-differentiation preference, cell fate, and the behavior of anterior-like cells in Hsl/Hs2 and G+/G- mixtures. Differentiation 32:1-9. Bonner, J. T. 1952. The pattern of differentiation in ameboid slime molds. Am. Nut. 86:79-89. Brazill, D. T., D. F. Lindsey, J. D. Bishop, and R. H. Gomer. 1998. Cell density sensing mediated by a G proteincoupled receptor activating phospholipase C. ]. Biol. Chem. 2739161-8 168. Brefeld, 0. 1869. Dictyostelium mucuroides. Ein neuer Organismus aus der Verwandtschaft der Myxomyceten. Abhandlungen der Senckenbergischen Naturforschenden Gesellschaft 7: 85-1 07. Brock, D. A., F. Buczynski, T. P. Spann, S. A. Wood, J. Cardelli, and R. H. Gomer. 1996. A Dictyostelium mutant with defective aggregate size determination. Development 122:25692578. Brock, D. A., and R. H. Gomer. 1999. A cell-counting factor regulating structure size in Dictyostelium. Genes Dev. 13~1960-1969. Brock, D. A., R. D. Hatton, D.-V. Giurgiutiu, B. Scott, R. Ammann, and R. H. Gomer. 2002. The different components of a multisubunit cell number-counting factor have both unique and overlapping functions. Development 129:3657-3668. Brock, D. A., R. D. Hatton, D.-V. Giurgiutiu, B. Scott, W. Jang, R. Ammann, and R. H. Gomer. 2003. CF45-1, a secreted protein which participates in group size regulation in Dictyostelium. Eukaryot. Cell 2:788-797. Brookman, J. J., K. A. Jermyn, and R. R. Kay. 1987. Nature and distribution of the morphogen DIF in the Dictyostelium slug. Development 100:119-124. Burdine, V., and M. Clarke. 1995. Genetic and physiologic modulation of the prestarvation response in Dictyostelium discoideum. Mol. Biol. Cell 6:3 11-325. Casademunt, E., T. R. Varney, J. Dolman, C. Petty, and D. D. Blumberg. 2002. A gene encoding a novel anti-adhesive protein is expressed in growing cells and restricted to anterior-like cells during development of Dictyostelium. Differentiation 70~23-35. Cavender, J. C., and K. B. Raper. 1968. The occurrence and distribution of Acrasiae in forests of subtropical and tropical America. Am. J. Bot. 55:504-513. Chen, Y., V. Rodrick, Y. Yan, and D. Brazill. 2005. PldB, a putative phospholipase D homologue in Dictyostelium discoideum mediates quorum sensing during development. Eukaryot. Cell 4:694-702. Chung, C. Y., and R. A. Firtel. 1999. PAKa, a putative PAK family member, is required for cytokinesis and the regulation of the cytoskeleton in Dictyostelium discoideum cells during chemotaxis. J. Cell Biol. 147559-576. Chung, C. Y., S. Funamoto, and R. A. Firtel. 2001. Signaling pathways controlling cell polarity and chemotaxis. Trends Biochem. Sci. 26557-566. Clarke, M., and R. H. Gomer. 1995. PSF and CMF, autocrine factors that regulate gene expression during growth and early development of Dictyostelium. Experientia 5'1:1124-1134.
449 Clay, J. L., R. A. Ammann, and R. H. Gomer. 1995. Initial cell type choice in a simple eukaryote: cell-autonomous or morphogen-gradient dependent? Dev. Biol. 172:665-674. Clow, P. A., T. Chen, R. L. Chisholm, and J. G. McNally. 2000. Three-dimensional in vivo analysis of Dictyostelium mounds reveals directional sorting of prestalk cells and defines a role for the myosin I1 regulatory light chain in prestalk cell sorting and tip protrusion. Development 127:2715-2728. Cohen, S. M., D. Knecht, H. F. Lodish, and W. F. Loomis. 1986. DNA sequences required for expression of a Dictyostelium actin gene. EMBO J. 5:3361-3366. Condeelis, J. 1992. Are all pseudopods created equal? Cell Motil. Cytoskeleton 22:l-6. Crowley, T. E., W. Nellen, R. H. Gomer, and R. A. Firtel. 1985. Phenocopy of discoidin I-minus mutants by antisense transformation in Dictyostelium. Cell 43:633-641. Dallon, J. C., and H. G. Othmer. 1997. A discrete cell model with adaptive signalling for aggregation of Dictyostelium discoideum. Philos. Trans. R. SOC. Lond. B 352:391-417. Dao, D. N., R. H. Kessin, and H. L. Ennis. 2000. Developmental cheating and the evolutionry biology of Dictyostelium and Myxococcus. Microbiology 146:1505-1512. Datta, S., R. H. Gomer, and R. A. Firtel. 1986. Spatial and temporal regulation of a foreign gene by a prestalk-specific promoter in transformed Dictyostelium discoideum. Mol. Cell. Biol. 6:811-820. Deery, W. J., T. Gao, R. Ammann, and R. H. Gomer. 2002. A single cell-density sensing factor stimulates distinct signal transduction pathways through two different receptors. J. Biol. Chem. 27731972-31979. Deery, W. J., and R. H. Gomer. 1999. A putative receptor mediating cell-density sensing in Dictyostelium. J. Biol. Chem. 274~34476-34482. de la Roche, M., and G. Cote. 2001. Regulation of Dictyostelium myosin I and 11. Biochim. Biophys. Acta 1525:245261. De Lozanne, A., and J. A. Spudich. 1987. Disruption of the Dictyostelium myosin heavy chain gene by homologous recombination. Science 236:1086-1091. Devine, K. M., and W. F. Loomis. 1985. Molecular characterization of anterior-like cells in Dictyostelium discoideum. Dev. Biol. 107:364-372. Devreotes, P. 1989. Dictyostelium discoideum: a model system for cell-cell interactions in development. Science 245:10541058. Dharmawardhane, S., V. Warren, A. L. Hall, and J. Condeelis. 1989. Changes in the association of actin-binding proteins with the actin cytoskeleton during chemotactic stimulation of Dictyostelium discoideum. Cell Motil. Cytoskeleton 1357-63. Dormann, D., B. Vasiev, and C. J. Weijer. 1998. Propagating waves control Dictyostelium discoideum morphogenesis. Biophys. Chem. 72:21-35. Eichinger, L., J. Pachebat, G. Glockner, M. Rajandream, R. Sucgang, M. Berriman, J. Song, R. Olsen, K. Szafranski, Q. Xu, B. Tunggal, S. Kummerfeld, M. Madera, B. Konfortov, F. Rivero, A. Bankier, R. Lehmann, N. Hamlin, R. Davies, P. Gaudet, P. Fey, K. Pilcher, G. Chen, D. Saunders, E. Sodergren, P. Davis, A. Kerhornou, X. Nie, N. Hall, C. Anjard,
450 L. Hemphill, N. Bason, P. Farbrother, B. Desany, E. Just, T. Morio, R. Rost, C. Churcher, J. Cooper, S. Haydock, N. van Driessche, A. Cronin, I. Goodhead, D. Muzny, T. Mourier, A. Pain, M. Lu, D. Harper, R. Lindsay, H. Hauser, K. James, M. Quiles, M. Madan Babu, T. Saito, C. Buchrieser, A. Wardroper, M. Felder, M. Thangavelu, D. Johnson, A. Knights, H. Loulseged, K. Mungall, K. Oliver, C. Price, M. Quail, H. Urushihara, J. Hernandez, E. Rabbinowitsch, D. Steffen, M. Sanders, J. Ma, Y. Kohara, S. Sharp, M. Simmonds, S. Spiegler, A. Tivey, S. Sugano, B. White, D. Walker, J. Woodward, T. Winckler, Y. Tanaka, G. Shaulsky, M. Schleicher, G. Weinstock, A. Rosenthal, E. Cox, R. Chisholm, R. Gibbs, W. Loomis, M. Platzer, R. Kay, J. Williams, P. Dear, A. Noegel, B. Barrell, and A. Kuspa. 2005. The genome of the social amoeba Dictyostelium discoideum. Nature 435:43-57. Elson, E. L., S. F. Felder, P. Y. Jay, M. S. Kolodney, and C. Pasternak. 1999. Forces in cell locomotion. Biochem. SOC. S y m p . 65:299-314. Esch, R. K., P. K. Howard, and R. A. Firtel. 1992. Regulation of the Dictyostelium CAMP-induced, prestalk-specific DdrasD gene-identification of cis-acting elements. Nucleic Acids Res. 20:1325-1332. Fields, S., Q. Arana, J. Heuser, and M. Clarke. 2002. Mitochondrial membrane dynamics are altered in cluA-mutants of Dictyostelium. J. Muscle Res. Cell Motil. 235329438. Firtel, R. A. 1996. Interacting signaling pathways controlling multicellular development in Dictyostelium. Curr. Opin. Genet. Dev. 6545-554. Fukuzawa, M., T. Abe, and J. G. Williams. 2003. The Dictyostelium prestalk cell inducer DIF regulates nuclear accumulation of a STAT protein by controlling its rate of export from the nucleus. Development 130:797-804. Funamoto, S., K. Milan, R. Meili, and R. Firtel. 2001. Role of phosphatidylinositol3' kinase and a downstream pleckstrin homology domain-containing protein in controlling chemotaxis in Dictyostelium. J. Cell Biol. 153:795-810. Gao, T., D. Knecht, L. Tang, R. D. Hatton, and R. Gomer. 2004. A cell number counting factor regulates Akt/Protein Kinase B to regulate group size in Dictyostelium discoideum group size. Eukaryot. Cell 3:1176-1184. Garrod, D. R., and J. M. Ashworth. 1972. Effect of growth conditions on development of the cellular slime mould Dictyostelium discoideum. J. Embryol. Exp. Morphol. 28:463479. Gilman, A. G. 1987. G proteins: transducers of receptor-generated signals. Annu. Rev. Biochem. 56:615-649. Gomer, R. H. 1998. Antisense: a key tool for cell and developmental studies in Dictyostelium, p. 135-141. In J. K. Setlow (ed.), Genetic Engineering, vol. 20. Plenum Press, New York, NY. Gomer, R. H., and R. A. Firtel. 1987a. Cell-autonomous determination of cell-type choice in Dictyostelium development by cell-cycle phase. Science 237:758-762. Gomer, R. H., and R. A. Firtel. 1987b. Tissue morphogenesis in Dictyostelium discoideum, p. 373-383. In Molecular Approaches to Developmental Biology. Alan R. Liss, Inc., New York, NY. Grabel, L., and W. F. Loomis. 1978. Effector controlling accumulation of N-acteylglucosaminidase during development of Dictyostelium discoideum. Dev. Biol. 64:203-209.
ANALOGOUS SYSTEMS Gross, J. D., J. Bradbury, R. R. Kay, and M. J. Peacey. 1983. Intracellular pH and the control of cell differentiation in Dictyostelium discoideum. Nature 303:244-245. Harwood, A. J., N. A. Hopper, M. N. Simon, D. M. Driscoll, M. Veron, and J. G. Williams. 1992. Culmination in Dictyostelium is regulated by the CAMP-dependent protein kinase. Cell 69:615-624. Hofer, T., and P. K. Maini. 1997. Streaming instability of slimemold amebas-an analytical model. Phys. Rev. 56:20742080. Huang, Y. E., M. Iijima, C. A. Parent, S. Funamoto, R. A. Firtel, and P. Devreotes. 2003. Receptor-mediated regulation of PI3Ks confines PI(3,4,5)P3 to the leading edge of chemotaxing cells. Mol. Biol. Cell 14:1913-1922. Insall, R., A. Kuspa, P. J. Lilly, G. Shaulsky, L. R. Levin, W. F. Loomis, and P. Devreotes. 1994. CRAC, a cytosolic protein containing a pleckstrin homology domain, is required for receptor and G protein-mediated activation of adenylyl cyclase in Dictyostelium. 1. Cell Biol. 126:1537-1545. Iranfar, N., D. Fuller, and W. Loomis. 2003. Genome-wide expression analyses of gene regulation during early development of Dictyostelium discoideum. Eukaryot. Cell 2:664670. Jain, R., I. S. Yuen, C. R. Taphouse, and R. H. Gomer. 1992. A density-sensing factor controls development in Dictyostelium. Genes Dev. 6:390400. Jang, W., B. Chiem, and R. H. Gomer. 2002. A secreted cellnumber counting factor represses intracellular glucose levels to regulate group size in Dictyostelium. ]. Biol. Chem. 277:3 1972-3 1979. Jang, W., and R. H. Gomer. 2005. Exposure of cells to a cellnumber counting factor decreases the activity of glucose6-phosphatase to decrease intracellular glucose levels in Dictyostelium. Eukaryot. Cell 4:991-998. Jermyn, K. A., K. T. Duffy, and J. G. Williams. 1989. A new anatomy of the prestalk zone in Dictyostelium. Nature 3 40: 144-1 46. Jermyn, K. A., and J. G. Williams. 1991. An analysis of culmination in Dictyostelium using prestalk and stalk-specific cell autonomous markers. Development 111:779-787. Julien, B., A. D. Kaiser, and A. Garza. 2000. Spatial control of cell differentiation in Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 97:9098-9103. Kamboj, R. K., T. Y. Lam, and C. H. Siu. 1990. Regulation of slug size by the cell adhesion molecule gp80 in Dictyostelium discoideum. Cell Regul. 1:715-729. Kay, R. R., M. Berks, and D. Traynor. 1989. Morphogen hunting in Dictyostelium. Development 107(Suppl.):8 1-90. Kay, R. R., and K. A. Jermyn. 1983. A possible morphogen controlling differentiation in Dictyostelium. Nature 303~242-244. Kellerman, K. A., and J. G. McNally. 1999. Mound-cell movement and morphogenesis in Dictyostelium. Dev. Biol. 208~416-429. Kessin, R. H. 2001. Dictyostelium-Evolution, Cell Biology, and the Development of Multicellularity. Cambridge University Press, Cambridge, United Kingdom. Knecht, D. A., J. H. Jung, and L. Matthews. 1990. Quantification of transformation efficiency using a new method for
25. A EUKARYOTIC NEIGHBOR: D. DISCOIDEUM clonal growth and selection of axenic Dictyostelium cells. Dev. Genet. 11:403-409. Kumagai, A., J. A. Hadwiger, M. Pupillo, and R. A. Firtel. 1991. Molecular genetic analysis of two G-alpha protein subunits in Dictyostelium. J. Biol. Chem. 266:1220-1228. Kumagai, A., M. Pupillo, R. Gundersen, R. Make-Lye, J?. N. Devreotes, and R. A. Firtel. 1989. Regulation and function of G a protein subunits in Dictyostelium. Cell 57:265275. Kuspa, A., T. Dingermann, and W. Nellen. 1995. Analysis of gene function in Dictyostelium. Experientia 51:1116-1123. Kuspa, A., and W. F. Loomis. 1992. Tagging developmental genes in Dictyostelium by restriction enzyme-mediated integration of plasmid DNA. Proc. Natl. Acad. Sci. USA 89: 8 803-8 807. Kuwayama, H., S. Ishida, and P. J. Van Haastert. 1993. Nonchemotactic Dictyostelium discoideum mutants with altered cGMP signal transduction. J. Cell Biol. 123:1453-1462. Kuwayama, H., and P. J. M. van Haastert. 1998. cGMP potentiates receptor-stimulated Ca2+ influx in Dictyostelium discoideum. Biochim. Biophys. Acta 1402:102-108. Kwong, L., and G. Weeks. 1989. Studies on the accumulation of the differentiation-inducing factor (DIF) in high-celldensity monolayers of Dictyostelium discoideum. Dev. Biol. 132554-558. Liang, W., L. Licate, H. Warrick, J. Spudich, and T. Egelhoff. 2002. Differential localization in cells of myosin I1 heavy chain kinases during cytokinesis and polarized migration. BMC Cell Biol. 3:19. Lilly, P. J., and P. N. Devreotes. 1995. Chemoattractant and GTP gamma S-mediated stimulation of adenylyl cyclase in Dictyostelium requires translocation of CRAC to membranes.]. Cell Biol. 129:1659-1665. Loomis, W. F. 1987. Genetic tools for Dictyostelium discoideum, p. 31-65. In J. A. Spudich (ed.),Methods in Cell Biology. Academic Press, Orlando, FL. Luck-Vielmetter, D., M. Schleicher, B. Grabatin, J. Wippler, and G. Gerisch. 1990. Replacement of threonine residues by serine and alanine in a phosphorylatable heavy chain fragment of Dictyostelium Myosin-11. FEBS Lett. 269:239-243. Mann, S. K. O., and R. A. Firtel. 1987. Cyclic AMP regulation of early gene expression in Dictyostelium discoideum: mediation via the cell surface cyclic AMP receptor. Mol. Cell. Biol. 7:458-469. Manstein, D. J. 1993. Myosin function in the motile behaviour of cells. Symp. Soc. Exp. Biol. 47:375-381. Maree, A. F. M., and P. Hogeweg. 2001. How amoeboids selforganize into a fruiting body: multicellular coordination in Dictyostelium discoideum. Proc. Natl. Acad. Sci. USA 98:3 8 79-3 8 83. Martiel, J. L., and A. Goldbeter. 198.5. Autonomous chaotic behaviour of the slime mould Dictyostelium discoideum predicted by a model for cyclic AMP signaling. Nature 313590-5 92. McDonald, S. A., and A. J. Durston. 1984. The cell cycle and sorting behaviour in Dictyostelium discoideum. J. Cell Sci. 66: 195-204. McRobbie, S. J., and P. C. Newell. 1983. Changes in actin associated with cytoskeleton following chemotactic stimulation
451 of Dictyostelium discoideum. Biochem. Biophys. Res. Commun. 115:351-359. Mehdy, M. C., and R. A. Firtel. 1985. A secreted factor and cyclic AMP jointly regulate cell-type-specific gene expression in Dictyostelium discoideum. Mol. Cell. Biol. 5:705713. Meili, R., C. Ellsworth, S. Lee, T. B. Reddy, H. Ma, and R. A. Firtel. 1999. Chemoattractant-mediated transient activation and membrane localization of Akt/PKB is required for efficient chemotaxis to cAMP in Dictyostelium. EMBO J. 18:2092-2105. Milne, J. L., and P. N. Devreotes. 1993. The surface cyclic AMP receptors, cAR1, cAR2, and cAR3, promote Ca2+ influx in Dictyostelium discoideum by a G alpha-2-independent mechanism. Mol. Biol. Cell 4:283-292. Nellen, W., C. Silan, and R. A. Firtel. 1984. DNA-mediated transformation in Dictyostelium discoideum: regulated expression of an actin gene fusion. Mol. Cell. Biol. 4:28902898. Newell, P. C. 1995. Calcium, cyclic GMP and the control of myosin I1 during chemotactic signal transduction of Dictyostelium. J. Biosci. 20:289-310. Noegel, A. A., and J. E. Luna. 199.5. The Dictyostelium cytoskeleton. Experientia 51:1135-1143. Otto, G. P., M. Y. Wu, M. Clarke, H. Lu, 0. R. Anderson, H. Hilbi, H. A. Shuman, and R. H. Kessin. 2004. Macroautophagy is dispensable for intracellular replication of Legionella pneumophila in Dictyostelium discoideum. Mol. Microbiol. 51:63-72. Pate, E. F., and G. M. Odell. 1981. A computer simulation of chemical signaling during the aggregation phase of Dictyostelium discoideum. J. Theor. Biol. 88:201-239. Pate, E. F., and H. G. Othmer. 1986. Differentiation, cell sorting and proportion regulation in the slug stage of Dictyostelium discoideum.]. Theor. Biol. 118:301-319. Pears, C . J., and J. G. Williams. 1987. Identification of a DNA sequence element required for efficient expression of a developmentally regulated and CAMP-iducible gene of Dictyostelium discoideum. EMBO J. 6:195-200. Rathi, A., S. C. Kayman, and M. Clarke. 1991. Induction of gene expression in Dictyostelium by prestarvation factor, a factor secreted by growing cells. Dev. Genet. 12532-87. Reymond, C. D., R. H. Gomer, W. Nellen, A. Theibert, P. Devreotes, and R. Firtel. 1986. Phenotypic changes induced by a mutated ras gene during the development of Dictyostelium transformants. Nature 323:340-343. Roisin-Bouffay, C., W. Jang, and R. H. Gomer. 2000. A precise group size in Dictyostelium is generated by a cell-counting factor modulating cell-cell adhesion. Mol. Cell 6:953-959. Rosenbluh, A., R. Nir, E. Sahar, and E. Rosenberg. 1989. Celldensity-dependent lysis and sporulation of Myxococcus xanthus in agarose beads. J. Bacteriol. 171:4923-4929. Sabry, J. H., S. L. Moores, S. Ryan, J. H. Zang, and J. A. Spudich. 1997. Myosin heavy chain phosphorylation sites regulate myosin localization during cytokinesis in live cells. Mol. Biol. Cell 8:2605-261.5. Saxe, C. L., 111, R. Johnson, P. N. Devreotes, and A. R. Kimmel. 1991. Multiple genes for cell surface cAMP receptors in Dictyostelium discoideum. Dev. Genet. 12:6-13.
452 Schaap, P., and M. Wang. 1986. Interactions between adenosine and oscillatory CAMP signaling regulate size and pattern in Dictyostelium. Cell 45:137-144. Shaffer, B. M. 1957. Variability of behavior of aggregating cellular slime moulds. Q. J. Microsc. Sci. 98:393-405. Shaulsky, G., R. Escalante, and W. F. Loomis. 1996. Developmental signal transduction pathways uncovered by genetic suppressors. Proc. Natl. Acad. Sci. USA 93:15260-15265. Siu, C. H., and R. K. Kamboj. 1990. Cell-cell adhesion and morphogenesis in Dictyostelium discoideum. Dev. Genet. 11:377-3 87. Solomon, J. M., A. Rupper, J. A. Cardelli, and R. R. Isberg. 2000. Intracellular growth of Legionella pneumophila in Dictyostelium discoideum, a system for genetic analysis of host-pathogen interactions. Infect. Immun. 68:2939-2947. Souza, G. M., S. J. Lu, and A. Kuspa. 1998. YakA, a protein kinase required for the transition from growth to development in Dictyostelium. Development 125:2291-2302. Spann, T. P., D. A. Brock, D. F. Lindsey, S. A. Wood, and R. H. Gomer. 1996. Mutagenesis and gene identification in Dictyosteliurn by shotgun antisense. Proc. Natl. Acad. Sci. USA 93~5003-5007. Steimle, P., S. Yumura, G. Cote, Q. Medley, M. Polyakov, B. Leppert, and T. Egelhoff. 2001. Recruitment of a myosin heavy chain kinase to actin-rich protrusions in Dictyostelium. Curr. Biol. 11:708-713. Sternfeld, J. 1988. Proportion regulation in Dictyostelium is altered by oxygen. Differentiation 37:173-179. Sternfeld, J., and C. N. David. 1982. Fate and regulation of anterior-like cells in Dictyostelium slugs. Dev. Biol. 93:lll-118. Sussman, M. 1987. Cultivation and synchronous morphogenesis of Dictyostelium under controlled experimental conditions, p. 9-29. In J. A. Spudich (ed.), Methods in Cell Biology, vol. 28. Academic Press, Orlando, FL. Taminato, A., R. Bagattini, R. Gorjao, G. K. Chen, A. Kuspa, and G. M. Souza. 2002. Role for YakA, CAMP, and protein kinase A in regulation of stress responses of Dictyostelium discoideum cells. Mol. Biol. Cell 13:2266-2275. Tanaka, K., H. Adachi, H. Konishi, A. Iwamatsu, K. Ohkawa, T. Shirai, S. Nagata, U. Kikkawa, and Y. Fukui. 1999. Identification of protein kinase B (PKB) as a phosphatidylinositol 3,4,5-trisphosphate binding protein in Dictyostelium discoideum. Biosci. Biotechnol. Biochem. 63:368-372. Tang, L., T. Gao, C. McCollum, W. Jang, M. G. Vickers, R. Ammann, and R. H. Gomer. 2002. A cell number-counting factor regulates the cytoskeleton and cell motility in Dictyostelium. Proc. Natl. Acad. Sci. USA 99:1371-1376. Theibert, A., and P. Devreotes. 1986. Surface receptor-mediated activation of adenylate cyclase in Dictyostelium regulation by guanine nucleotides in wild-type cells and aggregation deficient mutants. J . Biol. Chem. 261:15121-15125. Van Driessche, N., C. Shaw, M. Katoh, T. Morio, R. Sucgang, M. Ibarra, H. Kuwayama, T. Saito, H. Urushihara, M. Maeda, I. Takeuchi, H. Ochiai, W. Eaton, J. Tollett,
ANALOGOUS SYSTEMS J. Halter, A. Kuspa, Y. Tanaka, and G. Shaulsky. 2002. A transcriptional profile of multicellular development in Dictyosteliurn discoideum. Development 129:1543-1552. Van Haastert, P. J. M. 1984. Guanine nucleotides modulate cell surface CAMP-binding sites in membranes from Dictyostelium discoideum. Biochem. Biophys. Res. Commun. 124~597-604. Van Haastert, P. J. M., J. D. Bishop, and R. H. Gomer. 1996. The cell density factor CMF regulates the chemoattractant receptor CAR1 in Dictyostelium. J. Cell Biol. 134:15431549. Van Haastert, P. J. M., and H. Kuwayama. 1997. cGMP as second messenger during Dictyostelium chemotaxis. FEBS Lett. 4 10:25-2 8. Van Haastert, P. J. M., B. E. Snaar-Jagalska, and P. M. W. Janssens. 1987. The regulation of adenylate cyclase by guanine nucleotides in Dictyostelium discoideum membranes. Eur. J . Biochern. 162:25 1-25 8. Wang, N., F. Soderbom, C. Anjard, G. Shaulsky, and W. F. Loomis. 1999. SDF-2 induction of terminal differentiation in Dictyostelium discoideum is mediated by the membranespanning sensor kinase DhkA. Mol. Cell. Biol. 19:47504756. Weijer, C. J. 1999. Morphogenetic cell movement in Dictyostelium. Semin. Cell Dev. Biol. 10:609-619. Weijer, C. J., G. Duschl, and C. N. David. 1984. A revision of the Dictyostelium discoideum cell cycle. J. Cell Sci. 70:lll131. Williams, J. G., A. Ceccarelli, S. McRobbie, H. Mahbubani, R.R. Kay, A. Farly, M. Berks, and K. A. Jermyn. 1987. Direct induction of Dictyostelium prestalk gene expression by DIF provides evidence that DIF is a morphogen. Cell 49: 185-192. Williams, R. S., L. Cheng, A. W. Mudge, and A. J. Harwood. 2002. A common mechanism of action for three moodstabilizing drugs. Nature 417:292-295. Wireman, J. W., and M. Dworkin. 1975. Morphogenesis and developmental interactions in myxobacteria. Science 189:516-523. Wood, S. A., R. R. Ammann, D. A. Brock, L. Li, T. P. Spann, and R. H. Gomer. 1996. RtoA links initial cell type choice to the cell cycle in Dictyostelium. Development 122:36773685. Wu, L. J., and P. N. Devreotes. 1991. Dictyostelium transiently expresses eight distinct G-protein alpha-subunits during its developmental program. Biochem. Biophys. Res. Commun. 179:1141-1147. Xie, Y. J., L. Kwong, and G. Weeks. 1991. A possible role for DIF-2 in the formation of stalk cells during Dictyostelium development. Dev. Biol. 145:195-200. Yuen, I. S., R. Jain, J. D. Bishop, D. F. Lindsey, W. J. Deery, P. J. M. Van Haastert, and R. H. Gomer. 1995. A densitysensing factor regulates signal transduction in Dictyostelium. .J.Cell Biol. 129:1251-1262.
Myxobacteria: Multicellularity and Differentiation Edited by David E. Whitworth 02008 ASM Press, Washington, D.C.
Paul E. Kolenbrander Nicholas S. Jakubovics Natalia I. Chalmers
Multispecies Interactions and Biofilm Community Development
Bacteria interact with other microorganisms and coexist naturally on surfaces as biofilms. Among the most intricate biofilms are the myxobacteria gliding across surfaces in search of nutrients and implementing developmental programs that culminate in fruiting bodies. Most studies of biofilms have focused on single species and on genes that control or are regulated by life on a surface. Comparative microarray analyses of gene expression by planktonic versus surface-attached bacteria have yielded numerous candidate genes for in-depth analysis. As more information is uncovered by studies of pure cultures, these data can be applied towards understanding the roles of specific genes in multispecies interactions. In nature, most bacteria live in multispecies settings and are immersed in a variety of stimuli. For any one of these stimuli to be effective in eliciting a response from a species, the duration of the stimulus must be in harmony with the function it regulates. A particular gene-specific response to a stimulus in a pure culture setting may also be critical at a specific time in mixed-species settings. Myxobacteria are renowned for their highly organized developmental patterns that take them from the
26
vegetative state to a fruiting body. Multispecies communal organization among their neighbors is analogous in many ways. In this chapter, most of the focus is on multispecies interactions among oral bacteria in biofilms: a few single-species biofilms are featured to discuss responses to environmental signals, including signals generated by the occupants within the biofilm.
SIGNALS AND COMMUNICATION Signals involved in cell-to-cell communication among biofilm cells include acyl homoserine lactones, oligopeptides, and autoinducer-2 (AI-2).Some intracellular signals such as the recently discovered second messenger cyclic bis(3’,5’)guanylic acid (cdiGMP) regulate exopolysaccharide production and thus affect biofilm development. Information is conveyed indirectly through rhamnolipid concentration gradients in Pseudornonas aeruginosa and directly by outer membrane lipoproteins in Myxococcus xanthus. Various modes of signaling and communication are the topic of numerous reviews (Fuqua
Paul E. Kolenbrander and Nicholas S. Jakubovics, Oral Biofilm Communication Unit, Oral Infection and Immunity Branch, National Institute of Dental and Craniofacial Research, National Institutes of Health, Bethesda, MD 20892. Natalia I. Chalmers, Oral Biofilm Communication Unit, Oral Infection and Immunity Branch, National Institute of Dental and Craniofacial Research, National Institutes of Health, Bethesda, MD 20892, and Department of Biomedical Sciences, University of Maryland School of Dentistry, Baltimore, MD 21201.
453
ANALOGOUS SYSTEMS
454 et al., 2001; Keller and Surette, 2006; Kolenbrander et al., 2002; Waters and Bassler, 2005), which reflects the high interest in understanding the mechanisms of community information exchange.
Acyl Homoserine Lactones and Oligopeptides The acyl homoserine lactones all share a common homoserine lactone moiety and differ in their acyl side chain. These signals are produced by gram-negative bacteria and are typically species specific in that the species that produces a particular acyl homoserine lactone is the species that responds to it. As progressively greater emphasis is placed on investigating mixed-species communication, examples of cross-species (Riedel et al., 2001; Steidle et al., 2001) interactions via acyl homoserine lactones as well as bacterium-eukaryote interactions mediated by acyl homoserine lactones (Tait et al., 2005) and other small-molecular signals (Dudler and Eberl, 2006) are uncovered. A role in biofilm development for one of these acyl homoserine lactones was first shown in P. aeruginosa (Davies et al., 1998). Biofilm cocultures of P. aeruginosa and Agrobacterium tumefaciens were dominated by pseudomonads, required flagella and type IV pili on the pseudomonads, and were enhanced by pseudomonad acyl-homoserine lactones (An et al., 2006). Speciesspecific signaling in gram-positive bacteria is mediated by oligopeptides, including competence-stimulating peptides that increase the frequency of, and in some examples are essential for, genetic transformation. Several species of human oral streptococci are naturally competent, and the competence-stimulating peptide (CSP) increases biofilm growth of Streptococcus intermedius (Petersen et al., 2004). Inactivation of comD, the histidine kinase of the two-component signal transduction system that binds CSP, by transposon mutagenesis in Streptococcus gordonii caused reduced biofilm growth (Loo et al., 2000). In agreement with these data, real time reverse transcriptase PCR quantification of S. gordonii mRNAs revealed that comD and comE (response regulator of the two-component signal transduction system) were upregulated in the biofilm phase of growth (Gilmore et al., 2003). In Streptococcus mutans NG8, mutants defective in comC, encoding CSP, in comD or comE, or in comX, encoding an alternate sigma factor, produced abnormal biofilms: the phenotype of comD and comE mutants was reduced biomass (Li et al., 2002). Others, using a different strain, S. mutans UA159, reported multilevel controls that include two signal-transducing twocomponent systems that integrate multiple environmental signals to coordinate competence development, acid tolerance, and biofilm formation (Ahn et al., 2006). Further, these authors noted that inactivation of comC,
which encodes CSP, did not change the frequency of transformation (Ahn et al., 2006). The broad range of activities of acyl homoserine lactones and oligopeptides has been reviewed in depth elsewhere (Senadheera et al., 2005; Visick and Fuqua, 2005; Waters and Bassler, 2005).
AI-2 In contrast to the first two classes of signals, AI-2 is proposed to be a universal signal that mediates interspecies communication (Schauder et al., 2001) and is thus targeted towards multispecies biofilms. AI-2 was first described in the bioluminescent marine bacterium Vibrio harveyi (Bassler et al., 1993). AI-2 is extracellular, is detected by an assay that measures induction of bioluminescence in the V. harveyi BB170 reporter strain in response to conditioned cell-free culture medium, and is present in culture supernatants from numerous grampositive and gram-negative bacteria. The AI-2 biosynthetic pathway involves a three-step enzymatic production of the AI-2 precursor 4,5-dihydroxy-2,3-pentanedione (DPD) from S-adenosyl-L-methionine, a major methyl donor for cellular methyltransferase reactions. LuxS, the AI-2 synthase, in the third step cleaves S-ribosyl-Lhomocysteine to homocysteine and DPD (Duerre et al., 1971). Spontaneous rearrangement of DPD in solution (Semmelhack et al., 2005) results in a collection of molecules all in equilibrium and called AI-2. The luxS gene is present in most bacteria, which indicates that AI-2 from several sources contributes to the extracellular stimuli of multispecies community environments. The observation that conditioned cell-free culture media from numerous species induce bioluminescence in V harveyi promoted the hypothesis that AI-2 is a universal interspecies bacterial signal (Schauder et al., 2001). Evidence to support this hypothesis is accumulating, and results from three studies provide strong support. Two structurally similar AI-2 molecules have been cocrystallized with their respective receptors: R-THMF from Salmonella enterica serovar Typhimurium (Miller et al., 2004) and S-THMF-borate from V harveyi (Chen et al., 2002). The two molecules are in equilibrium with other AI-2 forms and are interconvertible, that is, each can be released from its purified receptor, meld in the AI-2 equilibrium, and bind to the other’s receptor. Second, a two-way interconvertible AI-2 communication between V. harveyi and Escherichia coli and between Vibrio cholerae and E. coli in batch culture was shown: AI-2 produced by one species promotes major effects on gene expression in the partnered species (Xavier and Bassler, 2005). The local AI-2 concentration in dualspecies culture is modulated by the ability of enteric
26. MULTISPECIES INTERACTIONS AND BIOFILMCOMMUNITIES bacteria to consume the AI-2 that they produce and thus interfere with AI-2-mediated communication with nearby species. Third, chemically synthesized DPD complemented a luxS mutation in Streptococcus oralis and mediated mutualistic growth of the streptococcus and Actinomyces naeslundii in biofilms with saliva as the sole source of nutrient (Rickard et al., 2006). The wild type of each organism was unable to grow in monoculture solely on saliva. Indeed, the AI-2 signal is suited for multispecies community development in which the members of the consortia each have a role in the utilization of complex nutritional sources as growth substrates. An intriguing question emerges, “How can an identical diffusible molecule produced by many bacteria possess any specificity to elicit species-specific informational responses?” In biofilms, diffusible factors are swept away from cells by bulk fluid flow, and therefore flow is a critical factor that determines whether signal molecules reach their targets. The mutualism pair, S. oralis and A. naeslundii, offered an opportunity to investigate AI-2 as an interspecies signal (Rickard et al., 2006). The luxS gene was inactivated in S. oralis: this gene could not be genetically manipulated in A. naeslundii. Cell-free supernatants from broth culture-grown S . oralis or A. naeslundii exhibited AI-2 activity by the bioluminescence-based assay, and the S. oralis luxS mutant did not produce AI-2. In dual-species biofilms formed under flowing conditions in saliva-fed flowcells, the wild-type strains showed luxuriant interdigitated mutualistic growth. No mutualism was seen with the S. oralis luxS mutantlA. naeslundii dual-species biofilm. Genetic complementation (transformation with a luxS-containing plasmid) and chemical complementation (synthetic AI-2 in the form of DPD) restored mutualism as well as the high biomass characteristic for the wild-type dual-species biofilm. Importantly, an optimal concentration of DPD was critical for maximal biofilm development. Concentrations of synthetic DPD that were higher or lower than the optimal concentration yielded less biomass. In addition, the optimal concentration was 100-fold lower than the detection limit of the currently accepted AI-2 assay. Thus, AI-2 acts as a bona fide interspecies signal whose target is sensitive to its concentration. Presumably, higher or lower AI-2 concentrations might also be critical for other targets in multispecies communities. Postulating that different bacteria sense different threshold concentrations of AI-2 agrees with the proposal that AI-2 is a universal signal that mediates species-specific responses. In this way, the AI-2 produced by all members of the community could be an interspecies signaling molecule at each and all concentrations. The biofilm with the S. oralis and A. naeslundii dual-species community
455
growing solely on saliva illustrates a community responsive to a specific concentration of AI-2.
cdiGMP Intracellular signaling may also be important in biofilm development. For example, in Shewanella oneidensis, a mineral-reducing soil bacterium, a small-molecule signal, cdiGMP, is an intracellular regulator that controls biofilm stability (attachment and detachment) through regulating the synthesis of exopolysaccharides (Thormann et al., 2006), critical components of many biofilms including those formed by myxobacteria. The molecule cdiGMP is proposed to control biofilm stability in a concentration-dependent manner. Higher concentrations promote attachment of cells to the biofilm; lower concentrations promote detachment. As a secondary messenger molecule, cdiGMP may allosterically control exopolysaccharide production. Allosteric regulation of cellulose synthase by cdiGMP was noted initially for extracellular cellulose production in Gluconacetobacter xylinus (Ross et al., 1987, 1990) and is now noted for biofilm formation by Yersinia pestis (Kirillina et al., 2004) and I? cholerae (Tischler and Camilli, 2004) and exopolysaccharide formation by the heterotroph Thermotoga maritima in coculture with the methanogen Methanococcus jannaschii (Johnson et al., 2005). The P. aeruginosa PA0 1genome contains 3 8 genes encoding enzymes with putative domains characteristic of diguanylate cyclases (DGCs) and phosphodiesterases (PDEs), which synthesize and hydrolyze, respectively, cdiGMP (Kulasakara et al., 2006). Seventeen proteins have the DGC domain, 5 have the PDE domain, and 16 have both domains. Transposon insertions in all DGC and PDE domains were prepared, and enhanced biofilm formation was associated at least in part with increased cdiGMP levels. However, distinct phenotypes were observed with these insertions, suggesting that the cdiGMP levels are localized and specifically linked to a targeted function. Unlike freely diffusible extracellular AI-2, intracellular cdiGMP is apparently function-targeted and not freely diffusible. A connection between cdiGMP and AI-2 in regulating cellular behaviors including biofilm development is proposed in a review (Camilli and Bassler, 2006). As another example of small-molecule signals, cdiGMP brings additional emphasis on unraveling the mechanisms of communication among multispecies groups.
Rhamnolipids Movement on a surface occurs in biofilms. If the biofilm forms in a flowing system, the movement might occur by bacteria entering the bulk fluid and passively moving downstream. Alternatively, bacteria can actively move by
ANALOGOUS SYSTEMS
456 means of their flagella. Some flagellated bacteria swarm on semisolid agar and form tendril-like extensions radiating out from a central inoculation site (Caiazza et al., 2005). Bacteria do not occupy the spaces between tendrils, which suggests that a diffusible molecule modulates the spacing of the tendrils. When tendrils radiating from two nearby inoculation sites approach each other, they do not overlap; instead, they extend parallel to each other. An interesting and at first puzzling role for rhamnolipids in the swarming movement was discovered (Caiazza et al., 2005). Rhamnolipids are surfactants, and their sole role in swarming was thought, previously, to be overcoming surface tension and promoting motility. How then can rhamnolipids both enhance motility and inhibit swarming? The answer to the puzzle was achieved by dissecting the biosynthetic pathway of rhamnolipids. The rhamnolipid precursors, 3-(3-hydroxyalkanoyloxy) alkanoic acids, are the primary surface-wetting agents for swarming, but rhamnolipids coordinate group behavior and modulate swarming. It is proposed that the secreted rhamnolipids form a gradient and the gradient maintains tendril formation in a radiating linear fashion, keeping tendrils separated from each other (Caiazza et al., 2005). Likewise, when approaching each other, tendrils from different inoculation sites change direction because the secreted rhamnolipid at the leading edge of the tendril signals the directional change.
regulator that controls retention of cells in biofilms is the above-mentioned cdiGMP (Kulasakara et al., 2006; Thormann et al., 2006). High concentrations of this regulator are proposed to enhance attachment of cells to a surface, and low concentrations permit dispersal of cells. Concentration-dependent responses to regulator and signaling molecules provide stability to biofilms and resilience against nutrient shock and other environmental perturbations. Enzymatic disruption of biofilms has been reported for an oral bacterium, Aggregatibacter (formerly Actinobacillus [Norskov-Lauritsen and Kilian, 20061) actinomycetemcomitans (Kaplan et al., 2003). A soluble P-N-acetyl-glucosaminidase (Kaplan et al., 2003) hydrolyzes the exopolysaccharide matrix of polymeric p-1,6N-acetyl-D-glucosamine (PGA), which is produced in biofilms by many species and depolymerized, dispersing cells in the process (Itoh et al., 2005; Kaplan et al., 2004). The enzyme termed dispersin B in A. actinomycetemcomitans (Kaplan et al., 2003) has been crystallized (Ramasubbu et al., 2005), and the gene cluster pgaABCD for biosynthesis of the enzyme substrate PGA has been identified in A. actinomycetemcomitans (Kaplan et al., 2004) and E. coli (Wang et al., 2004). PGAis not required for attachment of A. actinomycetemcomitans, indicating that dispersal of cells in a biofilm is a process distinct from attachment. However, both are part of the biofilm maintenance and development.
Outer Membrane Lipoproteins Signaling information is not limited to diffusible small molecules. Outer membrane lipoproteins Tgl and CglB of Myxococcus xanthus are part of the S-motility and the A-motility systems, respectively; these lipoproteins physically transfer between cells and facilitate organized myxobacterial gliding (Nudleman et al., 2005).
DISPERSAL OF BIOFILMS Most often, we think about the formation and development of biofilms as the continuous addition of biomass. However, continuous erosion of cells, probably those loosely attached, also occurs. Cells are removed from the biofilm by high shear forces (Stoodley et al., 2001) or by active detachment mechanisms, such as nutrientinduced dispersion of P. aeruginosa biofilms (Sauer et al., 2004). In the latter situation, a rapid increase in carbon substrate availability resulted in markedly different gene expression in biofilm cells and newly dispersed planktonic cells. Genes upregulated in dispersed cells included those encoding flagellar components, and pilin biosynthetic genes were upregulated in biofilm cells. One molecule proposed to be a key intracellular
MULTISPECIES BIOFILMS One site where natural multispecies biofilms are unusually accessible is the tooth surface in the human oral cavity. We use a retrievable enamel chip model system (Palmer et al., 2001b) that permits us to place three pieces of enamel ( 3 by 3 by 1 mm each) side by side in a groove cut into an acrylic stent that is placed bilaterally on the buccal (cheek side) surface of the lower dentition. Thus, at any time point, up to six replicates can be obtained. A volunteer wears a stent for 4 or 8 h, and then the enamel pieces are removed and prepared for confocal scanning laser microscopy. Antibodies against specific bacterial cell surface molecules can be conjugated directly to a fluorophore, and, in addition, a general nucleic acid stain can be included to label all bacteria. Coupling this procedure to confocal scanning laser microscopy gives single-cell resolution and allows investigations of oral bacterial communities in biofilms formed temporally on enamel. The predominant bacteria on enamel at 4 h after professional cleaning are several streptococcal species, as determined by scraping the enamel surface and plating
26. MULTISPECIES INTERACTIONS AND BIOFILM COMMUNITIES on agar medium (Nyvad and IGlian, 1987). Other bacteria including actinomyces are also found. Streptococci and actinomyces coaggregate in specific patterns: coaggregation is defined as specific cell-cell interactions between genetically distinct cell types. It is distinguished from aggregation, which occurs between genetically identical cells, and from agglutination, which occurs by the aggregation of cells with a soluble molecule such as an antibody. Hundreds of strains of human oral species from many genera have been examined for their ability to coaggregate in vitro by mixing pairwise dense cell suspensions of each cell type, and all strains have partners (Kolenbrander, 1988). While these coaggregations in vitro are the subject of numerous reports, investigations of coaggregation in vivo are fewer, because it has been difficult to distinguish between coaggregates formed in vivo and randomly juxtapositioned bacteria. It became possible to investigate coaggregates in vivo after it was shown that many coaggregations are mediated by the recognition of adhesins on the actinomyces type 2 fimbriae and complementary receptor polysaccharide molecules on the streptococci (Cisar et al., 1997; Klier et al., 1997). Thus, juxtaposition of specific antibodies against type 2 fimbriae and specific antibodies against streptococcal receptor polysaccharide would be strong evidence that coaggregation occurs in vivo. Indeed, by using fluorophore-conjugated antibodies and confocal scanning laser microscopy, such evidence was obtained with the retrievable enamel chip in vivo model to confirm that coaggregations are common in the development of human oral biofilms (Palmer et al., 2003). A second question that can be answered by using fluorophore-conjugated antibodies and confocal laser microscopy is whether initial biofilm communities on enamel are monospecies or multispecies. Do bacteria grow as pure-culture colonies on enamel? We showed unambiguously that in vivo growth of oral bacteria typically occurs as multispecies communities on enamel (Palmer et al., 2003). The significanceof these two results, in vivo coaggregation and multispecies communities, is a demonstration that only specific partnerships colonize a given surface, because collectively they have an advantage that evolved in that ecosystem. Accordingly, human oral bacteria do not thrive in soil, and myxobacteria do not thrive in the human oral cavity. Consistently observed multispecies communities suggest that cell-cell interactions are prerequisites for biofilm development. To investigate cell-cell interactions, we use an in vitro flow cell system to parallel the in vivo retrievable enamel chip system. Saliva is the sole source of nutrient. Three coaggregating species were examined as dual-species inocula and as single-species biofilms.
457
A. naeslundii and S. oralis (the mutualism pair described in the AI-2 section above) were unable to grow as biofilm monocultures on saliva, whereas Streptococcus gordonii did grow (Palmer et al., 2001a). In dual-species biofilms, S. gordonii grew equally well with or without A. naeslundii or S. oralis; neither of the last two species exhibited much growth, but importantly, both were retained in the biofilm. Retention is a critical feature because it positions the organism for growth under more favorable conditions such as changes in pH, oxygen tension, or the arrival of a more beneficial partnership. Indeed, when A. naeslundii and S. oralis were inoculated to form a dualspecies biofilm with saliva as the sole nutritional source, both grew abundantly and to a significantly higher biomass than was observed with S. gordonii growing alone or with coaggregation partners. The mutualism of A. naeslundii and S. oralis on saliva and the independent growth of S. gordonii emphasize the differences in cellcell interactions between species and resultant multispecies biofilm communities. These results suggest that signals mediate the success or failure of multispecies community development. To investigate cell-cell communication in multispecies community growth, an in vitro flow cell containing the coaggregation partners Veillonella atypica and S. gordonii was inoculated (Egland et al., 2004). These species are also metabolically linked. S. gordonii ferments carbohydrates to form lactic acid. V. atypica is unable to ferment sugars, but it uses lactic acid as a preferred fermentation substrate, thus completing a metabolic coupling of the two species. When the two species are spotted together on agar containing starch, a zone of hydrolysis is evident that is absent around monoculture spots. The increase in starch hydrolysis is due to increased expression of the S. gordonii alpha-amylase-encoding gene amyB. To monitor cell-cell communication between the species, a transcriptional fusion (PamyB-’gfp)consisting of promoterless gfp (encoding green fluorescent protein [GFP]) under control of the promoter/operator of amyB was constructed. A plasmid containing PamyB-’gfp was transformed into S . gordonii, which was cocultured with I? atypica in flow cells with saliva as the sole source of nutrient. After a 4-h incubation in this flowing system, single-cell resolution of the biofilm by confocal scanning laser microscopy revealed that only those streptococci in juxtaposition with V. atypica expressed GFP. T/: atypica colonies could not grow, and nearby colonies composed solely of S. gordonii were not green, indicating that the amyB promoter was not activated. Considering that the flow cell is an open system, it is possible that the signaling molecule was produced but diluted quickly by the bulk fluid saliva and washed out of the flow cell. This
ANALOGOUS SYSTEMS
458 possibility was examined by incubating the two species in a closed vessel but physically separating them by a dialysis membrane. S. gordonii containing the PamyB-'gfp reporter plasmid exhibited 20-fold-higher fluorescence levels than S. gordonii incubated alone. These results suggest that signal concentration can increase in a closed system, where cell-cell contact is not required. Furthermore, in a flowing environment, where signal cannot accumulate, completion of a signaling event in biofilms between the signal-producer and signal-receiver requires cell-cell contact or a distance of only 1 or 2 km. Coaggregation was reported to be indispensable for efficient syntrophic anaerobic propionate oxidation by the partnership of Pelotomaculum thermopropionicum (which produces hydrogen from propionate oxidation) and Methanothermobacter thermautotrophicus (which consumes hydrogen to produce methane) (Ishii et al., 2005). Anaerobic oxidation of volatile fatty acids is energetically unfavorable unless the hydrogen partial pressure is kept low, for example, by utilization of hydrogen for production of methane. In a flow system called upflow anaerobic sludge blanket, the distance between the two species for efficient interspecies hydrogen transfer was calculated to be approximately 2 pm. Coaggregated mixed species in close physical contact were observed, again indicating the need for juxtaposition to facilitate efficient transfer of diffusible molecules in a flowing system. A model to examine the development of a four-species biofilm was developed by inoculating into a saliva-fed flow cell the human oral coaggregation partners S. gordonii, A. naeslundii, V; atypica, and Fusobacterium nucleatum (Foster and Kolenbrander, 2004). The biofilm was inoculated in two ways, sequential inoculation or coaggregate inoculation. Flow cells inoculated sequentially produced biofilms with larger biovolumes than flow cells inoculated with coaggregates. Each species in the biofilms was identified by fluorescence in situ hybridization using species-specificoligonucleotide probes. The majority of cells in both sequentially and coaggregateinoculated biofilms were S. gordonii, regardless of the inoculation order. However, the representation of A. naeslundii and V; atypica was significantly higher in biofilms inoculated with coaggregates than in sequentially inoculated biofilms, suggesting that a mutualistic interaction occurred. These results indicate that the development of multispecies biofilm communities is influenced by coaggregations preformed in planktonic phase. Several genes of S. gordonii are involved in the formation of human oral dual-species biofilms with Porphyromonas gingivalis (Kuboniwa et al., 2006). S. gordonii is an early colonizer of the enamel surface, whereas
P. gingivalis is a later colonizer. Of interest were the functions that regulated the recruitment of P. gingivalis by already adherent streptococcal biofilm cells. A plasmid integration library of S. gordonii was screened and revealed putative functions that included intercellular or intracellular signaling, cell wall integrity and maintenance of adhesive proteins, capsule biosynthesis, and overall bacterial physiology. Collectively, this broad functional assemblage suggests that numerous environmental cues govern the formation of these dual-species biofilms. Biofilms are formed at interfaces. Usually we consider that biofilms are formed only on solid or semisolid substrata, such as steel pipes or agar, respectively. An example of the latter is the formation of a fruiting body by myxobacteria, gliding and aggregating on agar. Biofilms are also formed at liquid-liquid interfaces, and an example is the biofilm at the interface between polychlorinated biphenyls (PCB) and water (Macedo et al., 2005). This model to study degradation of PCBs over 31 days revealed that multispecies biofilms formed on the surface of the PCB droplet and that the biofilm developed in three stages: (i) bacteria accumulated on the droplet during the first 10 days; (ii) bacterial diversity was maximal by 24 days, and bacterial aggregates formed pockets in the PCB; and (iii) invasion of the biofilm resulted in a pock-marked PCB droplet. The apparent cooperation of many species in the degradation of PCB illustrates a remarkable example of why, in nature, many biofilms are multispecies.
CONCLUSIONS Biofilm communities are composed of multiple species that interact in an integrated way. Often, the integration is essential for the hydrolysis of complex substrates, a process in which each species of the community participates with an outcome whereby the members of the community benefit. Some communities such as the initial colonizers of the human enamel are repetitive on a twicedaily basis after oral hygiene procedures. The potential for metabolic communication is obvious in consortia that convert sugars to lactic acid, a fermentable substrate for other community members, as well as in syntrophic propionate oxidation. Many environmental cues and signals likely govern multispecies biofilms that form and disperse. One of the most important determinants is the distance between signal generator and signal receiver. This determinant is especially significant in biofilms that occur in flowing systems where diffusible signals are washed out by bulk flow. The role of AI-2 as a bona fide interspecies signal is supported by its activity as a concentration-dependent signal in an oral dual-species
26. MULTISPECIES INTERACTIONSAND BIOFILMCOMMUNITIES community developed under conditions relevant to nature. AI-2 mediates two-way communication between certain vibrios and enteric bacteria, and this crosscommunication between species further demonstrates that AI-2 can be produced by one species and recognized by another. Many bacteria, including myxobacteria and their neighbors, rely on highly localized communication by direct cell-to-cellcontact or within a few micrometers. Most commonly, bacteria exist as multispecies biofilms formed on a variety of surfaces and responsive to a variety of environmental cues. This research was supported by the Intramural Research Program of the National Institute of Dental and Craniofacial Research of the National Institutes of Health.
References Ahn, S. J., Z. T. Wen, and R. A. Burne. 2006. Multilevel control of competence development and stress tolerance in Streptococcus mutans UA159. Infect. Immun. 74:1631-1642. An, D., T. Danhorn, C. Fuqua, and M. R. Parsek. 2006. Quorum sensing and motility mediate interactions between Pseudomonas aeruginosa and Agrobacterium tumefaciens in biofilm cocultures. Proc. Natl. Acad. Sci. USA 103:38283833. Bassler, B. L., M. Wright, R. E. Showalter, and M. R. Silverman. 1993. Intercellular signalling in Vibrio harveyi: sequence and function of genes regulating expression of luminescence. Mol. Microbiol. 9:773-786. Caiazza, N. C., R. M. Shanks, and G. A. O’Toole. 2005. Rhamnolipids modulate swarming motility patterns of Pseudomonus aeruginosa. J. Bacteriol. 187:7351-7361. Camilli, A., and B. L. Bassler. 2006. Bacterial small-molecule signaling pathways. Science 311:1113-1116. Chen, X., S. Schauder, N. Potier, A. Van Dorsselaer, I. Pelczer, B. L. Bassler, and F. M. Hughson. 2002. Structural identification of a bacterial quorum-sensing signal containing boron. Nature 415545-549. Cisar, J. O., A. L. Sandberg, G. P. Reddy, C. Abeygunawardana, and C. A. Bush. 1997. Structural and antigenic types of cell wall polysaccharides from viridans group streptococci with receptors for oral actinomyces and streptococcal lectins. Infect. Immun. 655035-5041. Davies, D. G., M. R.Parsek, J. P. Pearson, B. H. Iglewski, J. W. Costerton, and E. P. Greenberg. 1998. The involvement of cell-to-cell signals in the development of a bacterial biofilm. Science 280:295-298. Dudler, R., and L. Eberl. 2006. Interactions between bacteria and eukaryotes via small molecules. Curr. Opin. Biotechnol. 17:268-273. Duerre, J. A., D. J. Baker, and L. Salisbury. 1971. Structure elucidation of a carbohydrate derived from S-ribosylhomocysteine by enzymatic cleavage. Fed. Proc. 30:1067. Egland, P. G., R. J. Palmer, Jr., and P. E. Kolenbrander. 2004. Interspecies communication in Streptococcus gordoniiVeillonella atypica biofilms: signaling in flow conditions
459
requires juxtaposition. Proc. Natl. Acad. Sci. USA 101: 16917-16922. Foster, J. S., and P. E. Kolenbrander. 2004. Development of a multispecies oral bacterial community in a saliva-conditioned flow cell. Appl. Environ. Microbiol. 70:4340-4348. Fuqua, C., M. R. Parsek, and E. P. Greenberg. 2001. Regulation of gene expression by cell-to-cell communication: acyl-homoserine lactone quorum sensing. Annu. Rev. Genet. 35~439-468. Gilmore, K. S., P. Srinivas, D. R. Akins, K. L. Hatter, and M. S. Gilmore. 2003. Growth, development, and gene expression in a persistent Streptococcus gordonii biofilm. Infect. Immun. 71:4759-4766. Ishii, S., T. Kosaka, K. Hori, Y. Hotta, and K. Watanabe. 2005. Coaggregation facilitates interspecies hydrogen transfer between Pelotomaculum thermopropionicum and Methanothermobacter thermautotrophicus. Appl. Environ. Microbiol. 71:7838-7845. Itoh, Y., X. Wang, B. J. Hinnebusch, J. F. Preston 111, and T. Romeo. 2005. Depolymerization of beta-1,6-N-acetyl-Dglucosamine disrupts the integrity of diverse bacterial biofilms. 1. Bacteriol. 187:382-387. Johnson, M. R., C. I. Montero, S. B. Comers, K. R. Shockley, S. L. Bridger, and R. M. Kelly. 2005. Population densitydependent regulation of exopolysaccharide formation in the hyperthermophilic bacterium Thermotoga maritima. Mol. Microbiol. 55:664-674. Kaplan, J. B., C. Ragunath, N. Ramasubbu, and D. H. Fine. 2003. Detachment of Actinobacillus actinomycetemcomitans biofilm cells by an endogenous beta-hexosaminidase activity. J. Bacteriol. 185:4693-4698. Kaplan, J. B., K. Velliyagounder, C. Ragunath, H. Rohde, D. Mack, J. K. Knobloch, and N. Ramasubbu. 2004. Genes involved in the synthesis and degradation of matrix polysaccharide in Actinobacillus actinomycetemcomitans and Actinobacillus pleuropneumoniae biofilms. J. Bacteriol. 186~8213-8220. Keller, L., and M. G. Surette. 2006. Communication in bacteria: an ecological and evolutionary perspective. Nut. Rev. Micro biol. 4:249-25 8. Kirillina, O., J. D. Fetherston, A. G. Bobrov, J. Abney, and R. D. Perry. 2004. HmsP, a putative phosphodiesterase, and HmsT, a putative diguanylate cyclase, control Hmsdependent biofilm formation in Yersinia pestis. Mol. Microbiol. 54~75-88. Klier, C. M., P. E. Kolenbrander, A. G. Roble, M. L. Marco, S. Cross, and P. S. Handley. 1997. Identification of a 95 kDa putative adhesin from Actinomyces serovar WVA963 strain PK1259 that is distinct from type 2 fimbrial subunits. Microbiology 143935-846. Kolenbrander, P. E. 1988. Intergeneric coaggregation among human oral bacteria and ecology of dental plaque. Annu. Rev. Microbiol. 42:627-656. Kolenbrander, P. E., R. N. Andersen, D. S. Blehert, P. G. Egland, J. S. Foster, andR. J. Palmer, J . 2002. Communication among oral bacteria. Microbiol. Mol. Biol. Rev. 66:486-505. Kuboniwa, M., G. D. Tribble, C. E. James, A. 0. Kilic, L. Tao, M. C. Herzberg, S. Shizukuishi, and R. J. Lamont. 2006. Streptococcus gordonii utilizes several distinct gene
460 functions to recruit Porphyromonas gingivalis into a mixed community. Mol. Microbiol. 60:121-139. Kulasakara, H., V. Lee, A. Brencic, N. Liberati, J. Urbach, S. Miyata, D. G. Lee, A. N. Neely, M. Hyodo, Y. Hayakawa, F. M. Ausubel, and S. Lory. 2006. Analysis of Pseudomonus aeruginosa diguanylate cyclases and phosphodiesterases reveals a role for bis-(3’-5’)-cyclic-GMP in virulence. Proc. Natl. Acad. Sci. USA 103:2839-2844. Li, Y. H., N. Tang, M. B. Aspiras, P. C. Lau, J. H. Lee, R. P. Ellen, and D. G. Cvitkovitch. 2002. A quorum-sensing signaling system essential for genetic competence in Streptococcus mutans is involved in biofilm formation. 1. Bacteriol. 184:2699-2708. Loo, C. Y., D. A. Corliss, and N. Ganeshkumar. 2000. Streptococcus gordonii biofilm formation: identification of genes that code for biofilm phenotypes. J. Bacteriol. 182:13741382. Macedo, A. J., U. Kuhlicke, T. R. Neu, K. N. Timmis, and W. R. Abraham. 2005. Three stages of a biofilm community developing at the liquid-liquid interface between polychlorinated biphenyls and water. Appl. Environ. Microbiol. 71~7301-7309. Miller, S. T., K. B. Xavier, S. R. Campagna, M. E. Taga, M. F. Semmelhack, B. L. Bassler, and F. M. Hughson. 2004. Salmonella typhimurium recognizes a chemically distinct form of the bacterial quorum-sensing signal AI-2. Mol. Cell 15:677-687. Norskov-Lauritsen, N., and M. Kilian. 2006. Reclassification of Actinobacillus actinomycetemcomitans, Haemophilus aphrophilus, Haemophilus paraphrophilus and Haemophilus segnis as Aggregatibacter actinomycetemcomitans gen. nov., comb. nov., and Aggregatibacter aphrophilus comb. nov. and Aggregatibacter segnis comb. nov., and emended description of Aggregatibacter aphrophilus to include V factordependent and V factor-independent isolates. Int. J. Syst. Evol. Microbiol. 56:2135-2146. Nudleman, E., D. Wall, and D. Kaiser. 2005. Cell-to-cell transfer of bacterial outer membrane lipoproteins. Science 309~125-127. Nyvad, B., and M. Kilian. 1987. Microbiology of the early colonization of human enamel and root surfaces in vivo. Scand. J. Dent. Res. 95:369-380. Palmer, R. J., Jr., S. M. Gordon, J. 0. Cisar, and P. E. Kolenbrander. 2003. Coaggregation-mediated interactions of streptococci and actinomyces detected in initial human dental plaque. J. Bacteriol. 185:3400-3409. Palmer, R. J., Jr., K. Kazmerzak, M. C. Hansen, and P. E. Kolenbrander. 2001a. Mutualism versus independence: strategies of mixed-species oral biofilms in vitro using saliva as the sole nutrient source. Infect. Immun. 695794-5804. Palmer, R. J., Jr., R. Wu, S. Gordon, C. G. Bloomquist, W. F. Liljemark, M. Kilian, and P. E. Kolenbrander. 2001b. Retrieval of biofilms from the oral cavity. Methods Enzymol. 337393-403. Petersen, F. C., D. Pecharki, and A. A. Scheie. 2004. Biofilm mode of growth of Streptococcus intermedius favored by a competence-stimulating signaling peptide. J. Bacteriol. 186:6327-633 1. Ramasubbu, N., L. M. Thomas, C. Ragunath, and J. B. Kaplan. 2005. Structural analysis of dispersin B, a biofilm-releasing
ANALOGOUS SYSTEMS glycoside hydrolase from the periodontopathogen Actinobacillus actinomycetemcomitans. J. Mol. Biol. 349:475-486. Rickard, A. H., R. J. Palmer, Jr., D. S. Blehert, S. R. Campagna, M. F. Semmelhack, P. G. Egland, B. L. Bassler, and P. E. Kolenbrander. 2006. Autoinducer-2: a concentrationdependent signal for mutualistic bacterial biofilm growth. Mol. Microbiol. 60:1446-1456. Riedel, K., M. Hentzer, 0. Geisenberger, B. Huber, A. Steidle, H. Wu, N. Hoiby, M. Givskov, S. Molin, and L. Eberl. 2001. N-acylhomoserine-lactone-mediated communication between Pseudomonas aeruginosa and Burkholderia cepacia in mixed biofilms. Microbiology 147:3249-3262. Ross, P., R. Mayer, H. Weinhouse, D. Amikam, Y. Huggirat, M. Benziman, E. de Vroom, A. Fidder, P. de Paus, L. A. Sliedregt, G.A.vanderMare1, and J.H. vanBoom. 1990. The cyclic diguanylic acid regulatory system of cellulose synthesis in Acetobacter xylinum. Chemical synthesis and biological activity of cyclic nucleotide dimer, trimer, and phosphothioate derivatives. J. Biol. Chem. 265:18933-18943. Ross, P., H. Weinhouse, Y. Aloni, D. Michaeli, P. WeinbergerOhana, R. Mayer, W. Braun, E. de Vroom, G. A. van der Marel, J. H. van Boom, and M. Benziman. 1987. Regulation of cellulose synthesis in Acetobacter xylinum by cyclic diguanylic acid. Nature 325:279-281. Sauer, K., M. C. Cullen, A. H. Rickard, L. A. Zeef, D. G. Davies, and P. Gilbert. 2004. Characterization of nutrient-induced dispersion in Pseudomonas aeruginosa PA01 biofilm. J. Bacteriol. 186:73 12-7326. Schauder, S., K. Shokat, M. G. Surette, and B. L. Bassler. 2001. The LuxS family of bacterial autoinducers: biosynthesis of a novel quorum-sensing signal molecule. Mol. Microbiol. 41:463-476. Semmelhack, M. F., S. R. Campagna, M. J. Federle, and B. L. Bassler. 2005. An expeditious synthesis of DPD and boron binding studies. Org. Lett. 7569-572. Senadheera, M. D., C. Levesque, and D. G. Cvitkovitch. 2005. Cell-density-dependent regulation of streptococcal competence, p. 233-267. In D. R. Demuth and R. J. Lamont (ed.), Bacterial Cell-to-Cell Communication: Role in Virulence and Pathogenesis. Cambridge University Press, Cambridge, United Kingdom. Steidle, A., K. Sigl, R. Schuhegger, A. Ihring, M. Schmid, S. Gantner, M. Stoffels, K. Riedel, M. Givskov, A. Hartmann, C. Langebartels, and L. Eberl. 2001. Visualization of N-acylhomoserine lactone-mediated cell-cell communication between bacteria colonizing the tomato rhizosphere. Appl. Environ. Microbiol. 675761-5770. Stoodley, P., S. Wilson, L. Hall-Stoodley, J. D. Boyle, H. M. Lappin-Scott, and J. W. Costerton. 2001. Growth and detachment of cell clusters from mature mixed-species biofilms. Appl. Environ. Microbiol. 675608-5613. Tait, K., I. Joint, M. Daykin, D. L. Milton, P. Williams, and M. Camara. 2005. Disruption of quorum sensing in seawater abolishes attraction of zoospores of the green alga Ulva to bacterial biofilms. Environ. Microbiol. 7:229-240. Thormann, K. M., S. Duttler, R. M. Saville, M. Hyodo, S. Shukla, Y. Hayakawa, and A. M. Spormann. 2006. Control of formation and cellular detachment from Shewanella oneidensis MR-1 biofilms by cyclic di-GMP. J. Bacteriol. 188~2681-2691.
26. MULTISPECIES INTERACTIONS AND BIOFILMCOMMUNITIES Tischler, A. D., and A. Camilli. 2004. Cyclic diguanylate (c-di-GMP) regulates Vibrio cholerae biofilm formation. Mol. Microbiol. 53:857-869. Visick, K. L., and C. Fuqua. 2005. Decoding microbial chatter: cell-cell communication in bacteria. 1.Bacteriol. 18755075519. Wang, X., J. F. Preston 111, and T. Romeo. 2004. The pgaABCD locus of Escherichia coli promotes the synthesis
461
of a polysaccharide adhesin required for biofilm formation.
1.Bacteriol. 186:2724-2734.
Waters, C. M., and B. L. Bassler. 2005. Quorum sensing: cellto-cell communication in bacteria. Annu. Rev. Cell. Dev. Biol. 2 1:3 19-346. Xavier, K. B., and B. L. Bassler. 2005. Interference with AI2-mediated bacterial cell-cell communication. Nature 437~750-753.
Mvxobacterial
Myxobactevia: MuLticeLLularity and Differentiation Edited by David E. Whitworth 02008 ASM Press, Washington, D.C.
Penelope I. Higgs John I? Merlie, Jr.
Myxococcus xanthus: Cultivation, Motility, and Development
This chapter presents a description of methods for cultivation of Myxococcus xanthus laboratory strains, as well as specific protocols for analysis of motility and development. A concise set of methods is complicated by variations necessary for cultivation of different laboratory “wild-type’’ or parent strains as well as research groupspecific variations in protocols that inevitably arise. Here we attempt to cover representative techniques, and to mention variations only occasionally. As several variations in protocols arise from using different laboratory strains, we should address the lineages of these strains here. Most of the current literature references three laboratory strains that were obtained directly, or were derived, from the stock collection at the University of California at Berkeley (UCB). One of these strains, DZ2, was obtained directly from the teaching collection at UCB and was immediately stored as a frozen stock (Campos and Zusman, 1975; David Zusman, personal communication). A second strain, FB (aka DZFl and DKlOl), was also obtained from UCB but was subsequently selected as a dispersed growing mutant (Dworkin, 1962). The mutation rendering this behavior was later determined to be piZQl ,
27
which results in a partial defect in social motility and cell cohesiveness (Wall et al., 1999). Finally, the third and most common parent strain is DK1622 (Wireman and Dworkin, 1975) from which the genome sequence was determined (Goldman et al., 2006). The lineage of DK1622, outlined by Wall et al., is as follows: strain FB was UV mutagenized to generate the nonmotile DK320 (agZB2 piZQ2) strain. DK1622 was generated by restoring DK320 to wild-type motility by phage transduction from YS (aka MD2 and DK1600), which was a spontaneous revertant of FB that restored motility (Wall et al., 1999). Some phenotypic differences are observed between these strains. DK1622 develops approximately 24 h more quickly than DZ2, and DZ2 does not develop well on strict starvation medium in the absence of pyruvate. In addition, DK1622 is more brightly yellow colored than DZ2 and is considerably more cohesive. While it is unknown precisely how strains DZ2, FB, and DK1622 differ genetically, Chen et al. determined that, unlike FB and presumably DZ2, DK1622 is deleted for a -220-kbp fragment of genomic DNA expected to consist mostly of Mx alpha prophage repeats (Chen et al., 1990).
~~
Penelope I. Higgs, Department of Ecophysiology, Max Planck Institute for Terrestrial Microbiology, Marburg, Germany 35043. John P. Merlie, Jr., Department of Molecular and Cell Biology, University of California-Berkeley, Berkeley, CA 94720.
465
MYXO BACTERIAL METHOD s
466
CULTIVATION In nature, communities of M . xanthus reside in the soil on decaying plant material or herbivore dung and obtain nutrients by secreting digestive enzymes (such as proteases and lysozyme) to digest macromolecules from prey microorganisms or decaying organic matter (Reichenbach, 1999). Communities of M. xanthus are thought to exhibit cooperative growth (the growth rate of cultures increases in the presence of higher cell numbers), which is dependent on the amount of secreted enzymes that break down complex macromolecules for subsequent uptake by the individual cells (Rosenberg et al., 1977). General Considerations In the laboratory, M . xanthus vegetative cells can be cultivated in liquid broth or on agar plates and maintained for long-term storage as freezer stocks with minor variations on standard microbiology procedures. However, there are some special considerations for cultivating 111. xanthus. The vegetative cells are somewhat fragile and can be sensitive to water purity and the presence of residual detergents on glassware. In addition, M. xanthus is slightly light sensitive and should be grown in the dark or low-light conditions and the cells are also sensitive to cold and should not be stored at 4°C. Finally, M. xanthus cells undergo spontaneous phase variation from yellow to tan coloration. Tan colonies should be avoided since cells in this phase do not develop normally (Laue and Gill, 1994,1995). Equipment Standard laboratory equipment is sufficient for cultivation of M. xanthus. Plate cultures are grown in incubator ovens maintaining temperatures at 32°C and capable of housing large plastic containers with lids. Liquid cultures are grown in shaking platform incubators capable of housing flat-bottomed (Erlenmeyer) flasks, heating at 32”C, and shaking at 200 to 300 rpm. Cell density measurements can be determined by using a Klett-Summerson colorimeter with a red filter (100 Klett equals approximately 4 X l o x cells ml-l) (Bretscher and Kaiser, 1978) or with standard l-cm-path-length spectrophotometer reading absorbance at 500 nm (A55o)(0.7 Ass0 equals approximately 4 X l o* cells ml-l). Strains are stored in cryoprotectant (see “Storing Strains” below) for long term in a freezer capable of maintaining - 80°C. More-specialized equipment is required for analysis of M . xanthus behavior including, at minimum, a stereomicroscope, a light microscope with phase contrast and 20,40, and 1OOX objectives, both connected to a camera or, ideally, a computer-controlled charge-coupled device (CCD)camera. Some protocols listed in this chapter also
require a fluorescence microscope and image/motility analysis software. Media Media recipes are listed at the end of this chapter. Rich medium is comprised of an enzymatic digest of protein (Casitone) supplemented with magnesium and buffered to slightly alkaline conditions. Several recipes for rich media are routinely used depending on the origin of the parent laboratory strain. Strains from the DK1622 lineage are typically grown in CTT medium, while strains of the DZ2 lineage are typically grown in Casitone yeast extract (CYE)medium buffered withMOPS (morpholinepropanesulfonic acid). The principal differences are the presence of yeast extract and the buffer component (Tris versus MOPS). Cultures growing in rich liquid medium with good aeration can reach generation times of 3 to 4 h. Defined media were initially developed to define nutritional requirements of M . xanthus (Bretscher and Kaiser, 1978; Dworkin, 1962; Hemphill and Zahler, 1968; Witkin and Rosenberg, 1970) and are comprised of a subset of amino acids and other supplements. A1 minimal medium supports growth with a generation time between 22 and 30 h; the fastest growth was reported at 29°C and 200 rpm. The observation that A1 medium supports the growth of several Myxococcus strains isolated directly from the soil suggests that this medium is generally applicable to all M . xanthus strains. An amino acid-salts media supplemented with glycogen was developed (Dworkin, 1962) which supported growth of M . xanthus with a generation time of 10h ( 30”C, 75 to 100 rpm). Additional defined media have been developed and reported to yield generation times of 6.5 h ( M l ) (Witkin and Rosenberg, 1970) and 9 to 12 h (LM) (Hemphill and Zahler, 1968); however, these conditions were defined for mutant M . xanthus strains adapted to minimal media conditions and the recipes are not included here.
Protocols for Growth Plate Cultures M . xanthus long-term stock cultures (see “Storing Strains” below) are recovered onto rich-media plates. The following is a detailed protocol for reviving strains from freezer stocks and generating plate cultures for short-term storage of M . xanthus.
1. Prepare rich medium culture plates (see “media” below), containing antibiotics as necessary, at least 1 day prior to use. 2. Prewarm plates at 32°C for 20 min. Inverted plates are placed in an incubator with lids cracked open to remove excess moisture from the plate.
27. M. XANTHUS: CULTIVATION, MOTILITY, AND DEVELOPMENT 3. Using a sterile wooden stick or pipette tip, quickly remove a large scrape of the frozen cells from a freezer stock and streak cells onto the surface of rich media plates. Note: Generally we do not attempt to streak for isolated colonies since the cells swarm across the plate and growth is facilitated by high inocula. An insufficient starting inoculation may not yield growth. 4. Place plates agar side down inside a sealed plastic container and place the container in an incubator oven set at 32°C. Note: Plate cultures are incubated in sealed boxes to reduce dehydration of the agar. 5. When sufficient growth is detected (approximately 2 to 4 days), remove plates from the incubator and store at room temperature or 18°C in the dark. Note: Plates can be stored approximately 1week at room temperature. Cells should be passed (restreaked onto fresh plate) no more than twice to avoid accumulation of mutations.
To obtain colonies from single cells, diluted vegetative cells are dispersed into top agar (rich medium containing 0.3% agar, autoclaved, and cooled to 55”C), which is then overlaid on standard agar plates. Colonies will be visible after approximately 3 to 5 days of incubation at 32°C. Cells on the surface of the agar overlay will swarm across the media, but cells embedded in the agar overlay are prevented from swarming and can easily be picked and restreaked on new plates.
Broth Cultures Assays for M. xanthus behavior generally require that cells be harvested from vegetatively growing broth cultures. The following is a standard protocol for generating liquid cultures of M. xanthus in preparation for subsequent assays. 1. With a sterile wooden stick, loop, or glass hook, inoculate a swab of cells from a fresh plate culture into 10 ml of rich broth medium in a sterile 100-ml flat-bottomed flask. Disperse cells as much as possible into the broth by vigorously shaking the stick in the broth. 2. Incubate the broth at 32°C with shaking at 250 rpm in low light or dark overnight. 3. Determine cell density by measuring Klett or absorbance at 550 nm (A,,o). Cells are harvested at 4 X l o8 cells ml-’ (100 Klett or 0.7 A,,o). Subculture cells if necessary to allow growth to the desired density. 4. Harvest cells for subsequent assays by transferring the required volume of cells to a centrifuge tube. Pellet cells at 7,600 to 8,000 X g for 5 min at
467
room temperature. Note: We routinely centrifuge M. xanthus cells at 7,600 to 8,000 X g (at room temperature) to avoid very densely packed cells that are difficult to resuspend. 5 . Remove supernatant and resuspend cells in an equivalent volume of wash buffer or media. Pellet again as above. 6. Remove supernatant and resuspend cells in assay buffer to desired cell concentration. Note: A cell density of 4 X l o 9cells ml-l is often used for developmental assays described later in this chapter.
Storing Strains M. xanthus can be maintained long term either at - 80°C (freezer stocks) in the presence of a cryoprotectant (glycerol or dimethyl sulfoxide [DMSO]) or as starvationinduced fruiting body spores.
Freezer Stocks We have found that concentrating exponentially growing cells approximately fivefold before storage at - 80°C in the presence of the cryoprotectant DMSO allows for faster revival.
1. Grow cells to approximately 4 X lo8 cells mlk’ (roughly 18 h). 2. Pellet 10 ml of cells for 10 min at 7,600 X g a t room temperature and suspend in 2 ml of rich media. 4. Prepare two tubes for freezer stocks by adding 1 ml of concentrated cells to 0.25 ml of autoclaved DMSO in a sterile screw-cap tube. Freeze in liquid nitrogen (optional) and store at -80°C. Note: For heavily used strains, it is advantageous to scale up and prepare at least five freezer tubes of strains to avoid possible accumulation of genetic variations by repeated generation of new freezer stocks. Alternatively, cells can first be converted into chemically induced spores by addition of sterile DMSO to 0.5 M after step 2 and continued shaking incubation overnight. Spores are then resuspended in 5 ml of starvation buffer treated for freezer stocks as above. We have found chemically induced spore freezer stocks to be more stable than cell freezer stocks. However, it is necessary to first determine whether mutant strains are capable of forming chemically induced spores. Spore Stocks Dried stocks of fruiting bodies can be generated by scraping cells grown on rich media onto sterilized filter paper placed onto clone fruiting (CF) starvation media (see “Media” below) and incubating at 32°C for 7 days to induce fruiting body formation. The filter paper is then
MYXOBACTERIAL METHODS
468 transferred to cryogenic tubes and dried at room temperature in a decompression desiccator for 2 days. Strips may then be stored at - 80°C or at room temperature.
Kaiser, 1978). CF agar medium contains low levels of Casitone and pyruvate which do not support vegetative growth but likely promote a more gradual transition into development (Hagen et al., 1978).
Reviving Stocks Freezer stocks of M. xanthus are recovered as described in “Protocols for Growth,” “Plate Cultures” above. From dried spore stocks, a portion of the dried filter paper containing fruiting bodies is removed, placed onto rich media, and incubated at 32°C. Spores will germinate and swarm out onto the medium surface over the course of 3 to 4 days.
Protocols for Induction of Development For these procedures, strains are typically grown in rich broth at 32°C with shaking at 250 rpm. Wild-type and mutant strains should be analyzed concurrently and harvested from cultures grown to similar density (typically 4 x lo 8cells rn1-l).
Starvation Plates DEVELOPMENT Initiation of the developmental program resulting in fruiting body formation requires (i) nutrient limitation, (ii)a solid surface, and (iii) sufficient population density (reviewed in chapter 3). Under laboratory conditions, induction of the developmental program is initiated by harvesting vegetatively growing cells from a liquid culture, resuspending the cells to a given concentration, spotting them on a nutrient-limited medium, and incubating them at 32°C. Fruiting body morphology and timing are greatly dependent on the medium surface. Therefore, solid media for development should be prepared the day before and plates should be cured (prewarmed) by incubating the plates with lids cracked open at least 20 min at 32°C just before use. Mutants should always be analyzed in parallel with parent strains, since minor day-to-day variation in fruiting body morphology is often observed. In addition, it should be noted that with some mutants, developmental phenotypes vary depending on the type of starvation media and the cell density at which development is induced (Kashefi and Hartzell, 1995; Rasmussen et al., 2006). Developmental phenotypes are generally viewed by examining the spot of cells with a stereomicroscope and recorded by a connected camera. Fruiting body morphology can also be analyzed under a light microscope with 20 to 40X objectives; this approach can be facilitated by developing cells on microscope slides covered by a thin layer of developmental agar. Alternatively, cells in submerged culture can by analyzed by an inverted light microscope.
This assay is a typical quick method to analyze M. xanthus developmental phenotypes with respect to timing and morphology of fruiting body formation. When characterizing mutants, development should be analyzed on strict starvation (TPM or MMC) and CF plates (see “Media” below). Our best results are obtained when plates are prepared the day before and cured for 20 min at 32°C just before use. Vegetative cells are grown as indicated in “Broth Cultures” under “Protocols for Growth” above. 1. Harvest 1 ml of vegetative cells, wash in 1 ml of starvation buffer, and resuspend to 4 X l o 9 cells ml-1 in starvation buffer. Cells must be fully resuspended, and no clumps of cells should be visible. 2. Optional: Prepare a series of twofold dilutions in starvation buffer. Note: Some mutants display variable phenotypes depending on cell density. 3 . Spot 10 or 20 pJ of concentrated cells for each strain on cured developmental medium plate. Place spots at a maximal distance apart. Note: Especially for CF plates which contain low levels of nutrients, spotted cells will spread out significantly. Under these conditions, spot no more than 4 or 5 spots per plate. Do not touch the pipette tip to the medium. Duplicate plates should be prepared. 4. Allow spots to dry into media (-20 min), invert plates, and incubate in sealed containers at 32°C for 3 to 5 days. 5. Examine cell spots with a stereomicroscope and record pictures with an attached camera every 12 to 18 h for at least 3 days.
Submerged Culture
Starvation Medium Strict starvation medium, such as TPM, contains no additional carbodenergy source, although it should be noted that commercial agar is contaminated with proteins that can be utilized by M . xanthus for growth (Bretscher and
M. xanthus may be induced to develop in submerged culture assays by allowing cells to settle in tissue culture wells overlaid with starvation buffer. This assay is particularly useful when cells must be subsequently harvested for further analyses as it allows quick
27. M. XANTHUS: CULTIVATION, MOTILITY, AND DEVELOPMENT
469
and quantitative recovery. It should be noted that the developmental phenotype of many mutants may be strikingly different when analyzed in submerged culture compared to agar plates. In addition, strains selected for dispersed growth (e.g., FB) do not develop well in submerged culture. These protocols are based on submerged cultures grown in 24-well (15.6-mm-diameter) tissue culture plates but can be scaled up or down in proportion to the change in area of the culture well or dish.
3. Incubate at 32°C in humid chamber or parafilm plates to prevent evaporation. Cells will settle onto the bottom surface of the well and begin to develop. 4. Record pictures of phenotype and/or harvest cells by using a pipette to resuspend the film of cells and transfer to a sterile microcentrifuge tube; rinse the well with an additional 0.5 ml of starvation buffer and transfer the buffer to the same microcentrifuge tube.
Method of Kuner and Kaiser (1982) It is necessary to develop strains of the DZ2 background by this method.
Rapid Spores 211. xanthus cells exponentially growing in well-aerated rich liquid broth can be rapidly and synchronously induced to form spores by the addition of glycerol to the media at a final concentration of 0.5 M (Dworkin and Gibson, 1964). Glycerol spores have intermediate resistance to environmental stresses (Sudo and Dworkin, 1969) and are lacking the thick spore coat associated with fruiting body spores and several proteins associated with fruiting body spores (Inouye et al., 1979). Several additional chemicals including 0.7 M DMSO and certain alcohols will also induce spore formation, although the kinetics and properties of the spores may differ (Komano et al., 1980; Sadler and Dworkin, 1966).
1. Harvest vegetative cells (follow “Broth Cultures” above to step 3) and directly dilute to 2 X lo7 cells ml-’ in fresh rich medium. Note: Calculate the harvest volume needed as the number of wells needed for each strain multiplied by l o7 cells. Calculate the total volume needed as 0.5 ml per well. 2. Add 0.5 ml of diluted cells per 15.6-mm-diameter tissue culture well (24-well plate). Incubate at 32°C for 24 h. Cells will settle onto the bottom surface of the well and form a semiadherent layer. 3. Gently and completely remove rich media by aspiration. Option: Gently add and then remove 1 ml of starvation buffer to rinse cells. 4. Gently add 1 ml of starvation buffer (MMC or MC7) to each well. Note: Avoid disrupting the film of cells by slowly adding the starvation buffer to the side of the well. 5. Incubate at 32°C in humid chamber or parafilm plates to prevent evaporation. 6. Record pictures of phenotype and/or harvest cells by using a pipette to resuspend the film of cells and transfer to a sterile microcentrifuge tube; rinse the well with an additional 0.5 ml of starvation buffer and transfer the buffer to the same microcentrifuge tube.
Method of Ssgaard-Andersen et al. (1996) Some strains of the DZ2 background do not develop under this protocol; use the Kuner and Kaiser (1982) protocol above instead.
1. Harvest vegetative cells, wash in an equivalent volume of starvation buffer, and resuspend cells to 4 x l o9 cells ml-l in MC7 starvation buffer (see “Broth Cultures” above). 2. Mix 25 ~1 of the above suspension with 375 pl of MC7 and transfer to 15.6-mm-diameter tissue culture well (24-well plate).
1. Inoculate cells into 20 ml of rich medium in a 250-ml Erlenmeyer flask and grow to 4 X lo8cells ml-’ at 32°C with good aeration. 2. Add sterile glycerol to 0.5 M and continue to incubate with aeration at 32°C overnight. Note: Round refractile spores can be detected by 120 min, but spores continue to mature over the course of 8 h.
Assays for Development
Sporulation Assay Development in M. xanthus culminates in the formation of spore-filled fruiting bodies. The timing and efficiency of spore production is thus a useful assay for characterizing the net effect of mutations on the developmental pathway. Spores can be distinguished from vegetative cells by increased resistance to heat and sonic disruption as well as by a change in morphology from rods to spherical refractile bodies. Spore viability can be determined by counting colonies arising from plating spores dispersed in top agar over rich media plates. The timing of spore formation can be ascertained by isolating spores from cells harvested at intervals through the developmental program.
1. Develop cells in triplicate on starvation agar (see “Starvation Plates” above) or in submerged culture for 120 h (see “Submerged Cultures” above).
MYXO BACTERIAL METHOD s
470 2. Harvest cells into a microcentrifuge tube containing 0.5 ml of sterile water. Note: Spots of developing cells can be scraped off the agar surface with a flame-sterilized spatula and resuspended in water. Cells in submerged cultures can be transferred to microcentrifuge tubes, pelleted, and resuspended in water. 3. Heat for 60 min at 50°C. 4. Sonicate 30 s (output 3,30% duty Branson sonifier) with a microtip. 5 . Enumerate round, refractile spores in a hemacytometer (counting chamber) suitable for bacterial enumeration according to the manufacturer’s instructions. If necessary, dilute spores in water to obtain statistically relevant counts. 6. Calculate sporulation efficiency as a percentage of the wild type.
Option 1 7. Prepare three serial 100-fold dilutions of spores in sterile water. 8. Add 100 pl of undiluted spores to 3 ml of molten top agar at 50”C, vortex well, and pour over rich media plates. Repeat for each dilution. 9. Incubate at 32°C for at least 5 to 8 days and count colonies arising from single germinating spores. Option 2 7. Prepare eight serial 10-fold dilutions of spores in sterile water. 8. Spot 3 pl of each dilution on rich medium. Spot mutants alongside wild-type samples on a plate. 9. Incubate at 32°C. Growth is visible after 3 days as microcolonies arising from a single, germinating spore. 10. Calculate viable sporulation efficiency as a percentage of the wild type.
Extracellular Complementation Assays A hallmark of development in M. xanthus is the observation that certain nondeveloping mutants can be rescued for development by codevelopment with wild-type cells (Hagen et al., 1978). Five classes of mutants, termed B, A, D, E, and C, have been described that appear to be arrested in development at successive temporal positions (reviewed in Kaiser, 2004) as was determined by monitoring expression of developmental marker genes (Kroos et al., 1986). As such, the ability to rescue, or be rescued by, the five different classes of mutants can be employed as an assay to determine where in the developmental program a nonsporulating mutant exhibiting a developmental phenotype is blocked.
1. Harvest vegetative cells (see “Cultivation,” “Protocols for Growth” above) of the strain of interest bearing a selectable marker and of a strain bearing a mutation in one of the five complementation classes. Note: The assay can be performed to check for the ability to (i) complement or (ii) be complemented by a second strain. The selectable marker should be in a strain which will be analyzed for sporulation efficiency. 2. Wash pelleted cells in 1volume of starvation buffer and then suspend the washed pellet to 5 X l o 9cells ml-’ in starvation buffer. 3. Mix cells in a 1:l ratio and spot 20 pl of mixed cells and 20 pl of each single strain on CF plates; let spots dry into agar, invert plates, and incubate at 32°C. 4. Record pictures of phenotype through development. 5. Harvest and enumerate spores as outlined in “Sporulation Assay,” steps 1 through 6. 6. Determine the proportion of spores germinating from each strain by plating serial dilutions of spores in parallel on rich medium (for total spores) and rich medium containing antibiotic (to enumerate the strain carrying the marker). 7. Incubate at 32°C for at least 5 to 8 days and count colonies formed on plates with and without selection; determine the proportion of spores on antibiotic selection to determine rescue efficiency.
Molecular Analysis of Development In addition to the phenotypic analyses described here, it is clear that great progress has been made in understanding the molecular details of the developmental program in M . xanthus (reviewed elsewhere in this book). In particular, observed phenotypic defects can be analyzed at the molecular level by examining the expression of temporally regulated marker genes (chapter 28) and the synthesis, modification, and localization of marker proteins.
MOTILITY M . xanthus powers its movement over solid surfaces by using two genetically distinct motility systems (Hodgkin and Kaiser, 1979). These systems also produce distinct types of motile behavior on the part of myxobacterial cells. Social (S) motility-the coordinated movement of cells in large groups-predominates on soft and moist surfaces (Shi and Zusman, 1993) and is directly mediated by the extension and retraction of polar type IV pili (Sun et al., 2000; Wu and Kaiser, 1995). Adventurous (A) motility-the movement of single isolated cellspredominates on harder and drier surfaces (Shi and
27. M. XANTHUS: CULTIVATION, MOTILITY, AND DEVELOPMENT Zusman, 1993) and has been proposed to be based on either a jet engine-like extrusion of carbohydrate slime (Wolgemuth et al., 2002), a twisting or inching-like motion on the part of the M. xanthus cell (Wolgemuth and Oster, 2004), or intracellular motors pushing against dynamic focal adhesion points within the cell (Mignot et al., 2007). It has been shown that under typical lab conditions using 1.5% agar nutrient-rich plates that the two systems interact to produce a faster rate of colony expansion than would be expected from the sum of the expansion rates of colonies using each system independently (Kaiser and Crosby, 1983). The following protocols represent a small subset of the many available qualitative and quantitative methods of analyzing these two motility systems. We have organized them in order of increasing quantitative power from colony level spreading assays to single-cell assays designed to enumerate several different parameters of cell motility under various conditions.
Colony Spreading Protocols The protocols in this section are designed to qualitatively measure the rate and morphology of the expansion of M. xanthus colonies on agar surfaces. This type of assay is a fast and straightforward method of analyzing the proficiency of the two motility systems in a strain of interest. As stated above, S-motility tends to predominate on soft (0.3 to 0.5 %) agar surfaces while A-motility is best assayed on harder (1.5y0)agar surfaces (Shi et al., 1993).
A-Motility Colony Spreading Assay 1. Transfer 1 ml of vegetative culture at 4 x lo 8 cells ml-’ into a sterile microcentrifuge tube and spin at 7,600 X g for 10 min at room temperature. 2. Place the required number of 1.5% agar nutrientrich plates at 32°C to prewarm for 20 min. Note: Plates should not have any visible moisture on the surface of the agar after 20 min. 3. Remove the supernatant and resuspend the cell pellet in a buffered solution such as MMC or TPM to a density of 4 X lo9 cells ml-l. Note: Make sure that the pellet is completely resuspended with no cell aggregates of any size visible by eye. 4. Spot 10 pl of cells per strain on prewarmed plates containing 1.5% agar. We place 3 to 5 spots, at most, per agar plate. Note: Avoid touching the pipette tip to the agar as the spot is placed. 5. Let the plates sit right side up for 10 to 30 min or until all spots are completely dry. Note: Avoid disturbing the plates during this period, as this can cause the spots to run.
471
6. Measure the initial diameter of the newly dry spots either with a ruler or by microscopy. 7. Incubate the plates upside down at 32°C. 8. Take microscopic images and colony diameter measurements every 24 h for up to 5 days.
S-Motility Colony Spreading Assay Colony spreading assays favoring S-motility are performed as per A-motility assays (above),except that cells are spotted onto “soft” agar plates containing reduced agar levels. Levels of 0.3 to 0.5% agar work well for social motility assays. However, 0.5% agar is less subject to contamination and to shaking during high-magnification (40X objective) microscopy. Soft-agar plates should be prepared within 1 day of the assay and should be cured (agar side down) at room temperature overnight, or for at least 12 h. They are incubated agar side down.
A-/S- Motility Test The classic method for assigning genetic mutations to one of the two M. xanthus motility systems is to independently combine the mutation in question with known “reference” mutations in A- and S-motility and to then characterize the colony edges of the resulting doublemutant strains on hard (1.5%) agar nutrient-rich plates. A- strains show only large groups of cells at the colony edge under such conditions. S- strains show only single cells and no large groups. A mutation of interest that is combined with a known mutation in A- or S-motility either will show the same phenotype as the known mutation, if the mutation of interest belongs to the same motility system, or will show a completely nonmotile colony phenotype if it belongs to the other motility system. For example, a newly discovered S-motility mutation should produce a typical S-A’ phenotype when combined with a mutation in S-motility and should produce a completely nonmotile S-A- phenotype when combined with a mutation in A-motility.
Media Types and Effects on Motility Most nutrient-rich media used for growth of M. xanthus support both hard- and soft-agar swarming. Soft-agar swarming is more robust when nutrient concentrations are higher (Hillesland and Velicer, 2005). For example, cells show severalfold-greater expansion on soft-agar CYE or CTT plates than on soft-agar CF, MMC, or TPM plates. Also, of critical importance for the abovementioned colony expansion assays is the observation (Victor Bustamante, personal communication) that in the DZ2 strain background S-motility mutations will actually prevent the efficient expansion on hard agar of otherwise A+ colonies if nutrient levels are high enough.
472 For example, S- M. xanthus strains on 1.5% agar CYE or CTT plates display almost no colony expansion even over 5 days of incubation at 32°C. However, the same strain incubated on 1.5% agar 0.5X CTT plates over the same time period show nearly normal levels of expansion. The molecular basis of this nutrient effect is still being explored.
Single-Cell Assays Single-cell assays are designed to characterize and quantify the motile behavior of single cells. Such characterizations are useful for several applications: (i) confirming the assignment of a mutation to one of the two motility systems, (ii) determining the specific parameter or parameters of locomotion affected by such mutations, and (iii) quantifying the tactic behavior of cells. Methods have been devised for tracking single isolated A- or S-motile cells. In both cases, tracking is performed on a microscope slide at high magnification (20 to 40X objective) with time-lapse acquisition of images. Adventurous Motility There are several protocols for filming isolated cells moving via A-motility: filming cells on (i) 1.5% agar pads, (ii) 1.5% agar slabs, and (iii) 1.5% agar-filled gasket chambers. Pads are most useful for short-duration filming under condition of high magnification such as tracking of fluorescently labeled molecules during cell movement (Mignot et al., 2005). Slabs are thicker and more desiccation resistant and are more appropriate for longerduration, low-magnification movies intended to quantify cell motility parameters (Bustamante et al., 2004). Tracking of A-motile cells on 1.5% agar pads
1. Prepare the agar pads just before vegetatively growing cells are ready to harvest. 2. Flame sterilize two clean glass slides per straidcondition to be tested. Also prepare two slides wrapped in parafilm for each strainkondition. Note: Performing the following steps with the slides inside a petri dish will help to prevent desiccation of the agar. 3 . Transfer 750 pl of melted low-nutrient agar (CF or 0.5X CTT) to each slide and immediately cover each with a parafilm-wrapped slide. Note: Apply gentle pressure to the parafilm-wrapped slide as you place it on top of the agar layer. This will flatten and even out the agar. 4. Allow agar to solidify for 5 min. Carefully remove the parafilm-wrapped slide and, working quickly to prevent desiccation, cut away excess agar to
MYXOBACTERIAL METHODS leave a pad in the center of the slide with dimensions just smaller than a coverslip. 5. Immediately spot with 5 ~1 of 4 x l o 8 cells ml-' and cover with a coverslip. 6. Allow cells to sit on the agar at room temperature in the dark for 1 h before filming. 7. Film cells with 60 to 1OOX objective under oilimmersion, and under differential interference contrast or fluorescent illumination. Note: When selecting a region of the pad for filming, try to find cells that are moving well, that are well spaced, and that are free of culture debris. 8. Acquire images as necessary. Tracking of A-motile cells on 1.5% agar slabs
1. The day of the experiment, flame sterilize two clean glass slides per straidcondition to be tested. 2. Transfer 2 ml of melted low-nutrient agar (CF or 0.5 X CTT) to each slide and allow agar to solidify while covered for 1 h. 3. When a vegetatively grown culture reaches 4 X l o 8 cells ml-' in density, dilute in MMC or TPM to 4 x lo6 cells ml-l. Gently resuspend by pipetting up and down several times. 4. For each strain, transfer 2 1 0 - ~ volumes 1 of cells to two well-separated spots on each of two agar slabs and cover with a YSI brand oxygen-permeable membrane (YSI Incorporated, Model 5793). 5. Allow cells to sit on the agar at room temperature in the dark for 1 h before filming. 6 . Film cells with 20 to 40X objective under phasecontrast illumination. Note: When selecting a region of the slab for filming, try to find cells that are moving, in focus throughout the field of view, and well spaced from one another. 7. For the quantification of most motility parameters, acquire images every 15 s for 15 to 30 min. Social Motility (TFP-Dependent Movement) Because type IV pilus (TFP)-dependent movement requires cell-cell contact, it has traditionally been difficult to quantify the motile behavior of single cells using this motility system. Tracking on hard agar requires inactivation of the A-motility system in order to remove any confounding effects. Instead, cells can be tracked on soft agar, a surface on which the A-motor does not produce efficient motility. However, tracking cells on soft agar has been difficult in the past due to the issue of rapid water evaporation from the thin slabs used for filming. Both methods also suffer from the drawback that TFPdependent movement normally requires cell-cell contact
27. M. XANTHUS: CULTIVATION, MOTILITY, AND DEVELOPMENT in large groups; the presence of other cells in close proximity can make tracking difficult. Recently, Sun et al. (2000) devised a protocol for tracking the movement of single isolated cells on glass slides under a layer of methylcellulose solution. Through a mechanism that is not completely understood, methylcellulose dispenses with the requirement for cell-cell contact for TFP-dependent motility. For this reason, cell suspensions can be diluted to a very low density so that single isolated cells can be accurately tracked. A-motility does not appear to be highly active in this system. For this reason, cells can be tracked in either an A+ or Abackground and the results should be comparable. Methylcellulose assay for TFP-dependent motility
1. When a vegetatively grown culture reaches 4 X l o 8 cells ml-' in density, dilute in MMC or TPM to 6 X l o6 cells m1-I. Resuspend gently by pipetting up and down several times. Note: Be extremely gentle during cell resuspension, as high shear forces can remove TFP from the cell surface and significantly affect the motility phenotypes seen in the assay. 2. Flame sterilize two clean glass slides per strairdcondition to be tested. Note: Perform the following steps in a petri dish to avoid contamination. 3 . Place a sterile silicon gasket (Grace-Biolabs, JTR13R-0.5) in the center of each slide. 4. Transfer 10 pl of cells to the center of each gasket. Let cells sit on the slide for 10 min. 5. Overlay cells with 150 pl of 1%methylcellulose (4,000) containing 0.1 % Na-pyruvate. Cover gasket with a flame-sterilized coverslip. 6. Let cells sit in chambers at room temperature in the dark for 1 h before filming. 7. Acquire images every 15 s for 15 to 30 min with a 20 to 40X objective under phase-contrast illumination. Note: When selecting a region of the chamber for filming, try to find cells that are moving and well spaced.
Data Analysis In order to quantify the motility parameters of cells filmed using the above protocols, they must first be tracked. This can be done in two ways. First, the cells can be tracked by eye and their positions marked with pen on a transparency overlaid on the computer screen displaying the film of motile cells. The cells' tracks on the transparency can then be measured for cell movement distance, and this parameter can be used to calculate cell velocity. Cell reversals can be marked on these transparencies as they
473
occur and later be tabulated. Although inexpensive and straightforward, this method suffers from a relatively high level of inaccuracy and time commitment. We prefer to instead use computer software to track cell movement. Commercially available software, such as Metamorph, contains very efficient cell tracking programs that can quickly and accurately track the movement of many cells in a motility film. The resulting cell coordinate data can then be exported to a spreadsheet program such as Excel for the calculation of motility parameters. To automate the calculation of these parameters, we routinely use an Excel macro designed by Roy Welch and modified by Dave Astling (Astling et al., 2006). This macro takes the cell position data generated by the tracking program and calculates many parameters such as distance moved, velocity, number of cell pauses, length of cell pauses, and reversal frequency. This macro is available over the Internet from the Zusman Lab website http://www.mcb. berkeley.edu/labs/zusman/. Recently, Oleksii Sliusarenko has written a Matlabbased program that performs very similarly to Metamorph (Sliusarenko et al., 2006). This program contains a cell-tracking function that, like Metamorph, can export data to Excel. Alternatively, this program can independently calculate certain motility parameters such as distance moved and velocity, bypassing the need to use Excel.
Analysis of the Movement of Individual Cells in Groups The single-cell assays above are convenient for quantification of motility parameters. However, they all suffer from a common drawback. At best, they allow for an approximation of the behavior of 111. xanthus cells in their true physiological context: social groups. Even cells moving only by A-motility move predominantly in groups. Therefore, assays that track single isolated cells can never fully recapitulate the true physiological context of M. xanthus motility. To date, several methods have been developed for labeling single cells for the purpose of tracking them in larger groups. These include dye- and GFP-based labeling techniques. The dye-based protocols allow for relatively easy short-term labeling of cells. In contrast, the GFP-labeling protocol requires more preparatory effort but produces strains that are permanently marked by fluorescence. In this section we describe a dilution method that combines these labeling techniques with a gasket chamber technique developed by Rion Taylor and Roy Welch that allows one to quantify the behavior of single cells moving via S-motility as a part of large groups. Use of the gasket chamber technique is also appropriate for the
~
4 74 tracking of single cells moving in small A-motile groups or for the characterization of whole-group dynamics, such as colony expansion rates, via time-lapse filming of unlabeled cells.
GFP-Based Cell Tracking The GFP-labeling technique described here makes use of the construct created by Ellen Licking that fuses a copy of the piZA promoter to the coding sequence of GFP (pilAp-GFP) (Wall and Kaiser, 1998). By transforming M. xanthus with the chromosomal DNA of donor strains containing this construct, it is possible to produce cells that are sufficiently fluorescently labeled for tracking at low magnification (lox objective) in large groups. However, it has been reported that the original strain, created via insertion in the chromosome at the pilA locus, actually contained a high number of tandem repeats of this construct (Roy Welch, personal communication). When moving the pilAp-GFP construct from the original strain into a new M. xanthus strain, the number of repeats may change. Because the GFP fluorescence intensity is dependent on the number of repeats, care must be taken to check by microscopy the GFP fluorescence intensity of several different kanamycin-resistant clones that result from electroporation.
1. Prepare vegetative cultures of the strain of interest and of an isogenic strain carrying the pilApGFP construct. 2. Approximately 1h before the cultures reach 4 x l o 8 cells ml-l, begin preparing the gasket chambers. 3 . For each unlabeled/labeled strain pair, wrap two glass slides completely with parafilm. 4. Flame sterilize two 24- by 60-mm coverslips and using flame-sterilized forceps apply one coverslip to each parafilm-wrapped slide. The parafilm-wrapped slide acts as a backing to prevent damage to the thin coverslip during the following manipulations. 5. Using flame-sterilized forceps, apply one autoclaved silicon gasket (Grace-Biolabs, JTR13R0.5) to the center of each coverslip. Note: Use the end of the forceps to apply pressure all around the top surface of the gasket to ensure that it adheres strongly and evenly to the coverslip below. 6. Apply 300 p,l of melted 0.3% agar containing nutrient-rich media such as CTT or CYE to the interior of each gasket and immediately cover with a flame-sterilized glass slide. 7. Attach two mini-binder clips, one to the middle of each long side, to each gasket slide assembly. Incubate the gasket assemblies at 4°C for 20 min.
MYXOBACTERIAL METHODS 8. Prepare two additional flame-sterilized glass slides for each labeled/unlabeled strain pair. Apply one sterile silicon gasket to each slide. These will form the second half of the gasket chambers. 9. When the cultures reach 4 X l o xcells ml-l, pellet 1 ml of each culture at 7,600 x g for 10 min. 10. Resuspend the unlabeled strain in 100 ~1 of buffer (MMC or TPM) and the labeled isogenic strain in 1,000 ~1 of the same buffer. The unlabeled strain will now be at 4 X l o 9cells ml-', while the labeled strain will be at 4 x lo8 cells m1-l. 11. Prepare a further 10-fold dilution of the labeled strain in buffer to 4 X lo 7 cells mlt'. Note: During each resuspensiorddilution step, pipette up and down vigorously to ensure that all cells are completely resuspended and do not aggregate. 12. Mix the two strains by adding 2 pl of the 4 x lo7 cells ml-1 labeled strain suspension to 18 pl of the 4 X lo9 cells mlk' unlabeled strain suspension. The cell mixture should contain unlabeled cells and isogenic labeled cells in a 1,OOO:l ratio. 13. Retrieve the gasket chamber assemblies from 4°C and remove the mini-binder clips and the parafilm-wrapped backing slide. 14. Carefully slide one end of flame-sterilized forceps between the gasket and the remaining glass slide. While making sure to impart no levering or twisting forces, continue to slide the forceps arm further in between the two layers until the glass slide detaches from the rest of the gasket assembly. Note: This step can be subject to high rates of failure with the agar pad detaching from the coverslip when the sandwich is pried apart. The best results can be obtained by disassembling the gasket sandwich at 4°C after exactly 20 min of incubation. Still, it may be necessary to prepare twice as many gasket assemblies as actually needed. 15. Working quickly, resuspend the 1,OOO:l unlabe1ed:labeled cell mixture and apply 0.5 pl to the center of the agar pad inside the gasket. Note: Take care not to touch the pipette tip to the surface of the agar during this process. Also, avoid placing the spot of cells on or near any residual surface moisture on the agar. This will lead to spreading and dilution of the cells in the spot. 16. Immediately place the gaskedslide assembly on the gasket/coverslip assembly and press to form a seal between the two gaskets. Attach 1 minibinder clip to the middle of each long side of the assembly and let sit right side up for 10 min at room temperature.
27. M.XANTHUS: CULTIVATION, MOTILITY, AND DEVELOPMENT 17. Remove the binder clips and flip the assembly upside down and place on a slide warmer or other temperature-controlled flat surface at 32°C. Incubate for least 1 h before filming cells. 18. Place gasket assembly on the stage-mounted slide warmer of an upright microscope at 32°C with the coverslip side facing up. 19. Film cells with 1 0 x objective magnification, under phase-contrast or fluorescent illumination, using a CCD camera. Note: If filming only at certain points within a multiday incubation, keep the gasket assembly, coverslip side facing up, on the slide warmer at 32°C in the dark.
Dye-Based Cell Tracking Several protocols also exist for the transient labeling of myxobacterial cells with dyes. Both protocols allow for at least multihour filming with no discernible effect on cell viability or motility. One protocol listed here makes use of the membrane-binding fluorescent molecule CFDA SE (Molecular Probes V-12883). This dye appears to be significantly retained for at least 24 to 48 h after staining under either vegetative or developmental conditions (Rion Taylor and Oleksii Sliusarenko, personal communication). Because it is fluorescent, it is appropriate for tracking cells in very large groups and can be used as a substitute for GFP labeling in the gasket chamber protocol described above. It can also be used with any other protocol in this chapter in which movement in groups occurs. The other dye-based protocol listed here makes use of a tetrazolium dye that is efficiently internalized by live cells (Shi et al., 1996). This protocol provides useful levels of dye retention for at least 20 h after staining. Using bright-field microscopy, the tetrazolium-treated cells can be identified by the dark red color imparted by the dye. Because bright-field microscopy is used for illumination, the cells bearing this dye are not as easy to visualize when in the interior of large, multilayered groups (especially those seen in mature S-motile colonies).
Staining and tracking cells using (CFDA SE) carboxy-fluorescein diacetate, succinimidyl ester (Molecular Probes, V-12883) 1. Prepare vegetative cultures of the relevant strains. 2. When the culture reaches 4 X l o 8cells ml-l, transfer 1 ml of culture to a microcentrifuge tube and pellet cells at 7,600 X g for 10 min at room temperature. Return the rest of the culture to 32°C to use as unlabeled cells in step 6 below. 3 . Remove the supernatant and resuspend the cell pellet in 1 ml of phosphate-buffered saline.
475
4.Add 1 p,l of a 10 mM CFDA SE stock (in DMSO) to the cell suspension. Incubate the cells for 15 min at 32°C with constant agitation. 5. Pellet cells as above, remove the supernatant, and resuspend the cell pellet in 1 ml of PBS. Incubate the cells for 30 min at 32°C with constant agitation. 6. Prepare a mixture of the unlabe1ed:labeled cells at a 1OOO:l or 500:l ratio and proceed with any of the protocols listed here in which cells move in groups. 7. Film cells using fluorescent illumination, a fluorescein filter set, and a CCD camera.
Staining and tracking cells using 2,3,5triphenyltetrazolium 1. Prepare two vegetative cultures of the following strains: a. the strain of interest b. the strain of interest in the above medium containing 0.002% 2,3,5-triphenyltetrazolium chloride (Sigma, T8877) 2. When the cultures reach 2 x l o 8cells mlk', remove them from the shaker and let sit at room temperature for 3 h. 3. Prepare a mixture of the unlabe1ed:labeled cells at a 1000:l or 500:l ratio and proceed with any of the protocols listed here in which cells move in groups. 4. Film cells using bright-field illumination and a CCD camera. Note: With a color CCD camera, labeled cells will appear red. With a black and white CCD camera, labeled cells will appear darker than unlabeled cells.
Molecular Analysis of Motility Several assays have been developed for analysis of known S-motility components including the surface levels and retraction of TFP (Wu and Kaiser, 1997; Li et al., 2003; Mignot et al., 2005), extracellular matrix (Shimkets, 1986; Ramaswamy et al., 1997; Behmlander and Dworkin, 1991; Kearns et al., 2002), and lipopolysaccharide 0-antigen (Gill and Dworkin, 1986). Molecular assays for A-motility are limited, as the structural components for this mechanism of motility have not yet been conclusively identified.
MEDIA Media recipes are for broth culture. For solid medium, agar (Difco)is added to 1.5% (standard culture plates and hardagar motility assays), 0.5% (soft-agar motility assays), or 0.3% (top agar and soft-agar motility assays). Media are sterilized by autoclaving under standard conditions.
MYXOBACTERIAL METHODS
476
Rich Media CTT (Hodgkin and Kaiser, 1977) 1%Bacto Casitone (BD 225930) 10 mM Tris-HC1 (pH 7.6) 1 mM KH,PO,-KHPO, (pH 7.6) 8 mM MgSO, Adjust pH to 7.6.
0.1 mg ml-' djenkolic acid 1mg ml-I each L-Leu, L-Phe, L-Trp, L-Ala 0.6 mg ml-' -Tyr 0.5 mg ml-l each L -Asp, L-Ile, L-Pro 0.25 mg mlk' L-LYS 0.1 mg ml-I each L-Arg, L-Ser, L-Thr, L-Val 0.05 mg ml-' each L-His, L-G~Y, L-Met
Starvation Media CTT-YE CTT plus 0.5% yeast extract
0.SX CTT CTT with Casitone reduced to 0.5%
CYE (Campos and Zusman, 1975) 1%Bacto Casitone (BD 225930) 0.5% yeast extract 10 mM MOPS (pH 7.6) 4 mM MgSO,
Defined Media A1 Defined Minimal Medium (Bretscher and Kaiser, 1978) 10 mM Tris-HC1 (pH 7.6) 1 mM KH,PO,-KHPO, (pH 7.6) 8 mM MgSO, 0.5 mg ml-' (NH,)2S04 100 pg ml-I each L - A s ~L-Ile, , L-Phe, L-Val 50 pg ml-' L-Leu 10 pg ml-' L-Met 0.5% sodium pyruvate (from 25% sodium pyruvate pH 7.6 stock) 0.5% potassium aspartate (from 25% aspartic acid, adjusted to pH 7.6 with KOH) 10 p M FeC1, 10 pM CaCI,
Clone Fruiting (CF) Medium (Hagen et al., 1978) 10 mM Tris (pH 7.6)* 1mM KH,PO,-K,HPO, (pH 7.6)+ 8 mM MgSO, 0.02% (NH4)2SO, 2% sodium citrate 0.015% Casitone 1.5% agar '10 mM MOPS (pH 7.6) can be substituted. +lmM KH,PO, can be substituted. Autoclave and cool to 60°C. Add sodium pyruvate to 0.1%.
TPM 10 mM Tris-HC1 (pH 7.6) 1 mM K,HPO, 8 mM MgSO,
MMC Medium MMC salts (below) 1.5% agar Autoclave and cool to 60°C. Add sodium pyruvate to 0.1%.
MMC Salts 10 mM MOPS (pH 7.6) 4 mM MgSO, 2 mM CaC1,
Adjust pH to 7.6. For solid media, add agarose to 0.8%. Autoclave and cool to 50°C. Add as filter-sterilized solution: 1 pg ml-' vitamin BI2.
Filter sterilize.
Amino Acids-Salts Medium (Dworkin, 1962)
MC7 Buffer
10 mM KH,PO,-KHPO, (pH 7.6) 1 mg ml-' MgSO, 3 mg ml-' glycogen
10 mM MOPS (pH 7.0) 1mM CaC1, Filter sterilize.
27. M. X A N T H U S : CULTIVATION, MOTILITY, AND DEVELOPMENT References Astling, D. P., J. Y. Lee, and D. R. Zusman. 2006. Differential effects of chemoreceptor methylation-domain mutations on swarming and development in the social bacterium Myxococcus xanthus. Mol. Microbiol. 59:45-55. Behmlander, R. M., and M. Dworkin. 1991. Extracellular fibrils and contact-mediated cell interactions in Myxococcus xanthus. J. Bacteriol. 173:7810-7820. Bretscher, A. P., and D. Kaiser. 1978. Nutrition of Myxococcus xanthus, a fruiting myxobacterium. J. Bacteriol. 133: 763-768. Bustamante, V. H., I. Martinez-Flores, H. C. Vlamakis, and D. R. Zusman. 2004. Analysis of the Frz signal transduction system of Myxococcus xanthus shows the importance of the conserved C-terminal region of the cytoplasmic chemoreceptor FrzCD in sensing signals. Mol. Microbiol. 53: 1501-1513. Campos, J. M., and D. R. Zusman. 1975. Regulation of development in Myxococcus xanthus: effect of 3 ‘5 ’-cyclic AMP, ADP, and nutrition. Proc. Natl. Acad. Sci. USA 72: 518-522. Chen, H., I. M. Keseler, and L. J. Shimkets. 1990. Genome size of Myxococcus xanthus determined by pulsed-field gel electrophoresis. J . Bacteriol. 172:4206-4213. Dworkin, M. 1962. Nutritional requirements for vegetative growth of Myxococcus xanthus. J. Bacteriol. 84:250-257. Dworkin, M., and S. M. Gibson. 1964. A system for studying microbial morphogenesis: rapid formation of microcysts in Myxococcus xanthus. Science 146:243-244. Gill, J. S., and M. Dworkin. 1986. Cell surface antigens during submerged development of Myxococcus xanthus examined with monoclonal antibodies. J. Bacteriol. 168505-51 1. Goldman, B. S., W. C. Nierman, D. Kaiser, S. C. Slater, A. S. Durkin, J. A. Eisen, C. M. Ronning, W. B. Barbazuk, M. Blanchard, C. Field, C. Halling, G. Hinkle, 0. Iartchuk, H. S. Kim, C. Mackenzie, R. Madupu, N. Miller, A. Shvartsbeyn, S. A. Sullivan, M. Vaudin, R. Wiegand, and H. B. Kaplan. 2006. Evolution of sensory complexity recorded in a myxobacterial genome. Proc. Natl. Acad. Sci. USA 103:15200-15205. Hagen, D. C., A. l? Bretscher, and D. Kaiser. 1978. Synergism between morphogenetic mutants of Myxococcus xanthus. Dev. Biol. 64:284-296. Hemphill, H. E., and S. A. Zahler. 1968. Nutrition of Myxococcus xanthus FBa and some of its auxotrophic mutants. J. Bacteriol. 95:1011-1017. Hillesland, K. L., and G. J. Velicer. 2005. Resource level affects relative performance of the two motility systems of Myxococcus xanthus. Microb. Ecol. 495.58-566. Hodgkin, J., and D. Kaiser. 1977. Cell-to-cell stimulation of movement in nonmotile mutants of Myxococcus. Proc. Natl. Acad. Sci. U S A 74:2938-2942. Hodgkin, J., and A. D. Kaiser. 1979. Genetics of glidingmotility in Myxococcus xanthus (Myxobacterales): two gene systems control movement. Mol. Genet. Genomics 171:171-191. Inouye, M., S. Inouye, and D. R. Zusman. 1979. Biosynthesis and self-assembly of protein S, a development-specific protein of Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 76:209-213.
477
Kaiser, A. D., and C. Crosby. 1983. Cell movement and its coordination in swarms of Myxococcus xanthus. Cell Motil. Cytoskeleton 3:227-245. Kaiser, D. 2004. Signaling in myxobacteria. Annu. Rev. Microbiol. 58:75-98. Kashefi, K., and I? L. Hartzell. 1995. Genetic suppression and phenotypic masking of a Myxococcus xanthus frzF-defect. Mol. Microbiol. 15:483-494. Kearns, D. B., P. J. Bonner, D. R. Smith, and L. J. Shimkets. 2002. An extracellular matrix-associated zinc metalloprotease is required for dilauroyl phosphatidylethanolamine chemotactic excitation in Myxococcus xanthus. J. Bacteriol. 184~1678-1684. Komano, T., S. Inouye, and M. Inouye. 1980. Patterns of protein production in Myxococcus xanthus during spore formation induced by glycerol, dimethyl sulfoxide, and phenethyl alcohol. J. Bacteriol. 144:1076-1082. Kroos, L., A. Kuspa, and D. Kaiser. 1986. A global analysis of developmentally regulated genes in Myxococcus xanthus. Dev. Biol. 117:252-266. Kuner, J. M., and D. Kaiser. 1982. Fruiting body morphogenesis in submerged cultures of Myxococcus xanthus. J. Bacterial. 151:458-461. Laue, B. E., and R. E. Gill. 1994. Use of a phase variation-specific promoter of Myxococcus xanthus in a strategy for isolating a phase-locked mutant. J. Bacteriol. 1765341-5349. Laue, B. E., and R. E. Gill. 1995. Using a phase-locked mutant of Myxococcus xanthus to study the role of phase variation in development. J . Bacteriol. 177:4089-4096. Li, Y., H. Sun, X. Ma, A. Lu, R. Lux, D. Zusman, and W. Shi. 2003. Extracellular polysaccharides mediate pilus retraction during social motility of Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 1005443-5448. Mignot, T., J. P. Merlie, Jr., and D. R. Zusman. 2005. Regulated pole-to-pole oscillations of a bacterial gliding motility protein. Science 3105355-857. Mignot, T., J. W. Shaevitz, P. L. Hartzell, and D. R. Zusman. 2007. Evidence that focal adhesion complexes power bacterial gliding motility. Science 315:853-856. Ramaswamy, S., M. Dworkin, and J. Downard. 1997. Identification and characterization of Myxococcus xanthus mutants deficient in calcofluor white binding. J. Bacteriol. 179:2863-2871. Rasmussen, A. A., S. Wegener-Feldbrugge, S. L. Porter, J. I? Armitage, and L. Ssgaard-Andersen. 2006. Four signalling domains in the hybrid histidine protein kinase RodK of Myxococcus xanthus are required for activity. Mol. Microbiol. 60525-534. Reichenbach, H. 1999. The ecology of the myxobacteria. Enviyon. Microbiol. 1:15-21. Rosenberg, E., K. H. Keller, and M. Dworkin. 1977. Cell density-dependent growth of Myxococcus xanthus on casein. J. Bacteriol. 129:770-777. Sadler, W., and M. Dworkin. 1966. Induction of cellular morphogenesis in Myxococcus xanthus. 11. Macromolecular synthesis and mechanism of inducer action. J. Bacteriol. 91~1.520-1525. Shi, W., T. Kohler, and D. R. Zusman. 1993. Chemotaxis plays a role in the social behaviour of Myxococcus xanthus. Mol. Microbiol. 9:601-611.
4 78 Shi, W., and D. R. Zusman. 1993. The two motility systems of Myxococcus xanthus show different selective advantages on various surfaces. Proc. Natl. Acad. Sci. USA 90:33783382. Shi, W., F. K. Ngok, and D. R. Zusman. 1996. Cell density regulates cellular reversal frequency in Myxococcus xantbus. Proc. Natl. Acad. Sci. USA 93:4142-4146. Shimkets, L. J. 1986. Correlation of energy-dependent cell cohesion with social motility in Myxococcus xanthus. J. Bacteriol. 166:837-841. Sliusarenko, O., J. Neu, D. R. Zusman, and G. Oster. 2006. Accordion waves in Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 103:1534-1539. Ssgaard-Andersen, L., F. J. Slack, H. Kimsey, and D. Kaiser. 1996. Intercellular C-signaling in Myxococcus xantbus involves a branched signal transduction pathway. Genes Dev. 10:740-754. Sudo, S. Z., and M. Dworkin. 1969. Resistance of vegetative cells and microcysts of Myxococcus xantbus. J. Bacteriol. 98~883-887. Sun, H., D. R. Zusman, and W. Shi. 2000. Type IV pilus of Myxococcus xanthus is a motility apparatus controlled by the frz chemosensory system. Curr. Biol. 10:11431146.
MYXO BACTERIAL METH oD s Wall, D., and D. Kaiser. 1998. Alignment enhances the cell-tocell transfer of pilus phenotype. Proc. Natl. Acad. Sci. USA 95:3054-3058. Wall, D., I?. E. Kolenbrander, and D. Kaiser. 1999. The Myxococcus xantbus pilQ (sglA)gene encodes a secretin homolog required for type IV pilus biogenesis, social motility, and development. J. Bacteriol. 181:24-33. Wireman, J. W., and M. Dworkin. 1975. Morphogenesis and developmental interactions in myxobacteria. Science 189:s16-523. Witkin, S. S., and E. Rosenberg. 1970. Induction of morphogenesis by methionine starvation in Myxococcus xanthus: polyamine control. J. Bacteriol. 103:641-649. Wolgemuth, C., E. Hoiczyk, D. Kaiser, and G. Oster. 2002. How myxobacteria glide. Curr. Biol. 12:369-377. Wolgemuth, C . W., and G. Oster. 2004. The junctional pore complex and the propulsion of bacterial cells. J. Mol. Microbiol. Biotechnol. 7:72-77. Wu, S. S., and D. Kaiser. 1995. Genetic and functional evidence that Type IV pili are required for social gliding motility in Myxococcus xanthus. Mol. Microbiol. 18547-558. Wu, S. S., and D. Kaiser. 1997. Regulation of expression of the pilA gene in Myxococcus xanthus. J. Bacteriol. 179: 7748-7758.
Myxobacteria: Multicellularity and Differentiation Edited by David E. Whitworth 0 2008 ASM Press, Washington, D.C.
Frank-Dietrich Miiller Jimmy Schouv Jakobsen
MVXOCOCCUS xanthus:
28
Expression Analysis
Analysis of gene expression is an important tool for the study of the program involved in development and sporulation of Myxococcus xanthus. A large number of genes have been fused to the P-galactosidase (la&) gene, and their expression profiles have been extensively studied using P-galactosidase assays (Kroos et al., 1986).Recently, two new methods have become available, namely, quantitative PCR (QPCR) and microarray analysis. Each of the three methods presented in this chapter has advantages and disadvantages. The P-galactosidase assay can be used to study the expression of only one gene at a time. But the sequence of this gene does not need to be known, since the lacZ fusion mostly is made by transposons that more or less randomly jump into DNA sequences. Therefore, the P-galactosidase assay has been the “gold standard” of expression studies of M. xanthus development until now. It will probably become less important in the future now that the complete genome sequence is available. lacZ fusions have also been made directly, but we think that this approach will become very rare with QPCR becoming more available. The two other methods require knowledge of the sequence. The QPCR method is feasible only for studying
the expression of a limited number of genes (10 to 20 genes), whereas the strength of microarrays is their ability to monitor global gene expression. Microarrays are not suitable or cost-effective for studying gene expression of single genes, but they provide trends of global changes. Since microarray experiments require large amounts of RNA (100 to 200 pg of high-quality RNA per sample), the logistics of growing and developing cells easily becomes a project in its own right. Since QPCR requires much smaller RNA amounts, this task becomes much easier. The amount of cells needed for the P-galactosidase assay is much smaller yet, and a whole developmental time course can be carried out in a few petri dishes or one microtiter plate. So, in short one can say that P-galactosidase assays are easy but can address the expression of only one gene at the time. QPCR is more time-consuming but makes it possible to analyze a number of genes at the same time. Microarrays are very time-consuming and expensive, but in the best cases, they can reduce the time that it takes to determine the consequences of changing conditions or deleting genes severely.
Frank-Dietrich Miiller, Department for Ecophysiology, Max Planck Institute for Terrestrial Microbiology, Karl-von-Frisch-Stra13e, D-35043 Marburg, Germany. Jimmy Schouv Jakobsen, 194 Chemin du Siege, Residence le Mirabaou, F-06140 Vence, France.
4 79
MYXOBACTERIAL METHOD s
480
Here we provide protocols for these three types of analysis. For QPCR and microarray experiments the RNA must be of good quality; it should therefore be carefully checked for purity and integrity before use. The QPCR protocol is a scaled-down version of the two-step kit from Applied Biosystems using SYBR Green. The microarray protocol is adapted from the standard protocol used in the Joseph DeRisi lab (UCSF) for probe preparation and from the Standard Operating Procedure for Microbial DNA Probe Hybridization from The Institute for Genomic Research (TIGR) on amino-silane-coated slides. Until recently, poly-L-lysinecoated slides covering 80% of the M . xanthus genes were available from the Stanford Functional Genomics Facility (Stanford, CA). Here, PCR products specific for M . xanthus genes are spotted and UV-cross-linked on the slide surface. These microarrays have been used successfully in several labs to determine expression levels of M . xanthus genes after certain treatments (Jakobsen et al., 2004; Diodati et al., 2006; Pham et al., 2006; Overgaard et al., 2006). However, these microarray slides have been discontinued. Therefore, we focus in this chapter on the hybridization protocol for amino silane-coated arrays obtained from TIGR (Bethesda, MD). The protocols should be used as starting points for these types of experiments, and some optimization will, especially for microarray experiments, be necessary. Furthermore, since the protocols are exactly as they work in our lab we have chosen to include which materials and kits we use. Many other materials, be it enzymes or kits, can be used, and we recommend testing better or cheaper alternatives once the methods are set up.
P-GALACTOSIDASE ASSAY FOR M. XANTHUS DEVELOPED IN SUBMERGED CULTURE OR ON AGAR Materials Z buffer 60 mM Na2HP04 40 mM NaH,PO, 10 mM KCl 1 mM MgSO, 50 mM P-mercaptoethanol (can be omitted) Adjust pH to 7.0 with 10 M NaOH
ONPG (o-nitrophenyl-P-D-galactopyranoside) 1 mg/ml in Z buffer (dissolve 100 mg ONPG in 1 ml dimethyl sulfoxide [DMSO] by vigorous agitation, bring up to volume [lo0 ml] with Z buffer). Store in the dark at 4°C.
Procedure 1. Submerged culture (SC) development is carried out with 25 or 40 pl of Klettlooocells (optical density at 550 nm of 7) in 400 pl of MC7 buffer (see chapter 27) in 24-well plates at 32°C. Freeze plates at each time point. Note: Alternatively, develop cells on TPM agar (plate) and scrape into 400 p1 of TPM buffer, freeze at -20°C. 2. Add 0.6 ml of MC7 or TPM buffer (see chapter 27) to each sample well to bring volume up to 1 ml for sonication. Sonicate each sample for 10 s with a microtip sonicator. Transfer to Eppendorf tubes and sonicate for 1.5 min in a cup sonicator (setting 10) or sonicate each sample three times for 10 s with microtip. Note: For plate: Sonicate each sample three times for 10 s with microtip sonicator or sonicate 3 min in cup sonicator. 3. Set up assays in glass tubes: 100 to 250 pl of sample (depending on expression level). Add Z-buffer with 1mg/ml ONPG to a total volume of 500 yl. Add 50 p1 of 0.1% sodium dodecyl sulfate (SDS). Optional: Add 50 pl of chloroform. Vortex and incubate tubes at 37°C and start timing. When samples have turned sufficiently yellow, the reaction is stopped by addition of 0.5 ml 1 M Na,C03. Record the total time of reaction. 4. Measure A420 of each sample versus a blank; ideally the reading (yellow) should be 0.6 to 0.9. Use a reaction mixture without ONPG in the Z buffer as the blank. 5. Determination of protein concentration: Take 100 pl of each sample and determine total protein concentration using the Bradford assay. Use immunoglobulin G to generate a standard curve. Make the following standard reactions: 0, 2.5, 5.0, 7.5, 10.0, 12.5, 15.0, 17.5, and 20.0 pg of immunoglobulin G per ml. Vortex each sample with Bradford reagent in a total volume of 1 ml, leave at room temperature for 10 min, and measure A,,,. The sample without protein can be used as a reference. Prepare protein standard curve. 6. Calculate P-galactosidase activity per milligram of total protein as: (213 X A,,,)/[(ml)(mg protein/ml)(min)] where ml is the volume of supernatant used in the assay, “mg proteirdml” is the protein concentration in the sample, and min is the number of minutes the 0-galactosidase assay was incubated before stopping the reaction.
28. M .
XANTHUS:
EXPRESSION ANALYSIS
PROTOCOL FOR QPCR Hot Phenol RNA Isolation This protocol refers to the two-step kit from Applied Biosystems; a number of suppliers have similar kits, and these can be used instead. Here we use 2 pg of total RNA per reverse transcription reaction, but good results can be achieved with as little as 0.5 pg. Note: This protocol describes small-scale RNA isolation. For large microarray experiments (e.g., time course experiments or several technical replicates), an upscaled version of this protocol is provided in the microarray section.
Materials Water bath or heating block at 65°C Phenol saturated in 0.01 to 0.03 M sodium acetate (NaAc) pH 4.5 containing 0.1% 8-hydroxyquinoline (antioxidant). Preheat aliquots of phenol in a water bath; use gloves and a fume hood! Solution 1: 0.3 M sucrose; 0.01 M NaAc, pH 4.5 Solution 2: 2% SDS; 0.01 M NaAc, pH 4.5; preheat in water bath at 65°C RNase-free H20 Liquid nitrogen Cooled centrifuge (4°C) Speed vacuum centrifuge RNase-free tubes Procedure
1. Cell lysis: resuspend frozen cell pellet in 300 PI of ice-cold solution 1 and transfer into a 1.5-ml microcentrifuge tube containing 300 ~1 of solution 2. Proceed immediately with phenol extraction. 2. Add 400 pl of phenol, mix, and incubate >3 min. at 65°C. Note: When handling multiple samples, these steps are repeated until all tubes are collected in the heating block. 3. Snap-freeze in liquid nitrogen for 15 s and centrifuge at maximum speed for 5 min. 4. Transfer the aqueous top layer to a fresh 1.5-ml microcentrifuge tube containing 600 pl of hot phenol. Note: It is important to avoid the organic phase in all steps. 5. Mix and incubate for 3 min at 65°C. Snap-freeze in liquid nitrogen for 15 s and centrifuge at maximum speed for 5 min. 6. Phenollchloroform extraction: Transfer the aqueous layer to a fresh 1.5-ml microcentrifuge tube containing 300 p1 of phenol and 300 pl of chloroform. Mix and centrifuge.
48 1
7. Chloroform extraction: Transfer the aqueous layer to a fresh 1.5-ml microcentrifuge tube containing 600 p1 of chloroform. Mix and centrifuge. 8. Ethanol precipitation: Transfer the aqueous layer to a fresh 1.5-ml microcentrifuge tube containing 40 pl of 3 of M NaAc (pH 4.5) + 900 11.1of 96% ethanol and incubate at -20" C for 15 min (or overnight if required). 9. Centrifuge at 20,000 x g for 20 min at 4°C. 10. Carefully remove the supernatant and wash the pellet with 200 ~l of ice-cold 70% ethanol. Centrifuge at 20,000 X g for 5 min at 4°C. 11. Carefully remove the supernatant and dry the pellet for a maximum of 5 min in a speed vacuum centrifuge. Note: Do not let the pellet dry completely, as a dry pellet will be difficult to dissolve in water. 12. Resuspend pellet in 50 pl of RNase-free H20. Spin down at 20,000 X g for a few seconds and transfer to a new tube. 13. Quuntitution: Take 2 pl of RNA sample and add to 1 ml of H20. Vortex and measure absorbance at 260 nm in a quartz cuvette. Concentration (pg/pI)= A,, X 20. Purity: Take 2 pl of RNA sample and add to 1 ml of 10 mM Tris-HC1, pH 7.5. Measure absorbance at 260 and 280 nm to determine A2,dA2so ratio. Good values are 1.8 to 2.1 for pure RNA. Integrity: Run an RNA sample on a 1 % denaturing agarose gel. Note: Two distinct bands of 16s and 23s rRNA confirm that your RNA did not suffer major degradation during preparation. However, degraded RNA will appear as a smear of smaller-sized RNAs. RNA samples are stored at - 80°C. Reverse Transcription Here we describe the cDNA synthesis by reverse transcription according to the cDNA Archive KIT (ABI) by an example of a reaction using 2 pg of total RNA. Materials
lox reverse transcription buffer. . . . . . . . . 25X dNTP.. ....................... 10X random primers . . . . . . . . . . . . . . . . . 2 pg of total RNA . . . . . . . . . . . . . . . . . . .
10 pl 4 Fl 10 pl 10 pl
MultiScribe Reverse Transcriptase 50U/pl ......................... 54 Nuclease-free water . . . . . . . . . . . . . . . . . . 61 pl Total
.............................
1OOpl
MYXOBACTERIAL METHODS
482 Procedure Incubate the reaction mix for 10 min at 25°C and then incubate for 120 min at 37°C. cDNA Synthesis QPCRs Using the SYBR GREEN PCR Master Mix (ABI) Perform the reaction in a 25-pl volume to reduce the amount of SYBR GREEN PCR Master Mix needed. Reactions should be done in triplicate and with different cDNA dilutions (start with l : l O , 1:100,1:1,000, and 1:10,000 if cDNA was synthesized from 2 pg of total RNA). Primer concentration must be optimized to avoid the formation of primer dimers; in general a final concentration of 100 nM for each primer works. Here we describe a single QPCR by an example. Prepare a Master Mix for all the reactions using the same primer pair. Materials SYBR GREEN PCR Master Mix 25X . . . 12.5 pl Primer l ( 1 0 pM). . . . . . . . . . . . . . . . . . . 0 . 2 5 ~ 1 Primer 2 (10 pM). . . . . . . . . . . . . . . . . . . 0.25 pl cDNA dilutions .................... 1Pl Nuclease-free water . . . . . . . . . . . . . . . . . 11pl Total ............................ 25p1 Procedure Run QPCR with the standard protocol: 10 min at 95”C, 15 s at 95”C, and 1 min at 60”C, for a total of 35 cycles. Note: Include a dissociation curve in order to analyze the nature of the synthesized product (single product, primer dimer, etc.). Determine C,values following the instructions for your instrument and calculate relative amounts of mRNA.
PROTOCOL FOR MICROARRAY EXPERIMENTS Microarray experiments need a careful design. This is not only because of the cost but also because of the time required. Moreover, reproducibility and correlation of results highly depend on standardized procedures and thorough work. Usually, one needs at least three independent biological replicate experiments to obtain statistically significant data. One of the best sources for an overview and for guidance on troubleshooting of microarray experiments is “ D N A Microarrays” (Cold Spring Harbor Press) (Bowtell and Sambrook, 2002).
Hot Phenol RNA Isolation Note: This protocol is an upscaled version of the Hot Phenol RNA isolation protocol for QPCR (described above) but includes some additional hints for processing larger samples. All steps are carried out in 50- or 15ml RNase-free plastic tubes. It is important to do cell lysis and first hot phenol extraction as fast as possible to avoid RNA degradation. Always wear gloves and use dedicated pipette tips and tubes. To protect yourself from toxic phenol fumes, use a fume hood. Materials Water bath at 65°C 4°C centrifuge for 50- and 15-ml conical plastic tubes Speed vacuum centrifuge Liquid nitrogen RNase-free tubes (15 ml) RNase-free H,O Stop solution: 5 % saturated phenol, pH <7, in ethanol Solution 1:0.3 M sucrose; 0.01 M NaAc, pH 4.5 Solution 2: 2% SDS; 0.01 M NaAc, pH 4.5 Phenol saturatedin 0.01 to 0.03 MNaAc, pH4.5, containing 0.1 % 8-hydroxyquinoline (antioxidant). Alternatively: saturated phenol (pH 6.6) (Ambion, catalog no. 9712) Acid phenol-chloroform-isoamyl alcohol (25:24:1, pH 4.5) Chloroform-isoamyl alcohol (24:1) 3 M NaAc (pH 4.5) Ethanol 96% Ethanol 75% Protocol The following protocol describes a total RNA extraction from 50 ml of vegetatively grown cells. Prepare five 15-ml tubes for every sample, containing:
2.5 ml of Solution 2, preheat at 65°C (water bath) 5 ml of phenol, preheat at 65°C (water bath) 5 ml of pheno1:chloroform:isoamyl alcohol 2.524: 1, pH 6.6 5 ml of ch1oroform:isoamyl alcohol 24:l 9 ml of ethyl alcohol EtOH + 400 pl of 3 M NaAc (pH 4.5) (freshly made) Additionally, prepare 5 ml of hot phenol for each sample and preheat at 65°C in water bath for first phenol extraction step. Prepare at least 10 ml of 75% EtOH per sample for washing the pellet and store at -20°C. Procedure
1. Harvesting cells: Prepare a 50-ml conical centrifugation tube containing 5 ml of ice-cold stop solution.
28. 211. XANTHUS: EXPRESSION ANALYSIS Fill up to 50 ml with cooled cell suspension (i.e., 1 volume of stop solution + 9 volume of culture). Keep on ice. Centrifuge at 4,500 x g, 4"C, 10 min. Remove the supernatant carefully and proceed immediately with RNA isolation or snap-freeze in liquid nitrogen and store at -80°C. 2. Cell lysis: Resuspend cell pellet (vortex or pipette to dissolve clumps) in2.5 ml of ice-cold solution 1.Transfer into a 15-ml tube containing 2.5 ml of hot (65°C) Solution 2. Mix gently by inversion five times. Proceed immediately with phenol extraction. 3. Hot phenol extraction: Add 5 ml of hot phenol (65°C) to each tube, mix gently by inversion, and incubate for 5 min at 65°C. Chill (do not freeze) tubes bottom up in liquid nitrogen for -10 s, keep on ice, and centrifuge at 4,500 X g for 5 min at 4°C. Note: If the aqueous top layer is cloudy after centrifugation, let the tube sit for 1 to 2 min or try to warm it a bit. Transfer the aqueous (top) layer to the 15-mi tube containing 5 ml of hot phenol. Note: It is important to avoid the organic phase in all steps. Mix gently by inversion and incubate for 5 min at 65°C. Chill (do not freeze) in liquid nitrogen for -10 s, bottom up, and centrifuge at 4,500 X g for 5 min at 4°C. 4. Phenollchloroformlisoamyl alcohol extraction: Transfer the aqueous (top) layer to the 15-ml tube containing 5 ml of pheno1:chloroform:isoamyl alcohol (25:24:1, pH 6.6). Mix gently by inversion and centrifuge at 4,500 X g for 5 min a t 4°C. 5. Chloroformlisoamyl alcohol extraction: Transfer the aqueous (top)layer to a 15-ml tube containing 5 ml of ch1oroform:isoamyl alcohol (24:1).Mix gently by inversion and centrifuge at 4,500 X g for 5 min at 4°C. 6. Ethanol precipitation: Transfer the aqueous (top) layer (-4 ml) to the 15-ml tube containing 400 p,l of 3 M NaAc (pH 4.5) + 9 ml of 96% EtOH. Mix gently by inversion. Incubate at -80°C for 30 min or at -20°C overnight. Note: Depending on the added volume, extended incubation at - 80°C may cause freezing. This should be avoided. Centrifuge tubes at 4,500 X g for 30 min at 4°C. 7. Washing: Carefully remove the supernatant and wash the pellet with 5 ml of ice-cold 75% EtOH. Centrifuge at 4,500 X g for 5 min at 4°C. Repeat this step. 8. Discard the ethanol and spin down. Remove the rest carefully using a Pasteur pipette. Air dry the pellet for <5 min. Note: Do not let the pellet dry completely, as a dry pellet will be difficult to dissolve in water.
483 9. Resuspend the pellet in 0.5 to 1 ml RNase-free H,O. Incubate at 60°C for 5 min only if the pellet does not dissolve at room temperature. This may cause some degradation. 10. Quality check: quantitation, purity, and integrity. RNA samples are stored at
-
80°C.
Probe Preparation Materials RNase-free water lox DNase I buffer (Ambion) 0.1 M dithiothreitol (DTT) 2 Up1 of DNase I (Ambion catalog no. 2222) RNase inhibitor (Ambion catalog no. 2694) Qiagen RNeasy Mini Kit Random Hexamer pd(N), (Amersham catalog no 27-2166-01) 100 mM each nucleotide (dATP, dGTP, dCTP, dTTP) 100 mM aminoallyl-dUTP (aa-dUTP) Reverse transcriptase (Stratascript Reverse Transcriptase) with 1OX buffer reverse transcription buffer) and DTT 1 M sodium hydroxide (NaOH) 0.5 M EDTA Zymo DNA purification columns (25 pg) or other column suitable for purification of small amounts of DNA 1 M phosphate buffer, pH -8.5 96% ethanol 0.1 M sodium bicarbonate buffer (pH 9.3) Monofunctional NHS-ester Cy3 or Cy5 dye (Pharmacia, Uppsala, Sweden) DMSO
Notes: Chemicals and enzymes of other suppliers should work as well, but concentrations and volumes have to be adapted. You will need one RNA sample for each dye, therefore 2 samples for every slide. DNase I Treatment of RNA Samples All contaminating genomic DNA has to be removed to avoid reduced labeling efficiency. 1OX DNase I buffer (Ambion) . . . . . . . . . . 10 pl RNase inhibitor (Ambion catalog no.2694) ........................ 5 Pl DNase I 2 U/pl (Ambion catalog 110.2222) ........................ 5 Pl Total RNA ........................ . l o 0 pl Water (RNase-free)to: . . . . . . . . . . . . . . . . 100 pl
MYXO BACTERIAL METHOD s
484
Mix and subject to centrifugation. Incubate in heating block at 37°C for 40 min. Note: Here you may lose 20 to 40% of your starting amount of RNA.
Add 14.5 pl of cocktail to each RNA tube and incubate reaction mixtures at 37°C for 10 min, at 42°C for 1 h and 40 min, and then at 50°C for 10 min. Samples may then be frozen here at -20°C if desired.
RNA Cleanup/Purification Using Qiagen RNeasy Mini Kit
Hydrolysis
This step eliminates DNase I and purifies RNA selectively. Column capacity is 100 pg. Follow the instructions of the manufacturer. Quantify RNA and test integrity as described in “Hot Phenol RNA Isolation” above before proceeding with the labeling reactions. Note: The silica-gel-basedmembrane has low affinity for RNA molecules shorter than 200 bp. Loss of RNA will be 15 to 20%. Protocol for Reverse Transcription and Amino-Ally1 Coupling of cDNA
RT reaction Concentrate RNA to >1.4 pgpl in speed vacuum centrifuge, if necessary. Random prime RNA: pd(N), ( 5 pg/pl) ..................... 2 Pl 20 pg of total RNA (>1.4 pg/pl) . . . . . . . . 5 Pl Water (RNase-free) to . . . . . . . . . . . . . . . .15.5 pl Incubate RNA and oligo pd(N), at 70°C for 10 min. Chill on ice 10 min.
cDNA synthesis Mix according to the schemes: 50X aa-dUTP mix (2:3 ratio) dATP (100 mM) ..................... dGTP (100 mM) ..................... dCTP (100 mM) ..................... dTTP (100 mM) ..................... aa-dUTP (100 mM) . . . . . . . . . . . . . . . . . . . Total
..............................
5 PJ.1 5 PI 5 PI 2 PJ.1 3 Pl 20p1
1. Add and mix 10 pl of 1N NaOH and 10 pl of 0.5 M EDTA. Note: Use fresh NaOH every time. Aliquots can be stored frozen. 2. Incubate 15 min at 65°C. Neutralize with 10 pl of 1MHCl. Note: Neutralization with 1M HCl is not usually necessary if using a Zymo column for cleanup. Using 1ml of binding buffer for the Zymo binding step should obviate the need for any neutralization step. Cleanup To continue with the amino-ally1 dye coupling procedure, all contaminants must be removed from the reaction to prevent the monofunctional NHS-ester Cy-dyes from coupling to free amine groups from the buffer or unincorporated amino-ally1groups. Note: Cleanup can be carried out with commercial spin columns and the associated buffers. Since one does not know the exact composition of these buffers, and since you want to avoid any contamination with free amines, we recommend to prepare your own washing and elution buffers. This provides for higher labeling efficiency. Phosphate buffers can be stored at 4°C for 2 weeks. Use autoclaved deionized (DI) water only (see “Materials” in “Protocol for Hybridization of AminoSilane-Coated Microarray (TIGR)” below). 1M phosphate buffer stock solution, pH 8.5
-
........................ .16.5 g ........................ .0.68 g Autoclaved DI water . . . . . . . . . . . . . . . . 100ml l00ml Total ............................ K,HPO, KH2P0,
Cocktail, no. of pl per aliquot plper 4 1 0 buffer ~ (Stratagene) . . . . . . . . . . . . 3 (13.5) 50 X aa-dUTP mix . . . . . . . . . . . . . . . . 0.6 (2.7)
DTT(0.1M) .................... Stratascript RT (50 U/pl) . . . . . . . . . . .
3(13.5) 3 (13.5) 0.5 (2.3)
RNase inhibitor (Ambion) (20 U/pl) . . Water (RNase-free) . . . . . . . . . . . . . . . 4.5 (20.2)
Total
.........................
.14.6 (65.7)
Phosphate wash buffer, pH 8.0 5 mH KPO,, 80% EtOH) Ethanol ( 9 6 % ) . . . . . . . . . . . . . . . . . . . .42.13 ml Autoclaved DI water . . . . . . . . . . . . . . . 7.63 ml 1M phosphate buffer (add dropwise). . . 0.25ml Total
...........................
50ml
28. 211. XANTHUS: EXPRESSION ANALYSIS
485
Phosphate elution buffer, pH 8.5 (4 mM KPO,)
Cleanup
Autoclaved DI water . . . . . . . . . . . . . . . .49.8 ml 1 M phosphate buffer. . . . . . . . . . . . . . . . 0.2 ml
To remove unincorporated Cy-dyes, proceed with Zymo Spin Column. Alternatively, the cleanup can be done with a QiaQuick purification kit (Qiagen). The eluate can be concentrated in speed vacuum dryer.
Total
............................
50ml
Note: Single-stranded DNA has a lower affinity to the column matrix. To reduce loss of cDNA, add at least 7 volumes of binding buffer to the sample.
1. Load 500 pl of sample into a Zymo Spin Column and place column in a 2-ml collection tube. Centrifuge at full speed for 10 s and discard flowthrough. Repeat loading with remaining sample. Centrifuge again. 2. Add 200 pl of wash buffer to column and centrifuge at full speed for 30 s. Repeat wash and centrifuge again at full speed for 1min. 3. Add 30 p1 of elution buffer to column matrix, incubate for 1min, and elute by centrifuging at full speed for 1 min into a fresh 1.5-ml tube. Note: You can store the cDNA here at -20°C. Before going on, you should measure the amount of cDNA generated by the reaction. Hybridization of too little cDNA will result in weak signals and high background levels.
Coupling This reaction is performed in bicarbonate buffer to ensure the appropriate pH for specificity and efficiency of coupling. The sodium bicarbonate can be stored frozen, but use a fresh tube every time.
1. Resuspend monofunctional NHS-ester Cy3 or Cy5 dye: If using a fresh tube of Cy3/Cy5, contents of the dye tube may be resuspended in 10 pl of DMSO and coupled immediately by transferring 2 pl of dye solution into cDNA + sodium bicarconate buffer. Note: Alternatively, resuspend the entire tube in 10 pl of DMSO, transfer 2-pl aliquots into screw-cap tubes, and immediately dry in speed vacuum dryer without heat. Freeze instantly. 2. Concentrate cDNA in speed vacuum dryer until it is almost dry (-1 1.1). Add freshly prepared 0.1 M sodium bicarbonate, p H 9.3, to a final volume of 8 pl and mix. Add 2 p1 of DMSO-dissolved dye. Incubate for 2 h in the dark or use a shaker wrapped in aluminum foil to shelter from light. Note: Adding more than 2 pl of dye is possible but usually not necessary to achieve complete labeling.
1. Add 500 pl of DNA Binding Buffer to each tube. Load sample into a Zymo Spin Column and place column in a 2-ml collection tube. Centrifuge at full speed for 10 s and discard flowthrough. 2. Add 200 p1 of Wash Buffer (from kit) to the column and centrifuge at full speed for 30 s. Repeat wash with another 200 p1 of Wash Buffer and centrifuge for 1min. Repeat washing one more time. Note: The matrix of the spin columns should now be of distinct color: blue for Cy5- and red for Cy3labeled samples. 3 . Add 30 pi of H,O to column matrix and elute by centrifuging for 10 s into a fresh 1.5-ml tube. 4. Measure and record labeling efficiency with spectrophotometer. Notes: Measure absorbance at 550 nm for Cy3 and 650 nm for Cy.5. You can also take readings at 260 nm, but have in mind that you measure modified cDNA. In our lab, we use a NanoDroplOOO spectrophotometer that allows for simultaneous measurement of all three wavelengths in a l - k l sample volume. If labeling did not work properly, go back to fix the problem or else hybridization results may not be satisfying and expensive arrays will be wasted. 5. Concentrate labeled cDNA until it is almost dry (-1 1.1) in speed vacuum dryer. Store labeled cDNA in the dark at room temperature (or for longer storage at -20°C).
Protocol for Hybridization of Amino-SilaneCoated Microarrays (TIGR) Materials 20X SSC Buffer (Invitrogen; catalog no. 15557-044) ( 1 X SSC is 0.15 M NaCl plus 0.015 M sodium citrate) 10% (wthol) SDS Solution (Roth; catalog no. 2326.2) Bovine serum albumin (BSA) (Albumin Fraction V, Roth; catalog no. 8076.2) Formamide (Roth; catalog no. 6749.1; store desiccated) 2-Propanol (isopropyl alcohol) (Roth; catalog no. 6752.2) Lifterslip cover glasses 25 X 60 (IMPLEN; catalog no. 2260 or Erie Scientific Company; 25x601-2-4789)
MYXO BACTERIAL METHOD s
486 Hybridization chamber(s) (Corning; catalog no. 2551) Millipore or Corning Brand Mini-Miser Filter-Top Tube 0.22 p,m (Fisher Scientific; catalog no. 09761-34) Sheared salmon sperm DNA (10 mg/ml) (Ambion; catalog no. 9680; store at -20°C) Surfactant-free cellulose acetate syringe filters (0.45 p,M) (Millex or W R ; catalog no. 28196-114) Glass staining dishes (Fisher catalog no. 08-812) Metal slide racks with holders Autoclaved DI water Microarray printed slides (TIGR) Cy3/CyS-labeled cDNA probes (see above; store at - 80°C) l-mi tuberculin syringe with slip tip (VWR; catalog no. BD309602) Aluminum foil
Notes: Materials provided by other suppliers should work as well but have not yet been tested in our lab. Cy dyes, particularly Cy5, are sensitive to light and oxygen radicals. At ALL times they must be kept in darkened containers, as light exposure will cause photo bleaching of the dye. W and carbon-filtered MQ water (ELGA or NANOpure Infinity W)may be used as an alternative to autoclaved DI water. However, water must be completely filtered to eliminate any residual free radicals. Free radicals compete with Cy dyes, drastically inhibiting efficient coupling and hybridization. We recommend using only autoclaved DI- or ELGA-water. Prehybridized slides must be used immediately following prehybridization to ensure optimal hybridization efficiency. Prehybridized slides cannot be stored for later use. Therefore, prepare labeled cDNA first (see above), and then proceed with prehybridization.
2 0 SSC ~ ........................ 1 0 % S D S . . ...................... DI water ........................ Total
...........................
42°C water bath Rotary shaker Centrifuge with a flat four-plate adaptor 95°C heat block Microcentrifuge tube centrifuge (Optional) Clean compressed air You will need to have or prepare the following buffers and reagents before beginning this procedure: Low-stringency wash solution (2X SSC, 0.1% SDS)
10ml 890 ml lliter
Heat to 42°C. Note: The low-stringency wash solution has a high amount of salt, which may precipitate out if left on a bench too long. Higher background may be observed if the solution is not properly mixed. To prevent the precipitation, it is recommended that the solution be kept at 42°C at all times. Medium-stringency wash solution (0.1X SSC, 0.1 % SDS) 2 0 SSC ~ ........................ 1 0 % S D S . . ...................... DI water ........................ Total
...........................
5 ml 10ml 985 ml lliter
High-stringency wash solution (0.1x SSC)
........................ ........................ ...........................
2OXSSC DI water. Total
5ml 995 ml lliter
Procedure Prehybridization (i)Prepare prehybridization solution Prehybridization solution (5x SSC, 0.1% SDS, 1% BSA) 20X SSC ........................ 10%SDS .......................
............................ ........................ Total ...........................
BSA
DI water Equipment Required You will need the following instruments for this procedure:
100 ml
125 ml 5ml
5g
330 ml 500ml
Dissolve BSA by stirring on a magnetic stirrer for at least 0.5 h. Filter the prehybridization buffer with a 0.22-pm Mini-Miser (CA) filter. Transfer the solution to a clean glass dish and preheat at 42°C for about 30 min. (ii) Prehybridize the array slide Note: It is extremely important that slides be perfectly clean, or else you will have background problems. Wear gloves. Try not to scratch or touch the printed area. Do not let the slides dry out at any time.
28. M.XANTHUS: EXPRESSION ANALYSIS 1. Place the printed slide(s) carefully and not too tightly in a slide holder and transfer it in the dish with preheated prehybridization buffer (no more than 10 slides per dish-less, if possible, to reduce scratching). 2. Incubate at 42°C for at least 1h. (Recommended: 2 h.) Occasionally sway the rack gently. (iii)Washing prehybridized slides 1. Fill a glass staining dish with autoclaved DI water for every five slides you will wash. Place a clean slide holder in every dish. 2. Using forceps, carefully grip the slides by the label and then remove the slides from the prehybridization solution into the slide holder placed in the waterfilled dish. Note: Moving the whole slide holder would carry over too much SDS and BSA. No more than five slides should be used per holder. Do not place slides in the first or last slot of the holder and do not put slides together so that they touch. Do not let the slides dry. Slides must be completely submerged in water. 3. Attach the metal handle to the slide holder and shake the rack in the water for about 10 s. 4. Place the entire staining dish apparatus on top of a rotor shaker and let shake for approximately 2 min. 5. Change the water inside the staining dish every 2 min or take the slide holder into a new dish with fresh water and let shake. 6. Continue to wash until you have replaced the wash water at least five times. 7. Empty the staining dish and fill with isopropyl alcohol. (Or transfer rack into another dish with isopropanol.) Wash the slides in isopropanol for 2 min on the rotary shaker. 8. Leave the slides in the alcohol and take them immediately to the centrifuge. (iv) Drying slides Note: DO NOT let the slides dry before putting them in the centrifuge. Allowing the slides to slowly air dry will cause a rise in background signal.
1. Take the glass slide holder with the slides out of the isopropyl alcohol and remove the metal handle from the holder. Put the slides into a centrifuge with a flat plate-holder adaptor lined with paper towels. Place another slider holder with the same number of slides as a counterbalance. Make sure the centrifuge is clean inside. Make sure the centrifuge is at room temperature before centrifuging. 2. Centrifuge the slides at -1,000 rpm for 10 min at room temperature.
487 3. Hold slides up to the light to check for any degree of streaking or spots. If any of these appear, the slides must be rewashed and respun. Hybridization (i)Making the hybridization buffer
1. Prepare the following 1X hybridization buffer fresh for every 5 slides: 1X hybridization buffer (50% formamide, 5 X SSC, 0.1% SDS, 0.6 pg salmon sperm DNA) Formamide
.......................
20xSSC.. ........................ 1 0 % SDS.. .......................
......................... ............................
DI water. Total
500 pl 250pl lop1 240 pl 1ml
2. Using a 1-ml syringe, draw up the hybridization solution and filter it through a 0.45-pin-pore-size filter. 3. Add 60 pl of salmon sperm DNA. Set the hybridization solution aside at room temperature for later use. (ii) Preparing the probe for hybridization
1. Add 22 pl of 1X hybridization buffer to the labeled probes prepared previously (Cy3Ky.5 probes: see above). Do not throw away excess hybridization solution! 2. Resuspend the probe by vortexing at very low speed, pipetting, or finger flicking for -1 min. 3. Mix the corresponding Cy3Ky.5 samples. Final volume will be -45 pl. 4. Heat the probe mixture at 95°C for 3 min. (iii)Apply labeled probe mixture t o array
1. Prepare clean coverslips for every slide: dust with compressed air or clean with 70% ethanol and a dustfree wipe. Note: Ensure that the coverslip is at least 60 mm long, as anything smaller will not cover the entire print area. If using Lifterslips, ensure that the raised side of the coverslip faces down. The lifter slip creates a space to contain the hybridization solution and ensures uniform hybridization. The side containing the lifters can be determined by the reflection of light at the edges of the slip; the lifters do not reflect light, whereas glass does. Record the barcode and the corresponding sample that you are going to hybridize for each array. 2. Place a prehybridized microarray slide (array side up) in the bottom half of a hybridization chamber. 3. Pipette the entire labeled probe mixture (-45 ~ 1 ) onto the array (printed) area of the slide. Try to keep bubbles to a minimum.
MYXO BACTERIAL METHOD s
488 4. Place the coverslip over the printed area of the slide. A pipette tip may help to place the coverslip slowly and carefully. Don’t let the probe mixture squeeze out from under the coverslip! Notes: Alternatively, first place slip carefully on slide so that the array is covered and the lifters are facing down. Then, apply the boiled probe to the array: place the pipette tip on the surface of the slide, not touching the Lifterslip, and expel a small droplet. Move the tip and the droplet slowly to one corner of the Lifterslip. Capillary action should draw the liquid under the slip. Slowly dispense more of the probe into the same corner of the Lifterslip. Going slowly and having a clean array and Lifterslip will prevent the formation of bubbles. Once the probe has fully covered the surface of the array, dispense the remaining probe at the remaining three corners. The position of the Lifterslip on the slide can be adjusted with a pipette tip. Work any large bubbles toward the edge by gently pressing the coverslip surface. Remember that the coverslips are glass and will break with rough handling.
(iv) Slide incubation 1. To the small wells at each end of the chamber, add 20 p,1 of unused hybridization solution. This step is to prevent the prehybridization solution from drying under the coverslip. 2. Place the top of the chamber on and seal it. Note: Do not tilt or flip the slide chamber once the chamber is sealed with the slide inside. 3 . Incubate in a 42°C water bath for 16 to 20 h in the dark. (v) Posthybridization washes Note: At this point all the staining dishes used should be covered in aluminum foil to prevent light exposure. Do not let the slides dry out at any time! The probe will streak the surface, causing background problems, if the slides dry. 1. Preheat the low-stringency buffer to 42°C. 2. Prepare two clean glass dishes filled with warmed low-stringency wash buffer and place an empty slide holder in one of these dishes. 3 . After the incubation in “Slide incubation” above, remove the hybridization chamber from the water bath. Remove the slides from the chamber, taking care not to disturb the coverslip. 4. To remove the coverslip, grab the slide label with forceps (or gloved fingertips if the forceps prove difficult) and submerge it in the dish without the slide holder. Keep the array level when submerging in low-stringency wash solution. Once submerged, tilt the slide. With time the coverslip will slide free of the slide surface. It may be
necessary to lightly swish slides under solution to dislodge the slip and gently remove it. Take care not to scratch the printed area with the loose coverslip! Wash the slide for 10 s and place it in the slide holder in the second dish to prevent from drying. Proceed with the next slide until five slides are in the rack. When all slides are in the rack, plunge the rack up and down 10 to 20 times. Note: Change the low-stringency buffer in the first dish for every five slides. Do not place more than five slides in the same rack. Instead, use a new dish (or fresh buffer) for every five slides. The slides should be spread out so they do not touch each other or the sides of the holder (as above). 5. Cover the dish with aluminum foil and put it on a rotary shaker. Let the arrays gently shake for 5 min. 6. After the 5 min, transfer the rack to a dish with new low-stringency buffer (preheated to 42°C) and agitate on a rotary shaker for 5 min. Repeat. 7. Transfer the slides to a slide dish with mediumstringency buffer at room temperature and agitate 5 min. Repeat. 8. After the 5 min, transfer the slides to a slide dish with high-stringency buffer at room temperature and agitate 5 min. Repeat. Note: You can continue with a quick wash in DI water or proceed directly. 9. Without removing the slides from the buffer, carry them to the centrifuge and spin them down as in “Drying slides” above.
Note: Keep the slides in the dark until they can be scanned. Scan array within hours of washing, as the Cy dyes are unstable and will degrade. When scanning, note that the top of the print area begins on the part of the slide farthest from the barcode. A properly scanned slide, therefore, should have a legible barcode at the bottom of the scanned area. Analysis In our lab we use a GenePix 4000B microarray scanner with GenePix Pro 6.0 image analysis software. Normalization and data analysis are carried out with Acuity 4.0X software package. For statistical analysis we use the SAM Software from Stanford University (http://www-stat. stanford.edu/-tibs/SAM/index.html). However, it should be noted that there are other systems available too. References Bowtell, D., and J. Sambrook (ed.). 2002. D N A Microarrays: A Molecular Cloning Manual. Cold Spring Harbor Press, Cold Spring Harbor, NY. Diodati, M. E., F. Ossa, N. B. Caberoy, I. R.Jose, W. Hiraiwa, M. M. Igo, M. Singer, and A. G. Garza. 2006. Nla18, a key regulatory protein required for normal growth and development of Myxococcus xantbus. ]. Bacteriol. 188:1733-1743.
28. M.XANTHUS: EXPRESSION ANALYSIS Jakobsen, J. S., L. Jelsbak, L. Jelsbak, R. D. Welch, C. Cummings, B. Goldman, E. Stark, S. Slater, and D. Kaiser. 2004. oS4enhancer binding proteins and Myxococcus xanthus fruiting body development. ].Bacterial. 186:43614368. Kroos, L., A. Kuspa, and D. Kaiser. 1986. A global analysis of developmentally regulated genes in Myxococcus xanthus. Dev.Biol. 117:252-266.
489 Overgaard, M., S. Wegener-Feldbriigge, and L. SsgaardAndersen. 2006. The orphan response regulator DigR is required for synthesis of extracellular matrix fibrils in Myxococcus xanthus. 1. Bacteriol. 188:4384-4394. Pham, V. D., C. W. Shebelut, I. R. Jose, D. A. Hodgson, D. E. Whitworth, and M. Singer. 2006. The response regulator PhoP4 is required for late developmental events in Myxococcus xanthus. Microbiology 152:1609-1620.
Myxobacteria: Multicellularity and Differentiation Edited by David E. Whitworth 0 2008 ASM Press, Washington, D.C.
IGmberly A. Murphy Anthony G. Garza
Genetic Tools for Studying Myxococcus xanthus Biology
Myxococcus xanthus has a remarkable repertoire of complex multicellular behaviors, including social gliding motility, predation, rippling, and fruiting body formation. There are a few genetic tools such as an autonomously replicating plasmid and a defined library of M . xanthus mutant strains that have yet to be developed to study these complex behaviors. However, because of the efforts of several prominent lab groups over the last 30 to 40 years many, if not most, of the tools available to study model organisms such as Escherichia coli or Bacillus subtilis are now available to study M . xanthus biology. This large arsenal of genetic tools has allowed research on 111. xanthus to expand beyond single gene studies; we can now examine the relationships between multiple genes in the same genetic pathway and the interactions between different genetic pathways. This chapter contains a description of M. xanthus genetic tools and their practical applications.
GENERALIZED TRANSDUCTION Generalized transducing particles form when by mistake a bacteriophage head or capsid assembles around a fragment of a donor bacterium's chromosomal DNA or
29
around a plasmid instead of the phage genome. When these particles infect a recipient host, the donor bacterium's DNA is inserted into the recipient host and is free to undergo homologous recombination with the host cell chromosome.
Mx4 and Mx8 Transducing Phages In M. xanthus, generalized transducing phages are typically used for genetic mapping and for strain constructions. Generalized transduction between 111. xanthus strains is accomplished with lytic phage Mx4 (Campos et al., 1978; Geisselsoder et al., 1978) or the temperate phage Mx8 (Martin et al., 1978). Mx4 and Mx8 can package about 50 kb of DNA (Kaiser, 1984), which corresponds to about 0.5 % of the M . xanthus genome (Goldman et al., 2006). A temperature-sensitive derivative of Mx4 phage (Mx4 ts-27htf hrm-1)that transduces at high frequency in M . xanthus is commonly used (Geisselsoder et al., 1978). Because Mx4 ts-27htf hrm-l phage lysates prepared at 25°C do not replicate when introduced into an M . xanthus strain grown at 32"C, this phage is well suited for use in generalized transductions. In the case of phage Mx8, the clear plaque derivative Mx8 clp2 is often used for generalized transduction (Martin et al., 1978).
Kimberly A. Murphy and Anthony G. Garza, Department of Biology, Syracuse University, 130 College Place, Syracuse, NY 13244-1220.
49 1
-~
MYXOBACTERIAL METHODS
492
Genetic Mapping Generalized transducing phages such as Mx4 and Mx8 have been used to identify the sites of chromosomal point mutations by genetically linking them to antibiotic resistance markers. When two genetic markers are sufficiently close on the chromosome, a single transducing phage can transfer both genetic markers from the donor cell to the recipient cell. Genes that are transferred together to the recipient cell are referred to as coinherited genes. Typically, the phenotype associated with one genetic marker is selected (e.g., kanamycin resistance), and transductants that inherit this marker will be screened for the phenotype associated with inheritance of the second genetic marker (e.g., a defect in gliding motility). Cotransduction frequency is the number of transductants that coinherit both genetic markers divided by the total number of transductants. By applying the Wu equation (Wu, 1966), cotransduction frequency can be used to approximate the physical distance between two chromosomal markers. cotransduction frequency
=
[l- (distance between two markedlength of transduced DNA)I3
Thus, the shorter the distance between two chromosomal markers, the more frequently they will be coinherited by. generalized transduction. However, a limitation of genetic mapping by cotransduction frequency is that the two markers must be in close enough physical proximity to be packaged in the same phage particle. In addition, this approach for genetic mapping requires that a selection exists for at least one of the markers. Typically, one marker imparts antibiotic resistance to the bacterial cells that carry it. To facilitate genetic mapping in M . xanthus, Kuner and Kaiser (1981) developed a procedure to link chromosomal mutations to the selectable marker Tn5, a transposable genetic element that confers resistance to kanamycin (Fig. 1).To do this, a library of independent Tn5 insertions in the M. xunthus chromosome is generated by shuttling Tn5 from E. coli to M. xunthus by using phage P1. Mx4 or Mx8 phage are grown on a pool of strains from the M. xanthus insertion library, and the phage stocks are used to transduce the Tn5 insertions from the pooled cells into the appropriate mutant strain. Kanamycin-resistant transductants are then screened for
Motility- mutant
Motility- mutanl
m \
P1::TnS
Wild-type M xanthus strain
Library of independent Tn5 insertions in M xanthus chromosome
Grow Mx4 or Mx8 on strains from the library
I
Transduce motility- strain using Mx phage
I0/Mot+
Select for Kad, screen for gliding motility
\
or Mx8 on Mot' strain
Transduce motility- strain using Mx phage
1 0I
Mot'
Select for Kan', screen for gliding motility
Figure 1 Procedure for linking chromosomal mutations to Tn5 . The diagram shows a Tn5 insertion being genetically linked to a mutation (*) that causes a gliding motility defect (Mot-), but it is applicable to any chromosomal mutation that produces a distinguishable phenotype. Open rectangles represent M . xanthus cells, and the ovals inside the rectangles represent the M. xanthus chromosome. The small, shaded rectangles denote Tn5 chromosomal insertions that impart kanamycin resistance to recipient cells. Hexagons represent the indicated phages.
29. GENETIC TOOLS FOR STUDYINGM.XANTHUS coinheritance of the wild-type allele of the mutation of interest. To determine the genetic linkage between the mutation of interest and Tn.5, an Mx4 or Mx8 phage stock is generated from kanamycin-resistant transductants that carry the wild-type allele of the mutation, the phage stock is used to transduce the original mutant strain, and kanamycin-resistant transductants are screened for coinheritance of the wild-type allele (Fig. 1). Using a Tn5 library such as the one generated by Kuner and Kaiser (1981), it is possible, for example, to determine whether point mutations that produce similar phenotypes are near a particular Tn.5 marker; close physical proximity to the same Tn5 insertion implies that the mutations are in the same genetic locus in the chromosome. Furthermore, it is possible to identify the site of a mutation because close proximity to Tn5 and its kanamycin resistance cassette facilitates cloning, which in turn allows the region of DNA carrying the mutation to be subjected to DNA sequence analysis. By comparing this DNA sequence to that of the corresponding wildtype region in the M. xanthus genome sequence, the site of the point mutation can be identified. Recently, magellan-4, a transposon that confers kanamycin resistance, was used to create M. xanthus insertion libraries (Youderian et al., 2003; Youderian and Hartzell, 2006; Chavira et al., 2006). The magellan-4 transposon recognizes a TA dinucleotide (Plasterk et al., 1999) and therefore has the capacity to integrate into a large number of sites on the chromosome. It also has features that make cloning chromosomal DNA flanking the insertion site relatively easy (see below). Hence, the magellan-4 transposon library should greatly facilitate the genetic mapping and identification of chromosomal point mutations in M. xanthus.
Strain Construction Generalized transduction has been used extensively to place mutant alleles in different genetic backgrounds, which has provided functional information about the genes carrying the mutations. For example, M . xanthus gliding motility is controlled by the A-motility and S-motility systems (Hodgkin and Kaiser, 1979a, 1979b), and it is common practice to use a transducing phage to transfer an uncharacterized motility mutation (e.g., a Tn.5 insertion mutation) from a donor strain into A-motility and S-motility mutant recipient strains. Since gliding motility is completely abolished only when both motility systems are inactivated, one can determine whether the insertion mutation in the donor strain affects the A-motility or the S-motility system by screening the kanamycin-resistant transductants, which are double mutants, for a complete loss
493 of gliding motility. Similarly, to characterize mutations that affect fruiting body development it has been common practice to transduce developmentally regulated TnSlacZ reporter gene fusions into the strains carrying the mutations. By comparing the lac2 expression profiles of the mutant strains to those of wild-type cells, the approximate time when the developmental program goes awry in the mutants is revealed. This has been a particularly useful strategy for dissecting the genetic pathways that control the process of fruiting body development.
TRANSFER OF PLASMID DNA TO M . XANTHUS Methods for transferring plasmid DNA from E. coli to M.xanthus using phage P1 have been developed (O’Connor and Zusman, 1983; Gill et al., 1988). These methods take advantage of the fact that P l y which is an E. coli phage, is unable to multiply in M. xanthus (Kuner and Kaiseq 1981),but in some instances it is able to package plasmid DNA into phage particles and introduce them into M. xanthus cells. Another method of transferringplasmid DNA to M. xanthus cells is electroporation, which is by far the most common method used today (Kashefi and Hartzell, 1995). Because there are no plasmids available that replicate autonomously in M. xanthus, plasmids are propagated in, and isolated from, E. coli before they are introduced into M. xanthus cells. Hence, they must contain E. coli origins of replication. These E. colibased plasmids are modified in a variety of ways for use in M. xanthus research. However, the one thing that all of the plasmids have in common is that once they are inside M. xanthus they must integrate into the chromosome in order to be passed onto progeny cells. To promote sitespecific integration in the M. xanthus chromosome, some plasmids used in M. xanthus research carry an Mx phage attachment (att)site and a phage integrase (int)gene. For complementation or expression studies, which are the type of analyses typically done with these Mx att plasmids, the gene or genes of interest and their corresponding promoter elements are ligated into the plasmids. The plasmids are introduced into M. xanthus cells, strains that carry the plasmid integrated into phage attachment sites in the chromosome are identified, and the appropriate assays are performed. Other plasmids contain M. xanthus DNA that allows them to integrate into the chromosome via homologous recombination. For example, an internal fragment of an M. xanthus gene can be placed into an E. coli plasmid in order to inactivate the chromosomal copy of the gene, a procedure that is described in more detail below.
MYXO BACTERIAL METHOD s
494
PLASMID INTEGRATION AT PHAGE ATTACHMENT SITES In M . xanthus, phage attachment sites are often used for genetic complementation experiments; a wild-type copy of a gene and its native promoter are introduced into the phage attachment site of a strain carrying a mutation in that gene. Phage attachment sites have also been used for mutational analysis of M. xanthus genes and promoters. In M. xanthus, site-specific integration of Mx8 occurs between attP, the preferred site on the Mx8 genome, and attB, the preferred site on the M. xanthus chromosome (Stellwag et al., 1985), which is shown in Fig. 2. Within the Mx8 genome, a minimal 2.2-kb fragment has been defined that contains the attP site and the integrase gene int required for integration at attB (Magrini et al., 1999). Several plasmids that contain the attP site and the int gene have been constructed. When M. xanthus genes are introduced into these plasmids, they seem to have a high degree of preference for integration into the chromosomal attB site. For example, Tojo et al. (1993)
generated a plasmid carrying the Mx8 attP site and int gene, as well as a 1.3-kb fragment of the M. xanthus ZonD gene. When this plasmid was transferred to M. xanthus cells, the frequency of site-specific recombination at the chromosomal attB site was about 300-fold higher than the frequency of homologous recombination at the native lonD locus in the chromosome (Tojo and Komano, 2003). In some cases, expression of M. xanthus genes and their native promoters at the Mx8 attB site is relatively low compared to expression at the native locus on the chromosome (Li and Shimkets, 1988; Fisseha et al., 1996; Whitworth et al., 2004). Hence, using the attB site for complementation analysis or other analyses in which gene expression levels are important may be problematic. The advantage of using the Mx8 attB site or another phage attachment site in the chromosome for complementation or expression studies, for example, is that only a single copy of the gene and promoter of interest is provided to M. xanthus cells, which alleviates potential concerns about multicopy effects. Recently, Julien (2003) generated plasmid vectors for integration into the Mx9 phage attachment site in the M . xanthus chromosome. The integration gene int, phage attachment site, and site of integration in the M. xanthus chromosome were identified and characterized. Mx9 integrates into the M. xanthus genome by site-specific recombination at one of two sites, attB1 or attB2, and requires a single protein, the product of the int gene. Julien (2003) found that integration of plasmids containing piZA or mgZ promoter fusions to lac2 at either Mx9 attB site is expressed at higher levels than the same fusions integrated at the Mx8 attB site, suggesting that the Mx9 attB2 and attB2 sites may be better suited for complementation and expression studies in some cases.
RANDOM MUTAGENESIS
attL
attR
Figure 2 Site-specific recombination at a chromosomal Mx8 phage attachment site. A plasmid carrying a gene conferring kanamycin resistanc (Kan'), the Mx8 phage attachment site attP (gray box) located within the coding sequence for the integrase gene int (stippled box), and the locus of interest (cross-hatched box) is shown. The plasmid integrates into the M. xanthus chromosomal Mx8 phage attachment site via sitespecific recombination between the attP and attB sites. The integration event produces the indicated DNA arrangement in the chromosome.
In M. xanthus, random mutations have been generated using conventional chemical mutagens such as nitrosoguanidine and ethyl methanesulfonate, UV irradiation, and insertion mutagenesis. The advantage of using chemical mutagens and W irradiation is that they produce single-base-pair changes. Therefore, conditional mutations or mutations in essential genes that only partially disrupt the function of their corresponding protein products can be generated. Although these types of mutations are difficult to generate using transposon or random plasmid insertions, insertions are particularly useful for generating a large pool of mutations that inactivate chromosomal genes and for tagging mutated genes so that they can be identified. Below is a brief description of transposable elements that are commonly used to
29. GENETIC TOOLSFOR STUDYING M.XANTHUS make insertions in M. xanthus, as well as a method for making random plasmid insertions.
T n 5 and Derivatives In M. xanthus, Tn5, a transposable genetic element that confers resistance to kanamycin, can be used to generate a library of insertions in the M. xanthus chromosome or to link a particular locus to an antibiotic resistance marker for genetic mapping or for cloning. Tn5 is introduced into M. xanthus by the E. coli transducing phage P1::TnS (Kuner and Kaiser, 1981). In M. xanthus this virus serves as a suicide vector; P1::TnS is unable to replicate inside M. xanthus cells, but kanamycin-resistant transductants are formed when Tn5 transposes into the M . xanthus chromosome. Once Tn5 integrates into the chromosome of M. xanthus cells, the frequency of transposition into new sites on the chromosome appears to be about per cell (Avery and Kaiser, 1983; Sodergren and Kaiser, 1983). Hence, P1::TnS generates mutants with relatively stable chromosomal insertions. This property and the fact that P1::TnS tends to generate mutants that carry only a single Tn5 insertion help prevent complications when trying to identify the mutated gene that is responsible for the desired phenotype.
495 Tn5-Tp (Sasakawa and Yoshikawa, 1987) and Tn.5132 (Rothstein et al., 1981; Avery and Kaiser, 1983) are derivatives of Tn5 that retain most of the right and left IS Tn5O elements, but in the central region the kanamycin resistance module has been replaced with genes conferring resistance to trimethoprim and tetracycline, respectively (Fig. 3A). For the most part, these elements have been used to replace the kanamycin resistance cassettes in existing Tn5 insertions with different selectable markers. To do this, the strain carrying the Tn5 insertion is infected with P1::TnS-Tp or Pl::Tn5-132 and transductants resistant to either trimethoprim or tetracycline are identified. If a double homologous recombination event occurs between the IS50 sequences in Tn5-Tp or Tn5-132 and the IS50 sequences in the resident Tn5 element in the chromosome, then the kanamycin resistance cassette of the resident Tn5 element will be replaced by tetracycline or trimethoprim modules. The resulting strain has a single transposon insertion at the same chromosomal location as the original Tn5, but the new Tn5 element will confer resistance to tetracycline or trimethoprim instead of kanamycin. With a tetracycline or trimethoprim marker tagging the insertion of interest, it is then possible to introduce a new insertion that
A. IS50,
IS50,
B. I
I
IS50,
IS50,
I
IS50,
IS50,
I
IS50,I
C. I
IS50,l
I
phoA Figure 3 Structures of Tn5 and its derivatives. (A) Tn5 containing the genes that confer resistance to kanamycin (Kan'), tetracycline (Tet'), or trimethoprim (Tmp') flanked by IS50 elements. (B) A derivative of Tn5 used to create lac2 transcriptional fusions. In this Tn5 derivative, a segment of the ISSO, element is replaced with a promoterless lucZ reporter gene. (C) A derivative of Tn5 used to create PhoA translational fusions. A segment of the ISSO, element is replaced with a fragment of the phoA gene. The phoA fragment lacks sequences corresponding to the PhoA translational start site and sequences needed for export of the PhoA protein across the cytoplasmic membrane.
496 confers kanamycin resistance into this strain. Having the flexibility to place two different insertions in the same strain would, for example, provide an opportunity to do genetic epistasis experiments or to genetically map the locations of the insertions relative to one another.
magellan-4 In M . xanthus studies, plasmid pMycoMar (Rubin et al., 1999) has been used as a donor for the transposition of magellan-4, a minitransposon derived from the mariner element Himarl. magellan-4 contains the npt gene from Tn5, which confers resistance to kanamycin, flanked by the ends of Himarl. Adjacent to the magellan-4 element in plasmid pMycoMar is the Himarl transposase gene, which is under control of the mycobacerial T6 promoter (Barsom and Hatfull, 1996). This mycobacterial promoter seems to function well in M . xanthus; transposition of magellan-4 in M. xanthus cells is efficient after plasmid pMycoMar has been introduced by electroporation (Youderian et al., 2003). Plasmid pMycoMar cannot replicate autonomously in M . xanthus. However, expression of the plasmid-borne transposase promotes transposition of the magellan-4 element into the chromosome. Presumably, transposition occurs by a “cut and paste” mechanism (Lampe et al., 1996) that does not require any M . xanthus proteins. To identify the chromosomal site in which magellan-4 has inserted, magellan-4 and flanking M . xanthus DNA can be circularized by digesting and ligating the chromosomal DNA of an insertion mutant. The circularized DNA is transferred and propagated in E. coli (magellan-4 contains an origin for replication in E. coli) and then isolated and subjected to DNA sequence analysis. In recent studies, magellan-4 was used to identify new genes for A-motility and S-motility. When placed on a solid surface, the A-motility system controls individual cell movements, while S-motility controls the movements of multicellular groups (Hodgkin and Kaiser, 1979a, 1979b).Using magellan-4, Youderian et al. (2003)implicated approximately 30 new genes in A-motility. Similarly, Youderian and Hartzell (2006) implicated about 30 new genes in S-motility. In both studies, the desired motility mutants were found with a frequency of about 1%, which is fivefold higher than the frequency obtained through similar approaches with transposon Tn.5 (Youderian et al., 2003; Youderian and Hartzell, 2006). Therefore, magellan-4 seems to have a much broader spectrum of target sites in M. xanthus than does Tn.5.
Plasmid Insertion Libraries Although M. xanthus insertion libraries have typically been created using site-specific transposable elements, they can also be created by homologous recombination
MYXO BACTERIAL METHOD s of random cloned fragments of M . xanthus DNA. For example, Cho and Zusman (1999a) generated a plasmid library containing approximately 500-bp random M . xanthus DNA fragments. When electroporated into wild-type M. xanthus cells, these Kan‘ plasmids were unable to replicate autonomously, but they did integrate into the chromosome via homologous recombination of the cloned M . xanthus DNA. Since the SOO-bp fragments of cloned DNA were smaller than most M. xanthus genes, many of the Kan‘ transformants contained inactivated copies of chromosomal genes. Thus, Cho and Zusman (1999a)were able to generate a random pool of M. xanthus insertion mutants. Phenotypic characterization of these insertion mutants led to the discovery of the espAB locus, which seems to control the timing of M. xanthus sporulation, and the asgD locus, a locus involved in production of an M. xanthus cell density signal called Asignal and nutrient sensing (Cho and Zusman, 1999a, 1999b). In a recent study, Lu et al. (2005)screened the random plasmid insertion library generated by Cho and Zusman (1999a)for M . xanthus mutants with defects in exopolysaccharide (EPS) production, which is required for S-motility, and identified two genetic regions essential for EPS biogenesis: the EPS synthesis (eps)region and the EPS-associated (eas)region.
DIRECTED MUTAGENESIS Targeted Plasmid Insertions In recent years, targeted plasmid insertions have been used to inactivate a iarietyof M . xanthus genes. To make a targeted plasmid insertion, a partial fragment of the target gene is generated using PCR, the PCR fragment is cloned into a suicide plasmid that carries an appropriate antibiotic resistance gene, and the plasmid is electroporated into wild-type M . xanthus cells. Although the plasmid is incapable of autonomous replication in M . xanthus, it can integrate into the chromosomal copy of the target gene by homologous recombination of the cloned PCR fragment. A single crossover yields kanamycin-resistant electroporants with two incomplete copies of the gene separated by vector DNA (Fig. 4).Therefore, the plasmid insertion is likely to inactivate the gene and its corresponding protein product. Homologous recombination between plasmid-borne gene fragments and genes that reside in the M. xanthus chromosome, which is essential for inactivation of chromosomal genes and hence for this technique to work, appears to be constrained by the size of the plasmidborne fragment. For example, in a recent study we used this technique to inactivate 28 ntrC-like activator (nla)
29. GENETIC TOOLSFOR STUDYING211.
497
XANTHUS
rn plasmid
I 5' end target gene
3' end target gene
Figure 4 Strategy for making targeted insertions. An internal fragment of the gene of interest is generated using PCR, the PCR fragment is cloned into a plasmid vector that confers resistance to an antibiotic such as kanamycin, and plasmid DNA is electroporated into wildtype M. xanthus cells. A single homologous crossover produces a tandem duplication of the internal fragment and incorporation of the vector into the chromosomal copy of the gene. The likely result of the crossover is an inactivated copy of the gene.
genes (Caberoy et al., 2003). We were frequently unable to generate insertions when the plasmid-borne n h gene fragments had about 350 bp of homology with their chromosomal nlu gene targets. When we increased the region of homology to about 500 bp, most of plasmid-borne nla fragments that we tested yielded insertions, and when we increased the region of homology to about 600 bp, all of the plasmid-borne nlu fragments that we tested produced insertions. In some studies, we have been able to generate insertions using plasmid-borne fragments as small as 250 bp. However, we were able to generate the insertion mutants only when we used a relatively high concentration of plasmid DNA for the electroporation and when we allowed electroporated cells to recover from electric shock in rich medium for 12 to 24 h instead of 4 to 8 h, which is standard procedure. With the availability of the M. xanthus genome sequence (Goldman et al., 2006) (GenBank accession number CP000113), it is now possible to inactivate whole gene families using targeted plasmid insertions. For example, the M. xanthus genome sequence revealed 53 genes encoding NtrC-like proteins, transcriptional activators that participate in a variety of adaptive responses in bacteria. Using information from the
M. xunthus genome sequence and the targeted plasmid insertion strategy, almost all of the 41 uncharacterized ntrC-like genes have been inactivated and characterized in the past 5 years (Caberoy et al., 2003; Jakobsen et al., 2004; Jelsbak et al., 2005). These studies led to the discovery of 10 NtrC-like proteins that are required for fruiting body development to progress normally.
In-Frame Deletions Insertions have the potential to block transcription of genes located downstream of the insertion sites via polar effects. Therefore, it is often difficult to determine whether the phenotype caused by an insertion is due to inactivation of the gene that carries the insertion or due to the effect the insertion has on downstream transcription. This problem is eliminated by constructing a strain that carries an in-frame deletion of the gene of interest. An M. xunthus mutant containing an in-frame deletion in the target gene can be constructed by gene replacement using plasmids that provide a gene for positive screening such as the kanamycin resistance gene, and a gene for negative screening such as sucB (Ried and Collmer, 1987; Wu and Kaiser, 1996) or gulK (Ueki et al., 1996). Although both sacB and gulK work for negative
MYXOBACTERIAL METHODS
498 screening, the galK gene has been used more frequently for construction of in-frame deletions in M. xanthus. To generate the appropriate plasmid-borne in-frame deletion, a reverse PCR on a plasmid containing genes for both positive (kanamycin resistance) and negative (sacB or galK) selection, as well as the target gene and flanking M. xanthus DNA, can be performed. Typically, the primers used for the PCR correspond to the 5’ end and 3’ end of the target gene and they have restriction sites engineered into their 5’ tails. The restriction sites engineered into the primers are unique sites not found in the plasmid template, allowing a plasmid containing an in-frame deletion of the target gene to be generated by digesting the PCR product with the appropriate restriction enzyme and then ligating the digested DNA. The inverse PCRs are usually performed with mixtures containing Taq and high-fidelity DNA polymerases, the plasmid template and primers carrying the appropriate restriction sites. An alternative strategy to reverse PCR is now possible with the availability of the M. xanthus genome sequence (Goldman et al., 2006) (GenBank accession number CP000113). In this approach standard protocols are used to generate two PCR products, one corresponding to DNA upstream of the gene to be deleted and the other corresponding to DNA downstream of the gene to be deleted. Because the primers are designed to produce upstream and downstream PCR products with compatible restriction sites a t their 3‘ and 5‘ ends, respectively, the PCR products can be digested with the appropriate restriction enzyme and then ligated together to create a DNA fragment carrying a deletion of the target gene. The primers can also be designed to yield an upstream PCR product with a restriction site at its 5’ end and a downstream PCR product with a restriction site at its 3’ end, which allows the deletion-carrying DNA fragment to be cloned into a positive-negative selection vector. After introducing the in-frame deletion plasmid into M. xanthus cells by electroporation, strains with the plasmid integrated in the chromosomal copy of the target gene locus by homologous recombination are identified by selecting for resistance to kanamycin. Strains carrying the integrated plasmid are sensitive to galactose because of the plasmid-borne galK gene or to sucrose because of the plasmid-borne sacB gene, depending on the positivenegative selection vector that is used. Strains in which a plasmid excision event has occurred are identified by their resistance to galactose or sucrose and their sensitivity to kanamycin. In some cases, a plasmid excision event gives rise to a strain that has an in-frame deletion of the target gene.
GENE FUSIONS Gene fusions have been used to assay the transcriptional/ translational regulation of genes/proteins and to examine protein localization in bacterial cells (Silhavy and Beckwith, 1985). Fusions belong to one of two classes: transcriptional fusions or translational fusions. In the case of a transcriptional fusion, the reporter gene lacks a promoter, but it possesses a functional ribosome-binding site. Hence, the reporter gene is transcribed with the endogenous gene, but the reporter gene product is not fused to the product of the endogenous gene. In the case of a translational fusion, the reporter gene lacks a promoter and a functional ribosome-binding site. Therefore, a fusion is created between the products of the reporter gene and of the endogenous gene.
Transcriptional Fusions: TnSZuc In M. xanthus, TnSlac (Kroos and Kaiser, 1984), a derivative of Tn5 that contains a promoterless lacZ reporter gene, has been used to generate transcriptional fusions (Fig. 3B). TnSlac is introduced into M. xanthus cells using phage P1::TnSlac (Kroos and Kaiser, 1984). Once inside an M . xanthus cell, TnSlac can transpose into a variety of sites in the chromosome. If the promoterless lacZ gene in TnSlac is in the same orientation as the gene in which it has inserted, then it will place lacZ under transcriptional control of the promoter for that gene. Using TnSlac, Kroos et al. (1986) generated a collection of developmentally regulated lac2 reporter fusions by screening for increases in P-galactosidase production during fruiting body formation. This collection of lacZ reporters has been used extensively to characterize M. xanthus developmental mutants. By introducing a representative panel of these lacZ fusions into a developmental mutant and monitoring lac2 expression, the approximate time that the developmental program goes awry in the mutant can be determined. Kroos et al. (1990) also used this collection of TnSlac fusions, which create insertion mutations, to identify M. xanthus mutants with severe developmental defects.
Translational Fusions: TnphoA TnphoA (Fig. 3C) is a version of the Tn.5 transposon with the phoA gene from E. coli inserted at its left end (Manoil and Beckwith, 1985). The product of the phoA gene is the enzyme alkaline phosphatase, which must be exported to the periplasm or beyond to be active (Michaelis et al., 1983; Hoffman and Wright, 1985). All sequences needed for transcription, translation, and export have been deleted from the phoA gene in TnphoA. When TnphoA transposes into the chromosomal copy of a gene in the appropriate way, a fusion between PhoA
29. GENETIC TOOLSFOR STUDYING211.
499
XANTHUS
and the product of the gene is created. If the protein to which PhoA is fused contains sequences that allow PhoA to be exported, then alkaline phosphatase activity will be detected. Hence, TnphoA is an excellent probe to identify endogenous exported proteins. TnphoA was used by Kalos and Zissler (1990) to detect genes whose products are involved in cell interactions. Three of these TnphoA mutants carried insertions in the cgl locus, a locus that is important for the normal function of the A-motility system that controls individual cell gliding (Hodgkin and Kaiser, 1977, 1979a). Subsequent work has confirmed that one of the products of the cgl locus, CglB, is localized to the outer membrane (Rodriguez and Spormann, 1999; Simunovic et al., 2003). Although Kalos and Zissler (1990) used TnphoA fusions successfully in their study, TnphoA usage has gained little traction in subsequent years, which perhaps is due to the fact that M. xanthus strains have multiple Pho systems and hence a high background of alkaline phosphate activity.
INDUCIBLE PROMOTER SYSTEMS As the name implies, inducible promoter systems allow gene transcription to be induced under the desired circumstances. They are particularly useful tools when trying to evaluate the effects of expressing relatively low levels or relatively high levels of a particular gene under a particular condition. At this time, the availability of inducible promoter systems to study M. xanthus is limited. However, given that data from DNA microarraybased promotedgene expression studies and molecular studies of M. xanthus promoter elements are beginning to accumulate, the number of inducible promoter systems is likely to increase in the near future. One inducible promoter system that has been used in several studies is the M. xanthus light-inducible carQRS system. When growing cultures of M. xanthus cells are kept in the dark, the carQRS promoter is almost completely inactive (Letouvet-Pawlek et al., 1990; Hodgson, 1993). In contrast, exposing cells to blue light causes the carQRS promoter to become highly active. Hence, under conditions of vegetative growth carQRS is an ideal inducible promoter system. However, the blue light needed to induce expression from the carQRS promoter inhibits the process of fruiting body development (Li et al., 1992). Therefore, for the most part use of this promoter system has been restricted to studies involving vegetatively growing cells. Furthermore, it has been reported that light induction of the carQRS promoter at the Mx8 phage attachment site attB in the chromosome increases activity only about 6-fold, while light induction
of this promoter at the native carQRS locus on the chromosome increases activity about 100-fold (Whitworth et al., 2004). Thus, induction of the carQRS promoter at the Mx8 phage attachment site in the chromosome appears to be attenuated. Another inducible promoter system that has been used in M. xanthus is the tac promoter from E. coli (Letouvet-Pawlek et al., 1990), a hybrid consisting of parts of the trp promoter and lac promoter/operator. In E. coli, transcription from the tac promoter is repressed when the product of the lac1 gene binds to the lac operator and derepressed when cells are provided with isopropyl-p-D-thiogalactopyranoside(IPTG), which binds to and inactivates LacI. When the trp promoter and the lac1 gene were introduced into M. xanthus and the cells were grown vegetatively, there was a low background level of activity from the tac promoter. With the addition of IPTG the activity of the tac promoter in growing cultures increased about eightfold. The activity of the tac promoter was not tested under the starvation conditions that induce fruiting body development. However, in a recent study Jelsbak and Kaiser (ZOOS) generated a system in which a native M. xanthus promoter element can be induced during fruiting body development by adding IPTG. In wild-type cells, the promoter for pilA, a gene that codes for the external structural subunit of type IV pili, is active during vegetative growth and during fruiting body development. For this study, the leader sequence of chromosomal copy of pilA was replaced with the lac operator and a ribosomal binding site from E. coli. In addition, the lac1 gene was placed under control of the pilA promoter and then introduced into the Mx8 phage attB site in the chromosome. When no IPTG was provided to cells carrying this promoter system, expression of pilA increased about twofold during development, indicating that LacI was not able to fully repress pilA expression. When IPTG was provided, pilA expression in cells carrying these promoter constructs increased about sixfold during development. Thus, while this system is not optimal because of the background expression in the absence of IPTG, it does provide a way to generate relatively high levels of gene expression in developing cells.
References Avery, L., and D. Kaiser. 1983. In situ transposon replacement and isolation of a spontaneous tandem duplication. Mol. Gen. Genet. 191:99-109. Barsom, E. K., and G. F. Hatfull. 1996. Characterization of Mycobacterium smegmatis gene that confers resistance to phages L5 and D29 when overexpressed. Mol. Microbial. 2 1:159-1 70.
500 Caberoy, N. B., R. D. Welch, J. S. Jakobsen, S. C. Slater, and A. G. Garza. 2003. Global mutational analysis of NtrC-like activators in Myxococcus xanthus: identifying activator mutants defective for motility and fruiting body development. ]. Bacteriol. 185:6083-6094. Campos, J. M., J. Geisselsoder, and D. R. Zusman. 1978. Isolation of bacteriophage Mx4, a generalized transducing phage for Myxococcus xanthus. J. Mol. Biol. 119:167-178. Chavira, M., N. Cao, K. Le, T. Riar, N. Moradshahi, M. McBride, R. Lux, and W. Shi. 20 October 2006. PD-Allose inhibits fruiting body formation and sporulation in Myxococcus xanthus. J. Bacteriol. doi:l0.1128/JB.00792-06. Cho, K., and D. R. Zusman. 1999a. Sporulation timing in Myxococcus xanthus is controlled by the espAB locus. Mol. Microbiol. 34:7 14-725. Cho, K., and D. R. Zusman. 199913. AsgD, a new two-component regulator required for A-signalling and nutrient sensing during early development of Myxococcus xanthus. Mol. Microbiol. 34:268-281. Fisseha, M., M. Gloudemans, R. E. Gill, and L. Kroos. 1996. Characterization of the regulatory region of a cell interactiondependent gene in Myxococcus xanthus. ]. Bacteriol. 178: 2539-2550. Geisselsoder,J., J. M. Campos, and D. R. Zusman. 1978. Physical characterization of bacteriophage Mx4, a generalized transducing phage for Myxococcus xanthus. J. Mol. Biol. 119~179-189. Gill, R. E., M. G. Cull, and S. Fly. 1988. Genetic identification and cloning of a gene required for developmental cell interactions in Myxococcus xanthus. J. Bacteriol. 1705279-5288. Goldman, B. S., W. C. Nierman, D. Kaiser, S. C. Slater,A. S. Durkin, J. Eisen, C. M. Ronning, W. B. Barbazuk, M. Blanchard, C. Field, C. Halling, G. Hinkle, 0. Iartchuk, H. S. Kim, C. Mackenzie, R. Madupu, N. Miller, A. Shvartsbeyn, S. A. Sullivan, M. Vaudin, R. Wiegand, and H. B.Kaplan. 2006. Evolution of sensory complexity recorded in a myxobacterial genome. Proc. Natl. Acad. Sci. USA 103:15200-15205. Hodgkin, J., and D. Kaiser. 1977. Cell-to-cell stimulation of movement in nonmotile mutants of Myxococcus. Proc. Natl. Acad. Sci. USA 74:2938-2942. Hodgkin, J., and D. Kaiser. 1979a. Genetics of gliding motility in Myxococcus xanthus (Myxobacterales): genes controlling movement of single cells. Mol. Gen. Genet. 171:167-176. Hodgkin, J., and D. Kaiser. 1979b. Genetics of gliding motility in Myxococcus xanthus: two gene systems control movement. Mol. Gen. Genet. 171:177-191. Hodgson, D. A. 1993. Light-induced carotenogenesis in Myxococcus xanthus: genetic analysis of the carR region. Mol. Microbiol. 7:471-48 8. Hoffman, C., and A. Wright. 1985. Fusions of secreted proteins to alkaline phosphatase: an approach for studying protein secretion. Proc. Natl. Acad. Sci. USA 825107-5111. Jakobsen, J. S., L. Jelsbak, L. Jelsbak, R. D. Welch, C. Cummings, B. Goldman, E. Stark, S. Slater, and D. Kaiser. 2004. d4enhancer binding proteins and Myxococcus xanthus fruiting body development. J. Bacteriol. 186:4361-4368. Jelsbak, L., and D. Kaiser. 2005. Regulating pilin expression reveals a threshold for S motility in Myxococcus xanthus. J. Bacteriol. 1872105-2112.
MYXO BACTERIAL METHOD s Jelsbak, L., M. Givskov, and D. Kaiser. 2005. Enhancer-binding proteins with a forkhead-associated domain and the d4 regulon in Myxococcus xanthus fruiting body development. Proc. Natl. Acad. Sci. USA 102:3010-3015. Julien, B. 2003. Characterization of the integrase gene and attachment site for the Myxococcus xanthus bacteriophage Mx9. J. Bacteriol. 185:6325-6330. Kaiser, D. 1984. Genetics of Myxobacteria, p. 163-184. In E. Rosenberg (ed.),Myxobacteria: Development and Cell Interactions. Springer-Verlag, New York, NY. Kalos, M., and J. Zissler. 1990. Transposon tagging of genes for cell-to-cell signaling in Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 878316-8320. Kashefi, K., and P. L. Hartzell. 1995. Genetic suppression and phenotypic masking of a Myxococcus xanthus fat;-defect. Mol. Microbiol. 15:483-494. Kroos, L., and D. Kaiser. 1984. Construction of TnSlac, a transposon that fuses lacZ expression to endogenous promoters, and its introduction into Myxococcus xanthus. Proc. Natl. Acad. Sci. USA 815816-5820. Kroos, L., A. Kuspa, and D. Kaiser. 1986. A global analysis of developmentally regulated genes in Myxococcus xanthus. Dev. Biol. 117:252-266. Kroos, L., A. Kuspa, and D. Kaiser. 1990. Defects in fruiting body development caused by TnSlac insertions in Myxococcus xanthus. J. Bacteriol. 172:484-487. Kuner, J., and D. Kaiser. 1981. Introduction of transposon TnS into Myxococcus for analysis of developmental and other nonselectable mutants. Proc. Natl. Acad. Sci. USA 78:425-429. Lampe, D., M. E. Churchill, and H. M. Robertson. 1996. A purified mariner transposase is sufficient to mediate transposition in vitro. E M B O J. 155470-5479. Letouvet-Pawlek, B., C. Monnier, S. Barray, D. A. Hodgson, and J. F. Guespin-Michel. 1990. Comparison of P-galactosidase production by two inducible promoters in Myxococcus xanthus. Res. Microbiol. 141:425-435. Li, S., B.-U. Lee, and L. J. Shimkets. 1992. csgA expression entrains Myxococcus xanthus development. Genes Dev. 6:401-410. Li, S. F., and L. J. Shimkets. 1988. Site-specific integration and expression of a developmental promoter in Myxococcus xanthus. J. Bacteriol. 1705552-5556. Lu, A., K. Cho, W. P. Black, X. Y. Duan, R. Lux, Z. Yang, H. B. Kaplan, D. R. Zusman, and W. Shi. 2005. Exopolysaccharide biosynthesis genes required for social motility in Myxococcus xanthus. Mol. Microbiol. 55:206-220. Magrini, V., C. Creighton, and P. Youderian. 1999. Site-specific recombination of temperate Myxococcus xanthus phage Mx8: genetic elements required for integration. J. Bacteriol. 181:4050-4061. Manoil, C., and J. Beckwith. 1985. Tn phoA: a transposon probe for protein export sequences. Proc. Natl. Acad. Sci. USA 82: 8 129-8 133. Martin, S., E. Sodergren, T. Masuda., and D. Kaiser. 1978. Systematic isolation of transducing phages for Myxococcus xanthus. Virology 88:44-53. Michaelis, S., H. Inouye, D. Oliver, and J. Beckwith. 1983. Mutations that alter the signal sequence of alkaline phosphatase in Escherichia coli. J. Bacteriol. 154:366-374.
TOOLSFOR STUDYINGM. 29. GENETIC
XANTHUS
O’Connor, I<. A,, and D. R. Zusman. 1983. Coliphage P1mediated transduction of cloned DNA from Escherichia coli to Myxococcus xanthus: use for complementation and recombinational analyses. J . Bacteriol. 155:317-329. Plasterk, R. H. A., Z. Izsvaak, and Z. Ivics. 1999. Resident aliens: the TcUmariner superfamily of transposable elements. Trends Genet. 15:326-332. Ried, J. L., and A. Collmer. 1987. An nptl-sacB-sacR cartridge for constructing directed, unmarked mutations in gramnegative bacteria by marker exchange-eviction mutagenesis. Gene 57:239-246. Rodriguez, A. M., and A. M. Spormann. 1999. Genetic and molecular analysis of cglB, a gene essential for single-cell gliding in Myxococcus xanthus. J. Bacteriol. 181:43814390. Rothstein, S. J., R. A. Jorgensen, J. C.-P. Yin, Z. Yong-Di, R. C. Johnson, and W. S. Reznikoff. 1981. Genetic organization of Tn5. Cold Spring Harbor Symp. Quant. Biol. 45: 99-105. Rubin, E. J., B. J. Akerley, V. N. Novik, D. J. Lampe, R. N. Husson, and J. J. Mekalanos. 1999. In vivo transposition of mariner-based elements in enteric bacteria and mycobacteria. Proc. Natl. Acad. Sci. USA 96:1645-1650. Sasakawa, C., and M. Yoshikawa. 1987. A series of Tn5 variants with various drug-resistance markers and suicide vector for transposon mutagenesis. Gene 56:283-288. Silhavy, T. J., and J. R. Beckwith. 1985. Uses of lac fusions for the study of biological problems. Microbiol. Rev 49:398418. Simunovic, V., F. C. Gherardini, and L. J. Shimkets. 2003. Membrane localization of motility, signaling, and polyketide synthetase proteins in Myxococcus xanthus. 1. Bacteriol. 185:5066-5075.
501 Sodergren, E., and D. Kaiser. 1983. Insertions of Tn5 near genes that govern stimulatable cell motility in Myxococcus. J. Mol. Biol. 167:295-310. Stellwag, E., J. M. Fink, and J. Zissler. 1985. Physical characterization of the genome of the Myxococcus xanthus bacteriophage Mx8. Mol. Gen. Genet. 199:123-132. Tojo, N., S. Inouye, and T. Komano. 1993. The lonD gene is homologous to the lon gene encoding an ATP-dependent protease and is essential for the development of Myxococcus xanthus. J. Bacteriol. 175:4545-4549. Tojo, N., and T. Komano. 2003. The IntP C-terminal segment is not required for excision of bacteriophage Mx8 from the Myxococcus xanthus chromosome. J. Bacteriol. 185:21872193. Ueki, T., S. Inouye, and M. Inouye. 1996. Positive-negative KG cassettes for construction of multi-gene deletions using a single drug marker. Gene 183:153-157. Whitworth, D. E., S. J. Bryan, A. E. Berry, S. J. McGowan, and D. A. Hodgson. 2004. Genetic dissection of the lightinducible carQRS promoter region of Myxococcus xanthus. J. Bacteriol. 186:7836-7846. Wu, S. S., and D. Kaiser. 1996. Markerless deletions of pi1 genes in Myxococcus xanthus generated by counterselection with the Bacillus subtilis sacB gene. J. Bacteriol. 1785817-5821. Wu, T. T. 1966. A model for three-point analysis of random generalized transduction. Genetics 54:405-410. Youderian, P., N. Burke, D. J. White, and P. L. Hartzell. 2003. Identification of genes required for adventurous gliding motility in Myxococcus xanthus with the transposable element mariner. Mol. Microbiol. 49555-570. Youderian, P., and P. L. Hartzell. 2006. Transposon insertions of magellan-4 that impair social gliding motility in Myxococcus xanthus. Genetics 172:1397-1410.
Myxobacteriu: Mctlticellctlarity and Differentiation Edited by David E. Whitworth 0 2008 ASM Press, Washington, D.C.
Anke Treuner-Lange Sabrina DOG Tina Knauber
Sorangium cellulosurn Methods
The industrial importance of Sorangium species, which produce a variety of bioactive compounds such as the antitumor agent epothilone (Gerth et al., 1996), the antifungal soraphens (Gerth et al., 1994) or ratjadons (Gerth et al., 1995; Koster et al., 2003), and the antibacterial sorangicins (Irschik et al., 1987), has led to a revival of research with these bacteria. Sorangium cellulosum has been the focal species of interest, and the genome sequence and function of strain So ce56 are being analyzed in a project initiated in 2001 within the “GenoMik” network funded by the German Ministry of Education and Research (BMBF) (Gerth et al., 2003). Knowledge of an organism’s genome is extremely valuable when applying modern molecular methods. Nonetheless, prior to the sequencing of its genome, S. cellulosum had already been extensively investigated with respect to its metabolism, enzyme repertoire, and capacity to form multicellular fruiting bodies. The goal of this chapter is to summarize research methods to encourage further research into the complex biology of S. cellulosum.
30
CULTIVATION S. cellulosum is a cellulolytic myxobacterium that can grow on simple mineral medium with KNO, as the sole nitrogen source and cellulose as the sole carbon source. In 1967 Coucke and Voets attempted to formulate a culture medium for optimal growth of a Polyangium cellulosum strain (Coucke and Voets, 1967). In older literature the name Polyangium cellulosum was used according to the 8th edition of Bergey’s Manual of Systematic Bacteriology of 1974. Although the name Sorangium cellulosum has still not been validated, most authors are using this name now following the suggestion to use the genus name Sorangium next to Polyangium to separate the cellulose-degrading S. cellulosum from other noncellulolytic Polyangium species (Reichenbach and Dworkin, 1992; Reichenbach, 1993; Sproer et al., 1999). Maximal growth, determined by the percentage of available cellulose that had been degraded, was observed in the presence of KNO,, MgSO,, CaCl,, K,HPO,, and FeSO, (Table 1) (Coucke and Voets, 1967). Neither manganese nor NaCl was found to be essential for growth, and significant cellulose degradation was observed in a pH range
Anke Treuner-Lange, Sabrina Dog, and Tina Knauber, Justus-Liebig-University Giessen, Department of Microbiology and Molecular Biology, Heinrich-Buff-Ring 26-32, D-35392 Giessen, Germany.
503
MYXOBACTERIAL METHODS
504 Table 1 Defined media for growth of S. cellulosuma
YOConcn in: Element C
N
Compound
cv
Cellulose G1u cose Cellobiose Maltose Mannose
2
KNO,
0.2
CIZ 1 1
CK6
Stanier
0.5 0.25
0.2
0.2
0.1 or
Asparagine
0.5
Mg
MgSO4.7H20
0.05
Ca
CaC1, CaC1,.2 H,O
0.125
K,HPO, FeSO,-7 H,O Fe3+-citrate FeC1,.7 H,O FeC1, Na-Fe-EDTA
0.025 0.001
HEPES PH
1 (also referred to as SG) 1 (also referred to as SC) 1 (also referred to as SM) 1 (also referred to as SMa)
0.1
(NH4)2S04
P Fe
S
0.3
0.15
0.18
0.15
0.025 0.002 0.002
0.025
0.02
0.05
0.01 0.05 0.1
0.006
0.0008 0.002 0.0008 5.6
7.0 -7.5
1.2 7.2
of 4.5 to 8.7 with an optimum of pH 5.6. The maximum growth rate was observed at 34 to 35°C. This medium has been slightly modified by other researchers (Irschik et al., 1987; Kleining et al., 1971),in particular by replacing cellulose with glucose as the carbon source (Table l). Peterson (1969) observed more robust growth both in liquid and on agar medium by using Stanier’s medium containing 0.25 % cellobiose rather than glucose (Table 1) (Peterson, 1969a, 1969b; Stanier, 1942). Krzemieniewski and Krzemieniewska (1937) and others (Kegler et al., 2006; Miiller and Gerth, 2006) have used asparagine rather than KNO, as a nitrogen source (Table 1). Coucke (1969) reported that S. cellulosum utilizes some carbon and nitrogen sources preferentially over others. (Although Coucke referred to a strain of S. compositum formerly called Polyangium cellulosum, the strain description in this publication suggests that he actually worked with a strain of S. cellulosum.) Carbon sources such as dextrin, mannose, maltose, starch, and cellobiose were shown to give high protein yields after similar growth periods. Growth on 2% glucose did not generate a high protein yield (Coucke, 1969), but it has been reported elsewhere that much lower glucose concentrations (slightly higher than 0.25%) are toxic to S. cellulosum cells (Peterson, 1969a). Carbon sources such as lactose, mannitol, dulcitol, dextran, chitin, and
melibiose do not support robust growth of S. cellulosum (Coucke, 1969). Nitrate, urea, ammonium, and the amino acids L-glutamate or L-asparagine can be used as nitrogen sources (Coucke, 1969). In Stanier’s medium or complex media such as Casitone broth (0.25% Casitone, 0.05% MgS04.7 H,O, 0.0025% K,HP04) or Casitone broth with 0.1% glucose, S. cellulosum doubling times of approximately 22 to 24 h have been reported (Sarao et al., 1985). Small amounts of peptone (0.05% peptone from a tryptic digest of casein) were found not only to improve growth but also to keep cells in a dispersed state during growth (Reichenbach and Dworkin, 1992). Generation times of 6 to 8 h have been reported for strain So ceS6 at 34°C in M or P medium (Table 2) (Pradella et al., 2002), but we frequently observe doubling times of 10 to 14 h when growing So ceS6 in these media (our unpublished data). For long-term growth experiments S. cellulosum cultures can be protected from contamination by adding gentamycin, neomycin, or kanamycin to cultures or plates, as the organism is naturally resistant towards these compounds (Reichenbach and Dworkin, 1992). Some authors (Reichenbach and Dworkin, 1992; Yan et al., 2003) reported the use of a trace element solution to enhance growth (Reichenbach and Dworkin, 1992), but we have never observed such an enhancement when
30. S .
CELLULOSUM
METHODS
Table 2
Different complex media”
505
% Concn in:
Element C
N
Compound
M medium
Maltose Glucose Mannose Cellobiose Starch
1.o
Soy peptone Peptone Yeast extract Probion
1.0
P medium
1 0.2
PM medium 0.35
0.8 0.2 0.2 0.4
0.05
0.05
(NH4)2S04
Mg
MgS04.7H,0
0.1
0.1
0.15
Ca
CaC1,
0.1
0.075
0.075
P
K,HP04
Fe
Na-Fe-EDTA
0.0008
0.0008
0.0008
HEPES Sodium dithionite PH
1.2
2.4
1.2 0.014
7.2
7.5
7.4
0.006
“Data from Pradella et al., 2002.
adding this trace solution to strain So ce56 (our unpublished data).
DEVELOPMENTAL ASSAYS Under appropriate starvation conditions, S. cellulosum cells initially accumulate in masses that form round structures. These primary cysts (-150 pm in diameter), or sori, are often connected in clusters by their adhesive slime walls. The sori are a red-brown color, in contrast to the bright orange color of vegetative colonies. At a later stage of development, circular to oval secondary cysts, or sporangia, with solid walls (8 to 15 pm in diameter) are formed within the primary cysts. This appears to be achieved by the secretion of slime that encompasses smaller cell masses. These sporangia contain the microcysts or spores. Sporangia can number from very few to hundreds within a single fruiting body. Sporangia can contain from 10 to more than 100 microcysts, with an average between 20 and 30 (Peterson, 1969a). These sporangia are of distorted polygonal shape (depending upon the pressure from surrounding sporangia) and have a quite rigid membrane that can resist several cycles of sonication. Reichenbach described the morphogenesis of Sorungiineae as rudimentary because the microcysts are hard to distinguish from vegetative cells (Reichenbach,
1974). Because of this, the terms “sporulation” or “formation of spores” were avoided early in the last century when describing Polyangium morphogenesis (Baur, 1905).The rod-shaped microcysts are described as being somewhat shorter than vegetative rods (1 to 3 pm by 0.5 to 0.8 pm and 2 to 4 pm by 0.5 to 0.8 pm, respectively) (Coucke, 1969; Lampky, 1976). The microcysts are not light refractile, and transmission electron microscopy analysis indicated the absence of a definite capsule (Lampky, 1976).Nonetheless, fruiting bodies of an S. cellulosum strain were shown to consist of a similar proportion of polysaccharides as fruiting bodies of Myxococcus xanthus (Sutherland, 1979). The only remarkable difference found between these species was a total lack of rhamnose in the exopolysaccharide fraction isolated from two different S. cellulosum strains (Sutherland, 1979) and the fact that vegetative slime of S. cellulosum does not bind Congo red as does slime from M. xanthus (Arnold and Shimkets, 1988; McCurdy, 1989).The cell membranes of the microcysts of S. cellulosum appear to develop irregular waves that were not observed on vegetative cells (Lampky, 1976). We have developed a reproducible protocol that induces fruiting body formation by S. cellulosum strain So ce56, which grows as dispersed cells in liquid culture (Knauber et al., in press). Following this protocol morphogenesis is completed within 90 to 100 h (Fig. 1).
Protocol for Development on P-Diff Agar 1. Cells are grown in M medium (Table2) to an optical density at 600 nm (OD,,, n m ) of -4.0 (-5 x lo9cells/ml). 2. Cells are harvested at 13,000 rpm for 4 min and washed once with M medium. 3. The cells are then resuspended in M medium to generate a concentrated cell suspension with an OD,,, nm of 25. 4. Development is initiated by spotting 50 pl of this suspension onto P-Diff agar (0.005% starch, 0.002% peptone, 0.001% Probion, 0.05% MgS0,.7 H,O, 0.05% CaCl,, 1.2% HEPES [pH 7.21, 1.5% agar). The wells of a 12-well plate are filled with 2 or 3 ml of P-Diff agar. 5. After spotting the cells, the spots are dried in a laminar-flow hood and incubated at 32°C in a humid incubator. Fruiting body formation of S . cellulosum strain So ce56 is not affected by light. Reproducible fruiting body formation is difficult to obtain when nutrients are removed quickly, for example, if cells are washed with buffer or spotted onto agar medium lacking nutrients immediately after growth in nutrient-rich medium. Also, cultures should not undergo more than two successive cycles of growth in liquid prior to initiation of development. If such cultures need to be
MYXOBACTERIAL METHODS
506
H
1 rnrn
Figure 1 Morphogenesis of S. cellulosum strain So ce56 on P-Diff agar. At time point zero the cells were spotted onto the agar. During phase 1 no visible changes occur. Phase 2 is the early aggregation phase. Primary cyst (sori) formation is achieved in phase 3. Here several slime-connected sori are formed. Within these sori, secondary cysts (sporangia) appear in phase 4. These sporangia contain 20 to 30 microcysts on average.
used, it is recommended that they be subjected to a growth cycle on an agar surface prior to regrowth in a new liquid culture. The Krzemieniewskis observed that microcyst formation depends on the type of carbon source. For the strain they examined, growth and microcyst formation occurred with dextrin, maltose, starch, glucose, and xylose but not with cellobiose, arabinose, and fructose (Krzemieniewski and Krzemieniewska, 1937). In contrast, strain So ce.56 has been observed to undergo fruiting body formation on agar medium containing cellobiose (A. Treuner-Lange, unpublished data). After exploring the carbon source preferences of S. cellulosum, Coucke concluded that cyst formation occurs on media in which the carbohydrates have been almost exhausted. Thus, rapid cyst formation occurs on media containing carbon sources that are consumed quickly (Coucke, 1969). Coucke also observed that fruiting body formation was inhibited below a threshold concentration of about 0.01 % KNO,, whereas fruiting bodies developed under the same conditions with 0.2% nitrate (Coucke, 1969). Distinct strains of S. cellulosum exhibit very different growth characteristics. As typical for myxobacteria,
most isolated strains of S. cellulosum do not naturally grow dispersed or submerged in the liquid medium itself but rather form flakes and clumps mostly at the airbroth interface. This property makes photometric methods with such isolates that grow in clumps impossible. However, with continuous growth and repeated transfers into fresh shaken liquid medium containing peptone (preferentially using the clump-free liquid as inoculum), strains exhibiting growth as dispersed cells are readily obtained. Such dispersed-growth strains are frequently defective at swarming and fruiting, and the evolution of these defects has been associated with the loss of fimbriae (Dobson et al., 1979) or changes in the slime (Grimm and Kuehlwein, 1973). The dispersed-growing So ce.56 strain shows severely reduced swarming on several agar surfaces but does form fruiting bodies when sufficiently concentrated cell suspensions are used (Fig. 2).On the lean growth agar VY/2 (0.5% baker’s yeast, 0.1YOCaCl,. 2 H,O, cyanocobalamin [0.5 p,g/ml], 1.5% agar) this strain barely moves, whereas the non-dispersed-growing S. cellulosum strain DSM14627 forms a big swarming colony. After swarming, fruiting body formation proceeds over several days in areas where nutrients have been
30. S. CELLULOSUM METHODS
50 7
Figure 2 Clusters of sporangia-filled sori of S. cellulosum So ce56 on P-Diff agar. The actual width of the image area is 1 mm.
depleted (Fig. 3 ) . Despite their very different swarming phenotypes, the fruiting bodies of the weak-swarmer So ce56 and the strong-swarmer DSM14627 look quite similar (Fig. 2 and 3). Swarming and morphology differences between the Soraphen A producer strain So ce26 and other related Sorungium species are described elsewhere (Yan et al., 2003). There is still no protocol for microcyst enumerations which gives reproducible numbers. This might be partly due to the observation that members of the suborder Sorangiineae do not readily produce swarms from single cells (Reichenbach and Dworkin, 1992). Also sporangia can be hard and tough, and the breaking of sporangia might affect microcyst viability. Nevertheless, with So ce56 it is even difficult to regrow a liquid culture when incubating several fruiting bodies in culture media. Therefore, more experiments need to be done to establish a germination and microcyst enumeration protocol.
frequency depends on the length of the cloned chromosomal fragment (Jaoua et al., 1992). Tenfold more transconjugants were obtained when integrating a 3.5-kb fragment rather than a 1.2-kb fragment (Jaoua et al., 1992). To our knowledge the smallest fragment successfully used to construct an S. cellulosum mutant was the 0.5-kb mglA fragment used to construct a So ce26 mutant (Zirkle et al., 2004). Depending on the plasmid used, media containing phleomycin (pSUP2021 [Simon et al., 19831 or pCIB132 [Schupp et al., 1995]), tetracycline (pSUP2021), and hygromycin (a modified derivate of pSUP102 [Simon et al., 19831 modified by a cloned hygromycin resistance gene [Pradella et al., 20021) can be used for selection of transconjugants. Genetic transformation of S. cellulosum is a low-efficiency process (Jaoua et al., 1992; Zirkle et al., 2004). The following protocol is used for transconjugation of So ce.56 in our laboratory.
Protocol for Conjugation of S. cellulosum So ce56
MUTAGENESIS Several mutants of S. cellulosum have been already constructed and described (Jaoua et al., 1992; Knauber et al., in press; Knauber, 2006; Dog, 2007; Kopp et al., 2004; Perlova et al., 2006; Pradella et al., 2002; Schupp et al., 1995; Zirkle et al., 2004). No plasmid that can replicate in S. cellulosum has yet been found. Broadhost-range plasmids such as RP4 or pME462 as well as the transposon Tn.5 failed to be successfully transferred and maintained (Jaoua et al., 1992). Nonetheless, plasmids harboring fragments of S. cellulosum chromosomal DNA can be integrated into the chromosome by homologous recombination. As expected, integration
1. S. cellulosum cells are grown in M medium up to an OD,,, nm of 10. 2. From this dense culture a 600-pl aliquot is used for conjugation with Escherichia coli. The best results have been obtained using E. coli strain S17-1 (Friedrich et al., 1981)transformed with pSupHyg-constructs for conjugation. These E. coli transconjugants are grown in LB medium overnight, diluted back to an OD,,, nm of 0.2, and harvested after they reach an OD,,, n n ~of 0.8. From these cultures, 200 pl is used for the conjugation. 3 . The E . coli cells are centrifuged, and after removal of the supernatant 600 pl of the So ce56 cell suspension is added. After another short centrifugation, 300 p1 of
MYXO BACTERIAL METHOD s
508
Figure 3 Morphogenesis of S. cellulosum strain DSM14627 on yeast VY/2 agar. (a) A young swarming colony. (b) The edge of the swarming colony photographed with a 1OOx objective. (c)The colony edge after 3 days; bar is 1 mm. (d) Enlargement of the right boxed area of panel c; bar is 0.1 mm. (e) Enlargement of the left boxed area of panel c; bar is 0.1 mm.
the supernatant is removed and the cells are gently resuspended in the remaining liquid. 4. The cell mixture is put on a Pc agar plate (0.50/, starch, 0.2% peptone, 0.1% Probion, 0.05% MgSO,. 7 H,O, 0.05% CaCl,, 1.2% HEPES [pH 7.2],1.5% agar) and incubated at 37°C for 2 days. 5. Afterwards the cells are scraped from the plate, resuspended in 1 ml of M medium, centrifuged, and after removal of 0.7 ml of supernatant, gently resuspended again. For each conjugation 100- and 2 0 0 - 4 aliquots are plated out on Pc agar plates supplemented with 100 pg/ml hygromycin and kanamycin. Following
this protocol it takes about 10 days for transconjugant colonies to appear.
So ce56 cells used for the conjugation should not have been previously passaged. Best results are obtained when using cultures initiated with cells coming directly from an M medium agar plate (S. Dog, unpublished data). This protocol is just one of several. Other groups have been successful using different E . coli strains and different cell numbers or preparations as well as different media for selecting transconjugants. For an overview of these methods see Table 3, and for further interesting details please refer to the cited publications. For example, Jaoua
30. S.
CELLULOSUM METHODS
509
et al. found out that a heat shock for 10 min at 50°C just before the conjugation of S. cellulosum SJ3 increased the conjugation frequency 10-fold (Jaoua et al., 1992). For random mutagenesis, a mariner-based transposon procedure has been developed (Julien and Fehd, 2003). A mariner-based transposon has been already used for M. xanthus that yields a high frequency of transposition and a more random distribution of insertions into the chromosome than does Tn5 (Youderian et al., 2003). The constructed plasmid pKOS183-3 (a conjugative plasmid harboring the mariner tnp gene and the mariner inverted repeats) inserted into the chromosome with a transposition frequency of to per cell and conferred resistance to phleomycin (Julien and Fehd, 2003). Interestingly, the transposition frequency was not increased by a heat shock of the So ce90 cells, nor did use of damand dcm-negative E. coli strains significantly affect the transposition frequency (for example, ET12567, which is often used for conjugation [Table 31). Another mariner-based transposon, pMycoMarHyg, that confers hygromycin resistance was reported to randomly integrate into So ce12 and So ce.56 (Kopp et al., 2004). We used the pMycoMarHyg transposon to construct developmental mutants of So ce.56, and some of the resulting mutant phenotypes are shown in Fig. 4. There is no literature concerning electroporation of S. cellulosum. We have never managed to electroporate So ce56 successfully by using electroporation protocols established for 111.xanthus.
BIOASSAYS The S. cellulosum strain So ce.56 produces the two known secondary metabolites chivosazole and etnangien (Gerth et al., 2003; Pradella et al., 2002). Qualitative bioassays
Table 3
are useful tools for analysis of physiological differentiation defects exhibited by mutants. However, such methods require knowledge of sensitive indicator strains. In the case of chivosazole, the sensitive yeast strains Hansenula anomala and Saccharomyces cerevisiae are used (Kopp et al., 2004; Perlova et al., 2006). The easiest way to perform a bioassay is to streak out the So ce56 wildtype strain and the mutants to test on a P medium agar plate. After sufficient growth of the strains the plates are overlaid with Myc-soft agar containing yeast (Perlova et al., 2006). After overnight incubation at 30°C chivosazole production can be estimated by comparing the growth inhibition zones above the colonies from the mutants and the wild type. A way to extract and concentrate secondary metabolites from S. cellulosum cultures is the use of XAD adsorber resins. The following protocol is used to test random So ce56/pMycoMar mutants for chivosazole production. Protocol for XAD Extraction from Different Stationary So ce56 Cultures So ce56 and mutants of interest are grown in 10 ml of Pro medium (0.8% starch, 0.2% glucose, 0.02% Probion, 0.1% CaC1,.2 H,O, 0.1% MgSO,.7 H,O, 1.2% HEPES, 0.0008% Na-Fe-EDTA, pH 7.4) containing 1 % XAD16. After the cultures have been shaken for 10 to 14 days at 32”C, the XAD is harvested by filtering. The XAD resin is dissolved in 1ml of methanol and incubated for 1 to 2 h. After letting the resin settle, the supernatant is further concentrated in a Speed-Vac concentrator to a final volume of 100 ~ 1 Two . small disks (0.5 cm) of Whatman paper for each tested strain are placed onto a Myc plate and covered with a layer of Myc-soft agar freshly inoculated with yeast. One microliter of the corresponding concentrated XAD-extract is then dropped
Comparison of different conjugation strategies S. cellulosum strains
E. coli strainshelper plasmid or strain
Plasmid to mobilize
Media for selection/ conjugation conditions
1992
SJ3 (So ce26 derivative)
W30UpME305
PSUP2021
S42-mediuma/40 h at 30°C
Jaoua et al., 1992
1995
SJ3
ED8767/pUZ8
PSUP2021
S42 medium
Schupp et al., 1995
2002
So ce56
ET12567/HB101 (RK600)
pSupHyg
P mediud40 h at 37°C
Pradella et al., 2002
2004
So ce26
ET12567/pUZ8002
PCIP132
S42 mediud40 h at 30°C
Zirkle et al., 2004
2004,2006
So ce56
ET12567/pUB307
pSupHyg
P mediud40 h at 37°C
Kopp et al., 2004; Perlova et al., 2006; Knauber et al., unpublished
Yr
Reference(s)
‘542 medium: 0.05% tryptone, 0.05% (NH,),SO,, 0.15% MgSO,.7H,O, 0.1% CaC1,.2 H,O, 0.006% KHJ’O,, 0.01% sodium dithionite, 0.0008% Na-Fe-EDTA, 0.35% glucose, 1.2% HEPES (pH 7.2), 1.5% agar, 3.5% (vol/vol)autoclaved supernatant of a stationary-phase culture of SJ3.
MYXOBACTERIAL METHOD s
510
S. cellulosurn
(W
S. ceNulosum S. cellulosum S. cel~ulosum S. cellulosurn S. cellulosum pMycoMar 31 pMycoMar 36
pMycoMar 17 pMycoMar 22 pMycoMar 24
differentiation phenotype
chivosazoleassay (ff. anomala) Figure 4 Developmental and physiological differentiation phenotypes of some So ce56 mariner mutants. The images of developing cells (first row) were taken after 7 days. The phenotypes shown are stable and did not change even with longer incubation. The second row shows the chivosazole assay phenotypes of the mutants compared to the wild type. Photographs were taken after overnight incubation. The experiments were performed according to the detailed protocols included in this chapter.
on these filters, and the plates are incubated overnight at 32°C. Chivosazole production can be estimated by comparing the growth inhibition zones around the filter disks containing XAD extracts from the mutants and the wild type (Fig. 4). When doing such experiments it is necessary to consider cultivation conditions and the medium used. Metabolite production can vary greatly as a function of medium type and growth conditions. For example, epothilone production in the native producer strain So ce90 was severely reduced in the presence of 12 mM ammonium whereas this concentration did not influence the growth of the cells (Regentin et al., 2003). Addition of 5 mM phosphate also reduced epothilone production (Regentin et al., 2003). Chivosazole production of So ce56 was reported to depend on temperature, aeration, nitrogen source, and carbon source (Miiller and Gerth, 2006). For example, in medium containing asparagines, chivosazole is produced strongly in the presence of 1% mannose, whereas the same concentration of glucose and maltose inhibits production (Miiller and Gerth, 2006). Interested readers are referred to this publication and to chapter 19 in this volume.
References Arnold, J. W., and L. J. Shimkets. 1988. Inhibition of cell-cell interactions in Myxococcus xanthus by Congo red. J. Bacteriol. 1705765-5770. Baur, E. 1905. Myxobakterien-Studien. Arch. Protistenk. 5~92-121.
Coucke, P. 1969. Morphology and morphogenesis of Sorangium compositum. J. Appl. Bacteriol. 32:24-29. Coucke, P., and J. P. Voets. 1967. The mineral requirements of Polyangium cellulosum. Z. Allg. Mikrobiol. 7:175-182. Dobson, W.J., H. D. McCurdy, and T. H. MacRae. 1979. The function of fimbriae in Myxococcus xanthus. 11. The role of fimbriae in cell-cell interactions. Can. J. Microbiol. 25:1359- 1372. DOG, S. 2007. Analyse der morphologischen und physiologischen Differenzierung in Sorangium cellulosum So ce56: Lon-Proteasen und Stickstoffmetabolismus. Ph.D. thesis. University of GieBen, GieBen, Germany. [Online: http://geb. uni-giessen.de/geb/volltexte/2007/4733/] Friedrich, B., C. Hogrefe, and H.G. Schlegel. 1981. Naturally occurringgenetic transfer of hydrogen-oxidizingability between strains of Alcaligenes eutrophus. J. Bacteriol. 147:198-205. Gerth, K., N. Bedorf, G. Hofle, H. Irschik, and H. Reichenbach. 1996. Epothilons A and B: antifungal and cytotoxic compounds from Sorangium cellulosum (Myxobacteria). Production, physico-chemical and biological properties. J . Antibiot. 49560-563. Gerth, K., N. Bedorf, H. Irschik, G. Hofle, and H. Reichenbach. 1994. The soraphens: a family of novel antifungal compounds from Sorangium cellulosum (Myxobacteria). I. Soraphen A1 alpha: fermentation, isolation, biological properties. ]. Antibiot. (Tokyo) 47:23-31. Gerth, K., S. Pradella, 0. Perlova, S. Beyer, and R. Miiller. 2003. Myxobacteria: proficient producers of novel natural products with various biological activities-past and future biotechnological aspects with the focus on the genus Sorangium. J. Biotechnol. 106:233-253. Gerth, K.,D. Schummer, G. Hofle, H. Irschik, and H. Reichenbach. 1995. Ratjadon: a new antifungal compound from Sorangium cellulosum (myxobacteria) production, physiochemical and biological properties. J. Antibiot. 48:973-976.
30. S.
CELLULOSUM
METHODS
Grimm, K., and H. Kiihlwein. 1973. Untersuchungen an spontanen Mutanten von Archangium violaceum (Myxobacterales) I. Bewegliche und unbewegliche Zellen von A. violaceum. Arch. Mikrobiol. 89:105-119. Irschik, H., R. Jansen, K. Gerth, G. Hofle, and H. Reichenbach. 1987. The sorangicins, novel and powerful inhibitors of eubacterial RNA polymerase isolated from myxobacteria. J. Antibiot. (Tokyo) 40:7-13. Jaoua, S., S. Neff, and T. Schupp. 1992. Transfer of mobilizable plasmids to Sorangium cellulosum and evidence for their integration into the chromosome. Plasmid 28:157-165. Julien, B., and R. Fehd. 2003. Development of a mariner-based transposon for use in Sorangium cellulosum. Appl. Environ. Microbiol. 69: 6299-6301. Kegler, C., K. Gerth, and R. Miiller. 2006. Establishment of a real-time PCR protocol for expression studies of secondary metabolite biosynthetic gene clusters in the G/C-rich myxobacterium Sorangium cellulosum So ceS6. J . Biotechnol. 121:201-212. Kleinig, H., H. Reichenbach, H. Achenbach, and J. Stadler. 1971. Carotenoid pigments of Sorangium compositum (Myxobacterales) including two new carotenoid glucoside esters and two new carotenoid rhamnosides. Arch. Mikrobiol. 78:224-233. Knauber, T. 2006. Der EinfluB der stringenten Kontrolle auf die morphologische und physiologische Differenzierung von Sorangium cellulosum So ce56. Ph.D. thesis. University of GieBen, GieBen, Germany. Knauber, T., S. DOSS,K. Gerth, 0. Perlova, R. Miiller, and A. Treuner-Lange. Mutation in the re1 gene of Sorangium cellulosum affects the morphological and physiological differentiation. Mol. Microbiol., in press. Kopp, M., H. Irschik, F. Gross, 0. Perlova, A. Sandmann, K. Gerth, and R. Miiller. 2004. Critical variations of conjugational DNA transfer into secondary metabolite multiproducing Sorangium cellulosum strains So ce12 and So ce56: development of a mariner-based transposon mutagenesis system. J. Biotechnol. 10729-40. Koster, M., S. Lykke-Andersen, Y. A. Elnakady, K. Gerth, P. Washausen, G . Hofle, F. Sasse, J. Kjems, and H. Hauser. 2003. Ratjadones inhibit nuclear export by blocking CRMU exportin 1. Exp. Cell Res. 286:321-331. Krzemieniewski, H., and S. Krzemieniewska. 1937. Die zellulosezersetzenden Myxobakterien. Bull. Int. Acad. Polon. Sci. Lettr. Classe Sci. Math. Nut. B (I):33. Lampky, J. R. 1976. Ultrastructure of Polyangium cellulosum. J. Bacteriol. 126:1278-1284. McCurdy, H. D. 1989. Order Myxococcales Tchan, Pochon and PrCvot 1948, 398, p. 2139-2170. In J. T. Staley, M. P. Bryant, N. Pfennig, and J. G. Holt (ed.), Bergey’s Manual of Systematic Bacteriology, vol. 3 . Williams & Wilkins, Baltimore, MD. Miiller, R., and K. Gerth. 2006. Development of simple media which allow investigations into the global regulation of chivosazol biosynthesis with Sorangium cellulosum So ce56. J. Biotechnol. 121:192-200. Perlova, O., K. Gerth, 0.Kaiser, A. Hans, and R. Muller. 2006. Identification and analysis of the chivosazol biosynthetic
51 1 gene cluster from the myxobacterial model strain Sorangium cellulosum So ce56. J. Biotechnol. 121:174-191. Peterson, E. A. 1969a. Isolation, cultivation and maintenance of the myxobacteria, p. 185-210. In J. R. Norris and D. W. Ribbons (ed.), Methods in Microbiology, vol. 3B. Academic Press, New York, NY. Peterson, J. E. 1969b. The fruiting myxobacteria: their properties, distribution and isolation. J. Appl. Bacteriol. 325-12. Pradella, S., A. Hans, C. Sproer, H. Reichenbach, K. Gerth, and S. Beyer. 2002. Characterisation, genome size and genetic manipulation of the myxobacterium Sorangium cellulosum So ce56. Arch. Microbiol. 178:484-492. Regentin, R., S. Frykman, J. Lau, H. Tsuruta, and P. Licari. 2003. Nutrient regulation of epothilone biosynthesis in heterologous and native production strains. Appl. Microbiol. Biotechnol. 61:451-455. Reichenbach, H. 1974. Die Biologie der Myxobakterien. Biol. Unserer Zeit 2:33-45. Reichenbach, H., and M. Dworkin. 1992. The Myxobacteria, p. 3416-3487. In A. Balows, H. G. Triiper, M. Dworkin, W. Harder, and K. H. Schleifer (ed.), The Prokaryotes. Springer Verlag, New York, NY. Reichenbach, H. 1993. Biology of the myxobacteria: ecology and taxonomy, p. 13-62. In M. Dworkin and D. Kaiser (ed.), Myxobacteria II. ASM Press, Washington, DC. Sarao, R., H. D. McCurdy, and L. Passador. 1985. Enzymes of the intermediary carbohydrate metabolism of Polyangium cellulosum. Can. J. Microbiol. 31:1142-1146. Schupp, T., C. Toupet, B. Cluzel, S. Neff, S. Hill, J. J. Beck, and J. M. Ligon. 1995. A Sorangium cellulosum (myxobacterium) gene cluster for the biosynthesis of the macrolide antibiotic soraphen A: cloning, characterization, and homology to polyketide synthase genes from actinomycetes. J. Bacteriol. 177:3 673-3 679. Simon, R., U. Priefer, and A. Puehler. 1983. A broad-host range mobilization system for in vivo genetic engineering: transposon mutagenesis in gram negative bacteria. BiolTechnology November:784-790. Sproer, C., H. Reichenbach, and E. Stackebrandt. 1999. The correlation between morphological and phylogenetic classification of myxobacteria. Int. J. Syst. Bacteriol. 49:1255-1262. Stanier, R. Y. 1942. The cytophaga group: a contribution to the biology of myxobacteria. Bacteriol. Rev. 6:143-196. Sutherland, I. W. 1979. Polysaccharides produced by Cystobacter, Archangium, Sorangium and Stigmatella species. J. Gen. Microbiol. 111:211-216. Yan, Z. C., B. Wang, Y. Z. Li, X. Gong, H. Q. Zhang, and P. J. Gao. 2003. Morphologies and phylogenetic classification of cellulolytic myxobacteria. Syst. Appl. Microbiol. 26:104-109. Youderian, P., N. Burke, D. J. White, and P. L. Hartzell. 2003. Identification of genes required for adventurous gliding motility in Myxococcus xanthus with the transposable element mariner. Mol. Microbiol. 49555-570. Zirkle, R., J. M. Ligon, and I. Molnar. 2004. Heterologous production of the antifungal polyketide antibiotic soraphen A of Sorangium cellulosum So ce26 in Streptomyces liuidans. Microbiol. Read. Engl. 150:2761-2774.
INDEX
Index Terms
Links
A A–/S– motility colony spreading assay
471
A-engine cooperation with S engine
94
polarity reversal
97
A-motility AglZ and
129
elasticotaxis and
109
frz gene regulation
126
genes required for
108
history of research
9
130
109
mechanism of
108
112
MglA protein and
116
118
regulation of gene transcription
118
slime secretion
112
113
surface lipoprotein exchange
116
117
swarming rates, surface effect on
25
26
swarming specialization
30
A-motility colony spreading assay A-signal
471 64
asg mutants
64
defining and quantifying
65
in fruiting body morphogenesis
79
history of research models of signaling system
6 65
linear phosphorelay system
66
network of several signals
67
reception and regulation
68
RodK
69
SasA
69
SasB
68 This page has been reformatted by Knovel to provide easier navigation.
123
Index Terms
Links
A1 defined minimal media
468
AbrB
364
act operon
476
7
71
180
292
ActA protein
84
173
ActB protein
84
156
ActC protein
84
ActD protein
84
Actin, in Dictyostelium discoideum
444
Actinomyces naeslundii biofilms
455
Acyl hornoserine lactones
454
Adenylyl cyclases, of Stigmatella auruntiaca
320
84
158
157
457
Adhesion of Caulobacter crescentus to surfaces
387
Adventurous motility, see A-motility Aggregatibacter actinomycetemcomitans
456
Aggregation C-signal and
79
83
Dictyostelium discoideum
439
443
FrzCD and
126
MXAN4899 and
86
overview
78
TodK and
86
AglA
112
AglU
112
AglW
112
AglZ
119
120
113
118
129
173
305
agm genes
288
Akinetes
410
Alanine, catabolism of
242
Alarmone
243
45
Aldehyde dehydrogenase, of Stigmatella aurantiaca
84
319
α Oxidation
249
alr genes
406
410
This page has been reformatted by Knovel to provide easier navigation.
86
130
Index Terms
Links
Altruism, in Dictyostelium discoideum
448
Amino acid catabolism alanine
242
243
arginine
242
243
asparagine
242
243
aspartate
242
cysteine
242
glutamine/glutamate
243
245
glycine
244
245
histidine
244
246
isoleucine
244
246
leucine
244
246
lysine
244
methionine
245
246
phenylalanine
245
247
proline
246
247
serine
246
247
threonine
246
tryptophan
247
tyrosine
245
valine
244
Amino acid-salts medium
476
Anabaena
399
Anabolic pathways
251
Anaeromyxobacter dehalogenans
173
Antarctica, myxobacterial species from
21
Antibiotics, production by Streptomyces
419
aphII gene
150
AppA
222
Arginine, catabolism of
242
Arthrospira platensis
411
246
405
408
410
174
243
asgA
64
66
asgB
64
66
asgC
64
66
asgD
52
53
asgE
64
66
This page has been reformatted by Knovel to provide easier navigation.
181
64
66
Index Terms
Links
Asparagine, catabolism of
242
Aspartate, catabolism of
242
Asticcacaulis biprosthecum
385
243
388
attB site Myxococcus xanthus
494
Stigmatella aurantiaca
322
Aurachin
289
Autoinducer -2(AI-2)
454
Avermectin
270
290
B B-signal
61
bsgA gene
61
BsgA protease substrates
63
nature of extracellular signaling
62
Bacillus subtilis multicellular behaviors
363
sporulation
364
Bacteriophages generalized transduction history of research BcsA
491 6 64
Bdellovibrio
351
ecology
354
hydrolytic enzymes
357
motility systems
355
phylogeny
352
predatory life cycle
351
type IV pili
355
β-Galactosidase assays
479
β-Lactamase
230
β-Methoxyacrylates
261
β Oxidation
249
480
This page has been reformatted by Knovel to provide easier navigation.
Index Terms
Links
Biofilms
453
dispersal
456
formation in M. xanthus
77
multispecies
456
signals and communication
453
acyl homoserine lactones
454
autoinducer-2 (AI-2)
454
cdiGMP
455
oligopeptides
454
outer membrane lipoproteins
456
rhamnolipids
455
Biogeography
21
bld genes
420
Blue light, induction of carotenogenesis
214
bofA/BofA genelprotein
375
brgE/BrgE gene/protein
71
Broth cultures
423
305
467
bsgA/BsgA genelprotein
61
150
157
81
96
C C-s ign a1 amplification loops
84
cis-acting DNA elements in C-signaldependent promoter regions
161
contact-dependent transmission
81
converging pathways on
85
in fruiting body morphogenesis
77
history of research
7
molecular identity of
80
reversal frequency regulation
96
synthesis
80
transduction pathway
7
transmission
7
Calcium, role in heterocyst development
400
This page has been reformatted by Knovel to provide easier navigation.
Index Terms
Links
CAMP
Dictyostelium discoideum aggregation and
439
in heterocyst signaling
401
synthesis
320
440
443
CarA
212
218
223
CarB
212
218
294
Carbohydrate utilization
249
217
221
glycolysis
250
phosphotransferase system
250
CarD
212
294
293
317 Cardiolipin
254
CarF
212
215
294
CarG
212
217
221
Carotenoid synthesis
211
che7 mutants
142
comparative genomics
294
212
215
294
212
215
219
215
219
294
history of research
5
induction by blue light
214
induction by copper
215
in non-Myxococcus species
221
signal reception and transduction
214
Stigmatella aurantiaca
322
structures genes and functions
212
CarQ
153
carQRS promoter
499
CarR
153 294
CarS
212
cas gene
290
Casein, density-dependent growth and
4
Catabolic pathways amino acid
242
carbohydrate
249
lipid
248 This page has been reformatted by Knovel to provide easier navigation.
Index Terms
Links
Catabolic pathways (Cont.) purine and pyrimidine Caulobacter crescentus
247 385
cell cycle
385
cell division
390
holdfast attachment
387
polar development
390
stalk function and synthesis
388
surface attachment
387
ccbP
399
cdiGMP
455
Cell-cell signaling in early development
60
historical overview
61
population starvation recognition
60
A-signal
64
B-signal
61
Cell density (quorum) sensing, in Dictyostelium discoideum
442
Cell division Caulobacter crescentus
390
Streptomyces coelicolor
430
Cell envelope
229
ECM (extracellular matrix)
233
biogenesis
236
regulation of EPS production
234
structure and function
233
LPS
232
outer membrane proteins
231
peptidoglycan
229
periplasmic motility apparatus
231
periplasmic proteins
231
pili
237
Cell tracking dye-based
475
GFP-based
474 This page has been reformatted by Knovel to provide easier navigation.
Index Terms
Links
Cell wall, of Streptomyces coelicolor
423
Ceramide biosynthesis
254
CF (clone fruiting) medium
476
cglB/CglB geneIprotein
10
426
100
116
119
139
288
231 Chaplins
427
Che1 chemosensory system, see Frz system Che2 chemosensory system, see dif/Dif genestproteins Che3 chemosensory system
52
137
Che4 chemosensory system
141
288
Che5 chemosensory system
141
Che6 chemosensory system
141
Che7 chemosensory system
142
Che8 chemosensory system
142
CheA
125
128
129
135
137
141
142
174
235
291
142
143
CheAY-like two-component signal transduction (TCST) systems CheB
174 125
136
141
144
174
183
CheC
144
145
235
Chemosensory system
135 139
Che3
52
137
Che4
141
288
Che5
141
Che6
141
Che7
142
Che8
142
Dif (Che2)
138
144
Frz (Che1)
123
138
genetic organization
138
regulation
144
143
This page has been reformatted by Knovel to provide easier navigation.
188
Index Terms
Links
Chemosensory system (Cont.) temporal control
145
two-component signal transduction (TCST) systems
135
Chemotaxis comparative genomics
291
Dictyostelium discoideum
444
in M. xanthus
126
predation and
27
CheR
125
128
136
141
125
128
135
141
183
235
125
128
135
137
141
142
173
174
177
235
334
144 Chew
CheY
CheZ
185
Chivosazols
260
261
276
337
340
509
Chondromyces crocatus, secondary metabolism of
262
Chromosome partitioning, in Streptomyces coelicolor
430
cis-acting DNA elements in C-signaldependent promoter regions
161
Cisplatin
447
Clone fruiting (CF) medium
476
Clp proteins
392
CodY
364
ComP
364
Comparative genomics
287
carotenogenesis
294
development
293
gas vesicle genes
292
This page has been reformatted by Knovel to provide easier navigation.
Index Terms
Links
Comparative genomics (Cont.) motility
287
secondary metabolism
288
signaling and chemotaxis
291
Competence-stimulating peptide (CSP)
454
ComX pheromone
364
Conditioned medium factor (CMF)
439
Copper, induction of carotenogenesis
215
countin gene
446
CrdA
442
52
58
137
156
157
175
213
218
294
7
24
49
70
71
79
84
96
97
157
292
CrdB
140
CrdC
141
CrtB
213
CrtC
214
CrtD
214
CrtE
213
crtl
212
CrtYc
213
CrtYd
213
csgA/CsgA gene/protein
287
CSP (competence-stimulating peptide)
454
cspA promoter, of Stigmatella aurantiaca
323
CtpB
376
CtrA
389
CTT media
476
CTT-YE media
466
476
Cultivation Myxococcus xanthus
466
equipment
466
general considerations
466
media
466
storing strains
467
Sorangium cellulosum
503
This page has been reformatted by Knovel to provide easier navigation.
139
Index Terms
Links
Cyanobacteria differentiation
410
heterocysts
397
developmental signals
398
maturation
406
pattern formation
401
structure and functions
397
predation on
26
secretion-mediated gliding in
109
symbiosis
401
112
Cyclic AMP, see CAMP Cylindrospermum
410
Cyrmenins
260
Cysteine, catabolism of
242
Cystobacter fuscus
254
Cystobacterineae
19
Cytoskeleton, Dictyostelium discoideum
261
276
444
D Database resources
108
ddc gene
337
Defined media
466
Deletions, in-frame
497
Density-dependent growth on casein Development
476
4 43
Caulobacter crescentus cell-cell signaling
385 60
comparative genomics density required
293 43
Dictyostelium discoideum
439
heterocysts
398
inducing conditions
44
initiation
44
methods, Myxococcus xantbus
468
extracellular complementation assay
470
induction protocols
468
This page has been reformatted by Knovel to provide easier navigation.
504
Index Terms
Links
Development (Cont.) molecular analysis
470
rapid spore formation
469
sporulation assay
469
starvation media
468
starvation recognition cellular
43 44
integration of cellular and population recognition pathways population
69 60
steps
43
Streptomyces
419
transcriptional regulation
149
44
Developmental timers
50
DevT protein
83
84
8
83
devTRS operon
292 Dictyostelium discoideum
439
aggregation
439
altruism
448
cell density (quorum) sensing
442
cell type choice
442
conditioned medium factor (CMF)
439
ease of working with
440
group size, regulation of
445
human disease studies
447
as model system
439
morphogenesis of groups of cells
446
motility
444
myxobacteria compared
448
starvation, sensing
441
443
442
dif/Dif genes/proteins comparative genomics
288
EPS production regulation
235
regulation
145
Y2H screen
305 This page has been reformatted by Knovel to provide easier navigation.
84
163
Index Terms
Links
Differential fluorescence induction Digestion, communal
323 29
Disorazol
343
Dispersal
22
DIV proteins
29
391
Diversity, of myxobacteria
19
DksA homologues in M. xantbus
47
DKxanthene
263
DNA-binding domains
182
dofA gene
84
dsp (dispersed growth) locus
8
Dworkin, Martin
5
Dye-based cell tracking
9
475
E EBPs, see Enhancer binding proteins (EBPs) ECF (extracytoplasmic function) sigmas
152
ECM (extracellular matrix) biogenesis
236
regulation of EPS production
234
structure and function
233
Ecology abundance
17
determinants of genotypic performance
25
distribution
17
diversity
19
local population
23
predation
26
research strategies for investigating
18
sociobiology
28
spatial structure in global population
22
21
sporulation efficiency and population density surface effect on swarming Elasticotaxis Elongation factor EF-TU
25
27
25
26
9
109
317
This page has been reformatted by Knovel to provide easier navigation.
235
288
Index Terms
Links
Enhancer binding proteins (EBPs) description
54
with early developmental defects
57
examples
156
ActB
156
157
CrdA
156
157
MrpB
156
158
Mx4885
156
159
Nla1
156
Nla4
156
159
Nla6
156
159
Nla18
156
159
Nla23
156
Nla24
156
Nla28
156
PiIR
156
SasR
156
158
SpdR
156
157
55
156
155
156
54
57
259
270
271
333
343
503
265
266
table of transcriptional activation with vegetative growth defects Epothilone
159
EPS, see Exopolysaccharide (EPS) Erythromycin synthesis Escherichia coli peptidoglycan
230
stringent response
45
two-component transduction (TCST) systems
136
EsgA
236
EshA protein
47
esp locus
51
Ether lipid biosynthesis
255
Etnangien
509
180
This page has been reformatted by Knovel to provide easier navigation.
236
273
Index Terms
Links
Evolution of cooperation
28
ecology and
17
escape from social defects
33
experimental
18
predation
26
reevolution of social swarming
34
research strategies for investigating
18
Exopolysaccharide (EPS) biofilm
453
biogenesis
236
function
233
genes involved in biogenesis
109
genomic screen
300
regulation of production
234
S motility and
108
as type IV pili anchor
233
Experimental evolution Expression analysis
455
456
114
18 479
β-galactosidase assays
479
480
microarray experiments
479
480
482
quantitative PCR
479
480
481
293
318
Extracellular complementation assays
470
Extracellular matrix, see ECM (extracellular matrix)
F fab genes
252
Fatty acid elongation
252
Fatty acid primer synthesis
251
fbf genes
292
fdgA gene
84
fibA gene diversity
24
mutants
234
fibR/FibR gene/protein
236
236
This page has been reformatted by Knovel to provide easier navigation.
319
Index Terms
Links
Fibrils, see also ECM (extracellular matrix) exopolysaccharide component of history of research Films, myxobacteria
114 8 4
Flaviolin
265
Freezer stocks
467
Frizilator
96
100
130
Frizzy mutants
11
71
81
84
161
163
303
see also Frz (frizzy) system fruA/IFruA gene/protein
DNA-binding domain
183
multiple partners for
184
NarXL-like signal system
177
receiver domains
173
fruE gene
293
Fruiting body packing spores in
30
sporulation without
30
Fruiting body development/morphogenesis C-signal, function of
77
Dictyostelium discoideum
445
fbf genes
318
history of research intracellular signaling during motility genes and
5 79 119
population density, effect of
25
signal integration
87
stages
78
Stigmatella aurantiaca
319
318
319
Frz (frizzy) system
95
123
C-signal and
83
frz operon
125
pathway output
129
predation, effect on
27
This page has been reformatted by Knovel to provide easier navigation.
96
Index Terms
Links
Frz (frizzy) system (Cont.) regulation
128
sequence similarities to chemotaxis proteins
125
FrzB
128
FrzCD
83
87
95
125
127
145
frzE/FrzE gene/protein
11
128
gliding reversal and
94
98
FrzF
83
97
FrzG
97
173
FrzS
99
115
FrzZ
178
FtsZ
365
Functional genomics
306
Fungi, predation on
26
methylation
Fusobacterium nucleatum
127
129
458
G galK gene
497
γ-Butyrolactones
424
Gas vesicle genes
292
GcrA
392
Gene density
11
Gene duplication
12
195
C-signal and
79
84
TodK and
86
Gene expression
transcriptional regulation mechanisms during development
149
Gene expression mapping
307
Gene families, paralog analysis of
302
Gene fusions
498
Gene organization, of Protein Ser/Thr kinases (PSTKs)
196
This page has been reformatted by Knovel to provide easier navigation.
125
87
173
Index Terms
Links
Gene regulation, in cellular response to starvation
53
General transduction
491
Genetic diversity, in global M. xanthus population
22
Genetic drift
22
Genetic mapping, using transducing phages Genome sequence
491 11
Genomics
299
application to experimental biology
300
comparative
287
database resources
108
functional
306
gene expression mapping
307
genomic screens using transposable elements
300
microarray, M. xanthus
306
new experimental technologies
305
paralog analysis of gene families
302
phylogenetic mapping
307
Y2H assays
304
GFP-based cell tracking
474
Gliding motility
103
see also A-motility; S-motility comparative genomics
287
genes involved
106
109
A-motility
108
112
S-motility
114
mechanisms
MglA protein, role of
116
multicellular development and
119
mutant classes
104
mutant phenotypes
105
overview
103
polar oscillation of TFP
115
regulation
118
107
This page has been reformatted by Knovel to provide easier navigation.
110
Index Terms
Links
Gliding motility (Cont.) reversal of direction surface lipoprotein exchange
93
285
Globomycin
140
Gluconacetobacter xylinus
455
Glucosaminidase, of Stigmatella aurantiaca
320
Glucose inhibition division (GidA) protein
231
Glutamine/glutamate, catabolism of Glycerol induction myxospores
127
129
407
411
116
GLIMMER (program)
Glucose, peptidoglycan-linked
124
4 243
245
3
Glycine, catabolism of
244
Glycogen consumption and sporulation
251
Glycolysis
250
GTP gene
46
Growth conditions, diversity of
21
245
GTP- binding protein, of Stigmatella aurantiaca
321
Guanosine-5´-(tri)di-3´-diphosphate see (P)PPGPP
H Halotolerant species
21
Heat shock proteins
231
hepA
411
hepN
410
HetC
407
Heterocysts
397
developmental signals
398
maturation
406
pattern formation
401
structure and functions
397
HetF
404
HetN
405
HetP
408
hetR/HetR gene/protein
400
319
408
402
This page has been reformatted by Knovel to provide easier navigation.
Index Terms
Links
hfa genes
387
388
hfs genes
387
High-mobility group A (HMGA) proteins
217
Histidine ammonia lyase (HAL)
338
Histidine, catabolism of
244
246
Histidine kinases
169
181
see also Two-component signal transduction (TCST) systems bifunctional
185
transmembrane
181
Holdfast, of Caulobacter crescentus
387
Homologous recombination
340
Horizontal gene transfer
24
Hot phenol RNA isolation
481
hsf genes
181
hspA/HspA gene/protein
293
HvrA protein
482
319
47
Hybridization of amino-silane-coated microarrays
485
Hydrolytic enzymes
357
Hyphomicrobium neptunium
385
I Icumazol
334
ihfA/IhfA gene/protein
218
In-frame deletions
497
Incompatibility, social
294
36
Indole
319
Inducible promoter systems
499
infB gene
317
Inositide degradation, in Stigmatella aurantiaca
320
Inositiol phospholipid synthesis, in Stigmatella aurantiaca
320
Insertion library plasmid
496
IPTG
499 This page has been reformatted by Knovel to provide easier navigation.
Index Terms
Links
Isoleucine, catabolism of
244
246
Isoprenoid biosynthesis
270
274
J Junctional pore complex
109
K kefC/KefC gene/protein Kin selection
141
181
30
KipI
184
Kiihlwein, Hans
5
L Legionella pneumophila
447
Leucine, catabolism of
244
246
Leupyrrins
260
261
Light-regulated carotenogenesis
214
221
Linkage disequilibrium
24
Lipases
248
Lipid biogenesis
251
ceramide and sphingolipid biosynthesis
254
fatty acid elongation
252
fatty acid primer synthesis
251
phospholipid biosynthesis
253
Lipid catabolism α oxidation
249
lipases
248
Lithium
447
litR/LitR gene/protein
222
litS/LitS gene/protein
222
lonD/LonD gene/protein
150
152
181
494
LPS (lipopolysaccharide)
124
232
301
302
Lysine, catabolism of
244
Lysophosphatidylethanolamine
322
This page has been reformatted by Knovel to provide easier navigation.
Index Terms
Links
M Magellan-4 transposon mutagenesis
107
300
Malonyl-coenzyme A
265
266
mariner-based transposon mutagenesis
509
510
masK gene
118
236
MBHA
231
MC7 buffer
476
MCP, see Methyl-accepting chemosensory proteins (MCP) Media
466
Metabolic pathway
241
amino acid catabolism
242
carbohydrate utilization
249
lipid biogenesis
251
lipid catabolism
248
purine and pyrimidine salvage
247
spore-specific products
255
Methanococcus jannaschii
455
Methanothermobacter thermautotrophicus
458
Methionine, catabolism of
245
Methods, Myxococcus xanthus
465
cultivation
475
246
466
equipment
466
general considerations
466
media
466
storing strains
467
development
468
extracellular complementation assay
470
induction protocols
468
molecular analysis
470
rapid spore formation
469
sporulation assay
469
starvation media
468
expression analysis β-galactosidase assays
479 479
480
This page has been reformatted by Knovel to provide easier navigation.
496
Index Terms
Links
Methods, Myxococcus xanthus (Cont.) microarray experiments
479
480
482
quantitative PCR
479
480
481
95
125
135
104
116
118
genetic tools
491
directed mutagenesis
496
gene fusion
498
general transduction
491
inducible promoter systems
499
plasmid DNA transfer
493
plasmid integration at phage attachment site
494
random mutagenesis motility –
494 470
–
A /S motility colony spreading assay
471
A-motility colony spreading assay
471
dye-based cell tracking
475
GFP-based cell tracking
474
media recipes
475
molecular analysis
475
S-motility colony spreading assay
471
single-cell assays
472
social motility (TFP-dependent movement)
472
Methods, Sorangium cellulosum
503
bioassays
509
cultivation
503
developmental assays
505
mutagenesis
507
Methyl-accepting chemosensory proteins (MCP) mglA/MglA gene/protein EPS production and
236
gliding reversal and
93
97
mglB/MglB gene/protein
97
116
Microarray, M. xanthus
306
Mkaps
200 This page has been reformatted by Knovel to provide easier navigation.
118
144
Index Terms
Links
MMC medium
476
mmrA
236
Motility, see also A-motility; S-motility C-signal dependent response
86
comparative genomics
287
Dictyostelium discoideum
444
history of research
8
A-motility
9
elasticotaxis
9
fibrils
8
reversal
10
S-motility
8
stimulation
10
methods, Myxococcus xanthus
470
A–/S– motility colony spreading assay
471
A-motility colony spreading assay
471
dye-based cell tracking
475
GFP-based cell tracking
474
media recipes
475
molecular analysis
475
S-motility colony spreading assay
471
single-cell assays
472
social motility (TFP-dependent movement)
472
swarming rates, surface effect on Movies, myxobacteria
25
26
4
mrp locus
58
mrpA/MrpA gene/protein
83
84
158
180
mrpB/MrpB gene/protein
83
84
156
158
83
84
153
159
160
180
303
289
321
180 mrpC/MrpC gene/protein
mta genes
This page has been reformatted by Knovel to provide easier navigation.
Index Terms
Links
Multicellularity Bacillus subtilis
363
Dictyostelizdm discoideum, initiation in
443
genetics of
11
motility genes and
119
Multicopy single-stranded DNA (msDNA)
316
Murein lytic transglycosylases (MLTs)
359
Mutagenesis directed
496
random
494
Sorangium cellulosum
507
Stigmatella aurantiaca
322
transposon
Mutation rate
6
106
300
340
341
387
406
424
494
509
272
290
289
291
36
Mx4 and Mx8 transducing phages
491
Mx4885
156
Mxa213
59
Mxa296
58
MXAN0206
81
MXAN2902
53
MXAN4899
86
MXAN5366
173
Myosin, in Dictyostelium discoideum
444
Myxalamides
263
159
84
270
291 Myxochelin
270
272
Myxochromides
270
275
Myxococcus stipitatus
254
Myxococcus virescens
248
Myxococcus xanthus carotenogenesis
211
cell envelope
229
chemosensory signal transduction system
135
contact-dependent signaling
77
This page has been reformatted by Knovel to provide easier navigation.
Index Terms
Links
Myxococcus xanthus (Cont.) Frz chemosensory system
123
genome sequence
285
genomics
285
gliding motility
103
299
methods cultivation
466
development
468
expression analysis
479
genetic tools
491
motility
470
phosphatases
202
polarity reversal
93
protein Ser/Thr kinases
191
secondary metabolism
263
207
transcriptional regulation during development Myxonemata
149 9
Myxospores Cystobacterineae
19
glycerol induction
3
Sorangiineae Myxothiazol
19
21
270
276
289
344
70
N Nannocystaceae
19
Nannocystineae
21
30
NarXL-like two-component signal transduction (TCST) systems ndk gene
177 46
Negative regulation
154
nif genes
405
Nitrogen fixation
397
nlaZ/Nlal gene/protein
156
236
nla4/Nla4 gene/protein
50
54
57
156
159
181
407
This page has been reformatted by Knovel to provide easier navigation.
Index Terms
Links
nla6/Mla6 gene/protein
59
156
159
181
nla18/Nla18 gene/protein
50
54
57
70
156
159
306
nla19/Nla19 gene/protein
236
nla23/Nla23 gene/protein
156
236
nla24/Nla24 gene/protein
119
156
236
nla28/Nla28 gene/protein
59
156
159
260
263
265
274
334
337
Nostoc
400
405
408
Nozzles
112
113
nrrA
400
Nsd
52
70
ntcA
399
402
Nonribosomally made peptides
NtrBC-like two-component signal transduction (TCST) systems Nucleases
176 359
Nutrient sensors
50
AsgD
53
Che3 operon
52
model of nutrient sensing
60
MXAN2902
53
Nsd
52
SigC
60
O O-antigen, lipopolysaccharide (LPS)
124
Oligopeptides, biofilms and
454
232
Oligotrophic environment, survival strategies for
385
ONPG
480
Oral biofilms
456
Oscillations
130
Oscillin
109
Outer membrane proteins
231
456
This page has been reformatted by Knovel to provide easier navigation.
408
269
410
Index Terms
Links
P P-Diff agar
505
Paralog analysis of gene families
302
Paralogous proteins
11
PAS domain
181
PatA
405
407
PatB
406
407
PatS
403
408
Pattern formation, in heterocysts
401
Pelotomaculum thermopropionicum
458
Peptidoglycan
3
Peripheral rods
123
Periplasmic proteins
231
PgpH protein
229
359
47
Phenylalanine ammonia lyase (PAL), encoding genes Phenylalanine, catabolism of
338 245
247
Pheromone Bacillus subtilis
364
Stigmatella aurantiaca
317
PhoA
140
PhoB
175
389
PhoR
175
390
Phormidium
109
112
11
251
254
322
445
Phosphatases, see Protein phosphatases Phosphatidylethanolamine
322 Phosphatidylglycerol
322
Phosphatidylinositol
254
Phosphoaspartate phosphatases
185
Phosphohistidine phosphatases
185
Phospholipid biosynthesis
253
in Stigmatella aurantiaca
322
Phosphorelays
177 This page has been reformatted by Knovel to provide easier navigation.
255
Index Terms
Links
Phosphorylation down-regulation
184
bifunctional histidine kinases
185
phosphoaspartate phosphatases
185
phosphohistidine phosphatases
185
Phosphotransferase system
250
phoU
176
phrA gene
294
Phycocyanobilin lyase
142
Phylogenetic mapping
307
Phylogeny (phylogenetics) pi1 genes of Bdellovibrio pilA gene
19
180
23
287
23
24
34
99
114
119
120
142
183
185
499
355
pilH gene
120
Pili
237 see also Type IV pili (TFP)
PilQ
10
99
116
PilR
119
156
183
PIS
119 114
115
PiIT
8
Pkn8
83
Pkn14
83
PktA2/Pknl
198
PktA5/PknDl
199
PktB8/PknD2
199
PktC2/Pkn8-PskA5/Pkn14 cascade
200
PktD1/Pkn4-PFK cascade
199
PktD7/Pkn5
198
PktD9/Pkn2
198
Plasmids DNA transfer to M. xanthus
493
insertion libraries
496
integration at phage attachment site
494
targeted insertions
496
Plate cultures
466 This page has been reformatted by Knovel to provide easier navigation.
185
Index Terms
Links
Ple proteins
390
PodJ
392
Polar oscillator of type IV pili
115
Polarity, reversal of
93
Polyelectrolyte slime
112
Polyketides
260
263
265
274
334
337
PoPC
80
Population density, sporulation efficiency and
25
Porphyromonas gingivalis
458
27
(P)PPGPP genes affecting
49
as internal starvation signal
45
role in development
47
stringent response and
45
ppk gene
46
PpsR
222
ppx gene
46
Predation by Bdellovibrio
351
chemotaxis and
27
diversity of
26
ecological effects of
26
evolution
26
prey lysis
27
searching rate, effect of
28
Proline, catabolism of
246
Promoter systems, inducible
499
Promoter trap vector
323
Proteases
358
Protein phosphatases
202
CPTP family
207
list of
205
LMMPTP family
207
overview
202
247
This page has been reformatted by Knovel to provide easier navigation.
269
Index Terms
Links
Protein phosphatases (Cont.) PhoRP family
203
phosphoaspartate
185
phosphohistidine
185
Pph1
203
PPM family
204
PPP family
204
Protein Ser/Thr kinases (PSTKs)
12
classification
192
domain structure
193
EBPs and
157
eukaryotic-like
192
gene duplication and
195
gene organization
196
genes, table of
194
identification in genome
192
in motility and development
202
mutational analysis
202
overview
191
paralog analysis of gene families
302
roles in M. xanthus life cycle
197
Mkaps
200
PktA2/Pkn1
198
PktASRknD1
199
PktBWPknD2
199
PktC2/Pkn8-PskA5/Pkn14 cascade
200
PktD1/Pkn4-PFK cascade
199
PktD7/Rkn5
198
PktD9/Pkn2
198
PskA2/Pkn6
198
PskA3/MasK
199
PskA4/Pkn9
198
two-component systems compared Protoporphyrin IX
207
159
160
191 5
pru gene
191
214
293
Pseudomonads, myxobacteria predation on
27
This page has been reformatted by Knovel to provide easier navigation.
Index Terms
Links
Pseudomonas aeruginosa biofilms
454
455
Pseudomonas putida
276
344
PskA2/Pkn6
198
PskA3/MasK
199
PskA4/Pkn9
198
PstSCAB
176
Public goods, genetic conflict over
180
456
389
31
Purine salvage
247
pykA mutant
319
Pyrimidine salvage
247
Pyrrolnitrin
333
Pyruvate kinase, of Stigmatella aurantiaca
319
320
Q Quantitative PCR
479
480
481
Quorum sensing A-signaling system Dictyostelium discoideum
65 442
R ram genes
425
RasA
236
reaA/ReaA gene/protein
153
Recombination
24
RedCDEF proteins
178
RegA
447
184
Regulation carotenoid synthesis
211
chemosensory systems
135
transcriptional regulation mechanisms during development
149
two-component signal transduction (TCST) systems
169
Reichenbach, Hans
4
relA gene
6
relAspoT-like homoIogues in M. xanthus
46
46
This page has been reformatted by Knovel to provide easier navigation.
48
Index Terms
Links
Retroelements, of Stigmatella aurantiaca
316
Reversal, gliding
93
124
Reverse transcription
481
484
Rhamnolipids
455
Rhodobacter
222
Rich media
466
476
C-signal and
79
83
overview
78
127
129
84
86
86
Rippling
RNA isolation protocol
481
482
RNA polymerase, see also Sigma factors Bacillus subtilis
365
Stigmatella aurantiaca
316
RodK
51
69
71
RpoN
54
59
389
RpoS homologue
59
150
rrpA
236
338
RsrA
153
S S-engine cooperation with A-engine
94
polarity reversal
99
S-motility ECM (extracellular matrix) function in evolutionary maintenance
233 30
exopolysaccharide and
114
frz gene regulation
126
FrzS and
129
genes required for
109
kin selection and
30
loss of
31
mechanism of
114
MglA protein and
116
110
34
118
This page has been reformatted by Knovel to provide easier navigation.
Index Terms
Links
S-motility (Cont.) regulation of gene transcription
118
surface lipoprotein exchange
116
117
swarming rates, surface effect on
25
26
swarming specialization
30
type IV pili
114
S-motility colony spreading assay
471
sacB gene
497
124
498
Saccharomyces cerevisiae, functional genomics of
306
saf genes
291
Saframycin
291
SapB
426
Sarcina lutea
9
sasA/SasA gene/protein
69
84
180
232
288 sasB
68
sasN/SasN gene/protein
71
158
180
SasR
58
156
158
180
SasS
158
364
Scout cells
123 159
184
263
265
269
263
265
269
sdeK/SdeK geneIprorein
71
85
305 Secondary metabolism
12
259
biosynthetic gene clusters
267
273
comparative genomics
288
identification of volatile compounds
261
in M. xanthus
263
molecular biology
265
nonribosomally made peptides
260 274
overview
259
polyketides
260 274
This page has been reformatted by Knovel to provide easier navigation.
Index Terms
Links
Secondary metabolism (Cont.) regulation
276
Sorangium celhilosum
333
509
Stigmatella aurantiaca
261
269
275
276
410
321 Ser/Thr protein kinases, see Protein Ser/Thr kinases (PSTKs) Serine, catabolism of
246
Sga15
189
sglK/SglK gene/protein
119
shd gene
247
236
46
Shewanella oneidensis
455
sigA/SigA gene/protein
150
151
316
sigB/SigB gene/protein
151
316
410
sigC/SigC gene/protein
60
151
316
410
sigD/SigD gene/protein
59
71
150
256
410 sigE/SigE gene/protein
151
410
sigF/SigF gene/protein
119
151
sigG/SigG gene/protein
151
371
374
316
410
Sigma factors Anabena
410
Bacillus subtilis
365
E
369
F
366
G
371
H
σ
365
σK
374
σ
σ σ
ECF (extracytoplasmic function) sigmas
152
SigA-G
150
Stigmatella aurantiaca
316
Signal peptidase II pathway
140
This page has been reformatted by Knovel to provide easier navigation.
Index Terms
Links
Signaling A-signal
6
biofilm communication C-signal
453 7
chemosensory systems
135
comparative genomics
291
history of research in Streptomyces coelicolor
6 424
SixA
185
Slime secretion
112
smlA gene
445
socD
141
181
49
70
SocE protein
113
Social motility, see S-motility Social motility (TFP-dependent movement) assays
472
Sociobiology
28
benefits of group living
29
cheating
31
communal digestion
29
group motility
30
incompatibility
36
physiological cost of cooperation
31
social divergence in natural populations
35
stress mitigation
29
Sorangicin Sorangiineae Sorangium cellulosum
334
503
19
21
329
503
bioassays
509
cell morphology
331
conjugation
507
cultivation
503
developmental assays
505
distribution
330
enrichment and isolation
330
fruiting bodies
332 This page has been reformatted by Knovel to provide easier navigation.
Index Terms
Links
Sorangium cellulosum (Cont.) genome size
337
history
329
molecular biology
335
mutagenesis
507
physiology
332
plant-like genes
337
secondary metabolism
269
swarms
332
taxonomy and systematics
329
276
333
509
174
184
186
333
334
344
57
63
156
157
84
87
two-component signal transduction system
173
XAD extraction of secondary metabolites Soraphens SpdR Spheroplasts, conversion to Sphingolpid biosynthesis spi gene
509
3 254 71
Spore stocks
158
467
Spores akinetes
410
ether lipid biosynthesis
255
Streptomyces
419
trehalose biosynthesis
256
Sporulation Bacillus subtilis
364
benefit of group
30
C-signal and
79
83
efficiency and population density
25
27
glycogen consumption
251
motility genes and
119
MXAN4899 and
86
Stigmatella aurantiaca
319
Streptomyces
419
TodK and
86
without fruiting body
30
431
This page has been reformatted by Knovel to provide easier navigation.
Index Terms
Links
Sporulation assay
469
SpoT
46
sqs gene
290
ssg genes
433
Stalks, of Caulobacter crescentus
388
70
Starvation C-signal activation fruiting body development types
79
84
5 44
Starvation media
468
Starvation plates
468
Starvation recognition cellular
476
43 44
developmenta1 timers
50
global gene regulation during response
53
model of overall response
60
nutrient sensors
50
stringent response
45
integration of cellular and population recognition pathways population
69 60
A-signal
64
B-signal
61
sti genes
290
Stigmatella aurantiaca
315
adenylyl cyclases
320
attB locus
322
carotenoids
322
CsgA
318
cspA promoter
323
fatty acid elongation
253
fatty acid primer synthesis
251
fbf genes
318
genome sequence
286
genome size
316
genomics
285
319
This page has been reformatted by Knovel to provide easier navigation.
Index Terms
Links
Stigmatella aurantiaca (Cont.) glucosaminidase
320
growth conditions
322
GTP-binding protein
321
HspA
319
inositide degradation
320
inositiol phospholipid synthesis
320
lipids of
322
mutagenesis
322
overview
315
pheromone
317
proteome 2D analysis of
323
retroelements
316
RNA polymerase
316
secondary metabolism
261
269
288
321
sporulation induction
319
transcription and translation factors
317
transcription factors
317
tryptophan catabolism
247
275
276
two-component signal transduction system Stigmatellin
173
174
184
186
270
272
276
290
291 Stigmolone
261
292
317
stk/Stk gene/protein
119
235
236
Storing strains
467
STPK (Ser/Thr protein kinases), see Protein Ser/Thr kinases (PSTKs) Streptococcus biofilms
454
455
457
Streptomyces avermitilis
223
263
425
433 Streptomyces coelicolor
222
cell division and chromosome partitioning
430
cell wall modifications
426
germination and apical growth
422
419
This page has been reformatted by Knovel to provide easier navigation.
430
Index Terms
Links
Streptomyces coelicolor (Cont.) life cycle overview
419
metabolic triggers
423
secondary metabolism
263
signaling and regulation
424
Stringent response
273
45
conditions inducing
44
in Escherichia coli
45
homologues in M. xanthus
46
(P)PPGPP and
45
Submerged culture
48
468
Surface type predation, effect on
27
swarming rate, effect on
25
swarming specialization
30
26
Swarming, see also Motility dominance
36
reevolution of social
34
reversal
10
Symbiosis, of nitrogen-fixing cyanobacteria
401
Synechococcus elongatus
399
T ta genes
291
TacA
389
TAL (tyrosine ammonia lyase)
338
tan mutants
263
Tap
144
Tar
135
Taxonomy
144
19
TCST, see Two-component signal transduction (TCST) systems TFP, see Type IV pili (TFP) Tgl/Tgl gene/protein
10
Themotoga maritima
455
Thermus thermophilus
223
99
This page has been reformatted by Knovel to provide easier navigation.
116
231
Index Terms
Links
Threonine, catabolism of
246
TipN
393
Tn5 transposable element
6
300
485
todK/TodK gene/protein
49
71
86
181
TPM media
476
tps gene
84
157
Transcription factors, of Stigmatella aurantiaca Transcription regulation activation
317 149 153
155
cis-acting DNA elements in C-signaldependent promoter regions
161
EBPs
155
FruA
161
MrpC
160
overview
155
of carotenogenesis
218
negative regulation
154
sigma factors
149
ECF (extracytoplasmic function) sigmas
152
SigA-G
150
Transcriptional activators, NtrC-like
302
Transduction, general
491
Translation initiation factor 2 (IF2)
317
Transmembrane histidine kinases
181
Transposon mutagenesis
156
6
106
300
340
341
387
406
424
494
509
Trehalose biosynthesis
256
Trg
144
Tryptophan, catabolism of
247
Tsr
144
tufB gene
317
This page has been reformatted by Knovel to provide easier navigation.
Index Terms
Links
Two-component signal transduction (TCST) systems
169
CheAY-like
174
complex systems
183
domain topology
136
functions input
181
output
182
gene neighborhood
180
gene organization
179
of M. xanthus
174
NarXL-like
177
NtrBC-like
176
overview
135
PhoBR-like
174
phosphorelays
176
phosphorylation down-regulation
184
bifunctional histidine kinases
185
phosphoaspartate phosphatases
185
phosphohistidine phosphatases
185
receiver and transmitter domains
173
signals
181
of Stigmatella aurentiaca
173
synteny of genes
180
table of
171
169
186
Type IV pili (TFP) Bdellovibrio
355
EPS production and
235
extension
114
genomic screen
300
gliding reversal
93
kin selection and
30
polar oscillator
115
retraction
114
S-engines
8
structure
124
99
124
237 This page has been reformatted by Knovel to provide easier navigation.
233
237
Index Terms
Links
Tyrosine ammonia lyase (TAL)
338
Tyrosine, catabolism of
245
V Valine, catabolism of
244
246
Vancomycin
423
Vaproic acid
447
Veillonella atypica
457
458
Vibrio biofilms
454
455
VxInsight
308
W whi genes
424
wzm gene
233
wzt gene
233
432
X XanthusBase
308
Y Y2H assays
304
YakA
442
Z Z buffer
480
This page has been reformatted by Knovel to provide easier navigation.
COLORPLATES
Time (min)
A
C
WT
F B
D R
6180" 690"
P
Color Plate 1 (chapter 6 ) The FrzS-Gfp protein may oscillate between poles along a helical filament. FrzS may track along a helical filament. (A) Time-lapse fluorescence microscopy of a dynamic FrzS cluster. The cell was filmed for 5 min. White arrows point to the observed dynamic FrzS spot. (B) Trajectory of the moving complex. (Left) Enhanced view of the cell shown in panel A after 2.5 min (top) and schematics of the fluorescence signal (bottom). (Top right) The images shown in panel A were overlaid, and the spots observed a t different times were linked to obtain a trajectory. The numbers refer to the times at which the foci were seen at a particular subcellular location. (Bottom right) Schematics of the trajectory (blue line) overlaid on the proposed coiled track (red line). (C) FrzS,,,~,,, is defective for vegetative swarming. Motility phenotypes of the wild-type (WT) and ~YzS,~~,.,~, strains on S-motilityspecific Casitone-yeast-extract-rich medium 0.3 % agar plates. (D) Subcellular localization of FrzS,,,.,,,. The localization pattern of FrzS,,,+,, was determined by immunostaining using the FrzS-specific antiserum. R, raw image; P, processed image. (Right) Clockwise 90" rotations of the reconstructed volume of the segment boxed in the processed image. Scale bar, 2 pni. Reprinted with permission from Science (Mignot et al., 2005).
COLORPLATES
LEADING POLE
Color Plate 2 (chapter 7) Model for the regulation of the Frz chemosensory pathway and its control of A- and S-motility. The input module (shaded in blue), a two-component signal transduction pathway, signals the output module (shaded in pink), a complex related in part to small GTPases, to trigger a reversal in both the S- and A-motility systems. The input module comprises the cytoplasmic MCP, FrzCD; the methyltransferase-TPR hybrid protein, FrzF; the Chew coupling protein, FrzA; and the CheA-CheY hybrid protein, FrzE. One or more unknown signals interact with FrzCD and/or interact with the TPR domain of FrzF, causing site-specificmethylation and activation of FrzCD. FrzG is not represented, as its regulatory role remains unclear. Activated FrzCD induces autophosphorylation of FrzE, which is under negative regulation by the C-terminal CheY domain. A CheY-CheY-like fusion protein, FrzZ, accepts phosphate from FrzE and propagates signaling to the output module. Phospho-FrzZ mediates an unknown interaction with components of the MglA complex, which independently and coordinately signals the S-motility system and the A-motility system to trigger a reversal and define a new leading pole. MglA shares homology with small GTPases and interacts with FrzS and AglZ, which are important components of the S- and A-motility systems, respectively.
Sequence
Similarity
Phylogenornic
Protein 1 Protein 2
+
Protein 1 Protein 2
Protein 3
Protein 3 Multiple genornes
d
OrdinationNisualization
Color Plate 3 (chapter 17) Phylogenomic mapping. ( a ) Sequence data from partial, single, or multiple genomes are used to produce protein predictions, which are aligned against a database of proteins from hundreds of completely sequenced genomes. (b) A phylogenomic raw bit score matrix, with each row corresponding to a specific protein and each column corresponding to a bacterial species, is then created. A heat map visualization of the matrix with array elements colored according to their corresponding shades of red is also shown. (c)From the phylogenomic matrix, a similarity matrix using Spearman's rank correlation is calculated. (d) Finally, a combination of multidimensional scaling and force-directed placement is used to transform the similarity matrix into a two-dimensional ordination. This is then visualized in three dimensions using the computer program VxInsight.
~1
COLORPLATES
Color Plate 4 (chapter 17) A total of 15 ORFs are selected from five different clusters on the map. Colored boxes denote the specific clusters selected for inactivation. The specific proteins which were selected for disruption are colored in each medium-resolution view.
COLORPLATES
Color Plate 5 (chapter 18) A drawing from Thaxter’s sketchbook: Polycephalum aurantiacum, later designated as Stigmatella aurantiaca. Reproduced from the original preserved in the Archives of the Farlow Library of Cryptogamic Botany, Harvard University, Cambridge, MA.
COLORPLATES
Color Plate 6 (chapter 18) Stigmatella cells on their way into an aggregation center (left). Fruiting bodies of S. aurantiaca DW4/3-1. A fruiting body is about 100 km in height (right). Reprinted, with permission, from the cover of Molecular Microbiology, volume 56, issue no. 5.
COLORPLATES
d Color Plate 7 (chapter 19) (a)Filter paper after 2 weeks of incubation. Growth of Sorungium occurs in the orange zone of the white filter paper. In the next inner circle the cellulose fibers are increasiiigly lysed. In the center of the filter, square red-brown fruiting bodies appear. (b) Typical Sorangium swarms with cable-like veins and rings growing outside the filter paper on the agar surface. ( c ) Sporangioles suspended in buffer. (d) Typical fruiting bodies from S. celldosum. Sporangioles are packed together into dense parcels. Light is reflected by a slime cover.
COLORPLATES
a
flaviolin
b
.nr
~
A[
,
59
zoo
,1o;o
,
,
,
250 300 350 40 450 nm
~?-;--, l5,O
P. pufida::rppA time, min
Color Plate 8 (chapter 19) Heterologous production of flaviolin in P. putidu. (a) Flaviolin biosynthetic pathway; (b) phenotypes of wild-type P. putidu and a mutant strain expressing the rppA gene from S. cellulosum; (c) HPLC analysis of culture extracts of P. putidu producing flaviolin.
COLORPLATES
i
i ,
Color Plate 9 (chapter 22) Protein localization in C. crescentus. The subceIlular localization patterns of regulatory proteins during the course of the cell cycle. For proteins that localize to more than one site, the sizes of the dots indicate the predominant localization. Lines across the cell indicate diffuse cytoplasmic protein. Colored lines around the periphery of the cell indicate diffuse membrane-bound protein.
COLORPLATES
Color Plate 10 (chapter 2 3 ) An image of fluorescence of Anabaena sp. expressing the gene for obelin, an indicator of calcium ions. With coelenterazine present, the intensity of blue fluorescence generated by obelin is linearly related to the concentration of calcium ions. Filaments of Anabaena sp. incubated with coelenterazine for 30 inin were excited by near-UV light and observed by fluorescence microscopy. The red fluorescence (>600 nm) is derived from the phycobiliproteiiis and chlorophyll of the vegetative cells. The blue fluorescence of the heterocysts measures their concentration of free calcium. Blue fluorescence of vegetative cells (wavelengths shorter than 600 nm) was much weaker than that of heterocysts. Calciumconcentration-dependent blue fluorescence increases in developing cells before morphological changes are observed (Zhao et al., 2005). Three light flecks were removed from the image by Adobe Photoshop CS.
COLORPLATES
t
Tip extension
4
Q
\
/
Nucleoid migration
Cell division Color Plate 11 (chapter 24) Apical growth in Streptomyces. ( a ) Staining of the sites of nascent peptidoglycan incorporation using fluorescently labeled vancomycin in vegetative hyphae of S. coelicolor. The fluorescence image is shown in inverted gray scale. Hyphal tips are indicated by a “t,” cross walls are indicated by arrowheads, and the spore from which the hyphae grew out is indicated by “Sp.” (b) Subcellular localization of DivIVAsc-EGFP (green color) overlaid on a phase-contrast image of nascent mycelium growing out of a spore (Sp). Bars, 5 kin. (c) Simplified illustration of polarized growth in Streptomyces hyphae. The apical cell is extending its cell wall only at the tip (green). Once this cell has divided by forming a new hyphal cross wall, the subapical daughter cell becomes unable to grow and eventually switches its polarity to generate a lateral branch with a new extending tip. A consequence of tip growth is that DNA, which replicates along most of the hyphal length, has to move towards the tip and into new branches-a process designated nucleoid migration (Flardh, 2003b). For clarity, only a few schematic nucleoids are drawn (red), and they are not meant to reflect the actual number of chromosomes per cell. Furthermore, individual nucleoids are typically not observed in vivo as separated bodies in growing hyphae. Reprinted from Flardh (2003b), with permission from Elsevier.
COLORPLATES
Color Plate 12 (chapter 24) Visualization of septation and nucleoid segregation in wild-type S. coelicolor. (a) Phase-contrast micrograph of aerial filaments. (b) Fluorescence image of the same filaments showing sites of cell wall synthesis stained with fluo-WGA. (c) Fluorescence image of the same filaments showing nucleoids stained with 7-AAD. The figure shows representative aerial hyphae from colonies that had developed for 3 days on mannitol MM plates before being prepared for microscopy. Bar, 10 p,m. Reproduced with permission from Flardh et al. (1999).