Thermophiles: Biology and Technology at High Temperatures

Thermophiles Biology and Technology at High Temperatures

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Thermophiles Biology and Technology at High Temperatures Edited by

Frank Robb Garabed Antranikian Dennis Grogan Arnold Driessen

Boca Raton London New York

CRC Press is an imprint of the Taylor & Francis Group, an informa business

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CRC Press Taylor & Francis Group 6000 Broken Sound Parkway NW, Suite 300 Boca Raton, FL 33487-2742 © 2008 by Taylor & Francis Group, LLC CRC Press is an imprint of Taylor & Francis Group, an Informa business No claim to original U.S. Government works Printed in the United States of America on acid-free paper 10 9 8 7 6 5 4 3 2 1 International Standard Book Number-13: 978-0-8493-9214-6 (Hardcover) This book contains information obtained from authentic and highly regarded sources. Reprinted material is quoted with permission, and sources are indicated. A wide variety of references are listed. Reasonable efforts have been made to publish reliable data and information, but the author and the publisher cannot assume responsibility for the validity of all materials or for the consequences of their use. No part of this book may be reprinted, reproduced, transmitted, or utilized in any form by any electronic, mechanical, or other means, now known or hereafter invented, including photocopying, microfilming, and recording, or in any information storage or retrieval system, without written permission from the publishers. For permission to photocopy or use material electronically from this work, please access www.copyright.com (http:// www.copyright.com/) or contact the Copyright Clearance Center, Inc. (CCC) 222 Rosewood Drive, Danvers, MA 01923, 978-750-8400. CCC is a not-for-profit organization that provides licenses and registration for a variety of users. For organizations that have been granted a photocopy license by the CCC, a separate system of payment has been arranged. Trademark Notice: Product or corporate names may be trademarks or registered trademarks, and are used only for identification and explanation without intent to infringe. Library of Congress Cataloging-in-Publication Data Thermophiles : biology and technology at high temperatures / editors, Frank Robb ... [et al.]. p. ; cm. “A CRC title.” Includes bibliographical references and index. ISBN 978-0-8493-9214-6 (hardcover : alk. paper) 1. Thermophilic microorganisms. I. Robb, F. T. (Frank T.) II. Title. [DNLM: 1. Bacterial Physiology. 2. Adaptation, Physiological. 3. Archaea--genetics. 4. Archaea--physiology. 5. Bacteria--genetics. 6. Genetics, Microbial. 7. Heat. QW 52 T4107 2008] QR84.8T445 2008 579.3’1758--dc22

2007029416

Visit the Taylor & Francis Web site at http://www.taylorandfrancis.com and the CRC Press Web site at http://www.crcpress.com

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Contents Preface ......................................................................................................................................

vii

About the Editors ....................................................................................................................

ix

Contributors ............................................................................................................................

xi

PART I

Overview ..............................................................................................................

1

Chapter 1

Introduction .......................................................................................................... Frank T. Robb, Garabed Antranikian, Dennis W. Grogan, and Arnold J.M. Driessen

3

PART II

Molecular Basis of Thermostability .................................................................

7

Chapter 2

Compatible Solutes of (Hyper)thermophiles and Their Role in Protein Stabilization .......................................................................................................... Helena Santos, Pedro Lamosa, Tiago Q. Faria, Tiago M. Pais, Manuela López de la Paz, and Luis Serrano

Chapter 3

Relationships among Catalytic Activity, Structural Flexibility, and Conformational Stability as Deduced from the Analysis of Mesophilic–Thermophilic Enzyme Pairs and Protein Engineering Studies ....... Reinhard Sterner and Eike Brunner

9

25

Chapter 4

Membranes and Transport Proteins of Thermophilic Microorganisms .............. Sonja Verena Albers and Arnold J.M. Driessen

39

Chapter 5

Thermophilic Protein-Folding Systems ............................................................... Frank T. Robb and Pongpan Laksanalamai

55

Chapter 6

Physical Properties of Membranes Composed of Tetraether Archaeal Lipids .... Parkson Lee-Gau Chong

73

PART III

Heat-Stable Enzymes and Metabolism ............................................................

97

Chapter 7

Glycolysis in Hyperthermophiles ......................................................................... Peter Schönheit

99

Chapter 8

Industrial Relevance of Thermophiles and Their Enzymes ................................. 113 Garabed Antranikian v

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Contents

Chapter 9

Denitrification Pathway Enzymes of Thermophiles ............................................ 161 Simon de Vries and Imke Schröder

PART IV

Genetics of Thermophiles .................................................................................. 177

Chapter 10 DNA Stability and Repair .................................................................................... 179 Malcolm F. White and Dennis W. Grogan Chapter 11 Plasmids and Cloning Vectors for Thermophilic Archaea .................................. 189 Kenneth M. Stedman Chapter 12 Genetic Analysis in Extremely Thermophilic Bacteria: An Overview ............... 205 Dennis W. Grogan Chapter 13 Targeted Gene Disruption as a Tool for Establishing Gene Function in Hyperthermophilic Archaea ............................................................................ 213 Haruyuki Atomi and Tadayuki Imanaka Chapter 14 Nanobiotechnological Potential of Viruses of Hyperthermophilic Archaea ....... 225 Tamara Basta and David Prangishvili PART V

Minimal Complexity Model Systems ................................................................ 237

Chapter 15 Master Keys to DNA Replication, Repair, and Recombination from the Structural Biology of Enzymes from Thermophiles ............................. 239 Li Fan, R. Scott Williams, David S. Shin, Brian Chapados, and John A. Tainer Chapter 16 DNA Replication in Thermophiles ...................................................................... 265 Jae-Ho Shin, Lori M. Kelman, and Zvi Kelman Chapter 17 DNA-Binding Proteins and DNA Topology ........................................................ 279 Kathleen Sandman Chapter 18 Structure and Evolution of the Thermus thermophilus Ribosome ....................... 291 Steven T. Gregory and Albert E. Dahlberg Chapter 19 Protein Phosphorylation at 80°C and Above ........................................................ 309 Peter J. Kennelly Chapter 20 Archaeal 20S Proteasome: A Simple and Thermostable Model System for the Core Particle ..................................................................... 333 Joshua K. Michel and Robert M. Kelly Index .......................................................................................................................................... 347

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Preface We, as scientists, find ourselves in an era of retooling both the technology and the concepts of biology on account of access to global information on an unprecedented scale, through whole genome analysis and community sequencing. This revolution has affected the field of extremophiles, perhaps to a greater extent than most areas of microbiology, due to the challenges posed by carrying out many routine investigations at high temperature. As a result, the database-induced change in modus operandi has pushed back the frontiers of discovery on thermophiles rapidly over the last decade. We, the editors, felt that this book would be a timely contribution to sum up some of the most exciting areas of growth in the field. We have focused on thermophiles, those bacteria and archaea adapted primarily to heat, although many thermophilic species also withstand high osmolarity, high hydrostatic pressure, low or high pH, toxic metals, or organic solvents. Many of the species in common use in this field were isolated and characterized by the pioneers of the field, including Karl-Otto Stetter, Carl Woese and the late Wolfram Zillig. Many of the extraordinary microbial species “in captivity” as a result of their efforts represent a very important window into the microbiology of marine hydrothermal vents and geothermal systems. Hydrothermal vent circulation is becoming more relevant as awareness of global climate change increases. Current estimates suggest that the circulation of seawater through the oceanic crust accounts for 34% of the heat input into the global oceans, about 25% of the globe’s total heat input. Hydrothermal vents may regulate the chemistry of the global oceans and could be responsible for the elemental composition of seawater, and we are aware from pioneering studies that autotrophic thermophiles are abundant and active in volcanic outflows. Much work remains to be done on the microbiology of hydrothermal systems in general and although this book is not focused on environmental microbiology, our knowledge of molecular insights into thermophilic lifestyles, and the development of new genetic tools are critical to the design of future field studies of thermophiles. Our main goal was to capture the excitement that currently prevails in the field of thermophile molecular biology. Examples of some of the topics include the description key adaptive mechanisms of hyperthermophiles and acidophiles, and the genetic methods that will be in the toolkit of all thermophile laboratories in future. Many practical applications of thermophiles and their enzymes have also recently matured and there has been a true “coming of age” of technology derived from thermophiles and other “extremophiles.” Our purpose was to capture some of the most exciting and innovative advances in the area of applications. We also sought to highlight areas where thermophiles provide model systems for complex cellular functions. These include DNA replication and repair, protein phosphorylation, osmoregulation and the enhancement of thermal stability by unusual compatible solutes, and protein folding systems. In many cases, thermophiles have provided critically important advantages to study complex multicomponent systems due to their inherent stability at high temperatures, leading to extreme durability and minimal thermal transitions at “normal” temperatures. An excellent example of the “thermophile advantage” is the resolution of the structure of the first 70S ribosome structure using Thermus thermophilus, a hot spring bacterium, as the source organism (see Chapter 18). In addition, the extremely thermophilic Archaea are “miniature” Eukarya in the sense that the foundational mechanisms of many complex functions such as DNA replication and transcription are identical, accomplished however with far fewer components in the Archaea. This leads to profound advantages in assigning functions to each component. vii

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We are greatly indebted to many people who saw the same vision and have provided essential help. The first to be thanked are without doubt the contributors who devoted valuable energy and time to their chapters. They are the authorities in the areas described above and without their unstinting help, the depth of review of our topics would have been much shallower. We are also most grateful for the patient and constructive help we received from Judith Spiegel and Amber Donley at Taylor & Francis, and the assistance of a number of reviewers in editing the initial drafts of the chapters. Frank T. Robb, Garabed Antranikian, Dennis W. Grogan, and Arnold J.M. Driessen

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About the Editors Frank T. Robb is professor at the Center of Marine Biotechnology, University of Maryland Biotechnology Institute. A South African, he received a BSc (Hons) from the University of Cape Town (UCT) in 1968 and a PhD from the University of California, Riverside in 1972. He completed postdoctoral studies at the University of California, San Diego and the University of Chicago. Before coming to the University in Maryland in 1988, he was associate professor in the microbiology department at UCT. Research projects in his laboratory include protein folding mechanisms in hyperthermophiles, genomic studies on extremophilic Bacteria and Archaea, and field studies in volcanic environments, including Iceland, New Zealand, Yellowstone National Park and the Kamchatka Peninsula, Eastern Siberia. He is a member of the Faculty of 1000. Garabed Antranikian studied biology as an undergraduate student at the American University in Beirut. At the University of Göttingen, he completed his PhD thesis in microbiology in 1980 in the laboratory of Professor Gerhard Gottschalk and qualified as a postdoctoral lecturer (Habilitation) in 1988. In 1989, he was appointed to a professorship in microbiology at the Hamburg University of Technology, where he has been the head of the Institute of Technical Microbiology since 1990. From 2000 to 2003, Professor Antranikian coordinated the Network Project Biocatalysis and has been coordinating the Innovation Center Biokatalyse (ICBio) since 2002. He is president of the International Society for Extremophiles and is a coeditor of several scientific journals. In 2004, he was awarded the prize for environment protection by the Federal Environmental Foundation of Germany (DBU). Since 2007, he has served as the coordinator of the “Biocatalysis 2021” Cluster of the Ministry of Education and Research. Dennis W. Grogan became interested in prokaryotic microorganisms during his undergraduate study at the University of Missouri, and participated in research on Synechococcus with Louis Sherman. He received his MS and PhD from the University of Illinois (Urbana-Champaign) in the laboratory of John E. Cronan, Jr., studying the genetics and physiology of cyclopropane fatty acids in Escherichia coli. During this period, he was exposed to the research of R.S. Wolfe, C.R. Woese, and others on Archaea and other microorganisms with diverse metabolic properties. He then spent several years of postdoctoral training in the laboratories of W. Zillig (Max-Planck-Institut für Biochemie, Martinsried-bei-München), Giuseppe Bertani (NASA Jet Propulsion Laboratory), and Robert P. Gunsalus (University of California, Los Angeles), focusing on Sulfolobus spp. and other Archaea from geothermal environments. In 1994, he joined the department of biological sciences at the University of Cincinnati, where he is currently full professor. Arnold J.M. Driessen was born in 1958 in Horst, the Netherlands. From 1997 to 1983, he studied biology at the University of Groningen, and in 1987 obtained his PhD cum laude on the thesis “Amino acid transport in lactic streptococci” under the supervision of Professor Dr. W.N. Konings. He then became scientific officer in the department of microbiology at the University of Groningen. In 1988, he was honored with the Kluyver Award of the Dutch Society of Microbiology. In 1989 to 1990, he went as postdoctoral student to the University of California at Los Angeles where he studied with Dr. W. Wickner working on the mechanism of bacterial protein translocation. After returning to the University of Groningen, he became associate professor in 1992 in the department of microbiology, ix

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About the Editors

and received the PIONIER award from the Netherlands Organization for Scientific Research (NWO), and in 1993, he was awarded the Federation of European Biochemical Societies (FEBS) Anniversary Prize of the Society for Biological Chemistry. In 1997, he became full professor in microbiology, and from 2000 to 2002, held a NWO-ALW Van der Leeuw Chair in microbiology in the same department. He now heads a group that works on the enzymatic and energetic mechanism of protein translocation in bacteria and archaea, and structural and functional studies on solute transport in microorganisms.

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Contributors

Sonja Verena Albers Department of Microbiology University of Groningen Groningen, the Netherlands Garabed Antranikian Institute of Technical Microbiology Hamburg University of Technology Hamburg, Germany Haruyuki Atomi Department of Synthetic Chemistry and Biological Chemistry Graduate School of Engineering Kyoto University Kyoto, Japan Tamara Basta Molecular Biology of the Gene in Extremophiles Unit Institute Pasteur Paris, France Eike Brunner Institute of Biophysics and Physical Biochemistry University of Regensburg Regensburg, Germany

Albert E. Dahlberg Department of Molecular Biology, Cell Biology, and Biochemistry Brown University Providence, Rhode Island Simon de Vries Department of Biotechnology Delft University of Technology Delft, the Netherlands Arnold J.M. Driessen Department of Microbiology University of Groningen Groningen, the Netherlands Li Fan Department of Molecular Biology The Scripps Research Institute La Jolla, California Tiago Q. Faria Institute of Chemical and Biological Technology New University of Lisbon Lisbon, Portugal

Brian Chapados Department of Molecular Biology The Scripps Research Institute La Jolla, California

Steven T. Gregory Department of Molecular Biology, Cell Biology, and Biochemistry Brown University Providence, Rhode Island

Parkson Lee-Gau Chong Department of Biochemistry Temple University School of Medicine Philadelphia, Pennsylvania

Dennis W. Grogan Department of Biological Sciences University of Cincinnati Cincinnati, Ohio

xi

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Tadayuki Imanaka Department of Synthetic Chemistry and Biological Chemistry Graduate School of Engineering Kyoto University Kyoto, Japan Robert M. Kelly Department of Chemical and Biomolecular Engineering North Carolina State University Raleigh, North Carolina

Contributors

Tiago M. Pais Institute of Chemical and Biological Technology New University of Lisbon Lisbon, Portugal David Prangishvili Molecular Biology of the Gene in Extremophiles Unit Institute Pasteur Paris, France

Lori M. Kelman Program in Biotechnology Montgomery College Germantown, Maryland

Frank T. Robb Center of Marine Biotechnology University of Maryland Biotechnology Institute Baltimore, Maryland

Zvi Kelman Center for Advanced Research in Biotechnology University of Maryland Biotechnology Institute Rockville, Maryland

Kathleen Sandman Department of Microbiology Ohio State University Columbus, Ohio

Peter J. Kennelly Department of Biochemistry Virginia Polytechnic Institute and State University Blacksburg, Virginia Pongpan Laksanalamai Center of Marine Biotechnology University of Maryland Biotechnology Institute Baltimore, Maryland Pedro Lamosa Institute of Chemical and Biological Technology New University of Lisbon Lisbon, Portugal Manuela López de la Paz European Molecular Biology Laboratory Heidelberg, Germany Joshua K. Michel Department of Chemical and Biomolecular Engineering North Carolina State University Raleigh, North Carolina

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Helena Santos Institute of Chemical and Biological Technology New University of Lisbon Lisbon, Portugal Peter Schönheit Institute for General Microbiology Christian Albrechts University in Kiel Kiel, Germany Imke Schröder Department of Microbiology, Immunology, and Molecular Genetics University of California–Los Angeles Los Angeles, California Luis Serrano European Molecular Biology Laboratory Heidelberg, Germany David S. Shin Department of Molecular Biology The Scripps Research Institute La Jolla, California

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Contributors

Jae-Ho Shin Center for Advanced Research in Biotechnology University of Maryland Biotechnology Institute Rockville, Maryland

John A. Tainer Department of Molecular Biology The Scripps Research Institute La Jolla, California

Kenneth M. Stedman Department of Biology Portland State University Portland, Oregon

Malcolm F. White Centre for Biomedical Sciences St. Andrews University St. Andrews, Scotland, United Kingdom

Reinhard Sterner Institute of Biophysics and Physical Biochemistry University of Regensburg Regensburg, Germany

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R. Scott Williams Department of Molecular Biology The Scripps Research Institute La Jolla, California

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Part I Overview

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1

Introduction Frank T. Robb, Garabed Antranikian, Dennis W. Grogan, and Arnold J.M. Driessen

CONTENTS Economic and Commercial Developments .................................................................................... Genetic Analysis ............................................................................................................................ Analysis of Complex Systems ........................................................................................................ References ......................................................................................................................................

4 4 5 5

Microorganisms have been found living in almost every place explored so far, from hydrothermal vents in the deepest trenches of the Pacific Ocean to frozen Antarctic ice, and deep in the earth’s lithosphere. Under conditions that represent the extreme ranges of physical and chemical conditions that permit cellular survival, microorganisms termed extremophiles represent the most radical adaptations that allow survival and growth. The concept of normal or mild conditions is of course relative. However, we can generalize that life, at least as we know it on earth, depends on the availability of liquid water as the most important solvent (Rothschild and Mancinelli, 2001). Thermophiles (literally heat lovers) are microorganisms that thrive at temperatures above the mesophilic range of 25°C to 40°C that characterizes the mainstream of life. While thermophiles are an eclectic bunch, these organisms share a common theme: they exist at the fringes, where high temperature excludes all but the hardiest of inhabitants. Every component of these small, prokaryotic cells (typically about 1 μm in diameter) is exposed continually to the high temperatures of their environments and must be adapted to function under these conditions. Thus, all molecules, ranging from cell surface complexes, cytoplasmic membrane (Itoh et al., 2001), and ribosomes, down to metabolic enzymes and intermediary metabolites (Robb and Maeder, 1998), must cope with the threat of unfolding or decomposition (Russell, 2003). Although certain of these molecular adaptations have been identified, many remain unknown. Furthermore, thermophiles are often adapted to additional extremes that combine with high temperature to threaten the structural integrity of their cells. For example, different groups of microbes are acidophilic and alkaliphilic, inhabiting extremely acidic or basic water or soil. These double or triple extremophiles may be able to extend the limit of one extreme because of the effects of another. For example, thermophilic piezophiles in the deep oceans or deep lithosphere survive at pressures hundreds of times greater than that of the earth’s atmosphere, and this may be because the thermal stability of many proteins is extended at high pressure (Madigan et al., 2003). Recent findings confirm that these organisms represent more than curiosities, microbial oddities in obscure and little explored ecological niches. They are a rich source of unexpected and unique adaptive mechanisms that fuel further understanding of the fundamental mechanisms of life (Huber et al., 2000). To date, over 450 thermophilic isolates are known. Representative species are shown in Figure 9.1 on the universal phylogenetic tree of small subunit ribosomal RNA sequences. They belong almost 3

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Thermophiles: Biology and Technology at High Temperatures

exclusively to the prokaryote domains, Bacteria and Archaea. Very few thermophilic eukaryotes exist (Kashefi and Lovley, 2003). An active controversy surrounds the upper temperature limit of terrestrial life. The most extreme named microbial species is Pyrolobus fumarii, which can multiply at temperatures up to 113°C and survive over 1 h at 121°C, under autoclave conditions (Blöchl et al., 1997). This strain grows on the walls of black smokers, undersea thermal vents which eject water at very high temperatures and pressures. Like many hyperthermophiles, P. fumarii is classified as a chemoautotroph; it synthesizes essential metabolites and carries out energy conservation using disequilibria between inorganic compounds in its environment. Recently debuted (Kashefi and Lovley, 2003), but not fully described yet, strain 121 is capable of prolonged survival, and possibly growth at 121°C. As we can see in the phylogenetic tree in Figure 9.1, p163, hyperthermophiles occupy the lower branches of the tree, while moderate and extreme thermophiles are widely distributed among the bacterial and archaeal taxa.

ECONOMIC AND COMMERCIAL DEVELOPMENTS We are entering a new era when microorganisms that were previously considered only as microbiological curiosities are being recognized as the basis for new and radically innovative biotechnology (Bull et al., 2000; Eichler, 2001). Starting with the invention of the polymerase chain reaction (PCR) amplification method and its reliance on thermostable DNA polymerases, thermophiles have contributed to many areas of economic development (food processing, biofuels, and so on) (Vieille and Zeikus, 2001).

GENETIC ANALYSIS Many of the microorganisms described in this volume require high temperatures for normal metabolism and reproduction. This fact focuses attention on their cellular components (RNA, DNA, enzymes, membranes, and so on) and the molecular modifications that enable these components to function adequately at high temperature. Analysis of thermophilic bacteria and archaea at this level benefits from microarray hybridization, high-throughput mass spectrometry, and other modern, sensitive techniques based on complete genome sequences. However, confirming the roles of specific proteins in specific processes and altering cellular properties for technological or experimental purposes require genetic manipulation of the microorganism, which remains an important challenge for thermophile research. The question of DNA repair at extremely high temperatures illustrates both the promise and challenge of this genetic analysis (Grogan, 1998). At the optimal growth temperatures of thermophiles, spontaneous decomposition of pure DNA in buffered solution is greatly accelerated, raising questions as to whether repair mechanisms successfully compensate for the increased load of damage predicted in vivo. As described in Chapter 10, this is a biochemically complex issue that reveals areas of commonality between bacteria and archaea, as well as areas of obvious difference, and promises to open a fascinating new perspective on the molecular diversity of DNA repair. Given the close association between DNA repair and genetic exchange processes, however, resolving the DNA repair processes in extreme thermophiles will require extensive genetic analysis and genetic manipulation, which remains technically challenging. The extent of these challenges is illustrated by the fact that many extreme thermophiles require special cultivation techniques and do not form isolated colonies readily on solid media, making even basic manipulation difficult. Thus, efforts toward genetic analysis have focused on bacteria and archaea which grow vigorously under aerobic, heterotrophic conditions and respond to routine microbiological manipulations. This has enabled researchers to adapt existing techniques of microbial genetics to these species, and to begin development of new techniques, as well. Thermus spp., for example, demonstrate how classical bacterial genetics can be extended to temperatures

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Introduction

5

above 70°C. These bacteria are sensitive to a number of antibiotics, and corresponding resistance genes have been adapted to high temperature and incorporated into plasmids that replicate in these hosts. Homologous recombination is efficient in Thermus spp., which enables selectable markers and engineered mutations to be transferred to the chromosome by simple transformation and selection, and at least one species is naturally competent. For thermophilic archaea, which grow at still higher temperatures, genetic methods derive less from the bacterial repertoire and more from strategies familiar to yeast geneticists. Two practical reasons for this trend are the intrinsic insensitivity of most archaea to most antibiotics (which renders the vast majority of bacterial selectable markers irrelevant) and the unanticipated difficulty of developing reliable vectors from natural archaeal plasmids. The most successful selectable markers in thermophilic archaea have been cloned genes that restore a selectable metabolic function in a corresponding mutant. The most popular strategy takes advantage of the sensitivity of many thermophilic archaea to the metabolite analog 5-fluoro-orotic acid (FOA), which selects for the loss of either of the two uridine monophosphate (UMP) biosynthetic enzyme activities. As in yeast, therefore, medium containing uracil plus FOA selects uracil auxotrophs of thermophilic archaea; these auxotrophs, in turn, provide a selection for a functional copy of the corresponding biosynthetic gene. Another selection, so far confined to Sulfolobus solfataricus, uses a chromosomal beta-glycosidase gene to restore growth of the corresponding mutant on lactose as sole carbon and energy source. The most useful vectors for thermophilic archaea to date have been modified versions of natural viruses, and most genetic engineering of thermophilic archaea has used homologous recombination to alter host genomes at defined loci (Chapter 11). In addition to their potential as genetic tools, viruses of thermophilic archaea provide a fascinating view of the molecular diversity of life on earth. In recent years, a remarkable series of morphologically and genetically novel viruses have been recovered from the archaeal communities in acidic hot springs around the world. Basta and Prangishvili summarize their recent progress in analyzing these novel viruses and developing the biotechnological potential they present.

ANALYSIS OF COMPLEX SYSTEMS An emerging theme in thermophile research is the recognition that complex cellular processes such as DNA replication must take place with minimal sets of components. This is leading to breakthroughs in understanding mechanisms of action in multicomponent cellular processes that thermophiles share with mesophiles. An additional advantage, pointed out in Chapter 15, is that complex cellular processes adapted to high temperatures can often be “frozen” in intermediate junctures that have very short lifetimes in mesophilic counterparts. This leads to advantages in obtaining insights into mechanisms from structures of intermediate ternary complexes. An example of this is the structure of the 70-S ribosome, which was first resolved in Thermus spp. and is described in Chapter 18 by Gregory and Dahlberg. The analysis of replication and repair systems described in Chapter 15 is another example of the inherent simplicity of thermophilic systems in which the proteins are reduced in size and more rigidly folded than they are in mesophiles.

REFERENCES Blöchl, E., R. Rachel, S. Burggraf, D. Hafenbradl, H. W. Jannasch, K. O. Stetter, 1997. Pyrolobus fumarii, gen. and sp. nov., represents a novel group of archaea, extending the upper temperature limit for life of 113 degrees C. Extremophiles, 1(1):14–21. Bull et al., 2000. Search and discovery strategies for biotechnology: the paradigm shift. Microbiology and Molecular Biology Reviews 64 no. 3:575–606. Eichler, J., 2001. Biotechnological uses of archaeal extremozymes. Biotechnol. Adv., 19:261–278. Grogan, D., 1998. Hyperthermophiles and the problem of DNA Instability. Mol. Microbiol., 28(6):1043–1049.

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Huber, R., H. Huber, and K. Stetter, 2000. Towards the ecology of hyperthermophiles: Biotopes, new isolation strategies and novel metabolic properties. FEMS Microbiol. Rev., 24:615–623. Itoh, Y., A. Sugai, I. Uda, and T. Itoh, 2001. The evolution of lipids. Adv. Space Res., 28(4):719–724. Kashefi, K. and D. R. Lovley, 2003. Extending the upper temperature limit for life science, Science, 301(5635):934. Madigan, M. T., J. M. Martinko, and J. Parker, 2003. Brock Biology of Microorganisms, Tenth Edition, Prentice Hall, Pearson Education, Inc., 1019 pp. Robb, F. and D. Maeder, 1998. Novel evolutionary histories and adaptive features of proteins from hyperthermophiles. Curr. Opin. Biotechnol., 9:288–291. Rothschild, L. and R. Mancinelli, 2001. Life in extreme environments. Nature, 409:1092–1101. Russell, A., 2003. Lethal effects of heat on bacterial physiology and structure. Sci. Prog., 86:115–137. Vieille, C. and G. Zeikus, 2001. Hyperthermophilic enzymes: Sources, uses, and molecular mechanisms for thermostability. Microbiol. Mol. Biol. Rev., 65(1): 1–43.

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Part II Molecular Basis of Thermostability

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Compatible Solutes of (Hyper)thermophiles and Their Role in Protein Stabilization Helena Santos, Pedro Lamosa, Tiago Q. Faria, Tiago M. Pais, Manuela López de la Paz, and Luis Serrano

CONTENTS Introduction .................................................................................................................................. Compatible Solutes Typical of (Hyper)thermophiles .................................................................. Protein Stabilization by Thermosolutes ....................................................................................... Kinetic Stabilization of Proteins by Thermosolutes ......................................................... Thermodynamic Stabilization of Proteins by Thermosolutes .......................................... Protein Conformational Stabilization by Thermosolutes ................................................. Understanding the Molecular Basis of Protein Stabilization ........................................... Effect of Thermosolutes on the Protein Unfolding Pathway ............................................ Impact of Thermosolutes on Protein Dynamics ............................................................... Effect of Thermosolutes on the Pathway of Fibril Formation .......................................... Concluding Remarks .................................................................................................................... Acknowledgments ........................................................................................................................ References ....................................................................................................................................

9 10 12 12 12 13 14 17 17 19 20 21 21

INTRODUCTION The discovery of hyperthermophilic microorganisms in the 1980s by Wolfram Zillig and Karl Stetter had a dramatic impact on the layman’s view that life was exclusive to mild environments. Their pioneering sampling expeditions showed that new microorganisms thrived wherever they searched, despite the extreme harshness of the habitats sought [1]. The remarkable ability of these organisms to overcome the challenges posed by extreme physical parameters can be an invaluable source of knowledge that allows us to understand how these forms of life have improved upon general protective strategies, taking them to their limits or devising new approaches [2]. Every organism, regardless of its optimal growth temperature, has to deal with the issue of thermal stability (both physical and functional) of its cell components. This statement is probably more patent in the case of proteins, entities that have to maintain a certain degree of flexibility to remain functional; therefore, the balance between stability and flexibility is a central feature in protein architecture [3–6]. This compromise has to be adapted to the environmental conditions in which each organism lives, namely temperature. In the case of thermophiles and hyperthermophiles 9

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[from this point on denominated as (hyper)thermophiles], proteins tend to display a higher intrinsic stability and are therefore assumed to have a lower flexibility, in other words, (hyper)thermophilic proteins would be more rigid to achieve a higher stability [7]. Much effort has been directed in the past two decades in understanding how (hyper)thermophiles are able to maintain structurally functional proteins at high temperature, and several structural features have been correlated with intrinsic protein stability (see Chapter 3 in this book). However, intriguing examples of proteins from (hyper)thermophilic origin displaying relatively low stability soon uncovered the need for some degree of extrinsic stabilization [8–12]. This added stability might be provided, in some cases, by the molecular crowding of the intracellular compartments. In fact, salts, high protein concentrations, coenzymes, substrates, and organic solutes greatly increase protein stability in vitro [13–17]. Protein stability is, therefore, the consequence of protein design (or intrinsic stability) and intracellular composition (or extrinsic factors). An organism adapted to grow optimally at a certain temperature will, in principle, have its components functionally adapted to that temperature in terms of the activity/flexibility/stability balance. Redesigning the structural architecture of cellular components to respond to fluctuations of the environmental temperature would be unfeasible. Instead, a change in the properties of the solvent (i.e., the composition of the cytoplasm) may provide, to a certain extent, the required extra stability to cope with an external temperature shift. It is in this context that organic solutes (or chemical chaperones) may come into play, as extrinsic stabilizers in a mechanism of adaptation to thermal stress. (Hyper)thermophiles that are also halophilic or halotolerant accumulate organic solutes not only in response to an increase in the external salinity, but also in response to supraoptimal growth temperatures [18,19]. Solute accumulation as an osmoregulatory strategy to cope with variations in the external water activity is a common trait among moderate halophiles and halotolerant microorganisms [20–22]. These solutes are highly soluble and can accumulate to high levels without disturbing cellular metabolism, hence the term “compatible solutes” [20]. Although the concept of compatible solute was initially restricted to osmoadaptation, it has recently been extended to account for a variety of stress conditions, which cause their intracellular levels to increase, namely, temperature [18,21]. At a first glance, compatible solute accumulation in response to heat stress seems rather puzzling, especially if we consider that the external water activity remains practically unaltered when the temperature is raised. On the other hand, if we consider that compatible solutes display a protective effect upon cellular structures (namely proteins) [8,14,15,17,23–25], then the suggestion of a link between compatible solute accumulation by (hyper)thermophiles and structural protection against heat damage is inevitable. If organic solute accumulation is indeed part of the heat stress adaptation in halophilic (hyper)thermophiles, then the type of solutes used, their accumulation patterns, and their ability to stabilize proteins should reflect that strategy. The main objectives of this chapter are: to review our knowledge on the nature of solutes typically associated with (hyper)thermophiles; to present an overview of the comparative performance of these compounds as protectors of protein structure against thermal denaturation; and to contribute to the elucidation of the complex molecular mechanisms underlying protein stabilization.

COMPATIBLE SOLUTES TYPICAL OF (HYPER)THERMOPHILES Only a decade ago little was known about compatible solute accumulation in hyperthermophilic organisms. Today the picture has changed considerably with many species examined and the identification of several newly discovered solutes [26]. Among the solutes occurring in (hyper)thermophiles, some, like trehalose, α-glutamate, or proline, are regularly found in nonthermophilic organisms; others, like di-myo-inositol phosphate (DIP) are restricted to (hyper)thermophiles; others still, like mannosylglycerate, are strongly associated with thermophily appearing only rarely in mesophiles. Compatible solutes restricted to or mainly found in (hyper)thermophiles will herein be named “thermosolutes” for the convenience of a short designation (Figure 2.1). In contrast to the solutes

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Role in Protein Stabilization -OOC OH

O

OH O

O

HO

-O

O P

P

O

HO

P

O-

O

HO

OH

HO O

OH

OH OH

OH O

O

OH

O-

O

P HO

OH

Di-myo -inositol phosphate

Cyclic-2,3-bisphosphoglycerate

O

OH

HO O

OH

O-

O

O

OP

OH

OH

Glycero-phospho-myo-inositol

OH

OH

Diglycerolphosphate

HO

CH2HO O OH

HO

HO

Mannosylglyceramide

CH2HO O OH HO CH2HO

CH2HO O

Mannosylglycerate C

NH2

O COO-

O

FIGURE 2.1

Compatible solutes primarily restricted to (hyper)thermophiles.

more usually found in mesophiles (bearing no net charge at physiological pH), thermosolutes are generally negatively charged, and most fall into two categories: hexose derivatives with the hydroxyl group at carbon 1 usually blocked in an α configuration, and polyol-phosphodiesters. The most representative compound in the first category is 2-α-O-mannosylglycerate (MG). MG was initially discovered in red algae of the order Ceramiales [27], but is currently acknowledged as one of the most widespread solutes among (hyper)thermophiles, occurring in members of bacteria and archaea belonging to distant lineages [28–32]. Variations on the MG theme include mannosylglyceramide (MGA), mannosylglucosylglycerate, and glucosylglucosylglycerate, compounds that have been found in Rhodothermus marinus, Petrotoga miotherma, and Persephonella marina, respectively. Another structurally related compound is glucosylglycerate (GG), which is relatively common in halotolerant mesophilic bacteria but also occurs in the thermophilic bacterium P. marina [33]. The most prominent member of the polyol-phosphodiester group is DIP. This solute is accumulated by hyperthermophilic bacteria (Thermotoga and Aquifex spp.) as well as archaea (members of the genera Pyrodictium, Pyrococcus, Thermococcus, Methanotorris, Aeropyrum, and Archaeoglobus), in response to supraoptimal growth temperatures [18]. Examples of other polyol-phosphodiesters include diglycerol phosphate (DGP), which has only been found in members of the genus Archaeoglobus, and glycerophospho-inositol, a structural chimera of DIP and DGP that was found in two distant hyperthermophilic genera [34]. Another thermosolute that does not fall into these two categories is cyclic 2,3-bisphosphoglycerate. Although present in many methanogens, this solute only accumulates to high levels in hyperthermophilic species, where its level responds to temperature increase [8,24]. Although (hyper)thermophiles use a variety of organic solutes during thermo- or osmoadaptation, some solutes are preferentially accumulated in response to heat stress whereas others are used mainly to counterbalance external osmolarity. MG, DGP, and amino acids tend to accumulate preferentially in response to an increase in salinity, while the level of DIP and DIP derivatives

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generally increases in response to heat stress [23,28,30–32,35]. Therefore, it is tempting to speculate that DIP (and derivatives) could be especially suited to confer extra protection to cell components against the disruptive effects of elevated temperatures.

PROTEIN STABILIZATION BY THERMOSOLUTES Hyperthermophiles are generally recognized by the high thermostability of most of their proteins. The concept of thermostability includes both thermodynamic and kinetic aspects. The kinetic (or long-term) stability is related to the processes that lead to irreversible loss of the protein activity; it is usually evaluated from the time course of activity loss and is characterized by a rate constant. The thermodynamic stability concerns the reversible unfolding reaction and is defined as the free energy of unfolding. In the case of thermal unfolding, the temperature value at which the populations of native and denatured conformations are equal is known as the melting temperature, Tm, often used to characterize the structural thermostability of macromolecules.

KINETIC STABILIZATION OF PROTEINS BY THERMOSOLUTES Although most studies on the effect of charged solutes on the enzymatic kinetic stability have been performed with model enzymes from mesophilic sources, their superior protective effect in comparison with neutral solutes was also demonstrated for enzymes isolated from (hyper)thermophiles. As an example, incubation of Methanopyrus kandleri formyltransferase for 60 min at 90ºC in the presence of 0.7 M cBPG, a solute accumulated up to molar concentrations in this organism, led to no activity loss, while 92% of the activity was lost in control experiments without solutes [24]. Mannosylglycerate was an effective protector against the thermal inactivation of alcohol dehydrogenase from Pyrococcus furiosus (PfADH) or glutamate dehydrogenase from Thermotoga maritima (TmGDH). The half-life of PfADH at 100ºC increased 10-fold and that of TmGDH at 85ºC increased fourfold in the presence of 0.5 M MG [14]. The stabilization rendered by DGP on Thermococcus litoralis glutamate dehydrogenase was also considerably higher than that exerted by glycerol, the neutral moiety in DGP [25]. The protecting ability of thermosolutes against the thermal inactivation of mesophilic enzymes was also compared with that of common mesophilic solutes like, trehalose or ectoine. As most solutes that accumulate in (hyper)thermophiles are negatively charged, it was relevant to examine their protecting efficiency on enzymes presenting either positive or negative net charge at the working pH. Rabbit muscle lactate dehydrogenase (LDH) is positively charged while pig heart malate dehydrogenase (MDH) has a negative net charge. Interestingly, MG was the best solute to protect both LDH and MDH against heat inactivation [15,36]. Unexpectedly, DIP had a harmful effect on both enzymes inducing a decrease of more than 50% in the half-life for thermal inactivation. In an attempt to obtain insight into the solute’s chemical features responsible for protein stabilization, the effect of compounds chemically related to the solutes found in (hyper)thermophiles was also investigated. In the presence of MGA, the thermal inactivation profile of LDH was identical to that observed in the absence of solutes, indicating that the negative charge in MG is very important to the protective action [15]. In accordance with this view, glycerate was a better stabilizer than glycerol. In another study, the effect of DGP on the kinetic stability of rubredoxins was evaluated [25]. The stabilization rendered by DGP was remarkable when compared with that of glycerol. Phosphate alone was also a good stabilizer but the stabilization conferred by DGP was even higher. The same conclusion was taken from the results obtained with rabbit LDH, baker’s yeast alcohol dehydrogenase and T. litoralis glutamate dehydrogenase [25]. The available data support the view that the negative charge found in most solutes from (hyper)thermophiles plays a fundamental role in their mechanism of stabilization.

THERMODYNAMIC STABILIZATION OF PROTEINS BY THERMOSOLUTES An important aspect in evaluating the stabilizing efficiency of compatible solutes is their effect on the protein unfolding thermodynamics. Once again, it was evident that the negative charge of MG

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Melting temperature of RNase A (ºC)

Role in Protein Stabilization 70 65 60 55 50 45 40 35 2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 pH

FIGURE 2.2 Melting temperature of RNase A as a function of pH in the absence of solutes (triangles and solid line), with 0.5 M mannosylglycerate (squares and dotted line) and with 0.5 M potassium chloride (circles and dashed line).

was a key parameter. The variation of the melting temperature, Tm, of bovine ribonuclease A (RNase A) in the presence and absence of MG depends clearly on the ionization state of the carboxylic group; at pH values higher than 5, when the solute is fully ionized, a significant increase on the RNase A melting temperature was observed (Figure 2.2). On the other hand, as the solute’s charge decreased, the protein melting temperature becomes identical to that observed in the absence of solutes [37]. The superiority of negatively charged solutes in protein stabilization was also evident from the comparison of the effect of a series of charged and uncharged compounds on the Tm of proteins with positive (staphylococcal nuclease, SNase) or negative net charge (pig heart malate dehydrogenase, MDH). The effect of MG, DGP, and DIP was compared with that induced by common uncharged solutes like trehalose, glycerol, ectoine, or hydroxyectoine. It was observed that the thermosolutes induced an increase on the Tms of both proteins, which was always higher than 7°C, while the increase in the presence of trehalose, the best performing neutral solute, was around 4°C for both proteins (Figure 2.3) [36]. The effect of potassium chloride (KCl) was also evaluated as a control for the ionic strength. The Tm of SNase was not affected by the presence of this salt, which caused an increase of only 3ºC on the Tm of MDH. Although the stabilization rendered by ionic solutes is always greater than that induced by neutral solutes, it depends on the particular protein/solute pair considered: DIP and MG were better stabilizers of MDH, but DGP induced a greater stabilization on SNase. It is worth noting that even when glycerol was used at concentration 10 times higher than that of MG, the stabilization rendered by MG was substantially higher [36].

PROTEIN CONFORMATIONAL STABILIZATION BY THERMOSOLUTES Early results showing the ability of compatible solutes to inhibit protein aggregation attracted a renewed interest to the growing knowledge on several debilitating diseases such as Alzheimer’s and Parkinson’s or familial amyloid polyneuropathies. The observation that amyloid fibrils are commonly found in tissues of patients afflicted by these pathologies [38,39] opened the possibility for a chemical chaperone-based therapy [40]. Knowledge on the mechanisms of fibril inhibition may pave the way to new approaches leading to disruption of these aggregates in vivo, or in preventing their formation.

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Thermophiles: Biology and Technology at High Temperatures 12 DIP

10 ΔTm of MDH (ºC)

MG GG

8 DGP

6 4 2

KCl Gly

MGA

Tre

Hect Ect

0 0

2

4

6

8

10

12

ΔTm of SNase (ºC)

FIGURE 2.3 Increment on the melting temperature (ΔTm) of staphylococcal nuclease (SNase) and pig heart malate dehydrogenase (MDH) in the presence of different solutes (0.5 M concentration). Abbreviations: DGP, diglycerol phosphate; DIP, di-myo-inositol phosphate; Ect, ectoine; GG, glucosylglycerate; Gly, glycerol; Hect, hydroxyectoine; KCl, potassium chloride; MG, α-mannosylglycerate; MGA, α-mannosylglyceramide; Tre, trehalose.

Studies to evaluate the effect of thermosolutes on amyloid fibril formation were carried out using the model hexapeptide, STVIIE. This peptide is a computer-aided designed model which rapidly self-associates to form amyloid fibrils: complete fibril maturation is achieved after one week incubation at pH 2.6 and room temperature [41]. The amyloid content was visualized by electron microscopy (EM) (Figure 2.4). The effect of thermosolutes (MG, MGA, DGP, and DIP) was studied in two types of experiments to assess either their ability to prevent fibril formation or to disrupt preformed fibrils [42]. In the first type of experiment, solutes were added to the sample immediately after peptide dissolution; in the second type, the peptide was allowed to form fibrils for a week prior to solute addition. In both types of experiments, solute effects were qualitatively evaluated by EM at one and seven days after solute addition. The effect of solute concentration was also examined. All four solutes caused strong inhibition of fibril formation, clearly apparent at the first time point (one day) (Figure 2.4). In addition, they were remarkably effective in disassembling preformed fibrils (Figure 2.5). The magnitude of the effect increased with the solute concentration in both types of experiments. Curiously, the content of fibrils remained unchanged when samples were analyzed by EM on the seventh day of solute contact. Mannosylglycerate and MGA were the most effective solutes as inhibitors of fibril formation, while DIP and MG were the best to disassemble preformed fibrils. It should be pointed out, however, that KCl alone (used as a control for ionic strength) was also able to inhibit fibril formation and promote disassembly, although to a lesser extent.

UNDERSTANDING THE MOLECULAR BASIS OF PROTEIN STABILIZATION The effect of small organic solutes on protein stability (either stabilizers or denaturants) has been known for many decades but, the molecular principles responsible for this phenomenon are still unknown and a subject of controversy. From basic thermodynamic considerations (Wyman equation) and also from experimental measurements (mainly from Timasheff’s group) it is known that protecting solutes must be preferentially excluded from the native proteins, whereas denaturing compounds bind preferentially to the unfolded protein conformation. However, this theory gives no insight into the molecular mechanisms of solute/protein interactions. Hence, a unique conceptual

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Role in Protein Stabilization

15

FIGURE 2.4 Mannosylglyceramide as an inhibitor of amyloid fibril formation. Electron microscopy photographs of negatively stained fibrils derived from the model peptide STVIIE (magnification 2200×). The peptide (0.5 mM) was dissolved in glycine-HCL buffer (pH 2.6) supplemented with 0.5 mM (a), 25 mM (b) and 50 mM (c) mannosylglyceramide and fibrils were allowed to form, for seven days, at room temperature.

framework to explain, at the molecular level, the action of stabilizers (trehalose, betaine, proline, sucrose) or denaturants (urea) has been sought for many years. The effect of solutes on protein stability can result from alterations in the solvent properties or from more direct protein/solute interactions [43–46]. Recently, several proposals for the molecular mechanism by which solutes modulate protein stability have been put forward. Bolen et al. [47–49] determined the transfer-free energies of amino acid side chains or peptide bond analogs from water to solutions of several neutral solutes. It was shown that the major contribution to the stabilization free energy arises from the peptide bond transfer. Additionally, a negative correlation between the backbone transfer-free energy and the fractional polar surface area of several neutral solutes was found [50]. In other words, the interaction of the solute’s polar groups with the peptide backbone is more favorable than the interaction with the nonpolar groups. Hence, solutes with a large nonpolar surface area are better stabilizers than those highly polar. Despite the merit of this model, and as pointed out by the authors, its simplicity disregards other types of solute/protein interactions, like side-chain interactions or solute binding, not allowing for reliable extrapolations, namely to comprise charged solutes. Another essential aspect in the issue of protein stabilization is the impact of solutes on the three-dimensional water structure, which indirectly affects the protein properties. In simple terms, proteins induce water molecules to be arranged into two domains: a more structured water shell in the vicinity of the protein surface, and a less structured water domain in the bulk solvent. The protein hydration shell is composed of less polarizable water molecules and thus, less efficient in solvating polar solutes, which are excluded from this shell. On the other hand, nonpolar compounds or ions with high charge density will disrupt the complex hydrogen bonding network of bulk water and thus are clustered in the protein hydration shell [51].

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FIGURE 2.5 Effect of di-myo-inositol phosphate (DIP) in the disruption of mature amyloid fibrils derived from the model peptide STVIIE. Electron microscopy photographs of negatively stained fibrils (magnification 2200×). The peptide was dissolved in glycine-HCl buffer (pH 2.6) and incubated at room temperature for seven days. After this time, 50 mM DIP was added. EM images were recorded at one day (a) and seven days (b) after DIP addition. In parallel, a control assay without DIP was run. (c) This shows an image of the control recorded at the same time point as (a), that is: total incubation time of eight days.

The effect of solutes on the water protein hydration shell was investigated using hemoglobin complexed with pyranine, a fluorescent probe that binds to the protein surface and is highly sensitive to the capacity of the solvent to accept hydrogen bonds [52]. Stabilizing sugars (like trehalose) are not driven to the protein hydration shell, which becomes more structured. Conversely, addition of urea leads to a decrease in the hydration shell hydrogen bonding network, a result consistent with the increase of urea concentration in the vicinity of the protein surface. To evaluate the effect of solutes on the water structure, Batchelor et al. [53] quantified the heat exchange observed when a solute is added to water. These heat quantities are related to the solute’s ability to enhance or disrupt the hydrogen bonding network of water. Curiously, no correlation was found between the stabilizing ability of different solutes and their effect on water structure. Nevertheless, it is worth pointing out that these experiments were performed in pure water/solute solutions (no protein was present) which may be unsuited to model the interactions between solutes, water, and proteins. Most of the studies available deal with neutral solutes, but it is believed that the molecular principles involved are not widely different in the case of charged solutes. The electrostatic contributions of the ionic groups, however, must be taken into account. In molecular terms, the interactions of ions with proteins are thought to be mainly indirect, that is, mediated by the effect of the ions on the hydrogen bond properties of water [45,54]. However, direct interactions in which weak binding of the ion to the protein structure occurs, may also play a role [43,55]. The ability of ions to stabilize (salting-out) or destabilize (salting-in) proteins has been known since long and ions were ranked in the Hofmeister series. Correlations between this series and different physicochemical properties of the ion solutions were attempted, but most of them failed probably because Hofmeister effects cannot be explained by a single factor [56]. It is the balance between peptide groups salting-in and nonpolar groups salting-out for each ion that will define the global salt effect on protein stability [45,57].

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Role in Protein Stabilization

In conclusion, powerful models and methodologies have been developed to investigate the molecular principles responsible for the effect of solutes on proteins, but we are still far from understanding the complexity of protein/solute/water interactions and from being able to make reliable predictions on the practical outcome.

EFFECT OF THERMOSOLUTES ON THE PROTEIN UNFOLDING PATHWAY Time-resolved fluorescence spectroscopy is a very powerful technique that allows discrimination and quantification of the different protein conformational states (folded or denatured). This technique was used to investigate the effect of MG on the unfolding pathway of staphylococcal nuclease [17]. The fluorescence decay times are signatures of the protein states. In the range of temperatures examined (20–90°C) the decay of SNase fluorescence was characterized by three life times, the longest assigned to the native state and the other two originating from unfolded conformations. The temperature dependence of the fluorescence decay times in the presence of MG was identical to that observed in the absence of solutes (Figure 2.6). Thus, MG has no influence on the SNase conformational states detectable by this technique in spite of the great stabilization rendered by this solute (the Tm increased more than 7°C in the presence of 0.5 M MG). Hence, it can be concluded that the effect of MG upon SNase induces no detectable changes on the nature of protein conformations during the process of thermal denaturation, that is, the protein unfolding pathway remains unchanged.

IMPACT OF THERMOSOLUTES ON PROTEIN DYNAMICS The molecular basis of protein stabilization by thermosolutes was first investigated in the framework of possible structural changes, capable of explaining the added stability. However, in the cases studied, no measurable structural changes could be detected by nuclear magnetic resonance (NMR). For instance, in the presence of 100 mM DGP, a concentration capable of producing a fourfold

Fluorescence life-time (ns)

6.0

5.0

4.0

3.0

2.0

1.0

0.0 20

30

40

50

60

70

80

90

Temperature (ºC)

FIGURE 2.6 Temperature dependence of the fluorescence decay times of staphylococcal nuclease in the presence of 0.5 M mannosylglycerate (solid symbols) and in the absence of solutes (open symbols). Native protein: circles; denatured states of the protein: triangles. Fluorescence decay times of N-acetyl-tryptophanamide (an analog of tryptophan in a peptide chain) as a function of temperature in dioxane (dashed line) or water (solid line) are also plotted. Dioxane mimics the hydrophobic interior of a protein.

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increase in the half-life for thermal denaturation of rubredoxin from Desulfovibrio gigas, no structural alteration was detected [16]. In related experiments using “mesophilic solutes,” no structural effect was observed in chymotrypsin inhibitor 2 and horse heart cytochrome c upon addition of 2 M glycine [58]. These results point towards a stabilization phenomenon that takes place through changes in the solvent structure and/or by changing dynamic properties of the protein rather than causing substantial alterations in the protein structure itself. Actually, as protein structure is not dissociable from its function, stabilizing solutes, inducing structural changes in proteins, might seriously hamper enzyme activity, rendering the stabilizing effect useless. As stated before, proteins are marginally stable entities, whose structural design is a compromise between flexibility and stability at a given temperature. This concept brought about the principle of “corresponding states” which means that homologous proteins in their respective physiological conditions tend to keep similar flexibility, kinetic stability, solvation, and function [4,5]. Within this line of thought, flexibility and stability should be inversely correlated, that is, the less flexible, the more stable a protein would be [6]. Taking this statement to its ultimate conclusion, if compatible solutes are stabilizing agents, then these should be able to rigidify protein structure. In fact, there is evidence supporting this hypothesis. For example, MG is able to increase the temperature for optimum activity of RNase A, which suggests that this protein acquired a less flexible behavior in the presence of MG [37]. In addition, evidence for the compaction of protein structure induced by the presence of MG or DGP, as inferred by the reduction of chemical shift temperature dependence, was found in a D. gigas rubredoxin mutant [59]. However, the way in which thermosolutes are able to slow down protein motions is still to be answered and requires a thorough analysis of changes in the protein dynamic behavior in the presence of stabilizing agents. The study of protein dynamics is not trivial as proteins experience a variety of motions covering a wide range of time scales and amplitudes, from fast local oscillations to slower motions of whole structural elements [60,61]. NMR is a valuable tool to explore protein dynamics as it can access several time scales and different types of motions from small atomic vibrational modes to the concerted motion of whole segments of the protein structure [62,63]. Amide H/D exchange experiments reflect the latter type of internal mobility because in order for the exchange reaction to occur, a structural opening reaction (involving the displacement of large protein segments) has to take place [63]; while NMR relaxation measurements carry information on the local fast atomic fluctuations [62,64], events that take place in the 10 –10 to 10 –5 s time scale. The correlation between exchange rates and protein internal mobility/stability has been established for quite some time. In 1979, exchange rates were measured in a series of nine highly homologous proteins (chemical modifications of the basic pancreatic trypsin inhibitor), and were found to increase with decreasing denaturation temperatures [65]. The addition of urea or guanidinium chloride increases the exchange rates in a number of proteins tested [58,66,67], and this observation is interpreted as a consequence of increased internal mobility of protein structure brought about by the loosening of internal cohesive forces upon denaturant addition. A number of other corroborating studies produced a wealth of results establishing that, in the appropriate experimental conditions, proton exchange studies can provide information, not only on the global stability and unfolding, but also on local transient unfolding reactions and local stability. Thermosolutes also cause a marked reduction in amide H/D exchange rates, an effect that has been observed for DGP and MG in several rubredoxins, SNase, and SNase mutants [16,68,69], meaning that these solutes are able to strongly restrict wide, low-frequency protein motions. In the other end of the time scale, 15N-NMR relaxation measurements were used to build dynamic models for D. gigas rubredoxin and an SNase mutant in the presence of DGP and MG [16,69]. These models show a protein rigidification expressed by a small but generalized increase in the order parameters upon solute addition. Moreover, this rigidification increases with solute concentration. As solutes slow down protein internal motions, they could promote favorable internal interactions. One example is the strengthening of hydrogen bonds in a D. gigas rubredoxin mutant as a function of the concentration of DGP [59].

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Role in Protein Stabilization

In order to change protein dynamic behavior, without measurable structural alterations, stabilizing compatible solutes might act through changes in the solvent structure. One hypothesis is that by affecting the properties of the bulk solvent (like viscosity and surface tension) thermosolutes could cause a tighter solvation water structure. This hypothesis would explain why the wider low-frequency motions are more affected than the smaller high-frequency ones and it is compatible with the “preferential exclusion theory” [44]. Another explanation might be that solutes interact specifically, but transiently, with selected sites on the protein surface, promoting internal stabilizing interactions like the optimization of surface charge distribution [68,70], which in turn would cause a higher protein compaction and reduce its flexibility. This could help to explain why the magnitude of the stabilizing effect seems so dependent on the protein/solute pair under examination [15,25,59,71].

EFFECT OF THERMOSOLUTES ON THE PATHWAY OF FIBRIL FORMATION Amyloid fibrils are highly ordered aggregates with extensive β-sheet content and often result from misfolded proteins or partially unfolded ones [72,73]. The typical structure of all mature fibrils is fairly similar, especially if compared with the structure of the different proteins from where they originate [41,74,75]. Although the specific steps of fibril formation and maturation are still a matter of intense debate, the overall picture of the pathway is generally accepted. It is proposed that fibrillogenesis starts with the destabilization of the native structure of the protein, followed by a nucleation–extension mechanism leading to the mature fibril [76]. Circular dichroism (CD) spectroscopy was used to investigate the effect of thermosolutes (MG, DIP, DGP, and MGA) in the pathway of amyloid fibril formation [42]. The model peptide STVIIE mentioned in the section “Protein Conformational Stabilization by Thermosolutes” was also used in these experiments. Considerable amount of data is available on the pathway of fibril formation for this particular peptide [41,77,78]. During the process of fibril formation, the peptide changes from a predominantly random coil state to an almost pure β-sheet conformation, these soluble aggregates are then polymerized into the mature fibril. CD spectra of the model peptide, run at different times of peptide association, show the disappearance of the band at 198 nm (indicative of random coil conformation) and the appearance of a new band at 218 nm, typical of β-sheet conformation (Figure 2.7).

10 8

7d

6 θ (degrees)

4 2

41 h

0 -2

24 h

-4 -6

4h

-8 -10 -12 190

0h 200

210

220 230 240 Wavelength (nm)

250

260

FIGURE 2.7 Kinetics of self-association of peptide STVIIE dissolved in glycine-HCl buffer (20 mM, pH 2.6) without solutes, as monitored by circular dichroism. Samples were incubated at room temperature and aliquots were analyzed at different time points after dissolution. Solid circles, time zero; open squares, 4 h; triangles, 24 h; crosses, 41 h; solid squares, seven days. Each curve results from the accumulation of 10 scans.

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Thermophiles: Biology and Technology at High Temperatures 0.6

Fraction of structural motif

0.5

0.4

0.3

0.2

0.1

0.0 α Helix

β Strand

Turn

Unordered

FIGURE 2.8 Effect of solutes on the relative content of secondary structure motifs in fibrils of the peptide STVIIE. The peptide was dissolved in glycine-HCl buffer (pH 2.6) supplemented with different solutes at several final concentrations. A control without solutes was also prepared. After seven day incubation at room temperature aliquots were analyzed by circular dichroism. Three algorithms were used to deconvolute spectra (see text). Bar patterns: ( ) control (no solute); ( ) di-myo-inositol phosphate; ( ) diglycerol phosphate; ( ) potassium chloride. For each solute, bars are presented from left to right according to increasing concentrations (0.5, 25, and 50 mM).

The relative populations of the secondary structure motifs (α-helix, β-strand, turn and random coil) during fibril formation, in the presence of thermosolutes, were determined from CD spectra using three different deconvoluting algorithms (CDSSTR, SELCON3, and CONTINLL) [79,80], yielding essentially the same result. Despite a slight reduction of β-sheet content and concomitant increase in α-helical population, the presence of the solutes seems to have a negligible impact on the relative population of each structural motif (Figure 2.8). In other words, it is apparent that our analysis of the CD data is unable to discriminate between a control sample with high mature fibril content (as visualized by EM) and a sample in which the fibril content was drastically reduced as a result of the action of thermosolutes. Therefore, the ratio of secondary structure motifs accessible from CD spectroscopy does not distinguish final mature fibrils from intermediate polymeric forms, EM-invisible, in which β-sheet is already the predominant motif. We propose that thermosolutes prevent the assembly of the incipient polymeric peptide forms into the following stages of fibril formation. Further work is required to test this hypothesis.

CONCLUDING REMARKS There is no doubt that (hyper)thermophiles isolated from marine environments use low-molecular mass organic solutes for osmo- and thermoadaptation that are rarely or never found in mesophiles. Like solutes from mesophiles, they are mainly polyol and sugar derivatives, but these neutral building blocks are linked to glyceric acid or esterified with phosphoric acid, leading to molecules with a net negative charge. This is the most distinctive feature of solutes from organisms adapted to hot environments. Interestingly, evaluating the effect of charged solutes on the thermal stability of proteins in vitro has revealed their superior protective ability. Therefore, it is tempting to speculate that the accumulation of charged solutes by (hyper)thermophiles is part of a mechanism of extrinsic

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thermal stabilization that evolved in organisms from hot marine environments. The mechanisms of protein stabilization by organic solutes are very complex, involve a large number of subtle contributions, and are still poorly understood. The importance of these studies, however, is unquestionable given the range of human activities that could benefit from understanding the molecular interactions underlying protein stabilization. In particular, the chaperone effect demonstrated by several solutes holds a tremendous potential for the treatment or prevention of many conformational diseases that afflict modern society.

ACKNOWLEDGMENTS This work was funded by the European Commission Contract COOP-CT-2003-508644 and Fundação para a Ciência e a Tecnologia and FEDER, Portugal, POCTI/BIA-PRO/57263/2004 and POCTI/BIA-MIC/59310/2004. P. Lamosa, and T.Q. Faria acknowledge grants from FCT, Portugal (BPD/26606/2006 and BPD/20352/2004). The experiments involving amyloid fibrils were performed by T.M. Pais at EMBL. The assistance of A. Esteras-Chopo is gratefully acknowledged.

REFERENCES 1. 2. 3. 4.

5. 6. 7. 8. 9.

10.

11. 12.

13.

14.

15.

9214_C002.indd 21

Stetter, K.O., History of discovery of the first hyperthermophiles, Extremophiles, 10, 357, 2006. Rothschild, L.J. and Mancinelli, R.L., Life in extreme environments, Nature, 409, 1092, 2001. Hensel, R., Proteins of extreme thermophiles, New Comp. Biochem., 26, 209, 1993. Jaenicke, R., Schurig, H., Beaucamp, N., and Ostendorp, R., Structure and stability of hyperstable proteins: glycolytic enzymes from hyperthermophilic bacterium Thermotoga maritima, Adv. Protein. Chem., 48, 181, 1996. Daniel, R.M. and Danson, M.J., Assaying activity and assessing thermostability of hyperthermophilic enzymes, Methods Enzymol., 334, 283, 2001. Fields, P.A., Protein function at thermal extremes: balancing stability and flexibility, Comp. Biochem. Physiol. A, 129, 417, 2001. Li, W.F., Zhou, X.X., and Lu, P., Structural features of thermozymes, J. Biotech. Adv., 23, 271, 2005. Hensel, R. and König, H., Thermoadaptation of methanogenic bacteria by intracellular ion concentration, FEMS Microbiol. Lett., 49, 75, 1988. Zwickl, P., Fabry, S., Bogedain, C., Haas, A., and Hensel, R., Glyceraldehyde-3-phosphate dehydrogenase from the hyperthermophilic archaebacterium Pyrococcus woesei: characterization of the enzyme, cloning and sequencing of the gene, and expression in Escherichia coli, J. Bacteriol., 172, 4329, 1990. Schmitz, R.A., Linder, D., Stetter, K.O., and Thauer, R.K., N5,N10 -Methylenetetrahydromethanopterin reductase (coenzyme F420-dependent) and formylmethanofuran dehydrogenase from the hyperthermophile Archaeoglobus fulgidus, Arch. Microbiol., 156, 427, 1991. Hensel, R., Fabry, S., Biro, J., Bogedain, C., Jakob, I., and Siebers, B., Glyceraldehyde-3-phosphate dehydrogenases from archaea: objects for studying protein thermoadaptation, Biocatalysis, 11, 151, 1994. Steen, I.H., Lien, T., and Birkeland, N.K., Biochemical and phylogenetic characterization of isocitrate dehydrogenase from a hyperthermophilic archaeon, Archaeoglobus fulgidus, Arch. Microbiol., 168, 412, 1997. Aálen, N., Steen, I.H., Birkeland, N.K., and Lien, T., Purification and properties of an extremely thermostable NADP+-specific glutamate dehydrogenase from Archaeoglobus fulgidus, Arch. Microbiol., 168, 536, 1997. Ramos, A., Raven, N.D.H., Sharp, R.J., Bartolucci, S., Rossi, M., Cannio, R., Lebbink, J., van der Oost, J., de Vos, W.M., and Santos, H., Stabilization of enzymes against thermal stress and freeze-drying by mannosylglycerate, Appl. Environ. Microbiol., 63, 4020, 1997. Borges, N., Ramos, A., Raven, N.D.H., Sharp, R.J., and Santos, H., Comparative study of the thermostabilizing properties of mannosylglycerate and other compatible solutes on model enzymes, Extremophiles, 6, 209, 2002.

11/5/2007 7:21:54 AM

22

Thermophiles: Biology and Technology at High Temperatures

16. Lamosa, P., Turner, D.L., Ventura, R., Maycock, C., and Santos, H., Protein stabilization by compatible solutes. Effect of diglycerol phosphate on the dynamics of Desulfovibrio gigas rubredoxin studied by NMR, Eur. J. Biochem., 270, 4606, 2003. 17. Faria, T.Q., Lima, J.C., Bastos, M., Maçanita, A.L., and Santos, H., Protein stabilization by osmolytes from hyperthermophiles: effect of mannosylglycerate on the thermal unfolding of recombinant nuclease A from Staphylococcus aureus studied by picosecond time-resolved fluorescence and calorimetry, J. Biol. Chem., 279, 48680, 2004. 18. Santos, H. and da Costa, M.S., Compatible solutes of organisms that live in hot saline environments, Environ. Microbiol., 4, 501, 2002. 19. Santos, H., Lamosa, P., Faria, T.Q., Borges, N., and Neves, C., The physiological role, biosynthesis and mode of action of compatible solutes from (hyper)thermophiles, in Physiology and Biochemistry of Extremophiles, Gerday, C. and Glandorff, N., Eds., ASM publishers, 2007. 20. Brown, A.D., Microbial water stress, Bacteriol. Rev., 40, 803, 1976. 21. da Costa, M.S., Santos, H., and Galinski, E.A., An overview of the role and diversity of compatible solutes in Bacteria and Archaea, Adv. Biochem. Eng. Biotechnol., 61, 117, 1998. 22. Oren, A., Bioenergetic aspects of halophilism, Microbiol. Mol. Biol. Rev., 63, 334, 1999. 23. Scholz, S., Sonnenbichler, J., Schäfer, H., and Hensel, R., Di-myo-inositol-1,1’-phosphate: a new inositol phosphate isolated from Pyrococcus woesei, FEBS Lett., 306, 239, 1992. 24. Shima, S., Hérault, D.A., Berkessel, A., and Thauer, R.K., Activation and thermostabilization effects of cyclic 2,3-diphosphoglycerate on enzymes from the hyperthermophilic Methanopyrus kandleri, Arch. Microbiol., 170, 469, 1998. 25. Lamosa, P., Burke, A., Peist, R., Huber, R., Liu, M.Y., Silva, G., Rodrigues-Pousada, C., LeGall, J., Maycock, C., and Santos, H., Thermostabilization of proteins by diglycerol phosphate, a new compatible solute from the hyperthermophile Archaeoglobus fulgidus, Appl. Environ. Microbiol., 66, 1974, 2000. 26. Santos, H., Lamosa, P., and Borges, N., Characterization of organic compatible solutes of thermophilic microorganisms, Methods Microbiol., 35, 171, 2006. 27. Bouveng, H., Lindberg, B., and Wickberg, B., Low-molecular carbohydrates in algae, Acta Chem. Scand., 9, 807, 1955. 28. Martins, L.O. and Santos, H., Accumulation of mannosylglycerate and di-myo-inositol-phosphate by Pyrococcus furiosus in response to salinity and temperature, Appl. Environ. Microbiol., 61, 3299, 1995. 29. Martins, L.O., Huber, R., Huber, H., Stetter, K.O., da Costa, M.S., and Santos, H., Organic solutes in hyperthermophilic Archaea, Appl. Environ. Microbiol., 63, 896, 1997. 30. Lamosa, P., Martins, L.O., da Costa, M.S., and Santos, H., Effects of temperature, salinity, and medium composition on compatible solute accumulation by Thermococcus spp., Appl. Environ. Microbiol., 64, 3591, 1998. 31. Gonçalves, L.G., Huber, R., da Costa, M.S., and Santos, H., A variant of the hyperthermophile Archaeoglobus fulgidus adapted to grow at high salinity, FEMS Microbiol. Lett., 218, 239, 2003. 32. Neves, C., da Costa, M.S., and Santos, H., Compatible solutes of the hyperthermophile Palaeococcus ferrophilus: osmoadaptation and thermoadaptation in the order Thermococcales, Appl. Environ. Microbiol., 71, 8091, 2005. 33. Lamosa, P., Rodrigues, M.V., Carr, J., Mingote, A., Ventura, R., Maycock, C., Raven, N.D., and Santos, H., Organic solutes in members of the Aquificales and the Thermotogales: occurrence of (1-6)glucosyl(1-2) glucosyl-glycerate in persephonella marina. In preparation, 2007. 34. Lamosa, P., Gonçalves, L.G., Rodrigues, M.V., Martins, L.O., Raven, N., and Santos, H., Occurrence of 1-glyceryl-1-myo-inosityl-phosphate in hyperthermophiles, Appl. Environ. Microbiol., 72, 6169, 2006. 35. Empadinhas, N., Marugg, J.D., Borges, N., Santos, H., and da Costa, M.S., Pathway for the synthesis of mannosylglycerate in the hyperthermophilic archaeon Pyrococcus horikoshii. Biochemical and genetic characterization of key enzymes, J. Biol. Chem., 276, 43580, 2001. 36. Faria, T.Q., Mingote, A., Siopa, F., Ventura, R., Maycock, C., and Santos, H., Mannosylglycerate as a lead compound for designing new protein stabilizers, unpublished data, 2007. 37. Faria, T.Q., Knapp, S., Ladenstein, R., Maçanita, A.L., and Santos, H., Protein stabilisation by compatible solutes: effect of mannosylglycerate on unfolding thermodynamics and activity of ribonuclease A, Chem. Bio. Chem., 4, 734, 2003.

9214_C002.indd 22

11/5/2007 7:21:54 AM

Role in Protein Stabilization

23

38. Selkoe, D.J., Aging, amyloid, and Alzheimer’s disease: a perspective in honor of Carl Cotman, Neurochem. Res., 28, 1705, 2003. 39. Dobson, C.M., Protein folding and misfolding, Nature, 426, 884, 2003. 40. Bernier, V., Lagacé, M., Bichet, D.G., and Bouvier, M., Pharmacological chaperones: potential treatment for conformational diseases, Trends Endocrinol. Metab., 15, 222, 2004. 41. López de la Paz, M., Goldie, K., Zurdo, J., Lacroix, E., Dobson, C.M., Hoenger, A., and Serrano, L., De novo designed peptide-based amyloid fibrils, Proc. Natl. Acad. Sci. USA, 99, 16052, 2002. 42. Pais, T.M., Esteras-Chopo, A., López de la Paz, M., Serrano, L., and Santos, H., Solutes from hyperthermophiles as inhibitors of amyloid fibril formation, unpublished data, 2007. 43. Ramos, C.H.I. and Baldwin, R.L., Sulfate anion stabilization of native ribonuclease A both by anion binding and the Hofmeister effect, Protein Sci., 11, 1771, 2002. 44. Timasheff, S.N., The control of protein stability and association by weak interactions with water: how do solvents affect these processes? Annu. Rev. Biophys. Biomol. Struct., 22, 67, 1993. 45. Baldwin, R.L., How Hofmeister ion interactions affect protein stability, Biophys. J., 71, 2056, 1996. 46. Davis-Searles, P.R., Saunders, A.J., Erie, D.A., Winzor, D.J., and Pielak, G.J., Interpreting the effect of small uncharged solutes on protein-folding equilibria, Annu. Rev. Biophys. Biomol. Struct., 30, 271, 2001. 47. Liu, Y. and Bolen, D.W., The peptide backbone plays a dominant role in protein stabilization by naturally occurring osmolytes, Biochemistry, 34, 12884, 1995. 48. Qu, Y., Bolen, C.L., and Bolen, D.W., Osmolyte-driven contraction of a random coil protein, Proc. Natl. Acad. Sci. U S A, 95, 9268, 1998. 49. Auton, M. and Bolen, D.W., Predicting the energetics of osmolyte-induced protein folding/unfolding, Proc. Natl. Acad. Sci. U S A, 102, 15065, 2005. 50. Street, T.O., Bolen, D.W., and Rose, G.D., A molecular mechanism for osmolyte-induced protein stability, Proc. Natl. Acad. Sci. U S A, 103, 13997, 2006. 51. Raschke, T., Water structure and interactions with protein structures, Curr. Opin. Struct. Biol., 16, 155, 2006. 52. Roche, C.J., Guo, F., and Friedman, J.M., Molecular level probing of preferential hydration and its modulation by osmolytes through the use of pyranine complexed to hemoglobin, J. Biol. Chem., 281, 38757, 2006. 53. Batchelor, J.D., Olteanu, A., Tripathy, A., and Pielak, G.J., Impact of protein denaturants and stabilizers on water structure, J. Am. Chem. Soc., 126, 1958, 2004. 54. Leberman, R. and Soper, A.K., Effect of high salt concentrations on water structure, Nature, 378, 364, 1995. 55. Makhatadze, G.I., Lopez, M.M., Richardson III, J.M., and Thomas, S.T., Anion binding to the ubiquitin molecule, Protein Sci., 7, 689, 1998. 56. Kunz, W., Lo Nostro, P., and Ninham, B.W., The present state of affairs with Hofmeister effects, Curr. Opin. Colloid Interface Sci., 9, 1, 2004. 57. Cacace, M.G., Landau, E.M., and Ramsden, J.J., The Hofmeister series: salt and solvent effects on interfacial phenomena, Q. Rev. Biophys., 30, 241, 1997. 58. Foord, R.L. and Leatherbarrow, R.J., Effect of osmolytes on the exchange rates of backbone amide protons in proteins, Biochemistry, 37, 2969, 1998. 59. Pais, T.M., Lamosa, P., dos Santos, W., LeGall, J., Turner, D.L., and Santos, H., Structural determinants of protein stabilization by solutes. The importance of the hairpin loop in rubredoxins, FEBS J., 272, 999, 2005. 60. Karplus, M. and McCammon, J.A., Dynamics of proteins: elements and function, Annu. Rev. Biochem., 52, 263, 1983. 61. Gerstein, M., Lesk, A.M., and Chothia, C., Structural mechanisms for domain movements in proteins, Biochemistry, 33, 6739, 1994. 62. Palmer, A.G., NMR probes of molecular dynamics: overview and comparison with other techniques, Annu. Rev. Biophys. Biomol. Struct., 30, 129, 2001. 63. Englander, S.W., Mayne, L., Bai, Y., and Sosnick, T.R., Hydrogen exchange: the modern legacy of Linderstrøm-Lang, Protein Sci., 6, 1101, 1997. 64. Lipari, G. and Szabo, A., Model-free approach to the interpretation of nuclear magnetic resonance relaxation in macromolecules. I. Theory and range of validity, J. Am. Chem. Soc., 104, 4546, 1982. 65. Wagner, G. and Wüthrich, K., Correlation between the amide proton exchange rates and the denaturation temperatures in globular proteins related to the basic pancreatic trypsin inhibitor, J. Mol. Biol., 130, 31, 1979.

9214_C002.indd 23

11/5/2007 7:21:55 AM

24

Thermophiles: Biology and Technology at High Temperatures

66. Loftus, D., Gbenle, G.O., Kim, P.S., and Baldwin, R.L., Effects of denaturants on amide proton exchange rates: a test for structure in protein fragments and folding intermediates, Biochemistry, 25, 1428, 1986. 67. Bai, Y., Englander, J.J., Mayne, L., Milne, J.S., and Englander, S.W., Thermodynamic parameters from hydrogen exchange measurements, Methods Enzymol., 259, 344, 1995. 68. Lamosa, P., Brennan, L., Vis, H., Turner, D.L., and Santos, H., NMR structure of Desulfovibrio gigas rubredoxin: a model for studying protein stabilization by compatible solutes, Extremophiles, 5, 303, 2001. 69. Pais, T.M., Lamosa, P., Turner, D.L., García-Moreno, B., and Santos, H., NMR study of the effect of mannosylglycerate on the dynamics of staphylococcal nuclease, unpublished data, 2007. 70. Spassov, V.Z., Karshikoff, A.D., and Ladenstein, R., The optimization of protein–solvent interactions: thermostability and the role of hydrophobic and electrostatic interactions, Protein Sci., 4, 1516, 1995. 71. Lippert, K. and Galinski, E.A., Enzyme stabilization by ectoine-type compatible solutes: protection against heating, freezing and drying, Appl. Microbiol. Biotechnol., 37, 61, 1992. 72. Dobson, C.M., Protein misfolding, evolution and disease, Trends Biochem. Sci., 24, 329, 1999. 73. Kelly, J.W., The alternative conformations of amyloidogenic proteins and their multi-step assembly pathways, Curr. Opin. Struct. Biol., 8, 101, 1998. 74. Jimenez, J.L., Guijarro, J.I., Orlova, E., Zurdo, J., Dobson, C.M., Sunde, M., and Saibil, H.R., Cryo-electron microscopy structure of an SH3 amyloid fibril and model of the molecular packing, EMBO J., 18, 815, 1999. 75. Nelson, R. and Eisenberg, D., Recent atomic models of amyloid fibril structure, Curr. Opin. Struct. Biol., 16, 260, 2006. 76. Higurashi, T., Yagi, H., Mizobata, T., and Kawata, Y., Amyloid-like fibril formation of co-chaperonin GroES: nucleation and extension prefer different degrees of molecular compactness, J. Mol. Biol., 351, 1057, 2005. 77. López de la Paz, M. and Serrano, L., Sequence determinants of amyloid fibril formation, Proc. Natl. Acad. Sci. USA, 101, 87, 2004. 78. López de la Paz, M., de Mori, G.M., Serrano, L., and Colombo, G., Sequence dependence of amyloid fibril formation: insights from molecular dynamics simulations, J. Mol. Biol., 349, 583, 2005. 79. Sreerama, N. and Woody, R.W., Estimation of protein secondary structure from circular dichroism spectra: comparison of CONTIN, SELCON, and CDSSTR methods with an expanded reference set, Anal. Biochem., 7, 252, 2000. 80. Sreerama, N., Venyaminov, S.Y., and Woody, R.W., Estimation of protein secondary structure from circular dichroism spectra: inclusion of denatured proteins with native proteins in the analysis, Anal. Biochem., 287, 243, 2000.

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Relationships among Catalytic Activity, Structural Flexibility, and Conformational Stability as Deduced from the Analysis of Mesophilic– Thermophilic Enzyme Pairs and Protein Engineering Studies Reinhard Sterner and Eike Brunner

CONTENTS Introduction .................................................................................................................................. Global and Local Probing of Structural Flexibility Correlated with Enzymatic Activity and Thermal Stability ....................................................................................................... Enzyme Engineering to Increase Stability or Catalytic Activity ................................................ Rational Design ................................................................................................................ Directed Evolution ............................................................................................................ Conclusions .................................................................................................................................. References ....................................................................................................................................

25 26 30 30 30 34 35

INTRODUCTION Naturally occurring enzymes must be stable to maintain their native structures even under unfavorable circumstances, for example at elevated temperatures or in the presence of harsh chemical conditions. On the other hand, enzymes also need to be sufficiently flexible to perform their various catalytic activities. As a consequence, conformational stability may have been partially sacrificed for functional reasons during the evolution of enzymes. This idea has been supported by the generation of active site substitutions that resulted in stabilized enzymes with reduced activities [1–5]. The comparison of enzymes from thermophiles and hyperthermophiles with their homologues from mesophiles can provide insights into the problem of enzyme activity and stability, as well as 25

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the correlation of these properties with protein dynamics. Extremely stable enzymes from hyperthermophiles, which optimally grow close to the boiling point of water, are often barely active at room temperature. However, they are as active as their thermolabile mesophilic counterparts at the corresponding physiological temperatures [6]. The differences in activity must be caused by subtle structural alterations in the protein chain, because the active site residues are conserved between homologous mesophilic and hyperthermophilic enzymes [7]. A wealth of experimental and theoretical evidence suggests that conformational dynamics is important for enzymatic catalysis [8]. Accordingly, it has been postulated that the low activity and high stability of enzymes from hyperthermophiles is due to a restricted structural flexibility, providing a high energetic barrier for both catalysis and unfolding. The conformational rigidity would be relieved at elevated temperatures, resulting in comparable activities and stabilities as observed for mesophilic enzymes at moderate temperatures (concept of “corresponding states” [9]). The present contribution briefly summarizes comparative investigations of homologous enzymes from hyperthermophiles and mesophiles as well as protein engineering approaches, which provided insights into the relationship between catalytic activity, structural flexibility, and conformational stability.

GLOBAL AND LOCAL PROBING OF STRUCTURAL FLEXIBILITY CORRELATED WITH ENZYMATIC ACTIVITY AND THERMAL STABILITY In agreement with the “corresponding states” concept, an inverse correlation between structural flexibility and conformational stability was monitored for a number of proteins by various techniques [10]. For example, the 3-phosphoglycerate kinase (PGK) from Thermus thermophilus is more stable than the homologous enzyme from yeast at a given temperature. In contrast, its catalytic activity is reduced and structural fluctuations of the protein scaffold as measured by the ability of acrylamide to access a buried tryptophan residue and quench its fluorescence are diminished. However, the flexibilities, stabilities, and catalytic activities of the two enzymes are similar when compared at the physiologically relevant temperatures which amount to 75°C for T. thermophilus and 25°C for yeast [11]. This result is relevant because PGK operates through a hinge-bending mechanism suggesting that its activity depends on large-scale conformational dynamics [12]. Likewise, a comparative study of three α-amylases showed that their thermal stability is inversely correlated with catalytic activity and conformational flexibility as measured by acrylamide-induced fluorescence quenching [13]. Reversible conformational fluctuations expose buried segments of the polypeptide chain to solvent molecules. When the protein is dissolved in D2O, hydrogen exchange measurements can be used to detect the solvent accessibility of backbone amide hydrogen atoms. A criterion for the rigidity of a protein is the rate constant of this exchange. Conformationally protected amide hydrogen atoms exchange with solvent hydrogen according to the general scheme kop

closed

´ kcl

kCh

open

Æ

exchanged

(3.1)

where kop, kcl, and kCh denote the rates for the conformational opening/closing process and the hydrogen exchange in the open state, respectively [14]. As an exchange can only occur after a conformational opening transition, the effective exchange rate, kex, detected in exchange experiments will be given by: kex = kopkCh/(kop + kcl + kCh).

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(3.2)

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Two different exchange regimes denoted as EX1 and EX2 must be distinguished. So-called EX1 conditions are present if the conformational opening step is rate-limiting in Equation 3.1, that is, if kop, kcl << kCh. Equation 3.2 then approximates to kex = kop. Very often, however, the rates of conformational opening and closing, kop and kcl, are significantly larger than kCh. These so-called EX2 conditions are usually observed for proteins. Equation 3.2 then yields: kex = ρ ⋅ kCh

(3.3)

where ρ = [open]/([open]+[closed]) denotes the fraction of molecules found in the open conformation [14]. The hydrogen exchange rate is given by kCh = {[H+] + 108 [OH−]}10[0.05 (θ−25)] s−1

(3.4)

where θ denotes the temperature in degrees centigrade [15]. Conventional hydrogen/deuterium (H/D) exchange experiments have been exploited for a long time [15] and were recently used to compare the flexibilities of thermophilic and mesophilic proteins. For example, hydrogen exchange kinetic studies were performed under EX2 conditions on the thermostable isopropylmalate dehydrogenase from T. thermophilus and its less stable homologue from Escherichia coli. The H/D exchange was monitored by Fourier-transform infra-red (FTIR) spectroscopy by observing the intensity decrease of the amide II band around 1550 cm–1 characteristic of N–H groups and the concomitant increase of the band around 1450 cm–1 characteristic of N–D groups as well as HDO molecules. The fraction of nonexchanged N–H groups, X, was recorded as a function of time and plotted in the form of the so-called relaxation spectra (X versus kCh⋅t) [16]. The shape of the relaxation spectra indicates that both enzymes do not expose their amide hydrogen atoms in a cooperative way. Instead, local movements with varying probabilities occur for the different parts of the protein. Overall, significant differences between the T. thermophilus and the E. coli enzyme were found at 25°C, indicating a considerably higher flexibility for the latter molecule at this temperature. However, the flexibilities are similar when comparing the two enzymes at their respective optimum activity temperatures of 70°C and 48°C; an observation completely in line with the “corresponding states” model [17]. On the other hand, isopropylmalate dehydrogenase from the psychrophile Vibrio sp. I5 was found to be more labile and more active but globally less flexible than its homologue from E. coli as indicated by H/D exchange measurements [18]; an observation which apparently contradicts the “corresponding states” model. Furthermore, comparative studies of two differently thermostable α-amylases from Bacillus amyloliquefaciens (melting temperature Tm = 86°C) and Bacillus licheniformis (Tm = 101°C) have shown that the picosecond timescale fluctuations detected by incoherent neutron scattering as well as the motions reflected by tryptophan fluorescence quenching are not related to the thermal stability or catalytic activity of the enzymes. Again, a higher flexibility is observed for the more thermostable protein from B. licheniformis in contrast to the predictions of the “corresponding states” model [19,20]. A problem of these studies is that only a localized (fluorescence) or a structurally averaged (H/D exchange monitored by FTIR, neutron scattering) quantity reflecting the conformational dynamics is monitored. In contrast to the aforementioned techniques, solution nuclear magnetic resonance (NMR) spectroscopy of 13C- and 15N-labelled proteins offers a spectral resolution which allows the assignment of the different backbone as well as side chain resonances caused by the various amino acid residues. Therefore, NMR spectroscopy allows the site-specific measurement of parameters influenced by the structure, conformation as well as internal mobility. Based on the common multidimensional NMR techniques [21], the structure of the cold-shock protein from the hyperthermophilic bacterium Thermotoga maritima could be determined at room temperature as well as at 70°C [22,23]. A comparison of the two structures revealed that the overall conformation is—apart

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FIGURE 3.1 Comparison of the backbone structure (ribbon plot) of the cold shock protein from Thermotoga maritima at 30°C (a) and 70°C (b, c). The arrow in the the 30°C structure points to a β-bulge. In the 70°C structure, the salt bridge Lys6-Asp24 (b) and the ion cluster around Arg2 (c) are indicated. It can be seen that the secondary structure elements are less defined at 70°C. For example, β1 is shortened at elevated temperature. Note the pronounced change of the loop between β4 and β5. In contrast, the structure remains to be well defined at high temperature in the neighborhood of the salt bridge and ion cluster. (Reprinted from Jung, A. et al., Protein Sci. 13, 342, 2004. With permission from Cold Spring Harbor Laboratory Press.)

from subtle changes—conserved at 70°C. However, the high-temperature conformation seems to be less well defined and more flexible than the room temperature conformation, except for a region stabilized by certain electrostatic interactions (Figure 3.1). These observations are fully in line with the predictions of the “corresponding states” concept. Besides three-dimensional structure determination, NMR spectroscopy also allows the site-specific measurement of other parameters influenced by the presence of intramolecular motions or conformational exchange processes. For example, a combination of NMR with the H/D exchange technique was used to measure stability parameters, in particular the free energy of unfolding, for individual amino acid residues [24]. Corresponding experiments were carried out and compared for the ribonuclease H from T. thermophilus and its mesophilic homologue from E. coli. Interestingly, a similar distribution along the amino acid sequence of the thus-defined stability parameters was found for both proteins being accompanied by a proportional increase of stability in the thermophilic protein. This observation led the authors to the conclusion that the increased stability of the thermophilic protein was due to a delocalized effect rather than the introduction of only few stabilizing elements. They therefore state the following: “although protein stability can be altered by single amino acid substitutions, evolution for optimal function may require more subtle and delocalized mechanisms” [24]. In addition to the H/D exchange method, NMR spectroscopic experiments were designed for the detection of pure 1H–1H exchange processes between backbone amide groups and the surrounding solvent molecules without the use of any deuterated compounds. A favorable experiment for such studies is the so-called CLEANEX-PM approach (phase-modulated CLEAN chemical exchange spin locking) combined with fast heteronuclear single quantum coherence (FHSQC) detection [25,26]. It removes the undesired intramolecular nuclear Overhauser effect as well as rotating frame Overhauser effect contributions to the exchange peaks and detects only chemically exchanging 1H nuclei. This approach was used to monitor differences in the conformational dynamics of the rubredoxins from the hyperthermophile Pyrococcus furiosus and the mesophile Clostridium pasteurianum in a spatially resolved manner [27]. The time window provided by this technique approximately

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corresponds to 0.1 s–1 < kex < 100 s–1, that is, it spans three orders of magnitude. Under alkaline conditions, kex will be proportional to [OH–] (see Equation 3.4). It is then useful to introduce the quantity kOH = kex/[OH–], which covers almost nine orders of magnitude [27]. Site-specific activation enthalpies for the amide exchange kinetics could be derived from the temperature dependence of kOH. Based on this technique, it could be shown that the conformational flexibility of a special multi-turn region in rubredoxin from the hyperthermophile P. furiosus exhibits a larger apparent flexibility below room temperature than in rubredoxin from the mesophile C. pasteurianum. This observation contradicts the expectation of increased rigidity in proteins from hyperthermophiles at lower temperatures. However, a reduced temperature dependence of the collective opening process in the multi-turn region was observed for the hyperthermophile compared with the mesophile rubredoxin. The resulting lower activation enthalpy would lead to a “flattening” of the Gibbs free energy versus temperature profile, which may account for the higher net temperature stability of the hyperthermophilic protein [27]. These observations could be further corroborated and extended by subsequent studies of rubredoxin hybrids making use of the same NMR spectroscopic techniques [28,29]. Other useful NMR spectroscopic parameters are, for example, the longitudinal and transverse self-relaxation rates R1 and R2 of nuclei such as 1H and 15N, cross-relaxation rates as well as heteronuclear 1H–15N nuclear Overhauser effect intensities. Based on such measurements as well as chemical shift analyses, the dynamics of an enzyme, human cyclophilin A (CypA), was studied during its catalytic action [30]. Strongly increased R2 values were observed for certain amino acid residues. This behavior could be explained by the presence of structural transitions between different molecular conformations on the time scale of hundreds of microseconds. The determination of the exchange contribution, Rex, to the transverse 15N relaxation rate R2 allowed a quantitative study of the underlying conformational exchange processes. This technique was applied in subsequent studies to understand the collective nature of conformational motions and their importance in biocatalysis [31,32]. It could, for example, be shown that intrinsic motions in CypA are already present in its substrate-free state and are a necessary precondition for the catalytic activity of the enzyme. Moreover, the frequencies of these internal motions, which are not restricted to the active site but suggest the existence of a dynamic network, coincide with the turnover rate [32]. Relaxation studies have also been performed with a pair of mesophilic and thermophilic adenylate kinases. The results indicated that the turnover numbers of both enzymes are limited by the dynamic opening/ closing of the nucleotide-binding lid, which is a prerequisite for the rate-determining release of the product. The frequency of this motion and—consequently—the turnover number is higher in the mesophilic than in the thermophilic enzyme, in agreement with the “corresponding states” concept [31]. In addition, Bae and Phillips (2006) have characterized various thermophilic–mesophilic adenylate kinase chimeras [33]. These studies suggested that the stability of adenylate kinase is determined by a special region (the so-called CORE) and is not influenced by the active site lid and adenosine monophosphate (AMP)-binding domains. In other words, local flexibility and mobility seem to control the catalytic activity of this enzyme without compromising its overall stability. Studies on alcohol dehydrogenase (ADH) have shown that hydrogen tunnelling, that is, the transfer of hydrogen from alcohols to enzyme molecules via the quantum-mechanical tunnel effect, is crucial for the dehydrogenation reaction [34,35]. The investigation of the ADH from the thermophilic bacterium Bacillus stearothermophilus revealed that hydrogen tunneling significantly contributes to its catalytic activity at the physiological temperature of 65°C [36]. In contrast to theoretical predictions, however, the influence of tunneling on catalysis drops down significantly below 30°C, suggesting that thermally excited fluctuations are a necessary condition for tunneling in ADH. In accordance with this hypothesis, the contribution of tunneling to hydrogen transfer correlates with the dynamics and flexibility of the ADH molecules as investigated by FTIRmonitored H/D exchange measurements [37]. In agreement with the predictions of the “corresponding states” model, the flexibilities of the ADH from the mesophilic yeast at 25°C and of B. stearothermophilus ADH at 65°C are similar [34,36].

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ENZYME ENGINEERING TO INCREASE STABILITY OR CATALYTIC ACTIVITY Naturally occurring enzymes have not evolved to maximum stability, flexibility, or catalytic activity, but represent a physiologically meaningful compromise between these properties. This opens the possibility to improve either enzyme stability or activity in the laboratory, and to measure the effect of this optimization on the other parameters. Along these lines, rational design and directed laboratory evolution have been used to deepen our understanding of the relationship between enzyme function, dynamics, and stability.

RATIONAL DESIGN In a rational design experiment, defined changes in the amino acid sequences of enzymes are planned on the basis of a preconceived idea, and then introduced by site-directed mutagenesis. In straightforward approaches, thermolabile enzymes could be stabilized by introducing residues found in their homologous counterparts from hyperthermophilic microorganisms [6,38] or by incorporating consensus or ancestral residues identified by multiple sequence alignments [39,40]. Whereas these mutational studies confirmed the view that proteins can be stabilized by various mechanisms [7,41,42], they emphasized the significance of salt bridges and long-range electrostatic interactions on the surface for the conformational integrity of proteins from hyperthermophiles [43,44]. These interactions had been considered largely irrelevant for protein stability on the basis of theoretical considerations and mutational studies of proteins from mesophiles [45]. With respect to function, the comparison of enzymes with similar structures and catalytic mechanisms allowed to alter substrate or co-factor specificities and stereoselectivities, and to establish novel catalytic activities on existing structural scaffolds by the exchange of only few amino acid residues [46,47]. In contrast to the “corresponding states” hypothesis, some rationally designed hyperstable enzymes retained high activity at low temperatures, as was for example shown for a triosephosphate isomerase [48] and a metallo-protease [39].

DIRECTED EVOLUTION A powerful alternative to rational design is the directed evolution of enzymes. With this technique, catalytic activity can be improved by exchanging only a few amino acid residues, given the availability of a powerful screening or selection system [49–52]. In a directed evolution approach, random mutagenesis is used to create a repertoire of modified enzymes, from which beneficial variants are isolated. The creation of genetic diversity is largely independent of the particular enzyme to be optimized. In contrast, finding the desired variants is unique for each biocatalyst and employs either selection (which couples the survival of the host with the newly acquired property of an enzyme and allows to test up to about 1010 variants), or screening (which quantifies the derived property of each enzyme variant individually and allows to test up to about 106 variants). One advantage of directed molecular evolution compared with rational design is that new properties can be generated without a detailed a priori knowledge of the structure or mechanism of the enzyme under study. Moreover, directed evolution is a largely unbiased approach because the molecular answer to a given selection pressure cannot be predicted. Instead, the experimental system is open to find solutions beyond the current knowledge and, therefore, can lead to unexpected insights. For example, the enantioselectivities and substrate specificities of aminotransferases and lipases were improved by the exchange of amino acids that were located far away from the active site [53–55]. Furthermore, directed evolution is well suited to elucidate the structural basis of protein thermostability because it ideally introduces only the minimum number of amino acid exchanges that are necessary for stabilization. The consequences of activation for stability and flexibility can then be analyzed in detail by comparing the purified wild type with the mutant proteins. In contrast, homologous proteins from mesophiles and thermophiles typically differ in many amino acid positions because of neutral genetic drift. As a consequence, it is more straightforward to deduce stabilization

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principles from an analysis of directed evolution experiments than from the comparison of natural proteins [56,57]. Various approaches of directed evolution have provided insights into the relationship between catalytic activity and thermal stability of enzymes. In one line of experiments, the growth of the extreme thermophilic bacterium T. thermophilus at elevated temperatures was coupled to the generation of stabilized variants of the homodimeric isopropylmalate dehydrogenase (LeuB) from Bacillus subtilis, which is a thermolabile enzyme involved in leucine biosynthesis. The LeuB gene was integrated into the T. thermophilus chromosome, coupling its growth at elevated temperatures (up to 70°C) on selective medium (without leucine) to the spontaneous acquisition of stabilizing mutations [58]. The best LeuB variant isolated by this approach was then further optimized by a combination of directed evolution and rational design, yielding an enzyme with nine amino acid exchanges that was stabilized by 23° compared with the wild-type LeuB and displayed almost unaltered catalytic activity [59]. In a similar approach, the thermostability of the homodimeric kanamycin nucleotidyltransferase (KNT) from Staphylococcus aureus was improved by random mutagensis, DNA shuffling and selection in T. thermophilus on agar plates containing kanamycin [60]. A stepwise increase of the selection pressure yielded a high-temperature version of the enzyme (HTK) containing 19 mutations. Its thermal denaturation, temperature was increased by 20°. Molecular dynamic simulations at 72°C and the kinetics of chemical cysteine modification at 37°C indicated that the flexibility within the two domains of each subunit is decreased in HTK, although its catalytic activity is unchanged compared with the wild-type KNT [60,61]. The selection for α-galactosidases (AraB1) from B. stearothermophilus that are active in T. thermophilus at 67°C resulted in the isolation of two enzyme variants [62]. Although their resistance to thermal inactivation was only marginally improved compared with the wild-type enzyme, their temperature of optimum catalytic efficiency was increased by 15° and 10°, respectively. Probably, their substrate affinity at elevated temperatures is much higher than that of the wild-type enzyme. The examples of LeuB, KNT (HTK), and AraB1 illustrate that selection for growth in a thermophilic host will lead to variants with improved in vivo activities at elevated temperatures. However, only in vitro measurements can reveal to which extent these improved activities are caused by an increased intrinsic stability and/or catalytic activity of a particular enzyme. A multi-step approach was taken to stabilize a thermolabile homo-tetrameric lactate oxidase [63]. In a first round of random mutagenesis and screening at elevated temperatures, six independent stabilizing amino acid exchanges were identified. These exchanges were then combined in a random manner, yielding a multiple mutant library from which variants with up to 5000-fold increased resistance against thermal inactivation were isolated. As the most stable variants, however, showed a drastically decreased catalytic activity (not tuned mutants in Figure 3.2), the six positions were individually subjected to saturation random mutagenesis. Those variants with the highest activities were selected from the stabilized members of each library. They were combined in a multiple mutant library. This approach allowed the isolation of variants whose gain in stability was accompanied by only a slight decrease in catalytic activity (fine-tuned mutants in Figure 3.2). An impressive example for the application of directed evolution and subsequent structural and computational analysis of the generated improved variants is p-nitrobenzyl esterase from B. subtilis. This enzyme is a monomer of about 490 amino acid residues. It was stabilized by a combination of random mutagenesis and screening [64]. The unfolding temperature of the best variant 8G8, which contained 13 amino acid exchanges, was elevated by 18° compared with the wild-type enzyme. Tryptophan phosphorescence lifetime measurements monitoring local conformational fluctuations around the solvent-sequestered aromatic chromophores suggest that the stabilized variants are less flexible than the wild-type enzyme [65]. Interestingly, the rigidifying mutations are remote from the tryptophan chromophores (especially tryptophan 102) and from two loop regions (residues 66–74 and 414–420), which are too mobile to be resolved in the wild-type enzyme but become visible in the x-ray structure of 8G8 [66]. These results suggest that the amino acid exchanges present in the mutants cause changes in long-range cooperative interactions within the protein. In molecular

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FIGURE 3.2 Fine-tuning allowed to isolate lactate oxidase mutants with increased thermal stability and only moderately decreased catalytic activity. The figure shows a plot of the time necessary to inactivate half of the enzyme molecules at 75°C versus the specific activity of multiple mutants. These were selected from libraries which were generated with either fine-tuned or not tuned mutations. (Reprinted from Hamamatsu, N. et al., Protein Eng. Des. Sel. 19, 483, 2006. With permission from Oxford University Press.)

dynamics (MD) simulations the time-averaged structure of 8G8 remained closer to its crystal structure than the wild-type enzyme while it experiences greater fluctuations about this average, especially for the residues around the mutations [67]. In agreement with this observation, the radius of gyration of 8G8 showed a small but statistically significant increase, indicating more extensive conformational “breathing” motions occurring in a concerted manner in a given protein region. The MD simulations suggested that the flexibility of 8G8 is decreased around tryptophan 102 and in the loop region comprising residues 414–420. This result supports the phosphorescence lifetime measurements and x-ray structure analysis cited before. Interestingly, the residues of the catalytic triad showed unchanged dynamics, although the enzymatic activity of 8G8 was increased threefold compared with the wild-type enzyme at 30°C. These results show that the term “flexibility” in the context of protein dynamics has to be specified in terms of location in the protein, time scales, and the amplitudes of the motions. Small amplitude local fluctuations, which were detected in the simulations as deviations from the average state, may contribute to thermostability, because they can increase the conformational entropy of the native state [68]. In contrast, large fluctuations from the lowest energy state as structurally manifested in the x-ray structure could initiate unfolding and will be destabilizing. A similar approach as with p-nitrobenzyl esterase from B. subtilis was followed to stabilize the monomeric and labile subtilisin from a psychrophilic Bacillus species [69,70]. Because the half-life of this subtilisin is Ca2+-dependent, mutant gene libraries were screened in the presence of low concentrations of Ca2+, to stabilize the enzyme by increasing its affinity for the bivalent ion. Indeed, the stability difference between the best isolated variant and the wild-type enzyme was almost 10 times greater at 1 mM Ca2+ than at 10 mM Ca2+, due to an almost 100-fold difference in Ca2+-affinity. Moreover, the turnover number of the evolved variant far surpassed that of wild-type subtilisin in the temperature range of 10°C to 60°C. This experiment underlines that the conformational stability of an enzyme can be modified in various ways, some of which do not influence catalytic activity. In analogy to improving the stability of a mesophilic enzyme by selection or screening under thermophilic conditions, the activity of a thermophilic enzyme at moderate temperatures can be increased by selection in a mesophilic host. Using this approach, the low catalytic activities of

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thermostable oligomeric sugar-degrading enzymes and an ornithine carbamoyltransferase were increased by random mutagenesis, followed by screening or selection in E. coli or yeast [71–73]. Along the same lines, the poor catalytic activity of a ribonuclease HI variant from T. thermophilus at 30°C was enhanced by random mutagenesis, followed by the selection for second-site revertants in E. coli. The introduction of the identified activating amino acid exchanges into the wild-type enzyme resulted in a 40-fold increase of catalytic efficiency accompanied by only a modest decrease in thermal stability [74]. In an inverse approach, the thermal stability of E. coli ribonuclease HI was improved by 20° without serious loss of enzymatic activity [75]. The dynamics of the wild-type enzyme and the stabilized mutant protein were compared by H/D exchange studies combined with 15N relaxation measurements. Although the stabilizing substitutions did not cause global changes in the backbone dynamics on fast (picoseconds to nanoseconds) or slow (microseconds to milliseconds) time scales, local effects on internal motions could be observed for residues close to the mutations [76]. The weak catalytic activity of indole glycerol phosphate (IGP) synthase from the hyperthermophile Sulfolobus solfataricus (sIGPS) was increased by a combination of random mutagenesis and selection in an auxotrophic E. coli strain on medium containing low concentrations of tryptophan [77]. Several of the isolated variants which had acquired mutations in the active site region were purified. Their activities, flexibilities, and thermostabilities were investigated. Compared with the wild-type enzyme, these variants had slightly increased (two- to fourfold) turnover numbers (kcat) but drastically increased (8–200-fold) Michaelis constants (KM). This led to a 3 to 60-fold decrease in catalytic efficiency (kcat/KM). These results revealed that the turnover number was the selected trait and that the intracellular substrate concentrations were saturated for all variants. Stopped-flow experiments performed at 25°C showed that the overall turnover in the wild-type enzyme was limited by the release of the product IGP from the active site. In the activated variants, product dissociation has become much faster, making the actual catalytic process of the reaction rate determining. Limited proteolysis experiments indicated that an extensive surface loop is more exposed in the activated

FIGURE 3.3 Temperature dependence of catalytic activity (upper panel) and of denaturation (as recorded by fluorescence emission; lower panel) for α-amylases from a psychrophile (o), a mesophile (Δ), and a thermophile (䊐). Note that the temperature of maximum activity of the psychrophilic enzyme is lower than its unfolding temperature, suggesting that the active site is less stable than the protein structure. (Reprinted from D’Amico, S. et al., J. Biol. Chem. 278, 7891, 2003. With permission from the American Society for Biochemistry and Molecular Biology.)

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sIGPS variants, suggesting that long-range perturbations caused by the exchanges loosen the phosphate-binding site and thereby support the release of IGP [77]. A comparable increase in flexibility might be caused by the high physiological temperatures of S. solfataricus, which would facilitate product release from wild-type sIGPS under physiological conditions. Some of the variants showed an increased rate of heat inactivation, whereas in others the improved activity was not traded off by a decreased thermal stability. Using a similar approach, the turnover number of LeuB from T. thermophilus was increased by random mutagenesis and selection in E. coli at 30°C [78–80]. The introduced amino acid exchanges caused enhanced reaction rates at low temperatures, either due to the destabilization of the Michaelis complex [78] or the stabilization of the transition state [80]. Although the variants did not show a decreased thermal stability, their catalytic activity at high temperatures was lower than that of the wild-type enzyme. These observations show that the mutations caused a decreased dependence of catalytic activity on temperature, corresponding to a reduced activation enthalpy ΔH‡ [79]. Interestingly, naturally evolved cold-adapted enzymes from psychrophilic microorganisms also display relatively small activation enthalpies which minimizes the negative effect of low temperature on their turnover numbers [81]. The higher reaction rate (kcat values) of psychrophilic compared with mesophilic enzymes corresponds to a lower Gibbs activation free energy, which appears to be determined by the lower ΔH‡ of the cold-adapted variants. However, the activation entropy ΔS‡ of psychrophilic enzymes is also decreased, resulting in an enthalpy–entropy compensation that limits their reaction rate (ΔG‡ = ΔH‡ – T ΔS‡). From these considerations, it has been concluded that only few enthalpy-based interactions have to be broken during the reaction (low ΔH‡) and that the highly flexible active site is rigidified upon formation of the transition state (low ΔS‡) [82]. In line with this concept of “localized flexibility” [83], which may lead to a localized destabilization, the active site of a psychrophilic α-amylase appears to “denature” at lower temperatures than the protein structure [13] (Figure 3.3).

CONCLUSIONS The case of psychrophiles illustrates the complex effect of dynamics on the conformational stability and catalytic activity of enzymes which also complicates the comparison of biocatalysts from mesophiles and thermophiles. A major problem for the generalization and rationalization of present experimental data is that numerous studies of protein dynamics performed in the past years made use of different techniques. These techniques either monitored global or local effects of different amplitudes and time constants. Nevertheless, it can be concluded that the rather general and global concept of “corresponding states” [9] remains to be useful for the comparative analysis of many mesophilic–thermophilic enzyme pairs. Recent experimental evidence, however, shows that this concept needs to be complemented and refined. Seemingly, nature has developed a variety of different and subtle strategies to accomplish the goal of fine-tuning the interplay between stability, flexibility, and catalytic activity. The cited directed evolution experiments and related recent investigations [84,85] have shown that an increase in stability is not necessarily traded off by a loss in catalytic activity, or vice versa. These observations suggest that the rarely found combination of high thermostability and high catalytic activity at low temperature in native enzymes is not based on physical chemistry but due to the lack of evolutionary constraints [70]. Enzymes in mesophilic organisms do not experience selective pressure toward higher stabilities at elevated temperature. Similarly, the activities of hyperthermophilic enzymes at low temperatures do normally not need to be high. Otherwise, hyperthermophilic enzymes would be much more efficient catalysts than their mesophilic homologs at the corresponding physiological temperatures due to the acceleration of chemical reactions with increasing temperature. An example are two (βα)8-barrel enzymes from the hyperthermophile T. maritima, which are extremely active at their physiological temperatures to outrun the spontaneous degradation of their thermolabile substrates [86,87].

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REFERENCES 1. Meiering, E.M., Serrano, L., and Fersht, A.R., Effect of active site residues in barnase on activity and stability, J. Mol. Biol. 225, 585, 1992. 2. Shoichet, B.K. et al., A relationship between protein stability and protein function, Proc. Natl. Acad. Sci. USA 92, 452, 1995. 3. Schreiber, G., Buckle, A.M., and Fersht, A.R., Stability and function: two constraints in the evolution of barstar and other proteins, Structure (London, England) 2, 945, 1994. 4. Kidokoro, S. et al., Remarkable activity enhancement of thermolysin mutants, FEBS Lett. 367, 73, 1995. 5. Wang, X., Minasov, G., and Shoichet, B.K., Evolution of an antibiotic resistance enzyme constrained by stability and activity trade-offs, J. Mol. Biol. 320, 85, 2002. 6. Sterner, R. and Liebl, W., Thermophilic adaptation of proteins, Crit. Rev. Biochem. Mol. Biol. 36, 39, 2001. 7. Jaenicke, R. and Böhm, G., The stability of proteins in extreme environments, Curr. Opin. Struct. Biol. 8, 738, 1998. 8. Benkovic, S.J. and Hammes-Schiffer, S., A perspective on enzyme catalysis, Science 301, 1196, 2003. 9. Jaenicke, R., Protein stability and molecular adaptation to extreme conditions, Eur. J. Biochem. 202, 715, 1991. 10. Jaenicke, R., Do ultrastable proteins from hyperthermophiles have high or low conformational rigidity? Proc. Natl. Acad. Sci. USA 97, 2962, 2000. 11. Varley, P.G. and Pain, R.H., Relation between stability, dynamics and enzyme activity in 3-phosphoglycerate kinases from yeast and Thermus thermophilus, J. Mol. Biol. 220, 531, 1991. 12. Balog, E., Laberge, M., and Fidy, J., The influence of interdomain interactions on the intradomain motions in yeast phosphoglycerate kinase. A molecular dynamics study, Biophys. J. 92, 1709, 2006. 13. D’Amico, S. et al., Activity–stability relationships in extremophilic enzymes, J. Biol. Chem. 278, 7891, 2003. 14. Hvidt, A. and Nielsen, S.O., Hydrogen exchange in proteins, Adv. Protein Chem. 21, 287, 1966. 15. Hvidt, A. and Wallevik, K., Conformational changes in human serum albumin as revealed by hydrogen– deuterium exchange studies, J. Biol. Chem. 247, 1530, 1971. 16. Zavodszky, P., Johansen, J.T., and Hvidt, A., Hydrogen-exchange study of the conformational stability of human carbonic-anhydrase B and its metallocomplexes, Eur. J. Biochem. 56, 67, 1975. 17. Zavodszky, P. et al., Adjustment of conformational flexibility is a key event in the thermal adaptation of proteins, Proc. Natl. Acad. Sci. USA 95, 7406, 1998. 18. Svingor, A. et al., A better enzyme to cope with cold. Comparative flexibility studies on psychrotrophic, mesophilic, and thermophilic IPMDHs, J. Biol. Chem. 276, 28121, 2001. 19. Fitter, J. and Haber-Pohlmeier, S., Structural stability and unfolding properties of thermostable bacterial α-amylases: a comparative study of homologous enzymes, Biochemistry 43, 9589, 2004. 20. Fitter, J. et al., Activity and stability of a thermostable α-amylase compared to its mesophilic homologue: mechanisms of thermal adaptation, Biochemistry 40, 10723, 2001. 21. Cavanagh, J., Fairbrother, W.J., Palmer III, A.G., and Skelton, N.J., Protein NMR Spectroscopy, Academic Press, San Diego, 1996. 22. Kremer, W. et al., Solution NMR structure of the cold-shock protein from the hyperthermophilic bacterium Thermotoga maritima, Eur. J. Biochem. 268, 2527, 2001. 23. Jung, A. et al., High-temperature solution NMR structure of TmCsp, Protein Sci. 13, 342, 2004. 24. Hollien, J. and Marqusee, S., Structural distribution of stability in a thermophilic enzyme, Proc. Natl. Acad. Sci. USA 96, 13674, 1999. 25. Hwang, T.-L., Mori, S., Shaka, A.J., and van Zijl, P.C.M., Application of phase-modulated CLEAN chemical EXchange spectroscopy (CLEANEX-PM) to detect water-protein proton exchange and intermolecular NOEs, J. Am. Chem. Soc. 119, 6203, 1997. 26. Hwang, T.L., van Zijl, P.C., and Mori, S., Accurate quantitation of water-amide proton exchange rates using the phase-modulated CLEAN chemical EXchange (CLEANEX-PM) approach with a fast-HSQC (FHSQC) detection scheme, J. Biomol. NMR 11, 221, 1998.

9214_C003.indd 35

11/5/2007 7:06:55 PM

36

Thermophiles: Biology and Technology at High Temperatures

27. Hernandez, G. and LeMaster, D.M., Reduced temperature dependence of collective conformational opening in a hyperthermophile rubredoxin, Biochemistry 40, 14384, 2001. 28. LeMaster, D.M. et al., Enhanced thermal stability achieved without increased conformational rigidity at physiological temperatures: spatial propagation of differential flexibility in rubredoxin hybrids, Proteins 61, 608, 2005. 29. LeMaster, D.M. and Hernandez, G., Additivity of differential conformational dynamics in hyperthermophile/mesophile rubredoxin chimeras as monitored by hydrogen exchange, Chembiochem 7, 1886, 2006. 30. Eisenmesser, E.Z. et al., Enzyme dynamics during catalysis, Science 295, 1520, 2002. 31. Wolf-Watz, M. et al., Linkage between dynamics and catalysis in a thermophilic–mesophilic enzyme pair, Nat. Struct. Mol. Biol. 11, 945, 2004. 32. Eisenmesser, E.Z. et al., Intrinsic dynamics of an enzyme underlies catalysis, Nature 438, 117, 2005. 33. Bae, E. and Phillips, G.N. Jr., Roles of static and dynamic domains in stability and catalysis of adenylate kinase, Proc. Natl. Acad. Sci. USA 103, 2132, 2006. 34. Cha, Y., Murray, C.J., and Klinman, J.P., Hydrogen tunneling in enzyme reactions, Science 243, 1325, 1989. 35. Nagel, Z.D. and Klinman, J.P., Tunneling and dynamics in enzymatic hydride transfer, Chem. Rev. 106, 3095, 2006. 36. Kohen, A. et al., Enzyme dynamics and hydrogen tunnelling in a thermophilic alcohol dehydrogenase, Nature 399, 496, 1999. 37. Kohen, A. and Klinman, J.P., Protein flexibility correlates with degree of hydrogen tunneling in thermophilic and mesophilic alcohol dehydrogenases, J. Am. Chem. Soc. 122, 10738, 2000. 38. Vieille, C. and Zeikus, G.J., Hyperthermophilic enzymes: sources, uses, and molecular mechanisms for thermostability, Microbiol. Mol. Biol. Rev. 65, 1, 2001. 39. van den Burg, B. and Eijsink, V.G., Selection of mutations for increased protein stability, Curr. Opin. Biotechnol. 13, 333, 2002. 40. Watanabe, K. et al., Designing thermostable proteins: ancestral mutants of 3-isopropylmalate dehydrogenase designed by using a phylogenetic tree, J. Mol. Biol. 355, 664, 2006. 41. Jaenicke, R. and Sterner, R., Protein design at the crossroads of biotechnology, chemistry, theory, and evolution, Angewandte Chemie Int. ed. 42, 140, 2003. 42. Razvi, A. and Scholtz, J.M., Lessons in stability from thermophilic proteins, Protein Sci. 15, 1569, 2006. 43. Sanchez-Ruiz, J.M. and Makhatadze, G.I., To charge or not to charge? Trends Biotechnol. 19, 132, 2001. 44. Perl, D. and Schmid, F.X., Some like it hot: the molecular determinants of protein thermostability, Chembiochem 3, 39, 2002. 45. Pace, C.N., Single surface stabilizer, Nat. Struct. Biol. 7, 345, 2000. 46. Penning, T.M. and Jez, J.M., Enzyme redesign, Chem. Rev. 101, 3027, 2001. 47. Woycechowsky, K.J., Vamvaca, K., and Hilvert, D., Novel enzymes through design and evolution, Adv. Enzymol. Relat. Areas. Mol. Biol. 75, 241, 2007. 48. Williams, J.C. et al., Structural and mutagenesis studies of leishmania triosephosphate isomerase: a point mutation can convert a mesophilic enzyme into a superstable enzyme without losing catalytic power, Protein Eng. 12, 243, 1999. 49. Farinas, E.T., Bulter, T., and Arnold, F.H., Directed enzyme evolution, Curr. Opin. Biotechnol. 12, 545, 2001. 50. Reetz, M.T., Combinatorial and evolution-based methods in the creation of enantioselective catalysts, Angew. Chem. Int. Ed. 40, 284, 2001. 51. Schmidt-Dannert, C., Directed evolution of single proteins, metabolic pathways, and viruses, Biochemistry 40, 13125, 2001. 52. Turner, N.J., Directed evolution of enzymes for applied biocatalysis, Trends Biotechnol. 21, 474, 2003. 53. Yano, T., Oue, S., and Kagamiyama, H., Directed evolution of an aspartate aminotransferase with new substrate specificities, Proc. Natl. Acad. Sci. USA 95, 5511, 1998. 54. Oue, S. et al., Redesigning the substrate specificity of an enzyme by cumulative effects of the mutations of non-active site residues, J. Biol. Chem. 274, 2344, 1999.

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Analysis of Mesophilic–Thermophilic Enzyme Pairs and Protein Engineering Studies

37

55. Liebeton, K. et al., Directed evolution of an enantioselective lipase, Chem. Biol. 7, 709, 2000. 56. Wintrode, P.L. and Arnold, F.H., Temperature adaptation of enzymes: lessons from laboratory evolution, Adv. Protein Chem. 55, 161, 2000. 57. Arnold, F.H. et al., How enzymes adapt: lessons from directed evolution, Trends Biochem. Sci. 26, 100, 2001. 58. Akanuma, S. et al., Serial increase in the thermal stability of 3-isopropylmalate dehydrogenase from Bacillus subtilis by experimental evolution, Protein Sci. 7, 698, 1998. 59. Akanuma, S. et al., Further improvement of the thermal stability of a partially stabilized Bacillus subtilis 3-isopropylmalate dehydrogenase variant by random and site-directed mutagenesis, Eur. J. Biochem. 260, 499, 1999. 60. Hoseki, J. et al., Directed evolution of thermostable kanamycin-resistance gene: a convenient selection marker for Thermus thermophilus, J. Biochem. (Tokyo) 126, 951, 1999. 61. Hoseki, J. et al., Increased rigidity of domain structures enhances the stability of a mutant enzyme created by directed evolution, Biochemistry 42, 14469, 2003. 62. Fridjonsson, O., Watzlawick, H., and Mattes, R., Thermoadaptation of α-galactosidase AgaB1 in Thermus thermophilus, J. Bacteriol. 184, 3385, 2002. 63. Hamamatsu, N. et al., Directed evolution by accumulating tailored mutations: thermostabilization of lactate oxidase with less trade-off with catalytic activity, Protein Eng. Des. Sel. 19, 483, 2006. 64. Giver, L. et al., Directed evolution of a thermostable esterase, Proc. Natl. Acad. Sci. USA 95, 12809, 1998. 65. Gershenson, A. et al., Tryptophan phosphorescence study of enzyme flexibility and unfolding in laboratory-evolved thermostable esterases, Biochemistry 39, 4658, 2000. 66. Spiller, B. et al., A structural view of evolutionary divergence, Proc. Natl. Acad. Sci. USA 96, 12305, 1999. 67. Wintrode, P.L. et al., Protein dynamics in a family of laboratory evolved thermophilic enzymes, J. Mol. Biol. 327, 745, 2003. 68. Stone, M.J., NMR relaxation studies of the role of conformational entropy in protein stability and ligand binding, Acc. Chem. Res. 34, 379, 2001. 69. Miyazaki, K. and Arnold, F.H., Exploring nonnatural evolutionary pathways by saturation mutagenesis: rapid improvement of protein function, J. Mol. Evol. 49, 716, 1999. 70. Miyazaki, K. et al., Directed evolution study of temperature adaptation in a psychrophilic enzyme, J. Mol. Biol. 297, 1015, 2000. 71. Lebbink, J.H. et al., Improving low-temperature catalysis in the hyperthermostable Pyrococcus furiosus beta-glucosidase CelB by directed evolution, Biochemistry 39, 3656, 2000. 72. Roovers, M. et al., Experimental evolution of enzyme temperature activity profile: selection in vivo and characterization of low-temperature-adapted mutants of Pyrococcus furiosus ornithine carbamoyltransferase, J. Bacteriol. 183, 1101, 2001. 73. Lönn, A. et al., Cold adaptation of xylose isomerase from Thermus thermophilus through random PCR mutagenesis. Gene cloning and protein characterization, Eur. J. Biochem. 269, 157, 2002. 74. Hirano, N. et al., Enhancement of the enzymatic activity of ribonuclease HI from Thermus thermophilus HB8 with a suppressor mutation method, Biochemistry 39, 13285, 2000. 75. Akasako, A. et al., High resistance of Escherichia coli ribonuclease HI variant with quintuple thermostabilizing mutations to thermal denaturation, acid denaturation, and proteolytic degradation, Biochemistry 34, 8115, 1995. 76. Yamasaki, K., Akasako-Furukawa, A., and Kanaya, S., Structural stability and internal motions of Escherichia coli ribonuclease HI: 15N relaxation and hydrogen–deuterium exchange analyses, J. Mol. Biol. 277, 707, 1998. 77. Merz, A. et al., Improving the catalytic activity of a thermophilic enzyme at low temperatures, Biochemistry 39, 880, 2000. 78. Suzuki, T. et al., Adaptation of a thermophilic enzyme, 3-isopropylmalate dehydrogenase, to low temperatures, Protein Eng. 14, 85, 2001. 79. Yasugi, M. et al., Analysis of the effect of accumulation of amino acid replacements on activity of 3-isopropylmalate dehydrogenase from Thermus thermophilus, Protein Eng. 14, 601, 2001. 80. Yasugi, M. et al., Cold adaptation of the thermophilic enzyme 3-isopropylmalate dehydrogenase, J. Biochem. (Tokyo) 129, 477, 2001.

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81. D’Amico, S. et al., Molecular basis of cold adaptation, Phil. Trans. R. Soc. Lond. B 357, 917, 2002. 82. Lonhienne, T., Gerday, C., and Feller, G., Psychrophilic enzymes: revisiting the thermodynamic parameters of activation may explain local flexibility, Biochim. Biophys. Acta 1543, 1, 2000. 83. Fields, P.A. and Somero, G.N., Hot spots in cold adaptation: localized increases in conformational flexibility in lactate dehydrogenase A4 orthologs of Antarctic notothenioid fishes, Proc. Natl. Acad. Sci. USA 95, 11476, 1998. 84. Hecky, J. and Müller, K.M., Structural perturbation and compensation by directed evolution at physiological temperature leads to thermostabilization of β-lactamase, Biochemistry 44, 12640, 2005. 85. Chiu, W.C. et al., Structure–stability–activity relationship in covalently cross-linked N-carbamoyl d-amino acid amidohydrolase and N-acylamino acid racemase, J. Mol. Biol. 359, 741, 2006. 86. Sterner, R. et al., Phosphoribosyl anthranilate isomerase from Thermotoga maritima is an extremely stable and active homodimer, Protein Sci. 5, 2000, 1996. 87. Henn-Sax, M. et al., Two (βα)8-barrel enzymes of histidine and tryptophan biosynthesis have similar reaction mechanisms and common strategies for protecting their labile substrates, Biochemistry 41, 12032, 2002.

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4

Membranes and Transport Proteins of Thermophilic Microorganisms Sonja Verena Albers and Arnold J.M. Driessen

CONTENTS Introduction .................................................................................................................................. Lipids in Bacterial and Archaeal Membranes .............................................................................. General Principles of Transport: Mechanisms and Transporter Classes ...................................... Distribution of Transport Systems in Thermophiles .................................................................... Channels ........................................................................................................................... Secondary Transporters .................................................................................................... ABC Transporters ............................................................................................................. Substrate-Binding Protein ...................................................................................... ATP-Binding Protein .............................................................................................. Concluding Remarks .................................................................................................................... References ....................................................................................................................................

39 40 41 42 44 45 46 47 49 50 51

Thermophilic microorganisms have developed specific adaptations to their cell surface to deal with the high environmental temperatures. The cytoplasmic membrane forms a barrier that separates the inside of the cell from the outside, allowing a unique ionic and macromolecule composition of the cytoplasm. Moreover, the cytoplasmic membrane fulfils a key role in energy transduction which depends on low endogenous ion- and proton permeability of the membrane. Various strategies are employed to reduce the ion permeability of the membranes to sustain viability, including the use of unique lipids that provide sufficient fluidity to the membrane to support membrane protein functioning while maintaining the barrier function of the membrane for small ions. Transport of solutes across the membrane of thermophiles involves primary ATP-driven transport systems or by proton or sodium motive force driven secondary transport systems. Unlike most bacteria, hyperthermophilic bacteria and archaea prefer primary uptake systems some of which exhibit a very high affinity for their substrates. This allows the organisms to efficiently scavenge solutes in nutrient-poor environments.

INTRODUCTION Since the discovery and cultivation of thermophilic microorganisms in the seventies of the last century, the number of identified extremophilic species has increased steadily. As soon as the first thermophilic organisms were characterized, speculation arose as to whether these microorganisms might be closely related to the organisms at the origin of life, as the early earth is envisioned as a hot and hostile environment similar to the conditions that thermophiles face nowadays. Not only 39

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from an evolutionary point of view this group of organisms has received a great amount of interest, but also fundamental research has resulted in novel insights in the mechanisms of adaptations that allow these organisms to grow at extreme temperatures. This is reflected in an altered protein and nucleic acid composition (see Chapter 3) but also in a different composition and organization of the cell envelope. Moreover, many thermostable enzymes have revolutionized methods commonly used in research such as polymerase chain reaction (PCR) or different production processes in biotechnology that can now be performed at temperatures that would destroy proteins from mesophiles. Thermophilic organisms have been able to adapt their proteins and nucleic acids composition in such a way that they function optimally at elevated temperatures. However, one major issue thermophiles also have to face is to sustain the integrity of their cytoplasmic membrane in order to perform energy transducing functions. The cytoplasmic membrane encloses the cell interior, the cytosol, and it functions as the main barrier to the outside medium that enables the cell to control the ionic and macromolecule composition of its cytoplasm. Moreover, the cytoplasmic membrane is crucial for the generation of metabolic energy by energy transduction. In this process, the energy of an electrochemical ion gradient across the membrane is transformed into other forms of energy or vice versa. Metabolic energy can also be obtained in the form of ATP and ADP by substrate level phosphorylation processes. Both metabolic energy-generating processes are closely linked and together they determine the energy status of the cell. The energy transducing systems are located in the cytoplasmic membrane. Specific membrane protein complexes translocate protons or sodium ions across the membrane into the external medium and this activity results in the generation of electrochemical gradients of protons or sodium ions. When protons are extruded, the resulting electrochemical gradient exerts a proton motive force (PMF). In analogy with the PMF, sodium ion pumps can generate a sodium motive force (SMF). The PMF or SMF can be utilized to drive energy-requiring processes such as ATP synthesis from ADP and phosphate, transport of specific solutes across the membrane, flagellar rotation, and maintenance of the intracellular pH and turgor. Obviously, this type of energy transduction can only operate if the transmembrane gradient of H+ c.q. Na+ can be maintained. A prerequisite for this maintenance is that the cytoplasmic membrane exhibits only a limited permeability for these ions. The intrinsic hydrophobic nature of the membrane ensures a low permeability towards protons, ions, and other polar molecules. Membranes typically consist of a bilayer of lipid molecules and of proteins. In nature, an enormous diversity of lipids is found. The membrane lipids have polar headgroups that stick into the water phase and long hydrophobic hydrocarbon chains that are oriented to the interior of the membrane. At the growth temperature of a given organism, the membranes are in a liquid crystalline state [1]. The structure of the membrane is mainly held together by noncovalent bonds such as van der Waals bonds and electric interactions. The barrier function of the cell membrane is critical for the functioning of the cell, as the membrane has to control the concentration of molecules and ions inside the cell. Most solutes can cross the membrane via specific transport proteins. The permeability of membranes for small solutes and ions is restricted due to the high energy that is required for the transfer of a hydrophobic solute or ion from the aqueous phase into the apolar interior of the membrane. Organisms control the fluidity and permeability of their cytoplasmic membrane [1]. The lipid layer forms a matrix for membrane proteins in which they need to function, and therefore it needs to be in the liquid-crystalline state while the more rigid crystalline state is incompatible with membrane protein functioning. The rate at which protons leak inward into the cells is determined by the intrinsic proton permeability and the magnitude of the PMF across the membrane. A proper balance between proton permeability and the rate of outward proton pumping is needed to sustain a viable PMF.

LIPIDS IN BACTERIAL AND ARCHAEAL MEMBRANES A detailed overview of the characteristics and features of lipids of thermophilic bacteria and archaea is presented in Chapter 6. The main difference between bacterial and archaeal lipid types is the

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41

composition and the linkage of the hydrophobic acyl chain to the polar head group. Whereas in bacterial lipids two fatty acid acyl chains are linked to glycerol via ester bonds, in archaeal lipids phytanyl chains are linked via ether bonds to glycerol or other alcohols like nonitol. The bacterial phospholipids are organized in a bilayer in which the carbon chains are directed towards the interior of the membrane. Halobacteria and most archaea growing under moderate conditions contain lipids which consist of a C20 diether lipid core [2–4]. These lipids form bilayers in a similar way as the esterlipids. However, membrane spanning (bolaform amphiphilic) tetraether lipids are found in extreme thermophiles and acidophiles [5]. These lipids have C40 isoprenoid acyl chains with two polar headgroups and can span the entire membrane to form a monolayer with the thickness of a regular lipid bilayer [6]. Freeze-fracturing of such membranes reveals that cleavage between the two leaflets of the membrane does not occur, which means that the water-facing sides of the membrane are connected and cannot be separated [6]. As mentioned earlier, one of the most essential feature of a biological membrane is that it needs to be in a liquid crystalline phase in order to function properly [1]. The phase transition temperature from the liquid crystalline to the liquid state of ether lipids is much lower than of ester lipids. Differential scanning calorimetry studies have shown that the phase transition point of the polar tetraether phytanyl lipids from Thermoplasma acidophilum is between −20°C and −15°C [7]. This is mostly due to the presence of the branched phytanyl groups that present a high ordering of the lipid acyl chains. In contrast, the phase transition point of ester lipids highly depends on the acyl chain length, the number of double bonds and the degree of saturation of the fatty acid acyl chain [8]. For instance, the ester lipids dipalmitoyl phosphatidyl choline and distearoyl phosphatidyl choline showed phase transition points of 41.5°C and 55°C, respectively. Therefore, bacteria have to adjust the composition of their membrane in response to temperature changes in the environment, whereas many archaea can maintain their membranes in a liquid crystalline phase over a wide range of temperatures of 0°C to 100°C. Hyperthermophilic archaea have also been shown to adapt their lipid composition upon heat stress: Sulfolobus and Thermoplasma increase the cyclization of the C40 isoprenoid chains to permit a more dense packing of the lipids. This results in a more restricted motion of the lipids and prevents that the membrane becomes too fluid [8]. Above 90% of the membrane lipids of these two archaea are formed by tetraether lipids, whereas in the euryarchaeote Methanocaldococcus jannaschii increasing temperatures induce the change from diether lipids to the more thermostable tetraether lipids [9]. Also in this case, the cyclization of the chains results in a decreased motion of the lipids and therefore contributes to acceptable membrane fluidity at elevated growth temperature.

GENERAL PRINCIPLES OF TRANSPORT: MECHANISMS AND TRANSPORTER CLASSES Depending on their molecular architecture and energy requirements solute transporters can be classified in five different groups (Figure 4.1): channels, primary transporters, secondary transporters, binding-protein-dependent transporters, and group-translocation transporters. The important characteristics of these five classes can be described briefly as follows. (i) Channels—well-known examples are the mechanosensitive (MS) channels, which can exhibit either large or small conductance in response to the turgor pressure. (ii) Primary transporters, which use light or chemical energy such as ATP to drive substrate translocation. Well-studied examples are ion-translocating respiratory chains, the light-driven proton pump bacteriorhodopsin, various types of ion-translocating ATPases and the ATP-binding cassette (ABC) transporters. ABC transporters have a typical modular domain structure which usually comprises two integral membrane proteins that form the permease domain and two cytoplasm-located ATPases which drive the transport of the substrate by the hydrolysis of ATP. In contrast to prokaryotes, ABC transporters in eukaryotes usually consist of a single polypeptide that encompasses all four domains. However, in bacteria and archaea, the membrane and ATPase domains may either exist as homo- or heterodimers, or even as separately

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Thermophiles: Biology and Technology at High Temperatures Channel

(a)

Secondary transporters H+ (Na+)

H+ (Na+)

H+

Out In Uniporter

Symporter

ABC transporter

Antiporter Binding-protein dependent transporter

PTS system

(b) Out EIIC

In

EIIB

ATP ADP + Pi

ATP ADP + Pi

~P

EIIA HPr EI PEP

Substrate efflux transporter

Binding-protein dependent transporter

FIGURE 4.1 Classes of transporters. Schematic drawings of: (a) channels and different modes of secondary transporters, and (b) ABC transporters and the PTS system. The membrane integral subunits of the transporters are shown in gray whereas the cytoplasmic or extracellular subunits are depicted in white. Open circle, transported substrate. The names in the different subunits of the PTS system refer to the mannitol transporter of E. coli, but PTS are generally absent from hyperthermophilic bacteria and archaea.

fused membrane or ATPase domains. The ABC export systems are often homo- or heterodimers of a membrane domain fused with an ATP domain. Bacterial and archaeal ABC uptake systems comprise a fifth domain which is an extracellular substrate-binding-protein that binds the substrate at the outside of the cell and delivers it to the permease domain. (iii) Secondary transporters which make use of the electrochemical gradient of either protons, sodium ions, or substrates to drive transport of substrates across the membrane. These are subdivided into three classes: (a) uniporters, which transport solutes without a coupling ion; (b) symporters, which translocate solutes together with a coupling ion, such as protons or sodium ions; and (c) antiporters, which transport two substrates/ions in opposite directions. Secondary transporters usually consist of one polypeptide chain typically with 12 transmembrane domains. (iv) Binding-protein-dependent secondary transporters which consist of a periplasmic-binding protein and a membrane domain. These systems use the PMF or SMF to drive uptake of solutes. Finally (v) group translocation systems, that is, the phosphoenolpyruvate (PEP) dependent phosphotransferase systems (PTS), which couple the transport of sugars to phosphorylation.

DISTRIBUTION OF TRANSPORT SYSTEMS IN THERMOPHILES The number of determined genome sequences of different microorganisms is increasing steadily. Paulsen and coworkers use these data to perform an analysis of the presence of different transport proteins classes (see TransportDB at http://www.membranetransport.org/). These data are summarized in Table 4.1 for the completely sequenced (hyper-)thermophiles. In general, PTS systems are not found in archaea, but they are also absent in the hyperthermophilic bacteria Thermotoga maritima and Aquifex aeolicus that deeply branch in the phylogenetic tree (Table 4.1). On the other hand, PTS are present in nearly all other bacteria, such as the moderate thermophile Streptococcus

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1.5 1.8 1.8

Bacteria Aquifex aeolicus VF5 Thermotoga maritima MSB8 Streptococcus thermophilus LMD-9‡

†

Data from http://www.membranetransport.org/ Including ABC and arsenate transporters, but not F1F0-ATPase. ‡Moderate thermophilic bacteria containing PTS.

*

2.18 1.67 1.66 1.89 1.75 0.5 1.77 1.91 1.8 2.22 1.55 2.23 2.99 2.81 1.56 2.09

Archaea Archaeoglobus fulgidus DSM4304 Aeropyrum pernix K1 Methanocaldococcus jannaschii DSM Methanopyrus kandleri AV19 Methanobacterium thermoautotrophicum ΔH Nanoarchaeum equitans Kin4-M Pyrococcus abyssi GE5 P. furiosus DSM3638 P. horikoshii OT3 Pyrobaculum aerophilum IM2 Picrophilus torridus DSM9790 Sulfolobus acidocaldarius DSM639 S. solfataricus P2 S. tokodaii strain7 Thermoplasma acidophilum DSM1728 Thermococcus kodakaraensis KOD1

Genome Size (Mb)

95 80 42

85 90 85 98 65 90 96 98 98 98 60 78 80 80 60 85 59 95 131

104 78 49 36 66 10 92 102 92 78 114 115 127 133 105 116

Optimum Growth Number of Temperature (ºC) Transporters

TABLE 4.1 Distribution of Transporter Classes in (Hyper-)Thermophiles*

18 56 50

33 41 14 10 18 2 29 39 33 44 22 28 37 31 25 42

ATP-Dependent Transporters†

6 4 7

6 5 7 5 6 3 6 5 4 4 4 6 7 4 4 5

Channels

0 0 18

0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0

PTS

29 32 47

56 27 26 19 30 4 52 51 54 25 84 76 81 90 71 61

Secondary Transporters

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thermophilus (Table 4.1). It thus seems that PTS-systems have evolved relatively late. The number of transporters present in the different archaeal and thermophilic bacterial genomes can be expressed as the number of transporters per megabase (Mb) of genomic DNA. The value for this parameter varies from 21.3 in Methanopyrus kandleri to 73 in Picrophilus torridus and Thermoplasma volcanicum, but in most archaea it is around 40. The two analyzed bacterial genomes of Thermotoga maritima and Thermus thermophilus both contain around 50 transporters per Mb genome. As there are no PTS systems present in archaea and thermophilic bacteria, only three classes of transporters remain to be considered, that is, ion channels, ATP-dependent transporters, and secondary transporters. The ion channels constitute the smallest class and amounts to only 5% to 8% of all transporters. Only M. kandleri and Nanoarchaeum equitans form an exception with a high incidence of ion channels, that is, 14% and 19%, respectively. Whereas N. equitans contains two small conductance MS ion channels, M. kandleri exhibits three candidates of voltage-gated ion channels. On average, more than 50% of the archaeal transporters belong to the class of secondary transporters, while the remainder, that is, about 37% are ATP-dependent transport systems. In thermophilic bacteria this relationship appears inverted, as ATP-dependent transport systems are most abundant (50% or higher) while the class of secondary transporters is present at around 40%. Only in the archaea Aeropyrum pernix and Pyrobaculum aerophilum are the ATP-dependent transport systems the predominant class. N. equitans contains only three ABC transport genes, of which only one has an identified membrane domain. The other two ATP-binding domains may have other functions for instance in the cytosol, as the ABC module is also used for transport unrelated activities such as FeS center assembly and transcription. Interestingly, in the group of acidophilic archaea (P. torridus, S. solfataricus, S. tokodaii, T. acidophilum, and T. volcanicum), the fraction of secondary transporters (~68%) is very high. The genomes of these organisms encode a multitude of members of the amino acid-polyamine-organocation (APC) family of secondary transporters. These systems mostly function as solute:cation symporters or solute:solute antiporters. Moreover, the genomic analysis of archaea thus far indicates that secondary transporters of the NiCoT (Ni2+/Co2+ transporter) and Nramp (metal-ion transporter) family are exclusively found in acidophiles. In general, these transport systems are involved in the uptake of heavy metals or iron. Acidophiles must be resistant to high concentrations of heavy metals present in the metal-rich acidic environments. Some of these transporters might be involved in the resistance mechanisms such as the energy-dependent excretion of the metal ions from the cell.

CHANNELS MS channels have been extensively studied in eukarya and bacteria [10]. In these organisms, MS channels play an important role in organ functions, such as hearing, touching, and cell swelling. In bacteria, MS channels play a main role in the survival of osmotic stresses experienced under hypotonic conditions. The physiological function of MS channels in archaea has not been studied, but it is likely to be similar to bacteria. Interestingly, the growth of an Escherichia coli strain that lacks the endogenous MS channels can be partially restored by expression of the MS channel of M. jannaschii in a medium with high osmolarity [11]. Archaeal MS channels were first described in Haloferax volcanii [12], and systems from Thermoplasma acidophilum, T. volcanicum, and Methanocaldococcus jannaschii were cloned and characterized [11,13,14]. These channels share many mechanistic features with their bacterial counterparts such as gating by osmotic stress, large conductance and low ion selectivity, and weak voltage dependence. The two MS channels from M. jannaschii, MscMJ and MscMJLR, differ significantly in conductance (0.3 and 2.0 ns, respectively) [14], but they appear highly selective for cations. In this respect, these channels functionally resemble the Ca2+ -channels of skeletal and heart muscle [15]. Two classes of ion channels can be discriminated that are opened either by ligand binding (e.g., neurotransmitters) or by the transmembrane potential (voltage-gated channels). Structural information for both classes of channels has been gathered from archaeal counterparts, that is, the calcium-gated K+ -channel, MthK of Methanobacterium thermoautotrophicum [16] and the

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voltage-dependent K+ -channel, KvAP of Aeropyrum pernix [17]. MthK is a tetramer, and each subunit contains two integral membrane domains and one C-terminal regulators of K+ conductance (RCK) domain, which is located in the cytoplasm. The RCK domains form a gating ring. When Ca+ binds, the pore of the channel opens to allow K+ to be transported. The voltage-dependent K+ -channel from A. pernix, KvAP, was first shown to be a functional and structural homolog of eukaryal voltage-gated K+ -channels [18]. Subsequent structural studies revealed the presence of “voltage-sensor paddles” [17,19], which may carry the charge across the membrane along the interface of the transporter and the membrane in order to open the channel. However, this model [17,19] is highly controversial as it opposes the conventional hypothesis in which the charge is transported along a helix in the core of the protein. Recent data suggests that the movement of the sensor paddles might only be very small [20,21] and insufficient to carry ions across the membrane. Very recently three different groups solved the structure of CorA, the Mg2+ transporter of T. maritima (Table 4.2) [22–24]. CorA is thought to be responsible for the Mg2+ homeostasis in bacteria and archaea [24]. The protein forms a cone shaped pentamer, in which each monomer has two C-terminal transmembrane domains and a cytosolic N-terminal domain containing the metalbinding sites. A second metal binding site was identified in a region of the transmembrane domains that is thought to function as selectivity filter [24].

SECONDARY TRANSPORTERS Although secondary transporters are an important class in thermophilic genomes, few studies have been performed on members of this class. A lactose transporter was identified in S. solfataricus by functionally complementing a mutant strain unable to grow on lactose [25]. Complementation required both the lacS gene (β-galactosidase) and a gene located upstream of lacS that encodes a secondary transporter [26]. The lactose transporter belongs to the major facilitator family (MFS) and is predicted to contain 12 transmembrane segments.

TABLE 4.2 Solute Transporters and Subunit Structures From (Hyper-)Thermophiles Organism

Gene Name

ABC Transporter Archaeoglobus fulgidus

Component

Substrate

Methanococcus jannaschii Methanococcus jannaschii Sulfolobus solfataricus Pyrococcus horikoshii Pyrococcus furiosus Thermococcus litoralis Thermococcus litoralis Thermotoga maritima Thermotoga maritima

MJ0796 MJ1267 SSO2850 PH0022 PF1938 TK1771 TK1775 TM0544 TM1223

Substrate-binding protein Glycine betaine, proline betaine, trimethylammonium ATP-binding protein ATP-binding protein ATP-binding protein Glucose ATP-binding protein Maltose/trehalose Substrate-binding protein Maltodextrin Substrate-binding protein Maltose, trehalose ATP-binding protein ATP-binding protein Substrate-binding protein β-1,4-mannobiose

Thermotoga maritima

TM0189

Substrate-binding protein Iron

References [47] [71] [77] [72] [78] [46] [42] [41] 1VPL 1VR5 [38] 2ETV

MIT (Metal Transporting Family) Thermotoga maritima TM0561

Full-length transporter

Mg2+

[23]

Secondary Transporter Archaeoglobus fulgidus Pyrococcus horikoshii Aquifex aeolicus

Full-length transporter Full-length transporter Full-length transporter

Ammonium Glutamate Leucine

[79] [29] [80]

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Methanosarcina thermophila TM-1 and Methanohalophilus portucalensis strain FDF1 were both shown to actively accumulate compatible solutes such as glycine-betaine. The transporter of M. thermophila is highly specific for glycine-betaine and transports this molecule with a high affinity (Kt of 10 μM) [27]. Studies with inhibitors suggest that transport is dependent on a PMF or SMF. However, since the gene(s) have not yet been identified, the exact mechanism remains to be determined. In M. portucalensis, growth on trimethylamine in the presence of 2.1 M NaCl was dramatically stimulated by the presence of glycine-betaine [28]. The transporter involved in this process is also specific for glycine-betaine and transports the molecule with a Kt of 23 μM [28]. Recently, the structure of a putative glutamate transporter, GltPph, from Pyrococcus horikoshii was elucidated (Table 4.2) [29]. This was the first report of a structure of a secondary transporter from a member of the archaea. This protein is homologous to high-affinity neurotransmitter transporters of the synaptic cleft in mammals. It now has been shown that GltP transports L-glutamate in symport with Na+ [30] and aspartate in symport with Na+, although aspartate is the preferred substrate [31].

ABC TRANSPORTERS ABC transporters are a major class of transporters in archaea and bacteria, and several systems have been studied in great detail. In all domains of life, the subunits of these transporters exhibit the typical consensus motifs that classify them as ABC transporters. The ATPase subunits contain the typical Walker sequences, the Q-loop, and the H-region. Archaeal and bacterial ABC transporters can be divided into at least two classes: the binding-protein-dependent uptake systems and the multidrug/antibiotic transporter which probably functions as exporters. The latter class is typically composed of two subunits, each with one integral membrane domain typically comprising six transmembrane segments and a cytoplasmic nucleotide-binding domain. They may exist as homoor heterodimers. In some cases, the two domains are fused into a single polypeptide chain. With ABC-type uptake systems, the two permease and cytoplasmic ATPase domains are complemented by an extracellular substrate-binding protein. The membrane domains of ABC transporters contain the EAAAx3Gx9IxLP motif which has been shown to be the site of interaction with the cytoplasmic ATPase subunit [32]. ABC sugar transporters were identified and characterized in T. litoralis, P. furiosus, and S. solfataricus, and the bacteria T. maritima and T. thermophilus. Transport of osmoprotectants has been studied primarily in methanogens. Bacterial sugar ABC uptake systems are divided into two main classes: the carbohydrate uptake transporters (the CUT class) and the di/oligopeptide transport class [33]. The archaeal systems involved in the uptake of mono- or disaccharides belong to the CUT class. However, ABC transporters that transport di- and oligosaccharides such as the cellobiose/ β-glucoside transport system of P. furiosus [34] and the maltose/maltodextrin transporter of S. solfataricus [35], are homologous to the di/oligopeptide class. The similarity of the two systems not only relates to their organization, which consists of two permeases and two cytoplasmic ATPases, but also includes the substrate-binding proteins which show homology on the primary sequence level to bacterial di/oligopeptide-binding proteins. By analogy to bacterial di/oligopeptidebinding proteins, the homologous archaeal sugar-binding proteins recognize a large range of oligosaccharides [34]. The operons encoding subunits of putative di/oligopeptide transporters appear highly abundant in hyperthermophilic archaea and bacteria. Strikingly, these operons are positioned in the vicinity of genes that encode sugar-degrading enzymes [36]. In Thermotoga maritima, some of these transporters were shown to be upregulated when the cells are grown on specific sugar substrates [37]. Moreover, several predicted oligopeptide-binding proteins from T. maritima have been shown to indeed bind oligosaccharides, rather than peptides [38]. It is advantageous for archaea and bacteria to accumulate shorter sugar oligomers in a single transport step as uptake of sugar monomers and oligomers presumably requires similar amounts of ATP. Uptake of oligomers thus minimizes the overall energy requirements for substrate acquisition.

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The best-studied example of an archaeal ABC uptake system is the trehalose/maltose transporter of Thermococcus litoralis. The trehalose/maltose binding protein (TMBP) binds its substrate with very high affinity (160 nM at 80°C) [39]; while transport is also a high-affinity process. The ATPase subunit of this transporter, MalK, was expressed in E. coli and characterized [40], and the structure was elucidated for both MalK ATPase and TMBP-binding protein [41,42]. The entire transporter was heterologously expressed in E. coli and purified [43]. Although the detergentsolubilized and purified transporter displayed an intrinsic ATPase activity, the activity was not stimulated by the addition of binding protein, in contrast to previous observations with the bacterial systems [44]. So far, archaeal ABC transporters have not yet been functionally reconstituted into proteoliposomes. Substrate-Binding Protein The binding protein captures the substrate at the outside of the cell and delivers it to the membraneembedded permease domains, whereupon it is translocated across the membrane. Substrate-binding proteins contain two large domains (lobes) that are linked by a flexible hinge region. Upon binding of the substrate, the two lobes close like a “venus-fly trap” mechanism [45]. The structures have been solved for TMBP of T. litoralis [42], the maltodextrin-binding protein of P. furiosus (PfuMBP) [46], and ProX, the glycine-betaine and proline-betaine binding protein of Archaeoglobus fulgidus [47]. Both TMBP and PfuMBP have high structural similarity to the maltose-binding protein of E. coli (EcMBP); even though the primary amino acid sequence identity is low. Structurally equivalent, but not identical, amino acids are involved in sugar binding in TMBP and the EcMBP [42]. In all of these binding proteins, two elongated patches of hydrophobic amino acids outline the binding pocket. No significant heat release or absorption occurs when maltose binds to PfuMBP, in contrast to endothermic binding of maltose to EcMBP, which is unexpected if the protein would function by the venus-fly trap mechanism. Both enzymes exhibit the same binding constant for maltose. On the other hand, binding of maltotriose to PfuMBP was strongly exothermic and occurs with an affinity (Kd ~ 2.7 nM) that is much higher than observed for the EcMBP (Kd ~ 1.3 μM) [46]. In comparison to EcMBP, the bound sugar seems less deeply buried in the binding cleft of PfuMBP. The latter structure is much less flexible, indicating that the sugar bound form occurs more in an “open” than in an “occluded” state. It has been suggested that this binding mode resembles more a “lock and key” mode rather than the “venus-fly trap” mode [46]. ProX from A. fulgidus binds glycine-betaine and proline-betaine with Kd values of 60 and 50 nM at room temperature, respectively [48]. These compounds act as thermoprotectants in this organism. The structure of ProX was determined as the apo form and in complex with glycinebetaine, proline-betaine, and trimethylammonium [47]. Upon binding of these compounds, the two lobes move closer to each other. The binding of the substrates is mediated by cation–pi interactions and nonclassical hydrogen bonds between four tyrosine residues, a main chain carbonyl oxygen, and the substrates. The mechanism of binding of the ligands occurs in a manner that is similar to the E. coli homolog. However, the A. fulgidus and E. coli proteins have low-sequence identity, and the residues involved in binding are structurally equivalent but not conserved [47]. Very recently a tungsten and molybdate-binding protein was identified in P. furiosus. The binding protein was expressed in E. coli and shown by isothermal titration calorimetry to bind tungsten and molybdate with a Kd values of 17 and 11 nM, respectively [49]. Structural studies on archaeal substrate-binding proteins have been performed with heterologously expressed proteins from E. coli. A common feature of the substrate-binding proteins and other extracellular, especially membrane-bound proteins from archaea is that they are extensively glycosylated in their native host [43,50–52]. Binding proteins isolated from P. furiosus contain glucose moieties [53,54], whereas mannose, glucose, galactose, and N-acetylglucosamine have been identified in binding protein of S. solfataricus [35]. Due to the extensive glycosylation, the archaealbinding proteins are readily isolated from solubilized membrane fractions by lectin affinity

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chromatography [34,35,43,55]. However, glycosylation does not appear to be essential for functionality as the heterologous, nonglycosylated proteins normally bind sugar [34,39,42,46,47,53]. Glycosylation may affect the stability of the proteins, protect them from proteolytic degradation or influence their interaction with the cell envelope. The distinction between the CUT and di/oligopeptide subclasses of ABC transporters is also evident for the protein-domain organization of the binding proteins (Figure 4.2a and b). Binding proteins belonging to the CUT class contain an N-terminal signal peptide followed by a stretch of hydroxylated amino acids that can be up to 60 residues long (the ST-linker) (Figure 4.2b). The STlinker might act as a flexible region between the membrane anchor and the substrate-binding domain. ST-linkers have also identified in a pullulanase [50] and a haloarchaeal S-layer protein [52]. In the latter case, this region was the site of O-linked glycosylation. As archaeal cells are surrounded by an S-layer with pores of 4–5 nm size [56], small molecules and proteins can easily diffuse into the medium. Therefore, extracellular proteins need to be attached to the archaeal cell envelope. In gram-positive bacteria, sugar-binding proteins are attached to the cytoplasmic membrane by a lipid anchor that is covalently attached to the protein after translocation across the membrane. In bacteria, a cysteine residue at the +1 position is first lipidated prior to signal sequence removal by a specific lipoprotein signal peptidase. However, archaeal genomes do not appear to contain a homolog of the bacterial enzyme, and it is currently unclear how lipidation takes place in archaea. Nevertheless, the N-termini of many euryarchaeal-binding proteins contain the same “lipobox” motif as bacterial proteins; a cysteine residue at the +1 position that is preceded by small hydrophobic residues [57]. Analysis of the halocyanin of the archaeon, Natronobacter pharaonis, by mass spectroscopy, revealed that the N-terminal cysteine residue is covalently linked to a C20 diphytanyl diether lipid [58]. Members of the CUT class of binding proteins from S. solfataricus contain type IV pilin-like signal peptides and do not contain secretory signal peptides [35,55,59]. The signal peptides are usually found in the subunits of bacterial pili and archaeal flagellins [60], and typically consist of a positively charged N-terminus followed by a hydrophobic domain. The hydrophobic domain usually acts as a scaffold for the assembly of the translocated subunits into multimeric structure. However, this process requires the positively charged N-terminus to be removed by a type IV pilin-like signal peptidase. Due to the presence of type IV pilin-like signal peptides, it has been hypothesized that the binding proteins of S. solfataricus multimerize into a structure, referred to as the “bindosome” [61]. In vitro assays demonstrated that the signal peptides of these binding proteins are processed by a specialized type IV prepilin signal peptidase, PibD, and not by the typical signal peptidase I [62].

FIGURE 4.2 Mode of anchoring of archaeal substrate binding proteins and domain structure. (a) Archaeal substrate-binding proteins are either bound to the membrane by a fatty acid modification of their N-terminus, or a hydrophobic domain at the C- or N-terminus. (b) Domain organization of substrate-binding proteins. N, N-terminus; C, C-terminus; SS, signal sequence; ST-linker, serine/threonine-rich amino acid stretch; filled circle, substrate of binding protein.

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This enzyme also processes the flagellin subunits before assembly into the flagellum [63]. Moreover, deletion mutant analysis in S. solfataricus showed that two proteins, a cytoplasmic ATPase and an integral membrane protein, which share homology to bacterial type II secretion systems, are essential for the functional assembly of the binding proteins exhibiting type IV pilin-like signal peptides (Zolghadr et al., 2007). The binding proteins of the di/oligopeptide class contain typical bacterial signal peptides that are processed by an archaeal homolog of signal peptidase I (Figure 4.2) [64,65]. In contrast to the CUT class binding proteins, these proteins harbor a hydrophobic region at the C-terminus that is preceded by an ST-linker. Therefore, it appears that the CUT-class binding proteins have a mirrorlike domain organization of that of the di/oligopeptide-class sugar-binding proteins. Due to the presence of the C-terminal hydrophobic domain, which is likely to function as a transmembrane domain to anchor these proteins to the cytoplasmic membrane, the di/oligopeptide-class binding proteins do not need to be lipid-modified at their N-termini. However, some pyrococcal-binding proteins contain a GGICG sequence motif immediately downstream of the hydrophobic domain at their C-terminus. Similar to some halophilic S-layer proteins that contain this motif, C-terminal lipid modification [66] may occur at these positions. Many archaea encode more binding proteins of the CUT class than other classes [57]. S. solfataricus is an exception in that the di/oligopeptide class of binding proteins are more abundant that the proteins belonging to the CUT class. On the other hand, methanogens do not contain any member of the di/oligopeptide class. This might imply that S. solfataricus grows on carbon sources derived from higher oligomeric substrates without the need of extracellular enzymatic “predigestion,” whereas methanogens do need to enzymatically process substrates before they can be transported into the cell. Very recently, two new archaeal-binding proteins have been characterized: the tungsten-binding protein, WtpA of P. furiosus [49] and an oligopeptide-binding protein of Aeropyrum pernix [67]. By isothermal titration it was demonstrated that WtpA binds tungstate and molybdate with very high affinities, Kd of 0.017 nM and 11 nM, respectively. Like other binding proteins of Pyrococcus, this protein contains a putative lipid-box motif at the N-terminus and might therefore be anchored to the membrane via a lipid anchor (see earlier). The only identified archaeal oligopeptide-binding protein from A. pernix was shown to bind 8 to 13 amino acids long oligopeptides, with a preference for the peptide xenopsin (EGKRPWIL) at high temperature (90°C) [67]. ATP-Binding Protein ATP-binding proteins of ABC transporters provide the driving force for substrate translocation. These proteins typically exist as homo- or heterodimers. Binding of ATP stabilizes the dimeric form of the isolated subunits, whereas hydrolysis of ATP causes the destabilization of the dimer. In short, upon ATP binding, the conformational change in the ATPase dimer affects the permease domain, and result in a conformational change of the membrane domains whereupon the substrate is transported. ABC ATPases are highly conserved proteins and present in all three domains of life. They share a number of motifs, such as the Walker A/B motifs [60] and the ABC signature motif, LSGGQ [68]. The residues of the Walker motifs have been shown to be important for binding of ATP and hydrolysis, whereas the ABC signature motif is essential for the dimerization process following ATP binding [69]. A number of archaeal ATP-binding proteins have been crystallized, such as LolD (MJ0769) and LivG (MJ1267) from M. jannaschii [70], MalK from the trehalose/maltose transporter of T. litoralis [14], and GlcV, the ATPase subunit of the glucose transporter of S. solfataricus [71] (Figure 4.3). Studies on LivG showed that the ABC signature motif is essential for dimer formation. Dimerization results in the completion of the catalytic ATP binding site, as both monomers contribute residues for the active site [69]. Although the ABC ATPase dimer contains two nucleotide-binding sites, hydrolysis of ATP at only one of the sites appears sufficient for transport [72]. The overall fold of the

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FIGURE 4.3 Structure of the ATP-binding subunit, GlcV of the glucose ABC transporter of S. solfataricus. The ATP-binding domain contains all the necessary residues for ATP hydrolysis. Bound ATP is shown in the structure (ball model). The function of the C-terminal domain is unknown.

archaeal ATP-binding domains is structurally the same as that of bacterial and eukaryal proteins. Interestingly, MalK and GlcV contain an additional domain at their C-terminus (Figure 4.3) that predominantly consists of β-sheets that are organized in an OB (oligonucleotide/oligosaccharide) fold. This C-terminal extension is also present in MalK, the ATPase of the E. coli maltose ABC transporter [73]. This domain interacts with a positive regulator of the mal operon, MalT [74]. When MalT is bound by MalK, activation of the mal operon cannot occur. However, when maltose is present in the medium, ATP is hydrolyzed by MalK, and MalT is released into the cytosol where it can activate the transcription of the mal operon. This is the only known example of an ATPase subunit that is involved in the regulation of expression of its own operon. MalT homologs have not been identified in archaea, and at this stage it is unclear if the C-terminal extension of the archaeal ABC ATPases are involved in regulation. Expression of the trehalose/maltose transporter of T. litoralis is repressed by the negative regulator, TrmB [75]. In P. furiosus, TrmB regulates both the trehalose/maltose transporter and the maltodextrin transporter [53,54,76]. This argues against the presence of a MalT-like regulation system in archaea. Interestingly, DNA microarray studies in P. furiosus revealed that the 16 kb region in the genome that is nearly identical to a portion of the T. litoralis genome, showed that all the transporter genes are upregulated during growth on maltose, whereas malK and trmB retain high levels of expression, even when cells are grown on peptides. This implies that there might be a regulatory role for MalK beside the direct regulation of the expression of the operon by TrmB, but how this regulation is conferred remains to be investigated.

CONCLUDING REMARKS Both from a structural and functional point of view, there is much interest in proteins from (hyper-) thermophiles. They appear particularly suitable for structural studies, presumably because at low temperatures they approximate a frozen state that tends to crystallize more readily than proteins isolated from mesophilic counterparts. Although this might be true for cytoplasmic proteins, with thermophilic membrane proteins only limited successes have been reported (Table 4.2). For protein crystallization, the protein of interest must, in particular, be stable in detergent solution and resist the effects of delipidation. In this respect, not only the protein composition, but also the lipid environment plays an important role in the thermostability of membrane proteins. Therefore, upon detergent extraction, membrane proteins from thermophiles may readily lose their thermostability unless the proteins are not completely delipidated.

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Another key element of membrane protein crystallization is the availability of sufficiently large amounts of purified membrane protein. Recent advances in the development of a genetic toolbox for protein expression and gene inactivation in hyperthermophilic archaea such as Thermus kodalaraensis and Sulfolobus solfataricus (see Chapters 11 and 12), now provide new directions for membrane proteins expression from thermophilic sources and will allow a more thorough functional analysis than has been possible until recently.

REFERENCES 1. Melchior, D. L. 1982. Lipid phase transitions and regulation of membrane fluidity in prokaryotes. Curr. Top. Membr. Transp. 17:263–316. 2. Kates, M. 1996. Structural analysis of phospholipids and glycolipids in extremely halophilic archaebacteria. J. Microbiol. Meth. 25:113–128. 3. Kates, M., N. Moldoveanu, and L. C. Stewart. 1993. On the revised structure of the major phospholipid of Halobacterium salinarium. Biochim. Biophys. Acta 1169:46–53. 4. Upasani, V. N., S. G. Desai, N. Moldoveanu, and M. Kates. 1994. Lipids of extremely halophilic archaeobacteria from saline environments in India: a novel glycolipid in Natronobacterium strains. Microbiology 140:1959–1966. 5. De Rosa, M., A. Trincone, B. Nicolaus, and A. Gambacorta. 1991. Archaebacteria: lipids, membrane structures, and adaptations to environmental stresses. In G. di Prisco (ed.), Life under extreme conditions. Springer-Verlag, Berlin Heidelberg, pp. 61–87. 6. Elferink, M. G. L., J. G. De Wit, R. Demel, A. J. M. Driessen, and W. N. Konings. 1992. Functional reconstitution of membrane proteins in monolayer liposomes from bipolar lipids of Sulfolobus acidocaldarius. J. Biol. Chem. 267:1375–1381. 7. Langworthy, T. A. 1982. Lipids of Thermoplasma. Methods Enzymol. 88:369–406. 8. De Rosa, M. and A. Gambacorta. 1988. The lipids of archaebacteria. Prog. Lipid Res. 27:153–175. 9. Sprott, G. D., M. Meloche, and J. C. Richards. 1991. Proportions of diether, macrocyclic diether, and tetraether lipids in Methanococcus jannaschii grown at different temperatures. J. Bacteriol. 173:3907–3910. 10. Hamill, O. P. and B. Martinac. 2001. Molecular basis of mechanotransduction in living cells. Physiol. Rev. 81:685–740. 11. Kloda, A. and B. Martinac. 2001. Molecular identification of a mechanosensitive channel in archaea. Biophys. J. 80:229–240. 12. Le Dain, A. C., N. Saint, A. Kloda, A. Ghazi, and B. Martinac. 1998. Mechanosensitive ion channels of the archaeon Haloferax volcanii. J. Biol. Chem. 273:12116–12119. 13. Kloda, A. and B. Martinac. 2001. Mechanosensitive channel of Thermoplasma, the cell wall-less archaea: cloning and molecular characterization. Cell Biochem. Biophys. 34:321–347. 14. Kloda, A. and B. Martinac. 2001. Structural and functional differences between two homologous mechanosensitive channels of Methanococcus jannaschii. EMBO J. 20:1888–1896. 15. Martinac, B. 2004. Mechanosensitive ion channels: molecules of mechanotransduction. J. Cell Sci. 117:2449–2460. 16. Jiang, Y., A. Lee, J. Chen, M. Cadene, B. T. Chait, and R. MacKinnon. 2002. Crystal structure and mechanism of a calcium-gated potassium channel. Nature 417:515–522. 17. Jiang, Y., A. Lee, J. Chen, V. Ruta, M. Cadene, B. T. Chait, and R. MacKinnon. 2003. X-ray structure of a voltage-dependent K+ channel. Nature 423:33–41. 18. Ruta, V., Y. Jiang, A. Lee, J. Chen, and R. MacKinnon. 2003. Functional analysis of an archaebacterial voltage-dependent K+ channel. Nature 422:180–185. 19. Jiang, Y., V. Ruta, J. Chen, A. Lee, and R. MacKinnon. 2003. The principle of gating charge movement in a voltage-dependent K+ channel. Nature 423:42–48. 20. Chanda, B., O. Kwame Asamoah, R. Blunck, B. Roux, and F. Bezanilla. 2005. Gating charge displacement in voltage-gated ion channels involves limited transmembrane movement. Nature 436:852–856. 21. Revell Phillips, L., M. Milescu, Y. Li-Smerin, J. A. Mindell, J. I. Kim, and K. J. Swartz. 2005. Voltagesensor activation with a tarantula toxin as cargo. Nature 436:857–860.

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22. Eshaghi, S., D. Niegowski, A. Kohl, M. D. Martinez, S. A. Lesley, and P. Nordlund. 2006. Crystal structure of a divalent metal ion transporter CorA at 2.9 angstrom resolution. Science 313:354–357. 23. Lunin, V. V., E. Dobrovetsky, G. Khutoreskaya, R. Zhang, A. Joachimiak, D. A. Doyle, A. Bochkarev, M. E. Maguire, A. M. Edwards, and C. M. Koth. 2006. Crystal structure of the CorA Mg2+ transporter. Nature 440:833–837. 24. Payandeh, J. and E. F. Pai. 2006. A structural basis for Mg2+ homeostasis and the CorA translocation cycle. EMBO J. 25:3762–3773. 25. Bartolucci, S., M. Rossi, and R. Cannio. 2003. Characterization and functional complementation of a nonlethal deletion in the chromosome of a beta-glycosidase mutant of Sulfolobus solfataricus. J. Bacteriol. 185:3948–3957. 26. Prisco, A., M. Moracci, M. Rossi, and M. Ciaramella. 1995. A gene encoding a putative membrane protein homologous to the major facilitator superfamily of transporters maps upstream of the β-glycosidase gene in the Archaeon Sulfolobus solfataricus. J. Bacteriol. 177:1614–1619. 27. Proctor, L. M., R. Lai, and R. P. Gunsalus. 1997. The methanogenic archaeon Methanosarcina thermophila TM-1 possesses a high-affinity glycine betaine transporter involved in osmotic adaptation. Appl. Environ. Microbiol. 63:2252–2257. 28. Lai, M. C., T. Y. Hong, and R. P. Gunsalus. 2000. Glycine betaine transport in the obligate halophilic archaeon Methanohalophilus portucalensis. J. Bacteriol. 182:5020–5024. 29. Yernool, D., O. Boudker, Y. Jin, and E. Gouaux. 2004. Structure of a glutamate transporter homologue from Pyrococcus horikoshii. Nature 431:811–818. 30. Raunser, S., M. Appel, C. Ganea, U. Geldmacher-Kaufer, K. Fendler, and W. Kuhlbrandt. 2006. Structure and function of prokaryotic glutamate transporters from Escherichia coli and Pyrococcus horikoshii. Biochemistry 45:12796–12805. 31. Boudker, O., R. M. Ryan, D. Yernool, K. Shimamoto, and E. Gouaux. 2007. Coupling substrate and ion binding to extracellular gate of a sodium-dependent aspartate transporter. Nature 445:387–393. 32. Dassa, E. and M. Hofnung. 1985. Sequence of gene malG in E. coli K12: homologies between integral membrane components from binding protein-dependent transport systems. EMBO J. 4:2287–2293. 33. Schneider, E. 2001. ABC transporters catalyzing carbohydrate uptake. Res. Microbiol. 152:303–310. 34. Koning, S. M., M. G. Elferink, W. N. Konings, and A. J. M. Driessen. 2001. Cellobiose uptake in the hyperthermophilic archaeon Pyrococcus furiosus is mediated by an inducible, high-affinity ABC transporter. J. Bacteriol. 183:4979–4984. 35. Elferink, M. G. L., S.-V. Albers, W. N. Konings, and A. J. M. Driessen. 2001. Sugar transport in Sulfolobus solfataricus is mediated by two families of binding protein-dependent ABC transporters. Mol. Microbiol. 39:1494–1503. 36. Nelson, K. E., R. A. Clayton, S. R. Gill, M. L. Gwinn, R. J. Dodson, D. H. Haft, E. K. Hickey, J. D. Peterson, W. C. Nelson, K. A. Ketchum, L. McDonald, T. R. Utterback, J. A. Malek, K. D. Linher, M. M. Garrett, A. M. Stewart, M. D. Cotton, M. S. Pratt, C. A. Phillips, D. Richardson, J. Heidelberg, G. G. Sutton, R. D. Fleischmann, J. A. Eisen, and C. M. Fraser. 1999. Evidence for lateral gene transfer between Archaea and bacteria from genome sequence of Thermotoga maritima. Nature 399:323–329. 37. Chhabra, S. R., K. R. Shockley, S. B. Conners, K. L. Scott, R. D. Wolfinger, and R. M. Kelly. 2003. Carbohydrate-induced differential gene expression patterns in the hyperthermophilic bacterium Thermotoga maritima. J. Biol. Chem. 278:7540–7552. 38. Nanavati, D. M., K. Thirangoon, and K. M. Noll. 2006. Several archaeal homologs of putative oligopeptide-binding proteins encoded by Thermotoga maritima bind sugars. Appl. Environ. Microbiol. 72:1336–1345. 39. Horlacher, R., K. B. Xavier, H. Santos, J. DiRuggiero, M. Kossmann, and W. Boos. 1998. Archaeal binding protein-dependent ABC transporter: molecular and biochemical analysis of the trehalose/ maltose transport system of the hyperthermophilic archaeon Thermococcus litoralis. J. Bacteriol. 180:680–689. 40. Greller, G., R. Horlacher, J. DiRuggiero, and W. Boos. 1999. Molecular and biochemical analysis of MalK, the ATP-hydrolyzing subunit of the trehalose/maltose transport system of the hyperthermophilic archaeon Thermococcus litoralis. J. Biol. Chem. 274:20259–20264.

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41. Diederichs, K., J. Diez, G. Greller, C. Muller, J. Breed, C. Schnell, C. Vonrhein, W. Boos, and W. Welte. 2000. Crystal structure of MalK, the ATPase subunit of the trehalose/maltose ABC transporter of the archaeon Thermococcus litoralis. EMBO J. 19:5951–5961. 42. Diez, J., K. Diederichs, G. Greller, R. Horlacher, W. Boos, and W. Welte. 2001. The Crystal Structure of a Liganded Trehalose/Maltose-binding Protein from the Hyperthermophilic Archaeon Thermococcus litoralis at 1.85 Å. J. Mol. Biol. 305:905–915. 43. Greller, G., R. Riek, and W. Boos. 2001. Purification and characterization of the heterologously expressed trehalose/maltose ABC transporter complex of the hyperthermophilic archaeon Thermococcus litoralis. Eur. J. Biochem. 268:4011–4018. 44. Reich-Slotky, R., C. Panagiotidis, M. Reyes, and H. A. Shuman. 2000. The detergent-soluble maltose transporter is activated by maltose binding protein and verapamil. J. Bacteriol. 182:993–1000. 45. Quiocho, F. A. and P. S. Ledvina. 1996. Atomic structure and specificity of bacterial periplasmic receptors for active transport and chemotaxis: variation of common themes. Mol. Microbiol. 20:17–25. 46. Evdokimov, A. G., D. E. Anderson, K. M. Routzahn, and D. S. Waugh. 2001. Structural basis for oligosaccharide recognition by Pyrococcus furiosus maltodextrin-binding protein. J. Mol. Biol. 305:891–904. 47. Schiefner, A., G. Holtmann, K. Diederichs, W. Welte, and E. Bremer. 2004. Structural basis for the binding of compatible solutes by ProX from the hyperthermophilic archaeon Archaeoglobus fulgidus. J. Biol. Chem. 279:48270–48281. 48. Holtmann, G. 2004. Thesis. University of Marbug, Germany. 49. Bevers, L. E., P. L. Hagedoorn, G. C. Krijger, and W. R. Hagen. 2006. Tungsten transport protein A (WtpA) in Pyrococcus furiosus: the first member of a new class of tungstate and molybdate transporters. J. Bacteriol. 188:6498–6505. 50. Erra-Pujada, M., P. Debeire, F. Duchiron, and M. J. O′Donohue. 1999. The type II pullulanase of Thermococcus hydrothermalis: molecular characterization of the gene and expression of the catalytic domain. J. Bacteriol. 181:3284–3287. 51. Hettmann, T., C. L. Schmidt, S. Anemuller, U. Zahringer, H. Moll, A. Petersen, and G. Schafer. 1998. Cytochrome b558/566 from the archaeon Sulfolobus acidocaldarius. A novel highly glycosylated, membrane-bound b-type hemoprotein. J. Biol. Chem. 273:12032–12040. 52. Sumper, M., E. Berg, R. Mengele, and I. Strobel. 1990. Primary structure and glycosylation of the S-layer protein of Haloferax volcanii. J. Bacteriol. 172:7111–7118. 53. Koning, S. M., W. N. Konings, and A. J. M. Driessen. 2002. Biochemical evidence for the presence of two α-glucoside ABC-transport systems in the hyperthermophilic archaeon Pyroccocus furiosus. Archaea 1:19–25. 54. Konings, S. M., S.-V. Albers, W.N. Konings, and A. J. M. Driessen. 2002. Sugar Transport in (Hyper-) thermophilic Achaea. Res. Microbiol. 153: 61–67. 55. Albers, S. V., M. G. Elferink, R. L. Charlebois, C. W. Sensen, A. J. M. Driessen, and W. N. Konings. 1999. Glucose transport in the extremely thermoacidophilic Sulfolobus solfataricus involves a highaffinity membrane-integrated binding protein. J. Bacteriol. 181:4285–4291. 56. Koenig, H. 1988. Archaeobacterial cell envelopes. Can. J. Microbiol. 34:395–406. 57. Albers, S. V., S. M. Koning, W. N. Konings, and A. J. M. Driessen. 2004. Insights into ABC transport in archaea. J. Bioenerg. Biomembr. 36:5–15. 58. Mattar, S., B. Scharf, S. B. Kent, K. Rodewald, D. Oesterhelt, and M. Engelhard. 1994. The primary structure of halocyanin, an archaeal blue copper protein, predicts a lipid anchor for membrane fixation. J. Biol. Chem. 269:14939–14945. 59. Albers, S. V., W. N. Konings, and A. J. M. Driessen. 1999. A unique short signal sequence in membrane anchored proteins of Archaea. Mol. Microbiol. 31:1595–1596. 60. Thomas, N. A., S. L. Bardy, and K. F. Jarrell. 2001. The archaeal flagellum: a different kind of prokaryotic motility structure. FEMS Microbiol. Rev. 25:147–174. 61. Albers, S. V. and A. J. M. Driessen. 2005. Analysis of ATPases of putative secretion operons in the thermoacidophilic archaeon Sulfolobus solfataricus. Microbiology 151:763–773. 62. Albers, S. V., Z. Szabó, and A. J. M. Driessen. 2003. Archaeal homolog of bacterial type IV prepilin signal peptidases with broad substrate specificity. J. Bacteriol. 185:3918–3925. 63. Bardy, S. L. and K. F. Jarrell. 2002. FlaK of the archaeon Methanococcus maripaludis possesses preflagellin peptidase activity. FEMS Microbiol. Lett. 208:53–59.

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64. Bardy, S. L., S. Y. Ng, D. S. Carnegie, and K. F. Jarrell. 2005. Site-directed mutagenesis analysis of amino acids critical for activity of the type I signal peptidase of the archaeon Methanococcus voltae. J. Bacteriol. 187:1188–1191. 65. Ng, S. Y. and K. F. Jarrell. 2003. Cloning and characterization of archaeal type I signal peptidase from Methanococcus voltae. J. Bacteriol. 185:5936–5942. 66. Konrad, Z. and J. Eichler. 2002. Lipid modification of proteins in Archaea: attachment of a mevalonic acid-based lipid moiety to the surface-layer glycoprotein of Haloferax volcanii follows protein translocation. Biochem. J. 366:959–964. 67. Palmieri, G., A. Casbarra, I. Fiume, G. Catara, A. Capasso, G. Marino, S. Onesti, and M. Rossi. 2006. Identification of the first archaeal oligopeptide-binding protein from the hyperthermophile Aeropyrum pernix. Extremophiles. 10:393–402. 68. Boos, W. and H. Shuman. 1998. Maltose/maltodextrin system of Escherichia coli: transport, metabolism, and regulation. Microbiol. Mol. Biol. Rev. 62:204–229. 69. Moody, J. E., L. Millen, D. Binns, J. F. Hunt, and P. J. Thomas. 2002. Cooperative, ATP-dependent association of the nucleotide binding cassettes during the catalytic cycle of ATP-binding cassette transporters. J. Biol. Chem. 277:21111–21114. 70. Yuan, Y. R., S. Blecker, O. Martsinkevich, L. Millen, P. J. Thomas, and J. F. Hunt. 2001. The crystal structure of the MJ0796 ATP-binding cassette: Implications for the structural consequences of ATP hydrolysis in the active site of an ABC-transporter. J. Biol. Chem. 276:32313–32321. 71. Verdon, G., S. V. Albers, B. W. Dijkstra, A. J. Driessen, and A. M. Thunnissen. 2003. Crystal Structures of the ATPase Subunit of the Glucose ABC Transporter from Sulfolobus solfataricus: Nucleotide-free and Nucleotide-bound Conformations. J. Mol. Biol. 330:343–358. 72. Verdon, G., S. V. Albers, N. van Oosterwijk, B. W. Dijkstra, A. J. M. Driessen, and A. M. Thunnissen. 2003. Formation of the productive ATP-Mg2+ -bound dimer of GlcV, an ABC-ATPase from Sulfolobus solfataricus. J. Mol. Biol. 334:255–267. 73. Chen, J., G. Lu, J. Lin, A. L. Davidson, and F. A. Quiocho. 2003. A tweezers-like motion of the ATP-binding cassette dimer in an ABC transport cycle. Mol. Cell 12:651–661. 74. Panagiotidis, C. H., W. Boos, and H. A. Shuman. 1998. The ATP-binding cassette subunit of the maltose transporter MalK antagonizes MalT, the activator of the Escherichia coli mal regulon. Mol. Microbiol. 30:535–546. 75. Lee, S. J., A. Engelmann, R. Horlacher, Q. Qu, G. Vierke, C. Hebbeln, M. Thomm, and W. Boos. 2003. TrmB, a sugar-specific transcriptional regulator of the trehalose/maltose ABC transporter from the hyperthermophilic archaeon Thermococcus litoralis. J. Biol. Chem. 278:983–990. 76. Lee, S. J., C. Moulakakis, S. M. Koning, W. Hausner, M. Thomm, and W. Boos. 2005. TrmB, a sugar sensing regulator of ABC transporter genes in Pyrococcus furiosus exhibits dual promoter specificity and is controlled by different inducers. Mol. Microbiol. 57:1797–1807. 77. Karpowich, N., O. Martsinkevich, L. Millen, Y. R. Yuan, P. L. Dai, K. MacVey, P. J. Thomas, and J. F. Hunt. 2001. Crystal structures of the MJ1267 ATP binding cassette reveal an induced-fit effect at the ATPase active site of an ABC transporter. Structure 9:571–586. 78. Ose, T., T. Fujie, M. Yao, N. Watanabe, and I. Tanaka. 2004. Crystal structure of the ATP-binding cassette of multisugar transporter from Pyrococcus horikoshii OT3. Proteins 57:635–638. 79. Andrade, S. L., A. Dickmanns, R. Ficner, and O. Einsle. 2005. Crystal structure of the archaeal ammonium transporter Amt-1 from Archaeoglobus fulgidus. Proc. Natl. Acad. Sci. U. S. A. 102:14994–14999. 80. Yamashita, A., S. K. Singh, T. Kawate, Y. Jin, and E. Gouaux. 2005. Crystal structure of a bacterial homologue of Na+/Cl− -dependent neurotransmitter transporters. Nature 437:215–223.

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Thermophilic ProteinFolding Systems Frank T. Robb and Pongpan Laksanalamai

CONTENTS Introduction .................................................................................................................................. Genome Size and Complexity of the Repertoires of Protein Chaperones ....................................................................................................... Chaperones and Thermotolerance ............................................................................................... Regulation of sHSP Expression ................................................................................................... Mechanistic Insights from Minimal Complexity ........................................................................ Structure and Subunit Composition of Archaeal Group II Chaperonins ......................... Prefoldins .......................................................................................................................... Small Heat Shock Proteins ............................................................................................... Nascent-Associated Proteins ............................................................................................ Protein-Folding Mechanism of Archaeal Group II Chaperonins ................................................ Perspectives: Next Five Years ...................................................................................................... Acknowledgment ......................................................................................................................... References ....................................................................................................................................

55 56 56 57 58 59 61 63 64 64 65 66 66

INTRODUCTION For survival and efficient growth under normal conditions, cells must be able to maintain the majority of their proteins in native conformations, and to recover or degrade proteins damaged during stress challenges. All organisms thus face a protein-folding problem, which is the requirement to convert their proteins from random coil or partially folded conformations into homogeneous, precisely folded states. Thermophiles grow at temperatures up to 113°C [1], and possibly as high as 121°C [2], and thermoacidophiles in solfataric environments at pH 0–1 at moderately high temperatures. Picrophilus torridus, which grows optimally at pH 0.8, has an intracellular pH of 4.6 unlike other thermoacidophiles, which maintain neutral pH. Therefore, in this case both extracellular and intracellular proteins must be stabilized against low pH in addition to temperature stress [3]. Protein folding is arguably more challenging in thermophiles than it is in mesophiles due to structural adaptations leading to enhanced protein thermostability. The compositional and conformational adaptations of proteins that result in intrinsic stability under moderate or extreme conditions are still the topic of active debate, however some principles have emerged. Protein adaptations include highly charged exterior surfaces, rigid folds maintained by multiple ion-pair networks [4], tight hydrophobic core packing, and overall compact protein structures achieved by increased packing density to minimize internal voids [5]. 55

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Adaptive amino acid substitutions have been characterized through large-scale comparisons of protein sequences and compositional variation in large data sets [6,7]. Identification of thermophilic species was done by the amino acid compositions deduced from their genomes [6]. High optimal growth temperature is accompanied by a proportionately increased content of the charged amino acids (lysine, arginine, glutamate, and aspartate) resulting in increased surface charge and ion pair formation. Other compositional changes include a higher average residue volume and a decrease in charged nonpolar amino acids on the surface of proteins [8,9] (see Chapter 3). In extremely stable proteins, the high intrinsic stability of the proteins requires the folding process to localize charged residues within the hydrophobic interior of the protein [10–12] raising questions regarding the energetics of folding at very high temperatures. Localization of a charged residue in the hydrophobic interior of a protein uses an additional increment of free energy, the desolvation penalty [12]. The recombinant expression of thermostable proteins in mesophilic hosts is often accompanied by misfolding, suggesting that thermophile-specific folding pathways might be operating. This chapter describes thermophile protein-folding pathways, including both heat shock proteins (HSPs) and constitutive protein chaperones.

GENOME SIZE AND COMPLEXITY OF THE REPERTOIRES OF PROTEIN CHAPERONES The published genomes of thermophilic species span a significant size range, from the 0.49-Mb genome of Nanoarchaeum equitans to the Symbiobacterium thermophilum genome of 3.57 Mb [13]. The hyperthermophiles, with optimal growth temperatures (Topt ) > 80°C, tend to have small genomes with many genes that exist as single paralogs that often occur in multiple copies in mesophiles [14]. Coding density in these compact circular genomes is high, with N. equitans, with the smallest genome, having the highest coding density among the hyperthermophiles [15]. Inventories of chaperones found in the genomes of archaea include representatives of several protein families they share, including the prefoldins, small heat shock protein (sHsp), and class II adenosine triphosphate (ATP)-dependent chaperonins (Table 5.1). Two major classes of eukaryal chaperones (Hsp100 and Hsp90/Hsp83) are absent in archaeal genome sequences. The chaperones that are shared by archaea and bacteria include the “Chaperone Machine” [16], which is composed of Hsp70 (DnaK), Hsp40 (DnaJ), and GrpE, only occur in the larger complete genome sequences of archaea [17,18], as well as the psychrophilic methanogen, Methanococcoides burtonii [19, R. Cavicchiolli, personal communication]. Hyperthermophiles represented by Pyrococcus spp, Sulfolobus spp, Pyrobaculum aerophilum, Methanocaldococcus jannaschii, Methanopyrus kandleri, Archaeoglobus fulgidus, N. equitans, and Picrophilus torridus [20] do not have Hsp90, DnaK, DnaJ, GrpE, Hsp33, and Hsp10 homologs (Table 5.1). The smaller archaeal genomes lack the highest molecular weight chaperones found in eukarya. The Hsp100/Clp protein family, which are absent from the genomes of the hyperthermophilic archaea, are present in several mesophilic and thermophilic archaea (Table 5.1). For example, the thermophilic methanogen, Methanothermobacter thermautotrophicus contains a ClpA/B homolog, which was probably acquired by lateral gene transfer from bacteria [21]. In Escherichia coli, degradation of denatured proteins is mediated by the cooperative functions of the ClpA and ClpP proteins. In addition to protein turnover, ClpA has protein remodeling functions and “protein repair” functions. In E. coli, ClpA alone can reactivate replication initiator protein, RepA, from an inactive RepA dimer to an active RepA monomer [22]; thermophilic bacteria encode Clp proteins, but post-translational remodeling by chaperones has not been characterized in any thermophile so far.

CHAPERONES AND THERMOTOLERANCE Organisms exposed to sublethal heat shock may develop short-term tolerance to otherwise lethal temperatures. This phenomenom is referred to as acquired thermotolerance. It has been well

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TABLE 5.1 Occurrence of Different Classes of Heat Shock Proteins (HSPs) in the Three Domains HSP HSP100s HSP90s HSP70s HSP60s sHSPs

Bacteria ClpA, ClpB, HslU HtpG DnaK GroEL/ES IbpA, IbpB (Escherichia coli)

Eukarya HSP100 Hsp90, Hsp83 Hsp70, Hsc70 TCP-1 sHSP, α-crystallin

Archaea Absent* Absent Hsp70† TF55, thermosome, Cpn60 sHSP

* ClpA/B homologs are found in Methanobacterium thermoautotrophicum. † Hsp70s are absent in most thermophiles and hyperthermophiles except Aeropyrum pernix, in which the putative mitochondrial HSP70 has been identified from the complete genome sequence.

established in a diverse range of organisms (e.g., Drosophila, yeast, E. coli) that heat shock protein (HSP) induction is responsible for acquired thermotolerance. In the archaea, evidence for an adaptive thermotolerance response linked to chaperone expression was first discovered in the hyperthermophilic archaeal species, Sulfolobus shibatae [23,24]. Acquired thermotolerance was achieved following heat shock at 88°C for 60 min which enabled the cells to survive normally lethal exposure at 95°C for 40 min [23]. Acquired thermotolerance was accompanied by the synthesis of high levels of the Hsp60, a class II chaperonin, also referred to as the thermosome, archaeosome, or rosettasome [25–27]. The gene for a putative AAA+ ATPase homolog (NP_579611) from the hyperthermophile, Pyrococcus furiosus, is up-regulated following heat shock at 105°C by induction of a repressor protein, Phr (heat shock regulator protein) [28]. A single phr gene is present in all three Pyrococcus genome sequences (P. furiosus, Pyrococcus abyssi, and Pyrococcus horikoshii), and encodes a 24-kDa basic protein. In P. furiosus, the promoters of the heat shock-inducible hsp20 (Pfu-shsp) and aaa+ ATPase genes have highly conserved dyad operator sites [29]. The expression of the phr gene was not induced by heat shock suggesting that the Phr protein may be required at both normal growth and heat shock temperatures. Repression is relieved by an unknown mechanism during heat shock, and a cis-acting regulatory sequence has been described that may be important for heat shock regulation [28]. Aligning the upstream regions of the AAA+ encoding genes from P. furiosus and P. abyssi enabled conserved regions to be identified which may be Phr-binding sites in both organisms [14]. The promoter region of the AAA+ gene from P. abyssi also has phr recognition motifs similar to the promoter of the heat-inducible, shsp gene from P. furiosus, indicating that these two species may be using a common heat shock regulatory mechanism [30]. HtpX is a putative membrane-bound metalloprotease in bacteria and is ubiquitous in the archaea, although it is annotated in many archaeal genomes as a conserved hypothetical protein. One copy of the htpX gene is present in the genomes of P. furiosus and P. abyssi, and two copies are present in each of the genomes of P. horikoshii and Sulfolobus solfataricus. Similar to AAA+ protein genes, htpX is heat inducible in M. jannaschii [31] and A. fulgidus [32,33] and P. furiosus [34 P. Laksanalamai, J. DiRuggiero, F. Robb, and T. Lowe, manuscript in preparation]. It is possible that htpX may have a cellular function that is similar to, or complements the heat shock inducible AAA+ ATPases.

REGULATION OF sHSP EXPRESSION The regulation of expression of sHSPs and α-crystallins has been well characterized in organisms from all three domains of life. Bacterial sHSPs such as those from a thermophilic cyanobacterium, Synechococcus vulcanus and a hyperthermophile, P. furiosus were not expressed under normal

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growth conditions [29,35–37]. In contrast, in mice, α-crystallin and HSP25 were highly expressed in several organs such as the developing eye lenses, heart, stomach, and lung under nonstress conditions [38,39]. Depending on the organism, sHsp expression control appears to be at transcriptional or translational levels. In transcriptional control, both positive and negative modes of regulation of shsp gene expression have been reported. In E. coli, the sHSP genes, ibpA and ibpB, form an operon and are positively controlled by δ32, the sigma factor that regulates multiple stress responsive genes [40]. However, an alternative mode of regulation has been suggested as there is some accumulation of the IbpA and IbpB proteins, even in a δ32-defective mutant of E. coli [41]. In several rhizobia, translational control of shsp gene expression has been reported. Repression of heat shock gene expression (ROSE) is a novel type of a regulatory system that functions by cis-acting hairpin element positioned within the 5′-untranslated region of the mRNA. The ribosome-binding site is sequestered by formation of the stem-loop from the 3′ region of ROSE and induction follows the disruption of the secondary structure by elevated temperature [42,43]. The thermophilic cyanobacterium S. vulcanus accumulates several HSPs including sHSP, GroEL, and GroES when cells are exposed to heat shock from 50°C to 63°C [35]. Unlike expression of other hsps, the regulation of shsp expression of cyanobacteria is not under the control of CIRCE (Controlling Inverted Repeat of Chaperone Expression) [36,44,45]. The putative DNA-binding protein appears to bind the DNA in this region more efficiently in nonheat shock than in heat shock conditions [44]. This result indicates that the shsp expression in thermophilic cyanobacteria may be similar to that found in Streptococcus albus. In addition, the recent study of HspA expression in the thermophilic cyanobacterium Synechococcus vulcanus suggested that the expression is also under translational control. The experiments were done by inserting an hspA gene into the lacZ gene with an inducible lac promoter and the constructs were transformed into E. coli as a surrogate system. The expression of HspA is enhanced significantly at 42°C compared with that 30°C as a result of thermally altered mRNA structure [46]. Hyperthermophilic archaea such as P. furiosus and Thermococcus KS-1 have single copy of sHSP that is significantly induced at heat shock temperatures [29,47]. In addition, sHsp of P. furiosus cells that were recovered at normal growth temperature (95°C) after heat shock (105°C) is rapidly degraded on return to growth permissive temperatures (Figure 5.1a). These results indicate that sHSP is not required at the optimal growth temperature even near 100°C. In P. furiosus, a 24 kDa putative heat shock regulator (Phr) and cis-acting regulatory sequence have been discovered [28]. The homologs of the Phr in P. horikoshii and P. abyssi are PH1744 and PAB0208, respectively. Double-stranded DNA is required as a binding substrate for Phr. The transcripts of the aaa+ ATPase and hsp20 genes are induced by heat shock [29]. The phr-regulated promoters of hsp20 (Pfu-shsp) and aaa+ ATPase show highly conserved regions. The Phr regulator regulates the expression of these genes negatively as it can inhibit the formation of mRNA polymerase complex, although the mechanism of derepression under heat shock conditions remains unknown. In A. fulgidus, AF1298 which is located upstream of the sHsp and cdc48 genes appears to have a cis-binding element that binds to HSR1, a product of AF1298 gene itself. It was suggested that HSR1 and Phr may be members of a very diverse protein family of archaeal repressors [33]. Temperature upshift is not the only factor that can induce HSP induction. We have shown in P. furiosus that when a mid-exponential phase (3 h) at 95°C was spiked with 5% ethanol, sHsp production was induced and sustained during growth (Figure 5.1b). Supplying 10% ethanol appears to inhibit growth and sHSP production completely (data not shown). We hypothesize that the sHsp may be significantly induced in response to unfolded proteins in cells, similar to the unfolded protein response in eukarya.

MECHANISTIC INSIGHTS FROM MINIMAL COMPLEXITY The chaperonins are ubiquitous molecular chaperones that form double-ring assemblies of subunits with a molecular mass of 60–70 kDa. The resulting structures have a large central cavity where

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FIGURE 5.1 Regulation of small heat shock protein (sHSP) in the hyperthermophile Pyrococcus furiosus. Evidence for the presence of an unfolded protein response. P. furiosus was grown at 95°C for 4 h. Absolute ethanol previously degassed to remove residual oxygen was added to P. furiosus cultures to the final concentration of 5%. Control is the culture without addition of ethanol. The cultures were further incubated at 95°C for 2 h. The cultures were then subjected to sodiumdodecylsulfate-polyacrylamide gel electrophoresis (SDSPAGE) and western blot analysis using Pf-Cpn and Pf-sHsp antibodies. (a) Heat shock induction and rapid turnover of sHSP following restoration of the culture to growth permissive temperatures. Lane 1, P. furiosus culture at 95°C in 20-l fermenter; lane 2, P. furiosus heat shocked at 105°C for 2 h; lanes 3 and 4, P. furiosus culture recovered at 95°C for 1 and 2 h, respectively, after heat shock. (b) Western blot visualization of the sHSP from P. furiosus expressed at a growth permissive temperature, 95°C, in the presence of ethanol. Lanes 1 and 2, Control without ethanol, 1 and 5 μl of extract loaded, respectively.

non-native proteins can undergo productive folding in an ATP-dependent manner [48,49]. The paradigm for chaperonin-assisted protein folding has been the group I GroEL/GroES system from E. coli, consisting of the GroEL chaperonin and associated GroES co-chaperonin, which are characteristic in bacteria and eukaryal organelles of bacterial origin [49,50].

STRUCTURE AND SUBUNIT COMPOSITION OF ARCHAEAL GROUP II CHAPERONINS The chaperonins form toroidal double rings with an eight- or nine-fold symmetry, consisting of homologous subunits [51]. The archaeal group II chaperonins are composed of up to five sequencerelated subunits. Sulfolobus species [52,53], Haloferax volcanii [54], Methanosarcina mazei [30], and M. burtonii [21, R. Cavicchioli, personal communication] contain three chaperonin genes. Table 5.2 lists the number of subunits per genome and subunit composition of chaperonins from characterized members of the archaea. Recently, it was found that there are five chaperonin subunits (Hsp60-1, -2, -3, -4, and -5) in Methanosarcina acetivorans. Among them, Hsp60-1, Hsp60-2, and Hsp60-3 have orthologs in Methanosarcinacea, but others, Hsp60-4 and Hsp60-5, occur only in M. acetivorans. Subunit composition is summarized in Table 5.3. The subunit composition of the chaperonin complexes in several archaea changes with growth temperature [53,55]. The chaperonin from the hyperthermophilic archaeon, Thermococcus sp. strain KS-1 (T. KS-1) is composed of two highly sequence-related subunits, α and β [56], that form a hetero-oligomer with variable subunit composition in vivo [55]. Expression of α- and β-subunits is regulated differently, and only the α-subunit is thermally inducible [55]. The proportion of the α-subunit in T. KS-1 chaperonin increases with temperature, and the β-subunit-rich chaperonin is more thermostable than the α-subunit rich-chaperonin [57]. In the hyperthermoacidophilic archaeon, S. shibatae, group II chaperonins encode three different subunits (α, β, and γ). Expression of the α- and β-subunits is increased by heat shock, and decreased by cold shock [53]. On the other hand,

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TABLE 5.2 Archaeal Chaperonin: Number of Subunits Encoded per Genome Organisms

Subunit Species

Rotational Symmetry

References

Crenarchaeota Aeropyrum pernix

2 (α, β)

NR

[104]

Pyrobaculum aerophilum

2 (α, β)

NR

[105,106]

Pyrodictium occultum

2 (α, β)

Sulfolobus acidocaldarius Sulfolobus shibatae

8

[25]

3 (α, β, γ)

NR

[52]

3 (α, β, γ)

9

Sulfobus solfataricus

3 (α, β, γ)

9

Sulfolobus tokodaii

3 (α, β, γ)

NR

[52,106] [53,107] [52]

Euryarchaeota Archaeoglobus fulgidus

2 (α, β)

Halobacterium sp. NRC-1

2 (α, β) 3 (CCT1, 2, 3) 1 1 1 1 I: 1 II: 5 (Hsp60-1, -2, -3, -4, -5) I: 1 II: 3 (1, 2, 3) I: 1 II: 3 (α, β, γ)

Haloferax volcanii Methanocaldococcus jannaschii Methanococcus thermolithotrophicus Methanococcus maripaludis Methanopyrus kandleri Methanosarcina acetivorans Methanosarcina barkeri Methanosarcina maeii Methanothermobacter thermautorophicus Picrophilus torridus Pyrococcus abyssi Pyrococcus furiosus Pyrococcus horikoshii Thermoplasma acidophilum

2 (α, β) 2 1 1 1

8

[32,106]

NR

[108]

NR NR 8 NR 8 NR NR NR NR NR 8? NR

[54] [109] [96] [97,110] [105,111] [21]

[112] [3] http://www.genoscope.cns.fr/Pab/ [113] [67] [27,114]

[21] [30]

2 (α, β)

NR NR NR NR 8

Thermoplasma volcanium

2 (α, β)

NR

[115]

Thermococcus kodakaraensis

2 (α, β)

NR

[116,117]

Thermococcus sp. strain KS-1

2 (α, β)

8

[56,57,100,118]

M. acetivorans, M. barkeri, and M. mazi contain both group I (refer to “I”) and group II (refer to “II”) chaperonins. CCT, chaperonin-containing t-complex polypeptide-1; NR, not reported.

expression of the γ-subunit gene is undetectable at heat shock temperatures and low at normal growth conditions, but induced by cold shock [53]. The halophilic archaeon H. volcanii has three group II chaperonins genes, cct1, cct2, and cct3, which are expressed constitutively but to differing levels [54]. Interestingly, deletion of cct3 has no effect on the activity of the chaperonin complex, but loss of cct1 leads to ~50% reduction in the purified chaperonin ATPase activity [18]. The precise functional properties and physiological significance of the heterologous subunit composition of archaeal group II chaperonin subunits is still the subject of active investigation. The crystal structure of the group II chaperonin is shown in Figure 5.2. This structure from the thermoacidophilic archaeon, Thermoplasma acidophilum, has shown that the subunit architectures

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FIGURE 5.2 The crystal structure of two adjacent subunits of the class II chaperonin from Thermoplasma acidophilum (from PDB: 1A6D). Functional domains are labeled. The figure was drawn with the threedimensional molecular viewer in the VectorNTI 10.0 package.

are very similar to group I chaperonins, except for differences in the helical protrusion region [58–60]. It is likely that the helical protrusion in group II chaperonins provides an equivalent functional role to the GroES subunit of group I chaperonins, by acting as a closure for the central cavity of the chaperonin complex [61–63] (Figure 5.2).

PREFOLDINS Prefoldins are universally present in eukarya and archaea, with similar structures, but are absent in bacteria. The prefoldins are “holdase” chaperones whose crystal structure was first resolved from the archaeon, Methanothermobacter thermoautotrophicum [64,65]. The chaperone has been likened to a jelly-fish in shape, with a globular “body” with six canonical, antiparallel coiled coils (the “tentacles”) with their N- and C-domains oriented outwardly from an oligomerization domain. The coiled-coil “tentacles” enclose a cavity lined with hydrophobic patches that clamp non-native target proteins [66]. The holding-and-release mechanism of the archaeal prefoldins has recently been elucidated [67,68]. In the archaea, with one exception, prefoldins are hexamers consisting of two α-subunits and four β-subunits, which act as generalized holding chaperones. The archaeal prefoldins bind to a wide range of non-native proteins in vitro, although their intracellular substrates are not known. Although similar in overall structure, the eukaryal prefoldins consist of six nonidentical subunits (two α-class and four β-class subunits) and in contrast to archaeal prefoldins, bind specifically to the ribosome-nascent forms of actins and tubulins [69]. Several recent lines of evidence indicate that prefoldins can act cooperatively with chaperonins, such as HSP60, and load non-native proteins into their cavity. The prefoldin tentacles are capable of flexing outwards to accommodate both small (14 kDa, lysozyme) and large (62 kDa, firefly luciferase) proteins in the cavity formed by the “tentacles” to prevent their aggregation [66,69]. In addition, the species-specific transfer of non-native substrates to chaperonins has been followed using surface plasmon resonance. Transfer takes place between the prefoldins and chaperonins from one species

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TABLE 5.3 Structural and Functional Characteristics of Archaeal Group II Chaperonins Subunit Species

Native or Recombinant

Rotational ATPase Arrest Symmetry Activity Activity*

Folding Activity

Sulfolobus shibatae Sulfobus solfataricus Sulfolobus tokodaii

3 3 3

9 9 NR NR

Trace Trace Trace Trace

+ + + +

NR + – –

[24,53] [52,94,95,107,119] [52,120]

Pyrodictium brockii Pyrodictium occultum

NR 2

Native Native Native Recombinant (α, β)† Native Native Recombinant (α, β) Recombinant (α + β)¶

8 8 8 8

NR + + +

NR NR + +

NR NR NR NR

[106] [105,106]

NR +

NR NR +

NR NR –§

[32,106] [54] [109]

Organism

References

Crenarchaeota

Euryarchaeota Archaeoglobus fulgidus Haloferax volcanii Methanococcus jannaschii Methanococcus thermolithotrophicus Methanococcus maripaludis Methanopyrus kandleri

2 3 1

Native Native Native

8 NR NR

1

Recombinant**

8

+

+

+

[96]

1

Recombinant

NR

+

+

+

[97,110]

1 3

NR + NR + (αβγ)¶¶

NR NR NR – (αβγ)

Pyrococcus horikoshii Thermoplasma acidophilum

1 2

8 + 8 + NR 8 (α:β: γ = 2:1:1) + 8 (αγ)‡ NR + 8 Trace 8 + 8 +

[111,121]

Methanosarcina mazei

Native Recombinant Native Recombinant

+ + NR Trace

+ NR NR Trace

[67] [27,98,114]

Thermococcus kodakaraensis Thermococcus sp. strain KS-1

2

NR

+

+ (β)§§

NR

[116,117]

NR 8

+ +

+ +

[56,100]

2

Recombinant Native Recombinant (α, β) Recombinant (α + β) Recombinant (α, β) Native Recombinant (α, β)

+ +

[30]

* “Arrest activity” means the binding activity to non-native proteins. † α- and β-subunits are separately expressed in Escherichia coli and purified. ¶ α- and β-subunits are co-expressed in E. coli and purified. § The measurement is carried out at 30°C. ** Reconstituted complex of the purified subunit. ‡ Reconstituted complex of α- and γ-subunits. ¶ ¶ Reconstituted complex of α-, β-, and γ-subunits. §§ Purified β-subunit prevents thermal inactivation of yeast alcohol dehydrogenase. NR, not reported.

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of Pyrococcus, but not when chaperones cloned and expressed from different Pyrococcus species are used [67]. The hyperthermophilic methanogen, M. jannaschii, encodes genes for the α- and β-subunits of prefoldin. However, a unique third prefoldin subunit is encoded by the pfdγ gene, and is heat shock-regulated, unlike the α- and β-subunits [31,70,71]. This system opens new questions regarding the functional assignments of this heat shock-inducible prefoldin and the sHSPs, as they appear to have overlapping chaperone activities in vitro.

SMALL HEAT SHOCK PROTEINS The sHSPs and α-crystallins have a monomeric molecular weight range of 15–40 kDa, and typically form polydisperse multimeric complexes in vivo. Putative shsp genes are present in all archaeal genome sequences including N. equitans. Both the sHSPs and vertebrate α-crystallins are holdase-type molecular chaperones [70–74]. However, among thermophiles, biochemical characterization is limited to thermophilic and hyperthermophilic organisms. Only two crystal structures of sHSPs from unrelated organisms, M. jannaschii [75] and Triticum aestivum (wheat) [76] have been reported. The sHSPs share amino acid sequence similarity with the central core of vertebrate eye lens α-crystallin proteins, which are conserved in this family of proteins through all domains of life. The sHSP proteins have relatively low amino acid sequence similarity and their quaternary structures are dissimilar. However, the monomeric structures of these proteins are almost identical. Their specific functional mechanisms may be determined by their individual quaternary structures and their cognate target proteins and chaperone partners. The archaeal sHSPs can prevent denatured proteins from aggregating under strong denaturing conditions, and in some cases, are able to refold denatured proteins [47,77,78]. The sequences of the N- and C-terminal domains of archaeal sHSPs differ, and this variability is responsible for the great variety of multisubunit structures that are formed. Although the N-terminal domain of the M. jannaschii sHSP16.5 is disordered in the crystal structure, low resolution features have been resolved by cryoelectron microscopy. This hydrophobic domain is essential for proper holdase function in sHSP16.5 [79]. The copy number of sHSP-encoding genes is variable among archaeal species. The thermophilic and hyperthermophilic archaea contain one, two, or three shsp homologs. Hyperthermophilic species growing optimally near 100°C have one shsp gene with the exception of P. aerophilum which has two homologs [29]. T. acidophilum and all these Sulfolobus species. represented by genome sequences each have three shsp homologs. However, one of the sHSPs in T. acidophilum appears to have domains that are similar to the two ATPase domains of ArsA from E. coli [80]. S. solfataricus and Sulfolobus tokodaii have one 14–15-kDa and two 20–21-kDa sHSPs each. The mesophilic methanogens, M. acetivorans and M. mazei GoE1 contain three and four shsp homologs, respectively. However, one of the two sHSPs from M. acetivorans (NP_619401) does not appear to belong to the α-crystallin-type HSPs. The genome sequence of Halobacterium NRC-1 has the highest paralogy among the archaea, encoding five sHSPs that all clearly belong to the α-crystallin family. It seems likely that the multiple sHSPs encoded in a single species perform a range of potentially overlapping cellular functions; however, this has not been experimentally assessed. The role of sHSPs in protein folding is still a topic of active investigation in both archaea and eukarya. They can maintain solubility of non-native proteins under physiological conditions indefinitely, for example in the eye lens, displaying a remarkable capacity for binding non-native target proteins present in greater concentration than the chaperones themselves. The binding capacity of eukaryal α, β crystallins for non-native proteins is greatly stimulated by serine phosphorylation of the sHSP, and the dynamic reordering of sHSP complexes is required for solubilization of non-native proteins [81]. Although archaeal systems for protein phosphorylation have been described (see Chapter 19), it is unknown whether archaeal chaperones are phosphorylated. Recently, reconstitution of a protein-refolding pathway in vitro was described [82,83]. Denatured Taq polymerase was reactivated cooperatively at 100°C by a mixture of sHSP or prefoldin with HSP60 from P. furiosus, in an ATP-dependent folding pathway. The cooperative protein salvage

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pathway is dependent on the presence of HSP60 and ATP for full activity. The sHSPs and prefoldins appear to fulfill similar roles in this system, namely to transfer denatured proteins to the HSP60 chaperonin, although the differences in their structures suggest that they transfer non-native proteins to chaperonins by different mechanisms. The rate of refolding of Taq polymerase was minimal when just the holdase chaperones were present, and was greatly increased when HSP60 and ATPMg2+ were added.

NASCENT-ASSOCIATED PROTEINS The nascent polypeptide-associated complex (NAC) was first isolated from bovine brain cytosol and recognized as an essential molecular chaperone. Multiple subunits of NAC were first characterized in yeast, and formation of NAC complexes with ribosomes appears to be critical for folding and export of eukaryal proteins [84]. NACs are found only in eukaryotes and archaea, however, unlike eukaryal NAC systems which are composed of α- and β-subunits, the archaeal NAC systems contain only an α-subunit [85]. Based on the complete archaeal genome sequences, all thermophilic archaea appear to have a single copy of NAC protein except that of N. equitans, a symbiotic archaeon with the smallest genome size where it is absent [86–89]. This suggests that NAC proteins may play significant roles in protein folding in thermophiles. The mechanisms of formation and dissolution of eukaryal NAC/polypeptide complexes is not well understood, although several hypotheses have been proposed, and studies are ongoing. The hypothesis that NAC proteins prevent inappropriate interaction between newly synthesized polypeptide chains and other cellular factors appears to be well supported. NAC functions in archaea may be similar to bacterial ribosme-associated chaperone trigger factor (TF), as TF homologs are absent from all archaeal genomes. Currently, a thermophilic TF has been characterized from a thermophilic bacterium, Thermus thermophilus. Surprisingly, the chaperone activity of T. thermophilus TF appears to be Zn2+-dependent [90]. Functions of NAC have been studied in eukaryotes suggesting that NAC acts on polypeptides nascent on the ribosome [85]. However, alternative roles of NAC have been reported. For instance, NAC proteins have been shown to be involved in translational control [91] and localization of Oskar mRNA [92]. The crystal structure of the archaeal NAC from a thermophilic methanogen, Methanothermobacter thermautotrophicus, revealed that the NAC subunit consists of two domains, the NAC domain and the ubiquitin-associated (UBA) domain (Figure 5.3) [93]. This in vitro functional analysis of archaeal NAC revealed that the protein is associated with ribosomes and also in contact with nascent chains on the ribosome [93]. At present, the hypothesis that the complex interacts with ubiquitin is speculative [93]. Putative ubiquitin homologs occur in several archaeal genomes but are missing from several others, and consequently the UBA domain, which is strongly conserved in all archaeal genomes, may have functions unrelated to ubiquitin binding.

PROTEIN-FOLDING MECHANISM OF ARCHAEAL GROUP II CHAPERONINS While nucleotide and amino acid sequences of many archaeal chaperonins have been reported, there are comparatively few reports on their functional characterization; these include the native chaperonin from S. solfataricus [94,95] and Thermococcus KS-1 [55], and recombinant chaperonins from Methanococcus thermolithotrophicus [67,96] P. horikoshii [67], Methanococcus maripaludis [97], T. acidophilum [56,98,99], and T. KS-1 [56]. The group II chaperonin from T. KS-1 has been studied in most detail. The T. KS-1, α- and β-subunits coassemble to form double-ring homo-oligomers (α- and β-chaperonins, respectively), and are able to capture denatured proteins and fold them in an ATP-dependent manner in vitro [56,100]. Taking advantage of this, significant progress has been made in defining functional mechanisms which have been compared to the operation of a two-stroke internal combustion motor [62,100,101].

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FIGURE 5.3 (See color insert following page 178.) The crystal structure of homodimeric nascent polypeptide associated complex (NAC) from Methanobacterium thermoautotrophicum (from PDB: 1TR8). NAC and ubiquitin-associated domain (UBA) domains are labeled. The figure was drawn with the three-dimensional molecular viewer in the VectorNTI 10.0 package.

PERSPECTIVES: NEXT FIVE YEARS Molecular chaperones are diverse and eclectic, and every species encodes a unique repertoire of different independent or cooperative protein-folding pathways. In archaea, stress-inducible chaperones have received the most attention thus far. Perhaps the most understandable cellular functions of chaperones occur during cell stress by salvaging non-native proteins and recruiting them to join the pool of stable proteins, to prevent their demise as intracellular aggregates. In the eukarya and bacteria, chaperones are also known to participate in many fundamental cellular processes in nonstressed cells including DNA replication, regulation of gene expression, cell division, membrane translocation, protein folding, and protein remodeling [102]. For example, the ClpA proteins in bacteria can mediate protein folding, unfolding, assembly, and disassembly without themselves being part of the final complex [103]. Open questions remain in the archaeal protein-folding systems, as to how post-translational modeling functions are partitioned between the known chaperones, and chaperones or co-chaperones that have not yet been discovered. The mechanisms of protein folding in more complex eukaryal pathways have become accessible through the analysis of chaperones from archaea, due to their simple architecture and exceptional stability. In the case of chaperonin and prefoldin, the compact nature of the protein-folding pathways in normal and stressed cells involves double duty assignments of the chaperonin which operates both under normal growth conditions, as well as carrying out the final step in stress-responsive protein-folding pathways [30]. New insights into post-translational processing and protein salvage will very likely emerge in the next five years as new developments, such as tractable genetic systems (see Chapters 11, 12, and 13), which are more widely applied to thermophiles.

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ACKNOWLEDGMENT The authors gratefully acknowledge grant support from the National Science Foundation, NSF MCB 98090352 and the US Air Force Office of Scientific Research FA9550-06-0020. This is contribution number 07-179 from the Center of Marine Biotechnology.

REFERENCES 1. Blochl, E. et al. Pyrolobus fumarii, gen. and sp. nov., represents a novel group of archaea, extending the upper temperature limit for life to 113°C. Extremophiles 1, 14–21 (1997). 2. Kashefi, K. and Lovley, D. R. Extending the upper temperature limit for life. Science 301, 934 (2003). 3. Futterer, O. et al. Genome sequence of Picrophilus torridus and its implications for life around pH 0. Proc Natl Acad Sci USA 101, 9091–6 (2004). 4. Robb, F. T. and Maeder, D. L. Novel evolutionary histories and adaptive features of proteins from hyperthermophiles. Curr Opin Biotechnol 9, 288–91 (1998). 5. Vetriani, C. et al. Protein thermostability above 100°C: a key role for ionic interactions. Proc Natl Acad Sci USA 95, 12300–5 (1998). 6. Kreil, D. P. and Ouzounis, C. A. Identification of thermophilic species by the amino acid compositions deduced from their genomes. Nucleic Acids Res 29, 1608–15 (2001). 7. Suhre, K. and Claverie, J. M. Genomic correlates of hyperthermostability, an update. J Biol Chem 278, 17198–202 (2003). 8. Macedo-Ribeiro, S., Darimont, B., and Sterner, R. Structural features correlated with the extreme thermostability of 1[4Fe-4S] ferredoxin from the hyperthermophilic bacterium Thermotoga maritima. Biol Chem 378, 331–6 (1997). 9. Haney, P. J. et al. Thermal adaptation analyzed by comparison of protein sequences from mesophilic and extremely thermophilic Methanococcus species. Proc Natl Acad Sci USA 96, 3578–83 (1999). 10. Elcock, A. H. The stability of salt bridges at high temperatures: implications for hyperthermophilic proteins. J Mol Biol 284, 489–502 (1998). 11. Elcock, A. H. Prediction of functionally important residues based solely on the computed energetics of protein structure. J Mol Biol 312, 885–96 (2001). 12. Elcock, A. H. and McCammon, J. A. Electrostatic contributions to the stability of halophilic proteins. J Mol Biol 280, 731–48 (1998). 13. Ueda, K. et al. Genome sequence of Symbiobacterium thermophilum, an uncultivable bacterium that depends on microbial commensalism. Nucleic Acids Res 32, 4937–44 (2004). 14. Laksanalamai, P., Whitehead, T. A., and Robb, F. T. Minimal protein-folding systems in hyperthermophilic archaea. Nat Rev Microbiol 2, 315–24 (2004). 15. Waters, E. et al. The genome of Nanoarchaeum equitans: insights into early archaeal evolution and derived parasitism. Proc Natl Acad Sci USA 100, 12984–8 (2003). 16. Macario, A. J. and Conway De Macario, E. The molecular chaperone system and other anti-stress mechanisms in archaea. Front Biosci 6, D262–83 (2001). 17. Conway de Macario, E., Maeder, D. L., and Macario, A. J. Breaking the mould: archaea with all four chaperoning systems. Biochem Biophys Res Commun 301, 811–12 (2003). 18. Lund, P. A., Large, A. T., and Kapatai, G. The chaperonins: perspectives from the archaea. Biochem Soc Trans 31, 681–5 (2003). 19. Galagan, J. E. et al. The genome of M. acetivorans reveals extensive metabolic and physiological diversity. Genome Res 12, 532–42 (2002). 20. Robb, F. T. Genomics of thermophiles, in Microbial Genomics, Fraser, C. M., Nelson, K. E., and Read, T., Eds., Humana Press, pp. 245–268 (2004). 21. Maeder, D. L., Macario, A. J., and de Macario, E. C. Novel chaperonins in a prokaryote. J Mol Evol 60, 409–16 (2005). 22. Squires, C. L., Pedersen, S., Ross, B. M., and Squires, C. ClpB is the Escherichia coli heat shock protein F84.1. J Bacteriol 173, 4254–62 (1991).

9214_C005.indd 66

11/5/2007 4:19:08 PM

Thermophilic Protein-Folding Systems

67

23. Trent, J. D., Gabrielsen, M., Jensen, B., Neuhard, J., and Olsen, J. Acquired thermotolerance and heat shock proteins in thermophiles from the three phylogenetic domains. J Bacteriol 176, 6148–52 (1994). 24. Trent, J. D., Nimmesgern, E., Wall, J. S., Hartl, F. U., and Horwich, A. L. A molecular chaperone from a thermophilic archaebacterium is related to the eukaryotic protein t-complex polypeptide-1. Nature 354, 490–3 (1991). 25. Kagawa, H. K. et al. The 60 kDa heat shock proteins in the hyperthermophilic archaeon Sulfolobus shibatae. J Mol Biol 253, 712–25 (1995). 26. Quaite-Randall, E., Trent, J. D., Josephs, R., and Joachimiak, A. Conformational cycle of the archaeosome, a TCP1-like chaperonin from Sulfolobus shibatae. J Biol Chem 270, 28818–23 (1995). 27. Waldmann, T., Lupas, A., Kellermann, J., Peters, J., and Baumeister, W. Primary structure of the thermosome from Thermoplasma acidophilum. Biol Chem Hoppe Seyler 376, 119–26 (1995). 28. Vierke, G., Engelmann, A., Hebbeln, C., and Thomm, M. A novel archaeal transcriptional regulator of heat shock response. J Biol Chem 14, 14 (2002). 29. Laksanalamai, P., Maeder, D. L., and Robb, F. T. Regulation and mechanism of action of the small heat shock protein from the hyperthermophilic archaeon Pyrococcus furiosus. J Bacteriol 183, 5198–202 (2001). 30. Klunker, D. et al. Coexistence of group I and group II chaperonins in the archaeon Methanosarcina mazei. J Biol Chem 278, 33256–67 (2003). 31. Boonyaratanakornkit, B. B. et al. Transcriptional profiling of the hyperthermophilic methanarchaeon Methanococcus jannaschii in response to lethal heat and non-lethal cold shock. Environ Microbiol 7, 789–97 (2005). 32. Emmerhoff, O. J., Klenk, H. P., and Birkeland, N. K. Characterization and sequence comparison of temperature-regulated chaperonins from the hyperthermophilic archaeon Archaeoglobus fulgidus. Gene 215, 431–8 (1998). 33. Rohlin, L. et al. Heat shock response of Archaeoglobus fulgidus. J Bacteriol 187, 6046–57 (2005). 34. Shockley, K. R. et al. Heat shock response by the hyperthermophilic archaeon Pyrococcus furiosus. Appl Environ Microbiol 69, 2365–71 (2003). 35. Roy, S. K., Hiyama, T., and Nakamoto, H. Purification and characterization of the 16-kDa heat-shockresponsive protein from the thermophilic cyanobacterium Synechococcus vulcanus, which is an alphacrystallin-related, small heat shock protein. Eur J Biochem 262, 406–16 (1999). 36. Roy, S. K. and Nakamoto, H. Cloning, characterization, and transcriptional analysis of a gene encoding an alpha-crystallin-related, small heat shock protein from the thermophilic cyanobacterium Synechococcus vulcanus. J Bacteriol 180, 3997–4001 (1998). 37. Sauer, U. and Durre, P. Sequence and molecular characterization of a DNA region encoding a small heat shock protein of Clostridium acetobutylicum. J Bacteriol 175, 3394–400 (1993). 38. Klemenz, R., Andres, A. C., Frohli, E., Schafer, R., and Aoyama, A. Expression of the murine small heat shock proteins hsp25 and alpha B crystallin in the absence of stress. J Cell Biol 120, 639–45 (1993). 39. Klemenz, R., Frohli, E., Steiger, R. H., Schafer, R., and Aoyama, A. Alpha B-crystallin is a small heat shock protein. Proc Natl Acad Sci USA 88, 3652–6 (1991). 40. Allen, S. P., Polazzi, J. O., Gierse, J. K., and Easton, A. M. Two novel heat shock genes encoding proteins produced in response to heterologous protein expression in Escherichia coli. J Bacteriol 174, 6938–47 (1992). 41. Laskowska, E., Wawrzynow, A., and Taylor, A. IbpA and IbpB, the new heat-shock proteins, bind to endogenous Escherichia coli proteins aggregated intracellularly by heat shock. Biochimie 78, 117–22 (1996). 42. Nocker, A. et al. A mRNA-based thermosensor controls expression of rhizobial heat shock genes. Nucleic Acids Res 29, 4800–7 (2001). 43. Nocker, A., Krstulovic, N. P., Perret, X., and Narberhaus, F. ROSE elements occur in disparate rhizobia and are functionally interchangeable between species. Arch Microbiol 176, 44–51 (2001). 44. Kojima, K. and Nakamoto, H. Specific binding of a protein to a novel DNA element in the cyanobacterial small heat-shock protein gene. Biochem Biophys Res Commun 297, 616 (2002). 45. Narberhaus, F. Negative regulation of bacterial heat shock genes. Mol Microbiol 31, 1–8 (1999).

9214_C005.indd 67

11/5/2007 4:19:08 PM

68

Thermophiles: Biology and Technology at High Temperatures

46. Kojima, K. and Nakamoto, H. Post-transcriptional control of the cyanobacterial hspA heat-shock induction. Biochem Biophys Res Commun 331, 583–8 (2005). 47. Usui, K., Yoshida, T., Maruyama, T., and Yohda, M. Small heat shock protein of a hyperthermophilic archaeum, Thermococcus sp. strain KS-1, exists as a spherical 24 mer and its expression is highly induced under heat-stress conditions. J Biosci Bioeng 92, 161–6 (2001). 48. Bukau, B. and Horwich, A. L. The Hsp70 and Hsp60 chaperone machines. Cell 92, 351–66 (1998). 49. Hartl, F. U. and Hayer-Hartl, M. Molecular chaperones in the cytosol: from nascent chain to folded protein. Science 295, 1852–8 (2002). 50. Sigler, P. B. and Horwich, A. L. Unliganded GroEL at 2.8 A: structure and functional implications. Philos Trans R Soc Lond B Biol Sci 348, 113–19 (1995). 51. Gutsche, I., Essen, L. O., and Baumeister, W. Group II chaperonins: new TRiC(k)s and turns of a protein folding machine. J Mol Biol 293, 295–312 (1999). 52. Archibald, J. M., Logsdon, J. M., and Doolittle, W. F. Recurrent paralogy in the evolution of archaeal chaperonins. Curr Biol 9, 1053–6 (1999). 53. Kagawa, H. K. et al. The composition, structure, and stability of a group II chaperonin is temperature regulated in a hyperthermophilic archaeon. Mol Microbiol (2003). 54. Large, A. T., Kovacs, E., and Lund, P. A. Properties of the chaperonin complex from the halophilic archaeon Haloferax volcanii. FEBS Lett 532, 309–12 (2002). 55. Yoshida, T., Ideno, A., Hiyamuta, S., Yohda, M., and Maruyama, T. Natural chaperonin of the hyperthermophilic archaeum, Thermococcus strain KS-1: a hetero-oligomeric chaperonin with variable subunit composition. Mol Microbiol 39, 1406–13 (2001). 56. Yoshida, T. et al. Structural and functional characterization of homo-oligomeric complexes of alpha and beta chaperonin subunits from the hyperthermophilic archaeum Thermococcus strain KS-1. J Mol Biol 273, 635–45 (1997). 57. Yoshida, T., Ideno, A., Suzuki, R., Yohda, M., and Maruyama, T. Two kinds of archaeal group II chaperonin subunits with different thermostability in Thermococcus strain KS-1. Mol Microbiol 44, 761–9 (2002). 58. Braig, K. Chaperonins. Curr Opin Struct Biol 8, 159–65 (1998). 59. Ditzel, L. et al. Crystal structure of the thermosome, the archaeal chaperonin and homolog of CCT. Cell 93, 125–38 (1998). 60. Klumpp, M. and Baumeister, W. The thermosome: archetype of group II chaperonins. FEBS Lett 430, 73–7 (1998). 61. Llorca, O. et al. Analysis of the interaction between the eukaryotic chaperonin CCT and its substrates actin and tubulin. J Struct Biol 135, 205–18 (2001). 62. Iizuka, R. et al. Role of the helical protrusion in the conformational change and molecular chaperone activity of the archaeal group II chaperonin. J Biol Chem 279, 18834–9 (2004). 63. Meyer, A. S. et al. Closing the folding chamber of the eukaryotic chaperonin requires the transition state of ATP hydrolysis. Cell 113, 369–81 (2003). 64. Siegert, R., Leroux, M. R., Scheufler, C., Hartl, F. U., and Moarefi, I. Structure of the molecular chaperone prefoldin: unique interaction of multiple coiled coil tentacles with unfolded proteins. Cell 103, 621–32 (2000). 65. Stirling, P. C., Lundin, V. F., and Leroux, M. R. Getting a grip on non-native proteins. EMBO Rep 4, 565–70 (2003). 66. Lundin, V. F. et al. Molecular clamp mechanism of substrate binding by hydrophobic coiled-coil residues of the archaeal chaperone prefoldin. Proc Natl Acad Sci USA 101, 4367–72 (2004). 67. Okochi, M., Matsuzaki, H., Nomura, T., Ishii, N., and Yohda, M. Molecular characterization of the group II chaperonin from the hyperthermophilic archaeum Pyrococcus horikoshii OT3. Extremophiles 9, 127–34 (2005). 68. Zako, T. et al. Facilitated release of substrate protein from prefoldin by chaperonin. FEBS Lett 579, 3718–24 (2005). 69. Leroux, M. R. et al. MtGimC, a novel archaeal chaperone related to the eukaryotic chaperonin cofactor GimC/prefoldin. Embo J 18, 6730–43 (1999). 70. Horwitz, J. Alpha-crystallin can function as a molecular chaperone. Proc Natl Acad Sci USA 89, 10449–53 (1992).

9214_C005.indd 68

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Thermophilic Protein-Folding Systems

69

71. Jacob, U., Gaestel, M., Katrin, E., and Buchner, J. Small heat shock proteins are molecular chaperones. J Biol Chem 268, 1517–20 (1993). 72. Muchowski, P. J., Hays, L. G., Yates, J. R., 3rd, and Clark, J. I. ATP and the core “alpha-crystallin” domain of the small heat-shock protein alphaB-crystallin. J Biol Chem 274, 30190–5 (1999). 73. Muchowski, P. J., Valdez, M. M., and Clark, J. I. AlphaB-crystallin selectively targets intermediate filament proteins during thermal stress. Invest Ophthalmol Vis Sci 40, 951–8 (1999). 74. Muchowski, P. J., Wu, G. J., Liang, J. J., Adman, E. T., and Clark, J. I. Site-directed mutations within the core ”alpha-crystallin” domain of the small heat-shock protein, human alphaB-crystallin, decrease molecular chaperone functions. J Mol Biol 289, 397–411 (1999). 75. Kim, K. K., Kim, R., and Kim, S. H. Crystal structure of a small heat-shock protein. Nature 394, 595–9 (1998). 76. van Montfort, R. L., Basha, E., Friedrich, K. L., Slingsby, C., and Vierling, E. Crystal structure and assembly of a eukaryotic small heat shock protein. Nat Struct Biol 8, 1025–30 (2001). 77. Kim, R., Kim, K. K., Yokota, H., and Kim, S. H. Small heat shock protein of Methanococcus jannaschii, a hyperthermophile. Proc Natl Acad Sci USA 95, 9129–33 (1998). 78. Laksanalamai, P., Jiemjit, A., Bu, Z., Maeder, D. L., and Robb, F. T. Multi-subunit assembly of the Pyrococcus furiosus small heat shock protein is essential for cellular protection at high temperature. Extremophiles 7, 79–83 (2003). 79. Kim, R. et al. On the mechanism of chaperone activity of the small heat-shock protein of Methanococcus jannaschii. Proc Natl Acad Sci USA 100, 8151–5 (2003). 80. Ruepp, A., Rockel, B., Gutsche, I., Baumeister, W., and Lupas, A. N. The chaperones of the archaeon Thermoplasma acidophilum. J Struct Biol 135, 126–38 (2001). 81. Shashidharamurthy, R., Koteiche, H. A., Dong, J., and McHaourab, H. S. Mechanism of chaperone function in small heat shock proteins: dissociation of the HSP27 oligomer is required for recognition and binding of destabilized T4 lysozyme. J Biol Chem 280, 5281–9 (2005). 82. Laksanalamai, P., Pavlov, A. R., Slesarev, A. I., and Robb, F. T. Stabilization of Taq DNA polymerase at high temperature by protein folding pathways from a hyperthermophilic archaeon, Pyrococcus furiosus. Biotechnol Bioeng 93, 1–5 (2006). 83. Robb, F. T., Izuka, R., and Yohda, M. In Archaea, molecular and cellular biology (ed. Cavicchioli, R.) 209–23 (ASM Press, Washington D.C., 2007). 84. Reimann, B. et al. Initial characterization of the nascent polypeptide-associated complex in yeast. Yeast 15, 397–407 (1999). 85. Wegrzyn, R. D. and Deuerling, E. Molecular guardians for newborn proteins: ribosome-associated chaperones and their role in protein folding. Cell Mol Life Sci 62, 2727–38 (2005). 86. Huber, H. et al. A new phylum of archaea represented by a nanosized hyperthermophilic symbiont. Nature 417, 63–7 (2002). 87. Huber, H., Hohn, M. J., Stetter, K. O., and Rachel, R. The phylum Nanoarchaeota: present knowledge and future perspectives of a unique form of life. Res Microbiol 154, 165–71 (2003). 88. Thomson, N. R., Sebaihia, M., Cerdeno-Tarraga, A. M., Holden, M. T., and Parkhill, J. Shrinking genomics. Nat Rev Microbiol 2, 11 (2004). 89. Waters, E. et al. The genome of Nanoarchaeum equitans: insights into early archaeal evolution and derived parasitism. Proc Natl Acad Sci USA 100, 12984–8 (2003). 90. Suno, R., Taguchi, H., Masui, R., Odaka, M., and Yoshida, M. Trigger factor from Thermus thermophilus is a Zn2+-dependent chaperone. J Biol Chem 279, 6380–4 (2004). 91. Rospert, S., Dubaquie, Y., and Gautschi, M. Nascent-polypeptide-associated complex. Cell Mol Life Sci 59, 1632–9 (2002). 92. Braat, A. K., Yan, N., Arn, E., Harrison, D., and Macdonald, P. M. Localization-dependent Oskar protein accumulation; control after the initiation of translation. Dev Cell 7, 125–31 (2004). 93. Spreter, T., Pech, M., and Beatrix, B. The crystal structure of archaeal nascent polypeptide-associated complex (NAC) reveals a unique fold and the presence of a ubiquitin-associated domain. J Biol Chem 280, 15849–54 (2005). 94. Guagliardi, A., Cerchia, L., Bartolucci, S., and Rossi, M. The chaperonin from the archaeon Sulfolobus solfataricus promotes correct refolding and prevents thermal denaturation in vitro. Protein Sci 3, 1436–43 (1994).

9214_C005.indd 69

11/5/2007 4:19:09 PM

70

Thermophiles: Biology and Technology at High Temperatures

95. Guagliardi, A., Cerchia, L., and Rossi, M. Prevention of in vitro protein thermal aggregation by the Sulfolobus solfataricus chaperonin. Evidence for nonequivalent binding surfaces on the chaperonin molecule. J Biol Chem 270, 28126–32 (1995). 96. Furutani, M., Iida, T., Yoshida, T., and Maruyama, T. Group II chaperonin in a thermophilic methanogen, Methanococcus thermolithotrophicus. Chaperone activity and filament-forming ability. J Biol Chem 273, 28399–407 (1998). 97. Kusmierczyk, A. R. and Martin, J. Nucleotide-dependent protein folding in the type II chaperonin from the mesophilic archaeon Methanococcus maripaludis. Biochem J 371, 669–73 (2003). 98. Bigotti, M. G. and Clarke, A. R. Cooperativity in the thermosome. J Mol Biol 348, 13–26 (2005). 99. Kusmierczyk, A. R. and Martin, J. Chaperonins—keeping a lid on folding proteins. FEBS Lett 505, 343–7 (2001). 100. Yoshida, T. et al. Archaeal group II chaperonin mediates protein folding in the cis-cavity without a detachable GroES-like co-chaperonin. J Mol Biol 315, 73–85 (2002). 101. Iizuka, R. et al. ATP binding is critical for the conformational change from an open to closed state in archaeal group II chaperonin. J Biol Chem 278, 44959–65 (2003). 102. Wickner, S. et al. A molecular chaperone, ClpA, functions like DnaK and DnaJ. Proc Natl Acad Sci USA 91, 12218–22 (1994). 103. Pak, M. and Wickner, S. Mechanism of protein remodeling by ClpA chaperone. Proc Natl Acad Sci USA 94, 4901–6 (1997). 104. Kawarabayasi, Y. et al. Complete genome sequence of an aerobic hyper-thermophilic crenarchaeon, Aeropyrum pernix K1. DNA Res 6, 83–101, 145–52 (1999). 105. Minuth, T. et al. The recombinant thermosome from the hyperthermophilic archaeon Methanopyrus kandleri: in vitro analysis of its chaperone activity. Biol Chem 380, 55–62 (1999). 106. Phipps, B. M., Hoffmann, A., Stetter, K. O., and Baumeister, W. A novel ATPase complex selectively accumulated upon heat shock is a major cellular component of thermophilic archaebacteria. Embo J 10, 1711–22 (1991). 107. Knapp, S. et al. The molecular chaperonin TF55 from the thermophilic archaeon Sulfolobus solfataricus. A biochemical and structural characterization. J Mol Biol 242, 397–407 (1994). 108. Ng, W. V. et al. Genome sequence of Halobacterium species NRC-1. Proc Natl Acad Sci USA 97, 12176–81 (2000). 109. Kowalski, J. M., Kelly, R. M., Konisky, J., Clark, D. S., and Wittrup, K. D. Purification and functional characterization of a chaperone from Methanococcus jannaschii. Syst Appl Microbiol 21, 173–8 (1998). 110. Kusmierczyk, A. R. and Martin, J. Nested cooperativity and salt dependence of the ATPase activity of the archaeal chaperonin Mm-cpn. FEBS Lett 547, 201–4 (2003). 111. Andra, S., Frey, G., Nitsch, M., Baumeister, W., and Stetter, K. O. Purification and structural characterization of the thermosome from the hyperthermophilic archaeum Methanopyrus kandleri. FEBS Lett 379, 127–31 (1996). 112. Smith, D. R. et al. Complete genome sequence of Methanobacterium thermoautotrophicum deltaH: functional analysis and comparative genomics. J Bacteriol 179, 7135–55 (1997). 113. Robb, F. T. et al. Genomic sequence of hyperthermophile, Pyrococcus furiosus: implications for physiology and enzymology. Methods Enzymol 330, 134–57 (2001). 114. Nitsch, M., Klumpp, M., Lupas, A., and Baumeister, W. The thermosome: alternating alpha and betasubunits within the chaperonin of the archaeon Thermoplasma acidophilum. J Mol Biol 267, 142–9 (1997). 115. Kawashima, T. et al. Archaeal adaptation to higher temperatures revealed by genomic sequence of Thermoplasma volcanium. Proc Natl Acad Sci USA 97, 14257–62 (2000). 116. Izumi, M., Fujiwara, S., Takagi, M., Kanaya, S., and Imanaka, T. Isolation and characterization of a second subunit of molecular chaperonin from Pyrococcus kodakaraensis KOD1: analysis of an ATPasedeficient mutant enzyme. Appl Environ Microbiol 65, 1801–5 (1999). 117. Yan, Z., Fujiwara, S., Kohda, K., Takagi, M., and Imanaka, T. In vitro stabilization and in vivo solubilization of foreign proteins by the beta subunit of a chaperonin from the hyperthermophilic archaeon Pyrococcus sp. strain KOD1. Appl Environ Microbiol 63, 785–9 (1997). 118. Yoshida, T., Kawaguchi, R., and Maruyama, T. Nucleotide specificity of an archaeal group II chaperonin from Thermococcus strain KS-1 with reference to the ATP-dependent protein folding cycle. FEBS Lett 514, 269–74 (2002).

9214_C005.indd 70

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71

119. Marco, S. et al. The molecular chaperone TF55. Assessment of symmetry. FEBS Lett 341, 152–5 (1994). 120. Nakamura, N. et al. Purification and molecular cloning of the group II chaperonin from the acidothermophilic archaeon, Sulfolobus sp. strain 7. Biochem Biophys Res Commun 236, 727–32 (1997). 121. Benaroudj, N., Zwickl, P., Seemuller, E., Baumeister, W., and Goldberg, A. L. ATP hydrolysis by the proteasome regulatory complex PAN serves multiple functions in protein degradation. Mol Cell 11, 69–78 (2003).

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Physical Properties of Membranes Composed of Tetraether Archaeal Lipids Parkson Lee-Gau Chong

CONTENTS Introduction .................................................................................................................................. Tetraether Archaeal Lipids .......................................................................................................... Membranes Made of Bipolar Tetraether Archaeal Lipids ........................................................... Formation of Liposomes ................................................................................................... Formation of Planar Monolayers ...................................................................................... Physical Properties of Tetraether Archaeal Liposomes ............................................................... Phase Behaviors ................................................................................................................ Phase Transitions in TPLE and P2 Liposomes Derived from Sulfolobus solfataricus ............................................................................. Phase Transitions in PLFE Liposomes Derived from Sulfolobus acidocaldarius ....................................................................... Membrane Stability .......................................................................................................... Solute Permeability ................................................................................................ Vesicle Aggregation and Fusion ............................................................................ Membrane Packing ........................................................................................................... Packing in PLFE Liposomes Derived from S. acidocaldarius ............................. Packing in Tetraether Liposomes Derived from S. solfataricus ............................ Structural Factors and Forces in Membrane Packing ............................................ Membrane Lateral Diffusion and Lateral Organization ................................................... Lateral Diffusion .................................................................................................... Lateral Organization .............................................................................................. Technological Applications of Tetraether Lipid Membranes ...................................................... Acknowledgment ......................................................................................................................... References ....................................................................................................................................

73 74 78 78 78 79 79 79 79 81 81 83 84 84 85 85 87 87 88 88 89 89

INTRODUCTION Studies of thermoacidophilic archaea are of great biological and technological interest. The harsh conditions (e.g., high temperature and acidic media) in their habitat suggest that these organisms must have extraordinarily stable membranes, proteins, and nucleic acids. The lipid composition of the plasma membrane of these extremophiles is distinctly different from that of eukaryotes and bacteria. The membrane lipids in thermoacidophilic archaea are dominated by tetraethers. These unusual lipids can form stable liposomes, planar membranes, and nonlamellar lipid assemblies. These tetraether lipid membranes or assemblies can serve as models for a better understanding of 73

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the structure–function relationship of the plasma membrane in thermoacidophiles and, in addition, can be used for technological applications. This chapter reviews the physical properties of tetraether lipid membranes. Special attention is focused on phase behaviors, solute permeation, vesicle aggregation and fusion, membrane structure and packing, membrane dynamics, and lateral organization.

TETRAETHER ARCHAEAL LIPIDS Traditionally, archaea are classified as halophiles, methanogens, and thermoacidophiles [1]. The most prominent chemotaxonomic markers of archaea are ether lipids (Figures 6.1 and 6.2) which are polar lipids composed of polyisoprenoid chains linked to either glycerol or calditol [2–5]. Polar lipids take up about 80% to 90% of the total lipids in the archaea [6,7]. Their glycerol backbone is in the sn-glycerol-1-phosphate stereochemistry, in contrast to the sn-glycerol-3-phosphate backbone found in bacteria and eukaryotes. The polyisoprenoid chains contain branched methyl groups and may possess cyclopentange rings (Figures 6.1 and 6.2). Two types of ether lipids, namely, diethers (Figure 6.1) and tetraethers (Figure 6.2), are found in archaea [3,4,7–11]. The polar lipids in halophiles are mainly composed of diphytanylglycerol diether lipids, also known as archaeols (Figure 6.1). A phytanyl chain is a polyisoprenoid chain containing 20 carbons. In a typical methanogen, the polar lipids consist of ~50–100% diether lipids (e.g., archaeols or their derivatives) (Figure 6.1) and ~0–50% dibiphytanylglycerol tetraether lipids with a caldarchaeol hydrophobic core (Figure 6.2). Caldarchaeol is also called glycerol-dialkyl-glycerol-tetraether (GDGT) (Figure 6.2). In low-temperature methanogens, the isoprenoid chains in the ether lipids usually contain no cyclopentane rings [7]. However, macrocyclic diether lipids with cyclopentane rings (Figure 6.1) have been found in certain methanogens such as the ones collected from a mud volcano in the Sorokin Trough, NE Black Sea [12]. In hyperthermophilic methanogens, such as Methanopyrus kandleri, the isoprenoid chains in the diether lipids can be either saturated or unsaturated [13,14]. In thermoacidophiles, the total polar lipid extract (TPLE) contains ~5–10% diphytanylglycerol diether lipids and ~90–95% dibiphytanylglycerol tetraether lipids [7], with either a caldarchaeol (GDGT) or a calditoglycerocaldarchaeol hydrophobic core (Figure 6.2). A biphytanyl chain is a polyisoprenoid containing 40 carbons due to the condensation of two phytanyl chains. Calditoglyce rocaldarchaeol is traditionally called glycerol-dialkyl-nonitol-tetraether (GDNT) (Figure 6.2). However, more recent findings suggest that the 9-carbon nonitol moiety may actually exist in a polyhydroxylated cyclopentanic form known as calditol [5,15,16]. For this reason, GDNT has been used as an abbreviation for both GDNT and glycerol-dialkyl-calditol-tetraether. The GDNT-based tetraether lipids are typically found only in the members of the order Sulfolobales and constitute

Diether Lipids Macrocyclic Diethers

Archaeols O

O

O

O

OR

OR R = H or a polar group

FIGURE 6.1 Illustrations of the molecular structures of diether lipids found in archaea. Archaeols are also called diphytanylglycerol diether.

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Physical Properties of Membranes Composed of Tetraether Archael Lipids Tetraether Lipids Glycerol Dialkyl Calditol Tetraether (GDNT)

HO

OH

O O

R3

O

HO

OH

O

O

O

R1–PO4 OH

HO HO

O

OH

O

R3

O O R1–PO4

O O

Glycerol Dialkyl Glycerol Tetraether (GDGT) R2 O

O

O

O

R1–PO4 O R2 O R1–PO4

O O

FIGURE 6.2 Illustrations of the molecular structures of bipolar tetraether lipids found in archaea. GDGT and GDNT are also called caldarchaeol and calditoglycerocaldarchaeol, respectively. Two structures of GDGT- and GDNT-based bipolar tetraether lipids are presented. Actually, the number of cyclopentane rings may vary from 0 to 4 in each biphytanyl chain. For the polar lipid fraction E (PLFE) lipid fraction from Sulfolobus acidocaldarius, R1 = inositol, R2 = β-D-galactosyl-D-glucose, and R3 = β-glucose.

70% to 80% or more of the total lipids of the thermoacidophiles [5,17,18]. A Metallosphaera sedula TA-2 strain from hot springs in Japan is an exception [19]. TA-2 is different in its lipid composition from other members of Sulfolobales. TA-2 has only GDGT-based lipids whereas the other members have both GDGT- and GDNT-derived tetraether lipids. This has raised a question whether GDNT is essential for survival under high temperature and acidic conditions [19]. In thermoacidophiles, the number of cyclopentane rings in each biphytanyl chain may vary from 0 to 4 with increasing growth temperature [2,20]. Tetraethers of Thermoplasma contain up to two cyclopentane rings per biphytanyl chain and of Sulfolobus two to four rings [7]. For tetraether lipids isolated from cells grown at a given temperature, the number of cyclopentane rings per dibiphytanyl chain is not fixed to a single integer value. Instead, the isolated tetraether lipids from the same batch contain a range of isoprenoid species differing in the number of cyclopentane rings [7,17,21]. At present, the data for the average value and the distribution of the number of cyclopentane rings per

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isoprenoid chain in a given cell growth condition are still rather limited; those data would be valuable for gaining a better biophysical understanding of archaeal tetraether liposomes. Various polar groups can be linked to the glycerol or calditol moieties of archaeal tetraether lipids. For example, the polar lipid fraction E (PLFE) is one of the main constituents in the plasma membrane of the thermoacidophilic archaeon Sulfolobus acidocaldarius which thrives at temperatures of 65°C to 85°C at a pH of ~2–3. PLFE contains a mixture of GDNT- and GDGT-based bipolar tetraether lipids (Figures 6.2 and 6.3). The GDNT component of PLFE (~90% of total PLFE) contains phospho-myo-inositol on the glycerol end and β-glucose on the calditol end, whereas the GDGT component (~10% of total PLFE) has phospho-myo-inositol attached to one glycerol and β-d-galactosyl-d-glucose to the other glycerol skeleton (Figure 6.2). Thus, in PLFE, both GDGT and GDNT components are bi-substituted in the polar headgroup regions; for this reason, they are designated as bipolar tetraether lipids (Figure 6.3). In thermoacidophiles, bipolar dibiphytanyl tetraether lipids are the dominating lipid species (~90–95%). The ability for thermoacidophiles to resist high temperature and low pH has been partly attributed to the unique structure of those lipids. In the Sulfolobus genus, S. solfataricus is another species that has been extensively studied. Four major fractions of the tetraether lipids in S. solfataricus have been isolated from the TPLE by silica gel column chromatograph [22]. The fractions P1, GL, and SL are monopolar tetraether lipids whereas the fraction P2 consists of bipolar tetraether lipids containing ~10% GDGT and ~90% GDNT with the same polar headgroups as PLFE from S. acidocaldarius (Figure 6.3). SL is a sulfurcontaining GDNT-based lipid. GL is a glycolipid. P1 is a GDGT-based phospholipid. The mean weight composition of TPLE has been reported as 10% P1, 30% GL, 7% SL, 48% P2, and ~5% monopolar diphytanyl glycerol (DPG) [23,24]. The P2 fraction is equivalent to the PLFE fraction from S. acidocaldarius. In addition to the aforementioned major fractions, an unusual acyclic tetraether lipid has been identified in S. solfataricus [2], where two oxygen atoms on opposite glycerol backbones are linked to one single biphytanyl chain crossing the hydrophobic core and each of the other two oxygen atoms is linked to a phytanyl chain (similar to the hydrophobic core of a-TEPC shown in Figure 6.4).

GDNT backbone

GL

Monopolar

SL

70%

P2 PLFE

30%

100% 100%

P1

Bipolar

GDGT backbone

90% 90%

10% 10%

FIGURE 6.3 Schematic structure of tetraether lipids from Sulfolobus solfataricus (GL, SL, P1, P2) and Sulfolobus acidocaldarius (PLFE). Wavy lines: hydrocarbon chains; small circles: unsubstituted glycerol OH; black square: nonitol; large circles: phosphomyoinositol; square: β-d-glucopyranose; double square: β-dglucopyranosyl-β-d-galactopyranose; triangle: β-d-glucopyranosyl sulfate. (From Gliozzi, A., Relini, A., and Chong, P.L.-G., J. Membr. Sci., 206, 131, 2002. With permission.)

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Physical Properties of Membranes Composed of Tetraether Archael Lipids

DPhPC _

P

_

O P

O

74.2

38.2

670

4.82 ± 0.30

70.8

39.3

710

0.78 ± 0.13

O

O

O

O O

O

70.2

_

O

O

39.2

2020

P O

+

O

O O

O

O

_

0.53 ± 0.17

O

O

N

P O

O

m -TEPC

+ N

D (10–8cm2/s)

O

a -TEPC

+ O

K (dyn/cm)

O

O

N

d (Å)

O

O

+ O

N

A (Å2)

P O _ O

N

+

O

FIGURE 6.4 Comparison of molecular dynamic calculations of membrane packing and dynamic properties in three ether lipid models. DPhPC: diphytanylphosphatidylcholine; a-TEPC: acyclic tetraether phosphatidylcholine; m-TEPC, macrocyclic tetraether phosphatidylcholine. A is the averaged molecular area (in Å); d is the peak-to-peak distance (in Å) of electron density profile (i.e., membrane thickness); K is the elastic area expansion modulus (in dyne/cm); D is the lateral diffusion coefficient in cm 2/s. (From Chong, P.L.-G., Ravindra, R., Khurana, M., English, V., and Winter, R., Biophys. J., 89, 1841, 2005. With permission.)

GDGT-based tetraether lipids are also abundant in nonthermophilic crenarchaea found in marine environments, soils, peat bogs, and low-temperature areas [25–30]. Many of those GDGTbased tetraether lipids contain branched biphytanyl chains but not in the form of isoprenoid, in contrast to the GDGT-based lipids found in thermophilic crenarchaea. A novel GDGT containing four cyclopentane rings and one cyclohexane ring was found in planktonic crenarchaea [27,28]. Like thermoacidophiles, marine crenarchaea can adjust the number of cyclopentane rings from zero to four in their branched GDGT tetraether lipids according to growth temperature [31]. For branched GDGT tetraether lipids found in soils, the number of cyclopentane rings (ranging 0–2) is primarily related to the pH of the soil while the number of branched methyl groups (ranging 4–6) is correlated with the annual mean air temperature [32]. Ether lipids are also present in the hyperthermophilic bacteria Aquificales and Thermotogales found in geothermally heated environments such as the pink streamer and vent biofilm from Octopus Spring in Yellowstone National Park [33]. Those ether lipids are diethers, such as diphytanylglycerol diether (archaeol, Figure 6.1) and C18,18-dialkyl glycerol diether, or monoethers such as C20- and C25-isoprenoid glycerol monoethers. An unusual glycerol monoether with a dimethyltriacontanyl chain has been identified in the hyperthermophilic bacterium Thermotoga maritima (growing between 55°C and 90°C) [34]. To our knowledge, no bipolar tetraether lipids have been reported in those hyperthermophilic bacteria. It appears that ether lipids, but not necessarily tetraethers, associate with extreme thermophiles. A number of acyclic and macrocyclic tetraether lipids have been synthesized [35–38]. None of them is identical to the natural P2 or PLFE archaeal lipids containing cyclopentane rings. However,

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synthetic tetraether lipids have provided many novel structural features that enable us to test the structure–property relationship in archaeal lipid membranes.

MEMBRANES MADE OF BIPOLAR TETRAETHER ARCHAEAL LIPIDS FORMATION OF LIPOSOMES In an aqueous phase, tetraether archaeal lipids may form stable (either small or medium size) unilamellar vesicles by sonication, detergent removing, or extrusion (~50–800 nm in diameter), multilamellar vesicles (MLVs) by shaking, and giant unilamellar vesicles (GUVs) by electroformation methods (~10–150 μm) [11,39–42]. However, not all the tetraether lipid fractions isolated from the archaea are able to form stable liposomes. According to Lo and Chang [40], PLFE is the only polar lipid fraction from S. acidocaldarius that is able to form stable liposomes. The P2 fraction from S. solfataricus, which is equivalent to the PLFE fraction from S. acidocaldarius, is also able to form vesicles in an aqueous phase [23]. It is possible to make closed vesicles from the TPLE of S. solfataricus [23], even though the theoretical calculation is not in favor of vesicle formation by these lipids [43]. Monopolar tetraether lipids (e.g., P1, SL, and GL from S. solfataricus) as well as hydrolyzed GDGT and hydrolyzed GDNT (with sugar and phosphate moieties removed) alone usually do not form closed vesicles [24,43,44]. Mixing different tetraether lipids or addition of monopolar diester (or diether lipids) to tetraether lipids may result in stable liposomes [23,43]. The lamellarity in the PLFE GUV liposomal membrane has been tested by Laurdan [6-lauroyl-2(dimethylamino)naphthalene] fluorescence [41]. The lack of Laurdan’s fluorescence intensity within the lipid domains observed by two-photon excitation fluorescence microscopy at low temperature indicated that the PLFE GUVs being studied were unilamellar. For MLVs, it would be extremely unlikely that the dark domains from one lipid layer match exactly with the domains from the others. In tetraether liposomes, lipids span the entire lamellar structure, forming a monomolecular thick membrane [7,45], in contrast to the bilayer structure formed by monopolar diester (or diether) phospholipids. The experimental evidence for a monolayer structure in tetraether liposomes initially came from freeze-fracture electron microscopy studies [40,45], which showed the lack of the fracture plane, while only cross-fracturing of the membrane was observed. The tetraether liposomes have allowed functional reconstitution of different integral membrane proteins [45–48]. In reconstitution studies of the cytochrome aa3-type quinol oxidase from S. acidocaldarius [48], for example, the quinol oxidase was able to generate and maintain a protonmotive force at temperatures (~70°C) close to the growth temperature of the archaeon. In addition, the quinol oxidase shows a higher turn-over number when reconstituted in S. acidocaldarius lipids (mainly tetraether lipids) as compared with Escherichia coli lipids (mainly diester lipids) [48]. These results suggest that tetraether liposomes are a useful membrane model for understanding the structure–function relationship of the plasma membrane in thermoacidophiles and that tetraether lipids provide an unusual lipid environment for the function of membrane proteins. However, the transmembrane orientation of the asymmetric tetraether lipids (e.g., PLFE lipids, Figures 6.2 and 6.3) and proteins in liposomes is hard to control, which could be a potential problem in interpreting the data from the reconstitution experiments. In contrast, in archaea cells, the phosphate moiety of bipolar tetraether lipids (Figure 6.2) has been reported to face mainly the cytoplasmic side whereas the sugar residues (e.g., galactose-glucose disaccharides) orient primarily to the extracellular environment [49,50].

FORMATION OF PLANAR MONOLAYERS The native archaeal tetraether lipids may form stable planar monomolecular films at the water–air interface [51,52]. In the monolayer film, the tetraether lipids may adopt two conformations. In the U-shaped conformation, the two polar headgroups end in the aqueous subphase. In the “upright” conformation, one polar end sits in the aqueous subphase while the other polar end hangs in the air.

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These two conformations may coexist and vary their proportions with spreading time and lateral pressure. A tetraether lipid with an “upright” conformation occupies an area of 0.72 –0.85 nm2 with a height ~4–5 nm in the monolayer film [45,51,53,54]. The occupied area per tetraether lipid molecule increases to 0.9–1.6 nm2 [52,55,56], and the height drops to 0.8–2.0 nm [53,54], when the U-shaped conformation dominates. The lack of cyclopentane rings in the bipolar tetraether lipids from Methanospirillum hungatei has been suggested to cause those lipids to adopt a U-shaped configuration in the monolayer film at the water–air interface [52]. The hydrolyzed GDNT and GDGT extracted from S. acidocaldarius or S. solfataricus do not form stable monolayer films at the air–water interface [57,58]. It was cautioned that the data derived from these monolayer films were dynamic rather than equilibrium values [58]. As pointed out by Kim and Thompson [35], the U-shaped conformation proposed for planar monolayers at the air–water interface may be misleading for describing the bipolar tetraether lipid conformation in liposomes. In liposomes, the two highly polar headgroups in bipolar tetraether lipids should be able to form extensive hydrogen bond networks on both surfaces of the liposomal membrane. The hydrogen bond networks plus the van der Waals and hydrophobic interactions should be able to maintain a stable “upright” monomolecular membrane-spanning arrangement without invoking a U-shaped conformation in tetraether liposomes [35].

PHYSICAL PROPERTIES OF TETRAETHER ARCHAEAL LIPOSOMES PHASE BEHAVIORS Phase Transitions in TPLE and P2 Liposomes Derived from Sulfolobus solfataricus Lipid membranes can undergo phase transitions via changes in temperature, pressure, and membrane composition. The temperature-induced phase transitions in lipid membranes made of the TPLE and the bipolar tetraether lipid fractions [22] from S. solfataricus have been characterized by x-ray diffraction, calorimetry, and other techniques [59,60]. Lipid assemblies made of TPLE exhibit complex polymorphic behaviors including a transition from the lamellar to the cubic phase at ~80°C [59–61]. Liposomes made of the P2 fraction show a strict lamellar structure [59–61]. Phase Transitions in PLFE Liposomes Derived from Sulfolobus acidocaldarius PLFE liposomes from S. acidocaldarius have been characterized by a variety of physical techniques. Differential scanning calorimetry (DSC) was employed [62] to detect the phase transitions involving significant enthalpy changes (ΔH). As shown in Table 6.1, the DSC data (summarized from three consecutive scans) from PLFE liposomes show an endothermic peak at 44.2°C to 46.7°C (ΔH = 3.5–4.2 kJ/mol, peak I) and at 57.1°C to 58.6°C (ΔH = 1.5–2.0 kJ/mol, peak II) and an exothermic peak at 78.5°C (ΔH = −23.2 kJ/mol, peak III) [62]. Peak I is in good agreement with the lamellarto-lamellar phase transition observed at ~50°C by small angle x-ray scattering (SAXS) (Figure 6.5) [63], ~46–48°C by infrared [63], ~48°C by perylene rotational rate [64], ~48°C by excimer fluorescence of pyrene-labeled phospholipids [65], ~45°C by pressure perturbation calorimetry (PPC) [62] and ~50°C by generalized polarization (GP) of Laurdan fluorescence [41]. Peak II, appeared at 57.1°C to 58.6°C in the DSC scans (Table 6.1), corresponds to the lamellar-to-lamellar phase transition at ~60–61°C detected by infrared and SAXS (Figure 6.5) [63]. As summarized in Table 6.1, there is a fairly good agreement from different techniques that PLFE liposomes derived from S. acidocaldarius undergo two thermal-induced lamellar-to-lamellar phase transitions: one at ~47.5 ± –2.5°C and the other at ~60°C. The exothermic transition at 78.5°C in the first heating DSC scan [62] (Table 6.1, peak III) corresponds to the phase transition from lamellar to probably inverted bicontinuous cubic phases (QIID and QIIP) as detected at ~74–75°C by SAXS [63]. The assignment to the coexistence of QIID and QIIP phases is based on the small angle x-ray diffraction pattern and the calculated ratio of the lattice

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TABLE 6.1 Temperature-Induced Phase Transitions in Polar Lipid Fraction E (PLFE) Liposomes Derived from Sulfolobus acidocaldarius Technique Used

Cell Growth Type of Vesicles T (°C)

Phase Transition Detected

References

I: 44.2–46.7°C (ΔH = 3.5–4.2 kJ/mol) II: 57.1–58.6°C (ΔH = 1.5–2.0 kJ/mol) (detected in the second and third scans) III: 78.5°C (ΔH = −23.2 kJ/mol) I: 51.0°C ( ΔH = 14.2 kJ/mol) II: not detected in the first scan III: 83.2°C ( ΔH = −18.0 kJ/mol)

[62]

Differential scanning calorimetry (DSC)

78

Multilamellar vesicles (MLVs) (pH 2.1)

DSC

65

MLVs (pH 2.1)

DSC

65

MLVs (pH 7.0)

I: 41.7–43.7°C ( ΔH = 10.0–16.0 kJ/mol) II: not detected in the first scan III: not reported

[62]

Pressure perturbation calorimetry (PPC)

78

MLVs (pH 2.1)

[62]

PPC

65

MLVs (pH 2.1)

PPC

65

MLVs (pH 7.0)

Small angle x-ray scattering

69–70

MLVs (pD 2.15)

IR CH2 symmetric stretching mode

69–70

MLVs (pD 2.15)

I: 45°C (ΔV/V = 0.10–0.14%); II: 57.5–60.0°C (ΔV/V = 0.08–0.09%) III: no significant peak I: 42.0°C (ΔV/V = 0.25%) II: not reported III: no significant peak I: 43.0°C (ΔV/V = 0.56%) II: not reported III: no significant peak I: 50°C (d ~ 50 Å) II: 60°C (d ~ 54 Å) III: 74–75°C (d ~ >56 Å) I: 46–48°C II: 61°C III: 74°C

Perylene fluorescence

69–70

MLVs (pH 7.2)

~48°C

[64]

Pyrene-phosphatidylcholine fluorescence

65–67

MLVs (pH 7.4)

~48°C

[65]

Laurdan fluorescence

69–70

Giant unilamellar vesicles (GUVs) (pH 2.9 and 7.2)

~50°C 44

[41]

[62]

[62]

[62]

[63]

[63]

constants of the cubic structures QIID and Q IIP [63]. This transition is broad, probably due to the coexistence of QIID and Q IIP and the chemical heterogeneity of PLFE. This exothermic transition disappears in the subsequent heating DSC scans [62], suggesting that the phase transition at ~78.5°C involves a metastable phase, which is irreversible at the scan rate used (20°C/h). PPC (pressure = 5 bar) has been used to study the thermal volume expansion coefficient, α, in PLFE liposome [62]. From the plot of α versus temperature, the phase transitions that involve significant volume changes were detected and the relative volume changes (ΔV/V) associated with the phase transitions were calculated. The lamellar-to-lamellar phase transitions of PLFE liposomes involve small volume changes (ΔV/V ~0.08–0.14%) (Table 6.1 and Figure 6.5), compared with the

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FIGURE 6.5 Lamellar repeat unit d of polar lipid fraction E (PLFE) liposomes in excess water (85 wt% D2O) as a function of temperature. Also shown are the volumetric properties at different temperature regions where either phase transitions are evident in the plot of the d-spacing (as revealed by small-angle x-ray scattering) versus temperature [63] or microdomain formation occurs [41].

ΔV/V value 3.0% for the main phase transition of dipalmitoylphosphatidylcholine (DPPC, a monopolar diester) [62]. The PPC scans do not reveal any significant peak in the 79°C to 83°C region, indicating that the lamellar-to-cubic phase transition does not involve a significant volume change (Table 6.1) [62]. High hydrostatic pressure (up to 150 kbar) also induces phase transitions in PLFE liposomes. The symmetric CH2 stretching vibrational wavelength reveals a variety of gel-like phases at elevated pressures under isothermal conditions [63]. The most prominent pressure-induced phase transition involves a ~2 cm–1 increase in wave number, which appears at 8.0 kbar at 60°C, 8.4 kbar at 43°C, and 10.8 kbar at 20°C, giving an unusual negative dT/dP value. The anomaly has been attributed to the temperature attenuation of the hydrogen bond networks in the polar headgroup region [63]. It is clear from all these phase transition studies that there is a rich polymorphism in tetraether liposomes.

MEMBRANE STABILITY Solute Permeability For permeation through monopolar diester bilayer membranes, solutes need to cross three membrane regions: the lipid head group (highly viscous, thus low permeation), hydrocarbon chain segment near the polar head group (semi-rigid), and hydrocarbon tail near the bilayer center (fluid, thus high permeation) [66]. This follows that solute permeation changes with membrane thickness, type of polar head groups, and membrane free volume. It has been proposed that small solutes move through the membrane by “hopping” between voids, often through the gauche-trans isomerization of the hydrocarbon chain [67].

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Most, but not all, of these considerations are applicable to bipolar tetraether liposomes. In the polar head group regions of bipolar tetraether liposomes, there is an extensive network of hydrogen bonds [46,68], which should generate a high electrical dipole potential, thus hindering solute permeation [44]. In bipolar tetraether liposomes, the biphytanyl hydrocarbon chains are linked covalently from one polar end to the other, lacking the midplane spacing. The cyclopentane rings in the dibiphytanyl chains also provide rigidity to the membrane. As such, bipolar tetraether lipids should exhibit rather limited gauche-trans isomerization in their hydrocarbon chains, thus a reduced rate for the “hopping” motion and a lower rate for solute permeation. Bartucci et al. [69] pointed out that the water permeability coefficient is determined by both the partition coefficient and the diffusion coefficient of water at different depths of the membrane. The partition coefficient is determined by the transmembrane polarity profile whereas the diffusion coefficient is dependent upon the hydrocarbon chain flexibility. Their spin-label data indicated that, although the transmembrane polarity profile is similar [69], the chain flexibility is reduced considerably in the P2 tetraether liposomes, compared to monopolar diester liposomes. It is then predicted [69] that the overall water permeability will be significantly lower for tetraether liposomal membranes than for normal monopolar diester liposomes. Indeed, using carboxyfluorescein fluorescence, Mathai et al. [70] demonstrated that water permeability across liposomes made of total lipid extract from Thermoplasma acidophilum (containing 90% bipolar tetraether lipids with two cyclopentane rings per molecule) is reduced by approximately fivefold as a result of the rigid and tight membrane packing due to the macrocyclic structure formed by dibiphytanyl chains and the glycerol backbones. A study on synthetic diether (rather than tetraether) archaea-like macrocyclic lipid membranes also showed that water permeation is significantly reduced by the cyclic ring structure in the lipid [71]. Like water, the passive permeation of urea, glycerol, and ammonium can be decreased several fold by tetraether lipids [70]. Since it has been proposed that proton permeation across lipid bilayers is mediated by the hydrogen-bonded chain of water [72], low water permeability across tetraether liposomal membranes implies a low proton permeability. Using pyranine fluorescence, Elferink et al. [73] demonstrated that the proton permeability in bipolar tetraether liposomes derived from S. acidocaldarius is lower and less temperature sensitive than that in liposomes composed of monopolar diester lipids. Using 5,6-carboxyfluorescein (5,6-CF) fluorescence, Komatsu and Chong [42] found similar results in PLFE liposomes derived from S. acidocaldarius. The proton permeability comparison of various liposomes reveals that the tight and rigid lipid packing is a major contributor of the low proton permeability in PLFE liposomes, whereas the inositol moiety and the branched methyl groups may contribute, but to a lesser extent [42]. Small PLFE unilamellar vesicles (SUVs, ~61 nm in diameter) exhibited even lower proton permeability than large PLFE unilamellar vesicles (LUVs, ~240 nm) and the proton permeability in PLFE SUVs was less sensitive to temperature, changing by <2 × 10 −10 cm/s from 25°C to 82°C [42]. This finding is surprising because curvature stress in SUVs is supposed to loosen membrane packing, consequently increasing proton permeation. This result may be caused by curvature-induced changes in membrane surface potential and lipid packing due to changes in the transmembrane distribution of negatively charged phosphate [42]. Membrane stability can also be monitored by determining the release of fluorescent dyes originally trapped in the intravesicular compartment of the liposomes. Tetraether liposomes exhibit unusual thermal stability against dye leakage [42,73,74]. The high stability has been attributed to the structure of tetraether lipids, the negative charges on membrane surface (zeta-potential = –34 mV in PLFE SUVs), and the tight and rigid lipid packing [42,73,74]. The rate of calcein release from liposomes made of TPLE from S. solfataricus is higher than that from P2 liposomes [23]. This result argues that membrane packing, rather than membrane thickness [75], is the main determinant of solute permeability in bipolar tetraether liposomes [23]. Membrane thickness in TPLE liposomes is known to be greater than that in P2 liposomes [59]. If membrane thickness is the main determinant, the thicker membrane (i.e., TPLE) would lead to a lower solute permeability [75].

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In summary, the low solute permeability in tetraether liposomes [42,70,73,74,76] is a direct consequence of the chemical structure of tetraether lipids and their monolayer organization, especially the cyclopentane rings and the hydrogen bond network between the sugar residues of the bipolar tetraether lipids exposed at the outer face of the membrane. Vesicle Aggregation and Fusion Membrane fusion is an important cellular process involved in membrane trafficking, fertilization, and virus infection. Bilayer membrane studies lead to a stalk-hemifusion-pore hypothesis for the mechanism of vesicle fusion [77,78]. According to this hypothesis, two juxtaposed bilayers form a stalk configuration at the contact point, followed by formation of a hemifusion intermediate and fusion pore. The fusion pore allows mixing of internal contents. While this hypothesis seems to work for bilayer membranes, it is not clear whether it is applicable to tetraether liposomes. Tetraether lipids span the entire membrane forming a monomolecular structure, which hinders the formation of the stalk and hemifusion intermediates. The fusogenic agent Ca2+ was able to induce fusion between S. solfataricus TPLE tetraether liposomes only above 60°C [23,24]. TPLE liposomes undergo a transition from the lamellar to the cubic phase at ~80°C [60] as mentioned earlier. It was suggested [23] that, in TPLE liposomes, local nonlamellar structures, such as cubic phases, may occur at temperatures as low as 60°C. The nonlamellar structure is of crucial importance for fusion to occur [79]. TPLE contains bipolar tetraether lipids (P2), monopolar tetraether lipids (P1, SL, GL) and monopolar diether lipids (DPG) (see section “Tetraether Lipids”) [11,44]. The presence of DPG promotes Ca2+-induced fusion [24] probably due to the fact that DPG molecules can be transformed from the lamellar structure to hexagonal or cubic phases [80]. However, only vesicle aggregation, not fusion, was observed in the vesicles made of the P2 fraction from the same archaeon S. solfataricus. P2 liposomes contain a strict lamellar structure [59], and fusion occurs only when this lamellar structure is perturbed [23,24]. Ca2+-induced tetraether liposome fusion occurs on the time scale of tens of minutes [23,24], which is much slower than that of monopolar diester liposomes [81]. Moreover, TPLE monolayers do not show any detectable changes in surface tension even in cases where fusion occurs [23]. In comparison, Ca2+ binding increases membrane surface tension in monopolar diester lipid membranes, which in turn induces vesicle fusion [82]. In tetraether liposomes, membrane surface tension does not seem to play a significant role in the fusion mechanism. Unlike Ca2+, polyethylene glycol (PEG), another fusogenic agent, induced fusion of TPLE liposomes at temperatures as low as 25°C [24], suggesting that PEG greatly disrupts or reduces the hydration layer on TPLE vesicles, thus promoting vesicle aggregation and fusion. Like Ca2+, PEG (20% w/v) did not induce fusion in P2 vesicles [24]. In addition, PEG induced only a very limited extent of fusion at a slow rate in TPLE/P2 mixed vesicles. However, once DPG (a monopolar diether lipid) or egg yolk phosphatidylcholine (egg-PC, a monopolar diester lipid) was added to tetraether liposomes, PEG-induced vesicle fusion occurred at faster rates and to a larger extent [24]. Ca2+ also induced aggregation and limited fusion between PLFE liposomes derived from S. acidocaldarius [83]. Aggregation is the first step involved in membrane fusion and lipid-mixing. In the absence of Ca2+, the diameter of PLFE liposomes changes little in six months, showing no signs of vesicle aggregation [83]. Addition of sufficient amounts of Ca2+ induces PLFE vesicle aggregation, which can be reversed by EDTA [83]. The reversibility of vesicle aggregation by EDTA decreases with increasing temperature and with increasing the incubation time of EDTA with [Ca2+]. After long-incubation (two weeks) with Ca2+, the PLFE vesicles have more than just aggregated, but have fused or coalesced, as revealed by freeze-fracture electron microscopy [83]. The Ca2+-induced aggregation of PLFE liposomes is slow, on the order of tens of minutes [83,84], as compared with the aggregation of negatively charged monopolar diester liposomes at comparable Ca2+ and lipid concentrations (on the order of seconds) [85]. The initial rate of Ca2+-induced PLFE vesicle aggregation increases sigmoidally with [Ca2+] [83,84]. From the sigmoidal curve, a threshold calcium concentration (Cr) for vesicle aggregation can be determined. Cr increases with increasing

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temperature but decreases with increasing pH [83,84]. The temperature variation of Cr has been attributed to the changes in membrane surface potential, which is –22.0 mV and –13.2 mV at 25°C and 40°C, respectively, at pH 6.6, as determined from 2-(p-toluidinyl) naphthalene-6-sulfonic acid (2,6-TNS) fluorescence [14,83,86,87]. Phosphate has also been reported to induce lipid mixing and vesicle fusion in tetraether liposomes [84]. In summary, bipolar tetraether liposomes possess many unusual properties with regard to membrane aggregation and fusion. Compared with monopolar diester or diether liposomes, aggregation/ fusion of tetraether lipsomes is limited and slow, implying a very large energy barrier for fusion of tetraether liposomes [24,84]. This trend holds true for influenza virus hemagglutinin-induced vesicle fusion [88]. To date, the fusion mechanism for tetraether liposomes still remains very much a mystery. Elferink et al. [84] suggested that the fusion of tetraether liposomes did not involve a distinct hemifusion intermediate, due to the fact that tetraether lipids cannot split in the mid-plane of the membrane to make two leaflets, like the monopolar diester lipids. If a fusion pore is involved, it is likely that the tetraether lipids adopt a U-shaped conformation with the polar head groups covering the interior of the fusion pore [89].

MEMBRANE PACKING The extraordinary stability of tetraether liposomes can be attributed to the tight and rigid membrane packing due to the presence of branched methyl groups, tetraether linkages, and cyclopentane rings in the dibiphytanyl chains as well as an extensive network of hydrogen bonds between the sugar and phosphate residues exposed at the outer face of tetraether liposomes [11,90]. The experimental studies on membrane packing in archaeal tetraether liposomes are summarized next, followed by a brief review of molecular dynamics (MD) calculations of the contributing factors to membrane packing. Packing in PLFE Liposomes Derived from S. acidocaldarius The rigid and tight membrane packing in PLFE liposomes has been revealed by different physical techniques. The vibrational frequency of the polyisoprenoid CH2 symmetric stretching mode detected by infrared spectroscopy showed an overall 2 cm–1 increase in wavenumber between 61°C and 74°C. This large change indicates a phase transition from a rigid, gel-like lipid conformation to a chain conformation with considerable disorder, as expected from cubic phases which appear above ~74°C [63]. The GP values of Laurdan fluorescence in PLFE GUVs were found to be low at all of the temperatures and pHs examined [41]. When excited with light polarized in the y direction, Laurdan fluorescence in the center cross section of the PLFE GUVs exhibited a photoselection effect showing much higher intensities in the x direction of the vesicles, a result opposite that observed on monopolar diester liposomes, which showed much higher intensities in the y direction [91,92]. This surprising result indicates that the chromophore of Laurdan in PLFE GUVs is aligned parallel to the membrane surface, whereas the chromophore of Laurdan in monopolar diester liposomes is aligned perpendicular to membrane surface [91,92]. This photoselection effect and the low GP values suggest that the Laurdan chromophore resides in the polar headgroup region of the PLFE liposomes in parallel with membrane surface, while the lauroyl tail inserts into the hydrocarbon core of the membrane. This unusual L-shaped disposition is presumably caused by the unique lipid structures and by the rigid and tight membrane packing in PLFE liposomes [41]. Bilayer membranes of diacylphosphatidylcholines (such as dimyristoylphosphatidylcholine, DMPC) entail extensive trans-gauche conformational changes along the polymethylene chains [93], thus possessing a large ΔH (e.g., ~25 kJ/mol for DMPC) through their main phase transitions. The ΔH value for the lamellar-to-lamellar phase transitions in PLFE liposomes is low (e.g., 3.5 kJ/ mol in Table 6.1). The low ΔH value for PLFE liposomes suggests that the lamellar-to-lamellar phase transition involves restricted trans-gauche conformational changes in the dibiphytanyl chains

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due to the presence of the cyclopentane ring, branched methyl groups, and the bipolar nature of the PLFE lipids, and due to the spanning of the dibiphytanyl chains through the membrane. Using a multi-excitation method, Khan and Chong detected an abrupt increase in the rotational rates of perylene in PLFE liposomes at ~48°C [64]. Perylene is a fluorescent probe presumably residing in the hydrophobic core. The data suggest that the hydrocarbon region of PLFE liposomes is rigid and tightly packed below ~48°C. Above ~48°C, the hydrocarbon core of PLFE membranes begins to gain appreciable membrane fluidity, which would be required in order for archaeal membranes to function. The polar headgroup region of PLFE membranes, on the other hand, may still be rigid and tightly packed through the hydrogen-bond network [46,68] at elevated temperatures (>48°C) so as to maintain a large proton gradient (pH 2–3 outside and pH 6.5 inside the cell) across the membrane under the growth condition. This proposition explains why low proton-permeability and appreciable membrane fluidity can occur at the same time in thermoacidophiles at high growth temperatures. Packing in Tetraether Liposomes Derived from S. solfataricus A spin-label study showed that the nonitol (more precisely, calditol) headgroup of tetraether lipids from S. solfataricus was relatively immobile, while the rotation of the spin label positional isomers of stearic acid (n-SASL, n = 5, 12, and 16) was anisotropic and restricted in the time scale of both conventional and saturation-transfer electron spin resonance (ESR) spectroscopy. An appreciable fluidity is gained only at temperatures close to the minimum growth temperature of the cells [94]. Using phosphatidylcholine spin-labeled at different acyl chain positions (n = 5, 7, 10, 12, 14, and 16), Bartucci et al. [69] studied the effect of temperature on lipid chain flexibility in the P2 tetraether liposomes derived from S. solfataricus. The lipid chain in P2 liposomes remains ordered and less flexible. Only at elevated temperature (~80°C) does the chain flexibility in the P2 liposomes reach the level normally found in fluid diester liposomes. This implies [69] that the plasma membrane of S. solfataricus just begins to gain appreciable fluidity needed for functionality at temperatures close to that of minimum cell growth, a result consistent with the previous tetraether liposome studies [65,94]. Structural Factors and Forces in Membrane Packing The structural factors and forces that are important for membrane packing in tetraether lipid membranes have been evaluated by MD calculations on various ether lipid membrane models. The calculations have helped to understand and predict the experimental results. Ether Linkage MD simulations showed that substitution of ether linkages for ester linkages reduces membrane dipole potential by a factor of two [95,96]. Reduced hydration of the ether functional group causes a decrease in lipid molecular area and an increase of the free energy barrier for hydrophilic molecules to move across the membrane [95]. Branched Methyl Groups The branched methyl groups in the polyisoprenoid chains are bulky. They make the limiting occupied area (A) of diphytanyl phosphatidylcholine (DPhPC, a monopolar diether lipid, Figure 6.4) about two times greater than that of saturated diacyl phosphatidylcholine (a monopolar diester). They also reduce the hydrocarbon chain–chain interaction energies that need to be overcome to spread the monolayer membrane to a larger area [97]. As a result, a DPhPC Langmuir membrane has an unusually low surface tension (32–37 mN/m at 20–70°C), in comparison with the values 54–56 mN/m for the conventional diester lipids [97]. Branched methyl groups also decrease chain– chain tangling, chain trans-gauche conformational changes, segmental order, and the wobbling, rotational, and translational motions [98,99]. An analysis of the cavity distribution in liposomes revealed that the branched chain in DPhPC bilayer had, compared with the acyl chains in DPPC

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bilayers, a relatively small and discrete free volume distribution in the hydrophobic core [100]. This implies that small solute molecules (e.g., water) have a lower rate of diffusion inside branch-chained lipid bilayers than inside DPPC bilayers [100]. The lower solute permeability in DPhPC bilayers than in DPPC correlates with the slower dynamics of the branched DPhPC [100]. Covalent Linkage of Polyisoprenoid Chains Between Two Polar Ends Another possible contributing factor to the remarkable stability of tetraether lipid membranes is that the hydrocarbon chains of the lipid molecules are covalently linked from one polar end to the other end, spanning the whole membrane. This structural feature leads to an increase in membrane lipid rigidity and the loss of mid-plane spacing that occurs in normal bilayer membranes. With regard to the impact of this structural factor on membrane packing and dynamics, the MD simulations carried out by Sinoda et al. [101] revealed several important points. They compared three ether lipids (Figure 6.4) with the same headgroup chemical structure and stereochemistry. DPhPC is a frequently studied diether lipid model compound. m-TEPC is a macrocyclic tetraether phosphatidylcholine with the hydrophobic core mimicking that of naturally occurring tetrather lipids (Figure 6.2). a-TEPC is acyclic tetraether phosphatidylcholine, which differs from m-TEPC only by missing one single C-C bond that links two phytanyl chains in the hydrophobic core (Figure 6.4). As shown in Figure 6.4, m-TEPC has a higher density (= A × d) in the membrane interior than a-TEPC, demonstrating that dibiphytanyl linkage in m-TEPC provides a tighter membrane packing than mono-biphytanyl linkage in a-TEPC. More strikingly is the result that the m-TEPC membrane shows an elastic area expansion modulus (K) about three times higher than a-TEPC and DPhPC (Figure 6.4). This indicates that flexibility of membrane area expansion is very sensitive to whether the whole tetraether molecule has a cyclic structure or not. Because K is related to tensile strength, the data also imply that among these three lipids, m-TEPC has the highest stability against external mechanical stress. Since tighter packing yields less membrane free volume for molecular motion, the lateral diffusion constant (D) of m-TEPC is the smallest among these three lipids (Figure 6.4) [101]. Cyclopentane Rings As discussed earlier, an increase in growth temperature is known to increase the number of cyclopentane rings in the dibiphytanyl chains of archaeal tetraether lipids, and the number of cyclopentane rings may vary from 0 to 4 in each biphytanyl chain. It has been long postulated [7] that the cyclopentane rings in the biphytanyl chains of tetraether lipids would reduce chain length and rotational freedom in the chain, therefore increasing rigidity. Because lipid rigidity and membrane thickness are known to be important physical determinants of membrane permeability, membrane packing and dynamics [75], it is expected that changes in the number of cyclopentane rings or the cell growth temperature have a significant impact on membrane properties. This idea has been tested by computer simulations and experiments as summarized subsequently. To calculate how the number of cyclopentane rings might affect membrane packing, Gabriel and Chong have conducted MD simulations on a membrane containing 4 × 4 GDNT molecules with polar headgroups similar to those found in PLFE lipids [102]. They found that an increase in the number of cyclopentane rings in the dibiphytanyl chains of GDNT from 0 to 8 makes GDNT membrane more tightly packed and the lipid–lipid interaction energy more negative (i.e., energetically more stable) [102]. Calorimetry experiments also suggested that the number of cyclopentane rings in the dibiphytanyl chains would affect membrane packing because PLFE liposomes derived from different cell growth temperatures showed different thermodynamics properties [62]. The DSC study showed that PLFE liposomes derived from cells grown at 78°C exhibited a lamellar-to-lamellar phase transition at 46.7°C with an unusually low enthalpy change (ΔH = 3.5 kJ/mol) (Table 6.1), as compared with that for the main phase transition of DMPC. The PPC scan revealed that, at this same phase transition, the relative volume change (ΔV/V) in the membrane is very little (~0.1%), much lower than the

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ΔV/V value 2.8% for the main phase transition of DMPC. The low ΔH and ΔV/V values may arise from the restricted trans-gauche conformational changes in the dibiphytanyl chain due to the presence of cyclopentane rings and branched methyl groups and due to the spanning of the lipid molecules over the whole membrane [62]. For PLFE MLVs derived from cells grown at 65°C, similar DSC and PPC profiles were obtained. However, the lower cell growth temperature yielded a higher ΔV/V (~0.25%) and ΔH (14 kJ/mol) value for the lamellar-to-lamellar phase transition. A lower growth temperature also generated a less negative temperature dependence of α. The changes in ΔV/V, ΔH, and the temperature dependence of α can be attributed to the decrease in the number of cyclopentane rings in PLFE due to the lower growth temperature [62]. A decrease in the number of cyclopentane rings appears to make the membrane less tight and less rigid, thus a higher ΔV/V value through the phase transition. Note that an earlier DSC study [21] on the hydrated main tetraether glycophospholipid (MPL) of T. acidophilium grown at two different temperatures (39°C and 56°C) also showed a different phase behavior between the two samples, probably due to the changes in the number of cyclopentane rings in the dibiphytanyl chains [11]. Hydrogen Bond Networks and Membrane Surface Charge Molecular modeling on a membrane containing 4 × 4 unhydrolyzed GDNT molecules with phospho-myo-inositol or sugar moieties attached to the glycerol or nonitol backbones [102] has revealed that, as the number of cyclopentane rings per molecule is increased from 0 to 8, the phosphate–phosphate distance is shortened and, as a result, the electrostatic interactions become less negative [102]. The van der Waals interactions also become less negative [102]. Only the hydrogen bonding and bonded interactions (e.g., harmonic bond stretching, theta expansion bond angle, and so on) become more negative from 0 to 8 rings [102]. Thus, even though the membrane containing GDNT with eight cyclopentane rings is more compact, the resulting energy lowering effect is not due to the decrease in polar headgroup separation or the changes in the van der Waals interactions. Instead, it is due to the more favorable hydrogen bonding and bonded interactions. According to the MD calculations, the hydrogen bonding energy changes from –5.8 kcal/mol for zero ring to −29 kcal/mol for eight cyclopentane rings [102]. In short, the molecular modeling study showed that an increase in the number of cyclopentane rings in the isoprenoid chains not only tightens the membrane packing in the hydrophobic core, but also strengthens the hydrogen bonding at the membrane surface.

MEMBRANE LATERAL DIFFUSION AND LATERAL ORGANIZATION Lateral Diffusion Vaz et al. [103,104] used fluorescence recovery after photobleaching (FRAP) to measure the lateral diffusion of a NBD-labeled GDGT derived from S. solfataricus in fluid monopolar diester lipid vesicles. The lateral diffusion coefficient of the membrane-spanning NBD-GDGT lipid is strongly dependent upon the viscosity of the medium bounding the membrane and is about 2/3 that for a NBD-labeled monopolar diester lipid, which spans only one half of the bilayer membrane. These studies demonstrate that, like a monopolar diester lipid, a tetraether lipid is able to undergo lateral diffusion in the plane of the membrane. Kao et al. [65], on the other hand, attempted to demonstrate that a monopolar diester lipid can diffuse in the membrane with tetraether lipids as the matrix. The data of the pressure dependence of excimer formation suggested that the lateral mobility of 1-palmitoyl-2-(10-pyrenyl)docanoyl)-snglycero-3-phosphatidylcholine (PyrPC) in PLFE liposomes is highly restricted and only becomes appreciable at temperatures close to the minimal growth temperature of the cell. This conclusion was supported by two dimensional exchange 31P-nuclear magnetic resonance (NMR) measurements on tetraether liposomes made of the TPLE from T. acidophilum, which has optimum growth conditions

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of pH 2 and 55°C to 59°C [105]. The NMR data showed that the lateral diffusion coefficient (D) is 2 × 10 −8 cm2/s at 55°C and decreases to 6–8 × 10 −9 cm2/s at 30°C. The D value at 30°C reflects a membrane viscosity considerably higher than that of monopolar diester liposomes in the liquid crystalline phase. The D value at 55°C suggests that the membrane viscosity of T. acidophilum tetraether liposomes reaches a level typical for liquid crystalline phase when the temperature for D measurements is near the minimum growth temperature of the archaeon. MD calculations also showed that tetraether lipids have limited lateral mobility, especially at low temperatures [101]. The calculated lateral diffusion coefficient (D) of macrocyclic tetraether phosphatidylcholine (m-TEPC) at 25°C (Figure 6.4) [101] is comparable with that of tetraether liposomes derived from T. acidophilum measured by 31P-NMR [105]. The D value for m-TEPC at 25°C is lower than that of DPhPC at 25°C by one order of magnitude (Figure 6.4) [101]. In fact, at low temperatures (<20–24°C), lipid microdomains in PLFE liposomes visualized by two-photon excitation Laurdan fluorescence intensity microscopy were laterally immobile in the plane of the membrane, implying that there was little free volume in PLFE liposomes at those low temperatures (Figure 6.5) [41]. Lateral Organization Membrane lateral organization such as domain segregation and regular distributions is an intensively studied subject in bilayer membranes composed of conventional diester phospholipids, cholesterol, and glycosphingolipids [106–108]. However, there are few studies on lateral organization in lipid membranes composed of archaeal tetraether lipids. The functional importance of lateral organization in archaeal membranes has not been explored to any great extent. Two-photon excitation of Laurdan fluorescence images of GUVs composed of PLFE tetraether lipids isolated from S. acidocaldarius showed an homogenous distribution of the probe Laurdan above 24°C [41]. However, below 24°C, fluorescence-free, snow flake-like microdomains were observed [41]. The shape of the domains varies with pH. Those microdomains, which were believed to result from lipid segregation, were laterally immobile on the vesicle surface, implying that very little free volume was involved in the PLFE vesicles at low temperatures (Figure 6.5). Small microdomains were visualized in Langmuir lipid films composed of tetraether lipids from the archaeon T. acidophilum [53]. Domains were observable immediately after the films were formed and enlarged with increasing pressure [52,53], but disappeared after 12 h of spreading without compression [53]. This implies that lateral organization in this archaeal membrane system may not be stabilized in 12 h and that tight packing in archaeal lipid membranes enhances domain formation, consistent with the results from the temperature study on PLFE liposomes [41].

TECHNOLOGICAL APPLICATIONS OF TETRAETHER LIPID MEMBRANES Archaeal tetraether lipids form liposomes that are remarkably stable against attack by phospholipase A1 and A2, elevated temperature and pressure, and mechanical stress, and provide a highly resistant barrier against solute and proton leakage [11,44,109]. The unusually low-proton permeability detected in PLFE SUVs (~64 nm in diameter) [42] is of interest in nano-technological applications, because temperature- and pH-stable liposomes are highly desirable for storage, sterilization, and drug delivery. Tetraether lipids have been shown to increase vesicle stability against autoclaving [110], a commonly used sterilization technique. PLFE-based stealth liposomes can sustain at least six cycles of autoclaving in terms of retaining the size and shape of the vesicles (Brown, English, Cook, and Chong, unpublished results). Tetraether liposomes can be stored for at least six months at room temperature without showing a significant change in vesicles size [83]. At physiological [Ca2+] (~1–2 mM), PLFE liposomes are stable and un-fused (or nonaggregated) [83]. More than 92% of

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C-labeled sucrose can be retained in archaeosomes for two weeks at 4°C [111]. Archaeosomes composed of total lipid extract from M. hungatei and T. acidophilum can entrap 90% 5(6)-carboxyfluorescein for seven days at 65°C [110]. These studies suggested that tetraether liposomes are remarkably stable and may have a good drug-loading efficiency. Yet, they are not too tightly packed to release the entrapped materials. Liposomes made of TPLE from archaea are safe and do not lead to cytotoxicity in animals [112–114]. These features make tetraether lipids or liposomes attractive for pharmaceutical applications such as a superior alternative for vaccine, gene, and drug delivery vehicles [18,38,109,114]. Tetraether lipids may also be used as a tool for bio-electronics [115]. Tetraether lipids have been deposited on S-layer ultrafiltration membranes [116,117], which may serve as membraneprotein based biosensors for DNA sequencing and high throughput screening and may also be used in the lab-on-a-chip technology [117]. The ability to form cubic phases makes PLFE lipids appealing for crystallization of membrane-bound proteins [118], especially those found in hyperthermophiles. The possible obstacle is that large scale archaeal cell cultures are difficult to set up, mainly due to growth medium evaporation, low growth rate, low biomass yield, and corrosion to the fermenter [114]. The procedure for isolating TPLE from archaea is simple enough to lend itself to mass production for industrial applications. However, total lipid extract contains a large number of different lipid molecules with heterogeneity in the polar headgroups and/or in the number of cyclopentane rings in the polyisoprenoid chains. Thus, the yield of isolating a particular type of tetraether lipids from dry cells could be low. A yield of only ~4% has been reported for PLFE lipids from the thermoacidophilic archaeon S. acidocaldarius [119]. Synthesis of novel but high-yield archaeal bipolar lipid mimics [38] may circumvent these problems to some extent.

ACKNOWLEDGMENT The author thanks Samantha Tran for technical assistance and the support from the US Army Research Office (DAAD19-02-1-0077) and National Science Foundation (MCB-9513669 and DMR-0706410).

REFERENCES 1. Woese, C.R. and Fox, G.E., Phylogenetic structure of the prokaryotic domain: The primary kingdoms, Proc. Natl. Acad. Sci. U.S.A., 74, 5088, 1977. 2. De Rosa, M., Gambacorta, A., Nicolaus, B., Chappe, B., and Albrecht, P., Isoprenoid ethers: Backbone of complex lipids of the archaebacterium Sulfolobus solfataricus, Biochim. Biophys. Acta, 753, 249, 1983. 3. Langworthy, T.A. and Pond, J.L., Membranes and lipids of thermophiles, in Thermophiles: General, Molecular, and Applied Microbiology, Brock, T.D., Ed., John Wiley & Sons, New York, 1986, pp. 107–134. 4. Koga, Y. and Morii, H., Recent advances in structural research on ether lipids from Archaea including comparative and physiological aspects, Biosci. Biotechnol. Biochem., 69, 2019, 2005. 5. Sugai, A., Sakuma, R., Fukuda, I., Kurosawa, N., Itoh, Y.H., Kon, K., Ando, S., and Itoh, T., The structure of the core polyol of the ether lipids from Sulfolobus acidocaldarius, Lipids, 30, 339, 1995. 6. Kates, M., The phytanyl ether-linked polar lipids and isoprenoid neutral lipids of extremely halophilic bacteria, Prog. Chem. Fats Other Lipids, 15, 301, 1978. 7. Langworthy, T.A., Tornabene, T.G., and Holzer, G., Lipids of archaebacteria, Zbl. Bakt. Hyg., I. Abt. Orig., C3, 228, 1982. 8. Lanzotti, V., Nicolaus, B., Trincone, A., De Rosa, M., Grant, W.D., and Gambacorta, A., An isopranoid ether analogue of phosphatidic acid from a halophilic archaebacterium, Biochim. Biophys. Acta, 1002, 398, 1989.

9214_C006.indd 89

11/5/2007 11:42:33 AM

90

Thermophiles: Biology and Technology at High Temperatures

9. Sprott, G.D., Meloche, M., and Richards, J.C., Proportions of diether, macrocyclic diether, and tetraether lipids in Methanococcus jannaschii grown at different temperatures, J. Bacteriology 173, 3907, 1991. 10. Kates, M., Archaebacterial lipids: structure, biosynthesis and function, in The Archaebacteria: Biochemistry and Biotechnology, Danson, M.J., Hough, D.W., and Lunt, G.G., Eds., Portland Press, London, 1992, pp. 51–72. 11. Gliozzi, A., Relini, A., and Chong, P.L.-G., Structure and permeability properties of biomimetic membranes of bolaform archaeal tetraether lipids, J. Membr. Sci., 206, 131, 2002. 12. Stadnitskaia, A., Baas, M., Ivanov, M.K., Van Weering, T.C.E., and Sinninghe Damste, J.S., Novel archaeal macrocyclic diether core membrane lipids in a methane-derived carbonate crust from a mud vocano in the Sorokin Trough, NE Black Sea, Archaea, 1, 165, 2003. 13. Hafenbradl, D., Keller, M., and Stetter, K.O., Lipid analysis of Methanopyrus kandleri, FEMS Microbiol., 136, 199, 1996. 14. Nishihara, M., Morii, H., Matsuno, K., Ohga, M., Stetter, K.O., and Koga, Y., Structural analysis by reductive cleavage with LiAlH4 of an allyl ether choline-phospholipid, archaetidylcholine, from the hyperthermophilic methanoarchaeon Methanopyrus kandleri, Archaea, 1, 123, 2002. 15. Untersteller, E., Fritz, B., Bleriot, Y., and Sinay, P., The structure of calditol isolated from the thermoacidophilic archaebacterium Sulfolobus acidocaldarius, C. R. Acad. Sci. Paris II C2, 429, 1999. 16. Gambacorta, A., Caracciolo, G., Trabasso, D., Izzo, I., Spinella, A., and Sodano, G., Biosynthesis of calditol, the cyclopentanoid containing moiety of the membrane lipids of the archaeon Sulfolobus solfataricus, Tetrahedron Lett., 43, 451, 2002. 17. De Rosa, M. and Gambacorta, A., The lipids of archaebacteria, Prog. Lipid Res., 27, 153, 1988. 18. Gambacorta, A., Gliozzi, A., and De Rosa, M., Archaeal lipids and their biotechnological applications, World J. Microbiol. Biotechnol., 11, 115, 1995. 19. Itoh, Y.H., Kurosawa, N., Uda, I., Sugai, A., Tanoue, S., Itoh, T., Horiuchi, T., and Itoh, T., Metallosphaera sedula TA-2, a calditoglycerocaldarchaeol deletion strain of a thermoacidophilic archaeon, Extremophiles, 5, 241, 2001. 20. De Rosa, M., De Rosa, S., Gambacorta, A., Minale, L., and Bu’lock, J.D., Chemical structure of the ether lipids of thermophilic acidophilic bacteria of the Caldariella group, Phytochemistry, 6, 1961, 1979. 21. Ernst, M., Freisleben, H.-J., Antonopoulos, E., Henkel, L., Mlekusch, W., and Reibnegger, G., Calorimetry of archaeal tetraether lipid--indication of a novel metastable thermotropic phase in the main phospholipid from Thermoplasma acidophilum cultured at 59°C, Chem. Phys. Lipids, 94, 1, 1998. 22. De Rosa, M., Gambacorta, A., and Gliozzi, A., Structure, biosynthesis, and physicochemical properties of archaebacterial lipids, Microbiol. Rev., 50, 70, 1986. 23. Relini, A., Cassinadri, D., Mirghani, Z., Brandt, O., Gambacorta, A., Trincone, A., De Rosa, M., and Gliozzi, A., Calcium-induced interaction and fusion of archaeobacterial lipid vesicles: a fluorescence study, Biochim. Biophys. Acta, 1194, 17, 1994. 24. Relini, A., Cassinadri, D., Fan, Q., Gulik, A., Mirghani, Z., De Rosa, M., and Gliozzi, A., Effect of physical constraints on the mechanisms of membrane fusion: bolaform lipid vesicles as model systems, Biophys. J., 71, 1789, 1996. 25. Fuhrman, J.F., McCallum, K., and Davis, A.A., Novel major archaebacterial group from marine plankton, Nature, 356, 148, 1992. 26. DeLong, E.F., Wu, K.Y., Prezelin, B.B., and Jovine, R.V., High abundance of archaea in antarctic marine picoplankton, Nature, 371, 695, 1994. 27. Schouten, S., Hopmans, E.C., Pancost, R.D., and Sinninghe Damste, J.S., Widespread occurrence of structurally diverse tetraether membrane lipids: evidence for the ubiquitous presence of low-temperature relatives of hyperthermophiles, Proc. Natl. Acad. Sci. U.S.A., 97, 14421, 2000. 28. Sinninghe Damste, J.S., Hopmans, E.C., Schouten, S., van Duin, A.C.T., and Geenevasen, J.A.J., Crenarchaeol: the characteristic core glycerol dibiphytanyl glycerol tetraether membrane lipid of cosmopolitan pelagic crenarchaeota, J. Lipid Res., 43, 1641, 2002. 29. Menzel, D., Hopmans, E.C., Schouten, S., Sinninghe Damste, J.S., Membrane tetraether lipids of planktonic crenarchaeota in pilocene sapropels of the Eastern Mediterranean Sea, Palaeogeogr. Palaeoclimatol. Palaeoecol., 239, 1, 2006.

9214_C006.indd 90

11/5/2007 11:42:33 AM

Physical Properties of Membranes Composed of Tetraether Archael Lipids

91

30. Weijers, J.W.H., Schouten, S., Spaargaren, O.C., and Sinninghe Damste, J.S., Occurrence and distribution of tetraether membrane lipids in soils: implications for the use of the TEX86 proxy and the BIT index, Org. Geochem., 37, 1680, 2006. 31. Schouten, S., Hopmans, E.C., Schefub, E., and Sinninghe Damste, J.S., Distributional variations in marine crenarchaeotal membrane lipids: a new organic proxy for reconstructing ancient sea water temperatures? Earth Planetary Sci. Lett., 204, 265, 2002. 32. Weijers, J.W.H., Schouten, S., van der Donker, J.C., Hopmans, E.C., Sinninghe Damste, J.S., Environmental controls on bacterial tetraether membrane lipid distribution in soils, Geochim. Cosmochim. Acta, 71, 703, 2007. 33. Jahnke, L., Eder, W., Huber, R., Hope, J.M., Hinrichs, K., Hayes, J.M., Des Marais, D.J., Cady, S.L., and Summons, R.E., Signature lipids and stable carbon isotope analyses of Octopus Spring hyperthermophilic communities compared with those of Aquificales representatives, Appl. Environ. Microbiol., 67, 5179, 2001. 34. De Rosa, M., Gambacorta, A., Huber, R., Lanzotti, V., Nicolaus, B., Stetter, K.O., and Trincone, A., A new 15,16-dimethyl-30-glyceryloxytriacontanoic acid from lipids of Thermotoga maritime, J. Chem. Soc. Chem. Comm., 19, 1300, 1988. 35. Kim, J. and Thompson, D.H., Tetraether bolaform amphiphiles as models of archaebacterial membrane lipids: synthesis, differential scanning calorimetry, and monolayer studies, Langmuir, 8, 637, 1992. 36. Eguchi, T., Ibaragi, K., and Kakinuma, K., Total synthesis of archaeal 72-membered macrocyclic tetraether lipids, J. Org. Chem., 63, 2689, 1998. 37. Benvegnu, T., Brard, M., and Plusquellec, D., Archaebacteria bipolar lipid analogues: structure, synthesis and lyotropic properties, Curr. Opin. Colloid Interface Sci., 8, 469, 2004. 38. Benvegnu, T., Rethore, G., Brard, M., Richter, W., and Plusquellec, D., Archaeosomes based on novel synthetic tetraether-type lipids for the development of oral delivery systems, Chem. Commun., 5536, 2005. 39. Lelkes, P.I., Goldenberg, D., Gliozzi, A., De Rosa, M., Gambacorta, A., and Miller, I.R., Vesicles from mixtures of bipolar archaebacterial lipids with egg phosphatidylcholine, Biochim. Biophys. Acta, 732, 714, 1983. 40. Lo, S.L. and Chang, E.L., Purification and characterization of a liposomal-forming tetraether lipid fraction, Biochem. Biophys. Res. Commun., 167, 238, 1990. 41. Bagatolli, L.A., Gratton, E., Khan, T.K., and Chong, P.L.-G., Two-photon fluorescence microscopy studies of bipolar tetraether giant liposomes from thermoacidophilic archaebacteria Sulfolobus acidocaldarius, Biophys. J., 79, 416, 2000. 42. Komatsu, H. and Chong, P.L.-G., Low permeability of liposomal membranes composed of bipolar tetraether lipids from thermoacidophilic archaebacterium Sulfolobus acidocaldarius, Biochemistry, 37, 107, 1998. 43. Cavagnetto, F., Relini, A., Mirghani, Z., Bertoia, D., Gambacorta, A., and Gliozzi, A., Molecular packing parameters of bipolar lipids, Biochim. Biophys. Acta, 1106, 273, 1992. 44. Gliozzi, A. and Relini, A., Lipid vesicles as model systems for archaea membranes, in Handbook of Nonmedical Applications of Liposomes, Barenholz, Y. and Lasic, D.D., Eds., CRC Press, Boca Raton, FL, 1996, Vol. 2, pp. 329–348. 45. Elferink, M.G., de Wit, J.G., Demel, R., Driessen, A.J., and Konings, W.N., Functional reconstitution of membrane proteins in monolayer liposomes from bipolar lipids of Sulfolobus acidocaldarius. J. Biol. Chem., 267, 1375, 1992. 46. Elferink, M.G., De Wit, J.G., Driessen, A.J., and Konings, W.N., Energy-transducing properties of primary proton pumps reconstituted into archaeal bipolar lipid vesicles, Eur. J. Biochem., 214, 917, 1993. 47. In’t Veld, G., Elferink, M.G., Driessen, A.J., and Konings, W.N., Reconstitution of the leucine transport system of Lactococcus Lactis into liposomes composed of membrane-spanning lipids from Sulfolobus acidocaldarius, Biochemistry, 31, 12493, 1992. 48. Gleissner, M., Elferink, M.G., Driessen, A.J.M., Konings, W.N., Anemuller, S., and Schafer, G., Generation of proton-motive force by an archaeal terminal quinol oxidase from Sulfolobus acidocaldarius, Eur. J. Biochem., 224, 983, 1994.

9214_C006.indd 91

11/5/2007 11:42:33 AM

92

Thermophiles: Biology and Technology at High Temperatures

49. De Rosa, M., Gambacorta, A., and Nicolaus, B., A new yype of cell membrane in thermophilic archaebacteria based on bipolar ether lipids, J. Membr. Sci., 16, 287, 1983. 50. Morii, H. and Koga, Y., Asymmetrical topology of diether- and tetraether-type polar lipids in membranes of Methanobacterium thermoautotrophicum cells, J. Biol. Chem., 269, 10492, 1994. 51. Strobl, C., Six, L., Heckmann, K., Henkel, B., and Ring, K., Physicochemical characterization of tetraether lipids from Thermoplasma acidophilum. II Film balance studies on the monomolecular organization of the main glycophospholipid in monofilms, Z. Naturforsch, 40C, 219, 1985. 52. Gliozzi, A., Relini, A., Rolandi, R., Dante, S., and Gambacorta, A., Organization of bipolar lipids in monolayers at the air-water interface, Thin Solid Films, 242, 208, 1994. 53. Bakowsky, U., Rothe, U., Antonopoulos, E., Martini, T., Henkel, L., and Freisleben, H., Monomolecular organization of the main tetraether lipid from Thermoplasma acidophilum at the water-air interface, Chem. Phys. Lipids, 105, 31, 2000. 54. Heathcock, C.H., Finkelstein, B.L., Aoki, T., and Poulter, C.D., Sterostructure of the archaebacterial C40 diols, Science, 229, 862, 1985. 55. De Rosa, M., Archaeal lipids: structural features and supramolecular organization, Thin Solid Films, 286, 13, 1996. 56. Tomoaia-Cotisel, M., Chifu, E., Zsako, J., Mocanu, A., Quinn, P.J., and Kates, M., Monolayer properties of archaeol and caldarchaeol polar lipids of a methanogenic archaebacterium, Methanospirillum hungatei, at the air/water interface, Chem. Phys. Lipids, 63, 131, 1992. 57. Rolandi, R., Schindler, H., De Rosa, M., and Gambacorta, A., Monolayers of ether lipids from archaebacteria, Eur. Biophys. J., 14, 19, 1986. 58. Dote, J.L., Barger, W.R., Behroozi, F., Chang, E.L., Lo, S., Montague, C.E., and Nagumo, M., Monomolecular film behavior of tetraether lipids from a thermoacidophilic archaebacterium at the air/ water interface, Langmuir, 6, 1017, 1990. 59. Gulik, A., Luzzati, V., De Rosa, M., and Gambacorta, A., Tetraether lipid components from a thermoacidophilic archaebacterium. Chemical structure and physical polymorphism, J. Mol. Biol., 201, 429, 1988. 60. Gulik, A., Luzzati, V., De Rosa, M., and Gambacorta, A., Structure and polymorphism of bipolar isopranyl ether lipids from archaebacteria, J. Mol. Biol., 182, 131, 1985. 61. Luzzati, V., Gulik, A., De Rosa, M., and Gambacorta, A., Lipids from Sulfolobus solfataricus, life at high temperature and the structure of membranes, Chemica Scripta, 27B, 211, 1987. 62. Chong, P.L.-G., Ravindra, R., Khurana, M., English, V., and Winter, R., Pressure perturbation and differential scanning calorimetric studies of bipolar tetraether liposomes derived from the thermoacidophilic archaeon Sulfolobus acidocaldarius, Biophys. J., 89, 1841, 2005. 63. Chong, P.L.-G., Zein, M., Khan, T.K., and Winter, R., Structure and conformation of bipolar tetraether lipid membranes derived from thermoacidophilic archaeon Sulfolobus acidocaldarius as revealed by small-angle X-ray scattering and high pressure FT-IR spectroscopy, J. Phys. Chem., 107, 8694, 2003. 64. Khan, T.K. and Chong, P.L.-G., Studies of archaebacterial bipolar tetraether liposomes by perylene fluorescence, Biophys. J., 78, 1390, 2000. 65. Kao, Y.L., Chang, E.L., and Chong, P.L.-G., Unusual pressure dependence of the lateral motion of pyrene-labeled phosphatidylcholine in bipolar lipid vesicles, Biochem. Biophys. Res. Commun., 188, 1241, 1992. 66. Marrink, S.J. and Berendsen, H.J.C., Permeation process of small molecules across lipid membranes studied by molecular dynamics simulations, J. Phys. Chem., 100, 16729, 1996. 67. Haines, T.H. and Liebovitch, L.S., A molecular mechanism for the transport of water across phospholipid bilayers, in Permeability and Stability of Lipid Bilayers, Disalvo, E.A. and Simon, S.A., Eds., CRC, Boca Raton, FL, 1996; pp. 123–136. 68. Vilalta, I., Gliozzi, A., and Prats, M., Interfacial air/water proton conduction from long distances by Sulfolobus solfataricus archaeal bolaform lipids, Eur. J. Biochem., 240, 181, 1996. 69. Bartucci, R., Gambacorta, A., Gliozzi, A., Marsh, D., and Sportelli, L., Bipolar tetraether lipids: chain flexibility and membrane polarity gradients from spin-label electron spin resonance, Biochemistry, 44, 15017, 2005. 70. Mathai, J. C., Sprott, G.D., and Zeidel, M.L., Molecular mechanisms of water and solute transport across archaebacterial lipid membranes, J. Biol. Chem., 276, 27266, 2001.

9214_C006.indd 92

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Physical Properties of Membranes Composed of Tetraether Archael Lipids

93

71. Dannenmuller, O., Arakawa, K., Eguchi, T., Kakinuma, K., Blanc, S., Albrecht, A. M., Schmutz, M., Nakatani, Y., and Ourisson, G., Membrane properties of archaeal macrocyclic diether phospholipids, Chem. A Eur. J., 6, 645, 2000. 72. Deamer, D.W. and Nichols, J.W., Proton flux mechanisms in model and biological membranes, J. Membr. Biol., 107, 91, 1989. 73. Elferink, M.G.L., de Wit, J.G., Driessen, A.J.M., and Konings, W.N., Stability and proton-permeability of liposomes composed of archaeal tetraether lipids, Biochim. Biophys. Acta, 1193, 247, 1994. 74. Chang, E.L. Unusual thermal stability of liposomes made from bipolar tetraether lipids, Biochem. Biophys. Res. Commun., 202, 673, 1994. 75. Paula, S., Volkov, A.G., Van Hoek, A.N., Haines, T.H., and Deamer, D.W., Permeation of protons, potassium ions, and small polar molecules through phospholipid bilayers as a function of membrane thickness, Biophys. J., 70, 339, 1996. 76. Van de Vossenberg, J.L.C.M., Ubbink-Kok, T., Elferink, M.G.L., Driessen, A.J.M., and Konings, W.N., Ion permeability of the cytoplasmic membrane limits the maximum growth temperature of bacteria and archaea, Mol. Microbiol., 18, 925, 1995. 77. Yang, L. and Huang, W.H. Observation of a membrane fusion intermediate structure, Science, 297, 1877, 2002. 78. Blumenthal, R., Clague, M.J., Durell, S.R., and Epand, R.M. Membrane fusion, Chem. Rev., 103, 53, 2003. 79. Siegel, D.P., Burns, J.L., Chestnut, M.H., and Talmon, Y., Intermediates in membrane fusion and bilayer/ nonbilayer phase transitions imaged by time-resolved cryo-transmission electron microscopy, Biophys. J., 56, 161, 1989. 80. Basanez, G., Nieva, J.L., Rivas, E., Alonso, A., and Goni, F.M., Diacylglycerol and the promotion of lamellar-hexagonal and lamellar-isotropic phase transitions in lipids: implications for membrane fusion, Biophys. J., 70, 2299, 1996. 81. Wilschut, J., Duzgunes, N., Fraley, R., and Papahadjopoulos, D., Studies on the mechanism of membrane fusion: kinetics of calcium ion induced fusion of phosphatidylserine vesicles followed by a new assay for mixing of aqueous vesicle contents, Biochemistry, 19, 6011, 1980. 82. Ohki, S.A., Mechanism of divalent ion-induced phosphatidylserine membrane fusion, Biochim. Biophys. Acta, 689, 1, 1982. 83. Kanichay, R., Boni, L.T., Cooke, P.H., Khan, T.K., and Chong, P.L.-G., Calcium-induced aggregation of archaeal bipolar tetraether liposomes derived from thermoacidophilic archaeon Sulfolobus acidocaldarius, Archaea, 1, 175, 2003. 84. Elferink, M.G.L., van Breemen, J., Konings, W.N., Driessen, A.J.M., and Wilschut, J., Slow fusion of liposomes composed of membrane-spanning lipids, Chem. Phys. Lipids, 88, 37, 1997. 85. Sundler, R. and Papahadjopoulos, D., Control of membrane fusion by phospholipid head groups. I. phosphatidate/phosphatidylinositol specificity, Biochim. Biophys. Acta, 649, 743, 1981. 86. Nagle, J.F. and Scott, H.L., Lateral compressibility of lipid mono- and bilayers. Theory of membrane permeability, Biochim. Biophys. Acta, 513, 236, 1978. 87. Olsen, G.J., Microbial ecology. Archaea, archaea, everywhere, Nature, 371, 657, 1994. 88. Bailey, A., Zhukovsky, M., Gliozzi, A., and Chernomordik, L.V., Liposome composition effects on lipid mixing between cells expressing influenza virus hemagglutinin and bound liposomes, Arch. Biochem. Biophys., 439, 211, 2005. 89. Melikyan, G.B., Matinyan, N.S., Kocharov, S.L., Arakelian, V.B., Prangishvili, D.A., and Nadareishvili, K.G., Electromechanical stability of planar lipid membranes from bipolar lipids of the thermoacidophilic archaebacterium Sulfolobus acidocaldarius, Biochim. Biophys. Acta, 1068, 245, 1991. 90. Sprott, G.D., Patel, G.B., and Krishnan, L., Archaeobacterial ether lipid liposomes as vaccine adjuvants, Methods Enzymol., 373, 155, 2003. 91. Bagatolli, L.A. and Gratton, E., Two-photon fluorescence microscopy observation of shape changes at the phase transition in phospholipid giant unilamellar vesicles, Biophys. J., 77, 2090, 1999. 92. Bagatolli, L.A. and Gratton, E., Two photon fluorescence microscopy of coexisting lipid domains in giant unilamellar vesicles of binary phospholipid mixtures, Biophys. J., 78, 290, 2000. 93. Thompson, T.E. and Huang, C., Composition and dynamics of lipids in biomembranes, in Membrane Physiology, Andreoli, T.E., Hoffman, J.F., Fanestil, D.D., and Schultz, S.G., Eds., Plenum Publishing, New York, NY, 1986; Chapter 2, pp. 25–44.

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94. Bruno, S., Cannistraro, A., Gliozzi, M., De Rosa, M., and Gambacorta, A., A spin label ESR and saturation transfer-ESR study of archaebacteria bipolar lipids, Eur. Biophys. J., 13, 67, 1985. 95. Shinoda, K., Shinoda, W., Baba, T., and Mikami, M., Comparative molecular dynamics study of etherand ester-linked phospholipid bilayers, J. Chem. Phys., 121, 9648, 2004. 96. Gawrisch, K., Ruston, D., Zimmerberg, J., Parsegian, V.A., Rand, R.P., and Fuller, N., Membrane dipole potentials, hydration forces, and the ordering of water at membrane surfaces, Biophys. J., 61, 1213, 1992. 97. Kitano, T., Onoue, T., and Yamauchi, K., Archaeal lipids forming a low energy-surface on air-water interface, Chem. Phys. Lipids, 126, 225, 2003. 98. Shinoda, W., Mikami, M., Baba, T., and Hato, M., Dynamics of a highly branched lipid bilayer: a molecular dynamics study, Chem. Phys. Lett., 390, 35, 2004. 99. Shinoda, W., Mikami, M., Baba, T., and Hato, M., Molecular dynamics study on the effect of chain branching on the physical properties of lipid bilayers: structural stability, J. Phys. Chem. B., 107, 14030, 2003. 100. Shinoda, W., Mikami, M., Baba, T., and Hato, M., Molecular dynamics study on the effects of chain branching on the physical properties of lipid bilayers: 2. Permeability, J. Phys. Chem. B., 108, 9346, 2004. 101. Shinoda, W., Shinoda, K., Baba, T., and Mikami, M., Molecular dynamics study of bipolar tetraether lipid membranes, Biophys. J., 89, 3195, 2005. 102. Gabriel, J.L. and Chong, P.L.-G., Molecular modeling of archaebacterial bipolar tetraether lipid membranes, Chem. Phys. Lipids, 105, 193, 2000. 103. Vaz, W.L.C., Hallmann, D., Clegg, R.M., Gambacorta, A., and De Rosa, M., A comparison of the translational diffusion of a normal and a membrane-spanning lipid in L α phase 1-palmitoyl-2-oleoylphosphatidylcholine bilayers, Eur. Biophys. J., 12, 19, 1985. 104. Vaz, W.L.C., Stumpel, J., Hallmann, D., Gambacorta, A., and De Rosa, M., Bounding fluid viscosity and translational diffusion in a fluid lipid bilayer, Eur. Biophys. J., 15, 111, 1987. 105. Jarrell, H.C., Zukotynski, K.A., and Sprott, G.D., Lateral diffusion of the total polar lipids from Thermoplasma acidophilum in multilamellar liposomes, Biochim. Biophys. Acta, 1369, 259, 1998. 106. Chong, P.L.-G. and Sugar, I.P., Fluorescence studies of lipid regular distribution in membranes, Chem. Phys. Lipids, 116, 153, 2002. 107. Kusumi, A., Koyama-Honda, I., and Suzuki, K., Molecular dynamics and interactions for creation of stimulation-induced stabilized rafts from small unstable steady-state rafts, Traffic, 5, 213, 2004. 108. Pucadyil, T.J. and Chattopadhyay, A., Role of cholesterol in the function and organization of G-protein coupled receptors, Progr. Lipid Res., 45, 295, 2006. 109. Patel, G.B. and Sprott, G.D., Archaeobacterial ether lipid liposomes (archaeosomes) as novel vaccine and drug delivery systems, Crit. Rev. Biotechnol., 19, 317, 1999. 110. Choquet, C.G., Patel, G.B., and Sprott, G.D., Heat sterilization of archaeal liposomes, Can. J. Microbiol, 42, 183, 1996. 111. Konings, W.N., Tolner, B., Speelmans, G., Elferink, M.G.L., de Wit, J.G., and Driessen, A.J.M., Energy transduction and transport processes in thermophilic bacteria, J. Bioenerg. Biomembr., 24, 601, 1992. 112. Freisleben, H., Bormann, J., Litzinger, D.C., Lehr, F., Rudolph, P., Schatton, M., and Huang, L., Toxicity and biodistribution of liposomes of the main phospholipid from the archaebacterium Thermoplasma acidophilum, J. Liposome Res., 5, 215, 1995. 113. Deamer, D.W. and Nichols, J.W., Proton-hydroxide permeability of liposomes, Proc. Natl. Acad. Sci. U.S.A., 80, 165, 1983. 114. Schiraldi, C., Giuliano, M., and De Rosa, M., Perspectives on biotechnological applications of archaea, Archaea, 1, 75, 2002. 115. De Rosa, M., Morana, A., Riccio, A., Gambacorta, A., Trincone, A., and Incani, O., Lipids of the archaea: A new tool for bio-electronics, Biosens. Bioelectron., 9, 669, 1994. 116. Pum, D., Wetzer, B., Schuster, B., and Sleytr, U.B., S-layers as patterning structures and supporting layers for biomimetic membranes, SPIE Proc., 2978, 53, 1997. 117. Schuster, B., Weigert, S., Pum, D., Sara, M., and Sleytr, U.B., New method for generating tetraether lipid membranes on porous supports, Langmuir, 19, 2392, 2003.

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118. Landau, E.M. and Rosenbusch, J.P., Lipidic cubic phases: a novel concept for the crystallization of membrane proteins, Proc. Natl. Acad. Sci. U.S.A., 93, 14532, 1996. 119. Chang, E.L. and Lo, S.L., Extraction and purification of tetraether lipids from Sulfolobus acidocaldarius, in Protocols for Archaebacterial Research, Maryland Biotechnology Institute, Baltimore, MD, 1991, pp. 2.3.1–2.3.14.

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Part III Heat-Stable Enzymes and Metabolism

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Glycolysis in Hyperthermophiles Peter Schönheit

CONTENTS Introduction ................................................................................................................................ Classical EM Pathway in Eukarya and Bacteria ....................................................................... Modified EM Pathways and Acetate Formation in Hyperthermophilic Archaea ..................... Unusual Enzymes of Modified EM Pathways in Hyperthermophilic Archaea .............. Glucose Phosphorylation to Glucose-6-Phosphate .............................................. Glucose-6-Phosphate Isomerization to Fructose-6-Phosphate ............................ Fructose-6-Phosphate Phosphorylation ............................................................... Cleavage of FBP .................................................................................................. GAP Oxidation .................................................................................................... Conversion of 3-Phosphoglycerate to PEP .......................................................... PEP Conversion to Pyruvate in Hyperthermophilic Archaea ............................. Enzymes of Pyruvate Conversion to Acetate ................................................................. Pyruvate Conversion to Acetyl CoA .................................................................... Acetyl-CoA Conversion to Acetate ...................................................................... Energetics of Modified EM Pathways ............................................................................ Classical EM Pathway and Acetate Formation in the Hyperthermophilic Bacterium Thermotoga maritima ................................................................................... Enzymes of the Classical EM Pathway in Thermotoga maritima ................................. Enzymes of Pyruvate Conversion to Acetate ................................................................. Energetics of Glucose Degradation to Acetate in Thermotoga maritima ...................... Conclusions ................................................................................................................................ References ..................................................................................................................................

99 100 101 103 103 103 104 104 105 105 105 105 105 106 106 106 107 107 108 108 110

INTRODUCTION Hyperthermophiles with an optimal growth temperature above 80°C and a maximum at 113°C [1] are considered to represent the phylogenetic most ancestral living organisms. According to their position in the single-stranded unit (SSU) ribosomal RNA (rRNA)-based phylogenetic tree, hyperthermophiles show the deepest branching and shortest lineages of all organisms analyzed. Most hyperthermophiles belong to the domain of archaea, only few hyperthermophilic genera of bacteria are known, which include, for example, Thermotogales. The metabolism of hyperthermophiles is diverse including both chemolithoautotrophs dependent on compounds derived from vulcanic activities, for example, H2, CO2, and various sulfur compounds, and chemo-organoheterotrophs growing, for example, on peptides or sugars [2]. Many sugar utilizing hyperthermophiles are strictly anaerobic fermentative organisms degrading glucose or glucose polymers (starch, maltose, cellobiose) to acetate as main fermentation product. These include, for example, the euryarchaeaota 99

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Pyrococcus furiosus, Thermococcus sp., and the crenarchaeon Desulfurococcus amylolyticus. Recently, the sulfate reducing hyperthermophilic archaeon Archaeoglobus fulgidus strain 7324 was found to degrade starch incompletely to acetate reducing sulfate to H2S [3]. The crenarchaeota Thermoproteus tenax and Pyrobaculum aerophilum completely oxidize sugars to CO2 with sulfur and O2 or nitrate, respectively, as external electron acceptors [4]. The best studied sugar utilizing hyperthermophilic bacterium is Thermotoga maritima; the organism ferments glucose and glucose polymers to acetate as main products as do fermentative hyperthermophilic archaea [5,6]. Comparative analysis of glycolytic pathways in hyperthermophiles revealed that all archaeal species do not degrade glucose to pyruvate via the classical Embden-Meyerhof (EM) and EntnerDoudoroff (ED) pathways operative in eukarya and bacteria, but instead use modified versions of these pathways. In thermoacidophilic aerobic archaea, in the hyperthermophilic Sulfolobus sp. in the moderate thermophilic Thermoplasma acidophilum and Picrophilus torridus, and to a small extent (<20%) in the anaerobic sulfur reducer T. tenax, glucose is metabolized via modifications of the ED pathway. For a recent detailed discussion of ED modification in (hyper)thermophilic archaea see References [4,7–11]. The hyperthermophilic fermentative anaerobes P. furiosus, Thermococcus sp., D. amylolyticus, the sulfur reducer T. tenax (>80% of glucose degraded via EM pathway), the sulfate reducer A. fulgidus strain 7324, and the microaerophilic P. aerophilum use different modifications of the EM pathway [4,11]. In this chapter, the modified EM pathways of selected hyperthermophilic archaea, P. furiosus, Thermococcus sp., A. fulgidus strain 7324 and the crenarchaeota D. amylolyticus, T. tenax, and P. aerophilum are described. In particular, unusual enzymes and enzyme families catalyzing the phosphorylation of glucose, the isomerization of glucose-6-phosphate, the phosphorylation of fructose-6-phosphate, the oxidation of glyceraldehyde-3-phosphate (GAP), and the formation of pyruvate will be discussed, as well as regulatory and energetic aspects. Since most of the hyperthermophilic archaea ferment sugars to acetate, the mechanism of acetate formation will also be analyzed. As it will be shown next, the enzymes of the EM pathways and of acetate formation in hyperthermophilic archaea differ considerably from the classical versions operative in eukarya and (mesophilic and moderate thermophilic) bacteria. It has been speculated that these differences might be adaptation to extreme high temperatures, near 100°C, the growth temperatures of hyperthermophilic archaea. Alternatively the differences might be due to the phylogenetic status of these hyperthermophiles being archaea. Thus, for comparison, the enzymes of glycolysis and of acetate formation were also described for the hyperthermophilic bacterium T. maritima. With an optimal growth temperature >80°C (Tmax ~90°C) T. maritima is a true hyperthermophile, which, however, belongs to the deepest branches of the domain of bacteria. Like fermentative hyperthermophilic archaea, the strictly anaerobic T. maritima ferments glucose and glucose polymers to acetate. The comparison of hyperthermophilic archaea and the hyperthermophilic bacterium T. maritima might give an indication to differentiate the influence of temperature and phylogeny on unusual enzymes of EM pathways and of acetate formation found in hyperthermophilic archaea. Detailed reviews on various aspects of sugar metabolism of archaea have recently appeared [2,4,10–13]. Before describing features of glycolysis of hyperthermophilic archaea and the bacterium Thermotoga, a short summary of the classical EM pathway operative in most eukarya and (mesophilic and moderate thermophilic) bacteria will be given.

CLASSICAL EM PATHWAY IN EUKARYA AND BACTERIA The EM pathway (synonym glycolysis) is the most common pathway in saccharolytic organisms catalyzing the degradation of glucose to pyruvate. The pathway (“classical EM pathway”), which has been analyzed in detail in eukarya and bacteria, is highly conserved with respect to reactions catalyzed and the enzymes (and enzyme families) involved, its regulatory properties, and its energetics [14]. The pathway comprises 10 enzymes, three of them catalyze irreversible reactions and are sites for allosteric control. Seven enzymes catalyze reversible reactions which therefore can be used in the reverse direction for gluconeogenesis.

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Glucose is taken up in eukarya either by facilitated diffusion or secondary (H+, Na+)/glucose symport systems. Secondary ion symport systems for glucose uptake are also found in many aerobic bacteria. The phosphorylation of glucose to glucose 6-phosphate in eukarya is catalyzed by ATPdependent hexokinases, which show a broad specificity for hexoses (glucose, fructose, and mannose). Some hexokinases show allosteric features and are inhibited by glucose-6-phosphate [15]. In bacteria, for example, Escherichia coli and Bacillus subtilis, glucose phosphorylation is catalyzed by ATP-dependent glucokinases, which belong either to the glucokinase or (ROK) repressor protein, open reading frame, sugar kinase glucokinase families both of which are highly specific for glucose and are usually not regulated by effectors [16]. Many anaerobic and facultative bacteria take up glucose via the phosphoenolpyruvate (PEP)-dependent phosphotransferase (PTS)-like transport system, generating glucose-6-phosphate as the result of group translocation [17]. The isomerization of glucose-6-phosphate to fructose-6-phosphate is catalyzed in almost all eukarya and bacteria by one type of phosphoglucose isomerase (PGI), which belong to the PGI superfamily. Fructose-6-phosphate is phosphorylated to fructose-1,6-bisphosphate by ATPdependent phosphofructokinases (ATP-PFK), which in eukarya and bacteria belong to the PFK-A family. The homotetrameric enzymes are usually allosterically regulated by the energy charge and compounds of intermediary metabolism. In bacteria, ATP-PFKs are activated by ADP and inhibited by PEP. In eukarya regulation of ATP-PFKs is more complex, including in addition, for example, citrate and fructose-2,6-bisphosphate as effectors. In few eukarya and bacteria fructose-6phosphate is phosphorylated by pyrophosphate (PP i)-dependent PFKs, which are also members of the PFK-A family. Fructose-1,6-bisphosphate (FBP) formed is cleaved to GAP and dihydroxyacetone phosphate (DHAP) by Schiff base-dependent Class I or metal-dependent Class II type FBP aldolases. Subsequent isomerization of DHAP to GAP via triosephosphate isomerase yields 2 mol of GAP. The oxidation of GAP to 3-phosphoglycerate involves two enzymes, phosphorylative glyceraldehyde-3-phosphate dehydrogenase (GAPDH) catalyzing the phosphate and NAD + -dependent GAP oxidation to 1,3-bisphosphoglycerate, which is converted to 3-phosphoglycerate by phosphoglycerate kinase yielding 1 ATP per mole of GAP by substratelevel phosphorylation. The isomerization of 3-phosphoglycerate to 2-phosphoglycerate involves two structural and mechanistical distinct types of phosphoglycerate mutases. One type, dPGM is dependent on the cofactor 2,3-bisphosphoglycerate for activity, whereas the second type, iPGM, is independent of this cofactor. Enolase catalyzes the dehydration of 2-phosphoglycerate to PEP. The subsequent conversion of PEP to pyruvate is catalyzed by pyruvate kinase (PK) and is coupled to ATP synthesis via substrate-level phosphorylation. PKs in eukarya and bacteria are usually homotetrameric enzymes that represent another site of allosteric control being activated by AMP or FBP or, as in some protists by fructose-2,6-bisphosphate, and are allosterically inhibited by ATP. The net ATP yield of the “classical EM pathway” is 2 mol of ATP per mole glucose. Pyruvate formed by glycolysis is converted to acetyl-CoA in eukarya and aerobic bacteria by pyruvate dehydrogenase complex, acetyl CoA is then oxidized to 2CO2 via the citric acid cycle. In many anaerobic bacteria, pyruvate is oxidized to acetyl-CoA via pyruvate ferredoxin oxidoreductase (POR). In several facultative and strictly anaerobic bacteria, pyruvate is converted to formate and acetyl-CoA by pyruvate formate lyase (PFL). Acetyl-CoA formed by POR or PFL is converted by bacteria under anaerobic conditions to acetate involving two classical enzymes, phosphate acetyl transferase and acetate kinase. In the following, the EM pathways as well as the enzymes in acetate formation will be described for selected hyperthermophilic archaea and the hyperthermophilic bacterium T. maritima.

MODIFIED EM PATHWAYS AND ACETATE FORMATION IN HYPERTHERMOPHILIC ARCHAEA In archaea, glucose is degraded to pyruvate via modified EM pathways shown in Figure 7.1. The modifications found in EM pathways in hyperthermophiles implicate either novel reactions

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Thermophiles: Biology and Technology at High Temperatures Archaea Pyrococcus, Thermococcus 2– Archaeoglobus 7324 (SO4 ) Glucose GLK

ADP AMP

Desulfurococcus

ATP ADP

Bacteria Pyrobaculum – (O2/NO3 )

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ATP

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cPGI ADP AMP

ATP ADP

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+

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+

Fdox

Fdox

Fdox

NAD

Fdred

Fdred

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GAPDH

1.3 BPG ADP

PGK

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ADP ATP

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ADP ATP

ADP ATP

ADP ATP

Pi CoA

Ac-CoA ACD Acetate

ADP + P

ADP + P

ATP + CoA

ATP + CoA

PTA

Ac - P ADP ATP

AK

FIGURE 7.1 Embden-Meyerhof (EM) pathways and the mechanism of acetate formation in hyperthermophilic archaea and the hyperthermophilic bacterium Thermotoga maritima. Selected enzymes (arrows) of the modified EM pathways in hyperthermophilic archaea, catalyzing glucose and fructose-6-phosphate phosphorylation, phosphoglucose isomerization, glyceraldehyde-3-phosphate oxidation, and phosphoenolpyruvate conversion to pyruvate are compared with the corresponding enzymes of the “classical” EM pathway in T. maritima. The conversion of acetyl-CoA to acetate in archaea is catalyzed by one enzyme, acetyl-CoA synthetase (ADP forming) (ACD) (acetyl-CoA + ADP + Pi ↔ acetate + ATP + CoA), in the bacterium T. maritima by two enzymes, phosphotransacetylase (PTA) and acetate kinase (AK); the oxidation of acetyl-CoA to 2CO2 in hyperthermophilic archaea Thermoproteus tenax with sulfur (S) and Pyrobaculum aerophilum with oxygen or nitrate (O2, NO − 3 ) as electron acceptors proceeds via the tricarboxylic acid cycle (TCA cycle) (not shown). Abbreviations: G-6-P, glucose-6-phosphate; F-6-P, fructose-6-phosphate; PFK, 6-phosphofructokinase; F-1,6-BP, fructose-1,6-bisphosphate; DHAP, dihydroxyacetone phosphate; GAP, glyceraldehyde3-phosphate; 3-PG, 3-phosphoglycerate; 2-PG, 2-phosphoglycerate; PEP, phosphoenolpyruvate; Fdox and Fdred, oxidized and reduced ferredoxin; GLK, glucokinase (ADP- or ATP-dependent); PGI, phosphoglucose isomerase (cPGI, cupin PGI; PGI/PMI, bifunctional phosphoglucose/phosphomannose isomerase); PFK, phosphofructokinase (PPi-, ADP- or ATP-dependent); GAPOR, glyceraldehyde-3 phosphate-ferredoxin oxidoreductase; GAPN, nonphosphorylative glyceraldehyde-3-phosphate dehydrogenase; PK, pyruvate kinase; GAPDH, phosphorylative glyceraldehyde-3-phosphate dehydrogenase, PGK, phosphoglycerate kinase; CoA, coenzyme A; Ac-CoA, acetyl coenzyme A; Ac-P, acetyl phosphate.

(ADP-dependent kinases and GAP-oxidizing enzymes) not found in the classical EM pathway. Further, several enzymes catalyzing formally identical reactions belong to different enzyme families and superfamilies. The modified EM pathways in the euryarchaeota Pyrococcus furiosus, Thermococcus sp., and A. fulgidus strain 7324 are similar, whereas larger variations of enzymes are found in EM pathways of the crenarchaeota D. amylolyticus, T. tenax, and P. aerophilum. All enzymes of a modified EM pathway similar to those of P. aerophilum were also found in the microaerophilic crenarchaeon Aeropyrum pernix except that the the latter contains nonphosphorylative

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glyceraldehyde-3-phosphate dehydrogenase (GAPN) rather than glyceraldehyde-3-phosphateferredoxin oxidoreductase (GAPOR) [18].

UNUSUAL ENZYMES OF MODIFIED EM PATHWAYS IN HYPERTHERMOPHILIC ARCHAEA In the following, the enzymes involved in phosphorylation of glucose and fructose-6-phosphate of glucose-6-phosphate isomerization of GAP oxidation and of pyruvate formation will be discussed. For references on triose phosphate isomerase, fructose-1,6-bisphosphate aldolase, phosphoglycerate mutases and enolases see References [4,11,12,19]. Glucose Phosphorylation to Glucose-6-Phosphate All hyperthermophilic archaea did not contain the bacteria-like PEP-dependent PTS-like transport systems. Thus, the initial step of archaeal EM pathways is the phosphorylation of glucose, which is catalyzed by variety of kinases, which differ in specificity for sugars and phosphoryl donors. In the euryarchaeota P. furiosus, Thermococcus sp., and A. fulgidus strain 7324, novel glucokinases were identified, which use ADP as phosphoryl donor forming AMP. The homodimeric (P. furiosus [20]) or monomeric (A. fulgidus [21]) proteins exhibit a high specificity for glucose. Structural analysis of the enzymes from Thermococcus litoralis and Pyrococcus horikoshii [4] indicate that ADPdependent glucokinases belong to the ribokinase superfamily. In the crenarchaeota A. pernix [22], P. aerophilum [23], and T. tenax [24] ATP-dependent glucokinases were identified. They constitute monomeric enzymes with an unusual broad specificity for the hexoses glucose, fructose, and mannose, as has been reported before only for hexokinases from eukarya. The archaeal ATPdependent glucokinases are members of the ROK family (repressor protein, open reading frame, sugar kinase), which belong to the actin-ATPase superfamily (SCOP) structural classification of proteins. Both ADP-dependent and ATP-dependent glucokinases from hyperthermophilic archaea are not regulated by effectors. Glucose-6-Phosphate Isomerization to Fructose-6-Phosphate The isomerization of glucose 6-phosphate to fructose-6-phosphate in almost all eukarya and bacteria is catalyzed by one type of PGI which constitutes the classical PGI superfamily (more than 1000 sequences known, Pfam). Homologs of the classical PGI were not found in sugar-degrading hyperthermophilic archaea; one homolog is, however, present in hyperthermophilic lithoautrophic methanogen Methanocaldococcus jannaschii probably as a result of lateral gene transfer from the hyperthermophilic bacterium T. maritima [25]. In the hyperthermophilic euryarchaeota P. furiosus, T. litoralis, and A. fulgidus a novel type of PGI has been identified and characterized. These PGIs designated cupin-PGIs (cPGIs) belong to the cupin superfamily and thus represent a convergent line of PGI evolution [26]. The crystal structure of cPGI from P. furiosus revealed a typical cupin fold, which contains a central domain composed of β-strands forming a small β-barrel called “cupin” [27,28]. Phylogenetic analysis suggests an euryarchaeotal origin of cPGIs [26]. Unlike all other known PGIs, cPGIs require divalent cations for activity, in vivo most likely Fe2+ [26]. In the crenarchaeota A. pernix and P. aerophilum, and the moderate thermophile Thermoplasma acidophilum, an unusual PGI was described, which differs from all knows PGIs by catalyzing the isomerization of both glucose-6-phosphate and mannose-6-phosphate at similar catalytic efficiency. Thus, the enzyme was designated as bifunctional phosphoglucose/phosphomannose isomerase (PGI/PMI) [29,30]. Usually, members of bacteria and eukarya use separate enzymes, showing either PGI or PMI activity. PGI/PMIs constitute a novel family within the PGI superfamily, which was proven by the crystal structure of the P. aerophilum enzyme, showing a typical PGI fold [31]. A structural basis for bifunctionality was proposed [32]. A PGI/PMI homolog is also present in T. tenax [33]. The structures of the two novel convergently evolved PGIs in hyperthermophilic archaea, of PGI/ PMI from P. aerophilum, as a member of the PGI superfamily (PGI fold), and of dimeric metal containing cPGI from P. furiosus (cupin fold) are shown in Figure 7.2.

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Fructose-6-Phosphate Phosphorylation The phosphorylation of fructose-6-phosphate to fructose-1,6-bisphosphate in hyperthermophilic archaea is catalyzed by kinases, which differ with respect to their phosphoryl donor ADP and ATP or PPi. The hyperthermophilic euryarchaeota P. furiosus, T. zilligii, and A. fulgidus 7324 contain ADP-dependent phosphofructokinases (ADP-PFK), which together with ADP-dependent glucokinases constitute the ADP-PFK/GLK family within the ribokinase superfamily. ADP-PFKs constitute homotetrameric enzymes (P. furiosus) or can be isolated as a mixture of functional tetrameric and dimeric forms (A. fulgidus, T. zilligii) [20,34,35]. A monomeric bifunctional ADP-GLK/PFK was found in the hyperthermophilic methanogen M. jannaschii [36]. ATP-PFK are present in the hyperthermophilic crenarchaeota D. amylolyticus and from A. pernix. These archaeal ATP-PFKs are homotetrameric enzymes, which belong to the PFK-B family [37–39]. The PPi-dependent PFK from T. tenax catalyzes a reversible reaction, and is functionally involved both in glycolysis and in gluconeogenesis [4]. The archaeal PPi-dependent PFK, which is a homotetrameric enzyme, is a member of the phosphofructokinase (PFK-A) family and thus is related to classical ATP-dependent PFKs of eukarya and bacteria [40]. All archaeal ADP-, ATP-, or PPi-dependent PFKs showed Michaelis Menten kinetics with respect to the substrate fructose-6-phosphate, indicating the absence of cooperative substrate binding. Further, allosteric regulation by classical effectors of bacterial and eukaryal ATP-dependent PFKs, was not observed indicating that these enzymes do not represent a site of allosteric control in archaeal glycolysis of hyperthermophiles.

Cleavage of FBP FBP is converted to GAP and DHAP by FBP aldolase. FBP aldolases in the hyperthermophilic archaea P. furiosus and T. tenax did not show significant sequence similarity to known eukaryal and bacterial Class I and II FBP aldolases and thus were designated as a novel family of archaeal type Class I FBP aldolases [41]. In P. aerophilum no obvious homologs of this archaeal Class I FPB aldolase could be identified. Triose phosphate isomerase converts DHAP to GAP. The enzyme is highly conserved in hyperthermophilic and other archaea, bacteria, and eukarya [12].

FIGURE 7.2 Convergent evolution of a glycolytic enzyme, phosphoglucose isomerase in hyperthermophilic archaea. (a) Crystal structure of dimeric metal-dependent cupin-phosphoglucose isomerase (PGI) from Pyrococcus furiosus (cupin fold). (From Swan, M.K. et al., J. Biol. Chem. 278, 47261, 2003). (b) Of dimeric PGI/phosphomannose isomerase (PMI) from Pyrobaculum aerophilum, a member of the PGI superfamily (PGI fold). (From Swan, M.K. et al., J. Biol. Chem., 279, 39838, 2004. With permission.)

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GAP Oxidation The oxidation of GAP in EM pathways of the hyperthermophilic archaea P. furiosus, A. fulgidus strain 7324, and P. aerophilum is catalyzed by GAPOR [4,18]. The enzyme catalyzes the phosphateindependent oxidation of GAP to 3-phosphoglycerate with ferredoxin as electron acceptor and is not coupled to ATP formation via substrate-level phosphorylation. GAPOR has been characterized from P. furiosus as a monomeric tungsten-pterin containing Fe/S protein, which belongs to the aldehyde ferredoxin oxidoreductase superfamily [42]. GAPOR activity is induced after growth on sugars, and for P. furiosus regulation of GAPOR on the level of transcription has been demonstrated [4]. In the modified EM pathway of T. tenax GAP is oxidized to 3-phosphoglycerate by GAPN, which catalyzes the irreversible oxidation of GAP with NAD(P)+. The enzyme was found to be allosterically regulated by the energy charge and by intermediates of the EM pathway [43]. GAPN is a member of the aldehyde dehydrogenase superfamily. Thus, in the modified EM pathways of hyperthermophilic archaea, the irreversible GAP oxidation to 3-phosphoglycerate catalyzed by one enzyme, GAPOR or GAPN, replaces the classical, reversible two-step conversion in the classical EM pathway of bacteria and eukarya involving phosphorylative GAPDH and phosphoglycerate kinase. It should be mentioned that both P. furiosus and T. tenax also contain low-anabolic activities of GAPDH and phosphoglycerate kinase, which, however, are operative during gluconeogenesis [44,45]. Gluconeogenesis in all hyperthermophilic archaea and all other organisms proceeds via reversible reactions of the EM pathway [4]. Conversion of 3-Phosphoglycerate to PEP The conversion of 3-phosphoglycerate to PEP in hyperthermophilic archaea is catalyzed by reversible phosphoglycerate mutase (PGM) and enolase [4,12]. As in bacteria and eukarya, two distinct PGM types, 2,3-bisphosphate-dependent dPGMs, and cofactor-independent iPGMs have been identified in hyperthermophilic archaea. iPGMs have been characterized, for example, in P. furiosus and A. fulgidus and an archaeal dPGM from the thermoacidophile T. acidophilum have been characterized [4,19]. PEP Conversion to Pyruvate in Hyperthermophilic Archaea All hyperthermophilic archaea contain PK catalyzing the irreversible formation of pyruvate from PEP coupled to ATP synthesis (pyruvate + ADP → PEP + ATP), PKs have been characterized from T. tenax, A. fulgidus, A. pernix, and P. aerophilum. In contrast to bacterial and eukaryal PKs, these hyperthermophilic archaeal PKs, which are all homotetrameric enzymes, exhibit reduced regulatory potential. Although cooperative binding of the substrates PEP and ADP was demonstrated for several of these hyperthermophilic PKs, classical heterotropic allosteric regulators of bacteria and eukarya, for example, AMP and fructose-1,6-bisphosphate did not affect PK activity [46,47]. Thus, PKs of the modified EM pathway in hyperthermophilic archaea apparently do not represent sites of (heterotrophic) allosteric control. Besides PK, PEP synthetase (PEP + P + AMP ↔ pyruvate + ATP), which represents an anabolic enzyme in bacteria, has recently been proposed to have a catabolic function in the modified EM pathway of P. furiosus [13] and Thermococcus kodakaraensis [48]. In T. tenax, pyruvate phosphate dikinase (ATP + Pi + pyruvate ↔ AMP + PPi + PEP) has also been suggested to have a glycolytic role in addition to PK [49].

ENZYMES OF PYRUVATE CONVERSION TO ACETATE Pyruvate Conversion to Acetyl CoA All hyperthermophilic archaea, both strictly anaerobic and microaerophilic species convert pyruvate to acetyl CoA via POR. POR from the hyperthermophiles P. furiosus and T. litoralis and A. fulgidus have been characterized as highly thermoactive heterotetramers [50].

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Acetyl-CoA Conversion to Acetate Several anaerobic fermentative hyperthermophilic archaea (Pyrococcus, Thermococcus, Desulfurococcus) degrade sugars to acetate as major fermentation product (Figure 7.1). In addition, the sulfate reducer A. fulgidus strain 7324 generates acetate via incomplete oxidation of starch, reducing sulfate to H2S. In these hyperthermophilic archaea the formation of acetate from acetyl CoA is catalyzed by a novel prokaryotic enzyme, acetyl-CoA synthetase (ADP forming) (ACD, acetyl-CoA + ADP + Pi ↔ acetate + ATP + CoA). This unusual synthetase catalyzes the conversion of acetyl CoA to acetate and couples this reaction with the formation of ATP from ADP and phosphate (Pi) by substrate-level phosphorylation [51]. Acetyl phosphate is not a free intermediate of this reaction. ACDs belong to the recently recognized superfamily of nucleotide diphosphate (NDP) forming acyl-CoA synthetases, which also include succinyl-CoA synthetases [52]. ACDs from several hyperthermophilic archaea, including P. furiosus and A. fulgidus, have been characterized [4].

ENERGETICS OF MODIFIED EM PATHWAYS The formal net ATP yields of all modified EM pathways, that is, the conversion of l mol glucose to 2 mol pyruvate, in hyperthermophilic archaea is zero, as nonphosphorylative GAPOR or GAPN are not coupled with ATP syntheses via substrate level phosphorylation (Figure 7.1). In T. tenax the ATP yield of the EM pathway might be up to 1 ATP, if it is assumed that PPi, which is formed in the cell by various anabolic reactions, is not hydrolyzed by pyrophosphatase but rather used by PPi-dependent PFK. In the EM pathway of P. furiosus and T. kodakaraensis additional ATP might be formed (up to 2 mol ATP/mol pyruvate), assuming that PEP conversion to pyruvate is catalyzed—in addition to PK—via reversible PEP synthetase operating in the glycolytic direction (PEP + AMP + P ↔ pyruvate + ATP). Recent disruption experiments of PEP synthetase gene in T. kodakaraensis strongly support a glycolytic role of PEP synthetase in this organism [48]. Another site of ATP formation has been proposed for P. furiosus in the course of H2 formation released by fermentation via membrane-bound ferredoxin-dependent hydrogenase. In P. furiosus H+-translocation and the formation of fractions of ATP via electron transport phosphorylation in the course of H2 formation have been demonstrated [53]. ATP formation in the modified EM pathway of P. furiosus has also been proposed on the basis of growth yield data [54]. In all fermentative hyperthermophilic archaea forming acetate, a major energy-conserving site is ADP-forming acetyl-CoA synthetase generating 1 mol ATP/mol acetate during acetyl-CoA conversion to acetate via substrate-level phosphorylation.

CLASSICAL EM PATHWAY AND ACETATE FORMATION IN THE HYPERTHERMOPHILIC BACTERIUM THERMOTOGA MARITIMA It has been speculated that, in hyperthermophilic archaea, (i) the presence of GAPOR, GAPN, and ACD, which all do not involve potentially thermolable intermediates during catalysis, that is, the phosphoacid anhydrides 1,3-bisphosphoglycerate or acetyl phosphate, and (ii) the presence of PFKs and PKs, which do not show allosteric control, might represent an adaptation to hyperthermophilic growth conditions. To test this hypothesis, the enzymes of the EM pathway and of acetate formation were analyzed in the hyperthermophilic bacterium T. maritima. T. maritima [5] belongs to the deepest branches of the bacterial domain. With a temperature optimum for growth above 80°C and a maximum around 90°C the organism is classified as a hyperthermophile. T. maritima utilizes a variety of sugars, including glucose as growth substrates [55]. Growing cultures of the strictly anaerobe ferment l mol glucose to 2 mol acetate, 2 mol CO2, and 4 mol H2 as products [6]. 13C-nuclear magnetic resonance (NMR) labeling experiments in cell suspensions and enzyme measurements indicate that glucose is degraded to pyruvate predominantly

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(85%) by the classical EM pathway. About 15% of glucose is degraded via the classical, phosphorylated ED pathway [56].

ENZYMES OF THE CLASSICAL EM PATHWAY IN THERMOTOGA MARITIMA Due to the absence of PTS system in T. maritima glycolysis starts with free glucose. The phosphorylation of glucose to glucose-6-phosphate is catalyzed by a ROK-type glucokinase. In contrast to the characterized archaeal ROK glucokinases from hyperthermophilic crenarchaeota A. pernix, P. aerophilum, and T. tenax, which are monomeric enzymes with broad hexose specificity, the bacterial ROK glucokinase from T. maritima is a dimeric enzyme showing high specificity for glucose a substrate [16]. Fructose and mannose were not utilized. The isomerization of glucose-6phosphate to fructose-6-phosphate is catalyzed by one type of PGI, the classical PGI, also found in eukarya and bacteria, which belongs to the PGI superfamily. The recombinant enzyme has been characterized as a homodimeric protein showing extremely high temperature optimum (87°C) and thermostability around 90°C [57], in accordance with the optimal growth temperature of the hyperthermophile. Homologs of the PGI/PMI family and cupin-type PGI, found in hyperthermophilic archaea, were not present in the bacterium T. maritima. The phosphorylation of fructose-6phosphate to FBP in T. maritima is catalyzed by an ATP-dependent PFK, which belongs to the PFK-A family, comprising almost all bacterial and eukaryal PFKs. The homotetrameric T. maritima enzyme showed a high temperature optimum and thermostability. Enzyme activity showed sigmoidal response to the substrate fructose-6-phosphate, indicating cooperative binding of the substrate. Further, the ATP-PFK showed the classical allosteric response to classical effectors; it was activated by AMP, and was inhibited by ATP. The allosteric response could only be detected at high temperature 75°C, near the temperature optimum of the enzyme [58]. T. maritima also contained a PP-dependent PFK [59], which, however, has a minor role in glycolysis [58]. The conversion of FBP to DHAP and GAP and the subsequent isomerization of DHAP to GAP are catalyzed by FBP aldolase and triose phosphate isomerase, which are present in high activities in cell extracts. In the genome of T. maritima, a gene for putative metal-dependent Class II aldolase was found rather than a homolog of archaeal Schiff base Class I aldolase found in hyperthermophilic archaea. The oxidation of GAP to 3-phosphogylcerate involves the classical two enzymes, phosphorylative NAD+ and phosphate-dependent GAPDH and phosphoglycerate kinase (Figure 7.1). Both enzymes are present in high activities in glucose-grown T. maritima cells [6] and the encoding gene are apparently transcriptionally regulated [55]. GAPDH and phosphoglycerate kinase have been purified [60,61]. Glucose-grown T. maritima cells contain high activities of a cofactor-dependent phosphoglycerate mutase catalyzing the 2,3-bisphosphate-dependent conversion of 3-phosphoglycerate to 2-phosphoglycerate [6]. In the genome of T. maritima, both dPGM and iPGM encoding genes are present. Glucose-grown cells also contained high enolase activity catalyzing the dehydration of 2-phosphoglycerate to PEP. The conversion of PEP to pyruvate is catalyzed by PK. The enzyme was characterized as homotetrameric, highly thermoactive, and thermostable protein with an extremely high melting temperature above 98°C, as analyzed by circular dichroism (CD) spectroscopy [47]. The enzyme showed sigmoidal rate dependence on the substrates PEP and ADP indicating cooperative binding of substrates. In contrast to PKs from hyperthermophilic archaea, the Thermotoga PK showed the classical response to allosteric effectors; it was activated by AMP and inhibited by ATP. Thus, both ATP-PFK and PK represent sites of allosteric control of EM pathway of T. maritima.

ENZYMES OF PYRUVATE CONVERSION TO ACETATE Pyruvate is converted to acetyl-CoA in T. maritima by POR, which has been characterized as heterotetrameric enzyme of high thermostability [50]. The formation of acetate from acetyl-CoA is catalyzed by the classical mechanism found in all acetate-forming bacteria analyzed so far. The conversion involves two enzymes, phosphotransacetylase (PTA) (acetyl-CoA + phosphate ↔

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acetyl-phosphate + CoA) and acetate kinase (AK) (acetyl-phosphate + ADP ↔ acetate + ATP) (Figure 7.1), which are highly conserved enzymes in all bacteria. PTA belongs to the isocitrate/ isopropylmalate dehydrogenase-like superfamily, AK to the actin-like ATPase domain superfamily (SCOP). AK and PTA from T. maritima have been characterized as homodimeric and homotetrameric enzymes, respectively, showing the highest temperature optimum for catalytic activity and the highest thermostability of all PTAs and AKs analyzed so far [62]. A homolog of ACD, catalyzing acetate formation in hyperthermophilic archaea, was not found in T. maritima.

ENERGETICS OF GLUCOSE DEGRADATION TO ACETATE IN THERMOTOGA MARITIMA Growing cultures of T. maritima convert 1 mol glucose to 2 mol acetate, 2 mol CO2, and 4 mol H2. Growth yield data indicate an in vivo ATP yield of 4 mol ATP per mole of 2 mol of acetate formed from glucose [6]. As 2 mol of ATP are formed per 2 mol of acetate in the AK reaction, 2 mol ATP have to be formed during glucose conversion to 2 mol pyruvate in accordance with the operation of the classical EM pathway involving GAP oxidation to 3-phosphoglycerate via GAPDH and PGK, in which 1 mol ATP/mol GAP is formed by substrate-level phosphorylation (Figure 7.1). In summary, T. maritima contains all enzymes of the classical EM pathway including GAPDH/ PGK couple catalyzing GAP oxidation and allosterically regulated ATP-PFKs and PKs. The enzymes are all well adapted to hyperthermophilic conditions showing extremely high temperature optima for catalytic activity and high thermostability. In addition, the allosteric behavior of PFK and PKs is also not impaired by the extremely high temperature. Thus, the classical EM pathway, which is highly conserved in bacteria and eukarya, is also operative under hyperthermophilic conditions indicating that extremely high temperatures do not cause a change in the enzyme set involved. In addition, the mechanism of acetate formation in hyperthermophilic bacterium involves the classical two-enzyme mechanism, via PTA and AK, typical for all acetate-forming bacteria. Thus, free phosphoacid anhydrides involved in glucose degradation to acetate in T. maritima, that is, 1,3-bisphosphate formed by GADPH, and acetyl phosphate formed by PTA, are apparently not thermolable in vivo.

CONCLUSIONS The comparative analysis of glycolytic pathway and enzymes of acetate formation in hyperthermophilic archaea and the bacterium T. maritima indicates that the variations found in modified EM pathways of archaea and the unusual one-enzyme mechanism of acetate formation do not represent an adaptation to a hyperthermophilic life style but rather are typical features of the archaeal domain. This was concluded from the findings that in T. maritima, which is also a hyperthermophile but belongs to the domain of bacteria, both the classical EM pathways and the classical two-enzyme mechanism of acetate formation are operative. In accordance with this conclusion are recent findings that several of the unusual glycolytic enzymes in hyperthermophilic archaea have homologs in mesophilic archaea, and also in few bacteria and eukarya. These include, for example, ADP-dependent glucokinases and ADP-dependent PFKs [20], cupin PGIs [26], PGI/PMIs, and GAPOR [4]. ACD, the unusual mechanism of acetate formation is also operative in mesophilic extreme halophilic archaea and has also been found in few anaerobic eukaryotic protists. Few enzymes, for example, cupin-PGI and PGI/PMI are also found in mesophilic bacteria probably due to lateral gene transfer events [26,29]. Recently, a functional ADP-dependent glucokinase has been characterized from mouse [63]. The reasons for the remarkable diversity of enzymes established in the modified EM pathway in hyperthermophilic archaea are not understood. During archaeal evolution several mechanisms might have been involved in generating glycolytic enzyme diversity, including convergent evolution, lateral gene transfer, and recruitment of existent enzyme families which attain a glycolytic function, for example, after gene duplication and diversification. A remarkable example for convergent

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evolution of a glycolytic enzyme is the metal-dependent cupin PGI predominantly found in euryarchaeota. The enzyme belongs to the cupin superfamily as proven by the crystal structure solved for P. furiosus (Figure 7.2a). In contrast, the second existing functional type of PGI in hyperthermophilic archaea is the bifunctional PGI/PMI, which belongs to the PGI superfamily as indicated by the PGI-fold (Figure 7.2b) In archaea, this PGI family developed a bifunctionality as PGI/PMI. ADP-GLKs and ADP-PFKs, which form a novel family within the ribokinase superfamily, evolved a novel specificity for ADP as phosphoryl donor. Finally, PP-dependent PFK in the archaeon T. tenax belongs to the PFK-A family, which comprise both ATP- and PP-dependent PFKs of eukarya and bacteria. The presence of PP-PFK in the hyperthermophilic archaeon T. tenax might best be explained by lateral gene transfer from bacterial PP-PFKs. Other PP-PFKs and ATP-PFK homologs are not found in hyperthermophilic archaea. Instead, functional ATP-PFKs in glycolysis of hyperthermophilic archaea are obviously recruited from existing PFK-B family, which belong to the ribokinase superfamily. Members of the PFK-B family comprise a large number of ATP-dependent sugar kinases, ubiquitous in all domains of life, which function as kinases for nucleosides and various sugars, rather than constituting functional ATP-dependent fructose-6-phosphate kinase [37]. Interestingly, in hyperthermophilic archaea PFK-B homologs with different substrate specificities have been characterized. The PFK-B from M. jannaschii is a true nucleoside (adenosine) kinase and does not phosphorylate fructose-6-phosphate [37]. The structure has been solved showing typical ribokinase fold, very similar to ribokinase from E. coli [64]. (Note that M. jannaschii contains an ADP-PFK which might be involved in glycogen degradation [36]). The PFK-B homolog in A. pernix exhibits similar activities for both, adenosine kinase and fructose-6-phosphate kinase, whereas the homolog in D. amylolyticus is a true ATP-PFK showing almost no nucleoside kinase activity [37]. Thus, in the archaeal domain PFK-B proteins evolved to the first functional ATP-PFKs within a glycolytic pathway. Further, GAPOR, which belongs to the aldehyde ferredoxin oxidoreductase family, constitutes a glycolytic enzyme oxidizing GAP to 3-phosphoglycerate. GAPN is a member of the aldehyde dehydrogenase superfamily being ubiquitously distributed in all domains of life catalyzing the oxidation of a variety of aldehydes [8]. In bacteria and eukarya, GAPN serves an anabolic role primarily generating NADPH coupled to GAP oxidation, whereas in the archaeal domain GAPN changed into a glycolytic enzyme catalyzing catabolic GAP oxidation to 3-phosphoglycerate. Finally, ADP-forming acetyl-CoA synthetase, a member of the NDP-forming acyl-CoA synthetase superfamily, which also contains ubiquitous succinyl CoA synthetase, represents in archaea a functional acetyl-CoA synthetase catalyzing acetyl-CoA conversion to acetate. This one-step conversion replaces the two-enzyme mechanism via PTA and AK, which are typical bacterial enzymes and which are not present in acetate-forming archaea. (The presence of AK/PTA in Methanosarcina species is probably due to lateral gene transfer from bacterial homologs). Although, the presence of genes and enzymes might in part be explained as described before, the functional involvement of several unusual enzymes in glycolysis of hyperthermophilic archaea cannot be explained. These include, for example, the ADP dependency of sugar kinases, the hexokinase-like sugar specificity of archaeal ROK glucokinases, the bifunctionality of PGI/ PMIs, as well as the absence of allosteric sites at the level of PFKs and PKs. In general, multifunctional enzymes provide an advantage of a higher metabolic flexibility comprising more than one enzyme functions, however, at the expense of a loss of independent regulation of the separate activities. Why hyperthermophilic archaea use GAPOR or GAPN to oxidize GAP to 3-phosphoglycerate instead of the classical two-enzyme mechanism of GAP-DH/PGK is not known. This is even more surprising as the latter two enzymes are also present in hyperthermophilic archaea, however, operating only in the direction of gluconeogenesis. Why GAPDH/PGK does not function in the glycolytic direction in hyperthermophilic archaea is not understood. The one-step conversion of GAP to 3-PGS not coupled to ATP synthesis via substrate-level phosphorylation might cause a higher, uncoupled, rate of GAP oxidation and thus a higher flux of archaeal glycolysis.

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With respect on the evolution of glycolysis it has been proposed that the primary function of the EM pathway is that of gluconeogenesis [12]. This has been concluded, for example, from distribution and phylogenies of several reversible enzymes of the EM pathway including enolase, phosphoglycerate mutase, phosphoglycerate kinase, phosphorylative GAPDH, and triosephosphate isomerase. These enzymes are highly conserved in all domains of life, suggesting gluconeogenesis to represent an ancestral highly conserved metabolic route. This is in accordance with a chemolithoautotrophic mode of life of most óf the ancestral hyperthermophilic organisms, which are living, for example, on compounds derived from vulcanic activities, such as H2, CO2, low O2 concentrations, and a variety of sulfur compounds. Later in evolution, when carbohydrates became available, for example, from biomass generated by chemolithoautotrophic growth, glycolysis appears to be invented several times as indicated by the highly diverse mosaic-like structure of archaeal glycolysis. This can be explained by convergent evolution of glycolytic genes, by lateral gene transfer of glycolytic genes from other organisms, and by changes of existing nonglycolytic genes (after duplication) to generate a specific function in glycolysis.

REFERENCES 1. Stetter, K.O., Hyperthermophiles in the history of life, Phil. Trans. R. Soc. B, 361, 1837, 2006. 2. Schönheit, P. and Schäfer, T., Metabolism of hyperthermophiles, World J. Microbiol. Biotechnol., 11, 26, 1995. 3. Labes, A. and Schönheit, P., Sugar utilization in the hyperthermophilic, sulfate-reducing archaeon Archaeoglobus fulgidus strain 7324: starch degradation to acetate and CO2 via a modified EmbdenMeyerhof pathway and acetyl-CoA synthetase (ADP-forming), Arch. Microbiol., 176, 329, 2001. Erratum in Arch. Microbiol., 177, 431, 2002. 4. Siebers, B. and Schönheit, P., Unusual pathways and enzymes of central carbohydrate metabolism in Archaea, Curr. Opin. Microbiol., 8, 695, 2005. 5. Huber, R. et al., Thermotoga maritima sp. nov. represents a new genus of unique extremely thermophilic eubacteria growing up to 90°C, Arch. Microbiol., 144, 324, 1986. 6. Schröder, C., Selig, M., and Schönheit, P., Glucose fermentation to acetate CO2 and H2 in the anaerobic hyperthermophilic eubacterium Thermotoga maritima: involvement of the Embden-Meyerhof pathway, Arch. Microbiol., 161, 460, 1994. 7. Ahmed, H., et al., The semi-phosphorylative Entner-Doudoroff pathway in hyperthermophilic archaea— a re-evaluation, Biochem. J., 390, 529, 2005. 8. Reher, M. and Schönheit, P., Glyceraldehyde dehydrogenases from the thermoacidophilic euryarchaeota Picrophilus torridus and Thermoplasma acidophilum, key enzymes of the non-phosphorylative EntnerDoudoroff pathway, constitute a novel enzyme family within the aldehyde dehydrogenase superfamily, FEBS Lett., 580, 1198, 2006. 9. Reher, M., Bott, M., and Schönheit, P., Characterization of glycerate kinase (2-phosphoglycerate forming), a key enzyme of the nonphosphorylative Entner-Doudoroff pathway, form the thermoacidophilic euryarchaeon Picrophilus torridus, FEMS Microbiol. Lett., 259, 113, 2006. 10. Danson, M.J. and Hough, D.W., Promiscuity in the Archaea. The enzymology of metabolic pathways, The Biochemist, 27, 17, 2005. 11. Verhees, C.H. et al., The unique features of glycolytic pathways in Archaea, Biochem. J., 375, 231, 2003. Erratum in: Biochem. J., 377, 819, 2004. 12. Ronimus, R.S. and Morgan, H.W., Distribution and phylogenies of enzymes of the Embden-MeyerhofParnas pathway from archaea and hyperthermophilic bacteria support a gluconeogenic origin of metabolism, Archaea, 1, 199, 2003. 13. Sakuraba, H., Goda, S., and Ohshima, T., Unique sugar metabolism and novel enzymes of hyperthermophilic archaea, Chem. Rec., 3, 281, 2004. 14. Fothergill-Gilmore, L.A. and Michels, P.A., Evolution of glycolysis, Prog. Biophys. Mol. Biol., 59, 105, 1993. 15. Cárdenas, M.L., Cornish-Bowden, A., and Ureta, T., Evolution and regulatory role of the hexokinases, Biochim. Biophys. Acta, 1401, 242, 1998. 16. Hansen, T. and Schönheit, P., ATP-dependent glucokinase from the hyperthermophilic bacterium Thermotoga maritima represents an extremely thermophilic ROK glucokinase with high substrate specifity, FEMS Microbiol. Lett., 226, 405, 2003.

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17. Deutscher, J., Francke, C., and Postma, P.W., How phosphotransferase system-related protein phosphorylation regulates carbohydrate metabolism in bacteria, Microbiol. Mol. Biol. Rev., 70, 939, 2006. 18. Reher, M., Gebhard, S., and Schönheit, P., Glyceraldehyde-3-phosphate ferredoxin oxidoreductase (GAPOR) and nonphosphorylating glyceraldehyde-3-phosphate dehydrogenase (GAPN), key enzymes of the respective modified Embden-Meyerhof pathways in the hyperthermophilic crenarchaeta Pyrobaculum aerophilum and Aeropyrum pernix, FEMS Microbial. Lett., 273, 196, 2007. 19. Johnsen, U. and Schönheit, P., Characterization of cofactor dependent and cofactor independent phosphoglycerate mutases from archaea, Extremophiles, 2007 in press. 20. Kengen, S.W.M. et al., ADP-dependent glucokinase and phosphofructokinase from Pyrococcus furiosus, Methods Enzymol., 331, 41, 2001. 21. Labes, A. and Schönheit, P., ADP-dependent glucokinase from the hyperthermophilic sulfate-reducing archaeon Archaeoglobus fulgidus strain 7324, Arch. Microbiol., 180, 69, 2003. 22. Hansen, T. et al., The first archaeal ATP-dependent glucokinase, from the hyperthermophilic crenarchaeon Aeropyrum pernix, represents a monomeric, extremely thermophilic ROK glucokinase with broad hexose specificity, J. Bacteriol., 184, 5955, 2002. 23. Hansen, T. and Schönheit, P., unpublished data. 24. Dörr, C. et al., The hexokinase of the hyperthermophile Thermoproteus tenax: ATP-dependent hexokinases and ADP-dependent glucokinases, two alternatives for glucose phosphorylation in Archaea, J. Biol. Chem., 278, 18744, 2003. 25. Rudolph, B., Hansen, T., and Schönheit, P., Glucose-6-phosphate isomerase from the hyperthermophilic archaeon Methanococcus jannaschiiI: characterization of the first archaeal member of the phosphoglucose isomerase superfamily, Arch. Microbiol., 181, 82, 2004. 26. Hansen, T. et al., Cupin-type phosphoglucose isomerases (cupin PGIs) constitute a novel metal dependent PGI family representing a convergent line of PGI evolution, J. Bacteriol., 187, 1621, 2005. 27. Swan, M.K. et al., Structural evidence for a hydride transfer mechanism of catalysis in phosphoglucose isomerase from Pyrococcus furiosus, J. Biol. Chem., 278, 47261, 2003. 28. Berrisford, J.M. et al, Crystal structure of Pyrococcus furiosus phosphoglucose isomerase. Implications for substrate binding and catalysis, J. Biol. Chem., 278, 33290, 2003. 29. Hansen, T., Wendorff, D., and Schönheit, P., Bifunctional phosphoglucose/phosphomannose isomerases from the Archaea Aeropyrum pernix and Thermoplasma acidophilum constitute a novel enzyme family within the phosphoglucose isomerase superfamily, J. Biol. Chem., 279, 2262, 2003. 30. Hansen, T., Urbanke, C., and Schönheit, P., Bifunctional phosphoglucose/phosphomannose isomerases from the hyperthermophilic archaeaon Pyrobaculum aerophilum, Extremophiles, 8, 507, 2004. 31. Swan, M.K. et al., A novel phosphoglucose isomerase (PGI)/phosphomannose isomerase from the crenarchaeon Pyrobaculum aerophilum is a member of the PGI superfamily: structural evidence at 1.16-A resolution, J. Biol. Chem., 279, 39838, 2004. 32. Swan, M.K. et al., Structural basis for phosphomannose isomerase activity in phosphoglucose isomerase from Pyrobaculum aerophilum: a subtle difference between distantly related enzymes. Biochemistry, 43, 14088, 2004. 33. Siebers, B. et al., Reconstruction of the central carbohydrate metabolism of Thermoproteus tenax by use of genomic and biochemical data, J. Bacteriol., 186, 2179, 2004. 34. Hansen, T. and Schönheit, P., ADP-dependent 6-phosphofructokinase, an extremely thermophilic, nonallosteric enzyme from the hyperthermophilic, sulfate-reducing archaeon Archaeoglobus fulgidus strain 7324, Extremophiles, 8, 29, 2004. 35. Ronimus, R.S. and Morgan, H.W., The biochemical properties and phylogenies of phosphofructokinases form extremophiles. Extremophiles, 5, 357, 2001. 36. Sakuraba, H. et al., ADP-dependent glucokinase/phosphofructokinase, a novel bifunctional enzyme from the hyperthermophilic Archaeon Methanococcus jannaschii, J. Biol. Chem., 277, 12495, 2002. 37. Hansen, T. et al., The phosphofructokinase-B (MJ0406) from Methanocaldococcus jannaschii represents a nucleoside with a broad substrate specifity, Extremophiles, 11, 105, 2007. 38. Hansen, T. and Schönheit, P., Purification and properties of the first-identified, archaeal, ATP-dependent 6-phosphofructokinase, an extremely thermophilic non-allosteric enzyme, from the hyperthermophile Desulfurococcus amylolyticus, Arch. Microbiol., 173, 103, 2000. 39. Hansen, T. and Schönheit, P., Sequence, expression, and characterization of the first archaeal ATPdependent 6-phosphofructokinase, a non-allosteric enzyme related to the phosphofructokinase-B sugar kinase family, from the hyperthermophilic crenarchaeote Aeropyrum pernix, Arch. Microbiol., 177, 62, 2001.

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40. Siebers, B., Klenk, H.P., and Hensel, R., PPi-dependent phosphofructokinase from Thermoproteus tenax, an archaeal descendant of an ancient line in phosphofructokinase evolution, J. Bacteriol., 180, 2137, 1998. 41. Siebers, B. et al., Archaeal fructose-1,6-bisphosphate aldolases constitute a new family of archaeal type class I aldolase, J. Biol. Chem., 276, 28710, 2001. 42. Roy, R., Menon, A.L., and Adams, M.W.W., Aldehyde oxidoreductases from Pyrococcus furiosus, Methods Enzymol., 331, 132, 2001. 43. Brunner, N.A. et al., NAD+ -dependent glyceraldehyde-3-phosphate dehydrogenase from Thermoproteus tenax. The first identified archaeal member of the aldehyde dehydrogenase superfamily is a glycolytic enzyme with unusual regulatory properties, J. Biol. Chem., 273, 6149, 1998. 44. Schäfer, T. and Schönheit, P., Gluconeogenesis from pyruvate in the hyperthermophilic archaeon Pyrococcus furiosus: involvement of reactions of the Embden-Meyerhof pathway, Arch. Microbiol., 159, 354, 1993. 45. Brunner, N.A. Siebers, B., and Hensel, R., Role of two different glyceraldehyde 3-phosphate dehydrogenases in controlling the reversible Embden-Meyerhof-Parnas pathway in Thermoproteus tenax: regulation of protein and transcript level, Extremophiles, 5, 101, 2001. 46. Schramm, A. et al., Pyruvate kinase of the hyperthermophilic crenarchaeote Thermoproteus tenax: physiological role and phylogenetic aspects, J. Bacteriol., 182, 2001, 2000. 47. Johnsen, U., Hansen, T., and Schönheit, P., Comparative analysis of pyruvate kinases from the hyperthermophilic archaea Archaeoglobus fulgidus, Aeropyrum pernix, and Pyrobaculum aerophilum and the hyperthermophilic bacterium Thermotoga maritima: unusual regulatory properties in hyperthermophilic archaea, J. Biol. Chem., 278, 25417, 2003. 48. Imanaka, H. et al., Phosphoenolpyruvate synthase plays an essential role for glycolysis in the modified Embden-Meyerhof pathway in Thermococcus kodakarensis, Mol. Microbiol., 61, 898, 2006. 49. Tjaden, B. et al., Phosphoenolpyruvate synthetase and pyruvate, phosphate dikinase of Thermoproteus tenax: key pieces in the puzzles of archaeal carbohydrate metabolism, Mol. Microbiol., 60, 287, 2006. 50. Schut, G.J., Menon, A.L., and Adams, M.W.W., 2-Keto acid oxidoreductase from Pyrococcus furiosus and Thermococcus litoralis, Methods Enzymol., 331, 144, 2001. 51. Schäfer, T., Selig, M., and Schönheit, P., Acetyl-CoA synthethase (ADP-forming) in archaea, a novel enzyme involved in acetate and ATP synthesis, Arch. Microbiol., 159, 72, 1993. 52. Sanchez, L.B., Acetyl-CoA synthetase from the amitochondriate eukaryote Giardia lamblia belongs to the newly recognized superfamily of acyl-CoA synthetases (nucleoside diphosphate-forming), J. Biol. Chem., 275, 5794, 2000. 53. Sapra, R., Bagramyan, K., and Adams, M.W., A simple energy-conserving system: proton reduction coupled to proton translocation, Proc. Natl. Acad. Sci. U S A, 100, 7545, 2003. 54. Kengen, S.W.M. and Stams A.J.M., Growth and energy conservation in batch cultures of Pyrococcus furiosus. FEMS Microbiol. Lett., 117, 305, 1994. 55. Conners, S.B. et al., Microbial biochemistry, physiology, and biotechnology of hyperthermophilic Thermotoga species, FEMS Microbial. Rev., 30, 872, 2006. 56. Selig, M. et al., Comparative analysis of Embden-Meyerhof and Entner-Doudoroff glycolytic pathways in hyperthermophilic archaea and the bacterium Thermotoga, Arch. Microbiol., 167, 217, 1997. 57. Schlichting, B. and Schönheit, P., unpublished data. 58. Hansen, T., Musfeldt, M., and Schönheit, P., ATP-dependent 6-phosphofructokinase from the hyperthermophilic bacterium Thermotoga maritima: characterization of an extremely thermophilic, allosterically regulated enzyme, Arch. Microbiol., 177, 401, 2002. 59. Ding, Y.H., Ronimus, R.S., and Morgan, H.W., Thermotoga maritima phosphofructokinases: expression and characterization of two unique enzymes, J. Bacteriol., 183, 791, 2001. 60. Wraba, A. et al., Extremely thermostable D-glyceraldehyde-3-phosphate dehydrogenase from the eubacterium Thermotoga maritima, Biochemistry, 29, 7584, 1990. 61. Crowhurst, G., McHarg, J. and Littlechild, J.A., Phosphoglycerate kinases from Bacteria and Archaea, Methods Enzymol., 331, 90, 2001. 62. Bock, A.-K. et al., Purification and characterization of two extemely thermostable enzymes, phosphate acetyltransferase and acetate kinase, from the hyperthermophilic eubacterium Thermotoga maritima, J. Bacteriol., 181, 1861, 1999. 63. Ronimus, R.S. and Morgan, H.W., Cloning and biochemical characterization of a novel mouse ADPdependent glucokinase, Biochem. Biophys. Res. Commun., 315, 652, 2004. 64. Arnfors, L. et al., Structure of Methanocaldococcus jannaschii nucleoside kinase: an archeal member of the ribokinase family, Acta Cryst. D 62, 1185, 2006.

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Industrial Relevance of Thermophiles and Their Enzymes Garabed Antranikian

CONTENTS Introduction ................................................................................................................................ Polymer Degrading Thermoactive Enzymes ............................................................................. Cellulose Hydrolyzing Enzymes .................................................................................... Endoglucanases .................................................................................................... β-Glucosidases ..................................................................................................... Xylanases ........................................................................................................................ Starch Processing Enzymes ............................................................................................ α-Amylases .......................................................................................................... β-Amylases .......................................................................................................... Glucoamylases ..................................................................................................... α-Glucosidases .................................................................................................... Pullulanases ......................................................................................................... CGTases ............................................................................................................... Branching Enzyme .............................................................................................. Amylomaltases ..................................................................................................... Pectin Degrading Enzymes ............................................................................................ Chitinolytic Enzymes ..................................................................................................... Proteases ......................................................................................................................... Biocatalysis with Nonpolymeric Compounds ............................................................................ Lipases and Esterases ..................................................................................................... Alcohol Dehydrogenases ................................................................................................ Glucose and Arabinose Isomerases ................................................................................ C–C Bond Forming Enzymes ........................................................................................ Nitrile-Degrading Enzymes ............................................................................................ DNA Processing Enzymes ......................................................................................................... Polymerase Chain Reaction ............................................................................................ DNA Sequencing ............................................................................................................ Ligase Chain Reaction .................................................................................................... Chemical Products ..................................................................................................................... Compatible Solutes ......................................................................................................... Other Compounds ........................................................................................................... Thermophiles as Cell Factories ................................................................................................. Biomining .......................................................................................................................

114 115 115 115 119 119 120 120 124 124 125 125 126 126 127 127 128 129 132 132 133 134 134 138 138 138 139 142 142 143 143 145 145 113

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Lipids and Peptides ......................................................................................................... Hydrogen Production ...................................................................................................... Outlook ...................................................................................................................................... References ..................................................................................................................................

145 146 146 147

Thermophilic archaea and bacteria that are able to grow at temperatures up to 110°C and at extremes of pH (thermoacidophiles and thermoalkaliphiles) are an interesting source of stable enzymes (extremozymes). These are in general superior to the traditional biocatalysts, because they provide proteins with unique properties and many show reasonable activity even at 120°C, at pH values between 0 and 3, pH 9 to 11 and in the presence of organic solvents (up to 99%). Aiming at the production of high-value products particularly for the chemical, pharmaceutical, cosmetic, food, feed, beverage, paper and textile industries, robust enzymes from thermophiles are gaining significant interest. There is an increasing interest in the utilization of renewable sources to satisfy the exponentially growing energy needs. Therefore, efficient enzyme systems are also needed for the breakdown of plant biomass, which contains complex substrates such a cellulose, hemicellulose, lignin, fats and oils. Microorganisms living in extreme habitats are an ideal source for polymer degraders, which allow to perform biotransformation reactions under nonconventional conditions under which many proteins are completely denatured. In this chapter a new generation of enzymes that are produced by thermophilic, thermoacidophilic and thermoalkaliphilic archaea and bacteria will be presented and their significance for industrial biotechnology will be highlighted.

INTRODUCTION The unique stability of enzymes from thermophiles at elevated temperatures (up to 110°C), extremes of pH and high pressure (up to 1000 bar) makes this group of organisms a valuable resource for industrial enzymes [1]. Of special interest is the thermoactivity and thermostability of these enzymes in the presence of high concentrations of organic solvents, detergents and alcohols. Therefore, they are expected to be a powerful tool in industrial biotransformation processes that run at harsh conditions [2,3]. For the successful exploitation of extremophiles and their enzymes a number of problems, however, have to be resolved. These include the development of efficient cloning and expression systems, especially for hyperthermophilic archaea, and improved cultivation techniques. It is a well known fact that the limitation of the fossil resources in the future will require a new approach to meet the challenges of the next decades. In order to ensure the supply of raw materials for the chemical and pharmaceutical industries and for fuel production a new strategy based on biomass has to be developed. New technologies should allow the efficient conversion of renewable resources which contain polymeric substrates, for example, cellulose, hemicellulose, lignin, and fats and oils to highvalue products. The application of robust enzymes and microorganisms for the sustainable production of chemicals, biopolymers, materials and fuels from renewable resources, also defined as industrial (white) biotechnology, will offer great opportunities for various industries. The utmost aim will be the reduction of waste, energy input and raw material and the development of highly efficient and environmentally friendly processes. Most of the industrial enzymes known to date have been derived from bacteria and fungi [4]. The global annual enzyme market has been estimated to be around 5 billion euros and the market for products derived from enzymes in more than 30-fold. In the case of thermophiles, only few enzymes, however, have found their way to the market. Concerted efforts and interdisciplinary approach from academia and industry are required in order to deliver tailor made industrial enzymes. In order to meet the future challenges, innovative technologies for the production of new generation of enzymes and bioprocesses are needed. In this chapter we will focus on thermophilic archaea and bacteria and their relevance for industrial biotechnology. Thermophilic microorganisms represent thermophiles (growth up to 60°C), extreme thermophiles (65–80°C) and hyperthermophiles (85–110°C).

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POLYMER DEGRADING THERMOACTIVE ENZYMES CELLULOSE HYDROLYZING ENZYMES Cellulose, which is the most abundant organic biopolymer in nature, consists of glucose units linked by β-1,4-glycosidic bonds with a polymerization grade of up to 15,000 glucose units. The molecular weight of cellulose has been estimated to vary from about 50,000 to 2,500,000 in different species. Naturally occurring cellulose is structurally heterogeneous and has both amorphous and highly ordered crystalline regions. The higher crystalline regions are more resistant to enzymatic hydrolysis. Therefore, complex enzyme systems with different specificities are needed to efficiently hydrolyze cellulose. Cellulose can be hydrolyzed into glucose by the synergistic action of different enzymes: endoglucanase (cellulase), exoglucanase (cellobiohydrolase), and β-glucosidase (cellobiase). Endoglucanase (E.C. 3.2.1.4) hydrolyzes cellulose in a random manner as endo-hydrolase producing various oligosaccharides, cellobiose, and glucose. Exoglucanases, (EC 3.2.1.91) hydrolyze β-1,4 d-glycosidic linkages in cellulose and cellotetraose, releasing cellobiose from the nonreducing end of the chain. β-glucosidases (EC 3.2.1.21) catalyze the hydrolysis of terminal, nonreducing β-d-glucose residues releasing β-d-glucose. Endoglucanases Several cellulose degrading enzymes from various thermophilic organisms including archaea and bacteria have been investigated (Table 8.1). Thermostable endoglucanases, which degrade β-1,4 or β-1,3 linkages of β-glucans and cellulose, have been identified in few archaea such as Pyrococcus furiosus, Pyrococcus horikoshii, and Sulfolobus solfataricus [1]. The purified recombinant endoglucanase from the hyperthermophilic archaeon P. furiosus is active at 100°C and hydrolyzes β-1,4 but not β-1,3 glycosidic linkages with the highest specific activity on cellopentaose and cellohexaose [5]. Another thermoactive glucanase (laminarinase) (Topt 100°C) from this strain catalyzes the hydrolysis of mixed-linked oligosaccharides with both β-1,4 and β-1,3 specificities [6]. The E170A mutant of the enzyme is additionally active as a glycosynthase, catalyzing the condensation of α-laminaribiosyl fluoride to different acceptors at pH 6.5 and 50°C [7]. Depending on the acceptor, the synthase generates either β-1,4 or β-1,3 linkage. A recombinant endoglucanase from P. horikoshii was also characterized [8]. This enzyme is active even towards crystalline cellulose. Its activity was recently improved by protein engineering [9]. This enzyme is expected to be useful in biopolishing of cotton products. Very recently, an acid-stable endoglucanase from the thermoacidophilic archaeon S. solfataricus P2 was cloned and expressed in Escherichia coli [10]. The purified recombinant enzyme with optimal activity at 80°C and pH 1.8, hydrolyses carboxymethylcellulose and cello-oligomers, with cellobiose and cellotriose as main products. This extracellular enzyme could be applicable for the large-scale hydrolysis of cellulose under acidic conditions. Bacteria belonging to the genera Thermotoga, Thermobifi da, Rhodothermus, and Clostridium are also good cellulose degraders (Table 8.1). Thermostable endoglucanases from Thermotoga maritima and Thermotoga neapolitana are rather small with a molecular mass of 27 kDa and are optimally active at 95°C to 106°C and between pH 6.0 and 7.0 [11]. Cellulase and hemicellulase genes have been found to be clustered together on the genome of the thermophilic anaerobic bacterium Caldocellum saccharolyticum, which grows on cellulose and hemicellulose as sole carbon sources [12]. The gene for one of the cellulases was isolated and was found to consist of 1751 amino acids. This is the largest cellulase gene sequenced to date. Another large cellulolytic enzyme with the ability to hydrolyze microcrystalline cellulose was isolated from the extremely thermophilic bacterium Anaerocellum thermophilum [13]. This enzyme has an apparent molecular mass of 230 kDa and exhibits significant activity towards Avicel. It is most active towards soluble substrates such as carboxymethylcellulose and β-glucan. Maximal activity is at pH 5 to 6 and 85°C to 100°C. The thermophilic bacterium Rhodothermus marinus produces a thermostable endoglucanase, with a temperature optimum of around 80°C [14]. A 100-kDa protein with endoglucanase

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Acidothermus cellulolyticus Alicyclobacillus acidocaldarius (CelA) Alicyclobacillus acidocaldarius (CelB) Anaerocellum thermophilum Aquifex aeolicus Caldocellum saccharolyticum Clostridium cellulovorans Clostridium thermocellum Pyrococcus furiosus (EglA) Pyrococcus furiosus (LamA) Pyrococcus horikoshii Rhodothermus marinus (struct.) Sulfolobus solfataricus MT4 (struct.) Sulfolobus solfataricus P2 Thermobifida fusca Thermotoga maritima Thermotoga neapolitana

Microbispora bispora Pyrococcus furiosus Pyrococcus horikoshii Sulfolobus acidocaldarius Sulfolobus shibatae Sulfolobus solfataricus (struct.) Thermosphaera aggregans (struct.) Thermotoga maritima Thermotoga neapolitana Thermus caldophilus Thermus nonproteolyticus Thermus sp. Z-1 Thermus thermophilus

β–Glucosidase (EC 3.2.1.21)

Strain

Cellulose-degrading enzymes Endoglucanase (EC 3.2.1.4)

Enzymes

3.6h at 100°C 5–7 6.0 5.6 7.0 7.0 95 80 90 70 60

56

49

49

0.2h at 90°C

48h at 85°C

15h at 85°C >130h at 80°C 6.5 95

240

224

6.2 5.0 6.0 7–8

<48h at 60°C 13h at 110°C 15h at 90°C

8h at 80°C and pH 1.8 >24h at 60°C >6h at 80°C >2h at 106°C

40h at 95°C 19h at 100°C >3h at 97°C

0.5h at 75°C 1h at 80°C 0.8h at 100°C 2h at 100°C

Thermostability (Half-Life)

60 102 >100

52 232

27–29 29–30

5–6 6.0 6.0 6–6.5 5.6 7.0 6.0 1.8 8.2 6–7.5 6–6.6

40–50

79 56 36 31 43–52 30 40 37 100 100 97 85 65 80 77 95 95–106

5.5 4.0 5–6.0 7.0

pH Opt

70 80 95–100 80

Topt °C

58 100 230 39

MW (kDa)

TABLE 8.1 Thermoactive Enzymes Involved in Cellulose, Hemicelullose, Chitin, and Pectin Degradation

Polymer degradation, color brightening, color extraction of juice, saccharification of agricultural and industrial wastes, animal feed, biopolishing of cotton products, bioethanol, synthesis of sugars, optically pure heterosaccharides

Possible Applications

167 168 169 3 3 3 170 171 172 173 174 175 176

160 16 15 13 161 12 162 163 5 7 9 14 164 10 165 11 166

References

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Clostridium stercorarium Clostridium thermocellum Thermomonospora fusca Thermotoga maritima Thermotoga sp. FjSS3-B.1.

Acidobacterium capsulatum Caldicellulosiruptor sp. Clostridium cellulovorans Clostridium thermocellum Dictyoglomus thermophilus Pyrodictium abyssi Rhodothermus marinus Sulfolobus solfataricus Thermoactinomyces sacchari Thermoactinomyces thalophilus Thermoanaerobacter saccharolyticum Thermoanaerobacterium sp. Thermobifida fusca Thermococcus zilligii AN1 Thermotoga maritima (XynA) Thermotoga maritima (XynB) Thermotoga neapolitana Thermus thermophilus

Clostridium cellulovorans Clostridium stercorarium Thermoanaerobacterium sp. JW/SL (A) Thermoanaerobacterium sp. JW/SL (B) Thermomonospora fusca

Thermoanaerobacter ethanolicus Thermotoga maritima

Pyrococcus furiosus Thermobifida fusca

Cellobiohydrolase (EC 3.2.1.91)

Xylan-degrading enzymes Endo-1,4-β-xylanase (EC 3.2.1.8)

Acetyl xylan esterase (EC 3.1.1.6)

1,4-β-xylosidase (EC 3.2.1.37)

β-D-Mannosidase (EC 3.2.1.25)

240 94

165

33 195 106 80

119

130 180 36 95 40,120

48 57

41 36 57 110

36

102 75 60

105 53

82

50 65 80 84

92–105 87 102 100

65 70 60 70 70–85 110 80 100 50 65 70 80 70

75 65 55 95 105

7.4 7.2

5–5.5

6.0 8.0 7.0 7.5 5.7

6.5 5.5 7.5 7.0 8.5 8.5–9 5.5 6.2 7.0 6.0 5–6 6.5 5.5 6.0

5.0 7.0 5.0

5.0 6.6 7–8 6–7.5 7–8

60h at 90°C 30h at 40°C

0.25h at 85°C

0.1h at 75°C 1h at 75°C 1h at 100°C

>3h at 70°C 4h at 95°C 22h at 90°C 8h at 90°C 2h at 100°C

2h at 65°C 1h at 75°C

1.6h at 80°C 0.8h at 90°C

0.6h at 70°C

>12h at 70°C

>16h at 55°C 0.5h at 95°C 1.2h at 108°C

1-2h at 75°C

Paper bleaching, animal feed

197 198

194 195 196

183 192 193 193

181 182 183 184 185 186 3 23 3 187 181 3 188 3 189 190 3 191

177 178 179 24 180

continued

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90 193 160 135

Thermococcus chitonophagus (Chi90) Thermococcus kodakaraensis (GlmA)

Thermococcus kodakaraensis (Tk-Dac)

Clostridium stercorarium Clostridium thermosulfurigenes Thermoanaerobacter italicus (PelA) Thermoanaerobacter italicus (PelB) Thermomonospora fusca Thermotoga maritima (PelA) Thermotoga maritima (PelB) Thermotoga maritima (exo-PG)

Chitobiase (EC 3.2.1.30)

Diacetylchitobiose Deacetylase

Pectin-degrading enzymes (EC 3.1.1.11)

51

135 251 56 151

35 55 42 50

Microbispora sp. V2 Pyrococcus furiosus (ChiB) Rhodothermus marinus Thermococcus chitonophagus (Chi50)

Exochitinase (EC 3.2.1.52)

53 40

350 332

65 75 80 80 60 90 80 95

75

80

60 90–95 70 80

60 90–95 80 70 85

85 70 90

6.0

7.0 5.5 9.0 9.0 10.45 9.0

8.5

6.0

3.0 6.0 4.5–5 6.0

4.5–6.5 6.0 9.0 7.0 5.0

5.5–7 6.0 7.0

5.8

65–75 80

120 38

70 134

Rhodothermus marinus Thermomicrobia sp. Thermotoga maritima

α-LArabinofuranosidase (EC 3.2.1.55) Chitin-degrading enzymes Endochitinase (EC 3.2.1.14)

5.0 5.4

pH Opt

80 85

Topt °C

40 113

MW (kDa)

Clostridium thermocellum Pyrococcus furiosus (ChiA) Streptomyces thermoviolaceus Thermococcus chitonophagus (Chi70) Thermococcus kodakaraensis (ChiA)

Caldocellulosyruptor sp. Rt8B.4. Dictyoglomus thermophilum Rhodothermus marinus Thermoanaerobacterium polysaccharolyticum Thermomonospora fusca

Strain

β-Mannanase (EC 3.2.1.78)

Enzymes

TABLE 8.1 (continued)

>5h at 90°C

2h at 95°C

0.2h at 70°C 0.5h at 70°C >1h at 80°C >1h at 80°C

3h at 90°C

24h at 50°C

10h at 60°C 1h at 120°C

8.3h at 85°C >1h at 70°C 2.7h at 100°C

>16h at 80°C >1h at 90°C

Thermostability (Half-Life)

Utilization of biomass of marine environment

Possible Applications

213 24 60 60 214 61 62 215

65

63, 64 212

210 67 211 63, 64

207, 208 67 209 63, 64 66

204 205 206

203

199 200 201 202

References

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activity was purified from Triton X-100 extract of cells of the thermoacidophilic Gram-positive bacterium Alicyclobacillus acidocaldarius [15]. The enzyme exhibits activity towards carboxymethylcellulose and oat spelt xylan with pH and temperature optima of pH 4 and 80°C, respectively. Remarkable stability was observed at pH values between 2 and 6 and 60% of activity was retained after incubation at 80°C for 1 h. Another glucanase purified from the same microorganism is less acidstable, having maximum activity at pH 5.5 [16]. β-Glucosidases Unlike endoglucanases, several β-glucosidases have been characterized from archaea. These enzymes have been detected in strains of the genera Sulfolobus, Pyrococcus, and Thermosphaera. The β-glucosidase from P. furiosus is very stable with optimal activity at 103°C and it also exhibits a β-mannosidase activity [17]. The β-glucosidase from S. solfataricus MT4 is very resistant to various denaturants with activity up to 85°C. The gene for this β-glucosidase has been cloned and expressed in E. coli and Saccharomyces cerevisiae [18]. Using a mixture of both β-glucosidases from P. furiosus and S. solfataricus an ultra high-temperature process for the enzymatic production of novel oligosaccharides from lactose was developed [19]. For the production of glucose from cellobiose a bioreactor system with immobilized recombinant β-glucosidase from S. solfataricus was developed [19]. The system runs at a high flow rate and has a high degree of conversion, productivity, and operational stability. The thermoactive β-glucosidase from P. horikoshii is active in organic solvents and it synthesizes a heterosaccharide with high optical purity [1]. There is a great demand for robust cellulolytic enzymes especially for the efficient bioconversion of plant material to utilizable monomeric and oligomeric sugar molecules. The bottleneck for the successful application of the enzymes is making the complex substrate available to the enzymes. This limiting step can be overcome by the application of chemical, physical and enzymatic methods including high pressure, temperature, ionic solvents, extraction with supercritical fluids and enzymes. The combination of these parameters will make the substrate accessible to enzymes and allow the conversion of renewable biomass (lignocelluloses) to high value products for various applications such as improvement of juice yield, effective color extraction of juices and fuel (ethanol) production. To date more than 40 million tons of ethanol is produced by fermentation with an estimated growth rate of 10% to 20% per year. Other suitable applications of cellulases include the pretreatment of cellulose biomass and forage crops to improve nutritional quality and digestibility. Due to the limitation of fossil resources it is expected that cellulases will be useful tools for the saccharification of agricultural and industrial wastes and production of fine chemicals. It has been estimated that around 40% of the bulk chemical produced to day can be derived from plant waste material.

XYLANASES There are a variety of thermophilic bacteria and archaea that are able to utilize xylan as carbon and energy source. Xylan is a heterogeneous molecule that constitutes the main polymeric compound of hemicellulose, a fraction of the plant cell wall, which is a major reservoir of fixed carbon in nature. The main chain of the heteropolymer is composed of xylose residues linked by β-1,4-glycosidic bonds. Approximately half of the xylose residues have substitution at O-2 or O-3 positions with acetyl-, arabinosyl-, and glucuronosyl-groups. The complete degradation of xylan requires the action of several enzymes. The endo-β-1,4-xylanase (EC 3.2.1.8) hydrolyzes β-1,4-xylosydic linkages in xylans, while β-1,4-xylosidase (EC 3.2.1.37) hydrolyzes β-1,4-xylans and xylobiose by removing the successive xylose residues from the nonreducing termini. To date only few hyperthermophilic archaea that are able to grow on xylan and secrete thermoactive xylanolytic enzymes (Table 8.1). Among the thermophilic archaea, a xylanase from Pyrodictium abyssi has been characterized with an optimum temperature of 110°C—one of the highest values reported for a xylanase [20]. The crenarchaeon Thermosphaera aggregans was

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shown to grow on heat-treated, but not native, xylan [21]. The xylanase from Thermococcus zilligii AN1 is active up to 100°C, can attack different xylans and is not active towards cellulose [22]. Recently, a thermoactive endoxylanase from S. solfataricus was purified and characterized [23]. The products of xylan hydrolysis are xylooligosaccharides and xylobiose. Thermophilic anaerobic bacteria such as Clostridium, Dictyoglomus, Rhodothermus, Thermotoga, and Thermus are also able to secrete heat stable xylanases (Table 8.1). These enzymes are active between 80°C and 105°C and are mainly cell associated and most probably localized within the toga, which covers the cells. Several genes encoding thermostable xylanases, for example, T. maritima, have been already cloned and expressed in E. coli. Other hemicellulases (glucoronidase, β-mannanase, β-mannosidase, galactosidase, acetyl xylan esterase, ferruloyl esterase, and α-arabinofuranosidase), isolated from extremophiles, are efficient enzymes for the complete saccharification of plant cell wall (Table 8.1). Robust xylanases are attractive candidates for various biotechnological applications and can be used also in combination with other depolymerases. Enzymes from bacteria and fungi are already produced on industrial scale and are used as food additives in poultry, for increasing feed efficiency diets and in wheat flour for improving dough handling and the quality of baked products. In the last decade, the major interest in thermostable xylanases was in enzyme-aided bleaching of paper. The chlorinated lignin derivatives generated by this process constitute a major environmental problem. Recent investigations have demonstrated the feasibility of enzymatic treatments as alternatives to chlorine bleaching for the removal of residual lignin from pulp. A treatment of craft pulp with cellulase-free thermostable xylanases leads to a release of xylan and residual lignin without undo loss of other pulp components. Xylanase treatment at elevated temperatures opens up the cell wall structure, thereby facilitating lignin removal in subsequent bleaching stages and thus enhance the development of environmentally friendly processes [24].

STARCH PROCESSING ENZYMES The starch processing industry, which converts starch into more valuable products such as dextrins, glucose, fructose, maltose, and trehalose, utilize microbial thermostable enzymes. In all starch converting processes, high temperatures are required to liquefy starch and make it accessible to enzymatic hydrolysis. The synergetic action of thermostable amylolytic enzymes brings an advantage to those processes, lowering the cost of sugar syrup production. The use of thermostable enzymes can lead to other valuable products, which include innovative starch-based materials with gelatin-like characteristics and defined linear dextrins that can be used as fat substitutes, texturizers, aroma stabilizers, and prebiotics [25]. Table 8.2 gives an overview on starch modifying enzymes from thermophilic archaea and bacteria. α-Amylases α-amylase (α-1,4-glucan-4-glucanohydrolase; EC 3.2.1.1), hydrolyzes linkages in the starch polymer, which leads to the formation of linear and branched oligosaccharides. The sugar reducing groups are liberated in the α-anomeric configuration. Most of starch hydrolyzing enzymes belongs to the α-amylase family that contains a characteristic catalytic (β/α)8-barrel domain. Throughout the α-amylase family, only eight amino acid residues are invariant, seven at the active site and a glycine in a short turn. It seems that the ability of hyperthermophilic archaea to utilize starch is more frequent than cellulose (Tables 8.1 and 8.2). Extremely thermostable α-amylases have been characterized from a variety of hyperthermophilic archaea belonging to the genera Methanocaldococcus, Pyrococcus, and Thermococcus [1]. The thermostability of the enzymes is often enhancing in the presence of divalent metal ions. The optimal temperatures for the activity of these enzymes range between 80°C and 100°C. The high thermostability of the pyrococcal extracellular α-amylase (thermal activity even at 130°C and after autoclaving for 4 h at 120°C) and α-amylase from Methanocaldococcus jannashii (temperature optimum 120°C, half-life of 50 h at

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Glucoamylase (EC 3.2.1.3)

β-Amylase (EC 3.2.1.2)

α-Amylase (EC 3.2.1.1)

Enzymes

Bacillus sp. Clostridium thermohydrosulfuricum Clostridium thermosaccharolyticum Methanocaldococcus jannaschii

Thermoanaerobacter thermosulfurigenes Thermotoga maritima

136 58 108

Alicyclobacillus acidocaldarius Anaerobranca gottschalkii (AmyA) Anaerobranca gottschalkii (AmyB) Desulfurococcus mucosus Dictyoglomus thermopilum Methanocaldococcus jannaschii Pyrococcus furiosus (intracellular) Pyrococcus furiosus (extracellular) Pyrococcus woesei (struct.) Pyrodictium abyssi Rhodothermus marinus Staphylothermus marinus Sulfolobus solfataricus Thermococcus aggregans Thermococcus celer Thermococcus hydrothermalis Thermococcus profundus (amyS) Thermococcus kodakaraensis Thermotoga maritima (AmyA) Thermototoga maritima (AmyC) (struct.) Thermus filimormis

75

180

60

49 43 49.5 61

240

76 100 68

75

MW (kDa)

Strain

70 75 70 80

5.0 4–6 5.0 6.5

6.0 4–5.5

6.0

95 70 95

6.5 5.5 5–5.5 5–6 6.5 7.0 8.5

3.0 8.0 6–6.5 5.5 5.5 5–8 7.0 5.5–6 5.5 5.0 6.5 5.0

pH Opt

95 90 75–85 80 90 85 90

75 70 55 100 90 120 92 100 100 100 85 100

Topt °C

TABLE 8.2 Enzymes from Thermophilic Microorganisms for Starch-Processing

3.8h at 70°C 1h at 85°C >6h at 70°C

2h at 80°C 0.5h at 90°C

8h at 65°C

4h at 90°C 3h at 80°C 24h at 70°C 4h at 80°C

13h at 98°C 11h at 90°C

50h at 100°C

48h at 70°C 0.2h at 70°C

Thermostability (Half-Life) Bread and baking industry, starch liquefaction and saccharification, production of glucose, textile desizing, paper industry, synthesis of oligosaccharides, detergent application, gelling, thickening in food industry

Possible Applications

222 223 31 30

24 24

221

28 216 216 3 3 217 3 3 3 3 218 3 3 3 3 3 3 3 219 220

References

continued

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Anaerobranca gottschalkii Caldicellulosiruptor saccharolyticus Clostridium thermohydrosulfuricum Desulfurococcus mucosus Fervidobacterium pennivorans Pyrococcus furiosus (pullulanase) Pyrococcus furiosus (pullulan-hydrolase) Pyrococcus woesei (struct.) Pyrodictium abyssi

Pullulanase (EC 3.2.1.41)

5.5 7.0 7.0 7.5

120 120 75 90

57 63

90

100 100

6.0 9.0

3 4

223 3 228 3 229 1h at 90°C 0.8h at 85°C 2h at 80°C 5.6–6 5.0 7.0 6.0 5.0 85 85 80 105 90 74 190 90 77

2h at 95°C

41 227 22h at 70°C 8.0 6.0

3, 226 21 225 36

70 85

48h at 50°C

6h at 80°C 39h at 85°C

37 21 3 225 33 34 32

98 96

110

60

2.4–3.5 5.5 5–5.5 5.5 4.5 5–5.5 5.5

55-60 115 110 85 105 75 65

5.5–6

90

250 57 125 90 313 80

Antranikian, unpublished

40h at 60°C

5.0

75

140

Ferroplasma acidiphilum Pyrococcus furiosus Pyrococcus woesei Sulfolobus shibatae Sulfolobus solfataricus Thermoanaerobacter ethanolicus Thermoanaerobacter thermosaccharolyticum Thermococcus hydrothermalis Thermococcus sp. AN1 Thermococcus zilligii Thermotoga maritima

29

24h at 90°C

2.0

90

141

26

29 29 Antranikian, unpublished 224 32

20h at 90°C 24h at 90°C 4h at 55°C 0.5h at 60°C 8h at 65°C

References

2.0 2.0 5.0 6.8 4–5.5

Possible Applications

90 90 50 60 65

Thermostability (Half-Life)

140 133 312 75 75

pH Opt

Picrophilus oshimae (extracellular) Picrophilus torridus (extracellular) Picrophilus torridus (intracellular) Thermoactinomyces vulgaris Thermoanaerobacter thermosaccharolyticum Thermoplasma acidophilum (extracellular) Thermoplasma acidophilum (intracellular) Sulfolobus solfataricus

Topt °C

MW (kDa)

Strain

α-Glucosidase (EC 3.2.1.20)

Enzymes

TABLE 8.2 (continued) 122 Thermophiles: Biology and Technology at High Temperatures

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80 87 53 57 66 66 72 74 78 72

Dictyoglomus thermophilus Pyrobaculum aerophilum Thermococcus aggregans Thermococcus litoralis (struct.) Thermotoga maritima Thermus aquaticus

Alicyclobacillus acidocaldarius Clostridium thermohydrosulfuricum

Anaerobranca gottschalkii Aquifex aeolicus Geobacillus stearothermophilus Rhodothermus obamensis Thermococcus kodakaraensis

Amylomaltase (EC 2.4.1.25)

Cyclomaltodextrinase (EC 3.2.1.54)

Branching enzyme (EC 2.4.1.18)

50 75 50 65 70

65

(struct.) – the protein has been crystallized and the three-dimensional structure is determined.

75

80 95 100 90 55–80 75

80 90–110

77 83

68

65–70 90–95 80–85

7.0 7.5–8 7.5 6–6.5 7.0

5.5 6.0

16h at 80°C up to 90°C

>0.5h at 70°C

2.5h at 90°C 24h at 80°C

20h at 90°C

0.33h at 100°C 0.66h at 110°C

5.5–6 5–5.5

46 46 48 49 47

236 237

235 27 Antranikian, unpublished 51 219 54

43 42

0.5h at 100°C

6.0 6.7 6.8 6.0 6.0 5.5–6

45 234 45

6–9 6.0 4.5–7

3.5h at 90°C 4.5h at 95°C 72h at 78°C

6.7h at 90°C

2.5h at 100°C

6.5 5.5 5.5 5.5 5.5 8.0 6.4 5.5

95 90 95 98 90 80–95 90 80–95 75 70

78

Anaerobranca gottschalkii Thermoanaerobacter sp. Thermoanaerobacterium thermosulfurigenes Thermococcus kodakaraensis Thermococcus sp.

93 83 65 80

128 119 43

232 4 3 3 3 225 34 34 34 233

2h at 80°C

6.0

75

180

231

0.5h at 65°C

100

218 230 32

5–5.5 6.0

80 90 65

109 150

Cyclodextrin glucosyltransferase (EC 2.4.1.19)

Rhodothermus marinus Thermoanaerobacter ethanolicus Thermoanaerobacter thermosaccharolyticum Thermoanaerobacter thermosulfurigenes Thermococcus aggregans Thermococcus celer Thermococcus hydrothermalis Thermococcus litoralis Thermococcus profundus (amyL) Thermococcus sp. ST489 Thermotoga maritima Thermus aquaticus Thermus caldophilus Thermus thermophilus

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100°C) makes these enzymes interesting candidates for industrial application [217]. The extreme marine hyperthermophilic archaeon P. abyssi can grow on various polysaccharides and also secretes a heat stable α-amylase [20]. α-amylases with lower thermostability have been isolated from the archaea Thermococcus profundus, Thermococcus kodakaraensis and the extreme thermophilic bacteria Dictyoglomus thermophilum, T. maritima, Thermus filiformis, and R. marinus (Table 8.2). In general the bacterial enzymes are less thermoactive than the archaeal enzymes. Similar to the amylase from Bacillus licheniformis, which is commonly used in liquefaction of starch in the industry, most of the enzymes from extreme thermophilic bacteria require calcium for activity. The use of α-amylases in detergents for medium-temperature laundering demands enzymes with high stability and activity at alkaline conditions. Therefore, extracellular enzymes from thermoalkaliphiles are good candidates for application in laundry and dish washing. The enzyme from the thermoalkaliphilic bacterium Anaerobranca gottschalkii is optimally active at pH 8.0 and has high transglycosylation activity on maltooligosaccharides. Interestingly, the enzyme exhibits also significant β-cyclodextrin glycosyltransferase (CGTase, EC 2.4.1.19) activity [216]. On the other hand, enzymes that are active at high temperatures but low pH are also of interest for application, for example, textile and beverage industries. An acid-stable amylase was purified and characterized from the thermoacidophilic bacterium Alyciclobacillus acidocaldarius [28]. This enzyme with a molecular mass of 160 kDa exhibits highest activity at pH 3.0 and 75°C. β-Amylases β-amylase (1,4-alpha-d-glucan maltohydrolase; EC 3.2.1.2), hydrolyzes 1,4-alpha-d-glucosidic linkages in polysaccharides removing successive maltose units from the nonreducing ends of the chains. For the efficient production of maltose syrups an additional debranching enzyme is needed. To date only few thermoactive bacterial β-amylases are known (Table 8.2). The enzyme from T. maritima is active in the absence of calcium at low pH (pH 4.3–5.5) and high temperature (95°C) [24]. The enzyme from Thermoanaerobacter thermosulfurigenes retains 70% of its activity at pH 4.0 and 70°C [24]. Glucoamylases Unlike α-amylase, the production of glucoamylase seems to be rare in archaea and bacteria (Table 8.2). Glucoamylases (EC 3.2.1.3) hydrolyze terminal α-1,4-linked-d-glucose residues successively from nonreducing ends of the chains, releasing β-d-glucose. An ideal catalyst for starch liquefaction should be optimally active at 100°C and pH 4.0 to 5.0 without requirement of calcium ions for the stabilization of the enzyme. Recently, it has been shown that also the thermoacidophilic archaea Thermoplasma acidophilum, Picrophilus torridus, and Picrophilus oshimae produce heat and acid stable extracellular glucoamylases. The purified archaeal glucoamylases are optimally active at pH 2 and 90°C. Catalytic activity is still detectable at pH 0.5 and 100°C [29]. These enzymes are more thermostable than already described glucoamylases from bacteria, yeasts, and fungi. They are of interest for application in the beverage industry. However, the lack of suitable genetic methods for thermoacidophiles have precluded structural studies aimed to discover their adaptation at very low pH. Recently, the gene encoding a putative glucoamylase from S. solfataricus, was cloned and expressed in E. coli, and the properties of the recombinant protein were examined in relation to the glucose production process [26]. This recombinant enzyme is extremely thermostable, with an optimal temperature at 90°C; however, it is most active in a slight acidic pH range from 5.5 to 6.0. The tetrameric enzyme liberates β-d-glucose from maltotriose, and the substrate preference for maltotriose distinguishes this enzyme from fungal glucoamylases. Genome analysis of other thermocidophiles revealed further putative glucoamylases, which were cloned and expressed in E. coli. Thus, the recombinant intracellular glucoamylase from M. jannashii is active at pH 6.5 and 80°C [30]. In our laboratory, the intracellular glucoamylases from the extreme thermoacidophiles P. torridus and T. acidophilum have been recently cloned and expressed in E. coli; the

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recombinant enzymes are optimally active at 50°C to 75°C and pH 5. Thermophilic anaerobic bacteria such as Clostridium thermohydrosulfuricum, Clostridium thermosaccharolyticum, and Thermoanaerobacterium thermosaccharolyticum produce glucoamylases, which have been purified and characterized [31,32]. α-Glucosidases α-glucosidases (EC 3.2.1.20) attack the α-1,4 linkages of oligosaccharides and unlike glucoamylases, α-glucosidase prefers smaller oligosaccharides, for example, maltose, maltotriose and liberates glucose with an α-anomeric configuration. An intracellular and an extracellular α-glucosidases have been purified and characterized from archaea, belonging to the genera Pyrococcus, Sulfolobus, and Thermococcus (Table 8.2) [3]. The enzymes exhibit optimal activity at pH 4.5 to 7.0 over a temperature range from 105°C to 120°C. An α-glucosidase gene and flanking sequences from S. solfataricus were cloned in E. coli and the product was characterized [33]. The purified recombinant enzyme with a calculated size of 80.5 kDa hydrolyzes p-nitrophenyl-d-glucopyranoside. At pH 4.5 it exhibits a pH optimum for maltose hydrolysis. Unlike maltose hydrolysis, glycogen was hydrolyzed efficiently at the intracellular pH of the organism (pH 5.5). The recombinant α-glucosidase exhibits greater thermostability than the native enzyme, with a half-life of 39 h at 85°C at a pH of 6.0. Less thermostable α-glucosidases were detected in the bacteria Thermoanaerobacter ethanolicus [34,35], T. thermosaccharolyticum [32], T. maritima [36], and archaea Ferroplasma acidiphilum [37]. The α-glucosidase from the extreme acidophilic archaeon has maximal activity at pH 2.4 to 3.5 (>70% activity at pH 1.5). Iron was found to be essential for enzymatic activity and His30 was shown to be responsible for iron binding. Pullulanases Enzymes capable of hydrolyzing α-1,6 glucosidic bonds in pullulan and branched oligosaccharides are defined as pullulanases. On the basis of substrate specificity and product formation, pullulanases have been classified into three groups: pullulanase type I, pullulanase type II and pullulan hydrolases (type I, II, and III). Pullulanase type I (EC 3.2.1.41) specifically hydrolyzes the α-1, 6-linkages in pullulan as well as in branched oligosaccharides (debranching enzyme), and its degradation products are maltotriose and linear oligosaccharides, respectively. Pullulanase type I is unable to attack α-1,4-linkages in α-glucans. Pullulanase type II (amylopullulanase) attacks α-1,6glycosidic linkages in pullulan and branched polysaccharides. Unlike pullulanase type I, this enzyme also attacks α-1,4-linkages in branched and linear oligosaccharides and is able to fully convert polysaccharides (e.g., amylopectin) to small sugars (e.g., glucose, maltose, and maltotriose) in the absence of amylases. Thermoactive pullulanase type II from the archaea Desulfurococcus mucosus, P. furiosus, Pyrococcus woesei, and Thermococcus hydrothermalis, have been reported to have temperature optima between 85°C and 105°C (Table 8.2), as well as remarkable thermostability even in the absence of substrate and calcium ions. In the presence of calcium ions pullulanase activity was also detected at 130°C to 140°C [38]. Site-directed mutagenesis performed on pullulanase from T. hydrothermalis reveals that the residues E291 and D394 are critical for the pullulanolytic and amylolytic activities of the pullulanase [39]. The crucial role of E291 as the catalytic nucleophile has been also confirmed for the pullulanase from P. furiosus [40]. The apparent catalytic efficiencies (Kcat/K m) of mutants E291Q and D394N on pullulan were 123 and 24 times lower than that of the native enzyme. The hydrolytic patterns for pullulan and starch were the same, while the hydrolysis rates differed as reported. Therefore, these data strongly suggest that the bifunctionality of the pullulanase type II is determined by a single catalytic center. Due to the dual specificity of pullulanases type II to degrade both α-1,4- and α-1,6-glucosidic linkages they cannot be used as debranching enzymes in maltose and glucose syrup production. The archaeal enzymes are promising candidates to optimize starch liquefaction for the production of maltose, maltotriose, and maltotetraose syrups.

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Interestingly, pullulanase type I has not been identified in archaea so far, whereas the enzyme has been characterized in several thermophilic bacteria belonging to the genera Fervidobacterium, Thermoanaerobacter, Thermotoga, and Thermus (Table 8.2). The aerobic thermophilic bacterium Thermus caldophilus GK-24 produces a thermostable pullulanase of type I when grown on starch [34]. The pullulanase is optimally active at 75°C and pH 5.5, is thermostable up to 90°C, and does not require calcium ions for either activity or stability. The first debranching enzyme (pullulanase type I) from anaerobic thermophilic bacteria was found in Fervidobacterium pennivorans. The recombinant enzyme forms long chain linear polysaccharides from amylopectin. A similar enzyme was also characterized from T. maritima. All known bacterial debranching enzymes are active in the acidic or neutral pH range. Until very recently, no reports have been presented on the ability of thermoalkaliphiles to produce heat and alkaline stable pullulanase type I. After sequencing the whole genome of the thermoalkaliphile A. gottschalkii, a pullulanase encoding gene was cloned and expressed in E. coli, this enzyme is optimally active at pH 8.0 and 70°C [41]. The third class of pullulan-hydrolyzing enzymes includes pullulan hydrolases type I, II, and III. Pullulan hydrolases type I and II are active towards α-1,4 linkages of amylose, starch, pullulan, but are unable to hydrolyze α-1,6 linkages. An exception is pullulan hydrolase type III. This enzyme attacks α-1,4 as well as α-1,6 linkages of pullulan. The enzymes from P. furiosus (AmyL), Thermococcus aggregans and T. profundus (Table 8.2) exhibit maximal activity at 90°C and pH 5.5 to 6.5 and are stable for several hours at 95°C to 100°C. In addition, the pullulan hydrolase from P. furiosus degrades β-cyclodextrin. CGTases Cyclodextrin glucosyltransferase (CGTase; EC 2.4.1.19) converts α-glucans into cyclodextrins, which are composed of 6 (α), 7 (β), or 8 (γ) α-1,4 linked glucose molecules. The internal cavities of cyclodextrins are hydrophobic and they can encapsulate hydrophobic molecules. Thermostable CGTases are generally found in bacteria and was recently discovered in archaea. The archaeal enzyme found in Thermococcus sp. is optimally active at 90°C to 110°C (Table 8.2). Incubation of the enzyme with 30% corn-starch (wt/vol) for 24 h at 96°C and pH 4.5 resulted in the production of α-cyclodextrin (69%), β-cyclodextrin (20%), and γ-cyclodextrin (11%) [42]. The major cyclodextrin formed by the action of the CGTase from T. kodakaraensis is β-cyclodextrin [43]. Bacterial CGTases, isolated mostly from the species of genus Bacillus, are already used in industry for the production of cyclodextrins [44]. The use of more thermoactive CGTases will allow to develop a single step process at temperatures above 90°C [19]. The CGTases active at the temperatures of 80°C to 90°C are produced by some anaerobic thermophilic bacteria (Table 8.2). After sequencing of the genome of the anaerobic bacterium A. gottschalkii the CGTase gene was cloned and expressed [45]. Branching Enzyme Branching enzyme (EC 2.4.1.18) catalyzes the formation of α-1,6 branching points from linear oligo- and polysaccharides, determining the final structures and properties of amylopectin and glycogen. Branching enzymes increase the solubility and stability of starch solutions and shelf life and loaf volume of baked goods [46]. The most thermoactive branching enzyme was isolated from the thermophilic bacterium Aquifex aeolicus (>90% activity after 10 min treatment at 90°C). The enzyme is also able to produce large cyclic glucans using amylopectin as substrate [48]. The branching enzyme of T. kodakaraensis KOD1 produces branched glucans that are 100 times larger than the substrate [47]. The branching activity of the enzyme from Rhodothermus obamensis is higher towards amylose than amylopectin [49]. The enzyme from the thermoalkaliphilic bacterium A. gottschalkii displays at 50°C high transglycosylation activity with extremely low hydrolytic activity [46].

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Amylomaltases Amylomaltases (EC 2.4.1.25, 4-α-glucanotransferase) catalyze the transfer of a segment of a α-1,4d-glucan to a new 4-position of an acceptor, which may be glucose or another α-1,4-d-glucan. Acting upon starch, amylomaltases can produce products of commercial interest, such as cycloamylose, a thermoreversible starch gel, which can be used as a substitute for gelatin. In combination with α-amylase the amylomaltase produces syrups of isomalto-oligosaccharides with reduced sweetness and low viscosity. The thermostable amylomaltase from the archaeon Pyrobaculum aerophilum produces a thermoreversible starch product with gelatin-like properties [27]. The enzyme from Thermococcus litoralis produces linear α-1,4-glucans and a cycloamylose (cyclic α-1,4-glucan) with a high degree (up to hundreds) of polymerization [50–52]. Recently, a heat-stable amylomaltase was characterized from the archaeon T. aggregans. The recombinant enzyme is stable at 90°C for more than 22 h (Antranikian, unpublished results). The combined use of the amylomaltase from the bacterium T. maritima with a maltogenic amylase results in the production of isomalto-oligosaccharides from starch [53]. Cyclic glucans can be produced using the thermostable bacterial amylomaltase from Thermus aquaticus [54,55]. The finding of novel thermostable starch-modifying enzymes will be a valuable contribution to the starch-processing industry. At elevated temperatures starch is more soluble (30 to 35% w/v) and the risk of contamination is reduced. This is of advantage when starch will be converted to high glucose and fructose syrup. The application of thermostable enzymes that are active and stable above 100°C and at acidic pH values can simplify the complicated multistage starch conversion process. The use of the extremely thermostable amylolytic enzymes can lead to other valuable products, which include innovative starch-based materials with gelatin-like characteristics and defined linear dextrins that can be used as fat substitutes, texturizers, aroma stabilizers, and prebiotics. CGTases are used for the production of cyclodextrins that can be used as a gelling, thickening, or stabilizing agent in jelly desserts, dressing, confectionery, and dairy and meat products. Due to the ability of cyclodextrins to form inclusion complexes with a variety of organic molecules, cyclodextrins improve the solubility of hydrophobic compounds in aqueous solution. This is of interest for the pharmaceutical and cosmetic industries. Cyclodextrin production is a multistage process in which starch is first liquefied by a heat-stable amylase and in the second step a less-thermostable CGTase from Bacillus sp. is used. The application of heat-stable CGTase from the Thermococcus species in jet cooking, where temperatures up to 105°C could be achieved, will allow liquefaction and cyclization to take place in one step [56]. Another promising application of an archaeal enzyme is the production of a disaccharide trehalose, a stabilizer of enzymes, antibodies, vaccines and hormones. The use of thermoactive enzymes in the process would eliminate problems associated with viscosity and sterility. The process was developed to produce trehalose from dextrins using Sulfolobus enzymes at 75°C in a continuous bioreactor, with a final conversion of 90% [19]. Recently, the trehalose biosynthetic pathway was identified in Sulfolobales and the responsible enzymes were cloned and expressed in E. coli [57–59].

PECTIN DEGRADING ENZYMES Pectin is an important plant material that is present in the middle lamellae as well as in the primary cell walls. This biopolymer is a branched heteropolysaccharide consisting of a main chain of α-1,4d-polygalacturonate, which is partially methyl esterified. Along the chain, l-rhamnopyranose residues are present that are the binding sites for side chains composed of neutral sugars. Pectin is degraded by pectinolytic enzymes, which can be classified into two major groups. The first group comprises methylesterases, which function is to remove the methoxy groups from pectin. The second group comprises the depolymerases (hydrolases and lyases), which attack both pectin and pectate (polygalacturonic acid). Interestingly, no reports are available on the production of pectinolytic enzymes by thermophilic archaea. Unlike archaea, few pectinolytic enzymes from thermophilic

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anaerobic bacteria have been reported (Table 8.1). The enzymes usually act at alkaline pH and are calcium dependent. Thus, a spore forming anaerobic microorganism Thermoanaerobacter italicus is able to grow at 70°C on citrus pectin and pectate. After growth on citrus pectin, two pectate lyases were induced, purified and biochemically characterized [60]. Both enzymes display similar catalytic properties and can function at temperatures up to 80°C. An increase in the enzymatic activity of both pectate lyases was observed after the addition of calcium ions. The ability of the hyperthermophilic bacterium T. maritima to grow on pectin as a sole carbon source coincides with the secretion of an extracellular pectate lyase. The corresponding gene was functionally expressed in E. coli as the first heterologously produced thermophilic pectinase [61]. Highest activity was demonstrated on polygalacturonic acid, whereas pectins with an increasing degree of methylation were degraded at a decreasing rate. The tetrameric enzyme requires calcium ions for stability and activity. The enzyme is highly thermoactive and thermostable, operating optimally at 90°C and pH 9.0, with a half-life for thermal inactivation of almost 2 h at 95°C, and an apparent melting temperature of 102.5°C. With polygalacturonic acid PelA has a unique eliminative exo-cleavage pattern liberating unsaturated trigalacturonate as the major product. T. maritima also produces exopolygalacturonase, which has been rarely described in bacteria [62]. Pectin degrading enzymes from the bacteria of the genus Clostridium are not very thermostable [24]. Enzymatic pectin degradation is widely applied in food technology processes, as in fruit juice extraction and wine making, in order to increase the juice yield, to reduce its viscosity, improve colour extraction from the fruit skin and to macerate fruit and vegetable tissues.

CHITINOLYTIC ENZYMES Chitin is the major structural component of most fungi and some invertebrates (crustacia und insects) and it is one of the most abundant natural biopolymer on earth. It has been estimated that the annual worldwide formation rate and steady state amount of chitin is in the order of 1010 to 1011 tons per [3]. Chitin is a linear β-1,4 homopolymer of N-acetyl-glucosamine residues. Particularly in the marine environment, chitin is produced in enormous amounts and its turnover is due to the action of chitinolytic enzymes. Chitin degradation is known to proceed with the endoacting chitin hydrolase (EC 3.2.1.14), the chitin oligomer degrading exoacting hydrolases (EC 3.2.1.52) and the N-acetyl-d-glycosaminidase (chitobiase; EC 3.2.1.30). Hyperthermophilic archaea, Thermococcus chitonophagus [63,64], T. kodakaraensis [65,66], and P. furiosus [67] have been shown to utilize chitin and produce chitinolytic enzymes (Table 8.1). The extreme thermophilic anaerobic archaeon T. chitonophagus possesses a multicomponent enzymatic system, consisting of an extracellular exochitinase (Chi50), a periplasmic chitobiase (Chi90) and a cell-membrane-anchored endochitinase (Chi70) [63]. The chitinolytic system is strongly induced by chitin, although a low-level constitutive production of the enzymes in the absence of any chitinous substrates was detected. The archaeal chitinase (Chi70) is a monomeric enzyme with an apparent molecular weight of 70 kDa and appears to be associated with the outer surface of the cell membrane. The enzyme is optimally active at 70°C and pH 7.0 and is thermostable, maintaining 50% activity at 120°C even after 1 h. The enzyme is not inhibited by allosamidin, the natural inhibitor of chitinolytic activity, and is also resistant to denaturation by urea and sodium dodecyl sulfate (SDS). The chitinase has a broad substrate specificity for several chitinous substrates and derivatives and has been classified as an endochitinase due to its ability to release chitobiose from colloidal chitin [68]. The purified recombinant chitinase from the hyperthermophile T. kodakaraensis is optimally active at 85°C and pH 5.0 and produces chitobiose as the major end product [66]. The thermostable chitinase from T. kodakaraensis is active in the presence of detergents and organic solvents and can be applied, for example, for the production of N-acetyl-chitooligosaccharides with biological activity [66]. This unique multidomain protein consists of two active sites with different cleavage specificities and three substrate-binding domains, which are related to two families of cellulose-binding domains [69]. A chitin-degrading pathway involves unique enzymes

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diacetylchitobiose deacetylase and exo-β-d-glucosaminidase. After the hydrolysis of chitin by chitinase diacetylchitobiose will be deacetylated and then successively hydrolyzed to glucosoamine [65]. P. furiosus was also found to grow on chitin, adding this polysaccharide to the inventory of carbohydrates utilized by this hyperthermophilic archaeon. Two open reading frames (ChiA and ChiB) were identified in the genome of P. furiosus, which encode chitinases with sequence similarity to proteins from the glycosyl hydrolase family 18 in less-thermophilic organisms [67]. The two chitinases share little sequence homology to each other, except in the catalytic region, where both have the catalytic glutamic acid residue that is conserved in all family 18 bacterial chitinases. The pH optimum of both recombinant chitinases is pH 6.0 with a temperature optimum between 90°C and 95°C. The chitinase A (ChiA) melts at 101°C, whereas the chitinase B (ChiB) has a melting temperature of 114°C. The ChiA exhibits no detectable activity towards chitooligomers smaller than chitotetraose, indicating that the enzyme is an endochitinase whereas the ChiB is a chitobiosidase, processively cleaving off chitobiose from the nonreducing end of chitin or other chitooligomers. The synergetic action of both thermoactive chitinases on colloidal chitin allows P. furiosus to grow on chitin as sole carbon source. Although a large number of bacterial chitin hydrolyzing enzymes has been isolated and their corresponding genes have been cloned and characterized, only few thermostable chitin-hydrolyzing enzymes are known. Those enzymes have been isolated from the thermophilic bacteria R. marinus, Microbispora sp. and Clostridium thermocellum (Table 8.1).

PROTEASES Proteases, which are involved in the conversion of proteins to amino acids and peptides, have been classified according to the nature of their catalytic site in three groups: serine, cysteine, aspartic, or metalloproteases. Proteases and proteasomes play a key role in the cellular metabolism of archaea and a variety of heat-stable proteases has been identified in hyperthermophilic archaea belonging to the genera Aeropyrum, Desulfurococcus, Sulfolobus, Staphylothermus, Thermococcus, Pyrobaculum, and Pyrococcus (Table 8.3). It has been found that most proteases from extremophilic archaea and bacteria belong to the serine type, and are stable at high temperatures even in the presence of high concentrations of detergents and denaturing agents [3,70]. Those properties of extracellular serine proteases are reported in a number of Thermococcus species and could be well illustrated by the extracellular enzyme from Thermococcus stetteri, which is highly stable (half-life of 2.5 h at 100°C) and resistant to chemical denaturation such as 1% SDS [4]. Heat-stable serine proteases were isolated from the cell-free supernatant of the hyperthermophilic archaea Desulfurococcus strain Tok12S1 and Desulfurococcus sp. SY [3]. A globular serine protease from Staphylothermus marinus was found to be extremely thermostable and is heat-resistant up to 125°C in the stalk-bound form [71]. A novel intracellular serine protease (pernisine) from the aerobic hyperthermophilic archaeon Aeropyrum pernix K1 is active at 90°C. The enzyme has a broad pH profile with an optimum at pH 9.0 for peptide hydrolysis [72,73]. A gene encoding a serine protease, named aereolysin has been cloned from P. aerophilum and the protein was modeled based on structures of subtilisin-type proteases [74]. Multiple proteolytic activities have been observed in P. furiosus. The cell-envelope associated serine protease of P. furiosus called pyrolysin was found to be highly stable with a half-life of 20 min at 105°C [75]. Proteases have also been characterized from the thermoacidophilic archaea S. solfataricus [76] and Sulfolobus acidocaldarius [74]. Thermostable serine proteases were also detected in a number of extreme thermophilic bacteria blonging to the genera Aquifex, Thermotoga, Thermus, and Fervidobacterium (Table 8.3). The enzyme from F. pennivorans is able to hydrolyze feather keratin forming amino acids and peptides. The enzyme, which has been named fervidolysin, is optimally active at 80°C and pH 10.0 [77]. The gene encoding fervidolysin was cloned and successfully expressed in E. coli. The gene encodes for a 73-kDa fervidolysin precursor, a 58-kDa mature fervidolysin, and a 14-kDa fervidolysin propeptide. Using site-directed mutagenesis, the active-site histidine residue at position 79 was replaced by

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Proteolytic enzymes (EC 3.4.21) Serine protease

Enzymes

Aeropyrum pernix Alicyclobacillus sendaiensis Aquifex aeolicus Aquifex pyrophilus Desulfurococcus mucosus Fervidobacterium islandicum Fervidobacterium islandicum AW-1 Fervidobacterium pennivorans (struct.) Geobacillus caldoproteolyticus Pyrobaculum aerophilum Pyrococcus abyssi Pyrococcus furiosus Staphylothermus marinus Sulfolobus acidocaldarius Sulfolobus solfataricus (struct.) Thermoactinomyces sp. Thermoactinomyces vulgaris

Strain

TABLE 8.3 Proteolytic Enzymes from Thermophiles

401 60 150 150 46–51 118 31 279

>200 58

34 37 54 43 43–54

MW (kDa)

80 85 95 80 100 80 70–80 >100 95 115 90 90 >90 85 60–65

90

Topt °C

8–9 3.9 8–8.5 7–9 7.5 8.0 9.0 10.0 8–9.0 7–9 9.0 6–9 9.0 2.0 6.5–8 11.0 7.5–9

pH Opt

0.33h at 105°C

1h at 80°C

1.5h at 100°C

>0.5h at 110°C 6h at 105°C 4.3h at 95°C

1h at 100°C

Thermostability (Half-Life)

Detergents, baking, brewing, amino acids production

Possible Applications

238 78 79 74 74 239 240 77 241 74 74 3 3 74 74 242 74

References

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46–51 45 52 128 79 95 320 170 37

Sulfolobus acidocaldarius Thermococcus kodakaraensis

Aeropyrum pernix Pyrococcus furiosus Pyrococcus furiosus Pyrococcus horikoshii OT3 Sulfolobus solfataricus Sulfolobus solfataricus Thermococcus sp. NA1

Acidic protease Thiol protease

Metalloprotease 100 100 75 >95 75 85 100

90 110

70 92 90 95 80 95 85 80 65 90–93 80 70

(struct.) – the protein has been crystallized and the three-dimensional structure is determined.

142 21.7 120 >669 281 178

44

Thermoanaerobacter keratinophilus Thermoanaerobacter yonseiensis Thermococcus aggregans Thermococcus celer Thermococcus kodakaraensis Thermococcus litoralis Thermococcus stetteri Thermomonospora fusca Thermoplasma acidophilum (struct.) Thermotoga maritima Thermus aquaticus Thermus sp.

>1.5h at 80°C

1h at 100°C

6.8 9.0 7.0 7.5 9.5 9.5 8.5–9 8.5 8.5 6–9.0 10.0 8.0 Detergents, baking, brewing, amino acid production

22h at 90°C 0.3h at 85°C

74 74 74 141 74 74 247

74 246

243 244 21 21 74 21 74 245 74 74 74 74

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an alanine residue. The resulting fervidolysin showed a single protein band corresponding in size to the 73-kDa fervidolysin precursor, indicating that its proteolytic cleavage resulted from an autoproteolytic process. From the thermoacidophile Alicyclobacillus sendaiensis, a novel thermostable collagenolytic member of the serine-carboxyl proteinase family was characterized [78]. This enzyme, with a molecular mass of 37 kDa, can be applied for the production of peptides from collagen. Organic solvents resistant aminopeptidase was described from the hyperthermophilic bacterium A. aeolicus [79]. In addition to serine proteases other subclasses of proteases have been identified in archaea (Table 8.3): aminopeptidases, metalloproteases, a thiol protease from T. kodakaraensis KOD1, an acidic protease from S. acidocaldarius, and a propylpeptidase and a new type of protease from P. furiosus. Indeed, the P. furiosus strain contains at least 13 different proteins with proteolytic activity. The amount of proteolytic enzymes produced worldwide on commercial scale is the largest. Heat stable proteases are useful enzymes, especially for the detergent industry. Serine alkaline proteases from thermophiles could be used as additives for laundering, where they have to resist denaturation by detergents and alkaline conditions. Proteases are also applied for peptide synthesis using their reverse reaction, mainly because of their compatibility with organic solvents. A number of heat-stable proteases are now used in molecular biology and protein chemistry. The protease S from P. furiosus is used to fragment proteins before peptide sequencing (TaKaRa Biomedicals). Carboxyand aminopeptidases from P. furiosus and S. solfataricus are used for protein N- or C-terminal sequencing [80,81].

BIOCATALYSIS WITH NONPOLYMERIC COMPOUNDS LIPASES AND ESTERASES Lipases hydrolyze triglycerides to glycerol and fatty acids and are also able to catalyze the reverse reaction in the presence of organic solvents. Lipases are an important group of biotechnologically relevant enzymes and they find applications in food, dairy, detergent, and pharmaceutical industries. Lipases produced by microbes and specifically bacterial lipases play a vital role in commercial ventures. Lipases are generally produced on carbon sources, such as oils, fatty acids, glycerol, or tweens in the presence of an organic nitrogen source. Bacterial lipases are mostly extracellular and are produced by submerged fermentation. Most lipases can act in a wide range of pH and temperature, though alkaline bacterial lipases are more common. Bacterial lipases generally have temperature optima in the range 30°C to 60°C. Lipases are serine hydrolases and have high stability in organic solvents. In addition, some lipases exhibit chemo-, regio-, and enantioselectivity. Very recently, five anaerobic thermophilic bacteria were found to produce extremely heat stable lipases. They are active at a broad temperature (50–95°C) and pH (3–11) range (unpublished results). In the field of industrial biotechnology, also esterases are gaining increasing attention because of their application in organic biosynthesis. In aqueous solution, esterases catalyze the hydrolytic cleavage of esters to form the constituent acid and alcohol, whereas in organic solutions, transesterification reaction is promoted. Both the reactants and the products of transesterification are usually highly soluble in the organic phase and the reactants may even form the organic phase itself. Several archaeal and bacterial esterases were successfully cloned and expressed in mesophilic hosts (Table 8.4). Esterases from archaea A. pernix, Pyrobaculum calidifontis, and Sulfolobus tokodaii exhibit high thermoactivity and thermostability and are active also in a mixture of a buffer and water-miscible organic solvents, such as acetonitrile and dimethyl sulfoxide [1]. The optimal activity for ester cleavage of the esterase from S. tokodaii strain 7 is at 70°C and pH 7.5 to 8.0. From the kinetic analysis, p-nitrophenyl butyrate is the better substrate than caproate and caprylate [81]. The P. furiosus esterase is the most thermostable (a half-life of 50 min at 126°C) and thermoactive (temperature optimum 100°C) esterase known to date [82]. A carboxylesterase from P. calidifontis, stable against heating and organic solvents, is active towards tertiary alcohol esters, a very rare

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feature among previously reported lipolytic enzymes [83]. A novel thermostable esterase from A. pernix K1 with an optimal temperature at 90°C exhibits additionally a phospholipase activity [84]. In our laboratory two thermoactive esterases from the thermoacidophilic archaeon P. torridus have been recently characterized after successful expression in E. coli (Antranikian, unpublished results). Both esterases are active at 50°C to 60°C and neutral pH. A gene coding the esterase from Archaeoglobus fulgidus was subjected to error-prone PCR in an effort to increase the low enantioselectivity towards the racemic mixture of p-nitrophenyl-2-chloropropionate to produce the S-2chloropropionic acid. This compound is an important intermediate in the synthesis of some optically pure compounds, such as a herbicide mecoprop [85]. A double mutant, Leu101Ile/Asp117Gly was obtained with increased preference in the opposite direction. The esterase from S. solfataricus P1 has been studied in detail for the chiral resolution of 2-arylpropionic esters [86]. Thus, the application of the esterase toward R,S-naproxen methyl ester yields highly optically pure S-naproxen [ee(p) > 90%] [86,87]. The enzyme is activated by DMSO to various extents, due to small changes in the enzyme structure resulting in an increase in its conformational flexibility. Thus, the addition of cosolvents, which is useful for solubilization of hydrophobic substrates in water, also serves as activators in applications involving thermostable biocatalysts at suboptimal temperatures [88]. Interestingly, experimental data on kinetic resolution of α-arylpropionic acid revealed that a carboxylesterase from S. solfataricus P2 hydrolyzes the R-ester of racemic ketoprofen methylester with enantiomeric excess of 80% [89]. A gene encoding a thermostable esterase was cloned from the bacterium Thermoanaerobacter tengcongensis and over-expressed in E. coli. The recombinant esterase, with a molecular mass of 43 kDa hydrolyzes tributyrin but not olive oil. The esterase is optimally active at 70°C (over 15 min) and at pH 9. It is highly thermostable, with a residual activity greater than 80% after incubation at 50°C for more than 10 h [90].

ALCOHOL DEHYDROGENASES Dehydrogenases are enzymes belonging to the class of oxidoreductases. Within this class, alcohol dehydrogenases (ADHs) (EC 1.1.1.1, also named keto-reductases) represent an important group of biocatalysts due to their ability to stereospecifically reduce prochiral carbonyl compounds. ADHs can be used efficiently in the synthesis of optically active alcohols, which are key building blocks in the synthesis of chirally pure pharmaceutical agent. From a practical point of view, ADHs that use NADH as cofactor are of particular importance, because they represent an established method to regenerate NADH efficiently. By contrast, for NADP-dependent enzymes the cofactor-recycling systems that are available are much less efficient [91]. The secondary specific ADH, which catalyzes the oxidation of secondary alcohols and, less readily, the reverse reaction (the reduction of ketones) has a promising future in biotechnology. Although ADHs are widely distributed among microorganisms, only few examples derived from hyperthemophiles are currently known (Table 8.4). The ADH from the archaeon S. solfataricus requires NAD as cofactor and contains Zn ions. In contrast to the enzyme from T. litoralis, it lacks metal ions and catalyzes preferentially the oxidation of primary alcohols, using NADP as cofactor. The enzyme is thermostable, having half-lives of 15 min at 98°C and 2 h at 85°C [92]. The pyrococcal ADH is the most thermostable short-chain ADH (half-life of 150 h at 80°C) known to date [93]. The NADP-dependent ADH from T. hydrothermalis oxidizes a series of primary aliphatic and aromatic alcohols preferentially from C2 to C8 but is also active towards methanol and glycerol and is stereospecific for monoterpenes [94]. The enzyme structure is pH-dependent, being a tetramer (45 kDa per subunit) at pH 10.5 (pH optimum for alcohol oxidation), and a dimmer at pH 7.5 (pH optimum for aldehyde reduction). Among the extreme thermophilic bacteria, T. ethanolicus 39E and Thermoanaerobacter brockii were shown to produce an ADH, whose gene was cloned and expressed in E. coli. Interestingly, a mutant has been found to posses an advantage over the wild type enzyme by using the more stable cofactor NAD instead of NADP [91]. An ADH was purified from an extremely thermophilic bacterium, Thermomicrobium roseum. The pI of the homodimeric enzyme (43 kDa/subunit) was determined

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to be 6.2, while its optimum pH and temperature are 10.0 and 70°C, respectively [95]. The enzyme oxidizes mainly primary aliphatic alcohols.

GLUCOSE AND ARABINOSE ISOMERASES Glucose isomerase or xylose isomerase (d-xylose ketol-isomerase; EC 5.3.1.5) catalyzes the reversible isomerization of d-glucose and d-xylose to d-fructose and d-xylulose, respectively. The enzyme has the largest market in the food industry because of its application in the production of highfructose corn syrup (HFCS). Glucose isomerases are widely distributed in mesophilic microorganisms and intensive research efforts were directed towards improving their suitability for industrial application. In order to reach fructose concentration higher than 55% the reaction must approach 110°C. Improved thermostable glucoses isomerases have been engineered from mesophilic enzymes [96]. Mostly thermophilic bacteria were found to produce glucose isomerases (Table 8.4). The gene encoding a xylose isomerase of Thermus flavus AT62 was cloned and the DNA sequence was determined. The enzyme has an optimum temperature at 90°C and pH 7.0; divalent cations are required for enzyme activity [97]. Thermoanaerobacterium strain JW/SL-YS 489 forms a xylose isomerase, which is optimally active at pH 6.4 and 60°C or pH 6.8 and 80°C. Like other xylose isomerases, this enzyme requires divalent cations for thermal stability (stable for 1 h at 82°C in the absence of substrate). The gene encoding the xylose isomerase of Thermoanaerobacterium strain JW/SLYS 489 was cloned and expressed in E. coli [24]. Comparison of the deduced amino acid sequence with sequences of other xylose isomerases showed that the enzyme has 98% homology with a xylose isomerase from a closely related bacterium Thermoanaerobacterium saccharolyticum B6A-RI. A thermostable glucose isomerase was purified and characterized from T. maritima. This enzyme is stable up to 100°C, with a half-life of 10 min at 115°C [24]. Interestingly, the glucose isomerase from T. neapolitana displays a catalytic efficiency at 90°C, which is 2 to 14 times higher than any other thermoactive glucose isomerases at temperatures between 60°C and 90°C [98]. Arabinose isomerase (EC 5.3.1.4) catalyzes the reversible isomerization of arabinose to ribulose. Thermoactive enzymes have been reported to convert d-galactose to d-tagatose, a novel and natural sweetener [99]. Such enzymes have been described from the thermophilic bacteria A. acidocaldarius, Geobacillus stearothermophilus, Thermoanaerobacter mathranii, T. maritima, and T. neapolitana (Table 8.4).

C–C BOND FORMING ENZYMES Synthetic building blocks bearing hydroxylated chiral centers are important targets for biocatalysis. C–C bond forming enzymes, such as aldolases and transketolases, have been investigated for new applications, and various strategies for the synthesis of sugars and related oxygenated compounds have been developed [100]. The use of aldolases in stereoselective C–C bond forming reactions is applicable for asymmetric synthesis of carbohydrates, leading to the development of new therapeutics and diagnostics. However, many aldolases display narrow specificity, often prefer phosphorylated substrates, which can limit the product range of chiral aldols. In contrast, an extremely thermostable aldolase (half-life 2.5 h at 100°C) from S. solfataricus, actively expressed in E. coli, possesses a broad specificity for nonphosphorylated substrates and has a great potential for use in asymmetric aldol reactions (Table 8.4) [2]. This aldolase represents a rare example of an enzyme that exhibits no diastereocontrol for the aldol condensation of its natural substrates pyruvate and glyceraldehyde. Recently, it was demonstrated that the stereoselectivity of the enzyme has been induced by employing the substrate engineering procedure [101]. The decameric transaldolases from Methanocaldococcus jannaschii retains full activity for 4 h at 80°C [102]. The aerobic bacterium T. aquaticus produces fructose aldolase, which is stable after heating at 90°C for 2 h [2].

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271 272 133 245 165 108 56 170 95 172 224 225 212 232 230

Methanocaldococcus jannaschii Pyrococcus furiosus Sulfolobus solfataricus (struct.) Thermoproteus tenax Thermus aquaticus (struct.)

Pseudonocardia thermophila Sulfolobus solfataricus (struct.)

Pyrococcus furiosus Pyrococcus horikoshii OT3 Thermococcus litoralis

Alicyclobacillus acidocaldarius Geobacillus stearothermophilus Thermoanaerobacter mathranii Thermotoga maritima Thermotoga neapolitana

Aldolase (EC 4.1.2.13) (EC 4.1.2.14)

Amidase (EC 3.5.1.4)

Aminoacylase (EC 3.5.1.14)

Arabinose isomerase (EC 5.3.1.3)

55 71 80.5 192 200

MW (kDa)

184 160 160 86

Strain

Aeropyrum pernix (struct.) Methanoculleus thermophilicus Pyrococcus furiosus Sulfolobus solfataricus (struct.) Thermococcus hydrothermalis Thermococcus litoralis Thermococcus sp. AN1 Thermococcus sp. ES-1 Thermococcus zilligii Thermoanaerobacter brockii (struct.) Thermoanaerobacter ethanolicus Thermomicrobium roseum

Alcohol dehydrogenase (EC 1.1.1.1)

Enzymes

TABLE 8.4 Thermoactive Enzymes of Industrial Relevance

10.0

90 70

65 80 65 90 85

100 95 85

70 95

6–6.5 7.5 8.0 7.5 7.5

6.5 7.5 8.0

4–8 7.5

7–8.5

7.0

85

80 50 100 50 80

7.5 7.5 7.5 8.8 7.0

pH Opt

90 70 90 95 80 80 85

Topt °C

4h at 80°C

10h at pH5.0

>48h at 90°C 1.7h at 85°C

1.2h at 70°C 25h at 80°C

>1h at 97°C

2.5h at 100°C

24h at 80°C

1.7h at 90°C

0.25h at 80°C

0.25h at 80°C 2h at 85°C

150h at 80°C

Thermostability (Half-Life)

Sweeteners in food industry

Pharmaceutical industry (production of stereoisomers)

Synthesis of fine chemicals

Synthesis of carbohydrates

Stereoselective transformation of ketones to pure chiral alcohols

Possible Applications

continued

99 253 254 255 256

138 141 140

103 3

102 250 251 250 252

248 91 93 91 94 92 249 92 91 91 92 95

References

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Thermotoga maritima

Pyrococcus woesei Thermotoga maritima Thermus sp. A4

β-Galactosidase (EC 3.2.1.23)

β-Glucuronidase (EC 3.2.1.31)

Thermotoga maritima Thermus sp.

α- Galactosidase (EC 3.2.1.22)

75

61

54

43

33 34

35 128

6.5

70–80

21

Pyrobaculum calidifontis Pyrococcus furiosus Sulfolobus acidocaldarius Sulfolobus shibatae Sulfolobus solfataricus P1 Sulfolobus solfataricus P2 Sulfolobus tokadaii Thermoanaerobacter tengcongensis

7.0 9.5

90 70 70

18 35.5

Aeropyrum pernix (struct.) Archaeoglobus fulgidus (struct.) Methanocaldococcus jannaschii (struct.) Picrophilus torridus

85

90 80 70

90–95 75

90 100 90 90 100 80 70 70

6.5

4.0 5.3 6.5

5–5.5

0.33h at 120°C

6.0 5–6 7.4 7.5–8 9.0

3h at 85°C

>2h at 85°C

1.2h at at 90°C 1h at 70°C

>10h at 50°C

0.66h at 80°C

2h at 100°C 2h at 120°C

1h at 100°C

>6h at 100°C

3h at 90°C

Thermostability (Half-Life)

7.0

7.5–8

Esterase (EC 3.1.1.1)

>60

65

Aeropyrum pernix

8.0

pH Opt

Cysteine synthase (EC 4.2.1.22)

90

Topt °C

178

MW (kDa)

Thermus brockianus

Strain

Catalase (EC 1.11.1.6)

Enzymes

TABLE 8.4 (continued)

Synthesis of oligosaccharides

Synthesis of oligosaccharides, production of dietary milk products

Sugar processing

Biotransfor-mation in organic solvents

Synthesis of sulfur-organic compounds

Industrial bleaching

Possible Applications

262

142 24 261

24 260

Antranikian, unpublished 83 86 3 3 86 89 81 90

84 258 259

144

257

References

136 Thermophiles: Biology and Technology at High Temperatures

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9214_C008.indd 137

90–95

92

53

28 28

23

84 87

60

Thermotoga maritima

Thermus thermophilus HB27

Caldanaerobacter subterraneus Thermoanaerobacter thermohydrosulfuricus

Pyrococcus horikoshii

Sulfolobus acidocaldarius Sulfolobus solfataricus

Pyrococcus abyssi

β-Fructosidase (EC 3.2.1.26)

Laccase (EC 1.10.3.2)

Lipase (EC 3.1.1.3)

N-Methyltransferase (EC 2.1.1.17)

Maltooligosyl trehalose synthase (EC 5.4.99.15)

Nitrilase (EC 3.5.5.1)

60–90

75 75

90–100

(struct.) – the protein has been crystallized and the three-dimensional structure is determined.

185 200

200

75 75

80 >100 97 70 90 90 90

200

200

65 70 80

200

Clostridium thermosulfurigenes Thermoanaerobacter ethanolicus Thermoanaerobacterium saccharolyticum Thermoanaerobacterium sp. Thermotoga maritima Thermotoga neapolitana Thermus aquaticus Thermus caldophilus (struct.) Thermus flavus Thermus thermophilus

Glucose isomerase (EC 5.3.1.5)

6–8

5.0 5.0

8.5

7.0 8.0

4.5–5.5

6h at 90°C

72h at 80°C 2h at 85°C

>2h at 100°C

2h at 80°C 2h at 85°C

>14h at 80°C

2h at 95°C

7.0 7.0

5.5

1h at 80°C 0.2h at 120°C 2h at 90°C 240h at 70°C

0.6h at 85°C

6.8 7.0 7.1 5.5

7.0 7–7.5

Production of mononitriles

Synthesis of phosphadylcholine for medicine and food

Biotrans-formation

Polymer synthesis, biosensors

Confectionery industry

Sweeteners in food industry

105

59 58

145

Antranikian, unpublished Antranikian, unpublished

267

24

264 24 98 24 265 97 266

24 263 24

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Thermophiles: Biology and Technology at High Temperatures

NITRILE-DEGRADING ENZYMES Nitrile-degrading enzymes are of considerable importance in industrial biotransformations, and to date several processes have been developed for chemical and pharmaceutical industries for the production of optically pure compounds, drugs, acrylic, and hydroxamic acids [103]. Nitrile degrading enzymes play also a significant role in the protection of the environment due to their capability to eliminate highly toxic nitriles. Thermostable amidases and nitrilases are gaining more attention, especially in enzymatic processes in mixtures of organic solvent or in the formation of highly pure products with a concomitant reduction of wastes. Amidases (EC 3.5.1.4) catalyze the conversion of amides to the corresponding carboxylic acids and ammonia. Nitrilases (EC 3.5.5.1) are thiol enzymes that convert nitriles directly to the corresponding carboxylic acids with release of ammonia. A number of bacterial amidases and nitrilases have been purified and characterized. Very little, however, is known on the enzymes that are active at high temperatures. Amidases are highly S-enantioselective, usually forming the optically pure acids with an enantiomeric excess above 99%. The only amidase derived from archaea is the amidase from the thermoacidophile S. solfataricus (Table 8.4). This enzyme is S-stereoselective with a broad substrate spectrum and is optimally active at 95°C [104]. Very recently, the first thermoactive and thermostable amidase from the thermophilic actinomycete Pseudonocardia thermophila has been purified and characterized [103]. The amidase is active at a broad pH (pH 4–9) and temperature range (40–80°C) and has a half-life of 1.2 h at 70°C. The amidase has a broad substrate spectrum, including aliphatic, aromatic, and amino acid amides. The amidase is highly S-stereoselective for 2-phenylpropionamide with an enantiomeric excess of >95% at 50% conversion of the substrate. Recently, the fi rst archaeal nitrilase from the hyperthermophile Pyrococcus abyssi, regiospecific towards aliphatic dinitriles, was cloned and characterized in our laboratory [105]. The enzyme is highly thermostable, having a half-life at 90°C for 6 h. Thermoactive nitrilases described so far were isolated from the bacteria Acidovorax facilis 72 W and Bacillus pallidus Dac521.

DNA PROCESSING ENZYMES POLYMERASE CHAIN REACTION Thermostable DNA polymerases (EC 2.7.7.7) play a major role in a variety of molecular biological applications, for example, DNA amplification, sequencing or labeling (Table 8.5). They are the key enzymes in the replication of cellular information present in all life forms. They catalyze, in the presence of Mg2+-ions, the addition of a deoxyribonucleoside 5′-triphosphate onto the growing 3′-OH end of a primer strand, forming complementary base pairs to a second strand. More than 100 DNA polymerase genes have been cloned and sequenced from various organisms, including thermophilic bacteria and archaea [106]. The first described polymerase chain reaction (PCR) procedure utilized the Klenow fragment of E. coli DNA polymerase I, which was heat-labile and had to be added during each cycle following the denaturation and primer hybridization steps. Introduction of thermostable DNA polymerases in PCR facilitated the automation of the thermal cycling part of the procedure. The DNA polymerase I from the bacterium T. aquaticus, called Taq polymerase, was the first thermostable DNA polymerase characterized and applied in PCR. Due to the absence of a 3′-5′-exonuclease activity, this enzyme is unable to excise mismatches and as a result, the base insertion fidelity is low. The use of high fidelity DNA polymerases is essential for reducing the increase of amplification errors in PCR products that will be cloned, sequenced and expressed. Several thermostable DNA polymerases with 3′-5′-exonuclease-dependent proofreading activity have been described and the error rates (number of misincorporated nucleotides per base synthesized) for these enzymes have been determined. Archaeal polymerases from Pyrococcus or Thermococcus species with stringent proofreading abilities are of widespread use. Archaeal proofreading polymerases such as Pwo pol from P. woesei, Pfu pol from P. furiosus, Deep Vent™ pol from Pyrococcus strain GB-D or Vent™ pol from T. litoralis have an error rate that is up to 10-fold lower than that of

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Taq polymerase. The 9°N-7 DNA polymerase from Thermococcus sp. strain 9°N-7 has a five-fold higher 3′-5′-exonuclease activity than T. litoralis DNA polymerase. However, Taq polymerase was not replaced by these DNA polymerases because of their low extension rates among other factors. DNA polymerases with higher fidelity are not necessarily suitable for amplification of long DNA fragments because of their potentially strong exonuclease activity. The recombinant KOD1 DNA polymerase from T. kodakaraensis KOD1 has been reported to show low error rates, high processivity and highest known extension rate, resulting in an accurate amplification of target DNA sequences up to 6 kb [107,108]. Recently, the PCR technique has been improved to allow low error synthesis of long amplificates (20–40 kb) by adding small amounts of thermostable archaeal proofreading DNA polymerases, containing 3′-5′-exonuclease activity, to Taq or other nonproofreading DNA polymerases. In this long PCR, the reaction conditions are optimized for long extension by adding different components such as gelatine, Triton X-100 or bovine serum albumin to stabilize the enzymes and mineral oil to prevent evaporation of water in the reaction mixture. In order to enhance specificity, glycerol or formamide are added. The supplement of the PCR reaction mixtures with recombinant P. woesei dUTPase improves the efficiency of the reaction and allows amplification of longer targets [109]. Low fidelity mutants of P. furiosus polymerase were also created for the performance in error-prone PCR [110]. The ssDNA-binding proteins are known to be involved in eliminating DNA secondary structure, and are key components in DNA replication, recombination and repair. The archaeal ssDNA-binding proteins derived from M. jannashii, Methanothermobacter thermoautotrophicum, and A. fulgidus are therefore useful reagents for genetic engineering and other procedures involving DNA recombination, such as PCR [111].

DNA SEQUENCING DNA sequencing by the Sanger method has undergone countless refinements in the last twenty years [112]. A major step forward was the introduction of thermostable DNA polymerases leading in the cycle sequencing procedure. This method uses repeated cycles of temperature denaturation, annealing and extension with dideoxy-termination to increase the amount of sequencing product by recycling the template DNA. Due to this “PCR like” amplification of the sequencing products several problems could have been overcome. Caused by the cycle denaturation, only fmoles of template DNA are required, no separate primer annealing step is needed and unwanted secondary structures within the template are resolved at high temperature elongation. The first enzyme used for cycle sequencing was the thermostable DNA polymerase I from Thermus thermophilus or T. flavus [113,114]. The enzyme displays 5′-3′-exonuclease activity that is undesirable because of the degradation of sequencing fragments. A combination of thermostable enzymes has been developed that produces higher quality cycle sequences. Thermo Sequenase DNA polymerase is a thermostable enzyme engineered to catalyze the incorporation of ddNTPs with an efficiency of several 1000-fold better than other thermostable DNA polymerases. Since the enzyme also catalyzes pyrophosphorolysis at dideoxy termini, a thermostable inorganic pyrophosphatase is needed to remove the pyrophosphate produced during sequencing reactions. T. acidophilum inorganic pyrophosphatase (TAP) is thermostable and effective for converting pyrophosphate to orthophosphate. The combination of Thermo Sequenase polymerase and TAP for cycle sequencing yields sequence data with uniform band intensities and allow the determination of longer, more accurate sequence reads. Uniform band intensities also facilitate interpretation of sequence anomalies and the presence of mixed templates. Sequencing PCR products of DNA amplified from heterozygous diploid individuals results in signals of equal intensity from each allele [115]. Another extremely stable inorganic pyrophosphatase was purified from S. acidocaldarius. The complete activity of the enzyme remained after incubation at 100°C for 10 min [116]. Highly thermostable alkaline phosphatases, which dephosphorylate linear DNA fragments, were also identified in archaea (Table 8.5). The alkaline phosphatase from P. abyssi dephosphorylates linear DNA fragments with efficiencies of 94% and 84% regarding to cohesive and blunt ends, respectively [117].

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104.8 100 101 97 90 90 98.9 90

Rhodothermus marinus Sulfolobus acidocaldarius Sulfolobus solfataricus Thermoanaerobacter yonseiensis Thermococcus aggregans Thermococcus kodakaraensis (struct.) Thermococcus litoralis Thermococcus sp. 9°N-7 Thermus aquaticus

KOD1

Vent pol

9°N-7 pol

Tay

Taq

Tca Thermus caldophilus

90 90.6 90

Pyrococcus furiosus Pyrococcus sp. GB-D Pyrococcus woesei

Pwo pol

Tfi

90

Pyrococcus abyssi

Deep Vent pol

75

75

70–80

8.7

9.0

8.8

6.5

6.7h at 95°C

2h at 100°C

1.2h at 90°C 12h at 95°C 6.8 7.5 70–80 75 70–80

0.1h at 90°C

0.25h at 87°C

2min at 90°C

4h at 95°C 8h at 100°C

5h at 100°C

>5h at 90°C

0.5h at 85°C 0.5h at 100°C

Thermostability (Half-Life)

7.5

9.0 8–9

7.3

pH Opt

75

65–75

55

72–78 70–80

70–80

70–80

90

Pyrobaculum islandicum

Pfu pol

Topt °C

60–70

MW (kDa) 108 88

Strain Aeropyrum pernix (pol I) Aeropyrum pernix (pol II) Carboxydothermus hydrogenoformans

DNA Polymerase (EC 2.7.7.7)

Enzymes

TABLE 8.5 DNA-Modifying Enzymes

24

Roche molecular biochemicals

3

3

272 273 274 3

3

271

3 3 3

270

269

268 268 Roche molecular biochemicals

References

140 Thermophiles: Biology and Technology at High Temperatures

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Pyrococcus abyssi Thermotoga maritima Thermotoga neapolitana Thermus sp. 3041 Thermus yunnanensis Aquifex pyrophilus Sulfolobus acidocaldarius Thermoplasma acidophilum (struct.) Archaeoglobus fulgidus Methanocaldococus jannashii Methanothermobacter thermoautothrophicum

Alkaline phosphatase (EC 3.1.3.1)

Inorganic pyrophosphatase (EC 3.6.1.1)

ssDNA-binding proteins

80–95 56

7.5–8 6.5

8–10.0

70–80

104 105 80

11.0 8.0 9.9

7.0 6–7.0 7.0 8.0

8–8.6

8.8

70 65 85

65 45–80 >55 50–70 65 100 65

70–75

108

52

80 62

82

87

(struct.) – the protein has been crystallized and the three-dimensional structure is determined.

Aquifex pyrophilus Pyrococcus furiosus Rhodothermus marinus Sulfolobus shibatae Thermococcus fumicolans Thermococcus kodakaraensis Thermus scodoductus

Thermus filiformis Thermus thermophilus Methanopyrus kandleri

DNA Ligase (EC 6.5.1.1)

DNA topoisomerase type I-group B

Tth

at 80–95°C

5h at 90°C 4h at 90°C

0.5h at 90°C

>1h at 95°C >1h at 95°C 0.1h at 90°C 0.15h at 90°C

111 111 111

279 116 115

117 276 24 277 278

124 Stratagene 125 123 122 121 125

Roche molecular biochemicals Roche molecular biochemicals 275

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LIGASE CHAIN REACTION A variety of analytical methods is based on the use of thermostable ligases. Of considerable potential is the construction of sequencing primers by high temperature ligation of hexameric primers, the detection of trinucleotide repeats through repeat expansion detection or DNA detection by circularization of oligonucleotides [118]. Up to now several archaeal DNA ligases, displaying nick joining and blunt-end ligation activities using either ATP or NAD+ as a cofactor, have been identified and characterized in detail (Table 8.5). Over the years several additional thermostable DNA ligases were discovered. The ligase from the archaeon Acidianus ambivalens is NAD+independent but ATP-dependent similar to the enzymes from bacteriophages, eukaryotes and viruses [119]. The DNA ligase from a hyperthermophilic archaeon T. kodakaraensis is also ATP-dependent [120]. Sequence comparison with previously reported DNA ligases indicated that the ligase is closely related to the ATP-dependent DNA ligase from Methanobacterium thermoautotrophicum H, a moderate thermophilic archaeon, along with putative DNA ligases from Euryarchaeota and Crenarchaeota. The optimum pH of the recombinant monomeric enzyme is 8.0, the optimum concentration of Mg2+ is 14 to 18 mM, and K+ is 10 to 30 mM. The protein does not display single-stranded DNA ligase activity. At enzyme concentrations of 200 nM, a significant DNA ligase activity is observed even at 100°C. Surprisingly, the protein displays a DNA ligase activity also when NAD+ is added as the cofactor [121]. The ability for DNA ligases, to use either ATP or NAD+, as a cofactor, appears to be specific of DNA ligases from Thermococcales. Also a DNA ligase from Thermococcus fumicolans displays nick joining and blunt-end ligation activity using either ATP or NAD+, as a cofactor [122]. The optima of temperature and pH of the ligase are 65°C and 7.0, respectively. The presence of MgCl 2 (optimally at 2 mM) is required for the enzymatic activity. In contrast to that the recombinant ATPdependent ligase from the thermoacidophilic crenarchaeon Sulfolobus shibatae is more active in the presence of Mn+2 ions than in the presence of other divalent cations such as Mg+2 or Ca+2 [123]. Splicing ligase activities were characterized from Aquifex pyrophilus [124], Thermus scotoductus and R. marinus [125]. The archaeal strains P. furiosus, Thermococcus marinus and Thermococcus radiotolerans are resistant to high levels of ionizing and ultraviolet radiation and therefore may have a unique method of removing damaged DNA [126]. A thermostable flap endonuclease from P. furiosus is described, which cleaves the replication fork-like structure endo/exonucleolytically [127]. The 06-methylguanine-DNA methyltransferase is the most common form of cellular defense against the biological effects of 06-methylguanine in DNA. The thermostable recombinant 06-methylguanine-DNA methyltransferase from T. kodakaraensis is functional in vivo and complements the mutant phenotype, making the cells resistant to the cytotoxic properties of the alkylating agent N-methylN′-nitro-N-nitrosoguanidine [128]. A thermostable type I group B DNA topoisomerase has been isolated and purified from the hyperthermophilic methanogen Methanopyrus kandleri [129]. The enzyme is active over a wide range of temperatures and salt concentrations and does not require magnesium or ATP for its activity, which makes manipulations on DNA more convenient and more efficient. Exploitation of the common features and the differences of topoisomerases will be important for modeling of novel drugs and understanding of the action of cancer chemotherapeutic agents.

CHEMICAL PRODUCTS In addition to above described applications, there is a great need for the biotechnological production of fine chemicals and building blocks thereby replacing or optimizing already existing chemical processes. Specially, the demand for the synthesis of optically pure compounds by specific enzymes for pharmaceutical and chemical industries is increasing.

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COMPATIBLE SOLUTES Accumulation of osmotically active substances, so-called compatible solutes, by uptake or de novo synthesis, enables microorganisms to reduce the difference between osmotic potentials of the cell cytoplasm and the extracellular environment. Those compounds are highly water-soluble sugars or sugar alcohols, other alcohols, amino acids, or their derivatives. They gained an increasing attention in biotechnology due to their action as stabilizers of biomolecules (enzymes, DNA, membranes, tissues) and stress-protecting agents (Table 8.6) [130]. Additionally, compatible solutes support the high-yield periplasmic production of functional active recombinant proteins in different expression systems [131]. Di-myo-inositol-1,1′-phosphate is the most widespread solute of hyperthermophilic archaea and was not detected in a mesophile. This thermoprotective compound was found in a variety of strains, belonging to the genera Methanococcus, Pyrococcus, Pyrodictium, Pyrolobus, and Thermococcus [132]. In most of these archaea, an increase of the solute concentrations is observed at growth temperatures above the optimum, reaching 20-fold in case of P. furiosus grown at 101°C. In contrast, the concentration of mannosylglycerate, detected in the euryarchaeotes of the genera Archaeoglobus, Pyrococcus, Thermococcus, Methanothermus fervidus and in the crenarchaeote A. pernix, increases concomitantly with the salinity of the medium and serves therefore as a compatible solute under salt stress. Mannosylglycerate has been observed to have a profound effect on thermoprotection and protection against desiccation of enzymes from mesophilic, thermophilic and hyperthermophilic microorganisms [1]. The biosynthetic routes for the synthesis of mannosylglycerate in the archaeon P. horikoshii and di-myo-inositol-1,1′-phosphate in P. woesei and Methanococcus igneus have been investigated [133–135]. The hyperthermophilic archaeon A. fulgidus accumulates a very rare compound diglycerol phosphate under salt and temperature stress. This solute demonstrated a considerable stabilizing effect against heat inactivation of various dehydrogenases and a strong protective effect on bacterial rubredoxins (with a four-fold increase in the half-lives) [136]. A compatible solute, cyclic 2,3-bisphosphoglycerate, has been detected only in methanogenic archaea such as M. kandleri. The thermoprotective role of this solute was proven by in vitro studies showing that the solute protects selected enzymes from M. kandleri against thermal denaturation [137].

OTHER COMPOUNDS Due to their chiral specificity in the synthesis of acylated amino acids, aminoacylases (EC 3.5.1.14) are attractive candidates for application in fine chemistry [138]. The l-aminoacylase from T. litoralis accepts a wide range of amino acid side chains and N-protecting groups and was recently commercialized [139]. The application of the thermostable enzyme reduces the process time, simplifies filtration procedure, improves substrate solubility and increases the enantiomeric excess to 99%. In contrast to the chemical process, the reaction completes overnight at 70°C, which avoids boiling in 20 equivalent volumes of 6 M HCl for two days [140]. Two thermostable zinc-containing aminoacylases were also characterized from Pyrococcus species (Table 8.6) [138,141]. Thermostable β-galactosidase e.g. from P. woesei is potentially useful for whey utilization and for the preparation of low-lactose milk and other dairy products or it can be used as a catalyst in the synthesis of galactooligosaccharides, using lactose as substrate and a nucleophile [142]. In recent years carotenoids have gained importance in nutraceutical field. These pigments have been shown to possess physiological function in the prevention of cancer and heart diseases, enhancing in vitro antibody production and as precursors for vitamins. The majority of carotenoids are synthesized from lycopene. The β-carotene, the precursor of vitamin A, is biosynthesized directly from lycopene by β-cyclization at both termini, and the reaction is catalyzed by lycopene β-cyclase. Recently, lycopene β-cyclase was predicted in the carotenogenic gene cluster in the genome of the thermoacidophilic archaeon S. solfataricus [143]. The recombinant expression of

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TABLE 8.6 Other Applications of Thermophiles Product

Strain

Application

References

Chaperones, chaperonins, maltodextrin-binding proteins, peptidyl-prolyl cis-trans isomerases

Methanothermococcus sp. Pyrococcus spp. Pyrococcus horikoshii Sulfolobus shibatae Thermococcus spp. Thermus spp.

Stabilization and solubilization of recombinant proteins

280–286

Compatible solutes (mannosylglycerate, mannosylglyceramide, diglycerol phosphate, di-myo-inositol-phosphate, N-acetyl-β-lysine, trehalose, 2-sulfotrehalose, cyclic-2,3bisphosphoglycerate)

Aeropyrum pernix Archaeoglobus spp. Methanococcus igneus Methanopyrus kandleri Methanosarcina thermophila Methanothermus fervidus Pyrobaculum aerophilum Pyrococcus spp. Pyrodictium occultum Pyrolobus fumarii Rhodothermus marinus Thermococcus spp. Thermus spp.

Cosmetics, biomolecules and tissue stabilizers, molecular biology

130–132

Cytochrome P450

Sulfolobus solfataricus Thermus thermophilus

Selective regio and stereospecific hydroxylations in chemical synthesis

24 287

Hydrogen gas (Ni-Fe hydrogenase)

Thermococcus kodakaraensis Caldicellulosiruptor saccharolyticus Carboxydothermus hydrogenoformans Fervidobacterium pennavorans Thermoanaerobacter tengcongensis Thermotoga elfii Thermotoga neapolitana

H2 production

154–159

S-layer proteins, lipids, liposomes

Methanobrevibacter smithii Methanococcus spp. Methanothermus spp. Staphylothermus marinus Sulfolobus solfataricus

Vaccine development, diagnostics, biomimetics, drugs, nanotechnology

149–151, 288–290

Whole cell biocatalysis

Thermococcus barophilus

Formation of gels and starch granules

291

Thermococcus gammatolerans Thermococcus marinus Thermococus radiotolerans Sulfolobus metallicus Pyrococcus furiosus

Detoxification of halogenated organic compounds and toxic chemicals, heavy metals, nuclear waste treatment Rubber recycling

126, 292

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the gene in E. coli resulted in the accumulation of lycopene β-carotene in the cells. Due to its great antioxidant activity canthaxanthin has been used as food and feed additive, in cosmetics and pharmaceuticals. Sulfur-containing organic compounds have been synthesized mainly chemically. Due to the side reactions, the chemical synthesis of those molecules results in the unavoidable production of impurities in the product and environmental pollution by the formation of by-products such as sulfur oxides. In order to overcome these problems, a method for the synthesis of sulfur-containing organic compounds using O-acetylserine sulfhydrylase has been proposed [144]. A recombinant cysteine synthase from A. pernix is highly stable within pH 6 to 10 and resistant to organic solvents. Due to its high heat resistance, the enzyme can act on highly concentrated substrate solutions compared to mesophilic thermolabile cysteine synthases. Phosphatidylethanolamine N-methyltransferase plays a key role in the synthesis of phosphatidylcholine, a main component of liposomal membrane, which is present in various foods as digestible surfactant. It plays an important role in medicine as a component of microcapsule for drugs. A phosphatidylethanolamine N-methyltransferase from P. horikoshii was cloned and expressed in E. coli. The enzyme is thermostable and is active in organic solvents. This opens the possibility to develop a new process for the synthesis of polar lipids with high optical purity [145].

THERMOPHILES AS CELL FACTORIES BIOMINING The development of industrial mineral processing has been established in several countries, such as South Africa, Brazil, and Australia. Iron- and sulfur-oxidizing microorganisms are used to release occluded gold from mineral sulfides. Most industrial plants for biooxidation of gold-bearing concentrates have been operated at 40°C with mixed cultures of mesophilic bacteria of the genera Thiobacillus or Leptospirillum. In subsequent studies a dissimilatosy iron-reducing archaea Pyrobaculum islandicum and P. furiosus were shown to reduce gold chloride to insoluble gold [146]. The potential of thermophilic sulfide-oxidizing archaea in copper extraction has attracted interest due to the efficient extraction of metals from sulfide ores that are recalcitrant to dissolution [19]. The acidophilic archaea Sulfolobus metallicus and Metallosphaera sedula tolerate up to 4% of copper and have been exploited for mineral biomining [147]. The efficiency of copper extraction from chalcopyrite by thermoacidophilic archaea was influenced by the characteristics of mineral concentrates. Between 40% and 60% copper extraction was achieved in primary reactors and more that 90% extraction in secondary reactors with overall residence times of about six days [147]. The handling and recycling of spent tyres are a significant and worldwide problem. The reuse of rubber material is preferable from an economic and environmental point of view. The anaerobic sulfate-reducing thermophilic archaeon P. furiosus was investigated for its capacity to desulfurize rubber. The tyre rubber treated with P. furiosus for 10 days was subsequently vulcanized with virgin rubber material (15% w/w). This results in the desulfurization of ground rubber and leads to a product with good mechanical properties [148]. The thermoacidophilic archaeon S. acidocaldarius has been also tested for desulfurization of rubber material [148].

LIPIDS AND PEPTIDES Liposomes are artificial spherical closed vesicles consisting of one or more lipid bilayers. Liposomes made from ether phospholipids have been studied extensively over the last thirty years as artificial membrane models with remarkable thermostability and tightness against solute leakage. Considerable interest has been generated for applications of liposomes in medicine, including their use as diagnostic agents, as carrier vehicles in vaccine formulations, or as delivery systems for drugs, genes or cancer imaging agents [149]. In general, archaeosomes (liposomes from archaea) demonstrate

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higher stability to oxidative stress, high temperature, alkaline pH, to attack of phospholipases, bile salts, and serum proteins. Some archaeosome formulations can be sterilized by autoclaving without problems of fusion or aggregation of the vesicles. The uptake of archaeosomes by phagocytic cells can be up to 50-fold higher than that of conventional liposomes [150]. Crystalline cell surface layers (S-layers) that are composed of protein and glycoprotein subunits are one of the most commonly observed cell envelope structures of bacteria and archaea. S-layers could be produced in large amounts by continuous cultivation of S-layer-carrying microorganisms and used as isoporous ultrafiltration membranes or as matrices for immobilization of biologically active macromolecules such as enzymes, ligands or mono- and polyclonal antibodies [151]. S-layers have been shown to be excellent patterning structures in molecular nanotechnology due to their high molecular order, high binding capacity and ability to recrystallize with perfect uniformity on solid surfaces, at the water/air interface or on lipid films. The two-dimensionally organized S-layers of S. acidocaldarius are suggested to be of practical use as biomimetic templates for material deposition and fabrication of advanced materials [151]. The production of antibiotic peptides and proteins is a near-universal feature of living organisms regardless of phylogenetic classification. Antimicrobial agents from bacteria and eucarya have been studied for more than 50 years. However, thermophilic archaea and bacteria are just in the beginning of investigation for the production of peptide antibiotics. A variety of halocins have been detected in halophilic archaea. These antimicrobial agents are diverse in size, consisting of proteins as large as 35 kDa and peptide “microhalocins” as small as 3.6 kDa [152]. Microhalocins with unclear mechanism of action are hydrophobic and robust, withstanding heat, desalting and exposure to neutral residues and are not cationic. The microhalocins S8 and R1 lack the biochemical and structural properties demonstrated by other antibiotics, suggesting that their mechanisms of action should be novel. The halocin H7 has been suggested for reducing injury during organ transplantation. Archaeocins are also produced by a thermoacidophilic Sulfolobus strain. The 20 kDa protein antibiotics are not excreted and are associated with small particles apparently derived from the cells S-layer [152].

HYDROGEN PRODUCTION There is an increasing interest in the utilization of renewable sources to satisfy the exponentially growing energy needs. Products of anaerobic fermentation include ethanol, methane, and hydrogen. Research on biological hydrogen production became attractive due to the possible use of biohydrogen as a clean energy carrier and raw material. The production of hydrogen in photobiological or heterotrophic fermentation routes depends on supply of organic substrates and could be therefore ideally suited for coupling energy production with treatment of organic wastes. A two-stage fermentation system was constructed for the production of biohydrogen from keratin-rich biowaste [153]. First, the bacterial strain B. licheniformis KK1 was employed to convert keratin-containing waste into a fermentation product that is rich in amino acids and peptides. In the next stage the thermophilic anaerobic archaeon T. litoralis was fermentated on the hydrolysate and hydrogen was produced. Also archaeal hydrogenases have been the target of intensive research. A cytosolic NiFe-hydrogenase from the hyperthermophilic archaeon T. kodakaraensis is optimally active at 90°C for hydrogen production with methyl viologen as the electron carrier [154]. A membrane bound NiFe-hydrogenase, responsible for hydrogen production, was also identified in the anaerobic bacterium T. tengcongensis [155,156]. Other thermophilic bacteria of the order Thermotogales have also demonstrated the ability to produce hydrogen [157–159].

OUTLOOK Owing to their properties such as activity over a wide temperature and pH range, substrate specificity, stability in organic solvents, diverse substrate range and enantioselectivity, extremophiles and their

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enzymes will represent the choice for future countless applications in industry. The growing demand for more robust biocatalysts has shifted the trend towards improving the properties of existing proteins for established industrial processes and producing new enzymes tailor-made for entirely new areas of application. The new technologies such as directed evolution and gene shuffling will provide valuable tools for improving and adapting enzyme properties to the desired requirements. These new technologies will allow to exploit the potential of extremophiles and elucidate their features in terms of stability, specificity and enzymatic mechanisms. The application of robust enzymes and microorganisms for the sustainable production of chemicals, biopolymers, materials and fuels from renewable resources, also defined as industrial (white) biotechnology, will offer great opportunities for various industries. The utmost aim will be the reduction of waste, energy input and raw material and the development of highly efficient and environmentally friendly processes. As a consequence already existing chemical processes will be optimized and new novel processes will be developed.

REFERENCES 1. Egorova, K. and G. Antranikian. Industrial relevance of thermophilic Archaea. Curr Opin Microbiol 8, 649–55, 2005. 2. Demirjian, D. C., F. Moris-Varas, and C. S. Cassidy. Enzymes from extremophiles. Curr Opin Chem Biol 5, 144–51, 2001. 3. Antranikian, G., C. Vorgias, and C. Bertoldo. Extreme environments as a resource for microorganisms and novel biocatalysts. Adv Biochem Eng Biotechnol 96, 229–62, 2005. 4. Niehaus, F., C. Bertoldo, M. Kahler, and G. Antranikian. Extremophiles as a source of novel enzymes for industrial application. Appl Microbiol Biotechnol 51, 711–29, 1999. 5. Bauer, M. W., L. E. Driskill, W. Callen, M. A. Snead, E. J. Mathur, and R. M. Kelly. An endoglucanase, EglA, from the hyperthermophilic archaeon Pyrococcus furiosus hydrolyzes beta-1,4 bonds in mixedlinkage (1—>3),(1—>4)-beta-D-glucans and cellulose. J Bacteriol 181, 284–90, 1999. 6. Gueguen, Y., W. G. Voorhorst, J. van der Oost, and W. M. de Vos. Molecular and biochemical characterization of an endo-beta-1,3-glucanase of the hyperthermophilic archaeon Pyrococcus furiosus. J Biol Chem 272, 31258–64, 1997. 7. van Lieshout, J., M. Faijes, J. Nieto, J. van der Oost, and A. Planas. Hydrolase and glycosynthase activity of endo-1,3-beta-glucanase from the thermophile Pyrococcus furiosus. Archaea 1, 285–92, 2004. 8. Ando, S., H. Ishida, Y. Kosugi, and K. Ishikawa. Hyperthermostable endoglucanase from Pyrococcus horikoshii. Appl Environ Microbiol 68, 430–3, 2002. 9. Kashima, Y., K. Mori, H. Fukada, and K. Ishikawa. Analysis of the function of a hyperthermophilic endoglucanase from Pyrococcus horikoshii that hydrolyzes crystalline cellulose. Extremophiles 9, 37–43, 2005. 10. Huang, Y., G. Krauss, S. Cottaz, H. Driguez, and G. Lipps. A highly acid-stable and thermostable endo-beta-glucanase from the thermoacidophilic archaeon Sulfolobus solfataricus. Biochem J 385, 581–8, 2005. 11. Liebl, W. Cellulolytic enzymes from Thermotoga species. Methods Enzymol 330, 290–300, 2001. 12. Te’o V. S., D. J. Saul, and P. L. Bergquist. celA, another gene coding for a multidomain cellulase from the extreme thermophile Caldocellum saccharolyticum. Appl Microbiol Biotechnol 43, 291–6, 1995. 13. Zverlov V., S. Mahr, K. Riedel, and K. Bronnenmeier. Properties and gene structure of a bifunctional cellulolytic enzyme (CelA) from the extreme thermophile “Anaerocellum thermophilum” with separate glycosyl hydrolase family 9 and 48 catalytic domains. Microbiology 144, 457–65, 1998. 14. Halldorsdottir S., E. T. Thorolfsdottir, R. Spilliaert, et al. Cloning, sequencing and overexpression of a Rhodothermus marinus gene encoding a thermostable cellulase of glycosyl hydrolase family 12. Appl Microbiol Biotechnol 49, 277–84, 1998. 15. Eckert, K. and E. Schneider. A thermoacidophilic endoglucanase (CelB) from Alicyclobacillus acidocaldarius displays high sequence similarity to arabinofuranosidases belonging to family 51 of glycoside hydrolases. Eur J Biochem 270, 3593–602, 2003. 16. Eckert K., F. Zielinski, L. Lo Leggio, and E. Schneider. Gene cloning, sequencing, and characterization of a family 9 endoglucanase (CelA) with an unusual pattern of activity from the thermoacidophile Alicyclobacillus acidocaldarius ATCC27009. Appl Microbiol Biotechnol 60, 428–36, 2002.

9214_C008.indd 147

11/5/2007 7:39:29 AM

148

Thermophiles: Biology and Technology at High Temperatures

17. Nagatomo, H., Y. Matsushita, K. Sugamoto, and T. Matsui. Preparation and properties of gelatinimmobilized beta-glucosidase from Pyrococcus furiosus. Biosci Biotechnol Biochem 69, 128–36, 2005. 18. Morana, A., M. Moracci, A. Ottombrino, M. Ciaramella, M. Rossi, and M. De Rosa. Industrial-scale production and rapid purification of an archaeal beta-glycosidase expressed in Saccharomyces cerevisiae. Biotechnol Appl Biochem 22, 261–8, 1995. 19. Schiraldi, C., M. Giuliano, and M. De Rosa. Perspectives on biotechnological applications of archaea. Archaea 1, 75–86, 2002. 20. Andrade, C. M., W. B. Aguiar, and G. Antranikian. Physiological aspects involved in production of xylanolytic enzymes by deep-sea hyperthermophilic archaeon Pyrodictium abyssi. Appl Biochem Biotechnol 91–93, 655–69, 2001. 21. Eichler, J. Biotechnological uses of archaeal extremozymes. Biotechnol Adv 19, 261–78, 2001. 22. Uhl, A. M. and R. M. Daniel. The first description of an archaeal hemicellulase: the xylanase from Thermococcus zilligii strain AN1. Extremophiles 3, 263–7, 1999. 23. Cannio, R., N. Di Prizito, M. Rossi, and A. Morana. A xylan-degrading strain of Sulfolobus solfataricus: isolation and characterization of the xylanase activity. Extremophiles 8, 117–24, 2004. 24. Vieille, C. and G. J. Zeikus. Hyperthermophilic enzymes: sources, uses, and molecular mechanisms for thermostability. Microbiol Mol Biol Rev 65, 1–43, 2001. 25. Gupta, R., P. Gigras, H. Mohapatra, V. Goswami, and B. Chauhan. Microbial alpha-amylases: a biotechnological perspective. Process Biochemistry 38, 1599–1616, 2003. 26. Kim, M. S., J. T. Park, Y. W. Kim, H. S. Lee, R. Nyawira, H. S. Shin, C. S. Park, S. H. Yoo, Y. R. Kim, T. W. Moon, and K. H. Park. Properties of a novel thermostable glucoamylase from the hyperthermophilic archaeon Sulfolobus solfataricus in relation to starch processing. Appl Environ Microbiol 70, 3933–40, 2004. 27. Kaper, T., B. Talik, T. J. Ettema, H. Bos, M. J. van der Maarel, and L. Dijkhuizen. Amylomaltase of Pyrobaculum aerophilum IM2 produces thermoreversible starch gels. Appl Environ Microbiol 71, 5098–106, 2005. 28. Matzke, J., B. Schwermann, and E. P. Bakker. Acidostable and acidophilic proteins: the example of the alpha-amylase from Alicyclobacillus acidocaldarius. Comp Biochem Physiol A Physiol, 118, 475–9, 1997. 29. Serour, E. and G. Antranikian. Novel thermoactive glucoamylases from the thermoacidophilic archaea Thermoplasma acidophilum, Picrophilus torridus and Picrophilus oshimae. Antonie Van Leeuwenhoek 81, 73–83, 2002. 30. Uotsu-Tomita, R. T. Tonozuka, H. Sakai, and Y. Sakano. Novel glucoamylase-type enzymes from Thermoactinomyces vulgaris and Methanococcus jannaschii whose genes are found in the flanking region of the alpha-amylase genes. Appl Microbiol Biotechnol 56, 465–73, 2001. 31. Specka, U., F. Mayer, and G. Antranikian. Purification and properties of a thermoactive glucoamylase from Clostridium thermosaccharolyticum. Appl Environ Microbiol 57, 2317–2323, 1991. 32. Ganghofner D., J. Kellermann, W. L. Staudenbauer, and K. Bronnenmeier. Purification and properties of an amylopullulanase, a glucoamylase, and an alpha-glucosidase in the amylolytic enzyme system of Thermoanaerobacterium thermosaccharolyticum. Biosci Biotechnol Biochem 62, 302–8, 1998. 33. Rolfsmeier, M., C. Haseltine, E. Bini, A. Clark, and P. Blum. Molecular characterization of the alphaglucosidase gene (malA) from the hyperthermophilic archaeon Sulfolobus solfataricus. J Bacteriol 180, 1287–95, 1998. 34. Bertoldo, C. and G. Antranikian. Starch-hydrolyzing enzymes from thermophilic archaea and bacteria. Curr Opin Chem Biol 6, 151–60, 2002. 35. Bertoldo, C. and G. Antranikian. Amylolytic enzymes from hyperthermophiles. Methods Enzymol 330, 269–89, 2001. 36. Raasch C., W. Streit, J. Schanzer, M. Bibel, U. Gosslar, and W. Liebl. Thermotoga maritima AglA, an extremely thermostable NAD+-, Mn2+-, and thiol-dependent alpha-glucosidase. Extremophiles 4, 189–200, 2000. 37. Ferrer, M., O. V. Golyshina, F. J. Plou, K. N. Timmis, and P. N. Golyshin. A novel alpha-glucosidase from the acidophilic archaeon, Ferroplasma acidiphilum Y with high transglycosylation activity and an unusual catalytic nucleophile. Biochem J 391(2), 269–276, 2005. 38. Brown, S. and R. Kelly. Characterization of amylolytic enzymes, having both alpha-1,4 and alpha-1,6 hydrolytic activity, from the thermophilic archaea Pyrococcus furiosus and Thermococcus litoralis. Appl Environ Microbiol 59, 2614–21, 1993.

9214_C008.indd 148

11/5/2007 7:39:29 AM

Industrial Relevance of Thermophiles and Their Enzymes

149

39. Zona, R., F. Chang-Pi-Hin, M. J. O’Donohue, and S. Janecek. Bioinformatics of the glycoside hydrolase family 57 and identification of catalytic residues in amylopullulanase from Thermococcus hydrothermalis. Eur J Biochem 271, 2863–72, 2004. 40. Kang, S., C. Vieille, and J. G. Zeikus. Identification of Pyrococcus furiosus amylopullulanase catalytic residues. Appl Microbiol Biotechnol 2004. 41. Bertoldo C., M. Armbrecht, F. Becker, T. Schafer, G. Antranikian, and W. Liebl. Cloning, sequencing, and characterization of a heat- and alkali-stable type I pullulanase from Anaerobranca gottschalkii. Appl Environ Microbiol 70, 3407–16, 2004. 42. Tachibana, Y., A. Kuramura, N. Shirasaka, Y. Suzuki, T. Yamamoto, S. Fujiwara, M. Takagi, and T. Imanaka. Purification and characterization of an extremely thermostable cyclomaltodextrin glucanotransferase from a newly isolated hyperthermophilic archaeon, a Thermococcus sp. Appl Environ Microbiol 65, 1991–7, 1999. 43. Rashid, N., J. Cornista, S. Ezaki, T. Fukui, H. Atomi, and T. Imanaka. Characterization of an archaeal cyclodextrin glucanotransferase with a novel C-terminal domain. J Bacteriol 184, 777–84, 2002. 44. Horikoshi, K. Alkaliphiles: some applications of their products for biotechnology. Microbiol Mol Biol Rev 63, 735–50, 1999. 45. Thiemann, V., C. Donges, S. G. Prowe, R. Sterner, and G. Antranikian. Characterisation of a thermoalkali-stable cyclodextrin glycosyltransferase from the anaerobic thermoalkaliphilic bacterium Anaerobranca gottschalkii. Arch Microbiol 182, 226–35, 2004. 46. Thiemann, V., B. Saake, A. Vollstedt, T. Schafer, J. Puls, C. Bertoldo, R. Freudl, and G. Antranikian. Heterologous expression and characterization of a novel branching enzyme from the thermoalkaliphilic anaerobic bacterium Anaerobranca gottschalkii. Appl Microbiol Biotechnol 12, 1–12, 2006. 47. Murakami, T., T. Kanai, H. Takata, T. Kuriki, and T. Imanaka. A novel branching enzyme of the GH-57 family in the hyperthermophilic archaeon Thermococcus kodakaraensis KOD1. J Bacteriol 188(16), 5915–24, 2006. 48. Takata, H., T. Takaha, S. Okada, S. Hizukuri, M. Takagi, and T. Imanaka. Structure of the cyclic glucan produced from amylopectin by Bacillus stearothermophilus branching enzyme. Carbohydr Res 295, 91–101, 1996. 49. Shinohara M. L., M. Ihara, M. Abo, M. Hashida, S. Takagi, and T. C. Beck. A novel thermostable branching enzyme from an extremely thermophilic bacterial species, Rhodothermus obamensis. Appl Microbiol Biotechnol 57, 653–9, 2001. 50. Imamura, H., B. Jeon, T. Wakagi, and H. Matsuzawa. High level expression of Thermococcus litoralis 4-alpha-glucanotransferase in a soluble form in Escherichia coli with a novel expression system involving minor arginine tRNAs and GroELS. FEBS Lett 457, 393–6, 1999. 51. Imamura, H., S. Fushinobu, B. S. Jeon, T. Wakagi, and H. Matsuzawa. Identification of the catalytic residue of Thermococcus litoralis 4-alpha-glucanotransferase through mechanism-based labeling. Biochemistry 40, 12400–6, 2001. 52. Jeon, B. S., H. Taguchi, H. Sakai, T. Ohshima, T. Wakagi, and H. Matsuzawa. 4-alpha-glucanotransferase from the hyperthermophilic archaeon Thermococcus litoralis--enzyme purification and characterization, and gene cloning, sequencing and expression in Escherichia coli. Eur J Biochem 248, 171–8, 1997. 53. Kaper, T., M. J. van der Maarel, G. J. Euverink, and L. Dijkhuizen. Exploring and exploiting starchmodifying amylomaltases from thermophiles. Biochem Soc Trans 32, 279–82, 2004. 54. Terada, Y., K. Fujii, T. Takaha, and S. Okada. Thermus aquaticus ATCC 33923 amylomaltase gene cloning and expression and enzyme characterization: production of cycloamylose. Appl Environ Microbiol 65, 910–5, 1999. 55. Fuji, K., H. Minagawa, Y. Terada, T. Takaha, T. Kuriki, J. Shimada, and H. Kaneko. Use of random and saturation mutageneses to improve the properties of Thermus aquaticus Amylomaltase for efficient production of cycloamyloses. Appl Environ Microbiol 71(10), 5823–27, 2005. 56. Biwer, A., G. Antranikian, and E. Heinzle. Enzymatic production of cyclodextrins. Appl Microbiol Biotechnol 59, 609–17, 2002. 57. de Pascale, D., I. Di Lernia, M. P. Sasso, A. Furia, M. De Rosa, and M. Rossi. A novel thermophilic fusion enzyme for trehalose production. Extremophiles 6, 463–8, 2002. 58. Fang, T. Y., X. G. Hung, T. Y. Shih, and W. C. Tseng. Characterization of the trehalosyl dextrin-forming enzyme from the thermophilic archaeon Sulfolobus solfataricus ATCC 35092. Extremophiles 8, 335–43, 2004.

9214_C008.indd 149

11/5/2007 7:39:29 AM

150

Thermophiles: Biology and Technology at High Temperatures

59. Gueguen, Y., J. L. Rolland, S. Schroeck, D. Flament, S. Defretin, M. H. Saniez, and J. Dietrich. Characterization of the maltooligosyl trehalose synthase from the thermophilic archaeon Sulfolobus acidocaldarius. FEMS Microbiol Lett 194, 201–6, 2001. 60. Kozianowski, G., F. Canganella, F. A. Rainey, H. Hippe, and G. Antranikian. Purification and characterization of thermostable pectate-lyases from a newly isolated thermophilic bacterium, Thermoanaerobacter italicus sp. nov. Extremophiles 1, 171–82, 1997. 61. Kluskens, L. D., G. J. van Alebeek, A. G. Voragen, W. M. de Vos, and J. van der Oost. Molecular and biochemical characterization of the thermoactive family 1 pectate lyase from the hyperthermophilic bacterium Thermotoga maritima. Biochem J 370, 651–9, 2003. 62. Kluskens, L. D., G. J. van Alebeek, J. Walther, A. G. Voragen, W. M. de Vos, and J. van der Oost. Characterization and mode of action of an exopolygalacturonase from the hyperthermophilic bacterium Thermotoga maritima. FEBS J 272, 5464–73, 2005. 63. Andronopoulou, E. and C. E. Vorgias. Multiple components and induction mechanism of the chitinolytic system of the hyperthermophilic archaeon Thermococcus chitonophagus. Appl Microbiol Biotechnol 65, 694–702, 2004. 64. Andronopoulou, E. and C. E. Vorgias. Isolation, cloning, and overexpression of a chitinase gene fragment from the hyperthermophilic archaeon Thermococcus chitonophagus: semi-denaturing purification of the recombinant peptide and investigation of its relation with other chitinases. Protein Expr Purif 35, 264–71, 2004. 65. Tanaka, T., T. Fukui, S. Fujiwara, H. Atomi, and T. Imanaka. Concerted action of diacetylchitobiose deacetylase and exo-beta-D-glucosaminidase in a novel chitinolytic pathway in the hyperthermophilic archaeon Thermococcus kodakaraensis KOD1. J Biol Chem 279, 30021–7, 2004. 66. Tanaka, T., S. Fujiwara, S. Nishikori, T. Fukui, M. Takagi, and T. Imanaka. A unique chitinase with dual active sites and triple substrate binding sites from the hyperthermophilic archaeon Pyrococcus kodakaraensis KOD1. Appl Environ Microbiol 65, 5338–44, 1999. 67. Gao, J., M. W. Bauer, K. R. Shockley, M. A. Pysz, and R. M. Kelly. Growth of hyperthermophilic archaeon Pyrococcus furiosus on chitin involves two family 18 chitinases. Appl Environ Microbiol 69, 3119–28, 2003. 68. Andronopoulou, E. and C. E. Vorgias. Purification and characterization of a new hyperthermostable, allosamidin-insensitive and denaturation-resistant chitinase from the hyperthermophilic archaeon Thermococcus chitonophagus. Extremophiles 7, 43–53, 2003. 69. Tanaka, T., T. Fukui, and T. Imanaka. Different cleavage specificities of the dual catalytic domains in chitinase from the hyperthermophilic archaeon Thermococcus kodakaraensis KOD1. J Biol Chem 276, 35629–35, 2001. 70. Bertoldo, C., R. Grote, and G. Antranikian. Biocatalysis under extreme conditions. Biotechnology 10, 61–103, 2001. 71. Mayr, J., A. Lupas, J. Kellermann, C. Eckerskorn, W. Baumeister, and J. Peters. A hyperthermostable protease of the subtilisin family bound to the surface layer of the archaeon Staphylothermus marinus. Curr Biol 6, 739–49, 1996. 72. Chavez Croocker, P. Y. Sako, and A. Uchida. Purification and characterization of an intracellular heatstable proteinase (pernilase) from the marine hyperthermophilic archaeon Aeropyrum pernix K1. Extremophiles 3, 3–9, 1999. 73. Sako, Y., P. C. Croocker, and Y. Ishida. An extremely heat-stable extracellular proteinase (aeropyrolysin) from the hyperthermophilic archaeon Aeropyrum pernix K1. FEBS Lett 415, 329–34, 1997. 74. Ward, D. E., K. R. Shockley, L. S. Chang, R. D. Levy, J. K. Michel, S. B. Conners, and R. M. Kelly. Proteolysis in hyperthermophilic microorganisms. Archaea 1, 63–74, 2002. 75. Voorhorst, W. G., R. I. Eggen, A. C. Geerling, C. Platteeuw, R. J. Siezen, and W. M. Vos. Isolation and characterization of the hyperthermostable serine protease, pyrolysin, and its gene from the hyperthermophilic archaeon Pyrococcus furiosus. J Biol Chem 271, 20426–31, 1996. 76. Burlini, N., P. Magnani, A. Villa, F. Macchi, P. Tortora, and A. Guerritore. A heat-stable serine proteinase from the extreme thermophilic archaebacterium Sulfolobus solfataricus. Biochim Biophys Acta 1122, 283–92, 1992. 77. Kluskens, L. D., W. G. Voorhorst, R. J. Siezen, R. M. Schwerdtfeger, G. Antranikian, J. van der Oost, and W. M. de Vos. Molecular characterization of fervidolysin, a subtilisin-like serine protease from the thermophilic bacterium Fervidobacterium pennivorans. Extremophiles 6, 185–94, 2002.

9214_C008.indd 150

11/5/2007 7:39:29 AM

Industrial Relevance of Thermophiles and Their Enzymes

151

78. Tsuruoka, N., T. Nakayama, M. Ashida, H. Hemmi, M. Nakao, H. Minakata, H. Oyama, K. Oda, and T. Nishino. Collagenolytic serine-carboxyl proteinase from Alicyclobacillus sendaiensis strain NTAP-1: purification, characterization, gene cloning, and heterologous expression. Appl Environ Microbiol 69, 162–9, 2003. 79. Khan, A. R., S. Nirasawa, S. Kaneko, T. Shimonishi, and K. Hayashi. Characterization of a solvent resistant and thermostable aminopeptidase from the hyperthermophillic bacterium, Aquifex aeolicus. Enzyme Microb Technol 27, 83–88, 2000. 80. Cheng, T. C., V. Ramakrishnan, and S. I. Chan. Purification and characterization of a cobalt-activated carboxypeptidase from the hyperthermophilic archaeon Pyrococcus furiosus. Protein Sci 8, 2474–8 1999. 81. Suzuki, Y., K. Miyamoto, and H. Ohta. A novel thermostable esterase from the thermoacidophilic archaeon Sulfolobus tokodaii strain 7. FEMS Microbiol Lett 236, 97–102, 2004. 82. Ikeda, M. and D. S. Clark. Molecular cloning of extremely thermostable esterase gene from hyperthermophilic archaeon Pyrococcus furiosus in Escherichia coli. Biotechnol Bioeng 57, 624–9, 1998. 83. Hotta, Y., S. Ezaki, H. Atomi, and T. Imanaka. Extremely stable and versatile carboxylesterase from a hyperthermophilic archaeon. Appl Environ Microbiol 68, 3925–31, 2002. 84. Wang, B., D. Lu, R. Gao, Z. Yang, S. Cao, and Y. Feng. A novel phospholipase A2/esterase from hyperthermophilic archaeon Aeropyrum pernix K1. Protein Expr Purif 35, 199–205, 2004. 85. Manco, G., G. Carrea, E. Giosue, G. Ottolina, G. Adamo, and M. Rossi. Modification of the enantioselectivity of two homologous thermophilic carboxylesterases from Alicyclobacillus acidocaldarius and Archaeoglobus fulgidus by random mutagenesis and screening. Extremophiles 6, 325–31, 2002. 86. Sehgal, A. C. and R. M. Kelly. Strategic selection of hyperthermophilic esterases for resolution of 2-arylpropionic esters. Biotechnol Prog 19, 1410–6, 2003. 87. Sehgal, A. C. and R. M. Kelly. Enantiomeric resolution of 2-aryl propionic esters with hyperthermophilic and mesophilic esterases: contrasting thermodynamic mechanisms. J Am Chem Soc 124, 8190–1, 2002. 88. Sehgal, A. C., R. Tompson, J. Cavanagh, and R. M. Kelly. Structural and catalytic response to temperature and cosolvents of carboxylesterase EST1 from the extremely thermoacidophilic archaeon Sulfolobus solfataricus P1. Biotechnol Bioeng 80, 784–93, 2002. 89. Kim, S. and S. B. Lee. Thermostable esterase from a thermoacidophilic archaeon: purification and characterization for enzymatic resolution of a chiral compound. Biosci Biotechnol Biochem 68, 2289– 98, 2004. 90. Zhang, J., J. Liu, J. Zhou, Y. Ren, X. Dai, and H. Xiang. Thermostable esterase from Thermoanaerobacter tengcongensis: high-level expression, purification and characterization. Biotechnol Lett 25, 1463–7, 2003. 91. Radianingtyas, H. and P. C. Wright. Alcohol dehydrogenases from thermophilic and hyperthermophilic archaea and bacteria. FEMS Microbiol Rev 27, 593–616, 2003. 92. Ma, K., F. T. Robb, and M. W. Adams. Purification and characterization of NADP-specific alcohol dehydrogenase and glutamate dehydrogenase from the hyperthermophilic archaeon Thermococcus litoralis. Appl Environ Microbiol 60(2), 562–68, 1994. 93. van der Oost, J., W. G. Voorhorst, S. W. Kengen, A. C. Geerling, V. Wittenhorst, Y. Gueguen, and W. M. de Vos. Genetic and biochemical characterization of a short-chain alcohol dehydrogenase from the hyperthermophilic archaeon Pyrococcus furiosus. Eur J Biochem 268, 3062–8, 2001. 94. Antoine, E., J.-L. Rolland, J.-P. Raffin, and J. Dietrich. Cloning and over-expression in Escherichia coli of the gene encoding NADPH group III alcohol dehydrogenase from Thermococcus hydrothermalis. Eur J Biochem 264, 880–89, 1999. 95. Yoon, S. Y., H. S. Noh, E. H. Kim, and K. H. Kong. The highly stable alcohol dehydrogenase of Thermomicrobium roseum: purification and molecular characterization. Comp Biochem Physiol B Biochem Mol Biol 132, 415–22, 2002. 96. Crabb, W. D. and J. Bolin. Protein technologies and commercial enzymes. Curr Opin Biotechnol 10, 321–3, 1999. 97. Park, B. C., S. Koh, C. Chang, S. W. Suh, D. S. Lee, and S. M. Byun. Cloning and expression of the gene for xylose isomerase from Thermus flavus AT62 in Escherichia coli. Appl Biochem Biotechnol 62, 15–27, 1997. 98. Vieille, C., J. M. Hess, R. M. Kelly, and J. G. Zeikus. xylA cloning and sequencing and biochemical characterization of xylose isomerase from Thermotoga neapolitana. Appl Environ Microbiol 61, 1867–75, 1995. 99. Lee, S. J., D. W. Lee, E. A. Choe, Y. H. Hong, S. B. Kim, B. C. Kim, and Y. R. Pyun. Characterization of a thermoacidophilic L-arabinose isomerase from Alicyclobacillus acidocaldarius: role of Lys-269 in pH optimum. Appl Environ Microbiol 71, 7888–96, 2005.

9214_C008.indd 151

11/5/2007 7:39:29 AM

152

Thermophiles: Biology and Technology at High Temperatures

100. Fessner, W. D. and V. Helaine. Biocatalytic synthesis of hydroxylated natural products using aldolases and related enzymes. Curr Opin Biotechnol 12, 574–86, 2001. 101. Lamble, H. J., M. J. Danson, D. W. Hough, and S. D. Bull. Engineering stereocontrol into an aldolasecatalysed reaction. Chem Commun (Camb) 124–6, 2005. 102. Soderberg, T. and R. Alver. Transaldolase of Methanocaldococcus jannaschii. Archaea 1, 255–62, 2004. 103. Egorova, K., H. Trauthwein, S. Verseck, and G. Antranikian. Purification and properties of an enantioselective and thermoactive amidase from the thermophilic actinomycete Pseudonocardia thermophila. Appl Microbiol Biotechnol 65, 38–45, 2004. 104. d’Abusco, A. S., S. Ammendola, R. Scandurra, and L. Politi. Molecular and biochemical characterization of the recombinant amidase from hyperthermophilic archaeon Sulfolobus solfataricus. Extremophiles 5, 183–92, 2001. 105. Mueller, P., K. Egorova, C. E. Vorgias, E. Boutou, H. Trauthwein, S. Verseck, and G. Antranikian. Cloning, overexpression, and characterization of a thermoactive nitrilase from the hyperthermophilic archaeon Pyrococcus abyssi. Protein Expr Purif 47(2),672–81, 2006. 106. Perler, F. B., S. Kumar, and H. Kong. Thermostable DNA polymerases. Adv Protein Chem 48, 377–435, 1996. 107. Takagi, M., M. Nishioka, H. Kakihara, M. Kitabayashi, H. Inoue, B. Kawakami, M. Oka, and T. Imanaka. Characterization of DNA polymerase from Pyrococcus sp. strain KOD1 and its application to PCR. Appl Environ Microbiol 63, 4504–10, 1997. 108. Hashimoto, H., M. Nishioka, S. Fujiwara, M. Takagi, T. Imanaka, T. Inoue, and Y. Kai. Crystal structure of DNA polymerase from hyperthermophilic archaeon Pyrococcus kodakaraensis KOD1. J Mol Biol 306, 469–77, 2001. 109. Dabrowski, S. and B. Kiaer Ahring. Cloning, expression, and purification of the His6-tagged hyperthermostable dUTPase from Pyrococcus woesei in Escherichia coli: application in PCR. Protein Expr Purif 31, 72–8, 2003. 110. Biles, B. D. and B. A. Connolly. Low-fidelity Pyrococcus furiosus DNA polymerase mutants useful in error-prone PCR. Nucleic Acids Res 32, e176, 2004. 111. Kowalczykowski, S., F. Chedin, and E. Seitz. Single stranded DNA binding proteins from archaea. USA patent 6,852,832, 2005. 112. Sanger, F., S. Nicklen, and A. R. Coulson. DNA sequencing with chain-terminating inhibitors. 1977. Biotechnology 24, 104–8, 1992. 113. Bechtereva, T. A., Y. I. Pavlov, V. I. Kramorov, B. Migunova, and O. I. Kiselev. DNA sequencing with thermostable Tet DNA polymerase from Thermus thermophilus. Nucleic Acids Res 17, 10507, 1989. 114. Rao, V. B. and N. B. Saunders. A rapid polymerase-chain-reaction-directed sequencing strategy using a thermostable DNA polymerase from Thermus flavus. Gene 113, 17–23, 1992. 115. Vander Horn, P. B., M. C. Davis, J. J. Cunniff, C. Ruan, B. F. McArdle, S. B. Samols, J. Szasz, G. Hu, K. M. Hujer, S. T. Domke, S. R. Brummet, R. B. Moffett, and C. W. Fuller. Thermo Sequenase DNA polymerase and T. acidophilum pyrophosphatase: new thermostable enzymes for DNA sequencing. Biotechniques 22, 758–62, 764–5, 1997. 116. Wakagi, T., C. H. Lee, and T. Oshima. An extremely stable inorganic pyrophosphatase purified from the cytosol of a thermoacidophilic archaebacterium, Sulfolobus acidocaldarius strain 7. Biochim Biophys Acta 1120, 289–96, 1992. 117. Zappa, S., J. L. Rolland, D. Flament, Y. Gueguen, J. Boudrant, and J. Dietrich. Characterization of a highly thermostable alkaline phosphatase from the euryarchaeon Pyrococcus abyssi. Appl Environ Microbiol 67, 4504–11, 2001. 118. Landegren, U., E. Schallmeiner, M. Nilsson, S. Fredriksson, J. Baner, M. Gullberg, J. Jarvius, S. Gustafsdottir, F. Dahl, O. Soderberg, O. Ericsson, and J. Stenberg. Molecular tools for a molecular medicine: analyzing genes, transcripts and proteins using padlock and proximity probes. J Mol Recognit 17, 194–7, 2004. 119. Kletzin, A. Molecular characterisation of a DNA ligase gene of the extremely thermophilic archaeon Desulfurolobus ambivalens shows close phylogenetic relationship to eukaryotic ligases. Nucleic Acids Res 20, 5389–96, 1992. 120. Nakatani, M., S. Ezaki, H. Atomi, and T. Imanaka. A DNA ligase from a hyperthermophilic archaeon with unique cofactor specificity. J Bacteriol 182, 6424–33, 2000. 121. Nakatani, M., S. Ezaki, H. Atomi, and T. Imanaka. Substrate recognition and fidelity of strand joining by an archaeal DNA ligase. Eur J Biochem 269, 650–6, 2002.

9214_C008.indd 152

11/5/2007 7:39:29 AM

Industrial Relevance of Thermophiles and Their Enzymes

153

122. Rolland, J. L., Y. Gueguen, C. Persillon, J. M. Masson, and J. Dietrich. Characterization of a thermophilic DNA ligase from the archaeon Thermococcus fumicolans. FEMS Microbiol Lett 236, 267–73, 2004. 123. Lai, X., H. Shao, F. Hao, and L. Huang. Biochemical characterization of an ATP-dependent DNA ligase from the hyperthermophilic crenarchaeon Sulfolobus shibatae. Extremophiles 6, 469–77, 2002. 124. Lim, J. H., J. Choi, S. J. Han, S. H. Kim, H. Z. Hwang, D. K. Jin, B. Y. Ahn, and Y. S. Han. Molecular cloning and characterization of thermostable DNA ligase from Aquifex pyrophilus, a hyperthermophilic bacterium. Extremophiles 5, 161–8, 2001. 125. Thorbjarnardottir, S. H., Z. O. Jonsson, O. S. Andresson, J. K. Kristjansson, G. Eggertsson, and A. Palsdottir. Cloning and sequence analysis of the DNA ligase-encoding gene of Rhodothermus marinus, and overproduction, purification and characterization of two thermophilic DNA ligases. Gene 161, 1–6, 1995. 126. Jolivet, E., E. Corre, S. L’Haridon, P. Forterre, and D. Prieur. Thermococcus marinus sp. nov. and Thermococcus radiotolerans sp. nov., two hyperthermophilic archaea from deep-sea hydrothermal vents that resist ionizing radiation. Extremophiles 8, 219–27, 2004. 127. Matsui, E., S. Kawasaki, H. Ishida, K. Ishikawa, Y. Kosugi, H. Kikuchi, Y. Kawarabayashi, and I. Matsui. Thermostable flap endonuclease from the archaeon, Pyrococcus horikoshii, cleaves the replication fork-like structure endo/exonucleolytically. J Biol Chem 274, 18297–309, 1999. 128. Leclere, M. M., M. Nishioka, T. Yuasa, S. Fujiwara, M. Takagi, and T. Imanaka. The O6-methylguanine-DNA methyltransferase from the hyperthermophilic archaeon Pyrococcus sp. KOD1: a thermostable repair enzyme. Mol Gen Genet 258, 69–77, 1998. 129. Slesarev, A. I., K. V. Mezhevaya, K. S. Makarova, N. N. Polushin, O. V. Shcherbinina, V. V. Shakhova, G. I. Belova, L. Aravind, D. A. Natale, I. B. Rogozin, R. L. Tatusov, Y. I. Wolf, K. O. Stetter, A. G. Malykh, E. V. Koonin, and S. A. Kozyavkin. The complete genome of hyperthermophile Methanopyrus kandleri AV19 and monophyly of archaeal methanogens. Proc Natl Acad Sci U S A 99, 4644–9, 2002. 130. Borges, N., A. Ramos, N. D. Raven, R. J. Sharp, and H. Santos. Comparative study of the thermostabilizing properties of mannosylglycerate and other compatible solutes on model enzymes. Extremophiles 6, 209–16, 2002. 131. Barth, S., M. huhn, B. Matthey, A. Klimka, E. Galinski, and A. Engert. Compatible-solute-supported periplasmic expression of functional recombinant proteins under stress conditions. Appl Environ Microbiol 66, 1572–1579, 2000. 132. Santos, H. and M. S. da Costa. Compatible solutes of organisms that live in hot saline environments. Environ Microbiol 4, 501–9, 2002. 133. Empadinhas, N., J. D. Marugg, N. Borges, H. Santos, and M. S. da Costa. Pathway for the synthesis of mannosylglycerate in the hyperthermophilic archaeon Pyrococcus horikoshii. Biochemical and genetic characterization of key enzymes. J Biol Chem 276, 43580–8, 2001. 134. Chen, L., E. T. Spiliotis, and M. F. Roberts. Biosynthesis of Di-myo-inositol-1,1’-phosphate, a novel osmolyte in hyperthermophilic archaea. J Bacteriol 180, 3785–92, 1998. 135. Scholz, S., S. Wolff, and R. Hensel. The biosynthesis pathway of di-myo-inositol-1,1’-phosphate in Pyrococcus woesei. FEMS Microbiol Lett 168, 37–42, 1998. 136. Lamosa, P., A. Burke, R. Peist, R. Huber, M. Y. Liu, G. Silva, C. Rodrigues-Pousada, J. LeGall, C. Maycock, and H. Santos. Thermostabilization of proteins by diglycerol phosphate, a new compatible solute from the hyperthermophile Archaeoglobus fulgidus. Appl Environ Microbiol 66, 1974–9, 2000. 137. Santos, H. and M. Da Costa. Organic solutes from thermophiles and hyperthermophiles. Methods Enzymol 334, 302–315, 2001. 138. Story, S. V., A. M. Grunden, and M. W. W. Adams. Characterization of an aminoacylase from the hyperthermophilic archaeon Pyrococcus furiosus. J Bacteriol 183, 4259–4268, 2001. 139. Toogood, H. S., E. J. Hollingsworth, R. C. Brown, I. N. Taylor, S. J. Taylor, R. McCague, and J. A. Littlechild. A thermostable L-aminoacylase from Thermococcus litoralis: cloning, overexpression, characterization, and applications in biotransformations. Extremophiles 6, 111–22, 2002. 140. Taylor, I., R. Brown, M. Bycroft, G. King, J. A. Littlechild, M. Lloyd, C. Praguin, and S. Taylor. Application of thermophilic enzymes in commercial biotransformation processes. Biochem Soc Trans 32, 290–292, 2004. 141. Ishikawa, K., H. Ishida, I. Matsui, Y. Kawarabayasi, and H. Kikuchi. Novel bifunctional hyperthermostable carboxypeptidase/aminoacylase from Pyrococcus horikoshii OT3. Appl Environ Microbiol 67, 673–9, 2001.

9214_C008.indd 153

11/5/2007 7:39:30 AM

154

Thermophiles: Biology and Technology at High Temperatures

142. Dabrowski, S., G. Sobiewska, J. Maciunska, J. Synowiecki, and J. Kur. Cloning, expression, and purification of the His-tagged thermostable beta-Galactosidase from Pyrococcus woesei in Escherichia coli and some properties of the isolated enzyme. Prot Exp Pur 19, 107–112, 2000. 143. Hemmi, H., S. Ikejiri, T. Nakayama, and T. Nishino. Fusion-type lycopene beta-cyclase from a thermoacidophilic archaeon Sulfolobus solfataricus. Biochem Biophys Res Commun 305, 586–91, 2003. 144. Ishikawa, K. and K. Mino. Heat resistant cysteine synthase. Japan patent 20040002075. January 1, 2004. 145. Matsui, I., K. Ishikawa, H. Ishida, Y. Kosugi, and Y. Tahara. Thermostable enzyme having phosphatidylethanolamine N-methyltransferase activity. Japan patent 6, 391, 604, 2002. 146. Lloyd, J., D. R. Loveley, and E. L. Macaskie. Biotechnological application of metal-reducing microorganisms. Adv Appl Microbio 53, 85–128, 2003. 147. Norris, P., N. Burton, and N. Foulis. Acidophiles in bioreactor mineral processing. Extremophiles 4, 71–76, 2000. 148. Bredberg, K., J. Persson, M. Christiansson, B. Stenberg, and O. Holst. Anaerobic desulfurization of ground rubber with the thermophilic archaeon Pyrococcus furiosus—a new method for rubber recycling. Appl Microbiol Biotechnol 55, 43–48, 2001. 149. Sprott, G. D., G. B. Patel, and L. Krishnan. Archaeobacterial ether lipid liposomes as vaccine adjuvants. Methods Enzymol 373, 155–72, 2003. 150. Patel, G. B. and G. D. Sprott. Archaeobacterial ether lipid liposomes (archaeosomes) as novel vaccine and drug delivery systems. Crit Rev Biotechnol 19, 317–57, 1999. 151. Sleytr, U. B., H. Bayley, M. Sara, A. Breitwieser, S. Kupcu, C. Mader, S. Weigert, F. M. Unger, P. Messner, B. Jahn-Schmid, B. Schuster, D. Pum, K. Douglas, N. A. Clark, J. T. Moore, T. A. Winningham, S. Levy, I. Frithsen, J. Pankovc, P. Beale, H. P. Gillis, D. A. Choutov, and K. P. Martin. Applications of S-layers. FEMS Microbiol Rev 20, 151–75, 1997. 152. O’Connor, E. and R. Shand. Halocins and sulfolobicins: The emerging story of archaeal protein and peptide antibiotics. J Indust Microbiol Biotechnol 28, 23–31, 2002. 153. Balint, B., Z. Bagi, A. Toth, G. Rakhely, K. Perei, and K. L. Kovacs. Utilization of keratin-containing biowaste to produce biohydrogen. Appl Microbiol Biotechnol 2005. 154. Kanai, T., S. Ito, and T. Imanaka. Characterization of a cytosolic NiFe-hydrogenase from the hyperthermophilic archaeon Thermococcus kodakaraensis KOD1. J Bacteriol 185, 1705–11, 2003. 155. Soboh, B., D. Linder, and R. Hedderich. A multisubunit membrane-bound [NiFe] hydrogenase and an NADH-dependent Fe-only hydrogenase in the fermenting bacterium Thermoanaerobacter tengcongensis. Microbiology 150, 2451–63, 2004. 156. Soboh, B., D. Linder, and R. Hedderich. Purification and catalytic properties of a CO-oxidizing:H2evolving enzyme complex from Carboxydothermus hydrogenoformans. Eur J Biochem 269, 5712–21, 2002. 157. Van Ooteghem, S. A., S. K. Beer, and P. C. Yue. Hydrogen production by the thermophilic bacterium Thermotoga neapolitana. Appl Biochem Biotechnol 98–100, 177–89, 2002. 158. Van Ooteghem, S. A., A. Jones, D. Van Der Lelie, B. Dong, and D. Mahajan. H(2) production and carbon utilization by Thermotoga neapolitana under anaerobic and microaerobic growth conditions. Biotechnol Lett 26, 1223–32, 2004. 159. Kadar, Z., T. De Vrije, M. A. Budde, Z. Szengyel, K. Reczey, and P. A. Claassen. Hydrogen production from paper sludge hydrolysate. Appl Biochem Biotechnol 105–108, 557–66, 2003. 160. Jin, R., S. Richter, R. Zhong, and G. K. Lamppa. Expression and import of an active cellulase from a thermophilic bacterium into the chloroplast both in vitro and in vivo. Plant Mol Biol 51(4), 493–507, 2003. 161. Kim, J. O., S. R. Park, W. J. Lim, S. K. Ryu, M. K. Kim, C. L. An, S. J. Cho, Y. W. Park, J. H. Kim, and H. D. Yun, Cloning and characterization of thermostable endoglucanase (Cel8Y) from the hyperthermophilic Aquifex aeolicus VF5. Biochem Biophys Res Commun 279, 420–6, 2000. 162. Han, S. O., H. Yukawa, M. Inui, and R. H. Doi. Molecular cloning and transcriptional and expression analysis of engO, encoding a new noncellulosomal family 9 enzyme, from Clostridium cellulovorans. J Bacteriol 187, 4884–9, 2005. 163. Zverlov, V. V., J. Kellermann, and W. H. Schwarz. Functional subgenomics of Clostridium thermocellum cellulosomal genes: identification of the major catalytic components in the extracellular complex and detection of three new enzymes. Proteomics 5, 3646–53, 2005. 164. Limauro, D., R. Cannio, G. Fiorentino, M. Rossi, and S. Bartolucci. Identification and molecular characterization of an endoglucanase gene, celS, from the extremely thermophilic archaeon Sulfolobus solfataricus. Extremophiles 5, 213–9, 2001.

9214_C008.indd 154

11/5/2007 7:39:30 AM

Industrial Relevance of Thermophiles and Their Enzymes

155

165. Posta, K., E. Beki, D. B. Wilson, J. Kukolya, and L. Hornok. Cloning, characterization and phylogenetic relationships of cel5B, a new endoglucanase encoding gene from Thermobifida fusca. J Basic Microbiol 44(5), 383–99, 2004. 166. Bok, J. D., D. A. Yernool, and D. E. Eveleigh. Purification, characterization, and molecular analysis of thermostable cellulases CelA and CelB from Thermotoga neapolitana. Appl Environ Microbiol 64, 4774–81, 1998. 167. Wright, R. M., M. D. Yablonsky, Z. P. Shalita, A. K. Goyal, and D. E. Eveleigh. Cloning, characterization, and nucleotide sequence of a gene encoding Microbispora bispora BglB, a thermostable betaglucosidase expressed in Escherichia coli. Appl Environ Microbiol 58, 3455–65, 1992. 168. Lebbink, J. H., T. Kaper, S. W. Kengen, J. van der Oost, and W. M. de Vos. beta-Glucosidase CelB from Pyrococcus furiosus: production by Escherichia coli, purification, and in vitro evolution. Methods Enzymol 330, 364–79, 2001. 169. Matsui, I., K. Ishikawa, H. Ishida, and Y. Kosugi. Methods for making and using a thermophilic enzyme as a beta-glycosidase. US patent 222866/1998, 2002. 170. Chi, Y. I., L. A. Martinez-Cruz, J. Jancarik, R. V. Swanson, D. E. Robertson, and S. H. Kim. Crystal structure of the beta-glycosidase from the hyperthermophile Thermosphaera aggregans: insights into its activity and thermostability. FEBS Lett 445, 375–83, 1999. 171. Yip, V. L., A. Varrot, G. J. Davies, S. S. Rajan, X. Yang, J. Thompson, W. F. Anderson, and S. G. Withers. An unusual mechanism of glycoside hydrolysis involving redox and elimination steps by a family 4 beta-glycosidase from Thermotoga maritima. J Am Chem Soc 126, 8354–5, 2004. 172. Park, T. H., K. W. Choi, C. S. Park, S. B. Lee, H. Y. Kang, K. J. Shon, J. S. Park, and J. Cha. Substrate specificity and transglycosylation catalyzed by a thermostable beta-glucosidase from marine hyperthermophile Thermotoga neapolitana. Appl Microbiol Biotechnol 69, 411–22, 2005. 173. Choi, J. J., E. J. Oh, Y. J. Lee, D. S. Suh, J. H. Lee, S. W. Lee, H. T. Shin, and S. T. Kwon. Enhanced expression of the gene for beta-glycosidase of Thermus caldophilus GK24 and synthesis of galactooligosaccharides by the enzyme. Biotechnol Appl Biochem 38, 131–6, 2003. 174. Wang, X., X. He, S. Yang, X. An, W. Chang, and D. Liang. Structural basis for thermostability of betaglycosidase from the thermophilic eubacterium Thermus nonproteolyticus HG102. J Bacteriol 185, 4248–55, 2003. 175. Akiyama, K., M. Takase, K. Horikoshi, and S. Okonogi. Production of galactooligosaccharides from lactose using a beta-glucosidase from Thermus sp. Z-1. Biosci Biotechnol Biochem 65, 438–41, 2001. 176. Fourage, L. and B. Colas. Synthesis of beta-D-glucosyl- and beta-D-fucosyl-glucoses using betaglycosidase from Thermus thermophilus. Appl Microbiol Biotechnol 56, 406–10, 2001. 177. Bronnenmeier, K., K. Kundt, K. Riedel, W. H. Schwarz, and W. L. Staudenbauer. Structure of the Clostridium stercorarium gene celY encoding the exo-1,4-beta-glucanase Avicelase, I.I. Microbiology 143, 891–8, 1997. 178. Zverlov, V. V., G. A. Velikodvorskaya, and W. H. Schwarz. A newly described cellulosomal cellobiohydrolase, CelO, from Clostridium thermocellum: investigation of the exo-mode of hydrolysis, and binding capacity to crystalline cellulose. Microbiology 148, 247–55, 2002. 179. Zhang, S., G. Lao, and D. B. Wilson. Characterization of a Thermomonospora fusca exocellulase. Biochemistry 34, 3386–95, 1995. 180. Ruttersmith, L. D. and R. M. Daniel. Thermostable cellobiohydrolase from the thermophilic eubacterium Thermotoga sp. strain FjSS3-B.1. Purification and properties. Biochem J 277, 887–90, 1991. 181. Beg, Q. K., M. Kapoor, L. Mahajan, and G. S. Hoondal. Microbial xylanases and their industrial applications: a review. Appl Microbiol Biotechnol 56, 326–38, 2001. 182. Morris, D. D., M. D. Gibbs, M. Ford, J. Thomas, and P. L. Bergquist. Family 10 and 11 xylanase genes from Caldicellulosiruptor sp. strain Rt69B.1. Extremophiles 3, 103–11, 1999. 183. Kosugi, A., K. Murashima, and R. H. Doi. Xylanase and acetyl xylan esterase activities of XynA, a key subunit of the Clostridium cellulovorans cellulosome for xylan degradation. Appl Environ Microbiol 68, 6399–402, 2002. 184. Kim, H., K. H. Jung, and M. Y. Pack. Molecular characterization of xynX, a gene encoding a multidomain xylanase with a thermostabilizing domain from Clostridium thermocellum. Appl Microbiol Biotechnol 54, 521–7, 2000. 185. Morris, D. D., M. D. Gibbs, C. W. Chin, M. H. Koh, K. K. Wong, R. W. Allison, P. J. Nelson, and P. L. Bergquist. Cloning of the xynB gene from Dictyoglomus thermophilum Rt46B.1 and action of the gene product on kraft pulp. Appl Environ Microbiol 64, 1759–65, 1998.

9214_C008.indd 155

11/5/2007 7:39:30 AM

156

Thermophiles: Biology and Technology at High Temperatures

186. Ramchuran, S. O., B. Mateus, O. Holst, and E. N. Karlsson. The methylotrophic yeast Pichia pastoris as a host for the expression and production of thermostable xylanase from the bacterium Rhodothermus marinus. FEMS Yeast Res 5, 839–50, 2005. 187. Kohli, U., P. Nigam, D. Singh, and K. Chaudhary. Thermostable, alkalophilic and cellulase free xylanase production by Thermoactinomyces thalophilus subgroup C. Enzyme Microb Technol 28, 606–610, 2001. 188. Irwin, D. C., M. Cheng, B. Xiang, J. K. Rose, and D. B. Wilson. Cloning, expression and characterization of a family-74 xyloglucanase from Thermobifida fusca. Eur J Biochem 270, 3083–91, 2003. 189. Wassenberg, D., H. Schurig, W. Liebl, and R. Jaenicke. Xylanase XynA from the hyperthermophilic bacterium Thermotoga maritima: structure and stability of the recombinant enzyme and its isolated cellulose-binding domain. Protein Sci 6, 1718–26, 1997. 190. Reeves, R. A., M. D. Gibbs, D. D. Morris, K. R. Griffiths, D. J. Saul, and P. L. Bergquist. Sequencing and expression of additional xylanase genes from the hyperthermophile Thermotoga maritima FjSS3B.1. Appl Environ Microbiol 66, 1532–7, 2000. 191. Lyon, P. F., T. Beffa, M. Blanc, G. Auling, and M. Aragno. Isolation and characterization of highly thermophilic xylanolytic Thermus thermophilus strains from hot composts. Can J Microbiol 46, 1029–35, 2000. 192. Donaghy, J. A., K. Bronnenmeier, P.-F. Soto-Kelly, and A. M. McKay. Purification and characterization of an extracellular feruloyl esterase from the thermophilic anaerobe Clostridium stercorarium. J Appl Microbiol 88(3), 458–66, 2000. 193. Shao, W. and J. Wiegel. Purification and characterization of two thermostable acetyl xylan esterases from Thermoanaerobacterium sp. strain JW/SL-YS485. Appl Environ Microbiol 61, 729–33, 1995. 194. Bachmann, S. L. and A. J. McCarthy. Purification and cooperative activity of enzymes constituting the xylan-degrading system of Thermomonospora fusca. Appl Environ Microbiol 57, 2121–2130, 1991. 195. Shao, W. and J. Wiegel. Purification and characterization of a thermostable beta-xylosidase from Thermoanaerobacter ethanolicus. J Bacteriol 174, 5848–53, 1992. 196. Xue, Y. and W. Shao. Expression and characterization of a thermostable beta-xylosidase from the hyperthermophile, Thermotoga maritima. Biotechnol Lett 26, 1511–5, 2004. 197. Bauer, M. W., E. J. Bylina, R. V. Swanson, and R. M. Kelly. Comparison of a beta-glucosidase and a beta-mannosidase from the hyperthermophilic archaeon Pyrococcus furiosus. Purification, characterization, gene cloning, and sequence analysis. J Biol Chem 271, 39, 23749–55, 1996. 198. Beki, E., I. Nagy, J. Vanderleyden, S. Jager, L. Kiss, L. Fulop, L. Hornok, and J. Kukolya. Cloning and heterologous expression of a beta-D-mannosidase (EC 3.2.1.25)-encoding gene from Thermobifida fusca TM51. Appl Environ Microbiol 69, 1944–52, 2003. 199. Gibbs, M. D., A. U. Elinder, R. A. Reeves, and P. L. Bergquist. Sequencing, cloning and expression of a beta-1,4-mannanase gene, manA, from the extremely thermophilic anaerobic bacterium, Caldicellulosiruptor Rt8B.4. FEMS Microbiol Lett 141, 37–43, 1996. 200. Gibbs, M. D., R. A. Reeves, A. Sunna, and P. L. Bergquist. Sequencing and expression of a betamannanase gene from the extreme thermophile Dictyoglomus thermophilum Rt46B.1, and characteristics of the recombinant enzyme. Curr Microbiol 39, 351–0357, 1999. 201. Politz, O., M. Krah, K. K. Thomsen, and R. Borriss. A highly thermostable endo-(1,4)-beta-mannanase from the marine bacterium Rhodothermus marinus. Appl Microbiol Biotechnol 53, 715–21, 2000. 202. Cann, I. K., S. Kocherginskaya, M. R. King, B. A. White, and R. I. Mackie. Molecular cloning, sequencing, and expression of a novel multidomain mannanase gene from Thermoanaerobacterium polysaccharolyticum. J Bacteriol 181, 1643–51, 1999. 203. Hilge, M., S. M. Gloor, W. Rypniewski, O. Sauer, T. D. Heightman, W. Zimmermann, K. Winterhalter, and K. Piontek. High-resolution native and complex structures of thermostable beta-mannanase from Thermomonospora fusca—substrate specificity in glycosyl hydrolase family 5. Structure 6, 1433–44, 1998. 204. Gomes, J., I. I. Gomes, K. Terler, N. Gubala, G. Ditzelmuller, and W. Steiner. Optimisation of culture medium and conditions for alpha-l-arabinofuranosidase production by the extreme thermophilic eubacterium Rhodothermus marinus. Enzyme Microb Technol 27, 414–422, 2000. 205. Birgisson, H., O. Fridjonsson, F.-K. Bahrani Mougeot, G. O. Hreggvidsson, J. K. Kristjansson, and B. Mattiasson. A new thermostable alpha-L-arabinofuranosidase from a novel thermophilic bacterium. Biotechnol Lett 26, 1347–51, 2004. 206. Miyazaki, K. Hyperthermophilic alpha-L-arabinofuranosidase from Thermotoga maritima MSB8: molecular cloning, gene expression, and characterization of the recombinant protein. Extremophiles 9, 399–406, 2005.

9214_C008.indd 156

11/5/2007 7:39:30 AM

Industrial Relevance of Thermophiles and Their Enzymes

157

207. Fukamizo, T. Chitinolytic enzymes: catalysis, substrate binding, and their application. Curr Protein Pept Sci 1, 105–24, 2000. 208. Zverlov, V. V., K. P. Fuchs, and W. H. Schwarz. Chi18A, the endochitinase in the cellulosome of the thermophilic, cellulolytic bacterium Clostridium thermocellum. Appl Environ Microbiol 68, 3176–9, 2002. 209. Kawase, T., A. Saito, T. Sato, R. Kanai, T. Fujii, N. Nikaidou, K. Miyashita, and T. Watanabe. Distribution and phylogenetic analysis of family 19 chitinases in Actinobacteria. Appl Environ Microbiol 70, 1135– 44, 2004. 210. Nawani, N. N., B. P. Kapadnis, A. D. Das, A. S. Rao, and S. K. Mahajan. Purification and characterization of a thermophilic and acidophilic chitinase from Microbispora sp. V2. J Appl Microbiol 93, 965–75, 2002. 211. Hobel, C. F., G. O. Hreggvidsson, V. T. Marteinsson, F. Bahrani-Mougeot, J. M. Einarsson, and J. K. Kristjansson. Cloning, expression, and characterization of a highly thermostable family 18 chitinase from Rhodothermus marinus. Extremophiles 9, 53–64, 2005. 212. Tanaka, T., T. Fukui, H. Atomi, and T. Imanaka. Characterization of an exo-beta-D-glucosaminidase involved in a novel chitinolytic pathway from the hyperthermophilic archaeon Thermococcus kodakaraensis KOD1. J Bacteriol 185, 5175–81, 2003. 213. Hla, Si. Si., J. Kurokawa, Suryani, T. Kimura, K. Ohmiya, and K. Sakka. A novel thermophilic pectate lyase containing two catalytic modules of Clostridium stercorarium. Biosci Biotechnol Biochem 69, 2138–45, 2005. 214. Stutzenberger, F. J. Inducible thermoalkalophilic polygalacturonate lyase from Thermomonospora fusca. J Bacteriol 169, 2774–80, 1987. 215. Parisot, J., A. Ghochikyan, V. Langlois, V. Sakanyan, and C. Rabiller. Exopolygalacturonate lyase from Thermotoga maritima: cloning, characterization and organic synthesis application. Carbohydr Res 337, 1427–33, 2002. 216. Ballschmiter, M., M. Armbrecht, K. Ivanova, G. Antranikian, and W. Liebl. AmyA, an alpha-amylase with beta-cyclodextrin-forming activity, and AmyB from the thermoalkaliphilic organism Anaerobranca gottschalkii: two alpha-amylases adapted to their different cellular localizations. Appl Environ Microbiol 71, 3709–15, 2005. 217. Kim, J. W., L. O. Flowers, M. Whiteley, and T. L. Peeples. Biochemical confi rmation and characterization of the family-57-like alpha-amylase of Methanococcus jannaschii. Folia Microbiol (Praha) 46, 467–73, 2001. 218. Gomes, I., J. Gomes, and W. Steiner. Highly thermostable amylase and pullulanase of the extreme thermophilic eubacterium Rhodothermus marinus: production and partial characterization. Bioresour Technol 90(2), 207–14, 2003. 219. Liebl, W., I. Stemplinger, and P. Ruile. Properties and gene structure of the Thermotoga maritima alphaamylase AmyA, a putative lipoprotein of a hyperthermophilic bacterium. J Bacteriol 179, 941–8, 1997. 220. Ballschmiter, M., O. Futterer, and W. Liebl. Identification and characterization of a novel intracellular alkaline alpha-Amylase from the hyperthermophilic bacterium Thermotoga maritima MSB8. Appl Environ Microbiol 72, 2206–11, 2006. 221. Egas, M. C., M. S. da Costa, D. A. Cowan, and E. M. Pires. Extracellular alpha-amylase from Thermus filiformis Ork A2: purification and biochemical characterization. Extremophiles 2(1), 23–32, 1998. 222. Gill, R. K. and J. Kaur. A thermostable glucoamylase from a thermophilic Bacillus sp.: characterization and thermostability. J Ind Microbiol Biotechnol 31, 540–3, 2004. 223. Hyun, H. H., G. J. Shen, and J. G. Zeikus. Differential amylosaccharide metabolism of Clostridium thermosulfurogenes and Clostridium thermohydrosulfuricum. J Bacteriol 164, 1153–61, 1985. 224. Ichikawa, K., T. Tonozuka, R. Uotsu-Tomita, H. Akeboshi, A. Nishikawa, and Y. Sakano. Purification, characterization, and subsite affinities of Thermoactinomyces vulgaris R-47 maltooligosaccharidemetabolizing enzyme homologous to glucoamylases. Biosci Biotechnol Biochem 68, 413–20, 2004. 225. Leveque, E., S. Janecek, B. Haye, and A. Belarbi. Thermophilic archaeal amylolytic enzymes. Enzyme Microbial Technol 26, 3–14, 2000. 226. Leveque, E., B. Haye, and A. Belarbi. Cloning and expression of an alpha-amylase encoding gene from the hyperthermophilic archaebacterium Thermococcus hydrothermalis and biochemical characterisation of the recombinant enzyme. FEMS Microbiol Lett 186, 67–71, 2000. 227. Albertson, G. D., R. H. McHale, M. D. Gibbs, and P. L. Bergquist. Cloning and sequence of a type I pullulanase from an extremely thermophilic anaerobic bacterium, Caldicellulosiruptor saccharolyticus. Biochim Biophys Acta 1354, 35–9, 1997.

9214_C008.indd 157

11/5/2007 7:39:30 AM

158

Thermophiles: Biology and Technology at High Temperatures

228. Bertoldo, C., F. Duffner, P. L. Jorgensen, and G. Antranikian. Pullulanase type I from Fervidobacterium pennavorans Ven5: cloning, sequencing, and expression of the gene and biochemical characterization of the recombinant enzyme. Appl Environ Microbiol 65, 2084–91, 1999. 229. Yang, S. J., H. S. Lee, C. S. Park, Y. R. Kim, T. W. Moon, and K. H. Park. Enzymatic analysis of an amylolytic enzyme from the hyperthermophilic archaeon Pyrococcus furiosus reveals its novel catalytic properties as both an alpha-amylase and a cyclodextrin-hydrolyzing enzyme. Appl Environ Microbiol 70, 5988–95, 2004. 230. Lin, F. P. and K. L. Leu. Cloning, expression, and characterization of thermostable region of amylopullulanase gene from Thermoanaerobacter ethanolicus 39E. Appl Biochem Biotechnol 97, 33–44, 2002. 231. Ramesh, B., P. R. Reddy, G. Seenayya, and G. Reddy. Effect of various flours on the production of thermostable beta-amylase and pullulanase by Clostridium thermosulfurogenes SV2. Bioresour Technol 76, 169–71, 2001. 232. Niehaus, F., A. Peters, T. Groudieva, and G. Antranikian. Cloning, expression and biochemical characterisation of a unique thermostable pullulan-hydrolysing enzyme from the hyperthermophilic archaeon Thermococcus aggregans. FEMS Microbiol Lett 190, 223–9, 2000. 233. Tomiyasu, K., K. Yato, M. Yasuda, T. Tonozuka, A. Ibuka, and H. Sakai. Cloning and nucleotide sequence of the pullulanase gene of Thermus thermophilus HB8 and production of the enzyme in Escherichia coli. Biosci Biotechnol Biochem 65, 2090–4, 2001. 234. Alcalde, M., F. J. Plou, M. Teresa Martin, I. Valdes, E. Mendez, and A. Ballesteros. Succinylation of cyclodextrin glycosyltransferase from Thermoanaerobacter sp. 501 enhances its transferase activity using starch as donor. J Biotechnol 86, 71–80, 2001. 235. Nakajima, M., H. Imamura, H. Shoun, S. Horinouchi, and T. Wakagi. Transglycosylation activity of Dictyoglomus thermophilum amylase A. Biosci Biotechnol Biochem 68, 2369–73, 2004. 236. Matzke, J., A. Herrmann, E. Schneider, and E. P. Bakker. Gene cloning, nucleotide sequence and biochemical properties of a cytoplasmic cyclomaltodextrinase (neopullulanase) from Alicyclobacillus acidocaldarius, reclassification of a group of enzymes. FEMS Microbiol Lett 183, 55–61, 2000. 237. Podkovyrov, S. M. and J. G. Zeikus. Structure of the gene encoding cyclomaltodextrinase from Clostridium thermohydrosulfuricum 39E and characterization of the enzyme purified from Escherichia coli. J Bacteriol 174, 5400–5, 1992. 238. Catara, G., G. Ruggiero, F. La Cara, F. A. Digilio, A. Capasso, and M. Rossi. A novel extracellular subtilisin-like protease from the hyperthermophile Aeropyrum pernix K1: biochemical properties, cloning, and expression. Extremophiles 7, 391–9, 2003. 239. Godde, C., K. Sahm, S. J. Brouns, L. D. Kluskens, J. van der Oost, W. M. de Vos, and G. Antranikian. Cloning and expression of islandisin, a new thermostable subtilisin from Fervidobacterium islandicum, in Escherichia coli. Appl Environ Microbiol 71, 3951–8, 2005. 240. Nam, G. W., D. W. Lee, H. S. Lee, N. J. Lee, B. C. Kim, E. A. Choe, J. K. Hwang, M. T. Suhartono, and Y. R. Pyun. Native-feather degradation by Fervidobacterium islandicum AW-1, a newly isolated keratinase-producing thermophilic anaerobe. Arch Microbiol 178, 538–47, 2002. 241. Chen, X. G., O. Stabnikova, J. H. Tay, J. Y. Wang, and S. T. Tay. Thermoactive extracellular proteases of Geobacillus caldoproteolyticus, sp. nov., from sewage sludge. Extremophiles 8, 489–98, 2004. 242. Gupta, R. and P. Ramnani. Microbial keratinases and their prospective applications: an overview. Appl Microbiol Biotechnol 70, 21–33, 2006. 243. Riessen, S. and G. Antranikian. Isolation of Thermoanaerobacter keratinophilus sp. nov., a novel thermophilic, anaerobic bacterium with keratinolytic activity. Extremophiles 5, 399–408, 2001. 244. Jang, H. J., C. H. Lee, W. Lee, and Y. S. Kim. Two flexible loops in subtilisin-like thermophilic protease, thermicin, from Thermoanaerobacter yonseiensis. J Biochem Mol Biol 35, 498–507, 2002. 245. Kim, T. and X. G. Lei. Expression and characterization of a thermostable serine protease (TfpA) from Thermomonospora fusca YX in Pichia pastoris. Appl Microbiol Biotechnol 68, 355–9, 2005. 246. Morikawa, M. and T. Imanaka. Thiol protease from Thermococcus kodakaraensis KOD1. Methods Enzymol 330, 424–33, 2001. 247. Lee, H. S., Y. J. Kim, S. S. Bae, J. H. Jeon, J. K. Lim, B. C. Jeong, S. G. Kang, and J. H. Lee. Cloning, expression, and characterization of aminopeptidase P from the hyperthermophilic archaeon Thermococcus sp. strain NA1.Appl Environ Microbiol 72, 1886–90, 2006. 248. Guy, E. J., M. N. Isupov, and J. A. Littlechild. The structure of an alcohol dehydrogenase from the hyperthermophilic archaeon Aeropyrum pernix. J Mol Biol 331, 1041–51, 2003. 249. Li, D. and K. J. Stevenson. Alcohol dehydrogenase from Thermococcus strain AN1. Methods Enzymol 331, 201–7, 2001.

9214_C008.indd 158

11/5/2007 7:39:31 AM

Industrial Relevance of Thermophiles and Their Enzymes

159

250. Lorentzen, E., B. Siebers, R. Hensel, and E. Pohl. Structure, function and evolution of the Archaeal class I fructose-1,6-bisphosphate aldolase. Biochem Soc Trans 32, 259–63, 2004. 251. Buchanan, C. L., H. Connaris, M. J. Danson, C. D. Reeve, and D. W. Hough. An extremely thermostable aldolase from Sulfolobus solfataricus with specificity for non-phosphorylated substrates. Biochem J 343 Pt 3, 563–70, 1999. 252. Izard, T. and J. Sygusch. Induced fit movements and metal cofactor selectivity of class II aldolases: structure of Thermus aquaticus fructose-1,6-bisphosphate aldolase. J Biol Chem 279, 11825–33, 2004. 253. Rhimi, M. and S. Bejar. Cloning, purification and biochemical characterization of metallic-ions independent and thermoactive l-arabinose isomerase from the Bacillus stearothermophilus US100 strain. Biochim Biophys Acta 1760, 191–9, 2005. 254. Jorgensen, F., O. C. Hansen, and P. Stougaard. Enzymatic conversion of D-galactose to D-tagatose: heterologous expression and characterisation of a thermostable L-arabinose isomerase from Thermoanaerobacter mathranii. Appl Microbiol Biotechnol 64, 816–22, 2004. 255. Lee, D. W., H. J. Jang, E. A. Choe, B. C. Kim, S. J. Lee, S. B. Kim, Y. H. Hong, and Y. R. Pyun. Characterization of a thermostable L-arabinose (D-galactose) isomerase from the hyperthermophilic eubacterium Thermotoga maritima. Appl Environ Microbiol 70, 1397–404, 2004. 256 Kim, B. C., Y. H. Lee, H. S. Lee, D. W. Lee, E. A. Choe, and Y. R. Pyun. Cloning, expression and characterization of L-arabinose isomerase from Thermotoga neapolitana: bioconversion of D-galactose to D-tagatose using the enzyme. FEMS Microbiol Lett 212, 121–6, 2002. 257. Thompson, V. S., K. D. Schaller, and W. A. Apel. Purification and characterization of a novel thermoalkali-stable catalase from Thermus brockianus. Biotechnol Prog 19, 1292–9, 2003. 258. Manco, G., E. Giosue, S. D’Auria, P. Herman, G. Carrea, and M. Rossi. Cloning, overexpression, and properties of a new thermophilic and thermostable esterase with sequence similarity to hormone-sensitive lipase subfamily from the archaeon Archaeoglobus fulgidus. Arch Biochem Biophys 373, 182–92, 2000. 259. Chen, S., A. F. Yakunin, E. Kuznetsova, D. Busso, R. Pufan, M. Proudfoot, R. Kim, and S. H. Kim. Structural and functional characterization of a novel phosphodiesterase from Methanococcus jannaschii. J Biol Chem 279, 31854–62, 2004. 260. Ishiguro, M., S. Kaneko, A. Kuno, Y. Koyama, S. Yoshida, G. G. Park, Y. Sakakibara, I. Kusakabe, and H. Kobayashi. Purification and characterization of the recombinant Thermus sp. strain T2 alphagalactosidase expressed in Escherichia coli. Appl Environ Microbiol 67, 1601–6, 2001. 261. Ohtsu, N., H. Motoshima, K. Goto, F. Tsukasaki, and H. Matsuzawa. Thermostable beta-galactosidase from an extreme thermophile, Thermus sp. A4: enzyme purification and characterization, and gene cloning and sequencing. Biosci Biotechnol Biochem 62, 1539–45, 1998. 262. Salleh, H. M., J. Mullegger, S. P. Reid, W. Y. Chan, J. Hwang, R. A. Warren, and S. G. Withers. Cloning and characterization of Thermotoga maritima beta-glucuronidase. Carbohydr Res 341, 49–59, 2006. 263. Erbeznik, M., S. E. Hudson, A. B. Herrman, and H. J. Strobel. Molecular analysis of the xylFGH operon, coding for xylose ABC transport, in Thermoanaerobacter ethanolicus. Curr Microbiol 48, 295–9, 2004. 264. Liu, S. Y., J. Wiegel, and F. C. Gherardini. Purification and cloning of a thermostable xylose (glucose) isomerase with an acidic pH optimum from Thermoanaerobacterium strain JW/SL-YS 489. J Bacteriol 178, 5938–45, 1996. 265. Chang, C., H. K. Song, B. C. Park, D. S. Lee, and S. W. Suh. A thermostable xylose isomerase from Thermus caldophilus: biochemical characterization, crystallization and preliminary X-ray analysis. Acta Crystallogr D Biol Crystallogr 55, 294–6, 1999. 266. Dekker, K., H. Yamagata, K. Sakaguchi, and S. Udaka. Xylose (glucose) isomerase gene from the thermophile Thermus thermophilus: cloning, sequencing, and comparison with other thermostable xylose isomerases. J Bacteriol 173, 3078–83, 1991. 267. Miyazaki, K. A hyperthermophilic laccase from Thermus thermophilus HB27. Extremophiles 9, 415– 25, 2005. 268. Cann, I. K., S. Ishino, N. Nomura, Y. Sako, and Y. Ishino. Two family B DNA polymerases from Aeropyrum pernix, an aerobic hyperthermophilic crenarchaeote. J Bacteriol 181, 5984–92, 1999. 269. Kahler, M. and G. Antranikian. Cloning and characterization of a family B DNA polymerase from the hyperthermophilic crenarchaeon Pyrobaculum islandicum. J Bacteriol 182, 655–63, 2000. 270. Dietrich, J., P. Schmitt, M. Zieger, B. Preve, J. L. Rolland, H. Chaabihi, and Y. Gueguen. PCR performance of the highly thermostable proof-reading B-type DNA polymerase from Pyrococcus abyssi. FEMS Microbiol Lett 217, 89–94, 2002. 271. Blondal, T., S. H. Thorbjarnardottir, J. Kieleczawa, S. Hjorleifsdottir, J. K. Kristjansson, J. M. Einarsson, and G. Eggertsson. Cloning, sequence analysis and functional characterization of

9214_C008.indd 159

11/5/2007 7:39:31 AM

160

272.

273. 274.

275. 276. 277.

278.

279. 280.

281. 282.

283.

284. 285. 286.

287.

288. 289.

290. 291. 292.

9214_C008.indd 160

Thermophiles: Biology and Technology at High Temperatures DNA polymerase I from the thermophilic eubacterium Rhodothermus marinus. Biotechnol Appl Biochem 34, 37–45, 2001. Nastopoulos, V., F. M. Pisani, C. Savino, L. Federici, M. Rossi, and D. Tsemoglou. Crystallization and preliminary X-ray diffraction studies of DNA polymerase from the thermophilic archaeon Sulfolobus solfataricus. Acta Crystallogr D Biol Crystallogr 54, 1002–4, 1998. Kim, D. J., H. J. Jang, Y. R. Pyun, and Y. S. Kim. Cloning, expression, and characterization of thermostable DNA polymerase from Thermoanaerobacter yonseiensis. J Biochem Mol Biol 35, 320–9, 2002. Bohlke, K., F. M. Pisani, C. E. Vorgias, B. Frey, H. Sobek, M. Rossi, and G. Antranikian. PCR performance of the B-type DNA polymerase from the thermophilic euryarchaeon Thermococcus aggregans improved by mutations in the Y-GG/A motif. Nucleic Acids Res 28, 3910–7, 2000. Margesin, R. and F. Schinner. Potential of halotolerant and halophilic microorganisms for biotechnology. Extremophiles 5, 73–83, 2001. Wojciechowski, C. L., J. P. Cardia, and E. R. Kantrowitz. Alkaline phosphatase from the hyperthermophilic bacterium Thermotoga maritima requires cobalt for activity. Protein Sci 11, 903–11, 2002. Ji, C. N., T. Jiang, M. Q. Chen, X. Y. Sheng, and Y. M. Mao. Purification, crystallization and preliminary X-ray studies of thermostable alkaline phosphatase from Thermus sp. 3041. Acta Crystallogr D Biol Crystallogr 57, 614–5, 2001. Gong, N., C. Chen, L. Xie, H. Chen, X. Lin, and R. Zhang. Characterization of a thermostable alkaline phosphatase from a novel species Thermus yunnanensis sp. nov. and investigation of its cobalt activation at high temperature. Biochim Biophys Acta 1750, 103–11, 2005. Hoe, H. S., H. K. Kim, and S. T. Kwon. Cloning, analysis, and expression of the gene for inorganic pyrophosphatase of Aquifex pyrophilus and properties of the enzyme. Mol Cells 13, 296–301, 2002. Laksanalamai, P., A. Jiemjit, Z. Bu, D. L. Maeder, and F. T. Robb. Multi-subunit assembly of the Pyrococcus furiosus small heat shock protein is essential for cellular protection at high temperature. Extremophiles 7, 79–83, 2003. Maruyama, T., R. Suzuki, and M. Furutani. Archaeal peptidyl prolyl cis-trans isomerases (PPIases) update 2004. Front Biosci 9, 1680–720, 2004. Suzuki, R., K. Nagata, F. Yumoto, M. Kawakami, N. Nemoto, M. Furutani, K. Adachi, T. Maruyama, and M. Tanokura. Three-dimensional solution structure of an archaeal FKBP with a dual function of peptidyl prolyl cis-trans isomerase and chaperone-like activities. J Mol Biol 328, 1149–60, 2003. Ideno, A., M. Furutani, T. Iwabuchi, T. Iida, Y. Iba, Y. Kurosawa, H. Sakuraba, T. Ohshima, Y. Kawarabayashi, and T. Maruyama. Expression of foreign proteins in Escherichia coli by fusing with an archaeal FK506 binding protein. Appl Microbiol Biotechnol 64, 99–105, 2004. Lund, P. A., A. T. Large, and G. Kapatai. The chaperonins: perspectives from the Archaea. Biochem Soc Trans 31, 681–5, 2003. Fox, J. D., K. M. Routzahn, M. H. Bucher, and D. S. Waugh. Maltodextrin-binding proteins from diverse bacteria and archaea are potent solubility enhancers. FEBS Lett 537, 53–7, 2003. Yan, Z., S. Fujiwara, K. Kohda, M. Takagi, and T. Imanaka. In vitro stabilization and in vivo solubilization of foreign proteins by the beta subunit of a chaperonin from the hyperthermophilic archaeon Pyrococcus sp. strain KOD1. Appl Environ Microbiol 63, 785–9, 1997. Yano, J. K., F. Blasco, H. Li, R. D. Schmid, A. Henne, and T. L. Poulos. Preliminary characterization and crystal structure of a thermostable cytochrome P450 from Thermus thermophilus. J Biol Chem 278, 608–16, 2003. Eichler, J. Facing extremes: archaeal surface-layer (glyco)proteins. Microbiology 149, 3347–51, 2003. DeLong, E. F., L. L. King, R. Massana, H. Cittone, A. Murray, C. Schleper, and S. G. Wakeham. Dibiphytanyl ether lipids in nonthermophilic crenarchaeotes. Appl Environ Microbiol 64, 1133–8, 1998. Upreti, R., M. Kumar, and V. Shankar. Bacterial glycoproteins: functions, biosynthesis and applications. Proteomics 3, 363–379, 2003. Gomes, J. and W. Steiner. The biocatalytic potential of extremophiles and extremozymes. Food Technol Biotechnol 42, 223–235, 2004. Jolivet, E., S. L’Haridon, E. Corre, P. Forterre, and D. Prieur. Thermococcus gammatolerans sp. nov., a hyperthermophilic archaeon from a deep-sea hydrothermal vent that resists ionizing radiation. Int J Syst Evol Microbiol 53, 847–51, 2003.

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Denitrification Pathway Enzymes of Thermophiles Simon de Vries and Imke Schröder

CONTENTS Introduction ................................................................................................................................ Phylogeny of Denitrifying Microorganisms .............................................................................. Analysis of Genes Involved in Denitrification of Thermophiles ............................................... Variation of Denitrification Respiratory Chains ........................................................................ Biochemical Properties of Purified Denitrification Pathway Enzymes ..................................... Nitrate Reductase ............................................................................................................ Nitrite Reductase ............................................................................................................ Nitric Oxide Reductase ................................................................................................... Nitrous Oxide Reductase ................................................................................................ Concluding Remarks .................................................................................................................. Acknowledgment ....................................................................................................................... References ..................................................................................................................................

162 162 163 166 167 167 169 170 170 171 171 171

Denitrification is an important part of the global nitrogen cycle, however, the extent that extremophilic bacteria and archaea contribute to this cycle is unclear as only few isolates are obtained in pure culture and even fewer genomes of these have been sequenced. This review focuses on the denitrification pathway of thermophilic bacteria and archaea. While thermophilic and mesophilic denitrifiers share the same type of pathway enzymes important differences exist with respect to the localization and cofactors of the denitrification pathway enzymes. The most significant difference is the exterior orientation of the archaeal nitrate reductase (Nar) that catalyzes the first step in the denitrification pathway. As a consequence of this orientation, archaeal Nars do not participate directly in the generation of the proton motive force. In the archaea, all denitrification pathway enzymes are membrane-associated. This is similar to the pathway enzymes from a gram-positive Bacillus sp., only one of which has been studied biochemically in more detail. This review will provide insight into the details of denitrification pathway genes and enzymes of selected thermophiles that have been characterized to date.

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INTRODUCTION –

Denitrification is the five-electron reduction of nitrate (NO3 ) to dinitrogen (N2) in which nitrite, nitric oxide, and nitrous oxide occur as intermediates according to: NO3 → NO2 → NO → N2O → N2. –

–

This series of redox reactions is catalyzed by Nar, nitrite reductase (Nir), NO reductase (Nor), and N2O reductase (Nos) [1–5]. Denitrification is an anaerobic dissimilatory process serving the bioenergetic needs of the cell. Since each intermediate potentially acts as a terminal electron acceptor of nitric oxide respiration, denitrification also serves to drain excess reducing equivalents from intermediary metabolism. The denitrification pathway plays an important role in the global nitrogen cycle as it is responsible for converting the majority of the fixed nitrogen to nitrogen gas. Because the pathway is often incomplete or the enzymes not well coupled, intermediate gases such as the highly reactive NO and the green house gas nitrous oxide are released into the environment. To date, the contribution of thermophiles or their mechanism of denitrification is poorly understood mainly due to the lack of isolates from various extreme environments. The denitrification pathway is distinct from the dissimilatory ammonification pathway in which nitrate is reduced to ammonium according to: NO3 → NO2 → NH 4 . –

–

+

Both pathways share the first enzyme, Nar. However, Nir that catalyzes the reduction of nitrite is distinct in both pathways. The ammonification pathway deploys a Nrf-type enzyme which catalyzes the six electron reduction from nitrite to ammonium [6,7]. While this pathway is also found in thermophiles, this chapter will focus on the enzymes of the denitrification pathway. All denitrification enzymes from proteobacteria are metallo-redox enzymes and have been characterized in great detail. High-resolution structures are available for all enzymes except for Nor [8–16]. The genetic organization of denitrification, and the many spectroscopic and functional studies performed on enzymes from denitrifying proteobacteria form an important source of understanding denitrification in other microorganisms including archaea. This review will highlight physiological, genetic, and biochemical aspects of anaerobic denitrification in bacteria and archaea highlighting recent findings in thermophiles.

PHYLOGENY OF DENITRIFYING MICROORGANISMS Denitrification is found among members of all three domains of life (Figure 9.1) [17–19]. Within the bacteria it is present in three phyla: the Proteobacteria [1,4,20,21], the Bacilli (e.g., Bacillus azotoformans [22], Bacillus halodenitrificans [23]), and the Aquifecales, the latter represented by the hyperthermophile Aquifex pyrophilus [24,25]. Among the bacteria, several strains are halotolerant, halophilic, and/or alkaliphilic (Paracoccus halomonas sp., Halomonas sp., and Thioalkalivibrio [26–28]), whereas some B. azotoformans strains have been reported to grow at temperatures up to 60°C [29]. To date, little is known about the denitrification enzymes from various thermophilic bacteria and some interesting features may yet be discovered that bridge to archaeal enzymes. As compared with bacterial denitrifiers only few archaea have been isolated thus far providing limited insight into the breath of denitrification in this domain. Within the Crenarchaeaota branch two denitrifyers are found: Pyrobaculum aerophilum, a facultative anaerobe with optimum growth temperature of 100°C, and Ferroglobus placidus, a strict anaerobic iron oxidizer that grows optimally at 85°C [30,31]. Examples for denitrifiers among the Euryarchaeota branch include the halophilic Haloferax mediterranei, Haloarcula marismortui, Haloarcula vallismortui, and Haloferax denitrificans [32–34].

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Bacteria

Eukarya

Archaea Fungi

Bacilli Deinococcus Proteobacteria Green non-sulphur

Crenarchaeota

bacteria

Cyanobacteria Green sulphur bacteria Bacteroidetes High-G+C Gram+ Bacteria

Pyrobaculum

Korarchaeota

Methanobacterium Archeoglobus

Spirochetes Planctomycetes Chlamydiae

Aquifecales

Sulfolobus

Ferroglobus Mc. jannaschii

Nanoarchaeota

Thermotogales

Haloarcula Haloferax Methanosarcina

Euryarchaeota LUCA

FIGURE 9.1 Phylogenetic tree highlighting the distribution of denitrifying organisms. The tree is adapted from Stetter [17] and combines recent findings [18,19]. Thick lines indicate lineages with hyperthermophilic organisms. Names of phyla that are underlined contain denitrifyers. LUCA is the abbreviation for last universal common ancestor assumed to be (hyper)thermophilic.

Denitrification has been observed at pH values as low as 2.8, but the responsible microorganisms were not obtained in pure culture. Growth at low pH appears to be limited by the toxicity of nitrite, which converts to the uncharged nitrous acid (pKa ~ 3.3) under acidic conditions and permeates the cell membrane [35].

ANALYSIS OF GENES INVOLVED IN DENITRIFICATION OF THERMOPHILES While only few denitrifying extremophiles have been isolated thus far, even fewer genome sequences of these microbes are publicly available and thus gene analysis will be confined to the archaeon P. aerophilum [36]. In the following section the denitrification genes of P. aerophilum will be compared with genes found in other thermophiles and few mesophilic denitrifying archaea. The extremophiles used for comparison are not denitrifiers per se as they contain genes for only one of the pathway enzymes indicating limited use of oxidized nitrogen compounds. The hyperthermophilic bacterium Thermus thermophilus, which reduces nitrate and secretes nitrite, for example, contains only the Nar genes [37,38]. Genes for Nar are also found in the hyperthermophilic, aerobic archaeon Aeropyrum pernix, whereas a Nor gene is present in the strict aerobic archaeon Sulfolobus solfataricus [39]. Whether all these genes are expressed is not known. However, gene comparison provides some insight into the organization and conserved features of denitrification pathway genes in thermophiles. Nars are encoded by the nar locus consisting of two highly conserved genes, narG and narH, a unique membrane anchor gene, and a private chaperone gene, narJ (Figure 9.2a). The narG and narH genes encode the molybdenum and [Fe-S]-containing, hydrophilic subunits, respectively, and are remarkably conserved throughout all prokaryotes sharing about 40% to 60% amino sequence identity. A clear distinction of archaeal NarG polypeptides is the presence of an N-terminal twin-arginine motif suggestive for translocation of the soluble subunits via the TAT translocation system and, thus, an exterior location of the archaeal Nar active site (Figure 9.2b) [40,41]. In contrast,

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A. pernix P. aerophilum H. marismortui H. mediterranei

narJ

petB

petB

petA

petA

T. thermophilus

narC

P. arcticus

narK

narG

narH

narM

narJ

narG

narH

narM

narJ

narG

narH

narG

narH

narM

narJ

narG

narH

narJ

narI

narG

narH

narJ

narI

narM

narJ

narK

narT

(b) Molybdopterin

MLKTTRRRMLAGVATITAAA DLTDDEGDSAGISRRDFVRGLGAASLLG DPPGDPVDADSGVSRRTFLEGIGVASLLG

Molybdopterin

P. aerophilum H. marismortui H. mediterranei

(S/T)RRXFLK

FIGURE 9.2 Organization of membrane-bound nitrate reductase gene clusters from extremophiles (a). Arrows designate the direction of transcription. The narG and narH genes in black indicated high amino acid sequence identity. The narK and narT genes encode nitrate/nitrite transporters. Archaeal NarGs contain an N-terminal twin-Arg motif that targets the enzymes to the TAT protein translocation machinery for export (b). Barrels indicate cofactor binding motifs; the shaded box designates the twin-Arg motif enlarged below as partial amino acid sequence.

the T. thermophilus NarG and all other bacterial NarGs lack a twin-arginine motif consistent with their cytoplasmic location. The bacterial membrane anchor gene narI is not conserved in any of the archaeal nar operons available to date (Figure 9.2a). Instead, all archaeal nar operons contain a nar-associated gene, termed narM, conserved also with few bacterial gamma subunit genes for selenate and chlorate reductases and for ethylbenzene and dimethylsulfide dehydrogenases. The narM gene encodes a hydrophobic protein annotated to bind one heme b in the bacterial homologs. Whether the narM gene product serves as the membrane anchor for the halophilic Nars is ground for speculation. For H. mediterranei, the petB gene product has been implicated to serve this function [42]. The petB gene encodes a hydrophobic cytochrome b/b6-like protein and is conserved in both halophiles, in T. thermophilus, where it is, however, not linked to the nar locus, and in many other bacteria. In all cases, petB forms a potential transcriptional unit with petA encoding a Rieske-type [2Fe-2S] protein. It is, thus, likely that petA and petB of H. mediterranei and H. marismortui function as bc1-type complex to provide reducing power to the denitrification pathway enzymes in these halophiles. A variation of a bacterial membrane anchor was found in T. thermophilus. Here, narC gene encodes a di-heme c-type cytochrome, which was shown by Zafra et al. to interact with NarI suggesting that

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Denitrification Pathway Enzymes of Thermophiles nirSc

nirSd

P. aerophilum UbiA prenyl Heme c transferase

RR

Heme d1

Uro III methyl transferase

nirS Bacteria

Heme c

Heme d1

FIGURE 9.3 The Pyrobaculum aerophilum nitrite reductase gene cluster, nirSc and nirSdI, in comparison with the canonical bacterial nirS gene. Arrows designate the direction of transcription. Barrels indicate cofactor binding motifs; the shaded box and RR designate the twin-Arg motif.

a NarCI complex forms the membrane anchor for the NarGH subunits in T. thermophilus [43]. The private chaperone, NarJ, is only distantly related between the two halophiles and P. aerophilum sharing 28% amino acid sequence identity. The low similarity with bacterial NarJ chaperones (less than 25%) suggests an organism-specific adaptation of this protein. Nirs are encoded by the nirS and nirK genes dependent on whether the enzyme contains cytochrome cd1 or Cu, respectively. The P. aerophilum nirS gene is split into a cytochrome c, nirSc, and a cytochrome d1 gene, nirSd, that are divergently transcribed (Figure 9.3). In contrast, bacterial NirS-type enzymes consist of a single polypeptide with an N-terminal cytochrome c and a C-terminal cytochrome d1 domain. The P. aerophilum NirSc subunit is distantly related to class I, soluble cytochrome c proteins and shares approximately 25% sequence identity with the cytochrome c domain of the one subunit cd1-type NirS nitrite reductase from bacteria. All cytochrome proteins have a conserved His and a distantly located (not conserved) Met in common that serve as ligands to heme c. NirS-type nitrite reductases appear to be confined to thermophilic crenarchaeota, in contrast to the halophilic euryarchaeota that contain the copper-dependent NirK nitrite reductase. Nor genes, nor, are present in P. aerophilum S. solfataricus, and H. marismortui. All archaeal nor genes encode the membrane-bound, quinone-dependent qNOR-type Nor (Figure 9.4). Bacterial cNOR-type enzymes consisting of the NorBC subunits appear to be absent in the archaea. The P. aerophilum qNor shares 42% amino acid sequence identity with the S. solfataricus enzyme and both are distantly related to the qNOR from H. marismortui and bacteria (about 25% amino acid sequence identity). The qnor and norB gene products contain six conserved His, three of which serve as ligands to the high- and low-spin heme b in bacteria or to the hemes of the Op-type found in P. aerophilum, and three to the nonheme Fe atom [2,3,44–47]. The N-terminal putative quinonebinding domain of qNORs exhibits some similarity to NorCs suggesting an evolutionary relatedness and functional divergence of the proteins to accommodate either quinol of small redox proteins as electron donors to the complex [45]. Nos encoded by nosZ have been thus far identified only in H. marismortui. The amino acid sequence of this and bacterial NosZs is highly conserved with an overall sequence identity of 47%. The N-terminus of the archaeal and most bacterial NosZ enzymes contains a twin-arginine motif targeting the protein for TAT-dependent protein translocation. In P. aerophilum a nosZ-type gene is absent. This is particularly puzzling as this archaeon is known to reduce nitrate to N2 gas and has Quinone

Heme Op1, Op2, FeB

Heme c

Heme b1, b2, FeB

P. aerophilum nor

Bacterial norBC

FIGURE 9.4 Comparison of gene architecture of the Pyrobaculum aerophilum qNor and bacterial cNORs that consist of the NorBC subunits. Barrels indicate cofactor binding motifs.

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NarK

B. azotoformans

NO2- NO3

NarK

(M)QH2 Nar GHI cyt. c/ps Az.

NO3-

Cu/cd1 Nir

Nar GHI

cyt. c/ps Az.

NO2NO

NO3- MQH2

q/c Nor

cyt. c

NO3-

H+ MQH2 /cyt. c

NO2-

Cu-Nir

qCuA Nor

Mem Cyt

Nar GHM

NO2-

Cu-Nir

MQH2 NO2-

halocyanin

NO

qNor

cd1Nir MQH2

NO

qNor

halocyanin

MQH2

Nos

Nos

Nos

N2O Peri

Nar GHM

MQH2 / cyt. c

Nos N2O

NO3- MQH2

halocyanin

MQH2 /cyt. c

NO

P. aerophilum

NO2-

MQH2 H+

H. marismortui

N2O Mem Cyt

N2O Mem Cyt

Mem Cyt

FIGURE 9.5 Variation of denitrification pathways in archaea and bacteria based on genetic, physiological, and biochemical data detailed in the text. In Bacillus azotoformans, Haloarcula marismortui, and Pyrobaculum aerophilum the four denitrfication enzymes and their respective electron donors are membrane bound. In pro– – teobacteria and B. azotoformans NarK facilitates NO3 and NO2 transport across the cytoplasmic membrane; nitrate is reduced to nitrite in the cytoplasm generating a proton electrochemical gradient (indicated by → H+). – In the archaea NO3 is reduced at the exterior face of the cytoplasmic membrane. Peri, Mem, and Cyt designate periplasm, cytoplasmic membrane, and cytoplasm, respectively. MQH2 is menaquinol; QH2, is ubiquinol, and ps Az means pseudo-azurin. Bacterial and archaeal menaquinones differ in ring substituents and the nature and length of the isoprene chain. Abbreviations for enzymes are explained in the text.

been shown to contain membrane-bound Nos activity [47,48]. The absence of this gene may be a result of the incomplete genome sequence for this archaeon [36].

VARIATION OF DENITRIFICATION RESPIRATORY CHAINS Microbial denitrification electron transfer chains display a great variety of electron mediators (menaquinone or ubiquinone, soluble or membrane bound c-cytochromes, blue copper proteins). In addition, the type and location of the specific denitrification enzymes can vary (Figure 9.5). In all microbes both Nar and Nor are membrane-bound enzymes. Nir and Nos from gram-negative bacteria are soluble, periplasmic enzymes, while they are membrane-bound in gram-positive bacteria and archaea. In contrast to bacteria, the reduction of nitrate to nitrite by thermophilic and other archaea occurs at the cell exterior. This architecture constitutes an important bioenergetic difference between the archaea and the bacteria. Bacteria transport nitrate by the NarK protein, which displays three activities: (i) import of nitrate in exchange for protons, (ii) import of nitrate in exchange for nitrite, and (iii) export of excess nitrite to rid the cell of the toxic intermediate [49,50]. After transport to the cytoplasm, nitrate is reduced to nitrite by the bacterial membrane-bound NarGHI complex using (mena)quinol [(M)QH2] as electron donor (Figure 9.5). The electrons from (M)QH2 traverse the cytoplasmic membrane to the interior-oriented NarG catalytic subunit. This arrangement leads to generation of a proton motive force. Nitrite is subsequently transported to the periplasm by NarK presenting it for further reduction to the periplasmic denitrification pathway enzymes [1]. In contrast, the nitrate reduction site of the membrane-bound archaeal NarGHM complex faces the S-layer [51]. The important

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167

bioenergetic consequence of this arrangement is that nitrate reduction does not contribute to the proton electrochemical gradient in archaea. In P. aerophilum all further reduction steps of the denitrification pathway are catalyzed by membrane-bound Nor that face the exterior face of the cytoplasmic membrane. While P. aerophilum contains a membrane-bound cytochrome cd1-type Nir the Cu-dependent nitrite Nirk-type reductases from H. marismortui and H. denitrificans were purified as soluble proteins inconsistent with an exterior localization of these proteins (Figure 9.5) [52,53]. Three different types of membrane-bound Nors have been characterized, the single subunit qNor present in bacteria and archaea, the two subunit cNor type only found in bacteria and the two subunit qCuANor, thus far only detected in B. azotoformans [1,2,54–57]. Although nitric oxide reduction is carried out by an integral membrane protein, the reaction does not contribute to the proton motive force because the nitric oxide reduction site and the electron donor are both located at the outer face of the cytoplasmic membrane [58]. The soluble Nos from proteobacteria are dimeric copper-containing enzymes located in the periplasm [1,3,16]. Interestingly, Nos activity in P. aerophilum has been shown to be membrane-bound [44,48]. To date, an Nos has not yet been purified from any thermophilic archaeon. Quinols seem to be the exclusive electron donor to the Nars. All other denitrification enzymes receive electrons from a variety of small electron carriers. Particularly in proteobacteria soluble and or membrane-associated c-type cytochromes or blue copper proteins, (pseudo)azurin, mediate electrons to Nir, Nor, and Nos. In contrast, menaquinol is the sole electron donor to all four denitrification enzymes in P. aerophilum [44,48]. In H. marismortui the membrane-bound blue copper proteins halocyanin and plastocyanin are the likely electron donors to Nir, Nor, and/or Nos. The Nir, Nor, and Nos from B. azotoformans are bifunctional, receiving electrons from menaquinol and various types of membrane bound c-cytochromes [22,59].

BIOCHEMICAL PROPERTIES OF PURIFIED DENITRIFICATION PATHWAY ENZYMES In contrast to the archaeal denitrification pathway enzymes, the proteobacterial denitrification enzymes have been studied in great detail and this provides the platform for comparing the biochemical properties of the enzymes involved.

NITRATE REDUCTASE The dissimilatory Nars from bacteria and archaea are highly similar with respect to two of their three subunits, NarG and NarH [5,7,43,51,60–62]. The smallest subunits, NarI in bacteria and NarM in archaea show no sequence similarity, but are responsible for anchoring the intact NarGH to the membrane. The different membrane anchor proteins reflect most likely the distinct lipid properties of the archaeal versus bacterial cytoplasmic membrane. In proteobacteria, Nars are dimeric enzymes containing a molybdopterin cofactor (bis-molybdopterin guanine dinucleotide or Mo-bisMGD) where reduction from nitrate to nitrite takes place. The Mo-bisMGD is located in the largest subunit, NarG (Figure 9.6). Tightly associated with Mo-bisMGD is a modified [4Fe-4S] cluster (Fe-S0) in which three cysteine residues and one histidine serve as ligands to the four iron atoms. The Fe-S0 cluster is the direct electron donor to the Mo-bisMGD (Figure 9.6) [9,10]. NarH contains a chain of four different iron–sulfur centers each separated by approximately 12–14Å (center-to-center) enabling very rapid electron transfer. Fe-S1, Fe-S4, and Fe-S2 are [4Fe-4S] clusters, Fe-S3 is a [3Fe-4S] cluster located most closely to the heme bp in NarI and usually has a relatively high reduction potential. Interestingly, Fe-S4 has a very low reduction potential (–420 mV), which will decrease the rate of electron transfer between this cluster and its neighbors. However, given their relative short distance, actual electron transfer could still be in the microseconds whereas substrate turnover occurs in the milliseconds, thus electron transfer through the Fe-S4 center may

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Thermophiles: Biology and Technology at High Temperatures NO2–+ H2O

NO3–+ 2H+

bis-MoMGD

NarG

Fe-S Fe-S00 Fe-S1 Fe-S4

NarH 2H+ + MQ

N2O+ H2O

Fe-S2

b

MQH2

2NO +

MQH2

Fe-S3

Op 1 Op 2

2H+

FeB

Membrane qNor

NarM Cytoplasm

FIGURE 9.6 Subunit structures and approximate cofactor location with respect to the membrane in archaeal nitrate reductase (NarGHM) and qNor (NorB). The nitrate and nitric oxide reduction sites are located at the exterior face of the cytoplasmic membrane, facing the S-layer. Nitrate reductase is probably present as a dimer—(NarGHM)2—of the three subunits, but only the monomer is shown. Nitrate is reduced by the Mo-pterin cofactor of NarG. NarG also harbors Fe-S0. Fe-S1–4 are located in NarH. NarM contains a single heme b proposed to be accessible by MQH2 from the exterior face of the cytoplasmic membrane. The direction of electron transfer from MQH2 via heme b, the Fe-S centers, and the Mo-pterin to nitrate is indicated by thin arrows. The qNor reduces nitric oxide to nitrous oxide at the heme Fe-FeB binuclear center. Electron flow in qNor is indicated as for NarGHM. MQH2 binds to the N-terminal domain of NorB. The location of the hemes Op1 or Op2, which are heme b types in bacteria, is indicated. FeB is the nonheme iron center.

not limit the overall activity [63]. A similar chain of Fe-S centers is found in succinate dehydrogenases and fumarate reductases, also containing one very low-potential Fe-S center. The precise catalytic function—perhaps the regulation of enzyme activity—of such a low-potential cluster remains to be established. In bacterial Nars, NarI contains five transmembrane α-helices, two phospholipids and two transmembrane-oriented hemes b, bp, and bd. Two NarI subunits associate by several hydrophobic interactions while NarG and NarH are connected through numerous hydrogen bonds and electrostatic interactions resulting in a (NarGHI)2 dimeric quaternary structure. (M)QH2 reduces heme bd first; from there electrons travel across the membrane—the energy conserving step—towards heme bp, and subsequently via the chain of Fe-S centers to the Mo-bisMGD crossing a distance of 125Å [9,13]. The Mo-bisMGD provides the molybdenum atom, which shuttles between MoIV, MoV, and MoVI during catalysis, with four cis-dithiolene sulfur ligands. In the oxidized enzyme the MoVI is also coordinated in a bidentate fashion by both carboxylate oxygens from a conserved aspartate residue [10]. The catalytic cycle of Nar is described by the reactions: (M)QH2 + MoVI-Asp → (M)Q + MoIV + 2H+ + unliganded-Asp MoIV + NO3– → MoVI ≡ O + NO2–

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(9.1) (9.2)

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Denitrification Pathway Enzymes of Thermophiles

(M)QH2 + MoVI ≡ O + 2H+ → (M)Q + MoIV + H2O second and next turnovers

169

(9.3)

During catalysis the aspartate must move away allowing nitrate to bind; this might occur upon reduction of MoVI to MoIV by menaquinol or ubiquinol (Equation 9.1). After binding of nitrate and reduction to nitrite, the oxo group is transferred to the molybdenum atom yielding MoVI⌶O (Equation 9.2). Upon reduction, the oxo group is released as water and the MoIV is ready to bind nitrate (Equation 9.3). According to this mechanism binding and rebinding of aspartate does not form part of the catalytic cycle. However, since this aspartate residue is highly conserved in Nars and certain other molybdoenzymes such as selenate reductase but not in dimethylsulfoxide (DMSO) reductases, formate dehydrogenases or the bacterial periplasmic Nars, Nap, its physiological function seems important but yet remains to be elucidated. To date archaeal Nars have been purified from P. aerophilum, H. marismortui, and H. mediterranei. The P. aerophilum enzyme was purified as a three-subunit enzyme containing heme b (NarGHM) [62]. The H. marismortui and H. mediterranei enzymes, which are stable in the absence of salt enabling their purification, were obtained as soluble, two-subunit enzymes with a (NarGH)2 quaternary structure [42,51,64]. The subunit masses of NarG, NarH, and NarM are approximately 120, 50, and 30 kDa, respectively. The archaeal enzymes contain molybdenum, most likely as Mo-bisMGD and similar amounts of iron and acid-labile sulfur per enzyme as the bacterial Nars. However, in the P. aerophilum enzyme the amount of heme b per NarGHM seems lower than two as for NarGHI and is closer to one per enzyme (Figure 9.6) [62]. The archaeal NarM lacks the four conserved histidine residues present in NarI, which serve as fifth and sixth ligands to the low spin heme. The archaeal Nars are, like the bacterial ones, highly active with nitrate and chlorate as substrate. Reduced heme b of NarGHM can be oxidized by nitrate as well as by chlorate [62]. The Nar activity of the H. marismortui enzyme is stimulated twofold in the presence of 2 M NaCl (143/s, Km = 79 μM); the activity of the H. mediterranei enzyme is independent of the salt concentration, but retains activity up to 70°C. The Nar from P. aerophilum has a very high activity (1130/s for nitrate, Km = 58 μM; 1300/s for chlorate Km = 140 μM, both at 75°C), which increases twofold at 95°C, the highest temperature experimentally accessible. Electron paramagnetic resonance (EPR) of the H. marismortui Nar revealed a MoV redox state containing a D2O exchangeable proton, suggesting coordination to an OH– ligand. EPR spectroscopy further showed resonances from Fe-S3 and Fe-S1 and the g = 5.7 and 5.0 signals from Fe-S0 [8,51,64]. The presence of other iron–sulfur centers, Fe-S2 and Fe-S4 is indicated by the broad unresolved peaks in the spectrum. The main distinction between archaeal and bacterial Nars (including the Nar from T. thermophilus) remains the active site orientation with respect to the membrane. The archaeal Nars have an exterior orientation while the bacterial Nars face the cytoplasm [51]. Electron transfer from the archaeal menaquinol to the heme b presumed to be located at the exterior face of the cytoplasmic membrane does not lead to transmembrane electron transport, and as a result does not generate a membrane potential [q/e = 0 (Figure 9.6)]. The reduced heme b likely donates its electron directly to the Fe-S3 center. Because of this arrangement archaeal Nars require only one heme per enzyme complex.

NITRITE REDUCTASE Enzymological data on the P. aerophilum membrane-bound cd1 Nir are presently not available. Copper-containing Nirs have been purified from H. denitrificans and H. marismortui [52,53]. The archaeal enzymes are homotrimers each subunit containing two copper sites. The type I blue copper site mediates electron transfer on the submillisecond time scale between the physiological electron donor and the type II (nonblue) copper site in the enzyme where reduction of nitrite to nitric oxide takes place. Interestingly, nitrite is oxygen-bonded to the type II copper [2,12] and the reaction is reversible [65]. Both the H. denitrificans and H. marismortui enzymes require 2M NaCl for optimal – activity, which amounts to approximately 600NO2 /s. Both halophlic NirKs were purified as soluble

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enzymes without the use of detergents. Whether these Nirs are truly soluble or attached to the cytoplasmic membrane via lipidation modifications such as seen for other halophile exterior proteins remains an open question.

NITRIC OXIDE REDUCTASE Nors and cytochrome oxidases (CcO) belong to the superfamily of heme–copper oxidases and have most likely evolved from a common ancestor that might have been an anaerobic, nitric oxidereducing enzyme [2,45,46,66]. The archaic function of Nors might have been mainly to rid cells of the toxic nitric oxide rather than catalyzing part of the denitrification pathway from nitrate to dinitrogen. The detoxification function of Nor is still preserved in some pathogenic and marine, nondenitrifying bacteria. Interestingly, in addition to nitric oxide reduction, Nors also catalyze the reduction of oxygen, albeit at approximately 20% of the nitric oxide reduction rate. CcOs, in particular the cbb3-type oxidases, are capable of reducing nitric oxide, though slowly [67]. These findings strongly suggest that the active sites of Nors and CcOs are similar, which was confirmed experimentally. Presently, three different bacterial Nors have been characterized, cNor, qNor, and qCuANor [47,54,56,57]. Only the qNor-type enzyme seems to occur in archaea and was recently purified from P. aerophilum. The purified qNor is an integral membrane protein consisting of a single subunit displaying MQH2: NO oxidoreductase (or qNor) activity [47]. The enzyme contains heme-iron and nonheme iron in a 2:1 stoichiometry (Figure 9.6). One of the hemes is low spin and involved in electron transfer from the MQH2, the other is high spin. In addition to two heme centers, the enzyme contains one nonheme iron center (FeB), which is the functional equivalent of CuB in CcOs. The FeB center and the high-spin heme form a functional binuclear iron–iron center where nitric oxide reduction occurs [2,68,69]. The enzyme activity is inhibited by unphysiologically high nitric oxide concentrations (Ki = 7 μM). However, this phenomenon of nitric oxide substrate inhibition is common to all Nors studied so far and is due to accumulation of an inactive heme ferric–nitrosyl intermediate. The steady-state nitric oxide reduction kinetics indicate a broad pH optimum between pH 7 and 9. The qNOR from P. aerophilum is thermostable with a half-life of 86 min at 100°C. Unlike cNor and qCuANor, the P. aerophilum qNor does not contain heme b. One heme is heme Op1, an ethenylgeranylgeranyl derivative of heme b, the other Op2, containing the hydroxyethylgeranylgeranyl modification (Figure 9.6) [47,70]. Thus far, the archaeal qNOR is the only example of a Nor containing modified hemes reminiscent of cytochrome bo3 and aa3 oxidases. The presence of the modified hemes allows extra hydrophobic interactions, which might contribute to the thermostability of the qNOR. Other factors increasing thermostability might be the relatively high content of branched-chain amino acids, the low cysteine content, and the smaller subunit size compared with proteobacterial qNors [ 47,71].

NITROUS OXIDE REDUCTASE Thus far, NosZ has not been purified from any thermophilic aquificales or archaea, only from mesophilic proteobacteria. The nosZ gene identified in the genome of H. marismortui contains all of the His, Cys, and Met residues required to serve as ligands for CuA and CuZ. CuA consists of two copper atoms 2.5Å apart, which are coordinated by two Cys, two His, one Glu and Met [2,16, 72–75]. CuA is located in the very C-terminal portion of the peptide and is the direct electron donor to CuZ. The CuZ center contains four Cu-atoms ligated by seven His and one inorganic sulfur atom. Reduction of nitrous oxide to nitrogen occurs at the CuZ site, which shuttles between the fully reduced (Cu1+)4 and the half-reduced (Cu1+–Cu2+)2 redox states during turnover [76]. While bacterial NosZ enzymes are soluble, we predict a membrane localization of this enzyme in thermophilic and other denitrifying archaea. Indeed, enzyme measurements in P. aerophilum linked Nos activity to the membrane [44].

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CONCLUDING REMARKS This overview indicates that our current insight in denitrifying processes in thermophilic bacteria and archaea is highly limited as compared with our in-depth knowledge of the pathway enzymes in denitrifying proteobacteria. This holds true for the physiology, regulation, molecular biology, and biochemistry of denitrification pathway enzymes. The currently available genomic and biochemical data display considerable sequence and functional homology between the pathway enzymes of all prokaryotes suggesting the possibility of lateral gene transfer. However, considerable diversity of the cellular organization of denitrification enzymes and electron mediators exists within different species affecting the bioenergetics of the cell and the protein transport and protein maturation machineries. The continued challenge in this field remains the isolation and characterization of thermophilic denitrifiers to gain more insight into enzyme variations and regulation of their genes an area that is virtually unexplored thus far. The intricate knowledge of thermophilic denitrifying microorganisms and their enzymes will aid in our understanding as to how this pathway may have evolved and how it contributes to the global nitrogen cycle in extreme environments present on this planet today.

ACKNOWLEDGMENT Imke Schröder was supported by NSF (MSB 0345037) and Simon de Vries by the Netherlands Organization for Scientific Research (NWO-700.54.003).

REFERENCES 1. Zumft, W. G., Cell biology and molecular basis of denitrification, Microbiol Mol Biol Rev 61, 533–616, 1997. 2. Wasser, I. M., de Vries, S., Moënne-Loccoz, P., Schröder, I., and Karlin, K. D., Nitric oxide in biological denitrification: Fe/Cu metalloenzyme and metal complex NO(x) redox chemistry, Chem Rev 102, 1201–34, 2002. 3. Richardson, D. J. and Watmough, N. J., Inorganic nitrogen metabolism in bacteria, Curr Opin Chem Biol 3, 207–19, 1999. 4. Philippot, L., Denitrifying genes in bacterial and archaeal genomes, Biochim Biophys Acta 1577, 355–76, 2002. 5. Cabello, P., Roldan, M. D., and Moreno-Vivian, C., Nitrate reduction and the nitrogen cycle in archaea, Microbiology 150, 3527–46, 2004. 6. Simon, J., Enzymology and bioenergetics of respiratory nitrite ammonification, FEMS Microbiol Rev 26, 285–309, 2002. 7. Richardson, D. J., Berks, B. C., Russell, D. A., Spiro, S., and Taylor, C. J., Functional, biochemical and genetic diversity of prokaryotic nitrate reductases, Cell Mol Life Sci 58, 165–78, 2001. 8. Rothery, R. A., Bertero, M. G., Cammack, R., Palak, M., Blasco, F., Strynadka, N. C., and Weiner, J. H., The catalytic subunit of Escherichia coli nitrate reductase A contains a novel [4Fe-4S] cluster with a high-spin ground state, Biochemistry 43, 5324–33, 2004. 9. Bertero, M. G., Rothery, R. A., Palak, M., Hou, C., Lim, D., Blasco, F., Weiner, J. H., and Strynadka, N. C., Insights into the respiratory electron transfer pathway from the structure of nitrate reductase A, Nat Struct Biol 10, 681–7, 2003. 10. Jormakka, M., Richardson, D., Byrne, B., and Iwata, S., Architecture of NarGH reveals a structural classification of Mo-bisMGD enzymes, Structure 12, 95–104, 2004. 11. Godden, J. W., Turley, S., Teller, D. C., Adman, E. T., Liu, M. Y., Payne, W. J., and LeGall, J., The 2.3 Å X-ray structure of nitrite reductase from Achromobacter cycloclastes, Science 253, 438–42, 1991. 12. Murphy, M. E., Turley, S., and Adman, E. T., Structure of nitrite bound to copper-containing nitrite reductase from Alcaligenes faecalis. Mechanistic implications, J Biol Chem 272, 28455–60, 1997.

9214_C009.indd 171

11/5/2007 7:24:31 AM

172

Thermophiles: Biology and Technology at High Temperatures

13. Bertero, M. G., Rothery, R. A., Boroumand, N., Palak, M., Blasco, F., Ginet, N., Weiner, J. H., and Strynadka, N. C., Structural and biochemical characterization of a quinol binding site of Escherichia coli nitrate reductase A, J Biol Chem 280, 14836–43, 2005. 14. Ellis, M. J., Dodd, F. E., Sawers, G., Eady, R. R., and Hasnain, S. S., Atomic resolution structures of native copper nitrite reductase from Alcaligenes xylosoxidans and the active site mutant Asp92Glu, J Mol Biol 328, 429–38, 2003. 15. Gordon, E. H., Sjögren, T., Löfqvist, M., Richter, C. D., Allen, J. W., Higham, C. W., Hajdu, J., Fülöp, V., and Ferguson, S. J., Structure and kinetic properties of Paracoccus pantotrophus cytochrome cd1 nitrite reductase with the d1 heme active site ligand tyrosine 25 replaced by serine, J Biol Chem 278, 11773–81, 2003. 16. Haltia, T., Brown, K., Tegoni, M., Cambillau, C., Saraste, M., Mattila, K., and Djinovic-Carugo, K., Crystal structure of nitrous oxide reductase from Paracoccus denitrificans at 1.6Å resolution, Biochem J 369, 77–88, 2003. 17. Stetter, K. O., Hyperthermophiles in the history of life, Ciba Found Symp 202, 1–10, 1996. 18. Huber, H., Hohn, M. J., Rachel, R., Fuchs, T., Wimmer, V. C., and Stetter, K. O., A new phylum of Archaea represented by a nanosized hyperthermophilic symbiont, Nature 417, 63–7, 2002. 19. Brochier, C. and Philippe, H., Phylogeny: a non-hyperthermophilic ancestor for bacteria, Nature 417, 244, 2002. 20. Philippot, L., Tracking nitrate reducers and denitrifiers in the environment, Biochem Soc Trans 33, 200–4, 2005. 21. Braker, G., Ayala-del-Rio, H. L., Devol, A. H., Fesefeldt, A., and Tiedje, J. M., Community structure of denitrifiers, bacteria, and archaea along redox gradients in Pacific Northwest marine sediments by terminal restriction fragment length polymorphism analysis of amplified nitrite reductase (nirS) and 16S rRNA genes, Appl Environ Microbiol 67, 1893–901, 2001. 22. Suharti and de Vries, S., Membrane-bound denitrification in the Gram-positive bacterium Bacillus azotoformans, Biochem Soc Trans 33, 130–3, 2005. 23. Yoon, J. H., Oh, T. K., and Park, Y. H., Transfer of Bacillus halodenitrificans Denariaz et al. 1989 to the genus Virgibacillus as Virgibacillus halodenitrificans comb. nov, Int J Syst Evol Microbiol 54, 2163–7, 2004. 24. Choi, I. G., Kim, S. S., Ryu, J. R., Han, Y. S., Bang, W. G., Kim, S. H., and Yu, Y. G., Random sequence analysis of genomic DNA of a hyperthermophile: Aquifex pyrophilus, Extremophiles 1, 125–34, 1997. 25. Burggraf, S., Olsen, G. J., Stetter, K. O., and Woese, C. R., A phylogenetic analysis of Aquifex pyrophilus, Syst Appl Microbiol 15, 352–6, 1992. 26. Sakurai, T., Nakashima, S., Kataoka, K., Seo, D., and Sakurai, N., Diverse NO reduction by Halomonas halodenitrifi cans nitric oxide reductase, Biochem Biophys Res Commun 333, 483–7, 2005. 27. Sorokin, D. Y., Lysenko, A. M., Mityushina, L. L., Tourova, T. P., Jones, B. E., Rainey, F. A., Robertson, L. A., and Kuenen, G. J., Thioalkalimicrobium aerophilum gen. nov., sp. nov. and Thioalkalimicrobium sibericum sp. nov., and Thioalkalivibrio versutus gen. nov., sp. nov., Thioalkalivibrio nitratis sp.nov., novel and Thioalkalivibrio denitrificancs sp. nov., novel obligately alkaliphilic and obligately chemolithoautotrophic sulfur-oxidizing bacteria from soda lakes, Int J Syst Evol Microbiol 51, 565–80, 2001. 28. Sorokin, D. Y., Kuenen, J. G., and Jetten, M. S., Denitrification at extremely high pH values by the alkaliphilic, obligately chemolithoautotrophic, sulfur-oxidizing bacterium Thioalkalivibrio denitrificans strain ALJD, Arch Microbiol 175, 94–101, 2001. 29. Kumaraswamy, R., van Dongen, U., Kuenen, J. G., Abma, W., van Loosdrecht, M. C., and Muyzer, G., Characterization of microbial communities removing nitrogen oxides from flue gas: the BioDeNOx process, Appl Environ Microbiol 71, 6345–52, 2005. 30. Vorholt, J. A., Hafenbradl, D., Stetter, K. O., and Thauer, R. K., Pathways of autotrophic CO2 fixation and of dissimilatory nitrate reduction to N2O in Ferroglobus placidus, Arch Microbiol 167, 19–23, 1997. 31. Vökl, P., Huber, R., Drobner, E., Rachel, R., Burggraf, S., Trincone, A., and Stetter, K. O., Pyrobaculum aerophilum sp. nov., a novel nitrate-reducing hyperthermophilic archaeum, Appl Environ Microbiol 59, 2918–26, 1993.

9214_C009.indd 172

11/5/2007 7:24:31 AM

Denitrification Pathway Enzymes of Thermophiles

173

32. Tindall, B. J., Tomlinson, G. A., and Hochstein, L. I., Transfer of Halobacterium denitrificans (Tomlinson, Jahnke, and Hochstein) to the genus Haloferax as Haloferax denitrificans comb. nov, Int J Syst Bacteriol 39, 359–60, 1989. 33. Lledo, B., Marhuenda-Egea, F. C., Martinez-Espinosa, R. M., and Bonete, M. J., Identification and transcriptional analysis of nitrate assimilation genes in the halophilic archaeon Haloferax mediterranei, Gene 361, 80–8, 2005. 34. Oren, A., Ginzburg, M., Ginzburg, B. Z., Hochstein, L. I., and Volcani, B. E., Haloarcula marismortui (Volcani) sp. nov., nom. rev., an extremely halophilic bacterium from the Dead Sea, Int J Syst Bacteriol 40, 209–10, 1990. 35. Baeseman, J. L., Smith, R. L., and Silverstein, J., Denitrification potential in stream sediments impacted by acid mine drainage: effects of pH, various electron donors, and iron, Microb Ecol 51, 232–41, 2006. 36. Fitz-Gibbon, S. T., Ladner, H., Kim, U. J., Stetter, K. O., Simon, M. I., and Miller, J. H., Genome sequence of the hyperthermophilic crenarchaeon Pyrobaculum aerophilum, Proc Natl Acad Sci USA 99, 984–9, 2002. 37. Ramirez-Arcos, S., Fernandez-Herrero, L. A., and Berenguer, J., A thermophilic nitrate reductase is responsible for the strain specific anaerobic growth of Thermus thermophilus HB8, Biochim Biophys Acta 1396, 215–27, 1998. 38. Kawarabayasi, Y., Hino, Y., Horikawa, H., Yamazaki, S., Haikawa, Y., Jin-no, K., Takahashi, M., Sekine, M., Baba, S., Ankai, A., Kosugi, H., Hosoyama, A., Fukui, S., Nagai, Y., Nishijima, K., Nakazawa, H., Takamiya, M., Masuda, S., Funahashi, T., Tanaka, T., Kudoh, Y., Yamazaki, J., Kushida, N., Oguchi, A., Kikuchi, H., and et al., Complete genome sequence of an aerobic hyper-thermophilic crenarchaeon, Aeropyrum pernix K1, DNA Res 6, 83–101, 145–52, 1999. 39. She, Q., Singh, R. K., Confalonieri, F., Zivanovic, Y., Allard, G., Awayez, M. J., Chan-Weiher, C. C., Clausen, I. G., Curtis, B. A., De Moors, A., Erauso, G., Fletcher, C., Gordon, P. M., Heikamp-de Jong, I., Jeffries, A. C., Kozera, C. J., Medina, N., Peng, X., Thi-Ngoc, H. P., Redder, P., Schenk, M. E., Theriault, C., Tolstrup, N., Charlebois, R. L., Doolittle, W. F., Duguet, M., Gaasterland, T., Garrett, R. A., Ragan, M. A., Sensen, C. W., and Van der Oost, J., The complete genome of the crenarchaeon Sulfolobus solfataricus P2, Proc Natl Acad Sci USA 98, 7835–40, 2001. 40. PohlSchröder, M., Hartmann, E., Hand, N. J., Dilks, K., and Haddad, A., Diversity and evolution of protein translocation, Annu Rev Microbiol 59, 91–111, 2005. 41. PohlSchröder, M., Gimenez, M. I., and Jarrell, K. F., Protein transport in archaea: sec and twin arginine translocation pathways, Curr Opin Microbiol 8, 713–19, 2005. 42. Lledo, B., Martinez-Espinosa, R. M., Marhuenda-Egea, F. C., and Bonete, M. J., Respiratory nitrate reductase from haloarchaeon Haloferax mediterranei: biochemical and genetic analysis, Biochim Biophys Acta 1674, 50–9, 2004. 43. Zafra, O., Cava, F., Blasco, F., Magalon, A., and Berenguer, J., Membrane-associated maturation of the heterotetrameric nitrate reductase of Thermus thermophilus, J Bacteriol 187(12), 3990–6, 2005. 44. de Vries, S. and Schröder, I., Comparison between the nitric oxide reductase family and its aerobic relatives, the cytochrome oxidases, Biochem Soc Trans 30, 662–7, 2002. 45. Hendriks, J., Oubrie, A., Castresana, J., Urbani, A., Gemeinhardt, S., and Saraste, M., Nitric oxide reductases in bacteria, Biochim Biophys Acta 1459(2–3), 266–73, 2000. 46. Hendriks, J., Gohlke, U., and Saraste, M., From NO to OO: nitric oxide and dioxygen in bacterial respiration, J Bioenerg Biomembr 30, 15–24, 1998. 47. de Vries, S., Strampraad, M. J., Lu, S., Moënne-Loccoz, P., and Schröder, I., Purification and characterization of the MQH2:NO oxidoreductase from the hyperthermophilic archaeon Pyrobaculum aerophilum, J Biol Chem 278, 35861–8, 2003. 48. Afshar, S., Kim, C., Monbouquette, H. G., and Schröder, I. I., Effect of tungstate on nitrate reduction by the hyperthermophilic archaeon Pyrobaculum aerophilum, Appl Environ Microbiol 64, 3004–8, 1998. 49. Wood, N. J., Alizadeh, T., Richardson, D. J., Ferguson, S. J., and Moir, J. W., Two domains of a dualfunction NarK protein are required for nitrate uptake, the first step of denitrification in Paracoccus pantotrophus, Mol Microbiol 44, 157–70, 2002. 50. Clegg, S., Yu, F., Griffiths, L., and Cole, J. A., The roles of the polytopic membrane proteins NarK, NarU and NirC in Escherichia coli K-12: two nitrate and three nitrite transporters, Mol Microbiol 44, 143–55, 2002.

9214_C009.indd 173

11/5/2007 7:24:31 AM

174

Thermophiles: Biology and Technology at High Temperatures

51. Yoshimatsu, K., Iwasaki, T., and Fujiwara, T., Sequence and electron paramagnetic resonance analyses of nitrate reductase NarGH from a denitrifying halophilic euryarchaeote Haloarcula marismortui, FEBS Lett 516, 145–50, 2002. 52. Inatomi, K. I. and Hochstein, L. I., The purification and properties of a copper nitrite reductase from Haloferax denitrificans, Curr Microbiol 32, 72–76, 1996. 53. Ichiki, H., Tanaka, Y., Mochizuki, K., Yoshimatsu, K., Sakurai, T., and Fujiwara, T., Purification, characterization, and genetic analysis of Cu-containing dissimilatory nitrite reductase from a denitrifying halophilic archaeon, Haloarcula marismortui, J Bacteriol 183, 4149–56, 2001. 54. Suharti, Strampraad, M. J., Schröder, I., and de Vries, S., A novel copper A containing menaquinol NO reductase from Bacillus azotoformans, Biochemistry 40, 2632–9, 2001. 55. Sakurai, N. and Sakurai, T., Isolation and characterization of nitric oxide reductase from Paracoccus halodenitrificans, Biochemistry 36, 13809–15, 1997. 56. Girsch, P. and de Vries, S., Purification and initial kinetic and spectroscopic characterization of NO reductase from Paracoccus denitrificans, Biochim Biophys Acta 1318, 202–16, 1997. 57. Cramm, R., Pohlmann, A., and Friedrich, B., Purification and characterization of the single-component nitric oxide reductase from Ralstonia eutropha H16, FEBS Lett 460, 6–10, 1999. 58. Hendriks, J. H., Jasaitis, A., Saraste, M., and Verkhovsky, M. I., Proton and electron pathways in the bacterial nitric oxide reductase, Biochemistry 41, 2331–40, 2002. 59. Suharti, Heering, H. A., and de Vries, S., NO reductase from Bacillus azotoformans is a bifunctional enzyme accepting electrons from menaquinol and a specific endogenous membrane-bound cytochrome c551, Biochemistry 43, 13487–95, 2004. 60. Martinez-Espinosa, R. M., Marhuenda-Egea, F. C., and Bonete, M. J., Assimilatory nitrate reductase from the haloarchaeon Haloferax mediterranei: purification and characterisation, FEMS Microbiol Lett 204, 381–5, 2001. 61. Ketchum, P. A., Denariaz, G., LeGall, J., and Payne, W. J., Menaquinol-nitrate oxidoreductase of Bacillus halodenitrificans, J Bacteriol 173, 2498–505, 1991. 62. Afshar, S., Johnson, E., de Vries, S., and Schröder, I., Properties of a thermostable nitrate reductase from the hyperthermophilic archaeon Pyrobaculum aerophilum, J Bacteriol 183, 5491–5, 2001. 63. Page, C. C., Moser, C. C., Chen, X., and Dutton, P. L., Natural engineering principles of electron tunnelling in biological oxidation-reduction, Nature 402, 47–52, 1999. 64. Yoshimatsu, K., Sakurai, T., and Fujiwara, T., Purification and characterization of dissimilatory nitrate reductase from a denitrifying halophilic archaeon, Haloarcula marismortui, FEBS Lett 470, 216–20, 2000. 65. Wijma, H. J., Canters, G. W., de Vries, S., and Verbeet, M. P., Bidirectional catalysis by copper-containing nitrite reductase, Biochemistry 43(32), 10467–74, 2004. 66. Richardson, D. J., Bacterial respiration: a flexible process for a changing environment, Microbiology 146, 551–71, 2000. 67. Giuffre, A., Stubauer, G., Sarti, P., Brunori, M., Zumft, W. G., Buse, G., and Soulimane, T., The heme-copper oxidases of Thermus thermophilus catalyze the reduction of nitric oxide: evolutionary implications, Proc Natl Acad Sci USA 96, 14718–23, 1999. 68. Moënne-Loccoz, P., Richter, O.-M. H., Huang, H. W., Wasser, I., Ghiladi, R. A., Karlin, K. D., and de Vries, S., Nitric oxide reductase from Paracoccus denitrificans contains an oxo-bridged heme/ non-heme diiron center, J Am Chem Soc 122, 9344–5, 2000. 69. Lu, S., Suharti, de Vries, S., and Moënne-Loccoz, P., Two CO molecules can bind concomitantly at the diiron site of NO reductase from Bacillus azotoformans, J Am Chem Soc 126, 15332–3, 2004. 70. Lübben, M. and Morand, K., Novel prenylated hemes as cofactors of cytochrome oxidases. Archaea have modified hemes A and O, J Biol Chem 269, 21473–9, 1994. 71. Kumar, S. and Nussinov, R., How do thermophilic proteins deal with heat? Cell Mol Life Sci 58, 1216–33, 2001. 72. Speno, H., Taheri, M. R., Sieburth, D., and Martin, C. T., Identification of essential amino acids within the proposed CuA binding site in subunit II of Cytochrome c oxidase, J Biol Chem 270, 25363–9, 1995. 73. Kroneck, P. M., Riester, J., Zumft, W. G., and Antholine, W. E., The copper site in nitrous oxide reductase, Biol Met 3, 103–9, 1990.

9214_C009.indd 174

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Denitrification Pathway Enzymes of Thermophiles

175

74. Charnock, J. M., Dreusch, A., Korner, H., Neese, F., Nelson, J., Kannt, A., Michel, H., Garner, C. D., Kroneck, P. M., and Zumft, W. G., Structural investigations of the CuA centre of nitrous oxide reductase from Pseudomonas stutzeri by site-directed mutagenesis and X-ray absorption spectroscopy, Eur J Biochem 267, 1368–81, 2000. 75. Blackburn, N. J., Barr, M. E., Woodruff, W. H., van der Oost, J., and de Vries, S., Metal-metal bonding in biology: EXAFS evidence for a 2.5Å copper–copper bond in the CuA center of cytochrome oxidase, Biochemistry 33(34), 10401–7, 1994. 76. Ghosh, S., Gorelsky, S. I., Chen, P., Cabrito, I., Moura, J. J., Moura, I., and Solomon, E. I., Activation of N2O reduction by the fully reduced μ4-sulfide bridged tetranuclear Cu Z cluster in nitrous oxide reductase, J Am Chem Soc 125(51), 15708–9, 2003.

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Part IV Genetics of Thermophiles

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DNA Stability and Repair Malcolm F. White and Dennis W. Grogan

CONTENTS Genetic Costs of Life at High Temperature ............................................................................... Preserving Secondary Structure of DNA in Vivo ...................................................................... Preserving Primary Structure of DNA in Vivo ......................................................................... Lessons from Genome Surveys ...................................................................................... Lessons from Genome Reduction ................................................................................... Links between DNA Repair and Transcription in Thermophiles .................................. Structural Studies of DNA Repair Enzymes from Thermophiles ............................................. Applications of Thermostable DNA Repair Enzymes ............................................................... References ..................................................................................................................................

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GENETIC COSTS OF LIFE AT HIGH TEMPERATURE The ability of certain bacteria and archaea to thrive in geothermal environments focuses attention on the means by which these prokaryotes preserve the integrity of their macromolecules at extremely high temperatures. The largest, and arguably most important, macromolecule present in all thermophiles is the single, circular DNA which serves as the genome, and which must be replicated completely and accurately before each cell division. This chapter summarizes mechanisms that help preserve the physical and genetic integrity of this DNA during growth at high temperature, and discusses some of the biological and technological implications of these mechanisms. Many DNA repair proteins discovered in mesophilic bacteria and eukaryotes have homologs in the extreme or “hyperthermophiles,” and these homologs may be presumed to play the corresponding roles at high temperature. Other DNA maintenance systems, however, seem to be absent from hyperthermophiles, or highly diverged in them, which raises questions about whether the corresponding functions are universally needed, or alternatively, can be performed by other proteins. Some of the threats to genome stability posed by high temperature can be demonstrated experimentally by heating duplex DNA in buffered solutions. For example, above a critical “melting” temperature, defined by properties of the solution and the DNA, the two complementary strands separate [1]. Other processes slowly degrade the covalent structure of duplex DNA; these include hydrolytic depurination (i.e., base loss), deamination, and backbone scission [2]. Nucleo-bases can also be altered by oxidation or covalent addition of reactive metabolites. All these processes occur spontaneously in mesophiles, but are accelerated tremendously at temperatures of 75°C to 100°C [3]. Thermophiles should also suffer additional types of DNA damage that occur commonly in cells and are not necessarily due to high temperature per se. These include the formation of doublestrand breaks during DNA replication and damage by environmental agents including toxic chemicals and ultraviolet (UV) light [4]. 179

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PRESERVING SECONDARY STRUCTURE OF DNA IN VIVO The parameters that determine the melting temperature of DNA in vitro suggest how strand separation can be prevented in vivo. These parameters include: (i) the concentration of dissolved salts or other ionic solutes, (ii) the mole% G + C of the DNA, and (iii), torsional constraints on the two DNA strands [1,5]. Most of the available evidence suggests that thermophiles depend heavily upon variations of strategy (i), that is, “extrinsic stabilization,” to keep the DNA double-stranded at normal growth temperatures. Some thermophiles, including methanogens and thermococci, have extremely high intracellular potassium concentrations [6]. Alternatively, bacteria of the genus Thermus, and thermophilic archaea, contain various novel polyamines [7], which are much more effective on a molar basis at increasing Tm than inorganic cations are. Finally, several thermophilic archaea have been shown to contain high levels of small, basic proteins having a high affinity for double-stranded DNA. The structures of several of these proteins and their effect on doublestranded DNA have been studied in some detail (see Chapter 17 by K. Sandman). It should be noted, however, that the biological roles of these ions and proteins probably extend well beyond thermal stabilization of duplex DNA. With respect to strategy (ii), the weight of the evidence argues against a general, intrinsic stabilization of thermophile DNA by high G + C content. Although some thermophilic bacteria indeed have high G + C contents, most do not, and in general, there is no correlation between optimal growth temperature (Topt ) and genomic mol% G + C among bacteria and archaea [8,9]. With respect to strategy (iii), positive supercoiling has been proposed to stabilize duplex DNA in vivo at extremely high temperature. Evidence for this includes the presence of genes encoding a reverse DNA gyrase (Rgy) in genomes of all bacteria and archaea growing optimally above about 75°C, absence of these genes from all other prokaryotic genomes [10], and the observation that the Tm of positively supercoiled DNA can be as high as 107°C [5]. However, negative supercoiling proves to be equally effective in stabilizing the duplex form of DNA in vitro [5]. In addition, bacterial thermophiles encode “normal” DNA gyrases in addition to Rgy homologs and maintain negatively supercoiled DNA in vivo [11]. Finally the rgy gene of Thermococcus kodakarensis has been shown to be nonessential for basic cell viability by deletion from the chromosome [12]. We stress, however, that these results do not exclude all thermal stabilization of the DNA duplex by G + C content or positive supercoiling. The Δrgy mutant of T. kodakarensis, for example, has a decreased range of growth temperature [12], suggesting that the enzyme does contribute to proper cellular function. Rather, it seems that mechanisms corresponding to strategies (ii) and (iii) cannot be considered single, decisive factors that make life possible at geothermal temperatures.

PRESERVING PRIMARY STRUCTURE OF DNA IN VIVO Spontaneous chemical (i.e., covalent) damage to DNA occurs at physiological temperatures of all cells, and poses a threat to their successful reproduction. By chemical standards, however, the rates are low and the damage difficult to detect. Thus, the acceleration of these reactions at high temperatures was used historically to estimate, by extrapolation, their significance for “normal” (i.e., mesophilic) cells [3,13]. For bacteria and archaea from geothermal environments, little extrapolation from the original measurements is needed and attention turns to questions such as (i) whether the rates of spontaneous damage are much lower in vivo than measured in vitro; (ii) what enzyme systems deal with the damage that does occur; and (iii) how these systems compare with those of the mesophilic bacteria and archaea. Question (i) concerns “passive stabilization” of DNA against covalent damage. Examples of this phenomenon have been reported for mesophiles; for example, high salt concentrations and small DNA-binding proteins have been shown to decrease the rates of strand breakage by hydrolysis and ionizing radiation, respectively [5,14]. Similarly, the ability of DNA in bacterial endospores to survive extreme heat is due to small, basic proteins which complex closely with the DNA [15]. The

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maximal protection (i.e., rate decrease) by either strategy seems limited to about 30-fold, however, which seems modest relative to the predicted 1000- to 10,000-fold acceleration of decomposition reactions at 80°C to 100°C [3,13]. Furthermore, ionizing radiation has been shown to generate the same number of double-strand breaks in two Pyrococcus species as are formed in Escherichia coli [16] suggesting that passive protection by DNA-binding proteins is not a major line of defense against strand breaks in vivo. Addressing question (ii), that of the biologically significant modes of “active” DNA repair in extremely thermophilic bacteria and archaea, requires an appreciation of the complexity of DNA repair in mesophilic microorganisms. A long history of experimental studies has identified at least eight biochemically distinct strategies for dealing with various forms of DNA damage (Figure 10.1). These major pathways include two that simply reverse the damage [alkyl transfer (AT) and photoreactivation (PR)], three that remove and resynthesize the affected DNA [base excision repair (BER), nucleotide excision repair (NER), and mismatch repair (MMR)], and three that allow genome replication to continue despite failure of the previous pathways to repair the lesion [double-strand break repair (DSBR), trans-lesion synthesis (TLS), and nonhomologous end-joining (NHEJ)] [17]. It should be noted that DSBR and TLS can be considered “damage-tolerance” mechanisms, and that TLS and NHEJ can be considered strategies of last resort, because they tend to create mutations [18].

LESSONS FROM GENOME SURVEYS Proteins mediating the same repair function in different organisms often share characteristic, conserved sequence motifs, which facilitates identification of their homologs in complete genome sequences of diverse bacteria and archaea [19]. Thus, four of the eight major pathways represented in Figure 10.1 can be found generally in thermophilic bacteria and archaea: AT, BER, DSBR, and TLS. A fifth pathway, PR, occurs more sporadically, primarily among aerobic species. Evidence of function has been documented for all these pathways in at least a few species of thermophiles [20,21]. The remaining three pathways are generally not evident in the hyperthermophilic archaea. For example, genes encoding potential NHEJ proteins have so far been identified only in Archaeoglobus fulgidus [18]. This rare occurrence seems more enigmatic than the limited distribution of of PR genes; PR is expected to benefit only those species exposed both to UV and to visible light, whereas NHEJ, which acts to rejoin the ends of double-stranded breaks without the aid of an intact copy of the broken sequence, would seem to have survival value for any microorganism, regardless of its environment.

FIGURE 10.1 Eight major DNA repair pathways. For a systematic and authoritative review, see Friedberg et al., DNA Rapair and Mutagenesis. ASM Press, Washington, D.C., 1995.

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Analogous questions apply to the MMR and NER pathways. MMR accounts for much of the accuracy of DNA replication in cellular organisms, and requires homologs of two E. coli proteins, MutS and MutL. These E. coli proteins (and their homologs in other organisms) cooperate to remove sections of the newly synthesized “daughter” strand that contains the mismatch, and appear to be universal among mesophilic and moderately thermophilic bacteria and mesophilic archaea. Paired MutS and MutL homologs are absent from thermophilic archaea, however, as judged from the entire genome sequences [21]. One of these archaea, Pyrobaculum aerophilum, has been reported to have extremely frequent spontaneous mutation at simple repetitive sequences between genes [22]. In contrast, a distant relative, Sulfolobus acidocaldarius, has been shown to have a remarkably low rate of spontaneous mutation at two biosynthetic genes [23]. Thus it remains unclear whether all hyperthermophilic archaea exhibit a high level of DNA replication fidelity, and whether S. acidocaldarius has alternative strategies that compensate for the lack of MutS and MutL homologs in this group. Another genetic function of these MMR proteins, the suppression of homologous recombination between nonidentical DNA sequences [24], has not been evaluated in hyperthermophilic archaea, but would also be relevant to genome stability. Evaluating the NER capabilities of hyperthermophilic archaea has been somewhat more complicated than evaluating other DNA repair functions in these organisms. This is due in part to the fact that bacterial and eukaryotic NER systems involve rather different sets of proteins, and that mesophilic archaea encode homologs of both sets. Most archaeal genomes encode a subset of the proteins used in eukarya to carry out NER, including the helicases XPB and XPD, and nucleases XPF and Fen1/XPG [25]. Structural studies of archaeal XPF (also known as Hef) [26–28] and XPB [29] have yielded important information relevant to the function of the homologous eukaryal proteins. Nevertheless it remains unclear whether these proteins cooperate to catalyze NER in archaea in vivo. One problem is that all of these proteins have alternative roles in DNA replication, repair and transcription, and the lack of strict co-conservation of the four proteins argues against the existence of an archaeal NER machine homologous to that found in eukarya. No one has yet succeeded in demonstrating that a NER-type patch repair mechanism actually exists in archaea. Another puzzle concerns the lack of obvious DNA damage detection proteins (homologs of eukaryal XPA and XPC) in archaea, though these are also absent from plants. Thus, a lesson learned from searching the genomes of thermophiles for homologs of known DNA repair genes has been that certain repair systems may be missing or highly diverged, particularly among the hyperthermophilic archaea. It seems significant that the “missing” repair systems are widely conserved among mesophiles; furthermore, cells that grow at high temperature would seem, logically, to need greater DNA repair capacity than those that grow at low temperature. The situation has thus prompted the hypotheses that the thermophiles in question have either replaced the conventional repair systems with functional alternatives, or preserved ancestral repair pathways that were discarded during evolution by mesophiles. Although testing these hypotheses remains a challenge, they have important implications for understanding the molecular diversity of DNA repair mechanisms across biology.

LESSONS FROM GENOME REDUCTION The patterns of genome sizes within bacterial clades indicates that when a member of a free-living lineage adapts to an obligately parasitic life strategy, it begins a rather rapid loss of many DNA sequences [30]. The gene content of the resulting, highly reduced genome provides valuable insight as to what important cellular functions cannot be supplied by the host. This type of analysis has now been made possible for hyperthermophilic archaea by Stetter et al., who recovered an obligately symbiotic prokaryote from a submarine hydrothermal environment [31]. The symbiotic archaeon, named Nanoarchaeum equitans, occurs as extremely small cells attached to cells of another archaeon, an obligately anaerobic, S-reducing Ignicoccus species growing at 90°C [31]. With a cellular volume of only about 0.03 cubic microns, N. equitans represents one of the smallest cellular

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organisms ever cultured, and its genome of 491 kb is the smallest cellular genome yet sequenced. As the set of cellular functions that cannot be provided to N. equitans by the host are expected to include DNA repair, the physiological and genomic properties of this system provides a strategic perspective on DNA repair functions required in geothermal environments [32]. N. equitans encodes homologs of four proteins required for DSBR: the strand exchange protein RadA, Holliday junction resolving enzyme Hjc, Mre11, and Rad50, emphasizing the importance of this pathway which may primarily be used for the rescue of stalled or collapsed DNA replication forks [33]. There are also homologs of XPF, XPB, and Fen1, with a potential role in NER, although XPD is absent. The genome encodes one O 6 -methylguanosine cysteine methyltransferase (OGT) for direct reversal of methylation. The complement of BER enzymes in N. equitans comprises four proteins in total: a single helix-hairpin-helix superfamily glycoslyase (EndoIII or Nth), which may detect and remove a variety of damaged bases; one endonuclease IV (nfo in E. coli) which may function as the main nuclease for abasic sites in this organism; one endonuclease V (Nfi) for removal of deoxyinosine, which results from deamination of deoxyadenosine; and one family-4 uracil DNA glycosylase for removal of uracil, which results from deamination of cytosine or incorporation of dUTP. This cohort reflects the lifestyle of N. equitans, which as a hyperthermophile is expected to suffer high rates of hydrolytic deamination of bases, and nonenzymatic methylation of bases by S-adenosyl methionine. It is notable that there is no obvious glycosylase for removal of oxidized guanines (OGG1 or AGOG), which may reflect the anaerobic lifestyle of N. equitans. In contrast, the aerobic hyperthermophile Sulfolobus solfataricus and the aerobic mesophile Halobacterium marismortui encode 10 and 11 BER enzymes, respectively [34]. Thus, oxidative damage of DNA may pose as big a challenge to organisms as does growth at elevated temperatures.

LINKS BETWEEN DNA REPAIR AND TRANSCRIPTION IN THERMOPHILES Transcription and DNA repair are linked intimately through both transcriptional responses to DNA damage and the fact that many DNA lesions are repaired following an encounter with a transcribing RNA polymerase molecule (transcription coupled repair). In most bacteria, DNA repair proteins such as the strand exchange protein RecA and the exinuclease UvrABC are under the control of the LexA repressor. Under normal growth conditions, where levels of DNA damage are low, transcription of repair control genes is repressed. When DNA damage is encountered the LexA repressor is destroyed and transcription of a large number of repair proteins is induced—the so-called “SOS Response” [35]. As we have discussed already, thermophiles are expected to encounter increased levels of DNA damage, and it is thus pertinent to consider whether an inducible model for the control of DNA repair would be appropriate. There is a scarcity of published data on this topic, though genome sequences provide some clues. Thus, for example, the genome of the bacterial thermophile Thermus thermophilus does not encode a LexA homolog, and therefore presumably lacks an SOS response, whereas the closely related mesophile Deinococcus radiodurans does encode LexA [36]. The genome of the thermophile Aquifex aeolicus also lacks LexA [37], however, a LexA homolog is present in the thermophile T. maritima [38]. These observations do not rule out the possibility that DNA proteins are under the control of a different transcriptional apparatus in species lacking LexA. Further experimental studies are necessary to address this question. While we know very little about the transcriptional control of DNA damage genes in bacterial thermophiles, we know next to nothing about the equivalent processes in archaeal thermophiles. Archaea lack LexA and therefore an SOS response, and as we have already seen they have a very different complement of DNA repair enzymes too. From the limited data available it appears that RadA (the archaeal RecA homolog) is expressed constitutively in thermophiles [39], whereas it is inducible following DNA damage in the temperature mesophile Halobacterium [40]. Microarray studies indicate that the expression levels of DNA repair genes are not induced following UV radiation in S. solfataricus (D. Götz and M.F. White, unpublished).

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STRUCTURAL STUDIES OF DNA REPAIR ENZYMES FROM THERMOPHILES The conservation observed between archaeal DNA repair enzymes and their eukaryal counterparts has been a real boon for structural biology, as detailed in Chapter 15. At a structural level, even proteins with a highly diverged primary sequence can adopt very similar folds (Figure 10.2). Many human DNA repair proteins have proven difficult to express and/or crystallize, or exist as components of large protein complexes that are difficult to study. For many proteins involved in human DNA repair, the first structural insights have come from an archaeal homolog. Examples include RadA (Rad51) [41] and Rad50-Mre11 [42] from Pyrococcus furiosus, the NER nuclease XPF from P. furiosus and Aeropyrum pernix [25,26], the XPB helicase from A. fulgidus [28], and numerous others. Thus, the study of thermostable DNA repair proteins (particularly from archaea) has increased our molecular understanding of human proteins important for the avoidance and treatment of cancer.

APPLICATIONS OF THERMOSTABLE DNA REPAIR ENZYMES The advent of the polymerase chain reaction (PCR) technique, which relies on a thermostable DNA polymerase to amplify target DNA, heralded a revolution in molecular biology and has since impinged on many other areas of science and life in general. However, there has not subsequently been a flood of new applications taking advantage of the wealth of thermostable DNA repair enzymes revealed by genome sequencing. Rather, novel repair enzymes have been used to refine and modify the technique of PCR. One such technique is helicase-dependent amplification (HDA), which provides an alternative to classical PCR. In this technique, genes are amplified by the action of a DNA polymerase together with a DNA helicase to synthesize and separate double-stranded DNA, respectively. This obviates the need for temperature cycling to separate duplex DNA products, and raises the possibility for development of simple portable amplification devices [43]. HDA has been shown to work best when a thermostable helicase, UvrD from Thermoanaerobacter tengcongensis, is used, allowing the amplification to be carried out at a constant temperature of 60°C to 65°C [44]. The amplification and sequencing of ancient DNA samples has become an important tool for the emerging discipline of molecular paleontology. A major limitation of this approach is that DNA accumulates damage very quickly over a geological timescale, even under very favorable conditions.

FIGURE 10.2 Comparison of the core oligonucleotide-binding fold of the single-stranded DNA-binding protein from Sulfolobus solfataricus (left) and Homo sapiens (right). Despite very weak conservation of the primary sequences the protein structures are strongly conserved.

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Likewise, DNA isolated for forensic analysis may have accumulated significant damage in a relatively short period of time. Such damage prevents DNA amplification by the polymerases normally used for PCR. In cells, DNA lesions are bypassed by using specialized polymerases such as Dpo4, a Y-family polymerase that can replicate through a wide spectrum of damage types. By using a combination of Taq polymerase with a thermostable archaeal Dpo4 polymerase, Woodgate et al. succeeded in amplifying UV-irradiated DNA that was resistant to amplification by Taq polymerase alone [45]. Such an enzyme mixture may therefore improve the chances of obtaining useful information from forensic and ancient DNA samples. One further refinement of the PCR technique has been the inclusion of a thermostable dUTP glycosylase (dUTPase) to remove uracil from the pool of deoxynucleotides during PCR. dUTP arises from the deamination of dCTP, a reaction that increases with temperature. Deamination of cytosine to uracil in DNA can lead to unwanted transition mutations, and archaeal polymerases have evolved a “read-ahead” domain that scans for the presence of uracil in the template DNA strand, causing the polymerase to stall [46]. Reducing the amounts of uracil in the nucleotide pool with dUTPase results in higher yields and more processive polymerization by archaeal polymerases such as Pfu [47]. An alternative approach has involved the mutagenesis of the uracilbinding pocket in the read-ahead domain of the archaeal polymerase, which prevents stalling at uracil [48].

REFERENCES 1. Marmur, J. and Doty, P. Determination of the base composition of deoxyribonucleic acid from its thermal denaturation temperature. J. Mol. Biol., 5, 109, 1962. 2. Lindahl, T. Instability and decay of the primary structure of DNA. Nature, 362, 709, 1993. 3. Lindahl, T. and Nyberg, B. Heat-induced deamination of cytosine residues in deoxyribonucleic acid. Biochemistry, 13, 3405, 1974. 5. Marguet, E. and Forterre, P. DNA stability at temperatures typical for hyperthermophiles. Nucleic Acids Res., 22, 1681, 1994. 6. Adams, M.W. Enzymes and proteins from organisms that grow near and above 100 degrees C. Annu. Rev. Microbiol., 47, 627, 1993. 7. Terui, Y. et al. Stabilization of nucleic acids by unusual polyamines produced by an extreme thermophile, Thermus thermophilus. Biochem. J., 388, 427, 2005. 8. Galtier, N. and Lobry, J.R. Relationships between genomic G+C content, RNA secondary structures, and optimal growth temperature in prokaryotes. J. Mol. Evol., 44, 632, 1997. 9. Hurst, L.D. and Merchant, A.R. High guanine-cytosine content is not an adaptation to high temperature: a comparative analysis amongst prokaryotes. Proc. Biol. Sci., 268, 493, 2001. 10. Forterre, P. A hot story from comparative genomics: reverse gyrase is the only hyperthermophilespecific protein. Trends Genet., 18, 236, 2002. 11. Guipaud, O. et al. Both DNA gyrase and reverse gyrase are present in the hyperthermophilic bacterium Thermotoga maritima. Proc. Natl. Acad. Sci. USA, 94, 10606, 1997. 12. Atomi, H., Matsumi, R. and Imanaka, T. Reverse gyrase is not a prerequisite for hyperthermophilic life. J. Bacteriol., 186, 4829, 2004. 13. Lindahl, T. and Nyberg, B. Rate of depurination of native deoxyribonucleic acid. Biochemistry, 11, 3610, 1972. 14. Isabelle, V. et al. Radioprotection of DNA by a DNA-binding protein: MC1 chromosomal protein from the archaebacterium Methanosarcina sp. CHTI55. Int. J. Radiat. Biol., 63, 749, 1993. 15. Fairhead, H., Setlow, B., and Setlow, P. Prevention of DNA damage in spores and in vitro by small, acidsoluble proteins from Bacillus species. J. Bacteriol., 175, 1367, 1993. 16. Gerard, E., Jolivet, E., Prieur, D., and Forterre, P. DNA protection mechanisms are not involved in the radioresistance of the hyperthermophilic archaea Pyrococcus abyssi and P. furiosus. Mol. Genet. Genomics 266, 72, 2001. 17. Friedberg, E.C., Walker, G.C., and Siede, W. DNA Repair and Mutagenesis. ASM Press, Washington, D.C., 1995.

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18. Weterings, E. and van Gent, D.C. The mechanism of non-homologous end-joining: a synopsis of synapsis. DNA Repair (Amst), 3, 1425, 2004. 19. Eisen, J.A. and Hanawalt, P.C. A phylogenomic study of DNA repair genes, proteins, and processes. Mutat. Res., 435, 171, 1999. 20. White, M.F. Archaeal DNA repair: paradigms and puzzles. Biochem. Soc. Trans., 31, 690, 2003. 21. Grogan, D.W. Stability and repair of DNA in hyperthermophilic Archaea. Curr. Issues Mol. Biol., 6, 137, 2004. 22. Fitz-Gibbon, S.T. et al. Genome sequence of the hyperthermophilic crenarchaeon Pyrobaculum aerophilum. Proc. Natl. Acad. Sci. USA, 99, 984, 2002. 23. Grogan, D.W., Carver, G.T., and Drake, J.W. Genetic fidelity under harsh conditions: analysis of spontaneous mutation in the thermoacidophilic archaeon Sulfolobus acidocaldarius. Proc. Natl. Acad. Sci. USA, 98, 7928, 2001. 24. Rayssiguier, C., Thaler, D.S., and Radman, M. The barrier to recombination between Escherichia coli and Salmonella typhimurium is disrupted in mismatch-repair mutants. Nature, 342, 396, 1989. 25. Kelman, Z. and White, M.F. Archaeal DNA replication and repair. Curr. Opin. Microbiol., 8, 669, 2005. 26. Newman, M. et al. Structure of an XPF endonuclease with and without DNA suggests a model for substrate recognition. EMBO J., 24, 895, 2005. 27. Nishino, T., Komori, K., Ishino, Y., and Morikawa, K. X-ray and biochemical anatomy of an archaeal XPF/Rad1/Mus81 family nuclease: similarity between its endonuclease domain and restriction enzymes. Structure (Camb), 11, 445, 2003. 28. Nishino, T., Komori, K., Ishino, Y., and Morikawa, K. Structural and Functional Analyses of an Archaeal XPF/Rad1/Mus81 Nuclease: Asymmetric DNA Binding and Cleavage Mechanisms. Structure (Camb), 13, 1183, 2005. 29. Fan, L. et al. Conserved XPB core structure and motifs for DNA unwinding: implications for pathway selection of transcription or excision repair. Mol. Cell., 22, 27, 2006. 30. Moran, N.A. Microbial minimalism: genome reduction in bacterial pathogens. Cell, 108, 583, 2002. 31. Huber, H. et al. A new phylum of Archaea represented by a nanosized hyperthermophilic symbiont. Nature, 417, 63, 2002. 32. Waters, E. The genome of Nanoarchaeum equitans: insights into early archaeal evolution and derived parasitism. Proc. Natl. Acad. Sci. USA, 100, 12984, 2003. 33. McGlynn, P. Links between DNA replication and recombination in prokaryotes. Curr. Opin. Genet. Dev., 14, 107, 2004. 34. White, M.F. DNA repair. In: Archaea: Evolution, Physiology and Molecular Biology. Garrett, R.A. & Klenk, H.P. (eds). Blackwell Publishing, Oxford, 2007. 35. Radman, M. Phenomenology of an inducible mutagenic DNA repair pathway in Escherichia coli: SOS repair hypothesis. In: Molecular and Environmental Aspects of Mutagenesis. Charles C. Thomas, Springfield Ill, 1974. 36. Henne, A. et al. The genome sequence of the extreme thermophile Thermus thermophilus. Nat. Biotechnol., 22, 547, 2004. 37. Deckert, G. et al. The complete genome of the hyperthermophilic bacterium Aquifex aeolicus. Nature, 392, 353, 1998. 38. Nelson, K.E., Eisen, J.A., and Fraser, C.M. Genome of Thermotoga maritima MSB8. Methods Enzymol., 330, 169, 2001. 39. Reich, C.I. et al. Archaeal RecA homologues: different response to DNA-damaging agents in mesophilic and thermophilic Archaea. Extremophiles, 5, 265, 2001. 40. Baliga, N.S. et al. Systems level insights into the stress response to UV radiation in the halophilic archaeon Halobacterium NRC-1. Genome Res., 14, 1025, 2004. 41. Shin, D.S. et al. Full-length archaeal Rad51 structure and mutants: mechanisms for RAD51 assembly and control by BRCA2. EMBO J., 22, 4566, 2003. 42. Hopfner, K.P. et al. Structural biochemistry and interaction architecture of the DNA double-strand break repair Mre11 nuclease and Rad50-ATPase. Cell, 105, 473, 2001. 43. Vincent, M., Xu, Y., and Kong, H. Helicase-dependent isothermal DNA amplification. EMBO Rep., 5, 795, 2004.

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44. An, L. et al. Characterization of a thermostable UvrD helicase and its participation in helicase-dependent amplification. J. Biol. Chem., 280, 28952, 2005. 45. McDonald, J.P. et al. Novel thermostable Y-family polymerases: applications for the PCR amplification of damaged or ancient DNAs. Nucleic Acids Res., 34, 1102, 2006. 46. Greagg, M.A. et al. A read-ahead function in archaeal DNA polymerases detects promutagenic templatestrand uracil. Proc. Natl. Acad. Sci. USA, 96, 9045, 1999. 47. Hogrefe, H.H., Hansen, C.J., Scott, B.R., and Nielson, K.B. Archaeal dUTPase enhances PCR amplifications with archaeal DNA polymerases by preventing dUTP incorporation. Proc. Natl. Acad. Sci. USA, 99, 596, 2002. 48. Fogg, M.J., Pearl, L.H., and Connolly, B.A. Structural basis for uracil recognition by archaeal family B DNA polymerases. Nat. Struct. Biol., 9, 922, 2002.

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Plasmids and Cloning Vectors for Thermophilic Archaea Kenneth M. Stedman

CONTENTS Introduction ................................................................................................................................ Plasmids .......................................................................................................................... Viruses ............................................................................................................................ Genetic Systems .............................................................................................................. Vectors ............................................................................................................................ Conjugative Plasmids ................................................................................................................. Sulfolobus Plasmid pNOB8 ............................................................................................ pING Family of Sulfolobus Plasmids ............................................................................. Other Conjugative Plasmids ........................................................................................... Conjugative Plasmids as Vectors .................................................................................... Nonconjugative Plasmids ........................................................................................................... Sulfolobus Plasmids ........................................................................................................ Plasmids pRN1 and pRN2 from Sulfolobus islandicus ....................................... Virus Plasmid pSSVx ........................................................................................... Plasmids of the pRN Family ................................................................................ Pyrococcal Plasmids ....................................................................................................... Rolling Circle Plasmids ....................................................................................... Vectors from pGT5 .............................................................................................. Thermococcal Plasmid Screening ....................................................................... Other Plasmids ..................................................................................................... Viral Vectors .............................................................................................................................. Pyrococcus Virus-Like Particles ...................................................................................... Crenarchaeal Viruses ...................................................................................................... SSV1 .................................................................................................................... SSV1-Based Vectors ............................................................................................ Other Fuselloviruses ............................................................................................ Other Vectors ............................................................................................................................. Summary and Future Directions ............................................................................................... Dedication .................................................................................................................................. Acknowledgments ...................................................................................................................... References ..................................................................................................................................

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INTRODUCTION There have been very few genetic studies in thermophiles in general and even less in thermophilic archaea. Much of this has been due to a lack of effective and efficient vectors for expression and 189

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transformation in these organisms. However, there has been considerable if halting progress in the last few years. This review will discuss archaeal plasmids and viruses and emphasize their role as genetic tools for the study of thermophilic archaea. Genetic tools for Thermus (thermophilic bacteria) species are discussed in Chapter 12. Gene knockout technologies for Thermococcus kodakarensis and Sulfolobus solfataricus are discussed in Chapter 13.

PLASMIDS A number of plasmids have been isolated from thermophiles, particularly thermophilic archaea. Most of the plasmids were isolated with the express purpose of developing molecular genetic tools for these organisms (e.g., [1]). The plasmids range in size from 846 bp for the cryptic Thermotoga plasmid pRQ7 [2] to over 40 kbp for the Sulfolobus pNOB8 conjugative plasmid [3,4]. Many of these plasmids have been found to be integrated in whole genome sequences [5,6]. A number of the larger plasmids are conjugative. For a more detailed recent review of archaeal plasmids, see Garrett et al. [7].

VIRUSES Most viruses of thermophilic archaea have linear genomes and are difficult to manipulate genetically. However, fuselloviruses of Sulfolobus with ca. 15 kbp circular double-stranded DNA genomes that replicate as episomes can be treated as plasmids and have been used as vectors [8,9]. The PAV1, STIV and ATV viruses, and virus-like particles also have double-stranded circular DNA genomes but have not yet been used as vectors. For a detailed review of viruses of thermophilic archaea see Chapter 14. For a review of viruses of archaea in general see Stedman et al. [10].

GENETIC SYSTEMS Conjugation and virus infectivity are very attractive features for vectors for extremely thermophilic archaea as very few antibiotic markers are available [11]. There are a few published exceptions, including hygromycin resistance [12], alcohol dehydrogenase [13], and potentially bleomycin resistance [14]. A number of auxotrophic genetic markers have been developed in Sulfolobus acidocaldarius, S. solfataricus, Pyrococcus abyssi, and T. kodakarensis [15–18]. Based in part on these markers, gene knockout systems have recently been developed in both Sulfolobus and Thermococcus [19,20]. For a review of these breakthroughs see Chapter 13.

VECTORS A few vectors have been developed from these plasmids and the Sulfolobus fusellovirus SSV1; most of these include Escherichia coli origins of replication as well as origins for replication in thermophilic archaea. These vectors are detailed next. Unfortunately, these vectors have yet to be used on a widespread basis. The furthest developed are SSV1-based vectors for S. solfataricus which are now in their third generation [21–23]. Vectors have been used to complement mutant strains of Sulfolobus [22,24,25] and Pyrococcus [18], for overexpression of exogenous genes in Sulfolobus [23,26] and, very recently, for gene expression studies in Sulfolobus [23].

CONJUGATIVE PLASMIDS SULFOLOBUS PLASMID PNOB8 The first conjugative plasmid of any thermophilic archaeon, pNOB8, was discovered in a Japanese Sulfolobus isolate in a screen for viruses [3]. It was found to spread throughout a culture when donor cells containing the plasmid were mixed with a vast excess of recipient cells, generally 1:1000 to 1:10,000, which did not contain the plasmid. Transcipient cells containing the plasmid grow very slowly and the plasmid is not stable when propagated [3]. The most common genetic change was

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found to be an 8 kbp deletion between an 85 bp direct repeat. After this deletion the plasmid was rapidly lost from the culture [4]. The 41,229-bp genome was found to contain 50 open reading frames (ORFs) of which 85% had no detectable sequence similarity to the then known proteins. The most striking putative homologs were to bacterial conjugation proteins in the TrbE/VirB4 and TraG/ VirD4 families and three potential partitioning proteins, two ParB homologs and one ParA homolog [4]. The ParA and one of the ParB homolog are missing in the unstable deletion variant leading to the hypothesis that these proteins are critical for plasmid maintenance [4]. A schematic diagram of pNOB8 and other conjugative plasmids is shown in Figure 11.1. PING

FAMILY OF SULFOLOBUS PLASMIDS

A group of smaller (ca. 25 kbp), closely related conjugative plasmids was isolated from an Icelandic Sulfolobus strain by transformation into S. solfataricus strain P1 [27]. These plasmids contained

pN O

Region A: Conjugation?

B

d 33 8-

tio ele

n

pING1/p KEF

9d ele tio n

4

/VirD

TraG

ase plrA integ r

Tr

bE

NG

/V

de

irB

4

riv ativ es

pNOB8 41229 bp

ORFs rearranged in pARN family plasmids

Region B: Origin? Insertion F63 0a

OR

I ll p a Sm

Region C: Replication Initiation? Regulation?

FIGURE 11.1 Conjugative plasmid pNOB8 with other conjugative plasmids overlaid. The pNOB8 genome open-reading frame (ORF) map is shown [4]. ORFs are shown as open (not conserved) and filled (conserved) single-headed arrows in the direction of translation. The ORFs that encode the putative homologs of the bacterial conjugation proteins TraG/VirD4 and TrbE/VirD4 are labeled together with the plrA gene and an ORF encoding an integrase. ORF 630a in which the lacS gene was inserted to make the first recombinant plasmid for thermophilic archaea is labeled with the site of insertion shown with an arrow [24]. Regions identified by Greve et al. [29] to be important for conjugation (region A), putative origin of replication (region B), and putative regulatory regions (region C) are shown as thick lines with double-headed arrows for regions A and C. Narrower lines in region A with double-headed arrows indicate regions deleted in the pNOB8-33 deletion mutant [4], missing in the pING [28] and pKEF [29] plasmids, and rearranged in the pARN [29] family plasmids.

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66% similar ORFs to pNOB8, indicating that these ORFs are required for conjugation, pair formation, and plasmid transfer [28]. However, like the pNOB8 deletion variant, these plasmids were also genetically unstable, by both recombination with the host chromosome and deletion (Figure 11.1). Deletion variants generated the much smaller (6–7 kbp) plasmids pING2 and pING3 that were not able to transfer conjugatively by themselves but could be transferred together with a complete conjugative plasmid. This indicated that the origin of conjugative transfer and possibly genes required for plasmid mobility were present in the region encompassed by these smaller plasmids [28].

OTHER CONJUGATIVE PLASMIDS Extensive characterization and sequencing of conjugative plasmids from Iceland together with pNOB8 and pING plasmids showed that they belong to two mutually compatible families [27,29]. They all contain three conserved segments, one apparently involved in conjugative transfer, one the putative origin of replication, and the third encoding putative replication proteins [29] (Figure 11.1). In one case, two different conjugative plasmids were found in the same Sulfolobus isolate. These pSOG1 and pSOG2 plasmids have about one-third identical sequence but otherwise appear to belong to the two different Sulfolobus conjugative plasmid families. Plasmid pSOG1 replicates at a high copy number and is unstable on propagation in “foreign” hosts, such as S. solfataricus strain P1, but pSOG2 appears to be stable and replicates at a low copy number [27,30,31]. Strangely, no free conjugative plasmids have been found in thermophilic archaea other than S. “islandicus” and its close relatives. It is not clear whether this is due to a lack of search or a specific feature of these strains or their environment (80°C, pH = 3). However, there are a number of apparently conjugative or degraded conjugative plasmids found in genome sequences of a number of thermophilic archaea [5]. It has been shown conclusively that DNA can be transferred conjugatively between strains of S. acidocaldarius in the absence of detectable free plasmid [15]. The genome sequence of S. acidocaldarius revealed an integrated conjugative plasmid which might be involved in this gene transfer process [32].

CONJUGATIVE PLASMIDS AS VECTORS Due to the paucity of selectable markers for thermophilic archaea and relatively low transformation frequencies, a self-spreading vector such as a conjugative plasmid is very attractive. The first successful recombinant vector for any thermophilic archaeon was made from the pNOB8 plasmid by inserting the S. solfataricus broad-spectrum β-glycosidase [33] lacS gene with the S12 ribsosomal protein gene promoter into the plasmid genome at a unique SalI restriction endonuclease site. This site had previously been found not to be critical for plasmid function [24]. The plasmid was then transformed by electroporation into a mutant strain of S. solfataricus, PH1, that contained a disrupted lacS gene [9,16,24]. Enzyme activity was detected in transformants on plates, but single colony isolates were not obtainable. This was probably due to either growth inhibition of transformants or recombination with the host genome [24]. The SalI site in pNOB8 is in ORF630a, which was annotated as a possible cell division protein involved in membrane binding [4]. This ORF and the area surrounding it is not conserved in any of the other conjugative plasmids [29]. Together these data indicate that pNOB8 ORF630a is not required for conjugative plasmid function and that this unconserved region in the conjugative plasmids can tolerate insertions of foreign DNA. A few of the other conjugative plasmids show promise as vectors. The pSOG2 plasmid has low copy number in S. solfataricus and seems to be stably maintained [31]. The identification of conserved sequences that appear to be required for conjugation, replication, and maintenance (Figure 11.1) could be used to design a minimal conjugative plasmid [4,28,29]. The two different compatible families of plasmids might allow multi-plasmid experiments. Finally the small pING deletion variants might be usable as vectors in strains that contain integrated conjugative plasmids [6,28].

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NONCONJUGATIVE PLASMIDS SULFOLOBUS PLASMIDS Pllasmids pRN1 and pRN2 from Sulfolobus islandicus The cryptic plasmids pRN1 and pRN2 from S. “islandicus” REN1/H1 were the first plasmids of Sulfolobus to be identified together with their relative pHE7 (also known as pHEN7) [1] (Figure 11.2a). They were present in the same strain of Sulfolobus, which has hindered their development as vectors, but they have been recently separated by Werner Purschke and seem to replicate normally [34]. Both have been sequenced [35,36] and extensive work has been done by Georg Lipps et al. on the function of their ORFs (reviewed in Lipps [37]) (Figure 11.2a). The best understood of these is the “repA” gene that encodes a novel helicase/primase/polymerase [38–40]. There is also a highly conserved gene of unknown function in both plasmids, annotated as “plrA” whose product binds to DNA [41]. Less conserved is a gene generally found directly upstream of the “repA” gene which shows slight similarity to CopG, a copy number control gene known from other plasmids [42]. The protein from pRN1 has been shown to bind to a inverted repeat sequence upstream of the gene. Thereby it presumably regulates both its own gene and the repA gene, as would be expected for a protein-regulating plasmid copy number [42]. Little progress has been made in using these plasmids as vectors for Sulfolobus, most likely due to the lack of suitable selectable markers and the original presence of both in the same strain. Introduction of the pyrEF selectable markers into the pRN plasmids in S. solfataricus has not resulted in transformation [22]. Virus Plasmid pSSVx A small virus-like particle was observed in virus preparations of the novel SSV virus SSV2, that appeared to harbor a small satellite-virus-like DNA [43]. This plasmid was sequenced and found to be very similar to the already characterized pRN plasmids but contain two ORFs in its variable region that were similar—but not identical to—two ORFs of unknown function in all sequenced SSV genomes [43–45] (Figures 11.2a and 11.3). The acquisition of these genes apparently allows the plasmid to be packaged in an infectious virion allowing it to spread in the presence of a fulllength SSV genome [43]. This discovery was important for two reasons. First, it showed that pRN plasmids could accept different DNA in parts of their genome and remain viable. Second, that this could provide a means for the spread of a plasmid DNA in a culture in the absence of a selectable marker. However, this spread is only possible in the presence of a complete SSV genome, confounding genetic tool development. It is possible that an integrated copy of the virus genome would be sufficient for this purpose. SSVs integrate specifically into their host’s genome and are also present as episomes [8,46,47]. However, some large SSV1-based vectors appear to mostly be present as a single integrated copy (see the “Viral Vectors” section later in this chapter) [22]. These strains are attractive hosts for pSSVx-based vectors. A similar small plasmid has been found together with SSV3 and is under investigation (W. Zillig and Q. She, personal communication). Plasmids of the pRN Family The pHE7 plasmid was found at the same time that pRN1 and pRN2, together with a plasmid apparently identical to the pDL10 plasmid (also known as pSL10) previously isolated from Acidianus ambivalens (also known as Sulfolobus ambivalens and Desulfurolobus ambivalens) [48]. These plasmids were shown to be similar to each other by southern hybridization and sequencing [1,49,50] (Figure 11.2a). A number of pRN-type plasmids appear to have integrated into the S. solfataricus P2 genome and Sulfolobus tokodai genomes [5,6,51]. Whether the presence of an integrated form of the plasmid will cause problems for vectors based on the pRN plasmids is not clear. It does demonstrate that a number of different plasmids can exist in Sulfolobus species, including those with sequenced genomes (see Table 11.1).

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h

SSO /d s

e

e as lic

o

plrA

pRN1 5030 bp

Va ria ble

copG

e as

me o ly /p

Re gion

rase

repA

A)

(plr

pr im

ORFA

sso

(b)

F2 OR

Rem

pGT5 3444 bp

Rep75

Rem oved in pYS2

oved in pAG-plasm ids

dso

pCSV1

FIGURE 11.2 Schematic diagram of “cryptic” plasmids from Sulfolobus, Acidianus, and Pyrococcus with vectors developed. (a) pRN1 family plasmids. The pRN1 genomic map is shown [36]. Open-reading frames (ORFs) are shown as open arrows. The annotated genes shared in almost all pRN family plasmids, repA, copG, and plrA are marked. Plasmid pTAU4 has an MCM homolog instead of repA. Plasmid pTIK4 has a different repA primase/polymerase domain than the other pRN family plasmids. There is no copG gene in pDL10 or pST3 and in its place is a conserved “ORFA” shown as an inserted ORF. This conserved ORF is present in addition to copG in plasmids pSSVx, pST1, and pXQ1 [7]. The plrA gene is adjacent to copG in plasmids pDL10 and pHEN7 and absent in plasmid pORA1 and in the integrated plasmids pXQ1, pST1, and pST3. The plrA gene is embedded in the variable region in plasmids pTAU4 and pTIK4 [53]. The variable region is in a different location in pHEN7, pDL10, pTAU4, pORA1, and pTIK4 [49,50,53]. The putative singleand double-stranded origin is marked with SSO/dso. This region is in different locations in the genome in various plasmids. (b) Plasmids and vectors based on Pyrococcus abysii plasmid pGT5. The pGT5 plasmid genome map is shown [55]. The insertion point of pUC19 in the pCSV1 vector is shown with a closed arrow. The region deleted in the pAG1, 2, and 21, and the pYS2 plasmids is indicated with a stippled line [13,18]. This region is replaced with an Escherichia coli plasmid origin of replication and additional marker genes (see Table 11.2 for details).

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TABLE 11.1 “Cryptic” pRN-Family Plasmids of Sulfolobus Name

Size

repA

copG

plrA

pRN1 pRN2 pHE(N)7 pSSVx pDL10 pIT3 pXQ1

5030 6959 7830 5705 7598 4967 7472*

Y Y Y Y Y Y Y†

Y Y Y Y Y Y Y

Y Y Y Y Y Y N

pST1 pST3 pTAU4 pORA1 pTIK4

6706* 6854* 7192 9689 13638

Y Y Y N Y‡

Y N Y Y Y

N N N Y Y

* † ‡

Notes Same strain as pRN2 Same strain as pRN1 SSV2 satellite virus Acidianus plasmid In Sulfolobus solfataricus genome In Sulfolobus tokodai genome In Sulfolobus tokodai genome

Conjugative?

References [1,35] [1,36] [1,49] [43] [50] [52] [49] [6] [6] [53] [53] [53]

Integrated plasmid. Disrupted by insertion (IS) element. Only homologus in the C-terminal domain.

A new pRN family plasmid from an Italian Sulfolobus isolate was published very recently [52]. This is the smallest of the pRN family plasmids but shares the same basic structure and conserved ORFs (Figure 11.2a). The host of this plasmid appears to be a new strain of S. solfataricus and the plasmid can transform the plasmid-free strain S. solfataricus strain G-theta [52]. It is an exciting new tool for vector development. Recently three new plasmids from Sulfolobus isolates from New Zealand have been sequenced [53]. Interestingly, two contain the replication protein gene, repA similar to the pRN family of plasmids, but one of these lacks the primase/polymerase domain (Figure 11.2a). One, pTAU4, completely lacks a repA homolog but contains a homolog of the putative replicative helicase from Sulfolobus, MCM [53,54]. All of these large ORFs are preceded by a copG homolog, however not all contain a putative copG binding site. The largest plasmid, pTIK4, contains a gene similar to the one involved in conjugation in Sulfolobus plasmids. This indicates that there are multiple mechanisms for replication of these plasmids and calls into question the necessity of plrA and repA for replication and maintenance. It also indicates that multiple plasmids may be compatible for complex genetic experiments.

PYROCOCCAL PLASMIDS Rolling Circle Plasmids Relative to Sulfolobus, only a few nonconjugative plasmids have been isolated from other thermophilic archaea. Plasmids pGT5 [55] and pRT1 [56] have been isolated and characterized from two strains of Pyrococcus: Pyrococcus abysii and Pyrococcus strain JT1. The two plasmids have 41% identical nucleotide sequences, are similar in size, and both contain two large ORFs. The largest ORF encodes a protein with slight sequence similarity to replication proteins of other plasmids that replicate via a rolling circle mechanism [55,56]. The other large ORF in pRT1 is slightly similar to the conserved ORF80 or plrA of pRN family plasmids, which encodes a protein shown to have DNA binding activity and may play a role in plasmid replication [41]. Strangely, this protein is not similar at all to the second large ORF in pGT5 [5]. Single-stranded DNA was observed in the replication of both plasmids, indicating that they replicate by a rolling circle mechanism.

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Vectors from pGT5 When combined with the pUC19 plasmid, plasmid pGT5 was shown to be maintained after transformation in both Pyrococcus furiosus and S. acidocaldarius cells [57] (Table 11.2, Figure 11.2b). It appeared that ORF2 was not required for plasmid replication as the pUC19 plasmid was inserted between the promoter and start codon for ORF2 [57]. In later constructs most of the gene was deleted [13]. However, it was very hard to detect the combined vector in these cells and it was only detectable with polymerase chain reaction (PCR) or retransformation into E. coli. There was also vector instability detected in E. coli [13]. Therefore, the alcohol dehydrogenase gene from S. solfataricus was added to the plasmid along with the pBR322 Rom/Rop gene, which together appeared to stabilize the plasmid [13]. This plasmid did not function with P. abysii, a close relative of P. furiosus [18]. However, a modified plasmid containing the pyrE gene from S. acidocaldarius, pYS2, was able to complement P. abysii uracil auxotrophs, albeit with relatively low transformation efficiency (Figure 11.2b) [18]. Thermococcal Plasmid Screening In a screen of about 190 novel Thermococcus and Pyrococcus isolates from deep sea hydrothermal vents, about 40% were found to contain plasmids [58,59]. There were five different plasmid types from 3 to 24 kbp. One was shown to be very closely related to pGT5 by southern hybridization. A set of three strains contained both a small (3 kbp) and large (24 kbp) plasmid all closely related to each other [59]. Unfortunately, no plasmids have been reported to date that replicate in T. kodakarensis, the thermophilic archaeon for which the best developed gene-knockout tools are available (see Chapter 13). Other Plasmids Additional plasmids have been characterized from Archaeoglobus profundus, pGS5 (2.8 kbp) [60] and a very small plasmid (846 bp) from the thermophilic bacterium T. maritima pRQ7 [2]. Mostly they have been characterized for their supercoiling states [61]. Plasmid pRQ7 has been used by the Noll lab to generate a number of replicating plasmids for the transformation of Thermatoga [62]. Original isolates of Picrophilus oshimae were found to contain about 8 kbp

TABLE 11.2 Plasmid Vectors for Thermophilic Archaea Name

Archaeal Origin

pCSV1

pGT5

pAG1, 2 and 21

pGT5

pYS2 pEXSs poriC-hphT pKMSD48,54,55,59,60 pSSV64 pMJ02 pMJ03,05,05-sor, 11 pSVA6,9,15,31

pGT5 SSV1* S.so “oriC” SSV1 SSV1 SSV1 SSV1 SSV1

Host Pyrococcus furiosus, Sulfolobus acidocaldarius Pyrococcus furiosus, Sulfolobus acidocaldarius Pyrococcus abyssi Sulfolobus solfataricus Sulfolobus solfataricus Sulfolobus solfataricus Sulfolobus solfataricus Sulfolobus solfataricus Sulfolobus solfataricus Sulfolobus solfataricus

Marker/Reporter

References

—

[57]

adh

[13]

pyrE hph, adh, lacS hph — pyrEF lacS pyrEF, various pyrEF, various

[18] [12,25,71,74] [79] [21] [22] [22] [22,23] [23]

* Fraction of SSV1, see Figure 11.3.

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197

plasmids [63]. A plasmid pTA1 was found in some Thermoplasma isolates [64] and was recently sequenced [65]. The 15-kbp plasmid contains no genes with similarities to known proteins with the intriguing exception of a cdc6 homolog that appears to be involved in chromosome replication [66]. Each of these are promising plasmids for the development of genetic tools, but none have come to fruition.

VIRAL VECTORS PYROCOCCUS VIRUS-LIKE PARTICLES The P. abysii strain GE23, that had previously been characterized as containing a plasmid [59], was also found to produce a virus-like particle, PAV1. Purified particles appeared to contain this 18 kbp double-stranded circular DNA [67]. Unlike the SSV-like viruses, this DNA did not appear to integrate into the host genome and an uninfected host could not be found [67]. At the date of writing, the DNA sequence of PAV1 was yet to be made available. A number of other virus-like particles have been observed in enrichment cultures of Pyrococcus and Thermococcus but none have been characterized further [68].

CRENARCHAEAL VIRUSES A vast array of viruses from thermophilic crenarchaea with novel morphology and genomes have been discovered by Wolfram Zillig and his collaborators (Chapter 14 discusses these viruses in more detail). The genomes of these viruses are both linear and circular, and range in size from 15 to 75 kbp. Those of the viruses STIV and ATV, having double-stranded circular DNA genomes, are attractive potential vectors. The best studied, however, are from the Fusellovirus family, particularly the virus SSV1. SSV1 The virus SSV1 was first found as a ultraviolet (UV)-inducible plasmid in a Sulfolobus isolate from Beppu, Japan [69]. This strain, B12, also known as S. acidocaldarius and S. solfataricus before being renamed Sulfolobus shibatae [70], was found to produce a virus-like particle, SSV1 (known at the time as SAV1) [8]. The virus particle is about 60 × 90 nm with a short tail at one end. It has a double-stranded circular 15,465 bp DNA genome. This genome could be transferred into S. solfataricus by electroporation and it generated infectious virus. This was a major breakthrough in the development of molecular genetics of thermophilic archaea both for transformation of S. solfataricus and the conclusive demonstration that SSV1 was a virus [9]. SSV1-Based Vectors The first vector for thermophilic archaea to be made based on the SSV1 virus was the plasmid pEXSs, using a 1.7-kbp fragment of the SSV1 genome encompassing repeat structures and divergent promoters [71,72] (Table 11.2, Figure 11.3). This putative origin was combined with a selected mutant hygromycin transferase gene from E. coli and the pGEM5Zf plasmid. Transformed S. solfataricus strain G-theta was resistant to hygromycin and plasmid isolated from these strains was detected by retransformation into E. coli. Other researchers have had difficulties with this selection in other S. solfataricus strains, however. Nevertheless, the same vector and host system have been used to complement lacS mutants and express resistance to benzaldehyde [25,73]. Using a novel partial digestion and serial genetic selection technique, a shuttle vector, pKMSD48 (Figure 11.3), was made using the whole SSV1 genome and the pBluescript plasmid from E. coli [21]. This plasmid was shown to integrate into the host genome and also be inducible by UV irradiation. Critical for its use as a vector, it spreads through a culture in the absence of selection [21].

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ovi rus es

int att P

ns Co

in all F use ll

ed erv

p2 p3 p1

pKMSD48 pSSV64 pKMSD59/60 pKMSD54/55 pMJ vectors pSVA vectors

s

or

i

SSV1

pE

XS

FIGURE 11.3 SSV1 genome with vectors developed from it. The SSV1 genome with its open-reading frames (ORFs) is shown, together with mapped transcripts [72,81]. The attachment site in the viral integrase gene is shown as “attP” [82,83]. Known viral genes are labeled. VP1, VP3, and VP2 are virus structural genes [84]. The putative origin of replication used in plasmid pEXSs is labeled as “pEXSs ori” [71]. Insertion points for full-length shuttle vectors are shown with arrows outside the viral genome [21–23]. Dotted ORFs have been shown to be not essential for virus function [21,76]. Diagonally striped ORFs appear to be important for virus function [21]. Vertically striped ORFs are conserved in pSSVx [43]. The genes conserved in all fusellovirus genomes are indicated by a stippled curve [45].

Using the location of random insertion as a guide, four other S. solfataricus–E.coli shuttle vectors, were made [21] (Table 11.2, Figure 11.3). Recently this information was used together with uracil auxotrophic strains of Sulfolobus [17] and the pyrE and pyrF genes to generate the selectable plasmid pSSV64 that complemented these mutants [22] (Table 11.2, Figure 11.3). In parallel, plasmid pMJ02 was made with pUC18 and SSV1 together with the lacS gene under the control of the heat-inducible TF55 promoter [22]. This plasmid complemented the S. solfataricus lacS mutant PH1 and single colony transformants could be obtained. However, the plasmid was lost over long periods of growth [22]. Therefore, the plasmid pMJ03 was constructed by adding the pyrEF genes to pMJ02 [22]. The copy number of these plasmids was very low; often only the integrated copy of the genome was present [22]. Nevertheless, inducible expression of the lacS gene from a heterologous promoter was demonstrated [22]. Moreover, unlike the previously described pNOB8-based vector, stable single colony isolates containing the vector could be obtained. More recently, a third generation of vectors based on the entire SSV1 genome have been developed by Sonja Albers et al. [23]. These vectors use two different inducible promoters and have been used to produce three different proteins from Sulfolobus and its relative Acidianus [23]. Importantly, the vectors have modified promoter sequences to allow for more facile cloning and the host used is S. solfataricus strain P2, one of the most widely used strains and one for which the entire genome sequence is available [74]. Two of the expressed proteins contained peptide tags, which allowed purification and localization. This currently represents the best vector-host system for any thermophilic archaea.

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Other Fuselloviruses In addition to SSV1, six additional fusellovirus genomes have been completely sequenced [10,44,45,75]. These viruses are only about 55% identical at the nucleotide level and differ in their sites of integration and UV inducibility [44,45]. A Sulfolobus–E. coli shuttle vector has been made from one of them, SSV-K1. Only about half of the virus genome is conserved in all of the viruses [45] (Figure 11.3), so vectors should be constructible using a minimal fusellovirus genome and designed to integrate into various sites in the host genome. Methods to create specific mutations and deletions in whole SSV genomes have recently been developed [76] and should aid this process.

OTHER VECTORS A vector based on a mobile intron from Desulfurococcus mobilis was used to transform S. acidocaldarius together with fragments of E. coli DNA. The vector spread through a culture of S. acidocaldarius but further work was hampered by a lack of recombination when using larger fragments [77]. A plasmid, poriC-hphT, was constructed from a putative cellular Sulfolobus origin of replication, oriC, with the previously characterized thermally adapted hygromycin-phosphotransferase gene [12,78]. Strangely, the origin did not correspond to those identified by other authors [79,80]. The reasons for this discrepancy are unclear. However, the strains used in the two studies were different and growth conditions may also have influenced the results.

SUMMARY AND FUTURE DIRECTIONS A number of plasmids from euryarchaeal and crenarchaeal thermophiles have been isolated and characterized. Impressive developments have been made, mutants have been complemented, and genes have been overexpressed. The S. solfataricus–SSV1 vector-host system is in its third generation. However, there are still a few hurdles to be overcome before the use of these vectors for molecular cloning becomes commonplace. Some of the vectors, particularly those based on the entire SSV1 genome and the conjugative plasmids, are rather large (18–20 kbp or larger), complicating manipulation. Additionally, the SSV1-based vectors do not seem to tolerate large insertions without selection. Low transformation efficiency is also an issue, although it can be addressed by using infectious or conjugative vectors. Vector stability continues to be a major issue for many of these vectors, but selection does seem to help. The dependence on shuttle vectors that also replicate in E. coli may be a hindrance for vector development. Copy number control is not understood for any of these plasmids, but there has been some recent promising work on basic plasmid biology, which may address these issues [37]. Host strains should be standardized, particularly for gene knockout and complementation studies. For example, there are four S. solfataricus strains that are regularly used; P2, the sequenced strain; P1, a strain used widely by the Zillig laboratory; G-theta, a derivative of the MT4 strain used by Bartolucci et al.; and 98/2, the strain used by the Blum et al. to develop gene-knockouts. These strains appear to have slightly but potentially critically different growth characteristics complicating experimental replication and comparison. Finally, many researchers isolate and sequence new plasmids and viruses that should serve as the basis for future vectors, complementing further developments and refinements of existing vectors. It is easy for the molecular geneticist working with thermophilic archaea to be envious of genetic tools available in other organisms, but they should be proud of the progress made in a relatively short period of time by a relatively small number of researchers. With the tools available some exciting experiments can and have been done. This should only improve in the future.

DEDICATION This chapter is dedicated to the recently deceased Wolfram Zillig, the pioneer in the field of plasmids and viruses of thermophilic archaea.

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ACKNOWLEDGMENTS Research in the Stedman lab was supported by grants from the American Heart Association, 0460002Z, the National Science Foundation, MCB-0132156, and DBI-0352224, and Portland State University. The author would like to thank Dennis Grogan, Adam Clore, James Laidler, and Melissa DeYoung for critical reading of the manuscript. The author would also like to thank Hans Peter Arnold, Gael Erauso, and Qunxin She for sharing unpublished results.

REFERENCES 1. Zillig, W., Kletzin, A., Schleper, C., Holz, I., Janekovic, D., Hain, J., Lanzendoerfer, M., and Kristjansson, J. K., Screening for Sulfolobales, their plasmids and their viruses in Icelandic solfataras, Syst Appl Microbiol 16(4), 609–28, 1994. 2. Harriott, O. T., Huber, R., Stetter, K. O., Betts, P. W., and Noll, K. M., A cryptic miniplasmid from the hyperthermophilic bacterium Thermotoga sp. strain RQ7, J Bacteriol 176(9), 2759–62, 1994. 3. Schleper, C., Holz, I., Janekovic, D., Murphy, J., and Zillig, W., A multicopy plasmid of the extremely thermophilic archaeon Sulfolobus effects its transfer to recipients by mating, J Bacteriol 177(15), 4417–26, 1995. 4. She, Q. X., Phan, H. E., Garrett, R. A., Albers, S. V., Stedman, K. M., and Zillig, W., Genetic profile of pNOB8 from Sulfolobus—the first conjugative plasmid from an archaeon, Extremophiles 2(4), 417–25, 1998. 5. She, Q., Brugger, K., and Chen, L., Archaeal integrative genetic elements and their impact on genome evolution, Res Microbiol 153(6), 325–32, 2002. 6. She, Q., Shen, B., and Chen, L., Archaeal integrases and mechanisms of gene capture, Biochem Soc Trans 32(Pt 2), 222–6, 2004. 7. Garrett, R. A., Redder, P., Greve, B., Brugger, K., Chen, L., and She, Q., Archaeal plasmids, in Plasmid Biology, Funnell, B. E. and Phillips, G. J. ASM Press, Washington, D.C., 2004, pp. 377–92. 8. Martin, A., Yeats, S., Janekovic, D., Reiter, W. D., Aicher, W., and Zillig, W., SAV-1 a temperate UV inducible DNA virus-like particle from the archaebacterium Sulfolobus acidocaldarius isolate B-12, EMBO J 3(9), 2165–8, 1984. 9. Schleper, C., Kubo, K., and Zillig, W., The particle SSV1 from the extremely thermophilic archaeon Sulfolobus is a virus: demonstration of infectivity and of transfection with viral DNA, Proc Natl Acad Sci USA 89(16), 7645–9, 1992. 10. Stedman, K., Prangishvili, D., and Zillig, W., Viruses of Archaea, in The Bacteriophages, 2nd edn., Calendar, R. Oxford Press, New York, 2005, pp. 499–516. 11. Noll, K. M. and Vargas, M., Recent advances in genetic analyses of hyperthermophilic archaea and bacteria, Arch Microbiol 168(2), 73–80, 1997. 12. Cannio, R., Contursi, P., Rossi, M., and Bartolucci, S., Thermoadaptation of a mesophilic hygromycin B phosphotransferase by directed evolution in hyperthermophilic archaea: selection of a stable genetic marker for DNA transfer into Sulfolobus solfataricus, Extremophiles 5(3), 153–9, 2001. 13. Aravalli, R. N. and Garrett, R. A., Shuttle vectors for hyperthermophilic archaea, Extremophiles 1(4), 183–91, 1997. 14. Brouns, S. J., Wu, H., Akerboom, J., Turnbull, A. P., de Vos, W. M., and van der Oost, J., Engineering a selectable marker for hyperthermophiles, J Biol Chem 280(12), 11422–31, 2005. 15. Grogan, D. W., Exchange of genetic markers at extremely high temperatures in the archaeon Sulfolobus acidocaldarius, J Bacteriol 178(11), 3207–11, 1996. 16. Schleper, C., Roder, R., Singer, T., and Zillig, W., An insertion element of the extremely thermophilic archaeon Sulfolobus solfataricus transposes into the endogenous beta-galactosidase gene, Mol Gen Genet 243(1), 91–6, 1994. 17. Martusewitsch, E., Sensen, C., and Schleper, C., High spontaneous mutation rate in the hyperthermophilic archaeon Sulfolobus solfataricus is mediated by transposable elements., J Bacteriol 182, 2574–81, 2000. 18. Lucas, S., Toffin, L., Zivanovic, Y., Charlier, D., Moussard, H., Forterre, P., Prieur, D., and Erauso, G., Construction of a shuttle vector for, and spheroplast transformation of the hyperthermophilic archaeon Pyrococcus abyssi, Appl Environ Microbiol 68(11), 5528–36, 2002.

9214_C011.indd 200

11/5/2007 7:42:38 AM

Plasmids and Cloning Vectors for Thermophilic Archaea

201

19. Worthington, P., Hoang, V., Perez-Pomares, F., and Blum, P., Targeted disruption of the alpha-amylase gene in the hyperthermophilic archaeon Sulfolobus solfataricus, J Bacteriol 185(2), 482–8, 2003. 20. Sato, T., Fukui, T., Atomi, H., and Imanaka, T., Targeted gene disruption by homologous recombination in the hyperthermophilic archaeon Thermococcus kodakaraensis KOD1, J Bacteriol 185(1), 210–20, 2003. 21. Stedman, K., Schleper, C., Rumpf, E., and Zillig, W., Genetic requirements for the function of the archaeal virus SSV1 in Sulfolobus solfataricus: construction and testing of viral shuttle vectors, Genetics 152(4), 1397–405, 1999. 22. Jonuscheit, M., Martusewitsch, E., Stedman, K. M., and Schleper, C., A reporter gene system for the hyperthermophilic archaeon Sulfolobus solfataricus based on a selectable and integrative shuttle vector, Mol Microbiol 48(5), 1241–52, 2003. 23. Albers, S. V., Jonuscheit, M., Dinkelaker, S., Urich, T., Kletzin, A., Tampe, R., Driessen, A. J., and Schleper, C., Production of recombinant and tagged proteins in the hyperthermophilic archaeon Sulfolobus solfataricus, Appl Environ Microbiol 72(1), 102–11, 2006. 24. Elferink, M. G., Schleper, C., and Zillig, W., Transformation of the extremely thermoacidophilic archaeon Sulfolobus solfataricus via a self-spreading vector, FEMS Microbiol Lett 137(1), 31–5, 1996. 25. Bartolucci, S., Rossi, M., and Cannio, R., Characterization and functional complementation of a nonlethal deletion in the chromosome of a beta-glycosidase mutant of Sulfolobus solfataricus, J Bacteriol 185(13), 3948–57, 2003. 26. Brouns, S. J. J., Ettema, T. J. G., Stedman, K. M., Walther, J., Smidt, H., Snijders, A. P. L., Young, M., Bernander, R., Wright, P. C., Siebers, B., and van der Oost, J., The hyperthermophilic archaeon Sulfolobus—from exploration to exploitation, in Geothermal Biology and Geochemistry in Yellowstone National Park: Proceeding of the Thermal Biology Institute Workshop, Yellowstone National Park, Wy, October 2003, Inskeep, W. P. and McDermott, T. R. Montana State University Press, Bozeman, Montana, 2005. 27. Prangishvili, D., Albers, S. V., Holz, I., Arnold, H. P., Stedman, K., Klein, T., Singh, H., Hiort, J., Schweier, A., Kristjansson, J. K., and Zillig, W., Conjugation in archaea—frequent occurrence of conjugative plasmids in Sulfolobus, Plasmid 40(3), 190–202, 1998. 28. Stedman, K., She, Q., Phan, H., Holz, I., Singh, H., Prangishvili, D., Garrett, R., and Zillig, W., The pING family of conjugative plasmids from the extremely thermophilic archaeon Sulfolobus islandicus: insights into recombination and conjugation in crenarchaeota, J Bacteriol 182(24), 7014–20, 2000. 29. Greve, B., Jensen, S., Brugger, K., Zillig, W., and Garrett, R. A., Genomic comparison of archaeal conjugative plasmids from Sulfolobus, Archaea 1(4), 231–9, 2004. 30. Zillig, W., Arnold, H. P., Holz, I., Prangishvili, D., Schweier, A., Stedman, K., She, Q., Phan, H., Garrett, R., and Kristjansson, J. K., Genetic elements in the extremely thermophilic archaeon Sulfolobus, Extremophiles 2(3), 131–40, 1998. 31. Erauso, G., Stedman, K., van de Werken, H. J. G., Zillig, W., and van der Oost, J., Two novel conjugative plasmids from a single strain of Sulfolobus, Microbiology 152(7), 1951–68, 2006. 32. Chen, L., Brugger, K., Skovgaard, M., Redder, P., She, Q., Torarinsson, E., Greve, B., Awayez, M., Zibat, A., Klenk, H. P., and Garrett, R. A., The genome of Sulfolobus acidocaldarius, a model organism of the crenarchaeota, J Bacteriol 187(14), 4992–9, 2005. 33. Moracci, M., Ciaramella, M., and Rossi, M., Beta-glycosidase from Sulfolobus solfataricus, Methods Enzymol 330, 201–15, 2001. 34. Purschke, W. G. and Schafer, G., Independent replication of the plasmids pRN1 and pRN2 in the archaeon Sulfolobus islandicus, FEMS Microbiol Lett 200(1), 97–102, 2001. 35. Keeling, P. J., Klenk, H. P., Singh, R. K., Feeley, O., Schleper, C., Zillig, W., Doolittle, W. F., and Sensen, C. W., Complete nucleotide sequence of the Sulfolobus islandicus multicopy plasmid pRN1, Plasmid 35(2), 141–4, 1996. 36. Keeling, P. J., Klenk, H. P., Singh, R. K., Schenk, M. E., Sensen, C. W., Zillig, W., and Doolittle, W. F., Sulfolobus islandicus plasmids pRN1 and pRN2 share distant but common evolutionary ancestry, Extremophiles 2(4), 391–3, 1998. 37. Lipps, G., Plasmids and viruses of the thermoacidophilic crenarchaeote Sulfolobus, Extremophiles 10(1), 17–28, 2006. 38. Lipps, G., Rother, S., Hart, C., and Krauss, G., A novel type of replicative enzyme harbouring ATPase, primase and DNA polymerase activity, EMBO J 22(10), 2516–25, 2003.

9214_C011.indd 201

11/5/2007 7:42:38 AM

202

Thermophiles: Biology and Technology at High Temperatures

39. Lipps, G., Weinzierl, A. O., von Scheven, G., Buchen, C., and Cramer, P., Structure of a bifunctional DNA primase-polymerase, Nat Struct Mol Biol 11(2), 157–62, 2004. 40. Lipps, G., The replication protein of the Sulfolobus islandicus plasmid pRN1, Biochem Soc Trans 32(Pt 2), 240–4, 2004. 41. Lipps, G., Ibanez, P., Stroessenreuther, T., Hekimian, K., and Krauss, G., The protein ORF80 from the acidophilic and thermophilic archaeon Sulfolobus islandicus binds highly site-specifically to doublestranded DNA and represents a novel type of basic leucine zipper protein, Nucleic Acids Res 29(24), 4973–82, 2001. 42. Lipps, G., Stegert, M., and Krauss, G., Thermostable and site-specific DNA binding of the gene product ORF56 from the Sulfolobus islandicus plasmid pRN1, a putative archael plasmid copy control protein, Nucleic Acids Res 29(4), 904–13, 2001. 43. Arnold, H. P., She, Q., Phan, H., Stedman, K., Prangishvili, D., Holz, I., Kristjansson, J. K., Garrett, R. A., and Zillig, W., The genetic element pSSVx of the extremely thermophilic crenarchaeon Sulfolobus is a hybrid between a plasmid and a virus, Mol Microbiol 34(2), 217–26, 1999. 44. Stedman, K. M., She, Q., Phan, H., Arnold, H. P., Holz, I., Garrett, R. A., and Zillig, W., Relationships between fuselloviruses infecting the extremely thermophilic archaeon Sulfolobus: SSV1 and SSV2, Res Microbiol 154(4), 295–302, 2003. 45. Wiedenheft, B., Stedman, K., Roberto, F., Willits, D., Gleske, A. K., Zoeller, L., Snyder, J., Douglas, T., and Young, M., Comparative genomic analysis of hyperthermophilic archaeal fuselloviridae viruses, J Virol 78(4), 1954–61, 2004. 46. Reiter, W. D., Palm, P., and Yeats, S., Transfer RNA genes frequently serve as integration sites for prokaryotic genetic elements, Nucleic Acids Res 17(5), 1907–14, 1989. 47. Reiter, W. D. and Palm, P., Identification and characterization of a defective SSV1 genome integrated into a tRNA gene in the archaebacterium Sulfolobus sp. B12, Mol Gen Genet 221(1), 65–71, 1990. 48. Zillig, W., Yeats, S., Holz, I., Bock, A., Gropp, F., Rettenberger, M., and Lutz, S., Plasmid-related anaerobic autotrophy of the novel archaebacterium Sulfolobus ambivalens, Nature 313(6005), 789–91, 1985. 49. Peng, X., Holz, I., Zillig, W., Garrett, R. A., and She, Q., Evolution of the family of pRN plasmids and their integrase-mediated insertion into the chromosome of the crenarchaeon Sulfolobus solfataricus, J Mol Biol 303(4), 449–54, 2000. 50. Kletzin, A., Lieke, A., Urich, T., Charlebois, R. L., and Sensen, C. W., Molecular analysis of pDL10 from Acidianus ambivalens reveals a family of related plasmids from extremely thermophilic and acidophilic archaea, Genetics 152(4), 1307–14, 1999. 51. She, Q., Peng, X., Zillig, W., and Garrett, R. A., Gene capture in archaeal chromosomes, Nature 409(6819), 478, 2001. 52. Prato, S., Cannio, R., Klenk, H. P., Contursi, P., Rossi, M., and Bartolucci, S., pIT3, a cryptic plasmid isolated from the hyperthermophilic crenarchaeon Sulfolobus solfataricus IT3, Plasmid 56(1), 35–45, 2006. 53. Greve, B., Jensen, S., Phan, H., Brugger, K., Zillig, W., She, Q., and Garrett, R. A., Novel RepA-MCM proteins encoded in plasmids pTAU4, pORA1 and pTIK4 from Sulfolobus neozealandicus, Archaea 1(5), 319–25, 2005. 54. McGeoch, A. T., Trakselis, M. A., Laskey, R. A., and Bell, S. D., Organization of the archaeal MCM complex on DNA and implications for the helicase mechanism, Nat Struct Mol Biol 12(9), 756–62, 2005. 55. Erauso, G., Marsin, S., Benbouzid-Rollet, N., Baucher, M., Barbeyron, T., Zivanovic, Y., Prieur, D., and Forterre, P., Sequence of plasmid pGT5 from the archaeon Pyrococcus abyssi: evidence for rollingcircle replication in a hyperthermophile, J Bacteriol 178(11), 3232–7, 1996. 56. Ward, D. E., Revet, I. M., Nandakumar, R., Tuttle, J. H., de Vos, W. M., van der Oost, J., and DiRuggiero, J., Characterization of plasmid pRT1 from Pyrococcus sp. strain JT1, J Bacteriol 184(9), 2561–6, 2002. 57. Aagaard, C., Leviev, I., Aravalli, R. N., Forterre, P., Prieur, D., and Garrett, R. A., General vectors for archaeal hyperthermophiles: strategies based on a mobile intron and a plasmid., FEMS Microbiol Rev 18, 93–104, 1996. 58. Prieur, D., Erauso, G., Geslin, C., Lucas, S., Gaillard, M., Bidault, A., Mattenet, A. C., Rouault, K., Flament, D., Forterre, P., and Le Romancer, M., Genetic elements of Thermococcales, Biochem Soc Trans 32(Pt 2), 184–7, 2004.

9214_C011.indd 202

11/5/2007 7:42:38 AM

Plasmids and Cloning Vectors for Thermophilic Archaea

203

59. Benbouzid-Rollet, N., Lopez-Garcia, P., Watrin, L., Erauso, G., Prieur, D., and Forterre, P., Isolation of new plasmids from hyperthermophilic archaea of the order Thermococcales, Res Microbiol 148(9), 767–75, 1997. 60. Lopez-Garcia, P., Forterre, P., van der Oost, J., and Erauso, G., Plasmid pGS5 from the hyperthermophilic archaeon Archaeoglobus profundus is negatively supercoiled, J Bacteriol 182(17), 4998–5000, 2000. 61. Guipaud, O., Marguet, E., Noll, K. M., de la Tour, C. B., and Forterre, P., Both DNA gyrase and reverse gyrase are present in the hyperthermophilic bacterium Thermotoga maritima, Proc Natl Acad Sci USA 94(20), 10606–11, 1997. 62. Yu, J. S., Vargas, M., Mityas, C., and Noll, K. M., Liposome-mediated DNA uptake and transient expression in Thermotoga, Extremophiles 5(1), 53–60, 2001. 63. Schleper, C., Puehler, G., Holz, I., Gambacorta, A., Janekovic, D., Santarius, U., Klenk, H. P., and Zillig, W., Picrophilus gen. nov., fam. nov.: a novel aerobic, heterotrophic, thermoacidophilic genus and family comprising archaea capable of growth around pH 0, J Bacteriol 177(24), 7050–9, 1995. 64. Yasuda, M., Yamagishi, A., and Oshima, T., The plasmids found in isolates of the acidothermophilic archaebacterium Thermoplasma acidophilum, FEMS Microbiol Lett 128(2), 157–61, 1995. 65. Yamashiro, K., Yokobori, S. I., Oshima, T., and Yamagishi, A., Structural analysis of the plasmid pTA1 isolated from the thermoacidophilic archaeon Thermoplasma acidophilum, Extremophiles, 10(4), 327–35, 2006. 66. Robinson, N. P. and Bell, S. D., Origins of DNA replication in the three domains of life, FEBS J 272(15), 3757–66, 2005. 67. Geslin, C., Le Romancer, M., Erauso, G., Gaillard, M., Perrot, G., and Prieur, D., PAV1, the first viruslike particle isolated from a hyperthermophilic euryarchaeote, “Pyrococcus abyssi”, J Bacteriol 185(13), 3888–94, 2003. 68. Geslin, C., Le Romancer, M., Gaillard, M., Erauso, G., and Prieur, D., Observation of virus-like particles in high temperature enrichment cultures from deep-sea hydrothermal vents, Res Microbiol 154(4), 303–7, 2003. 69. Yeats, S., McWilliam, P., and Zillig, W., A plasmid in the archaebacterium Sulfolobus solfataricus, EMBO J 1(9), 1035–8, 1982. 70. Grogan, D., Palm, P., and Zillig, W., Isolate B12, which harbours a virus-like element, represents a new species of the archaebacterial genus Sulfolobus, Sulfolobus shibatae, sp. nov, Arch Microbiol 154(6), 594–9, 1990. 71. Cannio, R., Contursi, P., Rossi, M., and Bartolucci, S., An autonomously replicating transforming vector for Sulfolobus solfataricus, J Bacteriol 180(12), 3237–40, 1998. 72. Palm, P., Schleper, C., Grampp, B., Yeats, S., McWilliam, P., Reiter, W. D., and Zillig, W., Complete nucleotide sequence of the virus SSV1 of the archaebacterium Sulfolobus shibatae, Virology 185(1), 242–50, 1991. 73. Contursi, P., Cannio, R., Prato, S., Fiorentino, G., Rossi, M., and Bartolucci, S., Development of a genetic system for hyperthermophilic archaea: expression of a moderate thermophilic bacterial alcohol dehydrogenase gene in Sulfolobus solfataricus, FEMS Microbiol Lett 218(1), 115–20, 2003. 74. She, Q., Singh, R. K., Confalonieri, F., Zivanovic, Y., Allard, G., Awayez, M. J., Chan-Weihere, C. C.-Y., Clausen, I. G., Curtis, B. A., De Moors, A., Erauso, G., Fletcher, C., Gordon, P. M. K., Heikamp-de Jong, I., Jeffries, A. C., Kozera, C. J., Medina, N., Peng, X., Thi-Ngoc, H. P., Redder, P., Schenk, M. E., Theriault, C., Tolstrup, N., Charlebois, R. L., Doolittle, W. F., Duguet, M., Gaasterland, T., Garrett, R. A., Ragan, M. A., Sensen, C. W., and van der Oost, J., The complete genome of the crenarchaeon Sulfolobus solfataricus P2, Proc Natl Acad Sci USA 98(14), 7835–40, 2001. 75. Stedman, K. M., Clore, A., and Combet-Blanc, Y., Biogeographical diversity of archaeal viruses, in SGM Symposium 66: Prokaryotic Diversity: Mechanisms and Significance, Logan, N. A., Pappin-Scott, H. M., and Oynston, P. C. F. Cambridge University Press, Cambridge, 2006, pp. 131–44. 76. Clore, A. J. and Stedman, K., Long inverse PCR using iProof ® polymerase, Biorad Bull 5337, Rev A, 2005. 77. Aagaard, C., Dalgaard, J. Z., and Garrett, R. A., Intercellular mobility and homing of an archaeal rDNA intron confers a selective advantage over intron cells of Sulfolobus acidocaldarius, Proc Natl Acad Sci USA 92, 12285–9, 1995. 78. Contursi, P., Pisani, F. M., Grigoriev, A., Cannio, R., Bartolucci, S., and Rossi, M., Identification and autonomous replication capability of a chromosomal replication origin from the archaeon Sulfolobus solfataricus, Extremophiles 8(5), 385–91, 2004.

9214_C011.indd 203

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204

Thermophiles: Biology and Technology at High Temperatures

79. Robinson, N. P., Dionne, I., Lundgren, M., Marsh, V. L., Bernander, R., and Bell, S. D., Identification of two origins of replication in the single chromosome of the archaeon Sulfolobus solfataricus, Cell 116(1), 25–38, 2004. 80. Lundgren, M., Andersson, A., Chen, L., Nilsson, P., and Bernander, R., Three replication origins in Sulfolobus species: synchronous initiation of chromosome replication and asynchronous termination, Proc Natl Acad Sci USA 101(18), 7046–51, 2004. 81. Reiter, W. D., Palm, P., Yeats, S., and Zillig, W., Gene expression in archaebacteria physical mapping of constitutive and UV-inducible transcripts from the Sulfolobus virus-like particle SSV1, Mol Gen Genet 209(2), 270–5, 1987. 82. Muskhelishvili, G., The archaeal SSV integrase promotes intermolecular excisive recombination in vitro, Appl Microbiol 16, 605–8, 1994. 83. Muskhelishvili, G., Palm, P., and Zillig, W., SSV1-encoded site-specific recombination system in Sulfolobus shibatae, Mol Gen Genet 237(3), 334–42, 1993. 84. Reiter, W. D., Palm, P., Henschen, A., Lottspeich, F., Zillig, W., and Grampp, B., Identification and characterization of the genes encoding three structural proteins of the Sulfolobus virus-like particle SSV1, Mol Gen Genet 206(1), 144–53, 1987.

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Genetic Analysis in Extremely Thermophilic Bacteria: An Overview Dennis W. Grogan

CONTENTS Bacterial Hosts for Genetics at High Temperature .................................................................... Significance .................................................................................................................... Genetic Phenomena ........................................................................................................ Genetic Tools .................................................................................................................. Research Themes ....................................................................................................................... Thermo-Adaptive Mechanisms ...................................................................................... Biotechnology ................................................................................................................. Anaerobes ....................................................................................................................... References ..................................................................................................................................

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BACTERIAL HOSTS FOR GENETICS AT HIGH TEMPERATURE SIGNIFICANCE Although the unusual cellular properties of archaea from geothermal environments generate interest in genetic analysis, they simultaneously necessitate and hamper development of genetic techniques for these organisms. However, for a broad spectrum of questions regarding enzyme and cellular function at extremely high temperature, bacteria from geothermal environments offer practical alternatives for genetic analysis and manipulation. Over the past 20 years, researchers have successfully developed a number of genetic tools which take advantage of robust, heterotrophic growth of particular species, and the ability of certain bacterial genes and selections found in mesophilic bacteria to function, after limited modification, at high temperatures. As a result, several important techniques familiar to bacterial geneticists and molecular biologists can be used at temperatures up to 85ºC, enabling an increasing number of biological and biochemical aspects of extreme thermophiles to be investigated experimentally. Much of the progress has involved bacteria of the genus Thermus, first described by Brock and Freeze [1]. Thermus cells are gram-negative rods that occur in geothermal springs, hot-water heaters, and similar habitats; the G + C contents of their DNA are about 70 mol%. Most Thermus spp. grow heterotrophically and aerobically, with optimal pH values slightly above 7.0 and optimal temperatures in the range of 60°C to 80°C. Phylogenetically, the group represents a deep branch within the bacterial domain, most closely related to the deinococci [2]. Growth rates and growth yields are both relatively high; thus, these bacteria can be manipulated much as well-studied mesophilic 205

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bacteria are, except for the extremely high incubation temperatures and accompanying technical complications, such as solidifying the medium and preventing excessive evaporation. Within this genus, one of the most popular species for developing genetic methods has been Thermus thermophilus, represented by two isolates from Japanese hot springs, designated HB8 and HB27 [3]. Both isolates grow at higher temperatures than do most other Thermus strains (up to 85ºC) and are naturally competent for transformation. The complete genomes of both isolates have been sequenced, and reveal extensive similarity, consisting of a large circular chromsomome of about 1.9 Mbp and a large plasmid (or small chromosome) of 0.23 Mb [4,5]. A relatively large proportion of the open-reading frames (ORFs) show similarity to genes in other bacteria, which facilitates tentative gene identification. Complete biosynthetic pathways for all 20 amino acids are evident, as are catabolic enzymes supporting growth on a range of carbon sources [4]. The genomes appear to encode diverse extracellular hydrolases, as well as numerous active transport systems for amino acids and other solutes. The gene inventory thus reinforces a picture of nutritional versatility based on efficient scavenging of organic materials from the environment [4]. This heterotrophic lifestyle underlies a number of the selections and native genetic markers developed for T. thermophilus, such as those involving resistance to amino acid analogs [6] and complementation of auxotrophic mutants [7].

GENETIC PHENOMENA The selection provided by amino acid auxotrophs led to the demonstration by Koyama et al. [8] that T. thermophilus is naturally competent. Transformation occurs in normal growth medium and at very high frequencies [8]. The rate of DNA uptake is rapid, and not limited to Thermus sequences or to linear DNA [9]. Uptake requires an energized cell and the products of at least eight genes, most of which have sequences similar to type-IV pilus genes of mesophilic bacteria [10]. The correlation between lack of pili in the corresponding mutants and lack of competence provides strong evidence implicating these type-IV pili in the transformation mechanism [10], as has been demonstrated in mesophilic bacteria. In addition, T. thermophilus exhibits efficient recombination of homologous DNA following uptake. The high frequencies (0.1–10%) of prototrophic transformants generated by chromosomal DNA [8] imply correspondingly high rates of homologous recombination. Furthermore, transformation frequencies obtained with chromosomal markers exceed those observed for selectable plasmids [11,12], indicating that plasmids are at a slight disadvantage relative to linear chromosomal fragments in transformation. This idea was reinforced by the effect of first establishing the unmarked parental plasmid in a recipient before transforming it with a marked construct derived from the same plasmid. Such preintroduction of the plasmid increased transformation frequency 10- to 20-fold [13]. Beginning with Thermus aquaticus, various Thermus isolates have provided type II restriction endonucleases useful for analysis of DNA, cloning, and related manipulations of molecular biology [14,15]. Analysis of the HB27 and HB8 genomes indicates that these strains may encode other restriction enzymes in addition to those commercially available (http://rebase.neb.com). In principle, native restriction systems of T. thermophilus could complicate genetic manipulation, although this has not emerged as a serious concern. The relative ease of preparing vector and target DNAs from T. thermophilus strains, combined with efficient transformation, obviates complete dependence on Escherichia coli as a host for the production of cloned Thermus DNA, for example.

GENETIC TOOLS Antibiotic-resistance genes that function at the growth temperatures of Thermus spp. have contributed greatly to the past and current progress in Thermus genetics. These markers, including those encoding resistance to kanamycin, bleomycin, or hygromycin, originated from mesophilic bacteria and were modified by various strategies of mutation and selection (see the following section on

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Biotechnology). The thermostable resistance proteins represent examples of successfully applying artificial evolution to solve a practical problem, and reveal specific structural changes that elevate the thermostability of these enzymes. Another useful selection for Thermus genetics is provided by the pyrimidine analog 5-fluoro-orotic acid (FOA), which is not toxic per se, but kills cells when converted to the uridine monophosphate (UMP) analog. As a result, growth medium containing FOA plus uracil selects mutants lacking either of the last two enzymes of the de novo pathway of UMP biosynthesis, and are, accordingly, pyrimidine (i.e., uracil) auxotrophs. This selection has proven very versatile in yeast genetics, and in the development of basic genetics for thermophilic archaea (see Chapter 13 by Atomi and Imanaka). Although reports of its use in Thermus remain less common [16], this selection has considerable potential for the analysis of mutation at extremely high temperatures, the development of “reusable” selectable markers, and other situations in which two opposing selections are important. The addition of selectable markers to small, naturally occurring plasmids of Thermus spp. has led to construction of a variety of practical plasmid vectors. These include shuttle vectors, which allow Thermus DNA to be cloned and manipulated in E. coli, then transferrred to Thermus for functional analysis, or vice versa [11]. More specialized vectors include those designed to clone and identify origins of replication [17], measure promoter activity [18,19], and drive high-level expression of cloned genes [11,20,21]. Alternatively, constructs incapable of replication in Thermus facilitate integration of a selectable marker near or within a gene of interest by homologous recombination [22,23]. Examples of genetic constructs developed for Thermus spp. are listed Table 12.1.

RESEARCH THEMES THERMO-ADAPTIVE MECHANISMS Part of the value of genetic analysis of Thermus spp. is to enable functional molecular features of these bacteria to be compared with those of well-studied mesophiles such as E. coli or Bacillus subtilis, thereby identifying possible adaptations of bacterial cells to life at geothermal temperatures. Because these comparisons span tremendous phylogenetic distances, however, many molecular features of the thermophiles may represent “neutral” divergence not directly related to the specific demands of life at high temperature. Thus, interpreting molecular features as being thermoadaptive should remain provisional while the larger picture of cellular structure and function emerges. Accordingly, one of the most significant patterns arguing for a thermo-adaptive role would be conservation of a feature among extreme thermophiles of both bacterial and archaeal domains but not among the corresponding mesophiles, as has been observed for genes encoding reverse DNA gyrase [24]. Conversely, the deep evolutionary divergence separating extremely thermophilic bacteria from mesophilic “model species” implies that molecular features found in both groups are likely to be conserved widely among all bacteria. An enduring theme of thermo-adaptation research has been investigating the molecular basis of the intrinsic stability of individual enzymes from thermophiles, as examined in Chapters 2 and 3. However, genetic studies have also begun to address mechanisms which involve relatively complex interactions of distinct proteins, illustrated by DNA repair and related genetic processes. Mechanisms for coping with DNA damage occur in all cells, but have particular relevance for extreme thermophiles because of the accelerated spontaneous decomposition of DNA predicted at high growth temperatures, combined with the lack of any intrinsic stabilization of DNA against such damage (see Chapter 10 by White and Grogan). RecA/Rad51-type proteins represent one of the few truly ubiquitous systems for coping with DNA damage (via homologous recombination), and recA gene homologs have been identified in a number of thermophilic bacteria. The T. thermophilus protein has been examined in structural terms [25], and the gene has been disrupted in vivo [26]. The mutant exhibited sensitivity to DNA damage, but was not evaluated for a defect in genetic recombination.

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Thermus thermophilus lacks a LexA homolog [4], and ultraviolet [UV] treatment does not induce transcription of DNA repair genes [27]. It is also one of the first hyperthermophiles (defined here as organisms that grow optimally near or above 80ºC) that lacks a reverse DNA gyrase [4]. This enzyme, a type I topoisomerase, correlates strongly with optimal growth temperatures above 70ºC among other bacteria, as well as archea [24]. T. thermophilus also encodes various DNA N-glycosylases, but none of the Ung type [4].

BIOTECHNOLOGY Much of the research involving Thermus genetics aims at practical applications of bacterial metabolism. Investigations of amino acid biosynthesis, for example, have included selection of analogresistant mutants of T. thermophilus which overproduce the corresponding amino acid [6]. An interesting result of research on amino acid metabolism in T. thermophilus has been demonstration of the amino adipic acid pathway for biosynthesis of lysine [28,29], which was previously known only in fungi, and not in bacteria. Another question of broad interest is the development of thermostable enzymes for industrial catalysis. This reflects the advantages of thermostable enzymes for various industrial conversions, and the difficulty of introducing such thermostability into mesophilic industrial enzymes based on existing theory. Thus, researchers have investigated T. thermophilus and other thermophilic bacteria as a context in which proteins can be expressed at extremely high temperatures, and improved stability can be selected genetically. Among the first enzymes to be improved (stabilized) by this approach have been those which confer antibiotic resistance and thus provide for the selection of genetic constructs in these bacteria. A kanamycin nucleotidyl transferase gene, obtained from a Staphylococcus aureus resistance plasmid, was originally adapted to function in vivo at temperatures up to 60ºC by introduction of multiple point mutations [30]. This level of thermostability allows effective selection in T. thermophilus only at the low end of its temperature range, but this accordingly has set the stage for selecting variants of the gene with enhanced performance at higher temperatures. One approach has been to increase expression, by fusing the 5′ regions of highly expressed,

TABLE 12.1 Components of Genetic Tools for Extremely Thermophilic Bacteria Category Selectable genes

Reporter genes

Example KanR BleoR Hph β-galactosidase

Cloning vectors

β-glucosidase pTT8 pNHK101 pLU, pMY series

Expression vectors

pYK134 pT8L2P70

Plasmid replicons

pMKE1 pTEX series

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Properties [References] Encodes thermostable nucleotidyl transferase [11,18,31] Encodes thermostable bleomycin-binding protein [32] Encodes thermostable hygromycin B phosphotransferase [33] Thermus-derived, used with deletion mutant [42,43] Thermus-derived, used with deletion mutant [27,42] Cryptic plasmid of strain HB8; 9328 bp [44] Cryptic plasmid of strain TK10; 1564 bp [45] Selectable marker: KanR, scorable marker: celA (encodes thermostable cellulase) [11] Selectable markers: KanR, trpB [13] Selectable marker: pyrE [35] Selectable marker: KanR; promoter: nitrate reductase [20] Selectable marker: KanR; scorable marker: β-gal; three native promoters available [43]

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protein-encoding Thermus genes to the 5′ end of the KanR gene. Examples include the 5′ regions of the highly expressed surface (S-) layer gene [11] and the L32 ribosomal protein [18], both of T. thermophilus. Other groups have further elevated the thermostability and catalytic efficiency of the enzyme by introducing additional substitutions into the coding region of the gene itself [31]. Another selectable marker for extreme thermophiles was developed recently by modifying the bleomycin-resistance gene of Streptoalloteichus hindustanus, which encodes a small protein that binds stoichiometrically to bleomycin. Brouns et al. [32] mutated the native gene and selected Thermus transformants with elevated resistance to bleomycin. A number of point mutations were identified, and several were combined into one synthetic mutant gene, which conferred a BleoR phenotype in Thermus grown at 77ºC, and also in E. coli grown at 37ºC. Furthermore, the mutant protein that complexed with bleomycin was structurally stable up to 100ºC. A hygromycin-resistance (phosphotransferase) gene has similarly been adapted to function in Thermus by iterations of mutation and selection [33]. The “artificial evolution” strategy used in these studies takes advantage of the strong genetic selection for enhanced function of the original (mesophilic) resistance protein expressed in the thermophilic host. The same strategy can be applied to other proteins if appropriate selections can be set up. Most proteins modified by this approach have been metabolic enzymes, which reflects both the industrial interest in enzymatic processing at high temperatures, and the necessity of inactivating the corresponding gene of the host, which requires it to be nonessential. Examples of enzymes stabilized by genetic selection include disaccharide hydolases [34], and amino acid biosynthetic enzymes [16,35]. An important, long-term challenge for this strategy of thermo-stabilization is to create a selection (or reliable, high-throughput screen) for enzymatic activities of interest that do not occur in Thermus spp.

ANAEROBES Anaerobic thermophiles have also generated interest in genetic manipulation; examples include the genera Thermotoga and Thermoanaerobacterium. Although these and related bacteria remain more difficult to manipulate genetically, they offer potential advantages for applications where anaerobic metabolism is desirable, as in conversions of biomass into ethanol or hydrogen. Thermotoga neopolitana and Thermotoga maritima are heterotrophic marine bacteria that grow optimally around 80ºC and neutral pH. The genome of T. maritima has been sequenced to completion [36]. Noll et al. [37] have isolated various selectable mutants of these species, and a very small native plasmid shows potential as a cloning vector [38]. Thermoanaerobacterium saccharolyticum is another extremely thermophilic bacterium of interest for biomass conversion. T. saccharolyticum grows optimally at about 60ºC and pH 6.0 and ferments glucose, xylose, starch, and xylan [39]. Wiegel et al. [40] have developed shuttle vectors for this species which support the expression of metabolic genes of interest, as demonstrated with a cellobiose hydrolase. Others have successfully altered the composition of fermentation products from this organism through genetic inactivation of l-lactate dehydrogenase [41].

REFERENCES 1. Brock TD, Freeze H. Thermus aquaticus gen. n. and sp. n., a nonsporulating extreme thermophile. J Bacteriol, 98(1), 289–97, 1969. 2. Hensel R, Dembarter W, Kandler O, Kroppenstadt RM, Stackebrandt E. Chemotaxonomic and moleculargenetic studies of the genus Thermus: evidence for a phylogenetic relationship of Thermus aquaticus and Thermus ruber to the genus Deinococcus. Int J Syst Bacteriol, 36, 444–53, 1986. 3. Oshima T, Imahori K. Description of Thermus thermophilus (Yoshida and Oshima), comb. nov., a nonsporulating thermophilic bacterium from a Japanese thermal spa. Int J Syst Bacteriol, 24, 102–12, 1974. 4. Henne A, Bruggemann H, Raasch C, Wiezer A, Hartsch T, Liesegang H, et al. The genome sequence of the extreme thermophile Thermus thermophilus. Nat Biotechnol, 22(5), 547–53, 2004.

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5. Bruggemann H, Chen C. Comparative genomics of Thermus thermophilus: plasticity of the megaplasmid and its contribution to a thermophilic lifestyle. J Biotechnol, 124(4), 654–61, 2006. 6. Kosuge T, Hoshino T. Construction of a proline-producing mutant of the extremely thermophilic eubacterium Thermus thermophilus HB27. Appl Environ Microbiol, 64(11), 4328–32, 1998. 7. Koyama Y, Furukawa K. Cloning and sequence analysis of tryptophan synthetase genes of an extreme thermophile, Thermus thermophilus HB27: plasmid transfer from replica-plated Escherichia coli recombinant colonies to competent T. thermophilus cells. J Bacteriol, 172(6), 3490–5, 1990. 8. Koyama Y, Hoshino T, Tomizuka N, Furukawa K. Genetic transformation of the extreme thermophile Thermus thermophilus and of other Thermus spp. J Bacteriol, 166(1), 338–40, 1986. 9. Schwarzenlander C, Averhoff B. Characterization of DNA transport in the thermophilic bacterium Thermus thermophilus HB27. FEBS J, 273(18), 4210–18, 2006. 10. Friedrich A, Prust C, Hartsch T, Henne A, Averhoff B. Molecular analyses of the natural transformation machinery and identification of pilus structures in the extremely thermophilic bacterium Thermus thermophilus strain HB27. Appl Environ Microbiol, 68(2), 745–55, 2002. 11. Lasa I, de Grado M, de Pedro MA, Berenguer J. Development of Thermus-Escherichia shuttle vectors and their use for expression of the Clostridium thermocellum celA gene in Thermus thermophilus. J Bacteriol, 174(20), 6424–31, 1992. 12. Mather MW, Fee JA. Development of plasmid cloning vectors for Thermus thermophilus HB8: expression of a heterologous, plasmid-borne kanamycin nucleotidyltransferase gene. Appl Environ Microbiol, 58(1), 421–5, 1992. 13. Hoshino T, Maseda H, Nakahara T. Plasmid marker rescue transformation in Thermus thermophilus. J Ferm Bioeng, 76(4), 276–9, 1993. 14. Barany F, Danzitz M, Zebala J, Mayer A. Cloning and sequencing of genes encoding the TthHB8I restriction and modification enzymes: comparison with the isoschizomeric TaqI enzymes. Gene, 112(1), 3–12, 1992. 15. Wayne J, Holden M, Xu SY. The Tsp45I restriction-modification system is plasmid-borne within its thermophilic host. Gene, 202(1–2), 83–8, 1997. 16. Tamakoshi M, Yaoi T, Oshima T, Yamagishi A. An efficient gene replacement and deletion system for an extreme thermophile, Thermus thermophilus. FEMS Microbiol Lett, 173(2), 431–7, 1999. 17. Wayne J, Xu SY. Identification of a thermophilic plasmid origin and its cloning within a new ThermusE. coli shuttle vector. Gene, 195(2), 321–8, 1997. 18. Maseda H, Hoshino T. Screening and analysis of DNA fragments that show promoter activities in Thermus thermophilus. FEMS Microbiol Lett, 128(2), 127–34, 1995. 19. Kayser KJ, Kwak JH, Park HS, Kilbane JJ, 2nd. Inducible and constitutive expression using new plasmid and integrative expression vectors for Thermus sp. Lett Appl Microbiol, 32(6), 412–18, 2001. 20. Moreno R, Zafra O, Cava F, Berenguer J. Development of a gene expression vector for Thermus thermophilus based on the promoter of the respiratory nitrate reductase. Plasmid, 49(1), 2–8, 2003. 21. Chen Y, Hunsicker-Wang L, Pacoma RL, Luna E, Fee JA. A homologous expression system for obtaining engineered cytochrome ba3 from Thermus thermophilus HB8. Protein Expr Purif, 40(2), 299–318, 2005. 22. Weber JM, Johnson SP, Vonstein V, Casadaban MJ, Demirjian DC. A chromosome integration system for stable gene transfer into Thermus flavus. Biotechnology (NY), 13(3), 271–5, 1995. 23. Lasa I, Caston JR, Fernandez-Herrero LA, de Pedro MA, Berenguer J. Insertional mutagenesis in the extreme thermophilic eubacteria Thermus thermophilus HB8. Mol Microbiol, 6(11), 1555–64, 1992. 24. Forterre P. A hot story from comparative genomics: reverse gyrase is the only hyperthermophilespecific protein. Trends Genet, 18(5), 236–7, 2002. 25. Yu X, Angov E, Camerini-Otero RD, Egelman EH. Structural polymorphism of the RecA protein from the thermophilic bacterium Thermus aquaticus. Biophys J, 69(6), 2728–38, 1995. 26. Castan P, Casares L, Barbe J, Berenguer J. Temperature-dependent hypermutational phenotype in recA mutants of Thermus thermophilus HB27. J Bacteriol, 185(16), 4901–7, 2003. 27. Ohta T, Tokishita S, Imazuka R, Mori I, Okamura J, Yamagata H. Beta-glucosidase as a reporter for the gene expression studies in Thermus thermophilus and constitutive expression of DNA repair genes. Mutagenesis, 21(4), 255–60, 2006. 28. Kosuge T, Hoshino T. Lysine is synthesized through the alpha-aminoadipate pathway in Thermus thermophilus. FEMS Microbiol Lett, 169(2), 361–7, 1998.

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29. Kosuge T, Hoshino T. The alpha-aminoadipate pathway for lysine biosynthesis is widely distributed among Thermus strains. J Biosci Bioeng, 88(6), 672–5, 1999. 30. Matsumura M, Yasumura S, Aiba S. Cumulative effect of intragenic amino-acid replacements on the thermostability of a protein. Nature, 323(6086), 356–8, 1986. 31. Hoseki J, Yano T, Koyama Y, Kuramitsu S, Kagamiyama H. Directed evolution of thermostable kanamycin-resistance gene: a convenient selection marker for Thermus thermophilus. J Biochem (Tokyo), 126(5), 951–6, 1999. 32. Brouns SJ, Wu H, Akerboom J, Turnbull AP, de Vos WM, van der Oost J. Engineering a selectable marker for hyperthermophiles. J Biol Chem, 280(12), 11422–31, 2005. 33. Nakamura A, Takakura Y, Kobayashi H, Hoshino T. In vivo directed evolution for thermostabilization of Escherichia coli hygromycin B phosphotransferase and the use of the gene as a selection marker in the host-vector system of Thermus thermophilus. J Biosci Bioeng, 100(2), 158–63, 2005. 34. Fridjonsson O, Watzlawick H, Mattes R. Thermoadaptation of alpha-galactosidase AgaB1 in Thermus thermophilus. J Bacteriol, 184(12), 3385–91, 2002. 35. Tamakoshi M, Nakano Y, Kakizawa S, Yamagishi A, Oshima T. Selection of stabilized 3-isopropylmalate dehydrogenase of Saccharomyces cerevisiae using the host-vector system of an extreme thermophile, Thermus thermophilus. Extremophiles, 5(1), 17–22, 2001. 36. Nelson KE, Clayton RA, Gill SR, Gwinn ML, Dodson RJ, Haft DH, et al. Evidence for lateral gene transfer between archaea and bacteria from genome sequence of Thermotoga maritima. Nature, 399(6734), 323–9, 1999. 37. Harriott OT, Huber R, Stetter KO, Betts PW, Noll KM. A cryptic miniplasmid from the hyperthermophilic bacterium Thermotoga sp. strain RQ7. J Bacteriol, 176(9), 2759–62, 1994. 38. Yu JS, Vargas M, Mityas C, Noll KM. Liposome-mediated DNA uptake and transient expression in Thermotoga. Extremophiles, 5(1), 53–60, 2001. 39. Lee YE, Jain MK, Lee CY, Lowe SE, Zeikus JG. Taxonomic distinction of saccharolytic thermophilic anaerobes—description of Thermoanaerobacterium xylanolyticum gen. nov. sp. nov, and Thermoanaerobacterium saccharolyticum, gen. nov., sp. nov., reclassification of Thermoanaerobium brockii, Clostridium thermosulfurogenes, and Clostridium thermohydrosulfuricum as Thermoanaerobacter brockii comb. nov., Thermoanaerobacterium thermosulfurigenes, comb. nov., and Thermoanaerobacter thermohydrosulfuricus, comb. nov., respectively and transfer of Clostridium thermohydrosulfuricum to Thermoanaerobacter ethanolicus. Int J Syst Bacteriol, 43, 41–51, 1993. 40. Mai V, Wiegel J. Advances in development of a genetic system for Thermoanaerobacterium spp.: expression of genes encoding hydrolytic enzymes, development of a second shuttle vector, and integration of genes into the chromosome. Appl Environ Microbiol, 66(11), 4817–21, 2000. 41. Desai SG, Guerinot ML, Lynd LR. Cloning of l-lactate dehydrogenase and elimination of lactic acid production via gene knockout in Thermoanaerobacterium saccharolyticum JW/SL-YS485. Appl Microbiol Biotechnol, 65(5), 600–5, 2004. 42. Koyama Y, Okamoto S, Furukawa K. Cloning of alpha- and beta-galactosidase genes from an extreme thermophile, thermus strain T2, and their expression in Thermus thermophilus HB27. Appl Environ Microbiol, 56(7), 2251–4, 1990. 43. Park HS, Kilbane JJ, 2nd. Gene expression studies of Thermus thermophilus promoters PdnaK, Parg and Pscs-mdh. Lett Appl Microbiol, 38(5), 415–22, 2004. 44. Takayama G, Kosuge T, Maseda H, Nakamura A, Hoshino T. Nucleotide sequence of the cryptic plasmid pTT8 from Thermus thermophilus HB8 and isolation and characterization of its high-copy number mutant. Plasmid, 51(3), 227–37, 2004. 45. Kobayashi H, Kuwae A, Maseda H, Nakamura A, Hoshino T. Isolation of a low-molecular-weight, multicopy plasmid, pNHK101, from Thermus sp. TK10 and its use as an expression vector for T. thermophilus HB27. Plasmid, 54(1), 70–9, 2005.

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Targeted Gene Disruption as a Tool for Establishing Gene Function in Hyperthermophilic Archaea Haruyuki Atomi and Tadayuki Imanaka

CONTENTS Introduction ................................................................................................................................ Isolation of a Uracil-Dependent Host Strain with a Deficiency in Pyrimidine Biosynthesis .... Strategy for Homologous Recombination .................................................................................. Construction of Various Sets of Auxotrophic Host Cells and Marker Genes ........................... Transformation Efficiency ......................................................................................................... Gene Disruption Based on Antibiotics and Resistance Genes .................................................. Gene Disruption as a Tool in Determining Gene Function ....................................................... Reverse Gyrase ............................................................................................................... Fructose-1,6-Bisphosphatase .......................................................................................... Pyruvate Kinase and Phosphoenolpyruvate Synthase .................................................... Pentose and Nucleotide Synthesis .................................................................................. Gene Disruption in Sulfolobus solfataricus .............................................................................. Future Perspectives .................................................................................................................... Acknowledgments ...................................................................................................................... References ..................................................................................................................................

213 214 215 215 217 217 218 218 219 220 220 220 221 221 221

INTRODUCTION In spite of the huge accumulation of sequence data and in vitro analyses on recombinant and native proteins, the lack of a genetic system in hyperthermophiles had been a major bottleneck in determining the physiological roles of genes. As demonstrated in a wide range of bacteria and eukaryotes, the examination of phenotypic changes brought about by deletion of a particular gene is a straightforward means for establishing gene function in vivo. Among the hyperthermophiles, gene disruption systems have been developed in Thermococcus kodakaraensis, a sulfur-reducing hyperthermophilic archaeon in the Euryarchaeota [1], and Sulfolobus solfataricus, an acidophilic hyperthermophilic archaeon belonging to the Crenarchaeota [2]. This chapter will deal with the recently developed gene disruption technology in hyperthermophilic archaea, focusing mainly on the system developed in T. kodakaraensis.

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ISOLATION OF A URACIL-DEPENDENT HOST STRAIN WITH A DEFICIENCY IN PYRIMIDINE BIOSYNTHESIS One disadvantage in developing a transformation system in hyperthermophiles is that the majority of antibiotic-resistant genes utilized in mesophilic organisms cannot be expected to be applicable. Even if the host cell were to be sensitive to a particular antibiotic, the resistance gene product would most likely not exhibit sufficient thermostability to function in hyperthermophiles. Therefore, a system based on auxotrophic host cells and marker genes (deriving from hyperthermophiles) that confer prototrophy can be regarded as the better choice. Gene disruption systems in yeast cells such as Saccharomyces cerevisiae utilize this strategy, and host cells exhibiting auxotrophy towards a wide range of compounds such as uracil, tryptophan, adenine, and histidine have been developed along with their complementary marker genes URA3, TRP1, ADE2, and HIS3. The first step in developing such a system would be to isolate a stable, auxotrophic host cell deficient in the function of a single gene in a particular biosynthetic pathway. The pyrimidine biosynthesis pathway provides an advantage as strains harboring defects in this pathway can be screened in a positive manner. Two key enzymes of this pathway orotate phosphoribosyltransferase (pyrE or URA5) and orotidine-5′-phosphate decarboxylase (pyrF or URA3) are responsible for the conversion of orotate to uridine-5′-phosphate (Figure 13.1). When 5-fluoro-orotate (5-FOA) is added to the medium, the function of the two enzymes result in the production of 5-fluorouridine5′-phosphate, a toxic compound that inhibits cell growth. On the other hand, the presence of uracil phosphoribosyltransferase activity will allow pyrimidine biosynthesis in the absence of PyrE or PyrF activity as long as uracil is added to the medium. Therefore, one can positively select mutant strains with a deficiency in PyrE or PyrF by isolating strains that can grow in the presence of 5-FOA and uracil (Figure 13.1).

O HN O

N H

PRPP

Upp PPi PRPP

O

carbamoyl phosphate aspartate

HN O

N H

O

COOH

PyrE

O HN N H

PRPP

N I

COOH

COOH

O

F

N I

COOH

Rib-5P 5-fluoroorotidine 5'-phosphate

O

PyrF

O

PPi HN

5-fluoro orotic acid

HN

Rib-5P orotidine 5'-phosphate

F

O

CO2

HN

orotic acid

O

O

PPi

pyrimidines N I

Rib-5P uridine 5'-phosphate O

CO2

F

HN O

N I

Rib-5P 5-fluorouridine 5'-phosphate

FIGURE 13.1 A schematic diagram illustrating the pyrimidine biosynthesis pathway, highlighting the reactions of orotate phosphoribosyltransferase (PyrE) and orotidine-5′-phosphate decarboxylase (PyrF). The conversion of uracil to uridine 5′-phosphate by uracil phosphoribosyltransferase (Upp) and the conversion of 5-fluoro-orotate to 5-fluorouridine 5′-phosphate by PyrE and PyrF is also shown.

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FIGURE 13.2 A schematic drawing of pUDT2, a plasmid designed for the disruption of trpETk via double cross-over homologous recombination. The homologous regions present on both the plasmid and chromosome are shaded. P represents the 5′-flanking region of a gene cluster which includes the pyrF gene.

This strategy was used to isolate uracil auxotrophs of T. kodakaraensis with defects in pyrE or pyrF [3]. After ultraviolet (UV) irradiation and screening on plate medium in the presence of both uracil and 5-FOA, a number of uracil auxotrophs were isolated. Among these, strain KU25, whose pyrF gene had a one base pair deletion at position 96, was used as a host cell for transformation experiments.

STRATEGY FOR HOMOLOGOUS RECOMBINATION Two types of plasmids harboring the wild-type pyrF gene were designed for single cross-over (pUDT1) and double cross-over (pUDT2; Figure 13.2) homologous recombination. The pyrF gene was under the control of a putative promoter region upstream of the gene cluster which includes pyrF. The target gene for disruption was the trpE gene of T. kodakaraensis (trpETk ) encoding the large subunit of anthranilate synthase. Examination of isolates exhibiting uracil prototrophy revealed that in T. kodakaraensis, double cross-over recombination occurred at much higher frequencies. One transformant (strain KW4) was selected and further examined. Southern blot analysis revealed that the pyrF marker gene was integrated into the trpE locus, thereby disrupting the gene. The absence of nonspecific integration of the plasmid into other regions of the chromosome was also confirmed. Growth of KW4 was no longer dependent on the presence of uracil, and instead exhibited tryptophan auxotrophy. The results indicated that in T. kodakaraensis, targeted gene disruption was possible via double cross-over homologous recombination [3].

CONSTRUCTION OF VARIOUS SETS OF AUXOTROPHIC HOST CELLS AND MARKER GENES As the efficient occurrence of double cross-over recombination became apparent, a plasmid designed to delete the pyrF gene was used to transform wild-type T. kodakaraensis KOD1 cells. The isolate, which was selected in media containing both 5-FOA and uracil, can be presumed not to harbor random mutations on its chromosome as was anticipated with strain KU25. Furthermore, the pyrF gene is almost entirely deleted from the chromosome, so that the possibilities of recombination between a

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FIGURE 13.3 Targeted disruption of pyrF, trpE, and hisD in Thermococcus kodakaraensis KOD1, KU216, and KW128 using the three disruption vectors pUDPyrF, pUDTrpE, and pUDHisD, respectively. Relevant regions of the chromosome are illustrated for (from the top) strains KOD1, KU216, KW128, and KH3.

heterologous pyrF marker gene and the native pyrF locus are negligible. This strain was designated KU216 (ΔpyrF) [4]. Using pUDT2, trpETk was once again disrupted from the host strain KU216 (Figure 13.3). The obtained isolate, KW128, exhibited uracil prototrophy and tryptophan auxotrophy, and was confirmed to harbor the genotype (ΔpyrF, ΔtrpE::pyrF). The trpE gene, fused to the promoter region used in the pyrF marker, was then used for gene disruption experiments with KW128 as a host cell. With hisD, a gene necessary for histidine biosynthesis, as the target for disruption, transformants displaying tryptophan prototrophy could be isolated. Growth of the transformants were dependent on the addition of exogenous histidine to the medium, and the genotype was confirmed to be as expected (ΔpyrF, ΔtrpE::pyrF, ΔhisD::trpE). These studies indicate that the host cells KU216 and KW128 can be used as stable host cells for gene disruption studies in T. kodakaraensis, using the wild-type pyrF and trpE genes as markers, respectively [4]. In yeast, a common method to reutilize a single marker gene has been developed using the counterselectable pyrF gene. This is based on the fact that pyrF is necessary for cell growth in the absence of uracil for pyrimidine biosynthesis, but its presence is detrimental when cells are grown in the presence of 5-FOA. As shown in Figure 13.4, an identical, second copy of the 3′-flanking region of the gene to be disrupted is inserted between the 5′-flanking region of the gene and the marker gene (pyrF). pyrF is thus sandwiched between two identical sequences. After transformation, screening is carried out for uracil prototrophy, and cells that have undergone double cross-over homologous recombination at the outermost 5′- and 3′-flanking regions can be selected. Using these transformants, a second round of selection is carried out in the presence of uracil and 5-FOA. Under these conditions, the growth of cells that harbor an active pyrF gene is completely inhibited, and thus only cells that have undergone a second recombination between the two identical regions flanking the pyrF marker gene can grow. As a result, both the gene targeted for disruption and the marker gene are removed from the chromosome, thereby allowing the consecutive use of the pyrF marker gene on the newly generated knockout strain. This methodology has been used in constructing the strain KUW1, which exhibits the genotype (ΔpyrF, ΔtrpE), and can therefore accommodate both the pyrF and trpE markers [4].

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FIGURE 13.4 Disruption of trpE followed by the excision of the pyrF marker by pop-out recombination. Construction of strain KUW1 (ΔpyrF ΔtrpE) using pop-out vectors harboring tandem repeats of the endogenous 3′-region of the target gene flanking pyrF on both sides. The regions shaded in gray indicate the tandem repeat regions. The changes in relevant genotypes are indicated on the left.

TRANSFORMATION EFFICIENCY The transformation efficiency of T. kodakaraensis was examined using plasmids harboring homologous regions of various lengths [4]. When the trpE marker gene was inserted between the 5′- and 3′-flanking regions of the target gene, the number of tryptophan prototrophs was in the range of 102 per microgram DNA per 4 × 108 cells with flanking regions of ~1000 bp. Linearization of the plasmid prior to transformation had little effect on transformation efficiency. When the flanking regions were shortened to ~500 bp, the transformation efficiency decreased to 101 per microgram DNA per 4 × 108 cells. No prototrophs were obtained with flanking regions of 100 bp. The efficiency of this system is sufficient for targeted gene disruption, but is too low for the introduction of DNA libraries for use in random mutagensis–complementation experiments. A 10-fold coverage of the genome (~2 Mbp) with 5-kbp fragments would require 4 × 103 colonies. Improvements in both transformation and plating efficiencies and the development of shuttle vector systems will have to be considered.

GENE DISRUPTION BASED ON ANTIBIOTICS AND RESISTANCE GENES As described before, conventional antibiotic resistance marker genes cannot be used in hyperthermophiles due to their lack of thermostability. However, a strategy based on inhibition of a particular endogenous protein by an antibiotic and relieving the inhibition by overexpressing the protein or by introducing a mutant protein insensitive to the antibiotic, is feasible. Simvastatin and mevinolin are specific inhibitors of the enzyme 3-hydroxy-3-methylglutaryl coenzyme A (HMG-CoA) reductase, the key enzyme of the mevalonate pathway. As isopentenyl diphosphate is the major precursor for membrane synthesis in the archaea, inhibition of HMG-CoA reductase can be presumed to have critical effects on the growth of all archaeal strains [5]. This has clearly been demonstrated in a number of halophilic archaea, and genetic systems utilizing mevinolin have been developed in these strains [6,7]. To develop such a system in T. kodakaraensis, overexpression cassettes were constructed for the endogenous HMG-CoA reductase gene from T. kodakaraensis (hmgTk) and the heterologous

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FIGURE 13.5 Design of the hmgTk and hmgPf overexpression cassettes using the 5′-upstream region of the glutamate dehydrogenase gene (Pgdh) from Thermococcus kodakaraensis.

gene from Pyrococcus furiosus (hmgPf) [8]. The promoter used was the 5′-upstream region of the glutamate dehydrogenase gene (Pgdh) from T. kodakaraensis. The wild-type strain, T. kodakaraensis KOD1, was sensitive to simvastatin and growth was completely inhibited for 5 days in the presence of 4 μM simvastatin. Plasmids designed for double cross-over homologous recombination were constructed using the overexpression cassettes (Pgdh-hmgTk, Pgdh-hmgPf) as marker genes (Figure 13.5). Transformants resistant toward simvastatin at concentrations higher than 4 μM were isolated and examined. All transformants displayed resistance against simvastatin, and were capable of growth even in the presence of 20 μM of simvastatin. Genotype analyses indicated that both overexpression cassettes were applicable for targeted gene disruption. However, Pgdh-hmgPf, with the heterologous HMG-CoA reductase gene, was found to be the more convenient of the two as unintended recombination at the native hmg locus, which was observed using Pgdh-hmgTk, did not occur. This system allows gene disruption in nutrient-rich media, and can be directly applied on the wild-type T. kodakaraensis KOD1 [8]. Besides representing a convenient alternative for gene disruption in this organism, it should be helpful in developing gene disruption systems in other hyperthermophilic archaea as there is no need for the initial development of auxotrophic host strains.

GENE DISRUPTION AS A TOOL IN DETERMINING GENE FUNCTION Using the transformation systems described before, a number of genes have been disrupted in T. kodakaraensis. The studies have led to a better understanding on the actual function of these genes in vivo [9–13], and will be described next.

REVERSE GYRASE Reverse gyrase was first identified in Sulfolobus acidocaldarius as an enzyme that introduces positive supercoils in covalently closed DNA [14]. Recent studies have shown that the enzyme also exhibits a number of additional activities toward DNA [15,16]. For example, the enzyme binds nicks and DNA ends in a cooperative manner, acting as a DNA chaperone that enhances the thermostability of DNA by decreasing the rate of strand breakage [17]. Reverse gyrase has attracted much attention not only due to its unique activity, but also because it is the only enzyme/gene that is present in all hyperthermophilic organisms but absent in all mesophilic organisms, that is, it is the one and only hyperthermophile-specific protein [18]. On the other

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hand, the structure of reverse gyrase suggests that it is not at all a primitive enzyme in terms of protein evolution. Reverse gyrase is formed by the association of two entirely different enzymes belonging to the DNA/RNA helicases and the topoisomerases, and thus could only have evolved after the diversification of the respective protein families [19]. If reverse gyrase were to be a prerequisite for life at extremely high temperatures, this would provide a convincing argument that contradicts with the hot origin of life, or life originating in hyperthermophilic organisms. The evolution of the two protein families could only have occurred in less thermophilic organisms [20]. Disruption of the reverse gyrase gene (rgyTk) and the subsequent phenotype examination have provided valuable insight on the importance of the enzyme for life at high temperatures [9]. Gene disruption was not lethal, but specific growth rates of the ΔrgyTk strain declined rapidly with increases in temperature above 80˚C compared with the host strain. The gene disruption strain was able to grow at 90˚C but not at higher temperatures. The results indicate that reverse gyrase provides a significant advantage for life at high temperatures (>80˚C), and helps in understanding why all organisms isolated from hyperthermophilic environments until now harbor a reverse gyrase. The results also revive the possibilities of a hot origin of life in which primitive hyperthermophiles without a reverse gyrase might have been the first organisms to evolve, most likely at temperatures below 90˚C.

FRUCTOSE-1,6-BISPHOSPHATASE Fructose-1,6-bisphosphatase (FBPase) is a key enzyme for gluconeogenesis in all three domains of life. The enzyme catalyzes the physiologically irreversible dephosphorylation of fructose1,6-bisphosphate (F16P) to fructose-6-phosphate (F6P). Its counterpart in glycolysis, phosphofructokinase (PFK), catalyzes the nucleotide-dependent phosphorylation of F6P to F16P, also in an irreversible manner. As the simultaneous function of PFK and FBPase would result in a futile cycle leading to energy dissipation, both enzymes are known to be under strict regulation responding to the growth conditions. One intriguing feature of the hyperthermophiles was that although these organisms can grow on gluconeogenic substrates, homologs of the classical FBPase genes (classes I–III) identified in bacteria and eukaryotes were not present on their genomes. As several hyperthermophiles even exhibited FBPase activity in their cell-free extracts, the presence of an FBPase with novel structure was strongly suggested. An inositol monophosphatase (IMPase) homolog from Methanocaldococcus jannaschii, whose catalytic pocket resembled those of the FBPases from higher eukaryotes, was found to exhibit FBPase activity in addition to its IMPase activity [21]. Corrresponding proteins from Archaeoglobus fulgidus [22] and Pyrococcus furiosus [23] also displayed FBPase activity, and these proteins were subsequently classified as the class IV FBPases. In T. kodakaraensis, still another protein was identified by purifying the FBPase activity found in cells grown on gluconeogenic substrates [24]. The gene encoding this protein ( fbpTk) and the gene from T. kodakaraensis encoding IMPase (impTk) were individually expressed in Escherichia coli, and examined for their phosphatase activities. Both recombinant enzymes displayed significant levels of FBPase activity, with ImpTk exhibiting a higher kcat/Km value than FbpTk [10]. This is a situation in which gene disruption provides a straightforward approach; two genes encoding proteins with comparable levels of identical activities in vitro. Gene disruption was performed on fbpTk and impTk using the host strain KW128 and the trpETk marker gene. The ΔimpTk strain displayed growth on both glycolytic and gluconeogenic substrates that were indistinguishable to the host strain. In contrast, while growth on glycolytic substrates was unaffected, ΔfbpTk cells could not grow on gluconeogenic substrates [10]. The results present strong evidence that the enzyme functioning for gluconeogenesis in T. kodakaraensis is FbpTk and not ImpTk. As the FbpTk homolog is present on most hyperthermophile genomes, the enzyme most likely represents a novel class of FBPases (class V) that function in hyperthermophiles.

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PYRUVATE KINASE AND PHOSPHOENOLPYRUVATE SYNTHASE Similar to the situation for FBPase, there were two enzyme/gene candidates for the conversion from phosphoenolpyruvate (PEP) to pyruvate, the final step of the modified Embden-Meyerhof (EM) pathway [25]. Pyruvate kinase (PYK) and PEP synthase (PPS) can be considered as isozymes when one focuses only on carbon conversion, but the cofactors and substrates involved are distinct and the free-energy differences of the reactions greatly differ. The PYK reaction can be considered only to function in the direction of PEP to pyruvate, coupled to the generation of adenosine triphosphate (ATP) from adenosine diphosphate (ADP). PPS can catalyze the reversible conversion between PEP, AMP and phosphate to pyruvate, ATP, and H2O [26]. The genes (pykTk and ppsTk) were individually disrupted and cell growth was examined under various growth conditions [11]. The most dramatic change in phenotype was that observed in the ΔppsTk grown under glycolytic conditions. No growth was observed for the ΔppsTk strain, indicating that PPS, and not PYK, is essential for glycolysis in the modified EM pathway. This is in contrast to the classical EM pathway of the eukaryotes and E. coli, in which PYK is responsible for the conversion of PEP to pyruvate. The results also provide valuable information on the energy metabolism of the modified EM pathway. As glyceraldehyde-3-phosphate dehydrogenase and 3-phosphoglycerate kinase are presumed to be substituted by the function of glyceraldehyde-3-phosphate:ferredoxin oxidoreductase [27]. ATP generation at the substrate level would be absent in the modified EM pathway if PYK were to be the main enzyme converting PEP to pyruvate. As this is not the case, the two ADP-dependent sugar phosphatases [28] convert two molecules of ADP to AMP per glucose, and PPS converts two molecules of AMP to ATP per glucose unit. In total, two molecules of ATP are generated from two molecules of ADP, indicating that the modified EM pathway is an energy-generating pathway at the substrate level.

PENTOSE AND NUCLEOTIDE SYNTHESIS The pentose phosphate pathway is a ubiquitous pathway in bacteria and eukaryotes that supplies the pentose precursors necessary for nucleotide biosynthesis. The pathway is also important for the generation of NADPH which is vital for reductive biosynthesis, and for the production of erythrose-4-phosphate, the precursor for aromatic amino acid biosynthesis. It had been pointed out that many of the archaeal genomes do not have complete sets of homologs of the pentose phosphate pathway, and that instead, homologs of the ribulose monophosphate pathway are present on many of these genomes [29]. Labeling experiments with the hyperthermophilic archaeon Methanocaldococcus jannaschii also indicated the involvement of this pathway in pentose synthesis [30]. The ribulose monophosphate pathway was originally identified in methylotrophic bacteria as an assimilation and detoxification pathway of formaldehyde. The pathway is composed of two enzymes; 3-hexulose-6-phosphate synthase (HPS) which converts ribulose-5-phosphate and formaldehyde to 3-hexulose-6-phosphate, and 6-phospho-3-hexuloisomerase (PHI) which converts 3-hexulose6-phosphate to F6P [31]. To confirm the involvement of the ribulose monophosphate pathway in pentose biosynthesis, the gene encoding the HPS/PHI fusion protein in T. kodakaraensis (hps/phiTk) was first expressed in E. coli. The recombinant protein exhibited activity in both formaldehyde-fixing and -releasing directions. The gene was then disrupted, and the Δhps/phiTk strain was found to grow only in the presence of exogenous nucleotides or nucleosides. The results indicate that the ribulose monophosphate pathway is involved in the biosynthesis of nucleosides/nucleotides in T. kodakaraensis by functioning in the formaldehyde-releasing direction to generate ribulose-5-phosphate from F6P [12].

GENE DISRUPTION IN SULFOLOBUS SOLFATARICUS The gene disruption system developed for S. solfataricus applies a host strain, PBL2002, which cannot utilize lactose due to a spontaneous transposition of IS1217 into the native lacS gene [32].

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A modified but functional lacS gene with a silent nucleotide substitution that disrupts a unique EcoRV site was used as the marker gene (lacS*). A plasmid aimed for double cross-over homologous recombination was constructed in which the lacS* gene, along with its 5′- and 3′-regions, was inserted into the α-amylase gene of S. solfataricus (amyA). Transformation of PBL2002 with this plasmid resulted in two classes of recombinants that were able to utilize lactose. Although the first class consisted of recombinants that had undergone recombination at the native lacS locus, the second class was composed of transformants whose amyA gene had been specifically disrupted via double cross-over recombination. Distinguishing the lacS* gene with the native lacS can be easily carried out by digesting polymerase chain reaction (PCR)-amplified products with EcoRV. The results of this study clarify that AmyA plays a major role not only in glycogen utilization but also in the breakdown of pullulan [32]. The power of the lacS* system in S. solfataricus has been confirmed through various examples of gene disruption, including the editing component of threonyl-tRNA synthetase [33], and mercuric reductase and its negative transcriptional regulator [34].

FUTURE PERSPECTIVES Although limited at present to T. kodakaraensis and S. solfataricus, targeted gene disruption is now possible in hyperthermophiles. The methodology will surely become applicable in a wider range of organisms in the near future. Gene disruption, together with the stable shuttle vectors already developed in a number of organisms (see Chapter 11) will accelerate our pace in solving the function of nonannotatable genes and confirming/correcting those of the annotated genes. The application of gene disruption can be expected to be particularly powerful in the functional examination of putative transcription factors and gene regulators in combination with transcriptome analyses. Proteins that are relatively difficult to examine in vitro such as the membrane proteins are also attractive targets. In terms of application, the gene disruption technology can also be used for the insertion of heterologous genes. Hyperthermophile cells can be used as host strains for overexpressing thermostable proteins that are difficult to be expressed in mesophilic host cells. They can also be utilized as host cells for screening random (mutant) libraries aimed to identify useful thermostable enzymes or increase the thermostability of proteins. These applications can also be performed with shuttle vector systems such as those described in Chapter 11, so one can choose the more convenient methodology depending on the needs of the particular study. Finally, combining the technologies of gene disruption/insertion and shuttle vectors will enable us to perform metabolic engineering and cell engineering in hyperthermophiles, and further examine the use of hyperthermophiles in whole cell biocatalysis at high temperatures.

ACKNOWLEDGMENTS The research on T. kodakaraensis in the Imanaka Lab was supported by Japan Science and Technology Corporation (JST) for Core Research for Evolutional Science and Technology (CREST), a Grant-in-Aid for Scientific Research (no. 14103011), and a Grant-in-Aid for Scientific Research on Priority Areas “Applied Genomics” (no. 18018026) from the Ministry of Education, Culture, Sports, Science, and Technology of Japan. The authors would like to thank Frank Robb and Dennis Grogan for critical reading of the manuscript.

REFERENCES 1. Atomi, H., Fukui, T., Kanai, T., Morikawa, M., and Imanaka, T. (2004) Description of Thermococcus kodakaraensis sp. nov., a well studied hyperthermophilic archaeon previously reported as Pyrococcus sp. KOD1, Archaea 1, 263–267. 2. Zillig, W., Stetter, K. O., Wunderl, S., Schulz, W., Priess, H., and Scholz, I. (1980) The Sulfolobus “Caldariella” group: taxonomy on the basis of the structure of DNA-dependent RNA polymerases, Arch. Microbiol. 125, 259–269.

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3. Sato, T., Fukui, T., Atomi, H., and Imanaka, T. (2003) Targeted gene disruption by homologous recombination in the hyperthermophilic archaeon Thermococcus kodakaraensis KOD1, J. Bacteriol. 185, 210–220. 4. Sato, T., Fukui, T., Atomi, H., and Imanaka, T. (2005) Improved and versatile transformation system allowing multiple genetic manipulations of the hyperthermophilic archaeon Thermococcus kodakaraensis, Appl. Environ. Microbiol. 71, 3889–3899. 5. Cabrera, J. A., Bolds, J., Shields, P. E., Havel, C. M., and Watson, J. A. (1986) Isoprenoid synthesis in Halobacterium halobium. Modulation of 3-hydroxy-3-methylglutaryl coenzyme A concentration in response to mevalonate availability, J. Biol. Chem. 261, 3578–3583. 6. Lam, W. L. and Doolittle, W. F. (1989) Shuttle vectors for the archaebacterium Halobacterium volcanii., Proc. Natl. Acad. Sci. USA 86, 5478–5482. 7. Wendoloski, D., Ferrer, C., and Dyall-Smith, M. L. (2001) A new simvastatin (mevinolin)-resistance marker from Haloarcula hispanica and a new Haloferax volcanii strain cured of plasmid pHV2, Microbiology 147, 959–964. 8. Matsumi, R., Manabe, K., Fukui, T., Atomi, H., and Imanaka, T. (2007) Disruption of a sugar transporter gene cluster in a hyperthermophilic archaeon using a host/marker system based on antibiotic resistance, J. Bacteriol., 189, 2683–2691. Published OnLine, doi:10.1128/JB.01692-06. 9. Atomi, H., Matsumi, R., and Imanaka, T. (2004) Reverse gyrase is not a prerequisite for hyperthermophilic life, J. Bacteriol. 186, 4829–4833. 10. Sato, T., Imanaka, H., Rashid, N., Fukui, T., Atomi, H., and Imanaka, T. (2004) Genetic evidence identifying the true gluconeogenic fructose-1,6-bisphosphatase in Thermococcus kodakaraensis and other hyperthermophiles, J. Bacteriol. 186, 5799–5807. 11. Imanaka, H., Yamatsu, A., Fukui, T., Atomi, H., and Imanaka, T. (2006) Phosphoenolpyruvate synthase plays an essential role for glycolysis in the modified Embden-Meyerhof pathway in Thermococcus kodakarensis, Mol. Microbiol. 61, 898–909. 12. Orita, I., Sato, T., Yurimoto, H., Kato, N., Atomi, H., Imanaka, T., and Sakai, Y. (2006) The ribulose monophosphate pathway substitutes for the missing pentose phosphate pathway in the archaeon Thermococcus kodakaraensis, J. Bacteriol. 188, 4698–4704. 13. Sato, T., Atomi, H. and Imanaka, T. (2007) Archaeal type III RuBisCOs function in a pathway for AMP metabolism, Science 315, 1003–1006. 14. Kikuchi, A. and Asai, K. (1984) Reverse gyrase—a topoisomerase which introduces positive superhelical turns into DNA, Nature 309, 677–681. 15. Nadal, M. (2007) Reverse gyrase: an insight into the role of DNA-topoisomerases, Biochimie, doi:10.1016/j.biochi.2006.12.010. 16. Hsieh, T. S. and Plank, J. L. (2006) Reverse gyrase functions as a DNA renaturase: annealing of complementary single-stranded circles and positive supercoiling of a bubble substrate, J. Biol. Chem. 281, 5640–5647. 17. Kampmann, M. and Stock, D. (2004) Reverse gyrase has heat-protective DNA chaperone activity independent of supercoiling, Nucleic Acids Res. 32, 3537–3545. 18. Forterre, P. (2002) A hot story from comparative genomics: reverse gyrase is the only hyperthermophilespecific protein, Trends Genet. 18, 236–237. 19. Confalonieri, F., Elie, C., Nadal, M., de La Tour, C., Forterre, P., and Duguet, M. (1993) Reverse gyrase: a helicase-like domain and a type I topoisomerase in the same polypeptide, Proc. Natl. Acad. Sci. USA 90, 4753–4757. 20. Forterre, P. (1996) A hot topic: the origin of hyperthermophiles, Cell 85, 789–792. 21. Stec, B., Yang, H., Johnson, K. A., Chen, L., and Roberts, M. F. (2000) MJ0109 is an enzyme that is both an inositol monophosphatase and the “missing” archaeal fructose-1,6-bisphosphatase, Nat. Struct. Biol. 7, 1046–1050. 22. Stieglitz, K. A., Johnson, K. A., Yang, H., Roberts, M. F., Seaton, B. A., Head, J. F., and Stec, B. (2002) Crystal structure of a dual activity IMPase/FBPase (AF2372) from Archaeoglobus fulgidus. The story of a mobile loop, J. Biol. Chem. 277, 22863–22874. 23. Verhees, C. H., Akerboom, J., Schiltz, E., de Vos, W. M., and van der Oost, J. (2002) Molecular and biochemical characterization of a distinct type of fructose-1,6-bisphosphatase from Pyrococcus furiosus, J. Bacteriol. 184, 3401–3405. 24. Rashid, N., Imanaka, H., Kanai, T., Fukui, T., Atomi, H., and Imanaka, T. (2002) A novel candidate for the true fructose-1,6-bisphosphatase in archaea, J. Biol. Chem. 277, 30649–30655.

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25. Verhees, C. H., Kengen, S. W., Tuininga, J. E., Schut, G. J., Adams, M. W., De Vos, W. M., and Van Der Oost, J. (2003) The unique features of glycolytic pathways in archaea, Biochem. J. 375, 231–246. 26. Sakuraba, H., Utsumi, E., Kujo, C., and Ohshima, T. (1999) An AMP-dependent (ATP-forming) kinase in the hyperthermophilic archaeon Pyrococcus furiosus: characterization and novel physiological role, Arch. Biochem. Biophys. 364, 125–128. 27. Mukund, S. and Adams, M. W. (1995) Glyceraldehyde-3-phosphate ferredoxin oxidoreductase, a novel tungsten-containing enzyme with a potential glycolytic role in the hyperthermophilic archaeon Pyrococcus furiosus, J. Biol. Chem. 270, 8389–8392. 28. Kengen, S. W., de Bok, F. A., van Loo, N. D., Dijkema, C., Stams, A. J., and de Vos, W. M. (1994) Evidence for the operation of a novel Embden-Meyerhof pathway that involves ADP-dependent kinases during sugar fermentation by Pyrococcus furiosus, J. Biol. Chem. 269, 17537–17541. 29. Soderberg, T. (2005) Biosynthesis of ribose-5-phosphate and erythrose-4-phosphate in archaea: a phylogenetic analysis of archaeal genomes, Archaea 1, 347–352. 30. Grochowski, L. L., Xu, H., and White, R. H. (2005) Ribose-5-phosphate biosynthesis in Methanocaldococcus jannaschii occurs in the absence of a pentose-phosphate pathway, J. Bacteriol. 187, 7382–7389. 31. Kato, N., Yurimoto, H., and Thauer, R. K. (2006) The physiological role of the ribulose monophosphate pathway in bacteria and archaea, Biosci. Biotechnol. Biochem. 70, 10–21. 32. Worthington, P., Hoang, V., Perez-Pomares, F., and Blum, P. (2003) Targeted disruption of the α-amylase gene in the hyperthermophilic archaeon Sulfolobus solfataricus, J. Bacteriol. 185, 482–488. 33. Korencic, D., Ahel, I., Schelert, J., Sacher, M., Ruan, B., Stathopoulos, C., Blum, P., Ibba, M., and Soll, D. (2004) A freestanding proofreading domain is required for protein synthesis quality control in archaea, Proc. Natl. Acad. Sci. USA 101, 10260–10265. 34. Schelert, J., Dixit, V., Hoang, V., Simbahan, J., Drozda, M., and Blum, P. (2004) Occurrence and characterization of mercury resistance in the hyperthermophilic archaeon Sulfolobus solfataricus by use of gene disruption, J. Bacteriol. 186, 427–437.

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Nanobiotechnological Potential of Viruses of Hyperthermophilic Archaea Tamara Basta and David Prangishvili

CONTENTS Acidianus Two-Tailed Virus ...................................................................................................... Acidianus Filamentous Virus 1 and Other Lipothrixviruses .................................................... Sulfolobus islandicus Rod-Shaped Virus 1 and Other Rudiviruses .......................................... Acidianus Bottle-Shaped Virus ................................................................................................. Spherical Viruses of Sulfolobus and Pyrobaculum ................................................................... Sulfolobus Spindle-Shaped and Droplet-Shaped Viruses ......................................................... Zipper Virus-Like Particles of Acidianus .................................................................................. Conclusions and Perspectives .................................................................................................... Acknowledgments ...................................................................................................................... References ..................................................................................................................................

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Viruses are particularly suitable for applications in nanobiotechnology because their structural components are in the nanoscale range and they have the intrinsic characteristic of self-assembly. In this chapter we describe specific features of several hyperthermophilic viruses and virus-like particles that could be of special interest for applications in nanobiotechnology. The viruses have been isolated from hot acidic springs, (>80°C and pH < 3) in different regions of active volcanism and tectonics and infect strains of the hyperthermophilic genera Sulfolobus and Acidianus from the third domain of life, the Archaea.

ACIDIANUS TWO-TAILED VIRUS Acidianus two-tailed virus (ATV) is the only virus known to day with a capability to undergo a major morphological transformation outside and independently of the host cell. The unique transformation process of ATV is initiated after the virions are extruded from infected cells. These particles are lemon-shaped with an average length of 243(±11) nm and a maximum width of 119(±2) nm. Subsequently, the lemon-shaped particles develop protrusions at each pointed end which are extended into tail-like appendices. Fully developed two-tailed virions show an average length of 744(±24) nm and maximum width of 85(±4) nm [1] (Figure 14.1a, and f). The ATV morphogenesis is independent of the presence of cells or any cofactors and can be initiated and proceed even in distilled water. The only requirement for the protrusion of tails is that the temperature of the environment is above 75°C. At 85°C to 90°C, the temperature range close to 225

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FIGURE 14.1 Tail development and virion structure of Acidianus two-tailed virus, ATV. Cryo-electron micrographs of tail-less (a) and fully developed two-tailed (f) virions schematically represented in (b) and (c), respectively. The inner filament in the tail tube (d) and terminal structure (e) were visualized by electron tomography of negatively stained particles [2]. Scale bars represent 50 nm. (Modified from Prangishvili, D. et al., J Mol Biol, 359, 1203–16, 2006.)

that of the host habitat, the rate of tail protrusion is fastest and according to rough estimate is equal to 250 nm/h. Lemon-shaped particles are stable in solution and can be kept at 4°C for at least several months without loosing the capability to grow tails. The tail-like appendices are hollow tubes of 27(±)3 nm in diameter with about 6(±1) nm thick wall [2]. A filament 2 nm in width with a repeat periodicity of about 11 nm resides inside the tube (Figure 14.1d). The tube ends in a narrow channel 2 nm in width and a terminal anchorlike structure formed by two furled filaments, each with a width of 4 nm (Figure 14.1e). Such terminal structure resembles in its appearance the furled protofilaments of the microtubule termini and could reflect a similar structural organization and/or mechanism of formation of the ATV “tails” and microtubules [3,4]. Protein 800 encoded on the viral genome is a likely candidate to be involved in extracellular morphogenesis. The purified recombinant form of P800 is capable to polymerase spontaneously at 4°C and form long fibers with the width of 2 nm. The filaments reveal the tendency to assemble longitudinally into clusters. Possibility of involvement of any of the ten other identified structural proteins in the tail formation has not yet been studied [2]. Despite the potential complexity of the system, it might be beneficial for some nanotechnological applications (e.g., synthesis of nanowires) to produce the tubes in vitro. This system would offer a possibility of self-assembly of tubes with lengths that could be “tuned” by temperature. A key property of ATV in terms of nanotechnological applications is its ability to undergo defined changes in shape, and thereby perform mechanical work, in response only to variations of the temperature. Most of the other natural linear molecular motors like RNA polymerase, kinesin, and myosin utilize chemical energy derived from hydrolysis of ATP to drive respective conformational changes [5,6]. Components of nanodevices have also been developed that can be triggered and powered by an external or internal light source [7]. ATV provides a new possibility to trigger and control linear movement on nanoscale by means of thermal energy. Furthermore, temperature can be easily controlled and also external control of a device could be envisaged. The “smart” behavior of ATV could be for example

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used as part of a switchable device for translocation of nanoscale matter, as switch to modulate flow of fluids or electricity, to create a new class of sensors, or to exert localized forces on nanostructures.

ACIDIANUS FILAMENTOUS VIRUS 1 AND OTHER LIPOTHRIXVIRUSES Members of the family Lipothrixviridae have enveloped filamentous virions and they were classified into four genera due to differences in core structure, terminal structures, genomes, and possible replication strategies. Taxonomic distribution and basic characteristics of lipothrixviruses are summarized in Table 14.1. Virions of lipothrixviruses are about 24 nm in width and they vary in length between approximately 900 and 1950 nm [8–10]. Their terminal structures are unusual and differ between species. Virions of Sulfolobus islandicus filamentous virus (SIFV) have tapered ends to which are attached six thin flexible fibers (Figure 14.2b). More complex are the termini of virions of Acidianus filamentous virus 1 (AFV1) and AFV2. The latter contain two sets of filaments arranged in collar-like manner resembling a bottle brush (Figure 14.2c) and the former resemble claws (Figure 14.2a). The most probable function of these unusual structures is in adsorption of virion to the host cells. This was clearly demonstrated for virions of AFV1. The claw-like structures of AFV1 have a diameter of about 20 nm and are identical at both ends (Figure 14.2a). The “claws” are connected to the virus body by appendages, 60 nm long and 12 nm wide and linking the appendages and the “claws” is a collar-like structure 12 nm in diameter and 8 nm thick [9]. In mature virions, the claws are in their open conformation, but they close upon the pili-like appendages of the host during the adsorption (Figure 14.2a, inset). Virions have also been observed clamping to the body of another virion suggesting that any filamentous structure could

TABLE 14.1 Lipothrixviruses Genus, Species

Virion Morphology, Size

Virion Termini

Host

References

Nonflexible filament, 410 × 38 nm

n.s.

Thermoproteus tenax

[8,9]

Flexible filament, 1950 × 24 nm Flexible filament, 1200 × 20 nm Flexible filament, 2500 × 30 nm

Tapered ends with mop-like structures n.s.

Sulfolobus

[10]

Thermoproteus

[8]

n.s.

Thermoproteus

[8]

Flexible filament, 900 × 24 nm

Claw-like termini

Acidianus

[11]

Flexible filament, 1100 × 24 nm

Collar-like structure with two sets of inserted filaments

Acidianus

[12]

Alphalipothrixvirus TTV1∗ Betalipothrixvirus SIFV TTV2∗ TTV3∗ Gammalipothrixvirus AFV1 Deltalipothrixvirus AFV2

*

Presently not available in laboratory collections. Abbreviations: n.s., not studied; AFV, Acidianus filamentous virus; TTV1, thermoproteus tenax virus 1.

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FIGURE 14.2 Electron micrographs of virions of Acidianus filamentous virus 1, AFV1, (a), Sulfolobus islandicus filamentous virus, SIFV, (b), Acidianus filamentous virus 2, AFV2, (c) and schematic representation of virions. (a) In insets, claw-like structures are shown in “open” and “closed” conformation. White arrow indicates a “claw” clamped around host pili and separated from the virion body, and black arrow indicates pili-like appendices of the host cell. Scale bars represent 100 nm.

serve for the adsorption of the virus provided that its diameter is not larger than the one of the claw (T.B., personal observation). The virions seem not to be able to attach directly to the surface of the host cells or to any of the isolated components of cellular envelopes. The two “claws” of a virion are identical in their function and can be used simultaneously by the virus for the attachment. The process of attachment appears to be irreversible. The contact between pili-like appendices and “claws” seem to be rather strong because “pili” often contain knob-like structures with 18 nm in diameter, representing the viral termini which have been separated from the virus body by mechanical forces (Figure 14.2, inset). The “gripping” movement of AFV1 “claws” is fundamentally different from classical linear or rotary movements produced by known natural nanoscale molecular motors [5,11]. AFV1 “claw” can be considered as biomolecular analog to a conventional functional device, a clamp, and could provide another gadget in the toolkit for the design of new nanodevices. The terminal structures of SIFV and AFV2 may also have a potential to serve for attachment of viruses onto surfaces, producing, for example, ordered three-dimensional arrays of viral particles.

SULFOLOBUS ISLANDICUS ROD-SHAPED VIRUS 1 AND OTHER RUDIVIRUSES Sulfolobus islandicus rod-shaped virus 1, SIRV1, Sulfolobus islandicus rod-shaped virus 2, SIRV2, and Acidianus rod-shaped virus 1, ARV1 are members of the crenarchaeal viral family Rudiviridae [12–14]. The virions are rather simple in their structure compared with any known linear DNA virus. They are nonenveloped rigid rods about 24 nm in width and ranging in length from about 600 to 900 nm. Particles contain central cavity with the diameter of around 6 nm that is plugged over the terminal 50 nm with a high-density structure. Three terminal fibers protrude from each end of the virion, and each fiber is about 10 nm in length and about 3 nm in width (Figure 14.3). The fibers

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FIGURE 14.3 Electron micrograph of a virion of Sulfolobus islandicus rod-shaped virus 2, SIRV2 and its schematic representation including a model of virion structure. In inset, virion end with three terminal fibers. Scale bar represents 200 nm. (Modified from Prangishvili, D., Stedman, K., and Zillig, W., Trends Microbiol, 9, 39–43, 2001.)

are used for the direct adsorption of virions both to the host cell surface and cellular appendages. The three structural components of virions, the body, plug-like structure, and the three fibers are distinct units and can be separated from each other by mechanical forces. Virion body is a nucleoprotein complex built of double-stranded DNA and a single basic, highly glycosylated virus-encoded protein of 15.8 kDa for SIRV1 and SIRV2, and 14.4 kDa for ARV1. The nucleoprotein is arranged in a regular helical manner with periodicity of about 4.3 nm. Apparently, it assumes the same conformation in all rudiviruses because the length of the virions is proportional to the size of the packaged DNA: SIRV2, 900 ± 50 nm long, 35.5 kb DNA; SIRV1, 830 ± 50 nm long, 32.3 kb DNA; ARV1, 610 ± 50 long, 24.7 kb DNA. The DNA packaged in this way is efficiently protected from degradation in extremely hot but also very acidic (pH 1.5–3) natural environment of rudiviruses. The resistance to very low pH implies that rudiviruses could be particularly suitable as vectors for DNA delivery in gene therapy and vaccination by oral administration. Virion structure of rudiviruses is strikingly similar to that of tobacco mosaic virus, TMV, a singlestranded (ss)RNA virus [15]. TMV virions are ~300 nm in length and ~18 nm in diameter with a distinct inner channel of ~4 nm and a single coat protein that self-assembles into the rod-like helical structure. By virtue of the highly ordered helical structure and uniform composition of the virion body, rod-shaped viruses such as TMV and ssDNA bacteriophage M13 represent excellent scaffolds for the construction of nanostructured materials. In the last several years these viruses have been extensively used for the production of nanowires of different metals, semiconductors, batteries, redox, and magnetic materials [16–24]. The potential for application of rudiviruses in bionanotechnology apparently includes all these possibilities. Moreover in certain aspects rudiviruses could have advantages due to their specific features. It is noteworthy that surfaces of proteins from hyperthermophiles are generally highly charged allowing protein stabilization through ion bonds [25]. Many of the protocols developed for the attachment of various ligands such as metals, fluorophores, polymers, enzymes, redox-active complexes, and so on, make use of ordered functional groups provided by charged amino acid residues on the surface of viruses. The increased proportion of charged amino acid residues of hyperthermophilic viruses should therefore offer a wider variety of nucleation sites for surface-controlled inorganic deposition. In addition, SIRV1 and SIRV2 are extremely stable at high temperatures and can be completely inactivated only after 50 min of autoclaving at 120°C [13]. High thermal stability in conjunction with stability at very low pH of rudiviruses provides the opportunity for the development of new functionalization approaches resulting in the synthesis of new nanomaterials.

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ACIDIANUS BOTTLE-SHAPED VIRUS The virion structure of the unique bottle-like virus Acidianus bottle-shaped virus (ABV) has been studied in some detail [26]. The virions are enveloped and have an overall length of 230 ± 20 nm with a width of 75 ± 5 nm at the broad end and 4 ± 1 nm at the pointed end. The broad end contains 20 ± 2 short filaments arranged in a circular manner and inserted into a disc or a ring (Figure 14.4a). The filaments do not appear to be involved in cellular adsorption and their function remains unclear but intriguing. The bottle-like shape of virions appears to be determined by the core and not by the outer envelope because the structural integrity of the core is maintained even after the outer envelope of the virion has been partially destroyed (Figure 14.4b). The core consists of torroidally supercoiled nucleoprotein filament (Figure 14.4c). One more separate structural unit of ABV is a pointed “stopper” inserted into the narrow end of the virion. In the partially disrupted particles of ABV the torroidally supercoiled nucleoprotein has been observed directly attached to “the stopper” and its role is most probably in adsorption and injection of viral DNA into host cell (Figure 14.4c). ABV genome encodes a putative gene for a small RNA with a predicted secondary structure highly similar to that of prohead RNA of bacteriophage phi29 [27]. This RNA is an essential part of packaging machinery of the phage which is responsible for the translocation of phage DNA from the cytoplasm of the cell into the preformed capsid (reviewed in [28]). Despite the similarity of the two RNAs, the packaging mechanism of ABV a priori appears to be different from the one of phi29. In the case of phi29 and other double-stranded (ds)DNA bacteriophages where this was studied, the naked DNA is transported into preformed capsid and it is organized there as the DNA solenoid [29]. DNA of ABV, however, is apparently packaged in a profoundly different manner, and is condensed into a cone-shaped structure. Such arrangement of the nucleoprotein may be an efficient way to compress DNA and stock the energy invested in the packaging process. DNA-packaging system of phi29 is a remarkably strong molecular motor and its maximum stall force used to package phage DNA is five times greater than that of myosin fibers [30]. Such a powerful nanomotor has a potential to be incorporated into nanodevices [28]. The packaging motor of ABV may be an interesting alternative offering a different way to condense and arrange DNA in constrained nanospace.

SPHERICAL VIRUSES OF SULFOLOBUS AND PYROBACULUM Known hyperthermophilic spherical viruses are represented by two isolates, Sulfolobus turreted icosahedral virus, STIV, and Pyrobaculum spherical virus, PSV. In addition a spherical virus-like

FIGURE 14.4 Electron micrographs of virions of Acidianus bottle-shaped virus, ABV, in their native state (a), and partially disrupted (b, c). Scale bars represent 100 nm. (Modified from Haring, M., Rachel, R., Peng, X., Garrett, R. A., and Prangishvili, D. J Virol 79, 9904–11, 2005.)

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FIGURE 14.5 Electron micrographs of Pyrobaculum spherical virus, PSV (a). (Modified from Haring, M. et al. Virology 323, 233–42, 2004.) Sulfolobus turreted icosahedral virus, STIV (b). Scale bars represent 100 nm. (Courtesy of Mark Young).

particle similar in its morphotype to PSV has been isolated recently and was named Thermoproteus tenax spherical virus 1, TTSV1 [31]. The virions of STIV are enveloped icosahedra 74 nm in diameter with an internal lipid layer consisting of a subpopulation of host lipids [32]. Virion structure of STIV, has been thoroughly studied and this resulted in the first reconstruction of an archaeal viral particle at 27Å resolution [33]. The virion capsid has unique appendages extending 13 nm from each of the fivefold vertices (Figure 14.5b). The appendages are five-sided turret-like structures that have an average diameter of 24 nm and a ~3 nm wide channel in their center. Their function is probably in host recognition and/or attachment and translocation of the viral DNA into the host cell. The virions are composed of one major ~37-kDa protein, eight minor viral, and two host-encoded proteins. The application spectrum of STIV in nanobiotechnology could be similar to that of cowpea chlorotic mottle and the cowpea mosaic viruses which have been extensively used as scaffolds for attachment of a wide variety of chemical and biological ligands giving rise to diverse nanostructured materials [16,34–38]. The virions of PSV could potentially be used in the same way. They are 100 nm in diameter and contain an envelope derived from host lipids (Figure 14.5a) [39]. The electron microscopy studies of mechanically disrupted particles showed that the envelope apparently covers a nucleocapsid with a helical symmetry and width of about 6 nm. Such structure is unique for an enveloped DNA virus [40].

SULFOLOBUS SPINDLE-SHAPED AND DROPLET-SHAPED VIRUSES Virions of Sulfolobus spindle-shaped viruses, SSVs, are enveloped and have a very short tail at one of the two pointed ends [41–43] (Figure 14.6a). The tail carries fibers which serve for adsorption to

FIGURE 14.6 Electron micrographs of Sulfolobus spindle-shaped virus 1, SSV1, (a), and Sulfolobus neozealandicus droplet-shaped virus, SNDV (b). Scale bars represent 200 nm. (Courtesy of Wolfram Zillig.)

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the host membrane. SSV virions measure about 55–60 × 80–100 nm and are much smaller than Sulfolobus tengchongensis spindle-shaped virus 1, STSV1, that measures around 230 × 107 nm and has a single tail of variable length (0–133 nm) suggesting an extracellular development similar to that of ATV [44]. A spindle-shaped virus-like particle (VLP) has also been isolated from a deep-sea strain of Pyrococcus abyssi [43]. The size and overall architecture of this VLP is very similar to that of SSVs but they are not related to each other on the genome level. The structure and assembly of spindle-shaped virions has not yet been studied in detail which makes it difficult to suggest how these viruses could be used for nanobiotechnology. This is also true for virions of Sulfolobus neozealandicus droplet-shaped virus, SNDV, that have a unique, droplet-like morphotype. The virion particles measure from 110 to 185 nm in length and from 95 to 70 nm in width and are densely covered by thin fibers at their pointed ends [45]. The core is protected by a beehive-like structure, the surface of which appears to be built up of components stacked in a helical manner (Figure 14.6b). The host strain in which SNDV could be replicated stably has not been found, therefore, the virus unfortunately no longer exists in laboratory collections.

ZIPPER VIRUS-LIKE PARTICLES OF ACIDIANUS Zipper virus-like particles, ZVLP, were initially observed in enrichment cultures of the original samples from hot springs in Yellowstone National Park. They are filamentous particles, with variable lengths of several to several hundred nanometers and zipper-like surface pattern consisting of equilateral triangular subunits with side size of about 15 nm [46]. Observation of ring-like structures probably representing detached structural subunits of ZVLPs suggested that they are hollow tubes with the internal diameter of about 5 nm (Figure 14.7). Producers of these intriguing virus-like particles were several Acidianus strains from the enrichment cultures. ZVLPs were detectable in the supernatants of these strains only if the cells were exposed to stress factors like freezing–thawing cycles, mitomycin C, and ultraviolet (UV) radiation. Such treatment induced the production of huge amounts of ZVLPs also inside the cells where they formed ordered arrays and seem to have completely filled up the cellular lumen (unpublished data). Also in solution ZVLPs have affinity for each other and they tend to assemble into regular flat sheets. Proteins self-assembling into regular structures, like bacterial and archaeal S-layers are considered to have broad application potential in biotechnology. This is based on their capability to serve as template for formation of regular arrays of bound molecules and particles. For example, functionalized S-layers are currently being used as novel affinity matrices, biosensors, biocompatible

FIGURE 14.7 Electron micrographs of zipper virus-like particles, ZVLPs, produced by strains of Acidianus. The surface pattern is illustrated schematically. Arrows indicate ring-like detached structural subunits of ZVLPs. Scale bars represent 200 nm. (Modified from Rachel, R. et al. Arch Virol 147, 2419–29, 2002.)

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surfaces, drug delivery and targeting systems and for molecular electronics and nonlinear optics (reviewed in [47]).

CONCLUSIONS AND PERSPECTIVES dsDNA viruses replicating in hyperthermophilic archaea have unique features as described before, that could be exploited for technological purposes. In addition, one has to bear in mind that all these viruses survive in nature in most aggressive environment with pH values lower than 3 and temperatures above 80°C. Correspondingly, purified virus particles and also viral proteins demonstrate high thermal stability in laboratory. Clearly, the viruses of hyperthermophilic archaea have much to offer to nanobiotechnologists. The potential of these viruses should be even more enhanced with better understanding of their biology and functional characterization of their proteins which are currently under way in several laboratories. The extraordinary diversity of viral morphotypes and genomes observed in geothermally heated terrestrial sources suggests that the characterized hyperthermophilic archaeal viruses represent only a “tip of the iceberg” and that many more of these fascinating viruses are waiting to be discovered in the future.

ACKNOWLEDGMENTS The authors thank Nicole Steinmetz and Dave Evans for helpful discussions. Tamara Basta was supported by Dr Roux postdoctoral fellowship from the Institut Pasteur.

REFERENCES 1. Haring, M. et al. Virology: independent virus development outside a host. Nature 436, 1101–2 (2005). 2. Prangishvili, D. et al. Structural and genomic properties of the hyperthermophilic archaeal virus ATV with an extracellular stage of the reproductive cycle. J Mol Biol 359, 1203–16 (2006). 3. Austin, J.R., 2nd, Segui-Simarro, J.M., and Staehelin, L.A. Quantitative analysis of changes in spatial distribution and plus-end geometry of microtubules involved in plant-cell cytokinesis. J Cell Sci 118, 3895–903 (2005). 4. Mandelkow, E.M., Mandelkow, E., and Milligan, R.A. Microtubule dynamics and microtubule caps: a time-resolved cryo-electron microscopy study. J Cell Biol 114, 977–91 (1991). 5. Vogel, P.D. Nature’s design of nanomotors. Eur J Pharm Biopharm 60, 267–77 (2005). 6. Wang, M.D. et al. Force and velocity measured for single molecules of RNA polymerase. Science 282, 902–7 (1998). 7. Astier, Y., Bayley, H., and Howorka, S. Protein components for nanodevices. Curr Opin Chem Biol 9, 576–84 (2005). 8. Arnold, H.P. et al. A novel lipothrixvirus, SIFV, of the extremely thermophilic crenarchaeon Sulfolobus. Virology 267, 252–66 (2000). 9. Bettstetter, M., Peng, X., Garrett, R.A., and Prangishvili, D. AFV1, a novel virus infecting hyperthermophilic archaea of the genus acidianus. Virology 315, 68–79 (2003). 10. Haring, M. et al. Structure and genome organization of AFV2, a novel archaeal lipothrixvirus with unusual terminal and core structures. J Bacteriol 187, 3855–8 (2005). 11. Soong, R.K. et al. Powering an inorganic nanodevice with a biomolecular motor. Science 290, 1555–8 (2000). 12. Peng, X. et al. Sequences and replication of genomes of the archaeal rudiviruses SIRV1 and SIRV2: relationships to the archaeal lipothrixvirus SIFV and some eukaryal viruses. Virology 291, 226–34 (2001). 13. Prangishvili, D. et al. A novel virus family, the Rudiviridae: structure, virus–host interactions and genome variability of the sulfolobus viruses SIRV1 and SIRV2. Genetics 152, 1387–96 (1999). 14. Vestergaard, G. et al. A novel rudivirus, ARV1, of the hyperthermophilic archaeal genus Acidianus. Virology 336, 83–92 (2005).

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15. Knipe, D.M. et al. Fields Virology (Lippincott Williams and Wilkins, 2007). 16. Flynn, C.E., Lee, S.-W., Peelle, B.R., and Belcher, A.M. Viruses as vehicles for growth, organization and assembly of materials. Acta Materialia 51, 5867–80 (2003). 17. Knez, M. et al. Spatially selective nucleation of metal clusters on the tobacco mosaic virus. Adv Func Mater 14, 116–24 (2004). 18. Mao, C. et al. Viral assembly of oriented quantum dot nanowires. Proc Natl Acad Sci USA 100, 6946–51 (2003). 19. Mao, C. et al. Virus-based toolkit for the directed synthesis of magnetic and semiconducting nanowires. Science 303, 213–17 (2004). 20. Nam, K.T. et al. Virus-enabled synthesis and assembly of nanowires for lithium ion battery electrodes. Science 312, 885–8 (2006). 21. Schlick, T.L., Ding, Z., Kovacs, E.W., and Francis, M.B. Dual-surface modification of the tobacco mosaic virus. J Am Chem Soc 127, 3718–23 (2005). 22. Shenton, W., Douglas, T., Young, M., Stubbs, G., and Mann, S. Inorganic-organic nanotube composites from template mineralization of tobacco mosaic virus. Adv Mater 11, 253–256 (1999). 23. Steinmetz, N.F., Lomonossoff, G.P., and Evans, D.J. Cowpea mosaic virus for material fabrication: addressable carboxylate groups on a programmable nanoscaffold. Langmuir 22, 3488–90 (2006). 24. Steinmetz, N.F., Lomonossoff, G.P., and Evans, D.J. Decoration of cowpea mosaic virus with multiple, redox-active, organometallic complexes. Small 2, 530–3 (2006). 25. Cambillau, C. and Claverie, J.M. Structural and genomic correlates of hyperthermostability. J Biol Chem 275, 32383–6 (2000). 26. Haring, M., Rachel, R., Peng, X., Garrett, R.A., and Prangishvili, D. Viral diversity in hot springs of Pozzuoli, Italy, and characterization of a unique archaeal virus, Acidianus bottle-shaped virus, from a new family, the Ampullaviridae. J Virol 79, 9904–11 (2005). 27. Peng, X., Basta, T., Haring, M., Garrett, A.R., and Prangishvili, D. Genome of Acidianus bottle-shaped virus and insights into the replication and packaging mechanisms. Virology 364, 237–43 (2007). 28. Guo, P. Bacterial virus phi29 DNA-packaging motor and its potential applications in gene therapy and nanotechnology. Methods Mol Biol 300, 285–324 (2005). 29. Jardine, P.J. and Anderson, D.L. in The Bacteriophages (ed. Calendar, R.), pp. 49–65 (Oxford University Press, New York, 2006). 30. Smith, D.E. et al. The bacteriophage straight phi29 portal motor can package DNA against a large internal force. Nature 413, 748–52 (2001). 31. Ahn, D.G. et al. TTSV1, a new virus-like particle isolated from the hyperthermophilic crenarchaeote Thermoproteus tenax. Virology 351, 280–90 (2006). 32. Maaty, W.S. et al. Characterization of the archaeal thermophile Sulfolobus turreted icosahedral virus validates an evolutionary link among double-stranded DNA viruses from all domains of life. J Virol 80, 7625–35 (2006). 33. Rice, G. et al. The structure of a thermophilic archaeal virus shows a double-stranded DNA viral capsid type that spans all domains of life. Proc Natl Acad Sci USA 101, 7716–20 (2004). 34. Blum, A.S. et al. An engineered virus as a scaffold for three-dimensional self-assembly on the nanoscale. Small 1, 702–6 (2005). 35. Medintz, I.L. et al. Decoration of discretely immobilized cowpea mosaic virus with luminescent quantum dots. Langmuir 21, 5501–10 (2005). 36. Raja, K.S., Wang, Q., and Finn, M.G. Icosahedral virus particles as polyvalent carbohydrate display platforms. Chembiochem 4, 1348–51 (2003). 37. Soto, C.M. et al. Fluorescent signal amplification of carbocyanine dyes using engineered viral nanoparticles. J Am Chem Soc 128, 5184–9 (2006). 38. Wang, Q., Raja, K.S., Janda, K.D., Lin, T., and Finn, M.G. Blue fluorescent antibodies as reporters of steric accessibility in virus conjugates. Bioconjug Chem 14, 38–43 (2003). 39. Haring, M. et al. Morphology and genome organization of the virus PSV of the hyperthermophilic archaeal genera Pyrobaculum and Thermoproteus: a novel virus family, the Globuloviridae. Virology 323, 233–42 (2004). 40. van Regenmortel, M.H.V. et al. Virus Taxonomy: The Classification and Nomenclature of Viruses. The Seventh Report of the International Committee on Taxonomy of Viruses (Academic Press, San Diego, 2000).

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41. Wiedenheft, B. et al. Comparative genomic analysis of hyperthermophilic archaeal Fuselloviridae viruses. J Virol 78, 1954–61 (2004). 42. Zillig, W. et al. Genetic elements in the extremely thermophilic archaeon Sulfolobus. Extremophiles 2, 131–40 (1998). 43. Geslin, C. et al. PAV1, the first virus-like particle isolated from a hyperthermophilic euryarchaeote, “Pyrococcus abyssi”. J Bacteriol 185, 3888–94 (2003). 44. Xiang, X. et al. Sulfolobus tengchongensis spindle-shaped virus STSV1: virus–host interactions and genomic features. J Virol 79, 8677–86 (2005). 45. Arnold, H.P., Ziese, U., and Zillig, W. SNDV, a novel virus of the extremely thermophilic and acidophilic archaeon Sulfolobus. Virology 272, 409–16 (2000). 46. Rachel, R. et al. Remarkable morphological diversity of viruses and virus-like particles in hot terrestrial environments. Arch Virol 147, 2419–29 (2002). 47. Sara, M., Pum, D., Schuster, B., and Sleytr, U.B. S-layers as patterning elements for application in nanobiotechnology. J Nanosci Nanotechnol 5, 1939–53 (2005). 48. Prangishvili, D., Stedman, K., and Zillig, W. Viruses of the extremely thermophilic archaeon Sulfolobus. Trends Microbiol 9, 39–43 (2001).

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Part V Minimal Complexity Model Systems

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Master Keys to DNA Replication, Repair, and Recombination from the Structural Biology of Enzymes from Thermophiles Li Fan, R. Scott Williams, David S. Shin, Brian Chapados, and John A. Tainer

CONTENTS Introduction ................................................................................................................................ Master Keys to DNA Replication, Repair, and Recombination ................................................ FEN-1, PCNA, Ligase, and Okazaki Fragment Maturation ........................................... XPB Helicase and Nucleotide Excision Repair .............................................................. Mre11/Rad50 Structural and Enzymatic Roles in DSB Recognition and Initiation of DSB Repair Pathways ...................................................................... Rad51 and HR DNA Strand Exchange ........................................................................... RuvB and HJ Branch Migration ..................................................................................... Conclusions and Perspectives .................................................................................................... Acknowledgments ...................................................................................................................... References ..................................................................................................................................

239 240 241 245 247 249 255 257 257 258

INTRODUCTION Microorganisms that are able to grow at temperatures above 90°C are defined as hyperthermophiles, the majority of which are classified as archaea [1]. Genes in these thermophiles encode proteins with high thermal stability even in the form of recombinant proteins expressed in bacteria, and thus provide advantages for characterizing protein interactions, conformations, and structures. In fact, most microbial responses to the environment involve reversible protein complexes and dynamic conformations that can be extremely challenging to study in mesophilic organisms but can often be kinetically trapped for hyperthermophiles. Such hyperthermophiles therefore not only aid crystallizations for x-ray structure determinations, but furthermore may provide a 1000-fold kinetic advantage for kinetically trapping the dynamic conformations and interactions responsible for most of the protein complexes and machines controlling microbial cell biology. Recently both x-ray structures and x-ray scattering in solution measurements or small angle x-ray scattering (SAXS) [2] of thermophilic proteins have dramatically improved our understanding of life and cell biology at the molecular level. This review focuses on our recent studies plus closely related literature results on 239

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thermophilic proteins that act in DNA replication, repair, and recombination as these are prototypical systems for understanding biologically important, dynamic protein interactions and conformations necessary for cells to survive, grow, and divide. We believe that an understanding of the structure and function of these thermophilic proteins is providing us with master keys [3] to the molecular mechanisms underlying reversible protein interactions and functional conformations critical for genome maintenance and cellular responses to endogenous and environmental stress.

MASTER KEYS TO DNA REPLICATION, REPAIR, AND RECOMBINATION The survival of all species requires accurate delivery of genetic information from parent to offspring through DNA replication [4]. The process of DNA replication is functionally and often structurally conserved in all domains of life and consists of three phases: initiation, elongation, and termination. Initiation of DNA replication starts at distinct DNA sequences, the origins of replication. Initiation steps include: (i) local melting of the DNA duplex at an origin of replication; (ii) synthesis of primers for bi-directional DNA replication. These primers are then extended by DNA polymerase to start DNA synthesis using the parental DNA strands as templates. During elongation, primer extension from each side of the origin creates replication forks moving away from the origin. Helicases are required for DNA unwinding at the replication fork. During this initial elongation stage, DNA synthesis occurs on only one strand, termed the leading strand, while the other parental DNA strand become single-stranded (ss) and is protected by single-stranded DNA binding (SSB) proteins. After extension reaches a certain distance, DNA synthesis starts at the other parental strand, the lagging strand, by a primase and DNA polymerase to produce an RNA–DNA fragment, termed an Okazaki fragment. Therefore, at the replication fork, only the leading strand DNA is used as a template for continuous 5′–3′ DNA synthesis by DNA polymerase, while the lagging strand DNA is periodically used for synthesis of Okazaki fragments. The movement of the replication fork controlled by helicases is coordinated with the leading strand DNA synthesis, and DNA syntheses on both the leading strand and lagging strand are also well coordinated to guarantee a harmonic elongation phase. To complete elongation, the RNA primers of Okazaki fragments are removed, a process involving flap endonuclease 1 (FEN-1) and DNA polymerase. The DNA fragments are then connected by ligase. It is essential for replicative DNA polymerases to have high fidelity and processivity to assure high accuracy and efficiency for genomic DNA replication. All the replicative DNA polymerases have 3′–5′ exonuclease activity for editing. They also have significant intrinsic processivity when compared with DNA polymerases involved in DNA repair. Additional factors called processivity factors, for example, the “sliding clamp” proliferating cell nuclear antigen (PCNA), are usually attached to replicative DNA polymerases to dramatically enhance their processivity. Genomic DNA is constantly attacked by numerous damaging agents arising from both inside the cell and outside the environment. The integrity of genomic information is guarded by a number of DNA repair pathways [5]. Different types of DNA damages are repaired by six major damage-specific DNA repair pathways: (i) direct repair (DR) for O6-alkyguanine; (ii) base excision repair (BER) for base damages caused by oxidation, depurination, or deamination; (iii) nucleotide excision repair (NER) for bulky DNA lesions caused by ultraviolet (UV)-radiation, chemicals, and protein-DNA adducts; (iv) mismatch repair (MMR) for base mismatches from replication errors and single base deletion or insertion; (v) recombinational repair (RER) including both homologous recombination (HR) and nonhomologous end-joining (NHEJ) pathways for repair of double-stranded DNA breaks (DSBs); and (vi) translesion synthesis (TLS) by TLS polymerases to bypass pyrimidine dimmers, 8-oxoG DNA lesion, and apurine sites. All these pathways follow the same overall strategy: damage localization, followed by elimination of damages, and final restoration of the normal DNA duplex. Table 15.1 lists the major protein factors involved in eukaryotic DNA repair pathways. We discuss here the structure and function of some of these key factors from thermophiles.

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TABLE 15.1 Key protein Factors in Eukaryotic DNA Repair Pathways NER DR AGT PHR1 MGT1

RER

BER

GGR

TCR

MMR

HR

NHEJ

TLS

MYH MAG1 UNG OGG1 NTG1 MTH1

DDB XPE XPC hHR23B

CSA CSB XPAB2

MSH2/3/6 MLH1/3 PMS1/2 EXO1

ATM

Ku70 Ku80 DNA-PKs

Polθ Polι Polη Polξ

Mre11 Rad50 Nbs1 Rad51 Rad52 Rad54 Rad55 Rad57 BRCA1 BRCA2 MMS4 MUS81

Mre11 Rad50 Nbs1

RFC PCNA Polδ/ε Ligase 1

Pol μ XRCC4 Ligase 4

APE1

RFC PCNA Polδ/ε FEN1 Ligase 1

PARP PAR Polβ XRCC1 Ligase 3

RPA XPA XPB XPD

XPG XPF-ERCC1

RFC PCNA Polδ/ε FEN1 Ligase 1

RFC PCNA RPA Polδ/ε Ligase

Abbreviations: For DNA repair pathways: DR, direct repair; BER, base excision repair; NER, nucleotide excision repair; GGR, general genomic repair; TCR, transcription-coupled repair; MMR, mismatch repair; RER, recombinational repair; HR, homologous recombination; NHEJ, non-homologous end-joining; TLS, translesion synthesis.

FEN-1, PCNA, LIGASE, AND OKAZAKI FRAGMENT MATURATION FEN-1 is a prototypic structure-specific nuclease that is central to both DNA replication and repair processes. During DNA replication and repair, a complex that includes both FEN-1 and PCNA removes RNA primers or damaged DNA, generating a product for ligation by DNA ligase I [6–8a]. Several lines of evidence underscore the importance of FEN-1 activity in DNA replication and repair pathways. FEN-1 homozygous knockouts are lethal in mice, and mice with haplo-insufficiency FEN-1 (FEN-1/null) exhibit accelerated tumor growth [9]. Deletions of FEN-1 in Saccharomyces cerevisiae (rad27) cause replication and repair defects, including increased sensitivity to UV light and chemical mutagens, genomic instability, increased tri-nucleotide repeat expansion, and destabilization of telomeric repeats (see review [10]). Unlike endonucleases that recognize a specific DNA sequence, FEN-1 recognizes a specific DNA structure, independent of the DNA sequence. Specifically, FEN-1 recognizes a branched DNA structure consisting of a single unpaired 3′ nucleotide (3′-flap) overlapping with a variable length region of 5′-single-stranded DNA (5′-flap) [11,12]. This “doubleflap” or “overlap-flap” structure results from DNA polymerase activity that displaces a RNA primer or damaged DNA creating an ssDNA 5′-flap. The newly synthesized DNA and the displaced region compete for base pairing with the template strand, resulting in the formation of the double-flap structure [13]. FEN-1 cleaves this substrate after the first base pair preceding the 5′-flap to remove the ssDNA 5′-flap [11,12,14] and creates a nicked DNA product for ligation. The FEN-1 class of structure-specific 5′-nucleases occurs in all domains of life [15,16]. Crystal structures are available for FEN-1 homologs from archaea Pyrococcus furiosus (Pf FEN-1) [17],

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Methanococcus jannaschii (MjFEN-1) [18], and Archaeglobus fulgidus (Af FEN-1) [19]. FEN-1 is a saddle-shaped, single-domain α/β protein (~60 Å × ~45 Å × ~40 Å) with a 20 Å deep groove along one face formed from the central seven-stranded β-sheet, an antiparallel β-ribbon, and two α-helical bundles. The C-terminal edge of the β sheet is identified as the substrate-binding region by the presence of catalytically important residues. However, the binding site for a 3′-flap and associated dsDNA is situated ∼25 Å away from the nuclease active site in the crystal structure of Af FEN1 bound to the 3′-upstream portion of a double-flap DNA substrate [19]. Af FEN-1 interacts with DNA through a surface-exposed “hydrophobic wedge.” The direct interactions between the hydrophobic wedge and the 3′-flap interrupt the DNA helix, likely preventing a continuous linear DNA conformation and separating the 5′-flap and associated duplex away from the 3′-flap. Additional experiments support a model in which FEN-1 binding kinks the DNA substrate by ~90° suggesting how 3′ flap binding might contribute to cleavage specificity [19]. Specific contacts to DNA minor groove and backbone atoms on both strands anchor the 3′-flap in a small pocket that sterically blocks binding of additional nucleotides. DNA binding is mediated by residues conserved in all known FEN-1 homologs. These residues emanate from two pairs of α-helices and the two loops. Hydrophobic packing and hydrogen bonding interactions with the 3′-terminal sugar, but not the associated base, allow sequence-independent recognition of 3′-nucleotides, consistent with the role of FEN-1 as a structure-specific endonuclease. The 3′ flap binding site locating about 25 Å away from the active site suggests that FEN-1 could track along the 5′ flap, but not efficiently catalyze phosphodiester cleavage until 3′ flap binding promotes the ordering of the helical clamp closing over the active site with the substrate properly positioned [19]. The predominant α-helical structure of the helical clamp region (Figure 15.1) observed in the FEN-1:DNA cocrystal structure is either disordered or adopts drastically different conformations in crystal structures of FEN-1 homologs determined without DNA [17,18]. Consistent with these crystal structures, biochemical and spectroscopic data indicated both conformational changes and an increase in α-helical content upon FEN-1 binding to DNA [20]. Mutational analyses of residues in this region suggest that conformational flexibility of the helical clamp is important for catalysis [21]. Together, these results suggest that ordering of the helical clamp region is coupled to FEN-1 conformational changes promoted by the specific recognition of the 3′-flap region of duplex DNA. In cells, FEN-1 forms a complex with PCNA, which exists as a ring-shaped homotrimer in solution [22]. Cocrystal structures [19] of both an Af FEN-1 peptide and consensus FEN-1 peptide bound to Af PCNA revealed two adjacent but structurally distinct motifs for PCNA interactions with FEN-1. FEN-1 interacts with PCNA mainly through a conserved, eight-residue PCNA-interaction motif (PIM: Q-X-X-L/I/M-X-X-F/Y/W-F/Y/W) located near the C-terminus. The C-termini of both FEN-1 peptides (TLERWF or TLDSFF) adopt a 310 helical conformation and bind within a hydrophobic pocket on PCNA formed by residues from the interdomain-connecting loop (IDCL) and nearby β−strands [19]. This is consistent with other PCNA:peptide cocrystal structures [23– 25]. The hydrophobic pocket on the PCNA surface functions as an anchor to attach replication and repair enzymes to the PCNA trimer. In addition, the peptide residues (KSTQA or KTTQS) preceding the conserved PCNA-binding motif form an antiparallel β-strand pair [19], termed β−zipper, with residues at the C-terminus of PCNA. The β−zipper formation causes a significant movement of the C-terminus of PCNA, and transforms the flexible unstructured C-terminus of AfPCNA into an ordered β-strand. The FEN-1 residues involved in the β−zipper connect a DNA-binding helix to the conserved PCNA-binding motif. These structural interactions are also conserved in interactions between human FEN-1 and human PCNA [26]. The cocrystal structures of the AfFEN-1:DNA complex and AfPCNA:FEN-1-peptide complex together support a specific model for FEN-1 localization at the DNA replication and repair locus [19]. The composite model positions FEN-1 on the polymerase binding front face of PCNA, with the upstream duplex DNA protruding through the central cavity of PCNA and the downstream DNA kinked ∼90o, orthogonal to the upstream duplex [19]. The FEN-1 hydrophobic wedge opens the DNA helix, enforcing a kink that facilitates 3′- and 5′-flap recognition. This kinked DNA

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FIGURE 15.1 (See color insert following page 178.) Archaeal protein structures prompted the proposal of a rotary-handoff mechanism mediated by proliferating cell nuclear antigen (PCNA) for sequential transition of DNA from DNA polymerase, to flap endonuclease (FEN)-1, and to ligase in Okazaki fragment maturation. SsPCNA (PDB code, 2HII) is presented in three colored surfaces (white gray, gray, and black). The backbone of DNA strands are in lines. To start DNA synthesis, PCNA is loaded to the 3′-end of a primer by the clamp loader (not shown). Binding of DNA polymerase (gray) to PCNA1 bends the template strand for DNA synthesis. When this complex meets the 5′-end of the adjacent Okazaki fragment, it displaces a short fragment to create a double-flap structure and hands the DNA over to FEN-1 (white gray glove shape) bound to PCNA2. FEN-1 cleaves the flapped 5′-fragment and hands over the nicked DNA to ligase (white gray C-shape) bound to PCNA3. DNA ligase then covalently connects the two Okazaki fragments together. This mechanism requires the kinked DNA rotates around the three PCNA subunits to interact with different enzymes at different stages of reactions. DNA polymerase, FEN-1, and ligase can bind to PCNA simultaneously as described for SsPCNA. The interactions of DNA with different enzymes are therefore regulated by the flexible interactions between distinct PCNA subunit and each enzyme through conformational changes. In other systems, these three enzymes may bind sequentially to PCNA to fulfill their distinct role during the process. The structure of AfFEN-1:DNA complex (PDB code, 1RXW) is presented in ribbon diagram with the helical clamp highlighted in magenta and DNA in sticks. The structure of SsLig (PDB code, 2HIV) is also presented in ribbon diagram with three domains colored differently. In addition, the structure of human ligase 1:DNA complex (PDB code, 1X9N) is presented in ribbon diagram with dsDNA in gray, and a bound adenosine monophosphate in the sphere at the active center.

conformation may be a feature of other PCNA complexes. For example, the gap-filling complexes of polymerase β [27], which is known to bind PCNA [28], show a kinked DNA topology that facilitates DNA end discrimination. The open kinked forms of DNA bound by FEN-1 and DNA polymerase may allow rotation of DNA substrates about the DNA phosphate bond opposite the 3′-flap. Rotation of the kinked DNA substrate would allow enzymes bound at any of the three binding sites on the PCNA trimer to access the kinked DNA intermediate (Figure 15.1). This observation suggests a possible PCNA-mediated, rotary-handoff mechanism [19].

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Most archaeal and eukaryotic PCNA homologs are homotrimers consisting of three identical subunits. However, PCNA from Sulfolobus solfataricus is a heterotrimer, consisting of three distinct subunits (SsPCNA1–3) [29]. Distinct SsPCNA subunits contact DNA polymerase, FEN-1, or DNA ligase, imposing a defined architecture at the lagging strand fork [29]. Crystal structures of SsPCNA reveal the physiochemical basis for assembling a heterotrimeric PCNA molecule [30–32]. Each SsPCNA subunit consists of two topologically similar domains that are arranged in a head-to-tail fashion and connected by a 10–15-residue IDCL. However, each of the three subunits differs significantly in electrostatic character near the intersubunit interface, creating complementary interactions that promote the formation of the heterotrimer [31]. Together, the distinctive curvatures of the three subunits create an asymmetric ring with a central hole [30]. Distinct SsPCNA subunits contact DNA polymerase, FEN-1, or DNA ligase, imposing a defined architecture at the lagging strand fork to tightly couple DNA synthesis and Okazaki fragment maturation [29]. DNA ligases catalyze DNA joining by a conserved, three-step reaction [33] in which the adenosine monophosphate (AMP) group of a high-energy cofactor (either ATP or NAD+) is transferred to an active site lysine (step 1) and then reacts with the phosphorylated 5′-end of DNA (step 2). The adenylate moiety serves to activate the 5′-phosphate for reaction with an adjacent 3′-OH end during step 3, with the release of AMP and the ligated DNA product from the enzyme. A single active site catalyzes these three different phosphoryl transfer reactions [34,35], and the flexible, multidomain structure of DNA ligases facilitates different conformations of the enzyme during the course of the reaction [35]. The catalytic core of ATP-dependent ligases comprises an adenylation domain (AdD), which harbors the adenylate group and most of the catalytic residues, and an OB-fold domain (OBD) that stimulates AMP transfers during step 1 and step 2 [36,37]. In complex with DNA, conserved residues on one face of the OBD engage the DNA substrate, whereas residues on the opposite face of the OBD (conserved motif VI) assist step 1 adenylation. This requires the OBD to swivel between two active conformations during the course of DNA end joining. An N-terminal DNA-binding domain (DBD) found in eukaryotic and archaeal DNA ligases binds nonspecifically to DNA and also positions the AdD and OBD on DNA to complete a ring-shaped protein structure encircling the DNA [38]. This closed conformation of the enzyme must open to permit the release of products and enable multiple turnovers. The crystal structure of the S. solfataricus DNA ligase (SsLig) revealed three domains arranged in an extended, “open” conformation (Figure 15.1), which is different from the compact ring-shaped arrangement of the homologous human enzyme DNA ligase I (hLig1) bound to nicked DNA [38]. The relative orientations of the AdD and DBD are invariant in the open and closed conformations of these DNA ligases, whereas the OBD is oriented differently on and off DNA. The OBD of the related enzyme hLig1 was shown to engage DNA in the minor groove opposite a nick situated between the AdD and DBD [38]. In the absence of DNA, the OBD of SsLig is turned away from the AdD. SAXS data support the open conformation of SsLig in solution. One molecule of SsLig binds to the SsPCNA trimer, forming a stable 1:1 complex [29] that can be purified by gel filtration chromatography. This binding stoichiometry results from the selective interaction of SsLig with the PCNA3 subunit of SsPCNA [29]. Residues within the DBD of SsLig contribute strongly to the binding interaction with SsPCNA. The amino acid sequence surrounding Phe110 and Leu111 resembles a canonical PIM that could insert into the hydrophobic pocket adjacent to the IDCL of PCNA3. These residues are important for the stimulation of DNA end-joining activity by SsPCNA. The PIMlike motif of SsLig has a single inserted residue (Ser104) not present in other PIMs that might help to target SsLig specifically to PCNA3. SAXS data suggested that SsLig remains the extended open conformation when associated with the heterotrimeric PCNA [30,8a]. The initial encounter with PCNA could tether ligase near the DNA and trigger a switch to the closed conformation of the enzyme wrapping around DNA that passes through the PCNA ring. The closed ring-shaped conformation of ligase catalyzes DNA end-joining reaction that is strongly stimulated by PCNA. This open-to-closed switch in the conformation of DNA ligase is accommodated by a malleable interface with PCNA that serves as an efficient platform for DNA ligation (Figure 15.1).

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XPB HELICASE AND NUCLEOTIDE EXCISION REPAIR The helicase XPB is a subunit of the general transcription factor TFIIH complex [39], which plays key roles in both transcription and nucleotide excision repair. The ATPase and helicase activities of XPB are required for the promoter DNA melting [40] and clearance [41] in transcription initiation by RNA polymerase II. TFIIH is a complex of 10 subunits including two helicases XPB and XPD [42]. Both XPB and XPD are required to unwind DNA duplex around lesions during nucleotide excision repair [43], which repairs a broad spectrum of DNA helix-distorting damage: for example, UV light-induced pyrimidine dimers and bulky chemical adducts [44,45]. The biological importance of XPB helicase is attested by clinically relevant XPB mutations [46]. Mutations in the human XPB gene are associated with three hereditary diseases: xeroderma pigmentosum (XP), Cockayne’s syndrome (CS), and trichothiodystrophy (TTD). These diseases are characterized by high skin and eye photosensitivity manifested at different levels, and neurological and developmental anomalies [47]. NER undergoes a sophisticated mechanism of dual incision DNA repair. There are two NER sub-pathways [48,49], DNA lesions on the transcribed strand of active genes may block the elongation of RNA polymerase II and are rapidly repaired by the so-called transcription-coupled repair (TC-NER) pathway. The stalled RNA polymerase II is recognized by CSB and XPG [50,51], which may remodel RNA polymerase II and facilitate the recruitment of TFIIH [50]. After TFIIH is recruited to the lesion, the XPB and XPD helicases unwind the DNA duplex around the lesion driven by ATP hydrolysis. The resulting DNA bubble is stabilized by XPA and RPA [52], and presents an optimal substrate for two endonucleases: XPG [53] and XPF-ERCC1 [54]. XPG and XPF incise the damaged DNA strand at 3′ and 5′ ends to the lesion, respectively. The resulting gapped DNA is refilled by DNA polymerase [55] and rejoined by DNA ligase. DNA lesions on other genomic regions are removed more slowly by the global genome NER (GG-NER). These lesions are first recognized by XPCHR23B [52]. Some studies also suggested that XPA and RPA are possibly involved in this step. TFIIH is then recruited to this “marked” lesion, and unwind the DNA duplex through the two helicase activities of XPB and XPD. The resulting DNA bubble recruits XPA and RPA, followed by XPG and XPF-ERCC1. The dual incision carried by XPG and XPF goes hand in hand with DNA re-synthesis by DNA polymerase [52]. Upon the final arrival of XPF-ERCC1, TFIIH is released and remains functionally active to participate not only in a new round of productive NER, but also in transcription mediated by RNA polymerase II as revealed by both in vivo and in vitro studies [52,56]. Recent developments in structural and biochemical characterization of XPB helicase started to address some key questions on the mechanism underlying the functions and roles of XPB in transcription and DNA repair [57–62]. Archaeoglobus fulgidus XPB homolog (AfXPB) shares 42% amino acid sequence similarity with the central region of human XPB, suggesting that the core XPB structure is conserved from archaea to humans. Crystal structures [57] are available for three AfXPB polypeptides including an N-terminal proteolytic fragment (N-AfXPB), the full-length AfXPB, and the C-terminal half construct (C-AfXPB). The combined structural information from these structures revealed unexpected details about the structure and functions of the AfXPB, possibly XPB helicase in general [57]. AfXPB consists of three consecutive domains (Figure 15.2) including the N-terminal domain and two RecA-like helicase domains (HD1 and HD2) bearing seven conserved helicase motifs characteristic of members of the helicase superfamily 2 [63]. The N-terminal domain shows a structural similarity to the mismatch recognition domain of the MER protein MutS [64]. This domain allows the N-AfXPB fragment to interact with some forms of damaged DNA; so it was named as the damage recognition domain (DRD) [57]. However, the DRD lacks the mismatch-specific Phe residue found in MutS. Instead of interacting with a specific lesion, the DRD of AfXPB likely recognizes distortions in the DNA, in agreement with the broad spectrum of DNA damage repaired by NER. The structure of C-AfXPB uncovers the presence of a thumb domain (ThM) inserted in helicase domain HD2. The ThM domain is predicted to bind DNA in a nonsequence-specific manner via the

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FIGURE 15.2 The structure and biochemistry of archaeal AfXPB supports XPB’s role in initiation of DNA unwinding at lesion sites during NER identifies functionally important conformations and damaged DNA recognition and supports its role in initiation of DNA unwinding at lesion site. Top, the structure of AfXPB (PDB code, 2FWR) and a proposed closed conformation are presented in gray ribbons with functional motifs and domains labeled. The structure of HCV helicase NS5:DNA complex (PDB code, 1A1V) is presented in light gray without the DNA shown. Bottom, cartoon presentation of the proposed mechanism for XPB to initiate dsDNA unwinding at the lesion site (see text for details).

phosphodiester backbone, based on its similarity with the ThM of DNA polymerases and several conserved positively charged amino acid residues at the interface between the ThM and HD2 domains [57]. The full-length structure of AfXPB revealed that N-AfXPB and C-AfXPB are connected by a long flexible loop. The AfXPB structure also reveals a highly conserved XPB-unique RED motif adjacent to helicase motif III. Mutational analysis of this motif suggests that the RED motif plays a critical role in DNA unwinding [57]. Large conformational changes in helicases are known to be required for translocation along the duplex DNA and are coupled by ATP hydrolysis [65–67]. AfXPB seems to follow this general trend. The relative orientation of the two helicase domains HD1 and HD2 observed in the full-length AfXPB is different from the “closed” conformation observed in crystal structures of nucleotide-bound helicases, suggesting that a significant

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reorientation of the two helicase domains would have to take place to bring the functional helicase motifs to the active cleft. These recent developments of structural and biochemical characterization on XPB lead to a proposed mechanism (Figure 15.2) for the involvement of XPB in the unwinding of duplex DNA at sites of DNA repair. When XPB is recruited to DNA, the DRD domain is proposed to recognize the distorted and damaged DNA. This interaction induces a reorientation of helicase domain HD2 via a rotation of ~170°, and allows XPB to wrap around the DNA. In this new “closed” configuration, the RED motif would be ideally placed at the helicase active site with the side chains intruding into the distorted DNA duplex. The ThM domain now “grips” one strand of the DNA above helicase domain HD2, whereas the other strand may lie in the groove on the opposite side of the RED motif. In this position, the RED motif would function as a “wedge” to unzip the DNA when ATP hydrolysis drives XPB to move along the duplex DNA during NER. It is noticed that DNA melting by XPB during transcription initiation is possibly mediated through an unconventional helicase mechanism [68], in which XPB functions as a molecular “wrench:” rotating downstream DNA relative to the fixed upstream protein-DNA interactions. Therefore, the conformation observed in the full-length AfXPB crystal structure may represent a “transcriptional mode” of XPB tuned for this action, whereas the domain reorientation is NER-specific and only occurs upon the interactions of the DRD with damaged DNA. If these mechanisms are true, the conformation of XPB will decide whether TFIIH functions as a transcription factor or a DNA repair factor. In other words, XPB will help TFIIH switch pathway selection for transcription or DNA repair whenever it is recruited to the DNA.

MRE11/RAD50 STRUCTURAL AND ENZYMATIC ROLES IN DSB RECOGNITION AND INITIATION OF DSB REPAIR PATHWAYS The phylogenetically conserved and essential Mre11/Rad50 complex (Mre11/Rad50/Nbs1) in higher eukaryotes is a key player in DNA HR repair, NHEJ, and telomere maintenance. Structural, biochemical, and cell biology data suggest that Mre11/Rad50/Nbs1 (MRN) function is elaborate, and serves in these diverse capacities by acting as a DNA damage sensor, an enzymatic effector in DNA damage repair, and as a transducer of critical signals to the cell-cycle checkpoint apparatus (see review [69]). Defects in the MRN complex cause cancer predispositions in humans and severe phenotypes in yeast, revealing the importance of the MRN three-member complex in cell biology. The importance of the MRN complex is further underscored by the fact that null mutations in any of the three proteins lead to embryonic lethality in mice [70–72], which is not surprising as the complex participates in nearly every facet of DNA DSB metabolism–DSB detection and processing, HR, NHEJ, telomere maintenance, and cell cycle checkpoint signaling. Biochemically, the MRN complex is an ATP-stimulated nuclease that acts on ssDNA and hairpins and resects dsDNA in a 3′ to 5′ direction, suggesting it may not be directly involved in generation of 3′ tails for HR although this possibility cannot be completely discounted [73,74]. The intensive ongoing search for a unifying function for MRN has led to evidence that the complex serves in part as a multipurpose DNA tether which acts to bridge severed DNA ends [75–78]. Our detailed understanding of MRN anatomy has precipitated from crystallographic snapshots of archael subcomplex components [77,79–81] and from electron and atomic force microscopic imaging of the intact archael and eukaryotic Mre11/Rad50 homologs [75,76] (Figure 15.3). The core Mre11-Rad50 (MR) complex exists as a heterotetrameric assembly (M2R2), whose morphology is divided into distinct regions designated as the head, coil, and hook domains (Figure 15.3). The globular DNA binding head is likely comprised of two Rad50-ATPase domains and the dimeric Mre11 nuclease that is physically bound to the base of the Rad50 coiled-coils [82]. Mre11 is the central protein–protein and protein–nucleic interaction module of the complex, as it binds Rad50, eukaryotic Nbs1, DNA and to itself via poorly understood mechanisms. Mre11 has Mn2+-dependent ssDNA and dsDNA endonuclease, dsDNA 3′ → 5′ exonuclease and DNA annealing

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FIGURE 15.3 (See color insert following page 178.) Archaeal protein structures revealed the architecture of the Mre11/Rad50 complex. Center: the Mre11/Rad50 complex assembly formed by heterotetramerization of Mre11/Rad50 (M2R2). Larger complexes 2X(M2R2) observed by negative stain electron microscopy through M2R2 intercomplex hook–hook interactions. Archael structures of the Mre11/Rad50 subcomplexes are highlighted by boxed regions: (a) structure of the Rad50 Zn-hook domain. CXXC motifs coordinate Zn2+ ions to bridge the apices of the Rad50 coiled coils and facilitate long-range DNA tethering; (b) structure of the Mre11 phosphoesterase domain bound Mn2+ and a 5′-adenosine monophosphate (AMP) nucleotide reaction product; (c) structures of ATP bound (top) and apo-Rad50 minimal ATPase domain. Nucleotide binding-induced dimerization of Rad50 ATPase halves within the M2R2 DNA-binding head. Hydrolysis causes dimeric ATPase release and a dramatic conformational twisting to the ATPase-N domain relative to ATPase-C domain.

and unwinding activities in vitro [73,74,80,83–87]. In vivo, Mre11 appears to participate in processing DNA ends needed for homologous recombination repair (HRR) in concert with other nucleases [74,76,83,85,86,88,89]. Mre11 nuclease activity is modulated through its interactions with Rad50 and Nbs1 [73,85,90–92]. Our P. furiosus Mre11 structure revealed five conserved phosphoesterase motifs located on amino-terminal end of the protein [79] (Figure 15.3). The PfMre11:Mn2+:dAMP complex structure suggests that Mre11 cleaves DNA ends 3′ → 5′, liberating 5′-phosphorylated nucleotides, consistent with the biochemically observed products. No significant 5′ → 3′ activity has been observed biochemically for Mre11 [83,86]. Our structure therefore supports the 3′ → 5′ nuclease direction as the main dsDNA exonuclease activity of Mre11. These results indicate that the generation of 3′ tails in HR in vivo requires an additional 5′ → 3′ nuclease, as suggested by genetic data [89], or the that nuclease direction of Mre11 is modulated in vivo by as yet uncharacterized factors. Each Rad50 polypeptide assembles with the intramolecular collapse of an expansive antiparallel coiled-coil which conspicuously emanates from the head domain, and measures ~500 Å long for eukaryotic Rad50 homologs. The protein has two classic ATP-binding cassettes (ABC), ATPase type Walker A and Walker B motifs. What makes the protein unusual is that these motifs are separated by a ~1200 Å (in humans) coiled-coil generated by heptad repeats [80,81,93]. The extreme Rad50 N- and C-terminal ends coalesce to form a bipartite ABC-ATPase. Coexpression

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of the N- and C-termini of the P. furiosus Rad50 protein demonstrated that these regions stably associate [81]. The bipartite ATPase domain can dimerize further in an ATP- and Mg2+-dependent manner. Two molecules of ATP bind at the dimeric interface, and are sandwiched between the catalytic and ATP binding ABC-ATPase conserved Walker A, Walker B, and signature motifs [81]. ATP hydrolysis liberates dimerization, and results in dramatic conformation twisting of the ATPase N-terminal half relative to the C-terminal half. The central Zn-coordinating hook domain of Rad50 is further crucial for activity of the MRX complex in yeast [77,94,95]. This domain adopts an about-turn to reverse directionality of each Rad50 coil, and caps the distal end of the coiled-coils with a CXXC Zn-hook motif that can mediate additional Zn2+-mediated Rad50– Rad50 combinatorial interactions to facilitate dynamic, long-range DNA tethering between M2R2 complexes to create M4R4 oligomeric assemblies during double-strand break repair. Despite significant advances, key questions regarding Mre11/Rad50 structure/function remain unresolved. Of paramount importance will be resolution of structural nature of the heterotetrameric Rad50/Mre11 assembly and its mode of interaction with DNA. These DNA scaffolding interactions mediate the critical expeditious double-strand break recognition, tethering, and cell cycle signaling responses following sensing of DNA damage [95,96]. The precise functional roles for Rad50 ATP-induced conformational rotations also remain obscure. Indeed, chemomechanical transmission of DNA and ATP-induced Rad50 conformational changes, and auxiliary interactions through the Zn-hook motifs within eukaryotic MRN complexes appear to lie at the core of eukaryotic DSB signaling activation process [78,94,95,97,98]. Understanding these Mre11/Rad50 ATPase-mediated structural transitions and the biochemical means for stimulating these motions will thus be key to understanding DSB sensing and downstream signaling. To date archaeal Mre11/Rad50 assemblies have solely provided the critical reagents for the elucidation of core Mre11/Rad50 architectures and structural biology. Extension of our understanding beyond this core to more complex eukaryotic Mre11/Rad50/Nbs1 assemblies will require delineation of structurally tractable eukaryotic MRN subcomplexes. Damage sensitivity observed in the absence of any MRN member makes MRN an attractive target for inhibitors to increase sensitivity of cells to ionizing radiation and other DNA-damaging agents. MRN structure–activity relationships therefore form a basis for a deeper understanding of MRN biological functions and of the possibilities for targeting MRN for future cancer therapies.

RAD51 AND HR DNA STRAND EXCHANGE Rad51 plays an essential role in the repair of DNA double-strand breaks that, if not repaired, can lead to programmed cell death, gross chromosomal rearrangements or chromosomal loss, thus threatening genome stability and leading to several human diseases including cancer [99–101]. DSBs are repaired with fidelity by HR, a repair pathway that uses homologous DNA segments as replication templates to facilitate rejoining of broken DNA ends in meiotic and mitotic cells [99,100,102]. Despite decades of genetic, biochemistry, and biophysics research, the mechanism of HR DNA strand exchange reaction is still poorly understood. Recent progress in structural work defining the roles of the proteins involved is partially attributed to the similarity between the central enzymes involved between the kingdoms and the simplicity, stability, and ease of use of hyperthermophilic proteins for biophysical and structural charachterizations [103]. The homologous recombination pathway is very complex (Figure 15.4a). Following the formation of a DNA double-strand break, individual Rad51 subunits form helical nucleoprotein filaments that catalyze DNA pairing and strand exchange in concert with other proteins termed mediators [104,105]. In most cells, DSBs are resected by a ssDNA exonuclease to yield 3′-ssDNA overhangs, which are then protected by proteins, such as single-strand DNA-binding protein (SSB) or replication protein A (RPA) [106]. In eukaryotes, Rad51 then displaces RPA, a process that is facilitated by Rad52. Rad54, a Swi2/Snf2 family member, is implicated in assembly [107] and disassembly [108] of Rad51 nucleoprotein filaments. Disassembly of Rad51:dsDNA filaments is

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FIGURE 15.4 The archaeal Rad51 ATPase structure, polymerization motif, and the homologous recombination pathway. (a) Homologous recombination pathway. Following DNA double-strand breakage, the broken ends are processed to yield 3′-overhangs, which are coated by single-strand binding proteins. The DNA strand exchange protein Rad51 or RadA (eukaryotes/archaea) then displace the protective single-strand binding proteins and with the help of mediator proteins, invade a homologous duplex whose complimentary strand will serve as a template for synthesis of new DNA. (b) Gene organization of DNA strand exchange proteins from archaea (Pf Rad51), human (HsRad51), yeast (ScRad51), and bacteria (EcRecA). Domains are colored differently, where ND = N-terminal domain (archaea and eukaryotes only), AD = ATPase domain, CD = C-terminal domain (bacteria only). Archaea and eukaryotes also have highly charged disordered N-terminal leaders. Walker A and B motifs of the ATPase domain are shown. Continued

thought to facilitate the later stages of HR. In archaea, the Rad51 or RadA proteins fill the role of eukaryotic Rad51. Following formation of the nucleoprotein filament, a homologous DNA segment is located and is invaded by the coated ssDNA overhang. The overhang is then paired with its complement DNA to form a joint molecule. The complement DNA is then used as a template for the synthesis of DNA of the correct sequence by a DNA polymerase system. In the thermophilic

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FIGURE 15.4 (Contiued) (c) A PfRad51 subunit and a EcRecA subunit show the major differences in domain organization. Both share the ATPase domain only. DNA binding regions (DNA), nucleotide active site, and polymeriztion motif (PM) are labeled.

archaea, the polD polymerase [109] has been suggested as one of the enzymes that performs this function. During the process, complex DNA cross-overs may occur, and these are resolved by Holliday junction (HJ) “resolvases.” Archaeal Rad51/RadA homologs [110] generally share more than 40% primary sequence identity with eukaryotic Rad51, and these enzymes share similar overall domain architecture (Figure 15.4b). Archaeal and eukaryotic Rad51 proteins are more conserved between themselves than with bacterial RecA, with which they share only ~20% sequence identity limited to the ATPase domain (AD). In addition, RecA has a C-terminal domain (CD) not present in Rad51/RadA proteins, and RecA lacks the Rad51 N-terminal domain (ND). RecA exists as a multisubunit polymeric filament [111,112], while Rad51/RadA exists primarily as polymeric rings. In the presence of DNA, both RecA and Rad51 subunits coat DNA to form helical nucleoprotein filaments, which are believed to be the recombination active form of the proteins [113–115]. However, some Rad51/RadA proteins also bind DNA as rings. The first full-length Rad51 x-ray crystal structure was derived from the hyperthermophilic archaeon P. furiosus PfRad51 [116]. PfRad51 has a relatively small N-terminal domain (residues 35–91) and a larger C-terminal ATPase domain (residues 112–349) (Figure 15.4c). These domains interact weakly with each other. A protruding 19-residue amino acid linker (residues 92–111) is bent by ~90o and connects the N- and C-terminal domains, possibly providing structural flexibility while at the same time stabilizing subunit:subunit interactions in Rad51 rings and DNA-bound filaments. The PfRad51 N-terminal domain is a four α-helix bundle similar to a helix-hairpin-helix or HhH motif [117], which binds DNA phosphate backbones. The structure of PfRad51 C-terminal ATPase domain is nearly identical to the yeast (ScRad51) [118] and human Rad51 ATPase domains (HsRad51) [119], but less similar to the ATPase domain of Escherichia coli RecA (EcRecA) [112]. The ATPase domain consists of a large, twisted central β-sheet sandwiched by α-helices (Figures 15.4c and 15.5a,b). It contains both the classic Walker A and B motifs and includes an unusual cis-linked peptide bond. The asymmetric unit of PfRad51 crystals consisted of a heptameric ring, which then forms a higher-order dimer by crystallographic symmetry. The N-terminal domain was only well-ordered in one of the seven Rad51 heptamer subunits, reflecting reduced translational motion due to crystal contacts. The dimer of heptamers was verified in solution by dynamic light scattering (DLS) and SAXS [116]. The dimensions of the biheptamer are 118 Å in diameter by 105 Å in height. The ATPase domains form a ring of pie-shaped wedges with a central ~21 Å diameter hole lined by loop hairpins (Figure 15.5a). Intersubunit contacts are made between the ATPase domains of neighboring subunits. The most prominent subunit interface feature is the extension of the central ATPase β-sheet through strand β3 by the elbow-like interdomain linker of the adjacent subunit. Thus in the polymeric

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form an extra β-strand β0 is formed from the linker creating a β-zipper. A second prominent feature of the PfRad51 intersubunit interface is the insertion of a conserved Phe residue in the interdomain linker (Phe97 in PfRad51), located immediately prior to β0, into a hydrophobic pocket formed by residues of the central β-sheet and one of the large α-helices of an adjacent subunit. This arrangement resembles a ball and socket (Figures 15.5a,b, and d). Together the intersubunit β-sheet or

FIGURE 15.5 Rad51 interactions, conformations, and the interface mimicry and interface exchange hypothesis. (a) Pf Rad51 heptameric ring angled with a slight tilt. Subunits are shaded differently in the front to distinguish subunits. The region at the beginning of the arrow pointing to panel C shows the polymerization motif (PM). It consists of an extended β-sheet made by the PM β0 of the front dark subunit with β3 of the right adjacent light subunit. The conserved Phe residue from the dark subunit buries itself into a pocket formed by the light subunit. The PM is located in the interdomain linker that tethers the N-terminal domain to the C-terminal domain of a single subunit. (b) A modeled Rad51 filament generated by rigid body docking Pf Rad51 crystal structures into SsRadA electron microscope density. The area at the beginning of the arrowhead pointing to panel D shows how the PM is retained. (c) Rad51 structural mimicry by BRCA2 BRC repeats. The combined solid/ribbon representation corresponds to the right light subunit in panel A. The PM of the Rad51 front dark subunit in panel A is shown as sticks. A BRC4 peptide is shown in stick form and its position is based on an overlay of the HsRad51 ATPase:BRC4 fusion protein with Pf Rad51. (d) Zoom view of the PM interaction with the neighboring subunit from panel B. The arrows point to the common PM interaction between structures shown in panels (a–d). Continued

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FIGURE 15.5 (Continued) (e) Overlay of the PfRad51 and SsRadA crystal structures. Flexibility of the linker that contains the PM is easily visualized between the two structures. The PfRad51 structure shows the relative positions of the domains in the ring form, while the SsRadA structure shows the positions in an extended filament form. (f) Engineered PfRad51 mutants are targeted to the nucleus of human cells following DNA damage. Top panels show the position of the mutant through fusion to GFP in the cell nuclei. Bottom panels show that the effect is dependent on BRCA2, as competing peptides consisting of BRC repeats 3 and 4 inhibit translocation of mutant PfRad51 into the nucleus.

β-zipper, and the ball and socket make up the Rad51 polymerization motif. The flexibility of the linker, in which the polymerization motif resides, allows the rings to exist in different oligomeric forms, depending on the species. The S. solfataricus RadA [120] (SsRadA) and human DMC1 [121] (HsDMC1) proteins were found to form octamers rather than heptamers. The recombination active form of Rad51 is a helical nucleoprotein filament. To characterize a filament, the PfRad51 structure was computationally docked into electron microscopy three-dimensional reconstruction density of the archaeal S. solfataricus RadA protein [116,120] (Figure 15.5b). Because the Rad51/RadA proteins have an N-terminal domain that resides on the opposite side of the ATPase domain in contrast to the RecA C-terminal domain, the Rad51 filament has a corresponding opposite polarity of lobes in the large outer groove of the filament relative to the polarity seen for bacterial RecA proteins [113,114,120]. This difference in polarity may explain the 5′ to 3′ polarity of Rad51 for strand exchange, which is opposite from the polarity of RecA [122,123]. The Rad51 x-ray crystal structures solved later from the yeast S. cerevisiae (ScRad51) [118] and the archaeon Methanococcus voltae [124] revealed filament forms in the absence of DNA. Both x-ray crystal structures confirmed the polarity predicted from the electron microscopy-derived models. In all Rad51/RadA structures, the polymerization motif remains intact despite changes in pitch from 0 Å for the PfRad51 and DMC1 structures to ~107 and ~130 Å for the ScRad51 and MvRadA structures, respectively, which are in accordance with the pitches observed by electron microscopy. The Phe residue of the ball and socket within the polymerization motif makes the greatest interface surface contact. The SsRadA structure crystallized in a very unusual form that had a P3121 space group resulting in the filament having only three subunits per helical turn. Electron microscopy reconstructions usually result in filaments having just over six subunits per turn, while other crystal structure filaments, being constrained by P61 symmetry crystal contacts, are slightly more tightly wound having exactly six subunits per turn. The pitch of the SsRadA structure was 99 Å, however, because it has fewer subunits per turn, the interface is quite different. Yet, the polymerization motif still tethers the polymer together. The significant rotation required for the transition can be seen in the overlay of the PfRad51 and SsRadA structures in Figure 15.5e. Together, these results support the use of the polymerization motif as a structural device for transition from rings and filaments and to extend and contract the filaments. Further analyses of the various structures reveal that the ATPase domains reorient themselves during the ring-to-filament transition and

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during expansion and contraction of filaments. This likely represents a switch mechanism that may regulate ATPase activity using DNA-induced conformational changes and support a nucleotidemediated handoff hypothesis that implies that nucleotide binding in the form of ATP/ADP and DNA interactions mediate steps of HR. In the helical Rad51 structures, positive electrostatic potential maps to the filament interior and the HhH motif, so these regions have potential to bind the DNA phosphate backbone. Biochemical experiments show that Rad51 binds ssDNA before dsDNA during strand exchange [123]. Electron microscopy reconstructions of Rad51 nucleoprotein filaments implicate the inner region as the primary ssDNA binding site [113,114]. Nuclear magnetic resonance (NMR) studies show that HsRad51 HhH residues Ala61-Glu69 may bind dsDNA [125], creating a potential secondary site. By superimposing HhH motifs bound to DNA from other structures, such as DNA polymerase β [27], with the PfRad51 structure a general dsDNA-binding mode of Rad51 was deduced [116]. This arrangement places the dsDNA in the wide outer groove of the protein filament. Analysis of the various electron microscopy reconstructions and crystal structures suggest that the two DNAs are able to come into contact in extended filaments. Furthermore, homology searches may be facilitated in the Rad51 filament by nucleotide-dependent movement of the Rad51 N-terminal domains relative to the C-terminal domains [114,115]. While the details of how DNA strand exchange are still being characterized, the utility of the first Rad51 structure derived from a hyperthermophile was extremely useful in determining possible mechanisms for the involvement of Rad51 in certain forms of cancer [103,116,119] as BRCA2 mediates Rad51 interactions in human cells [126–130]. In higher eukaryotes, the breast cancer susceptibility protein BRCA2 interacts with Rad51 and plays a role in HR. Women who carry a BRCA2 mutation have a 60% to 85% lifetime risk for developing breast cancer and a 10% to 12% lifetime risk for developing ovarian cancer [131]. BRCA2 polymorphisms may be associated with increased risk of other tumor types [132] and BRCA2 mutations are linked to Fanconi anemia-associated acute myeloid leukemia and squamous cell carcinoma [133]. Cells harboring BRCA2 truncations have an increased frequency of gross chromosomal rearrangements and DSBs and are sensitive to UV light, ionizing radiation, methyl methanesulfate, and other genotoxic agents [129,134–138]. Loss of function mutations in BRCA2 (or Rad51) cause embryonic lethality [139]. Furthermore, BRCA2 binds Rad51 and forms discrete nuclear foci in cells with DNA damage [136,140,141]. The central region of BRCA2 contains a set of noncontiguous but highly conserved repeat sequences of roughly 30 amino acids. Many tumorigenic polymorphisms map to these conserved BRC repeats, and a single mutation within a repeat can increase cancer risk [101,119]. The eight BRC repeats bind directly to the Rad51 filament; however BRC repeat-derived peptides prevent Rad51 polymerization into rings and nucleoprotein filaments in vitro [140,141] and prevent nuclear aggregates of Rad51 in vivo [136]. To determine how BRCA2 operates as an antagonist for Rad51 polymerization and a chaperone for DNA targeting, the BRC repeat 4 (BRC4) peptide from a HsRad51 ATPase domain:BRC4 fusion (HsRad51-AD:BRC4) structure [119] was analyzed as to how it would be accommodated by a full-length polymeric Rad51 protein using PfRad51 [116]. When HsRad51-AD:BRC4 is superimposed on one subunit of PfRad51 in the ring, a remarkable similarity is observed between PfRad51 residues 93–102 and BRCA2 residues 1520–1529 (Figure 15.5c). In essence, the BRC4 repeat mimics the polymerization motif by forming the extra β-strand and inserting its conserved Phe residue in to the Rad51-binding pocket. These structures indicate that a BRC repeat-derived peptide disrupts the Rad51:Rad51 intersubunit interaction [116]. Thus, when an intermolecular Rad51:BRC interaction occurs, the β-zipper binding interface that facilitates Rad51 polymerization is sequestered in the Rad51:BRCA2 interface. These data support an Interface Exchange Hypothesis for interactions between Rad51 and BRCA2, in that the Rad51:Rad51 interactions in a polymer are substituted by Rad51:BRC repeat interactions. In particular, the data suggest that prior to loading onto DNA, Rad51 forms a β-zipper with a BRC4 repeat, which is subsequently

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exchanged for a β-zipper with another Rad51 subunit as it loads onto DNA. The interaction and validity of the position of the BRC4 repeat binding site within the HsRad51-AD:BRC4 structure was verified by using a positive mutagenesis scheme, where a structure-based mutant PfRad51 binds BRC repeats and forms nuclear foci in human 293T cells in response to γ-irradiation-induced DNA damage (Figure 15.5f). Furthermore, coexpression of BRC repeats 3 and 4 block nuclear foci formation by mutant PfRad51. It should be noted that archaea do not carry BRCA2 proteins, thus the uniqueness of the thermophilic archaeal Rad51/RadA proteins, in terms of their stability, and strong sequence and fold similarities to those found in the human system combined with their different set of HR system proteins, unexpectedly played a significant role in our understanding part of the structural basis for a disease of high medical relevance allowing for therapeutic design.

RUVB AND HJ BRANCH MIGRATION RuvB is the ATP-driven motor for DNA homologous recombination by which organisms not only maintain genetic stability but also generate biological diversity, through rearrangements between homologous chromosomes during meiosis. Recent data also suggest that HR is required to restart progression of stalled replication forks [142]. DNA HR involves the formation of the universal DNA intermediate termed Holiday junction (HJ). Therefore, a common HJ resolution mechanism might be shared by all the species. The HJ is a dynamic structure and can adopt diverse conformations between two extremes: termed “open X” and “stacked X,” depending on its binding to proteins. In prokaryotes, the HJ DNA is recognized by RuvA, which forms a fourfold symmetric tetramer [143,144]. Each RuvA contains three domains (domains I, II, and III). It has been demonstrated that the RuvA core (domains I and II) is exclusively responsible for HJ binding whereas the highly mobile domain III directly interacts with RuvB to promote the loading of the hexameric RuvB motor proteins onto the HJ DNA [145]. They together are responsible for the ATP-dependent branch migration. The HJ structure is then resolved by the structure-specific endonuclease RuvC through a divalent-metal-dependent cleavage, generating two separate recombinant DNA duplexes [146]. RuvB is a member of the diverse AAA+ (ATPase associated with various cellular activities) ATPase family [147], which includes NSF, HslU, SV40 large T-antigen (Tag), and others. Electron microscopic analysis of RuvB from Thermus thermophilus indicates that RuvB is a heptamer, but converts to a hexamer upon dsDNA binding [148]. Crystallographic studies of RuvB from T. thermophilus and Thermotoga maritima reveal that RuvB has a crescent-like structure consisting of three domains (classified as domains N, M, and C) [149,150]. Domain N has a typical Rossman fold, composed of five paralleled β-strands and the surrounding α-helices. Domain M is composed of four α-helices connected by loops. Domains N and M are conserved among AAA+ ATPases and involved in ATP hydrolysis. An ATP analog AMPPNP (5′-adenyl-imido-triphosphate) and ADP molecule was observed at the interface between domains N and M in crystal structure of ThRuvB and TmRuvB, respectively. The nucleotide binding pocket is provided by the conserved Walker A and B motifs, located separately in domains N and M. Domain C consists of five α-helices and one β-hairpin, showing a “winged-helix” DNA-binding motif that is observed in many transcription factors and in nonspecific DNA-bind proteins such as linker histone H5 [151]. Structure–function analysis [152] of the ThRuvB protein indicated that domain N is involved in RuvA–RuvB and RuvB–RuvB interactions, domains N and M are required for ATP hydrolysis and hexameric formation, and domain C plays an essential role in DNA binding. TmRuvB forms a helix with six RuvB molecules per turn through crystal packing [149]. Protein– protein contacts in the helix use the same molecular interfaces that are observed in other hexameric AAA+ ATPases. Superimposition (Figure 15.6) of RuvB domain N onto the conserved domain of the large Tag structure [153] results in a polar hexamer containing a large lobe (domains N and M) and a small lobe (domain C) as observed by electron miscroscopy [148]. Both ThRuvB [150] and TmRuvB [149] hexamers constructed this way have a central hole large enough for dsDNA duplex

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accommodation. The fact that ADP-bound RuvB assembles into a helix with six subunits per turn in crystals rather than a hexameric ring suggests that conformational changes are induced upon ADP binding, and that not all RuvB molecules in the hexameric ring can exist in an ADP-bound conformation at the same time. Crystal structure of the RuvA–RuvB complex from T. thermophilus [145] reveals that two RuvA tetramers form a symmetric and closed octameric shell, where four RuvA domain IIIs spring out toward the four corners of a square from the center of the RuvA core structure composed of domains I and II of RuvA. These RuvA domain IIIs tether four RuvB molecules by interacting individually with a unique hydrophobic β-hairpin protruded from RuvB domain N. The tethered RuvB molecules are arranged in such a way that domain C with the winged-helix DNA motif is farthest from the RuvA octameric core while domain N lies closest to the core. The concave surfaces of the two RuvA tetramers face head-to-head, generating an empty space large enough to accommodate the missing HJ DNA. This is in good agreement with the crystal structure of Mycobacterium coli octameric RuvA–HJ DNA complex, where two RuvA tetramers bind to both sides of the junction [144]. A model of the RuvA–RuvB/junction DNA ternary complex, constructed by fitting the crystal RuvA–RuvB structure into the average electron microscopic images of the RuvA–RuvB/junction DNA complex indicated that two hexameric RuvB rings located at the opposite sides of the RuvA octameric shell with two RuvB molecules from each ring interacting with domain IIIs of two RuvA molecules. The binding of RuvA domain III may deform the β-hairpin (disordered in TmRuvB crystals) in RuvB domain N, and therefore induces a functional but less symmetric RuvB hexameric ring for branch migration. The six RuvB subunits within each ring are grouped into two semicircular rings, each of which consists of three subunits possibly in distinct nucleotide states (Figure 15.6). The two pairs of RuvA domain IIIs linking the RuvA octameric core and two RuvB rings are positioned on the same plane as that of open HJ DNA and parallel to the DNA arms passing through the central holes of the two RuvB hexameric rings. All dsDNA arms of the HJ DNA run into and out from the RuvA octamer and plausibly rotate along a spiral taxiway on the inner surface of the RuvA octamer without steric clash between DNA and protein. During branch migration, the acid pins at the central hole of each RuvA tetramer play a crucial role in separating DNA strands that are incoming to the junction center [143]. The two hexameric RuvB rings are responsible for pumping out

FIGURE 15.6 The Holliday junction molecular motor RuvB structure, domain interactions, and hexameric assembly. Functional RuvB is a hexameric ring with twofold symmetry. RuvB hexamer is built by superimposing the N-terminal domain of TtRuvB (PDB code, 1HQC) onto the large Tag-ATPase domain within the large Tag structure (PDB code, 1N25). One half of the ring is shown with three RuvB molecules in free (dark), ADP-bound (light), and ATP-bound (dark) forms, respectively.

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DNA duplexes by exerting a spiral rotation on each encircled dsDNA arm using the energy of ATP hydrolysis. A tight connection between RuvA domain III and RuvB seems essential for branch migration because all substitutions of various single amino acid residues involved in contacts between RuvA domain III and RuvB cause complete loss of ATP-dependent branch migration activity [154]. This implies that the RuvB rings are essentially fixed to the RuvA octameric core during branch migration. RuvA domain III interacting with RuvB is connected to the RuvA octameric core by a flexible loop to allow appropriate flexibility for coordination between the individual actions of rearranging base pairs by RuvA and pulling DNA duplexes by RuvB, therefore driving a smooth and consecutive branch migration. At the end of branch migration, one RuvA tetramer somehow comes off the octameric RuvA–HJ complex to form a tetrameric RuvA–HJ complex as observed in the crystal structure of E. coli RuvA–HJ complex [155]. The open HJ surface is then occupied by RuvC dimer for junction resolution [149,156,157].

CONCLUSIONS AND PERSPECTIVES Proteins from hyperthermophiles have aided numerous discoveries involving the critical dynamic and reversible complexes that act in DNA replication, recombination, and repair. The combination of x-ray crystal structures, SAXS solution measurements, and electron microscopy three-dimensional reconstruction of themophilic proteins in particular provide detailed molecular understanding from proteins to pathways for DNA replication, repair, and recombination. The structures of these thermophilic proteins, which are often the first ones in their individual classes and sometimes the only structures available, contribute tremendously to our understanding of the molecular mechanisms underlying these important biological processes from microbes to humans. Also in several cases structural and biophysical characterizations of these thermophilic proteins have identified functionally important conformational states and changes that eluded other approaches. Fortunately, the conservation observed between thermophilic proteins and their mesophilic prokaryotic and eukaryotic counterparts is characteristic of the key proteins for many important biological and cellular processes. As more than a dozen archaeal genomes have been sequenced in the past 10 years and more are expected to be sequenced soon, hyperthermophilic genomic information will provide a valuable resource for elucidating new structures and molecular mechanisms important to life sciences and industrial technology. Currently most target selections in structural studies of thermophilic proteins solely depend on sequence conservation with their counterparts in prokaryotes and eukaryotes. We predict that this will soon change. With new technologies being developed for culturing thermophiles in laboratories, many macromolecular assemblies important to fundamental biological processes such as those described here will be isolated directly from thermophilic organisms. Structural studies on these macromolecular assembles by both x-ray crystallography and other biophysical techniques such as SAXS will have profound impacts in life sciences. It is obvious that a complete realization of the tremendous value of hyperthermophiles for defining master keys to structural cell biology from microbes to humans is still ahead of us.

ACKNOWLEDGMENTS The synchrotron crystallographic and SAXS studies are dependent upon the SIBLYS beamline at the Advanced Light Source and the development of methods for the characterization of reversible protein complexes and conformations as supported by the National Institutes of Health (NIH) grants (CA92584, CA081967, and CA63503), and by the Office of Science, Office of Biological and Environmental Research, U.S. Department of Energy, under Contract Number DE-AC02-05CH11231.

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REFERENCES 1. Woese, C.R., Kandler, O., and Wheelis, M.L. Towards a natural system of organisms: proposal for the domains Archaea, Bacteria, and Eucarya. Proc Natl Acad Sci USA 87, 4576–9 (1990). 2. Tsutakawa, S.E., Hura, G.L., Frankel, K.A., Cooper, P.K., and Tainer, J.A. Structural analysis of flexible proteins in solution by small angle X-ray scattering combined with crystallography. J Struct Biol 158, 214–23 (2007). 3. Perry, J.J., Fan, L., and Tainer, J.A. Developing master keys to brain pathology, cancer and aging from the structural biology of proteins controlling reactive oxygen species and DNA repair. Neuroscience 145, 1280–99 (2007). 4. DePamphilis, M.L. (ed.) DNA Replication and Human Disease (Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York, 2006). 5. Larsen, N.B., Rasmussen, M., and Rasmussen, L.J. Nuclear and mitochondrial DNA repair: similar pathways? Mitochondrion 5, 89–108 (2005). 6. Bambara, R.A., Murante, R.S., and Henricksen, L.A. Enzymes and reactions at the eukaryotic DNA replication fork. J Biol Chem 272, 4647–50 (1997). 7. Maga, G. et al. Okazaki fragment processing: modulation of the strand displacement activity of DNA polymerase delta by the concerted action of replication protein A, proliferating cell nuclear antigen, and flap endonuclease-1. Proc Natl Acad Sci USA 98, 14298–303 (2001). 8. Matsumoto, Y. Molecular mechanism of PCNA-dependent base excision repair. Prog Nucleic Acid Res Mol Biol 68, 129–38 (2001). 8a. Nishada, et al. The closed structure of an archael DNA ligase from Pyrococcus furiosus. J Mol Biol 360, 956–67 (2006). 9. Kucherlapati, M. et al. Haploinsufficiency of Flap endonuclease (Fen1) leads to rapid tumor progression. Proc Natl Acad Sci USA 99, 9924–9 (2002). 10. Henneke, G., Friedrich-Heineken, E., and Hubscher, U. Flap endonuclease 1: a novel tumour suppresser protein. Trends Biochem Sci 28, 384–90 (2003). 11. Kaiser, M.W. et al. A comparison of eubacterial and archaeal structure-specific 5′-exonucleases. J Biol Chem 274, 21387–94 (1999). 12. Kao, H.I., Henricksen, L.A., Liu, Y., and Bambara, R.A. Cleavage specificity of Saccharomyces cerevisiae flap endonuclease 1 suggests a double-flap structure as the cellular substrate. J Biol Chem 277, 14379–89 (2002). 13. Reynaldo, L.P., Vologodskii, A.V., Neri, B.P., and Lyamichev, V.I. The kinetics of oligonucleotide replacements. J Mol Biol 297, 511–20 (2000). 14. Xie, Y. et al. Identification of rad27 mutations that confer differential defects in mutation avoidance, repeat tract instability, and flap cleavage. Mol Cell Biol 21, 4889–99 (2001). 15. Lieber, M.R. The FEN-1 family of structure-specific nucleases in eukaryotic DNA replication, recombination and repair. Bioessays 19, 233–40 (1997). 16. Shen, B., Qiu, J., Hosfield, D., and Tainer, J.A. Flap endonuclease homologs in archaebacteria exist as independent proteins. Trends Biochem Sci 23, 171–3 (1998). 17. Hosfield, D.J., Mol, C.D., Shen, B., and Tainer, J.A. Structure of the DNA repair and replication endonuclease and exonuclease FEN-1: coupling DNA and PCNA binding to FEN-1 activity. Cell 95, 135–46 (1998). 18. Hwang, K.Y., Baek, K., Kim, H.Y., and Cho, Y. The crystal structure of flap endonuclease-1 from Methanococcus jannaschii. Nat Struct Biol 5, 707–13 (1998). 19. Chapados, B.R. et al. Structural basis for FEN-1 substrate specificity and PCNA-mediated activation in DNA replication and repair. Cell 116, 39–50 (2004). 20. Kim, C.Y., Park, M.S., and Dyer, R.B. Human flap endonuclease-1: conformational change upon binding to the flap DNA substrate and location of the Mg2+ binding site. Biochemistry 40, 3208–14 (2001). 21. Storici, F. et al. The flexible loop of human FEN1 endonuclease is required for flap cleavage during DNA replication and repair. Embo J 21, 5930–42 (2002). 22. Krishna, T.S., Kong, X.P., Gary, S., Burgers, P.M., and Kuriyan, J. Crystal structure of the eukaryotic DNA polymerase processivity factor PCNA. Cell 79, 1233–43 (1994). 23. Gulbis, J.M., Kelman, Z., Hurwitz, J., O’Donnell, M., and Kuriyan, J. Structure of the C-terminal region of p21 (WAF1/CIP1) complexed with human PCNA. Cell 87, 297–306 (1996). 24. Matsumiya, S., Ishino, S., Ishino, Y., and Morikawa, K. Physical interaction between proliferating cell nuclear antigen and replication factor C from Pyrococcus furiosus. Genes Cells 7, 911–22 (2002).

9214_C015.indd 258

11/5/2007 4:32:06 PM

Master Keys to DNA Replication, Repair, and Recombination

259

25. Shamoo, Y. and Steitz, T.A. Building a replisome from interacting pieces: sliding clamp complexed to a peptide from DNA polymerase and a polymerase editing complex. Cell 99, 155–66 (1999). 26. Sakurai, S. et al. Structural basis for recruitment of human flap endonuclease 1 to PCNA. Embo J 24, 683–93 (2005). 27. Sawaya, M.R., Prasad, R., Wilson, S.H., Kraut, J., and Pelletier, H. Crystal structures of human DNA polymerase beta complexed with gapped and nicked DNA: evidence for an induced fit mechanism. Biochemistry 36, 11205–15 (1997). 28. Kedar, P.S. et al. Direct interaction between mammalian DNA polymerase beta and proliferating cell nuclear antigen. J Biol Chem 277, 31115–23 (2002). 29. Dionne, I., Nookala, R.K., Jackson, S.P., Doherty, A.J., and Bell, S.D. A heterotrimeric PCNA in the hyperthermophilic archaeon Sulfolobus solfataricus. Mol Cell 11, 275–82 (2003). 30. Pascal, J.M. et al. A flexible interface between DNA ligase and PCNA supports conformational switching and efficient ligation of DNA. Mol Cell 24, 279–91 (2006). 31. Dore, A.S. et al. Structure of an archaeal PCNA1-PCNA2-FEN1 complex: elucidating PCNA subunit and client enzyme specificity. Nucleic Acids Res 34, 4515–26 (2006). 32. Williams, G.J. et al. Structure of the heterotrimeric PCNA from Sulfolobus solfataricus. Acta Crystallograph Sect F Struct Biol Cryst Commun 62, 944–8 (2006). 33. Lehman, I.R. DNA ligase: structure, mechanism, and function. Science 186, 790–7 (1974). 34. Shuman, S. and Lima, C.D. The polynucleotide ligase and RNA capping enzyme superfamily of covalent nucleotidyltransferases. Curr Opin Struct Biol 14, 757–64 (2004). 35. Tomkinson, A.E., Vijayakumar, S., Pascal, J.M., and Ellenberger, T. DNA ligases: structure, reaction mechanism, and function. Chem Rev 106, 687–99 (2006). 36. Doherty, A.J. and Wigley, D.B. Functional domains of an ATP-dependent DNA ligase. J Mol Biol 285, 63–71 (1999). 37. Subramanya, H.S., Doherty, A.J., Ashford, S.R., and Wigley, D.B. Crystal structure of an ATP-dependent DNA ligase from bacteriophage T7. Cell 85, 607–15 (1996). 38. Pascal, J.M., O’Brien, P.J., Tomkinson, A.E., and Ellenberger, T. Human DNA ligase I completely encircles and partially unwinds nicked DNA. Nature 432, 473–8 (2004). 39. Schaeffer, L. et al. DNA repair helicase: a component of BTF2 (TFIIH) basic transcription factor. Science 260, 58–63 (1993). 40. Tirode, F., Busso, D., Coin, F., and Egly, J.M. Reconstitution of the transcription factor TFIIH: assignment of functions for the three enzymatic subunits, XPB, XPD, and cdk7. Mol Cell 3, 87–95 (1999). 41. Moreland, R.J. et al. A role for the TFIIH XPB DNA helicase in promoter escape by RNA polymerase II. J Biol Chem 274, 22127–30 (1999). 42. Zurita, M. and Merino, C. The transcriptional complexity of the TFIIH complex. Trends Genet 19, 578–84 (2003). 43. Sung, P., Guzder, S.N., Prakash, L., and Prakash, S. Reconstitution of TFIIH and requirement of its DNA helicase subunits, Rad3 and Rad25, in the incision step of nucleotide excision repair. J Biol Chem 271, 10821–6 (1996). 44. Reardon, J.T. and Sancar, A. Nucleotide excision repair. Prog Nucleic Acid Res Mol Biol 79, 183–235 (2005). 45. Hoeijmakers, J.H. Genome maintenance mechanisms for preventing cancer. Nature 411, 366–74 (2001). 46. Weeda, G. et al. A mutation in the XPB/ERCC3 DNA repair transcription gene, associated with trichothiodystrophy. Am J Hum Genet 60, 320–9 (1997). 47. Lehmann, A.R. DNA repair-deficient diseases, xeroderma pigmentosum, Cockayne syndrome and trichothiodystrophy. Biochimie 85, 1101–11 (2003). 48. Wood, R.D. Nucleotide excision repair in mammalian cells. J Biol Chem 272, 23465–8 (1997). 49. Hanawalt, P.C. Subpathways of nucleotide excision repair and their regulation. Oncogene 21, 8949–56 (2002). 50. Sarker, A.H. et al. Recognition of RNA polymerase II and transcription bubbles by XPG, CSB, and TFIIH: insights for transcription-coupled repair and Cockayne Syndrome. Mol Cell 20, 187–98 (2005). 51. Laine, J.P. and Egly, J.M. Initiation of DNA repair mediated by a stalled RNA polymerase IIO. Embo J 25, 387–97 (2006). 52. Riedl, T., Hanaoka, F., and Egly, J.M. The comings and goings of nucleotide excision repair factors on damaged DNA. Embo J 22, 5293–303 (2003).

9214_C015.indd 259

11/5/2007 4:32:06 PM

260

Thermophiles: Biology and Technology at High Temperatures

53. O’Donovan, A., Davies, A.A., Moggs, J.G., West, S.C., and Wood, R.D. XPG endonuclease makes the 3′ incision in human DNA nucleotide excision repair. Nature 371, 432–5 (1994). 54. Sijbers, A.M. et al. Xeroderma pigmentosum group F caused by a defect in a structure-specific DNA repair endonuclease. Cell 86, 811–22 (1996). 55. Shivji, M.K., Podust, V.N., Hubscher, U., and Wood, R.D. Nucleotide excision repair DNA synthesis by DNA polymerase epsilon in the presence of PCNA, RFC, and RPA. Biochemistry 34, 5011–17 (1995). 56. Hoogstraten, D. et al. Rapid switching of TFIIH between RNA polymerase I and II transcription and DNA repair in vivo. Mol Cell 10, 1163–74 (2002). 57. Fan, L. et al. Conserved XPB core structure and motifs for DNA unwinding: implications for pathway selection of transcription or excision repair. Mol Cell 22, 27–37 (2006). 58. Lin, Y.C., Choi, W.S., and Gralla, J.D. TFIIH XPB mutants suggest a unified bacterial-like mechanism for promoter opening but not escape. Nat Struct Mol Biol 12, 603–7 (2005). 59. Coin, F. et al. Phosphorylation of XPB helicase regulates TFIIH nucleotide excision repair activity. Embo J 23, 4835–46 (2004). 60. Lin, Y.C. and Gralla, J.D. Stimulation of the XPB ATP-dependent helicase by the beta subunit of TFIIE. Nucleic Acids Res 33, 3072–81 (2005). 61. Coin, F. et al. p8/TTD-A as a repair-specific TFIIH subunit. Mol Cell 21, 215–26 (2006). 62. Morgante, P.G. et al. Functional XPB/RAD25 redundancy in Arabidopsis genome: characterization of AtXPB2 and expression analysis. Gene 344, 93–103 (2005). 63. Gorbalenya, A.E. and Koonin, E.V. Helicases: amino acid sequence comparisons and structure-function relationships. Curr Opin Struct Biol 3, 419–29 (1993). 64. Obmolova, G., Ban, C., Hsieh, P., and Yang, W. Crystal structures of mismatch repair protein MutS and its complex with a substrate DNA. Nature 407, 703–10 (2000). 65. Caruthers, J.M. and McKay, D.B. Helicase structure and mechanism. Curr Opin Struct Biol 12, 123–33 (2002). 66. Korolev, S., Hsieh, J., Gauss, G.H., Lohman, T.M., and Waksman, G. Major domain swiveling revealed by the crystal structures of complexes of E. coli Rep helicase bound to single-stranded DNA and ADP. Cell 90, 635–47 (1997). 67. Soultanas, P. and Wigley, D.B. Unwinding the ‘Gordian knot’ of helicase action. Trends Biochem Sci 26, 47–54 (2001). 68. Kim, T.K., Ebright, R.H., and Reinberg, D. Mechanism of ATP-dependent promoter melting by transcription factor IIH. Science 288, 1418–22 (2000). 69. Stracker, T.H., Theunissen, J.W., Morales, M., and Petrini, J.H. The Mre11 complex and the metabolism of chromosome breaks: the importance of communicating and holding things together. DNA Repair (Amst) 3, 845–54 (2004). 70. Zhu, J., Petersen, S., Tessarollo, L., and Nussenzweig, A. Targeted disruption of the Nijmegen breakage syndrome gene NBS1 leads to early embryonic lethality in mice. Curr Biol 11, 105–9 (2001). 71. Luo, G. et al. Disruption of mRad50 causes embryonic stem cell lethality, abnormal embryonic development, and sensitivity to ionizing radiation. Proc Natl Acad Sci U S A 96, 7376–81 (1999). 72. Xiao, Y. and Weaver, D.T. Conditional gene targeted deletion by Cre recombinase demonstrates the requirement for the double-strand break repair Mre11 protein in murine embryonic stem cells. Nucleic Acids Res 25, 2985–91 (1997). 73. D’Amours, D. and Jackson, S.P. The Mre11 complex: at the crossroads of dna repair and checkpoint signalling. Nat Rev Mol Cell Biol 3, 317–27 (2002). 74. Assenmacher, N. and Hopfner, K.P. MRE11/RAD50/NBS1: complex activities. Chromosoma 113, 157–66 (2004). 75. de Jager, M. et al. DNA-binding and strand-annealing activities of human Mre11: implications for its roles in DNA double-strand break repair pathways. Nucleic Acids Res 29, 1317–25 (2001). 76. Chen, L., Trujillo, K., Ramos, W., Sung, P., and Tomkinson, A.E. Promotion of Dnl4-catalyzed DNA end-joining by the Rad50/Mre11/Xrs2 and Hdf1/Hdf2 complexes. Mol Cell 8, 1105–15 (2001). 77. Hopfner, K.P. et al. The Rad50 zinc-hook is a structure joining Mre11 complexes in DNA recombination and repair. Nature 418, 562–6 (2002). 78. Moreno-Herrero, F. et al. Mesoscale conformational changes in the DNA-repair complex Rad50/Mre11/ Nbs1 upon binding DNA. Nature 437, 440–3 (2005). 79. Hopfner, K.P. et al. Structural biochemistry and interaction architecture of the DNA double-strand break repair Mre11 nuclease and Rad50-ATPase. Cell 105, 473–85 (2001).

9214_C015.indd 260

11/5/2007 4:32:06 PM

Master Keys to DNA Replication, Repair, and Recombination

261

80. Hopfner, K.P. et al. Mre11 and Rad50 from Pyrococcus furiosus: cloning and biochemical characterization reveal an evolutionarily conserved multiprotein machine. J Bacteriol 182, 6036–41 (2000). 81. Hopfner, K.P. et al. Structural biology of Rad50 ATPase: ATP-driven conformational control in DNA double-strand break repair and the ABC-ATPase superfamily. Cell 101, 789–800 (2000). 82. Hopfner, K. et al. Structural biochemistry and interaction architecture of the dna double-strand break repair mre11 nuclease and rad50-atpase. Cell 105, 473–85 (2001). 83. Connelly, J.C., Kirkham, L.A., and Leach, D.R. The SbcCD nuclease of Escherichia coli is a structural maintenance of chromosomes (SMC) family protein that cleaves hairpin DNA. Proc Natl Acad Sci USA 95, 7969–74 (1998). 84. Furuse, M. et al. Distinct roles of two separable in vitro activities of yeast Mre11 in mitotic and meiotic recombination. Embo J 17, 6412–25 (1998). 85. Paull, T.T. and Gellert, M. The 3′ to 5′ exonuclease activity of Mrel1 facilitates repair of DNA doublestrand breaks. Mol Cell 1, 969–79 (1998). 86. Trujillo, K.M., Yuan, S.S., Lee, E.Y., and Sung, P. Nuclease activities in a complex of human recombination and DNA repair factors Rad50, Mre11, and p95. J Biol Chem 273, 21447–50 (1998). 87. Usui, T. et al. Complex formation and functional versatility of Mre11 of budding yeast in recombination. Cell 95, 705–16 (1998). 88. Lobachev, K.S., Gordenin, D.A., and Resnick, M.A. The Mre11 complex is required for repair of hairpin-capped double-strand breaks and prevention of chromosome rearrangements. Cell 108, 183–93 (2002). 89. Tsubouchi, H. and Ogawa, H. Exo1 roles for repair of DNA double-strand breaks and meiotic crossing over in Saccharomyces cerevisiae. Mol Biol Cell 11, 2221–33 (2000). 90. Carney, J.P. et al. The hMre11/hRad50 protein complex and Nijmegen breakage syndrome: linkage of double-strand break repair to the cellular DNA damage response. Cell 93, 477–86 (1998). 91. Paull, T.T. and Gellert, M. Nbs1 potentiates ATP-driven DNA unwinding and endonuclease cleavage by the Mre11/Rad50 complex. Genes Dev 13, 1276–88 (1999). 92. Moncalian, G. et al. The rad50 signature motif: essential to ATP binding and biological function. J Mol Biol 335, 937–51 (2004). 93. Melby, T.E., Ciampaglio, C.N., Briscoe, G., and Erickson, H.P. The symmetrical structure of structural maintenance of chromosomes (SMC) and MukB proteins: long, antiparallel coiled coils, folded at a flexible hinge. J Cell Biol 142, 1595–604 (1998). 94. Lobachev, K., Vitriol, E., Stemple, J., Resnick, M.A., and Bloom, K. Chromosome fragmentation after induction of a double-strand break is an active process prevented by the RMX repair complex. Curr Biol 14, 2107–12 (2004). 95. Wiltzius, J.J., Hohl, M., Fleming, J.C., and Petrini, J.H. The Rad50 hook domain is a critical determinant of Mre11 complex functions. Nat Struct Mol Biol 12, 403–7 (2005). 96. Hopfner, K.P., Putnam, C.D., and Tainer, J.A. DNA double-strand break repair from head to tail. Curr Opin Struct Biol 12, 115–22 (2002). 97. Lee, J.H. and Paull, T.T. Direct activation of the ATM protein kinase by the Mre11/Rad50/Nbs1 complex. Science 304, 93–6 (2004). 98. Lee, J.H. and Paull, T.T. ATM activation by DNA double-strand breaks through the Mre11-Rad50-Nbs1 complex. Science 308, 551–4 (2005). 99. Khanna, K.K. and Jackson, S.P. DNA double-strand breaks: signaling, repair and the cancer connection. Nat Genet 27, 247–54 (2001). 100. van Gent, D.C., Hoeijmakers, J.H. and Kanaar, R. Chromosomal stability and the DNA double-stranded break connection. Nat Rev Genet 2, 196–206 (2001). 101. Venkitaraman, A.R. Cancer susceptibility and the functions of BRCA1 and BRCA2. Cell 108, 171–82 (2002). 102. Symington, L.S. Role of RAD52 epistasis group genes in homologous recombination and double-strand break repair. Microbiol Mol Biol Rev 66, 630–70, table of contents (2002). 103. Shin, D.S., Chahwan, C., Huffman, J.L., and Tainer, J.A. Structure and function of the double-strand break repair machinery. DNA Repair (Amst) 3, 863–73 (2004). 104. Sung, P. Catalysis of ATP-dependent homologous DNA pairing and strand exchange by yeast RAD51 protein. Science 265, 1241–3 (1994). 105. Benson, F.E., Stasiak, A., and West, S.C. Purification and characterization of the human Rad51 protein, an analogue of E. coli RecA. Embo J 13, 5764–71 (1994).

9214_C015.indd 261

11/5/2007 4:32:07 PM

262

Thermophiles: Biology and Technology at High Temperatures

106. Sugiyama, T., Zaitseva, E.M., and Kowalczykowski, S.C. A single-stranded DNA-binding protein is needed for efficient presynaptic complex formation by the Saccharomyces cerevisiae Rad51 protein. J Biol Chem 272, 7940–5 (1997). 107. Mazin, A.V., Alexeev, A.A., and Kowalczykowski, S.C. A novel function of Rad54 protein: Stabilization of the Rad51 nucleoprotein filament. J Biol Chem 278(16), 14029–36 (2003). 108. Solinger, J.A., Kiianitsa, K., and Heyer, W.D. Rad54, a Swi2/Snf2-like recombinational repair protein, disassembles Rad51:dsDNA filaments. Mol Cell 10, 1175–88 (2002). 109. Hayashi, I., Morikawa, K., and Ishino, Y. Specific interaction between DNA polymerase II (PolD) and RadB, a Rad51/Dmc1 homolog, in Pyrococcus furiosus. Nucleic Acids Res 27, 4695–702 (1999). 110. Sandler, S.J., Satin, L.H., Samra, H.S., and Clark, A.J. recA-like genes from three archaean species with putative protein products similar to Rad51 and Dmc1 proteins of the yeast Saccharomyces cerevisiae. Nucleic Acids Res 24, 2125–32 (1996). 111. Datta, S. et al. Crystal structures of Mycobacterium tuberculosis RecA and its complex with ADPAlF(4): implications for decreased ATPase activity and molecular aggregation. Nucleic Acids Res 28, 4964–73 (2000). 112. Story, R.M., Weber, I.T., and Steitz, T.A. The structure of the E. coli recA protein monomer and polymer. Nature 355, 318–25 (1992). 113. Yang, S., VanLoock, M.S., Yu, X., and Egelman, E.H. Comparison of bacteriophage T4 UvsX and human Rad51 filaments suggests that RecA-like polymers may have evolved independently. J Mol Biol 312, 999–1009 (2001). 114. Yu, X., Jacobs, S.A., West, S.C., Ogawa, T., and Egelman, E.H. Domain structure and dynamics in the helical filaments formed by RecA and Rad51 on DNA. Proc Natl Acad Sci U S A 98, 8419–24 (2001). 115. VanLoock, M.S. et al. ATP-Mediated Conformational Changes in the RecA Filament. Structure (Camb) 11, 187–96 (2003). 116. Shin, D.S. et al. Full-length Archaeal Rad51 structure and mutants: mechanisms for RAD51 assembly and control by BRCA2. Embo J 22, 4566–76 (2003). 117. Thayer, M.M., Ahern, H., Xing, D., Cunningham, R.P., and Tainer, J.A. Novel DNA binding motifs in the DNA repair enzyme endonuclease III crystal structure. Embo J 14, 4108–20 (1995). 118. Conway, A.B. et al. Crystal structure of a Rad51 filament. Nat Struct Mol Biol 11, 791–6 (2004). 119. Pellegrini, L. et al. Insights into DNA recombination from the structure of a RAD51-BRCA2 complex. Nature 420, 287–93 (2002). 120. Yang, S., Yu, X., Seitz, E.M., Kowalczykowski, S.C., and Egelman, E.H. archaeal RadA protein binds DNA as both helical filaments and octameric rings. J Mol Biol 314, 1077–85 (2001). 121. Kinebuchi, T. et al. Structural basis for octameric ring formation and DNA interaction of the human homologous-pairing protein Dmc1. Mol Cell 14, 363–74 (2004). 122. Namsaraev, E.A. and Berg, P. Branch migration during Rad51-promoted strand exchange proceeds in either direction. Proc Natl Acad Sci U S A 95, 10477–81 (1998). 123. Sung, P. and Robberson, D.L. DNA strand exchange mediated by a RAD51-ssDNA nucleoprotein filament with polarity opposite to that of RecA. Cell 82, 453–61 (1995). 124. Wu, Y., He, Y., Moya, I.A., Qian, X., and Luo, Y. Crystal structure of archaeal recombinase RADA: a snapshot of its extended conformation. Mol Cell 15, 423–35 (2004). 125. Aihara, H., Ito, Y., Kurumizaka, H., Yokoyama, S. and Shibata, T. The N-terminal domain of the human Rad51 protein binds DNA: structure and a DNA binding surface as revealed by NMR. J Mol Biol 290, 495–504 (1999). 126. Sung, P. Function of yeast Rad52 protein as a mediator between replication protein A and the Rad51 recombinase. J Biol Chem 272, 28194–7 (1997). 127. Sugiyama, T. and Kowalczykowski, S.C. Rad52 protein associates with replication protein A (RPA)single-stranded DNA to accelerate Rad51-mediated displacement of RPA and presynaptic complex formation. J Biol Chem 277, 31663–72 (2002). 128. Yang, H. et al. BRCA2 function in DNA binding and recombination from a BRCA2-DSS1-ssDNA structure. Science 297, 1837–48 (2002). 129. Jasin, M. Homologous repair of DNA damage and tumorigenesis: the BRCA connection. Oncogene 21, 8981–93 (2002). 130. Kowalczykowski, S.C. Molecular mimicry connects BRCA2 to Rad51 and recombinational DNA repair. Nat Struct Biol 9, 897–9 (2002).

9214_C015.indd 262

11/5/2007 4:32:07 PM

Master Keys to DNA Replication, Repair, and Recombination

263

131. Nathanson, K.L., Wooster, R., Weber, B.L. and Nathanson, K.N. Breast cancer genetics: what we know and what we need. Nat Med 7, 552–6 (2001). 132. Easton, D.F.A.C., T.B.C.L. Cancer risks in BRCA2 mutation carriers.The Breast Cancer Linkage Consortium. J Natl Cancer Inst 91, 1310–6 (1999). 133. Howlett, N.G. et al. Biallelic inactivation of BRCA2 in Fanconi anemia. Science 297, 606–9 (2002). 134. Yu, V.P. et al. Gross chromosomal rearrangements and genetic exchange between nonhomologous chromosomes following BRCA2 inactivation. Genes Dev 14, 1400–6 (2000). 135. Patel, K.J. et al. Involvement of Brca2 in DNA repair. Mol Cell 1, 347–57 (1998). 136. Chen, P.L. et al. The BRC repeats in BRCA2 are critical for RAD51 binding and resistance to methyl methanesulfonate treatment. Proc Natl Acad Sci U S A 95, 5287–92 (1998). 137. Morimatsu, M., Donoho, G., and Hasty, P. Cells deleted for Brca2 COOH terminus exhibit hypersensitivity to gamma-radiation and premature senescence. Cancer Res 58, 3441–7 (1998). 138. Moynahan, M.E., Pierce, A.J., and Jasin, M. BRCA2 is required for homology-directed repair of chromosomal breaks. Mol Cell 7, 263–72 (2001). 139. Sharan, S.K. et al. Embryonic lethality and radiation hypersensitivity mediated by Rad51 in mice lacking Brca2. Nature 386, 804–10 (1997). 140. Wong, A.K., Pero, R., Ormonde, P.A., Tavtigian, S.V., and Bartel, P.L. RAD51 interacts with the evolutionarily conserved BRC motifs in the human breast cancer susceptibility gene brca2. J Biol Chem 272, 31941–4 (1997). 141. Davies, A.A. et al. Role of BRCA2 in control of the RAD51 recombination and DNA repair protein. Mol Cell 7, 273–82 (2001). 142. McGlynn, P. and Lloyd, R.G. Recombinational repair and restart of damaged replication forks. Nat Rev Mol Cell Biol 3, 859–70 (2002). 143. Ariyoshi, M., Nishino, T., Iwasaki, H., Shinagawa, H., and Morikawa, K. Crystal structure of the holliday junction DNA in complex with a single RuvA tetramer. Proc Natl Acad Sci USA 97, 8257–62 (2000). 144. Roe, S.M. et al. Crystal structure of an octameric RuvA-Holliday junction complex. Mol Cell 2, 361–72 (1998). 145. Yamada, K. et al. Crystal structure of the RuvA-RuvB complex: a structural basis for the Holliday junction migrating motor machinery. Mol Cell 10, 671–81 (2002). 146. Dickman, M.J. et al. The RuvABC resolvasome. Eur J Biochem 269, 5492–501 (2002). 147. Ogura, T. and Wilkinson, A.J. AAA+ superfamily ATPases: common structure--diverse function. Genes Cells 6, 575–97 (2001). 148. Miyata, T. et al. Two different oligomeric states of the RuvB branch migration motor protein as revealed by electron microscopy. J Struct Biol 131, 83–9 (2000). 149. Putnam, C.D. et al. Structure and mechanism of the RuvB Holliday junction branch migration motor. J Mol Biol 311, 297–310 (2001). 150. Yamada, K. et al. Crystal structure of the Holliday junction migration motor protein RuvB from Thermus thermophilus HB8. Proc Natl Acad Sci U S A 98, 1442–7 (2001). 151. Gajiwala, K.S. and Burley, S.K. Winged helix proteins. Curr Opin Struct Biol 10, 110–16 (2000). 152. Ohnishi, T., Hishida, T., Harada, Y., Iwasaki, H., and Shinagawa, H. Structure-function analysis of the three domains of RuvB DNA motor protein. J Biol Chem 280, 30504–10 (2005). 153. Li, D. et al. Structure of the replicative helicase of the oncoprotein SV40 large tumour antigen. Nature 423, 512–18 (2003). 154. Han, Y.W. et al. A unique beta-hairpin protruding from AAA+ ATPase domain of RuvB motor protein is involved in the interaction with RuvA DNA recognition protein for branch migration of Holliday junctions. J Biol Chem 276, 35024–8 (2001). 155. Hargreaves, D. et al. Crystal structure of E.coli RuvA with bound DNA Holliday junction at 6 A resolution. Nat Struct Biol 5, 441–6 (1998). 156. Davies, A.A. and West, S.C. Formation of RuvABC-Holliday junction complexes in vitro. Curr Biol 8, 725–7 (1998). 157. van Gool, A.J., Hajibagheri, N.M., Stasiak, A., and West, S.C. Assembly of the Escherichia coli RuvABC resolvasome directs the orientation of holliday junction resolution. Genes Dev 13, 1861–70 (1999).

9214_C015.indd 263

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DNA Replication in Thermophiles Jae-Ho Shin, Lori M. Kelman, and Zvi Kelman

CONTENTS Introduction ................................................................................................................................ Cell Cycle ................................................................................................................................... Origin of Replication ................................................................................................................. Initiator Proteins ........................................................................................................................ Structure of the Cdc6 Proteins ....................................................................................... Biochemical Properties of the Cdc6 Proteins ................................................................. Helicase ...................................................................................................................................... MCM Structure ............................................................................................................... Biochemical Properties of MCM .................................................................................... Mechanism of Helicase Assembly at the Origin ....................................................................... DNA Replication and Chromatin .............................................................................................. Concluding Remarks .................................................................................................................. Acknowledgments ...................................................................................................................... References ..................................................................................................................................

265 266 266 268 269 269 270 270 271 272 274 274 274 275

Since the sequencing of the first archaeal genome a decade ago, much attention has been focused on the study of the mechanism of DNA replication of these unique microorganisms. These studies revealed that although many of the archaeal DNA replication proteins are more similar to those of eukarya than bacteria, they are not simply “mini eukarya” but are rather a mosaic of the eukaryal and bacterial systems, with archaeal-specific features. Here our current understanding of the process of initiation of DNA replication and its interplay with chromatin and the cell cycle is summarized.

INTRODUCTION Chromosomal DNA replication is a complex process involving dozens of proteins and enzymes to ensure the accurate and timely duplication of the genetic information. The process is functionally, and often structurally, conserved in all life forms and is divided into three main stages: initiation, elongation, and termination. Replication starts at specific chromosomal regions called origins of replication. During the initiation process, origin-binding proteins (OBPs) bind to the origin and locally unwind the DNA duplex. The OBP recruits additional initiation factors to the origin to facilitate the initiation process. Next a helicase is recruited to the DNA to form the initial replication bubble. The single-stranded DNA (ssDNA) exposed behind the helicase is coated with ssDNA-binding protein (SSB). The polymerase and the rest of the replication machinery are associated with the SSB/origin complex to form the two replication forks and to initiate bidirectional DNA synthesis in the 265

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elongation phase. At termination, replication forks collide and are resolved, and the resulting daughter DNA molecules are completed and separated. The initiation process in thermophilic microorganisms is not well understood. Very limited information is available on the biochemical properties of thermophilic bacterial initiation proteins, but a few structures have been solved (Erzberger et al., 2002). Where pertinent, these studies will be discussed subsequently. The situation in archaea is different, however, and extensive effort has been put toward understanding the archaeal cell cycle and initiation of DNA replication. In particular, the structures and functions of the OBP proteins and the replicative helicase have been studied in detail, and are summarized subsequently. On the other hand, much is known about the elongation phase of DNA replication in thermophilic bacteria and archaea. Many of the enzymes and factors participating in the process were purified and biochemically characterized, and the three-dimensional structures of many were determined. For studies on these proteins, the reader is referred to detailed studies and reviews on the subject (Bruck et al., 2002; Bullard et al., 2002; Cann and Ishino, 1999; Forterre and Elie, 1993; Grabowski and Kelman, 2003; Kelman, 2000a; Perler et al., 1996).

CELL CYCLE Chromosomal DNA replication takes place during the S-phase of the cell cycle and is regulated to insure that DNA replication will take place accurately, in a timely fashion, and only once per cell cycle. Although it is beyond the scope of this chapter the cell cycle will be briefly summarized subsequently. For comprehensive reviews on the subject see Lundgren and Bernander (2005) and Bernander (2007). To date there has been no study of the cell cycle of thermophilic bacteria published. Like other features of the archaeal information processes, the cell cycle has characteristics reminiscent of both bacteria and eukarya. The laboratory of Rolf Bernander performed detailed cell cycle studies of four thermophilic archaeal species; Archaeoglobus fulgidus, Methanocaldococcus jannashcii, Sulfolobus solfataricus, and Sulfolobus acidocaldarius (Hjort and Bernander, 2001; Lundgren et al., 2004; Maisnier-Patin et al., 2002). Rather than the eukaryotic G1, S, G2, and M or the prokaryotic B, C, and D phases, the archaeal cell cycle is divided into prereplicative (G1), replicative (S), and postreplicative stages (G2/M), but the relative lengths of these phases varies in different species. The euryarchaeal A. fulgidus and crenarchaeal Sulfolobus species examined have a short prereplication period (<5% of the cell cycle), a replication period of intermediate length (30–40% of the cell cycle), and a long postreplication stage (55–65% of the cell cycle) (Maisnier-Patin et al., 2002). In M. jannashcii, however, stages were not so easy to define; the cells demonstrated asymmetric division and unequal distribution of chromosomal DNA to daughter cells (Maisnier-Patin et al., 2002). After DNA replication, DNA segregation occurs, and again, all archaea are not alike. In Methanothermobacter thermautotrophicus, genome segregation occurs at approximately the same time as DNA replication (Majernik et al., 2005), but in S. acidocaldarius there is a gap between replication and segregation (Poplawski and Bernander, 1997). Evidence of chromosome condensation and even some alignment in Sulfolobus species was also observed, suggesting a mitosis-like division (Poplawski and Bernander, 1997). Although the cell cycle of only a few archaeal organisms has been studied, it is clear that there is no general rule regarding the mechanism of cell division in this domain, and thus it is not possible to speculate about general characteristics of the cell cycle of thermophiles.

ORIGIN OF REPLICATION Both bacteria and archaea contain circular genomes. Although most organisms studied to date contain a single chromosome, multiple chromosomes have been noted in several thermophilic bacterial and archaeal species (Table 16.1).

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TABLE 16.1 Summary of Genome Organization and Initiation Proteins in Thermophilic Bacteria and Archaea Bacteria Thermus thermophilus strain HB27 Thermotoga maritima MSB8 Thermoanaerobacter tengcongensis MB4 Thermobifida fusca YX Thermosynechococcus elongatus BP-1 Streptococcus thermophilus CNRZ1066 Symbiobacterium thermophilum IAM 14863

Archaea Aeropyrum pernix K1 Archaeoglobus fulgidus DSM4304 Methanocaldococcus jannaschii DSM2661 Methanopyrus kandleri AV19 Methanothermobacter thermautotrophicus Nanoarchaeum equitans Kin4-M Natronomonas pharaonis sp Picrophilus torridus DSM 9790 Pyrobaculum aerophilum IM2 Pyrococcus abyssi GE5 Pyrococcus furiosus DSM 3638 Sulfolobus acidocaldarius DSM 639 Sulfolobus solfataricus P2 Sulfolobus tokodaii strain 7 Thermococcus kodakarensis KOD1 Thermoplasma acidophilum DSM 1728 Thermoplasma volcanium GSS1

Group

Optimal Temp. (°C)

Number of Chromosomes

DnaB

DnaA

Deinococcus-Thermus Thermotogae Firmicutes

68 80 75

2 1 1

1 1 1

1 1 1

Actinobacteria Cyanobacteria Firmicutes Actinobacteria

50–55 55 45 60

1 1 1 1

1 1 1 1

1 1 1 1

Group

Optimal Temp. (°C)

Number of Chromosome

MCM

Cdc6

Crenarchaeota Euryarchaeota Euryarchaeota

90–95 83 85

1 1 3

1 1 4

2 2 1

Euryarchaeota Euryarchaeota

98 65–70

1 1

2 1

N.I.* 2

Nanoarchaeota Euryarchaeota Euryarchaeota Crenarchaeota Euryarchaeota Euryarchaeota Crenarchaeota Crenarchaeota Crenarchaeota Euryarchaeota Euryarchaeota Euryarchaeota

80–90 86 60 100 103 100 70–75 85 80 85 59 60

1 3 1 1 1 1 1 1 1 1 1 1

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

1 2 1 1 2 2 3 3 3 2 2 5

* N.I., not identified, no homolog was detected by sequence alignment.

In all organisms, origins of replication share common features and are responsible for the initiation of bidirectional DNA synthesis via similar mechanisms. All origins are rich in A and T residues and contain one or more AT-rich stretches essential for origin function. They also contain purine/ pyrimidine stretches and inverted repeats of various sizes which facilitate initiator protein binding (Boulikas, 1996; Pearson et al., 1996). The first putative origins of replication in thermophilic archaea were identified using in silico skew analysis. This method is based on the observation of a strand-specific bias in nucleotide, oligomer, and codon frequencies between the leading and lagging strands. These observations permit the detection of an origin by plotting the skew content along the genome (Lobry, 1996). Other in silico analyses, such as Z-curve analysis, have also been used to identify archaeal origins

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(Zhang and Zhang, 2005). Most putative archaeal origins identified in these ways have been found to be located close to a Cdc6 gene encoding the archaeal OBP protein (discussed next). These in silico observations were followed by detailed in vivo studies of the replication origin of Pyrococcus abyssi (Matsunaga et al., 2001; Matsunaga et al., 2003; Myllykallio et al., 2000), S. solfataricus (Lundgren et al., 2004; Robinson et al., 2004), and S. acidocaldarius (Lundgren et al., 2004). Limited in vitro studies were also reported in A. fulgidus (Maisnier-Patin et al., 2002). Using pulsed-field gel electrophoresis (Myllykallio et al., 2000) and two-dimensional gel electrophoresis of replication intermediates (Matsunaga et al., 2001) to determine the early replicating regions of DNA, it was found that P. abyssi contains a single origin from which bidirectional DNA synthesis of the circular chromosome initiates. Replication terminates in a region of the chromosome located opposite the origin. Detailed studies using replication initiation point (RIP) mapping placed the replication initiation site within the origin region (Matsunaga et al., 2003). Using marker frequency analysis, a method of identifying early replicating DNA fragments, a putative single origin was also identified in the genome of A. fulgidus (Maisnier-Patin et al., 2002). Not all archaea, however, contain a single origin. In silico analysis identified two putative origins in the genomes of M. jannaschii (Zhang and Zhang, 2003; Zhang and Zhang, 2004) and in vivo studies identified three origins in the genome of Sulfolobus species. Using two-dimensional gel electrophoresis of replication intermediates and probes to sequences located in the vicinity of the three Cdc6 genes, it was found that S. solfataricus contains at least two origins upstream of Cdc6-1 and -3 (Robinson et al., 2004). These studies also demonstrated that bidirectional DNA synthesis initiated from both origins. Further studies identified the initiation sites within the large origin regions (Robinson et al., 2004). A complimentary study using a combination of marker frequency analysis and whole-genome microarrays was used to identify three origins in the genomes of S. solfataricus and S. acidocaldarius (Lundgren et al., 2004). Two of the origins are located in the vicinity of the Cdc6-1 and -3 genes, while the third is not near a gene encoding Cdc6 (Lundgren et al., 2004). The data and subsequent computer simulation suggested that all three origins fire simultaneously to initiate bidirectional DNA synthesis with similar rates of replication fork movement. It is not clear, however, whether all origins fire in all cells or whether different origins fire in different cells. The regulation of origin firing is another interesting and important question that needs to be addressed. Using two dimensional gel analysis, two origins of replication have also been identified in the genome of Aeropyrum pernix (Robinson and Bell, 2007). All archaeal origins identified to date are several hundred base pairs in length and have similar characteristics. They are all highly AT-rich, contain one or more long AT stretches and multiple copies of repeat elements. While the number of repeats varied between different organisms, the sequence is highly conserved [TxCAxxxGAAA, where x is any residue (Capaldi and Berger, 2004; Robinson et al., 2004)]. Many of the origins are located in close proximity to Cdc6, the gene encoding the OBP proteins (discussed subsequently), and several are located in regions that encode additional replication factors such as helicases, DNA polymerases, and polymerase accessory proteins (Bernander, 2000; Kelman and Kelman, 2004; Kelman, 2000b).

INITIATOR PROTEINS The study of the initiation of archaeal DNA replication only began several years ago, so limited information is available about the process. Primary amino acid sequence analysis of archaeal replication proteins together with biochemical and structural studies suggested that archaeal initiation is a combination of the bacterial and eukaryal systems. While all eukaryotic organisms from yeast to humans contain similar initiation factors (Bell, 2002; Bell and Dutta, 2002), each archaeal species contains a slightly different subset of the initiation proteins (Table 16.1). The difference between archaea and eukarya could have resulted from the extreme environmental conditions in which archaea have evolved or the greater genetic diversity among the various species in the prokaryotic domains.

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Nearly every archaeal species contains at least one homolog of the eukaryotic initiation protein Cdc6 and/or subunits of the origin recognition complex (ORC; the eukaryotic OBP; Table 16.1) (Myllykallio and Forterre, 2000). As the eukaryotic Cdc6 and subunits of ORC (Orc1, 4 and 5) are similar in primary amino acid sequence and it is not yet known whether the archaeal proteins are functional homologs of ORC, Cdc6, or both, they are referred to in the text as Cdc6. It is thought that one of the proteins functions in origin recognition while the other is a helicase loader, similar to the bacterial DnaA and DnaC proteins, respectively.

STRUCTURE OF THE CDC6 PROTEINS The structures of the Cdc6 protein from Pyrobaculum aerophilum (Liu et al., 2000) and A. pernix (Singleton et al., 2004) have been determined and revealed the expected two domain structure found in other members of the AAA+ family of ATPases (Neuwald et al., 1999; Ogura and Wilkinson, 2001). The protein also contains a C-terminal winged-helix (WH) domain. Domain I, at the N-terminus, has an α/β RecA-type fold that contains the nucleotide-binding pocket, which is linked to the α-helices of domain II. Domain III, at the C-terminal, is the WH domain. The structures and biochemical studies (discussed subsequently) also revealed that the Cdc6 proteins are tightly bound to ADP (Liu et al., 2000; Singleton et al., 2004). Interestingly, the archaeal Cdc6 structure is similar to that of the DnaA protein, which is the bacterial OBP (Erzberger et al., 2002). There is a slight difference in the structure of domain III, however. While Cdc6 has a WH-type fold, the bacterial DnaA protein contains a helix-turn-helix (HTH) motif. The difference may be due to the difference in DNA substrate to which the two proteins bind. While DnaA binds to a 9-bp DnaA box consensus within the bacterial origin the archaeal Cdc6 may bind to longer inverted repeats (≥11 base pairs) found within the archaeal origins (Capaldi and Berger, 2004; Lopez et al., 1999; Robinson et al., 2004). Based on primary amino acid sequence analysis, the archaeal Cdc6 proteins can be divided into two distinct subgroups, called I and II (Giraldo, 2003; Singleton et al., 2004). It is not yet clear whether these two groups of proteins have redundant or different functions. It may be that one is the functional homolog of the eukaryotic ORC while the other is the homolog of the eukaryotic Cdc6 protein. This hypothesis may be supported by the recent observations demonstrating that the two M. thermautotrophicus Cdc6 homologs bind differently to origin-specific and nonspecific sequences (described subsequently).

BIOCHEMICAL PROPERTIES OF THE CDC6 PROTEINS The first biochemical property reported for the archaeal Cdc6 proteins was the ability to undergo autophosphorylation on Ser residues, utilizing the γ-phosphate of ATP or dATP (De Felice et al., 2003; De Felice et al., 2004a; Grabowski and Kelman, 2001; Kasiviswanathan et al., 2005; Shin et al., 2003a). Autophosphorylation is inhibited in the presence of single-stranded (ss) or doublestranded (ds)DNA (Grabowski and Kelman, 2001). This indirect assay was the first experimental evidence for DNA binding by the archaeal Cdc6 proteins. Autophosphorylation is also regulated by minichromosome maintenance (MCM) binding to Cdc6 (Kasiviswanathan et al., 2005). While the role of autophosphorylation for Cdc6 function is not yet known, it may play a regulatory role during initiation, as all archaeal Cdc6 proteins studied to date exhibit this autophosphorylation activity. To prevent rereplication in eukaryotes, the Cdc6 protein is either degraded or exported out of the nucleus after initiation (Bell and Dutta, 2002). In archaea, autophosphorylation may be stimulated during the initiation process (e.g., upon helicase loading), which may lead to enzyme inactivation. The structures of the Cdc6 proteins revealed the presence of a WH domain at the C-terminus, suggesting a dsDNA binding activity. The role of the WH domain in DNA binding was initially demonstrated with the autophosphorylation assay of the Cdc6 proteins from M. thermautotrophicus. It was found that while autophosphorylation is inhibited by ss and dsDNA, no inhibition by dsDNA was observed when a truncated form of Cdc6, in which the WH domain was removed, was used

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(Grabowski and Kelman, 2001). ssDNA retains the ability to inhibit the autophosphorylation of the truncated enzyme, suggesting a role of the WH domain in dsDNA binding. Subsequent DNA binding studies with Cdc6 proteins from M. thermautotrophicus and S. solfataricus show that the proteins bind ss and dsDNA in a sequence-independent manner (Capaldi and Berger, 2004; De Felice et al., 2004a; Grabowski and Kelman, 2001; Grainge et al., 2003; Robinson et al., 2004) and that the DNA binding is inhibited by MCM binding to Cdc6 (Kasiviswanathan et al., 2006). As described previously, the archaeal origins contain a number of inverted repeats. In vitro and in vivo studies with the Cdc6 proteins of M. thermautotrophicus and S. solfataricus show that the proteins preferentially bind to these inverted repeats compared with random dsDNA sequences (Capaldi and Berger, 2004; Matsunaga et al., 2001; Robinson et al., 2004). Studies with the Cdc6 homolog from A. fulgidus demonstrated that the protein preferentially binds to forked or bubble DNA structures in comparison with ss or dsDNA (Grainge et al., 2003). In addition, studies with the two Cdc6 proteins from M. thermautotrophicus illustrated that they bind differently to ss and dsDNA, One of the two Cdc6 homologs found in the organism binds both ss and dsDNA but the other binds only dsDNA (Kasiviswanathan et al., 2006). These observations, along with the structural similarity to the bacterial OBP, DnaA, support the hypothesis that the archaeal Cdc6 proteins function in origin recognition (discussed subsequently). Although all archaeal Cdc6 proteins belong to the AAA+ family of ATPases, no catalytic ATPase activity has been reported for any Cdc6 protein to date. This finding may be explained by the observation that the proteins are so tightly bound to ADP that only protein denaturation can remove the bound nucleotide (Liu et al., 2000; Singleton et al., 2004).

HELICASE Genetic and biochemical studies strongly suggest that the eukaryotic MCM complex is the replicative helicase (Forsburg, 2004). In eukaryotes, MCM is a family of six essential proteins (MCM2–7) with highly conserved amino acid sequences. All thermophilic archaeal species contain at least one homolog of the MCM helicase (Tables 16.1 and 16.2). In bacteria, the helicase is the product of the DnaB gene and all thermophilic bacteria for which the genome is known to contain a DnaB homolog (Tables 16.1 and 16.2), making it the likely replicative helicase.

MCM STRUCTURE The structure of the archaeal MCM complex is not yet clear. The MCM homologs of S. solfataricus (Carpentieri et al., 2002; Pucci et al., 2004), A. fulgidus and A. pernix (Grainge et al., 2003), and Thermoplasma acidophilum (unpublished observation) form hexamers in solution while the M. thermautotrophicus enzyme appears to form dodecamers (Chong et al., 2000; Kelman et al., 1999; Shechter et al., 2000). The dodecameric form was also suggested by the crystal structure of the N-terminal portion of the M. thermautotrophicus MCM (Fletcher et al., 2003). However, electron microscope reconstructions of the full-length M. thermautotrophicus enzyme revealed hexameric (Pape et al., 2003), heptameric (Yu et al., 2002), dodecameric (Gomez-Llorente et al., 2005), filamentous (Chen et al., 2005), and open ring (Gomez-Llorente et al., 2005) structures. The helicase consists of two main parts. The C-terminal region contains the helicase catalytic domains. Studies with an intact N-terminal portion of M. thermautotrophicus showed that the region is needed for protein multimerization (Carpentieri et al., 2002; Chong et al., 2000; Fletcher et al., 2003; Kasiviswanathan et al., 2004; Poplawski et al., 2001; Pucci et al., 2004). A high-resolution three-dimensional structure of the N-terminal part of M. thermautotrophicus MCM revealed the protein to be a dumbbell-shaped double hexamer (Fletcher et al., 2003). Each monomer folds into three distinct domains. Domain A, at the N-terminus, is mostly α-helical. Domain B has three β-strands and contains a zinc-finger motif. Domain C, positioned between domains A and B, connects the N-terminal portion of the enzyme to the catalytic region.

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TABLE 16.2 Replication Proteins in Thermophilic Bacteria and Archaea Bacteria

Archaea

Origin Origin recognition Helicase loader Helicase SSB Primase Polymerase Clamp loader

Single DnaA N.I.* DnaB SSB (one subunit) DnaG (one subunit) Pol III core (three subunits) γ-complex (five subunits)

Single or multiple Cdc6 Cdc6 MCM SSB/RPA† (one or three subunits) Primase (two subunits) PolB/PolD‡ (one or two subunits) RFC (two subunits)

DNA sliding clamp

β clamp RNase H DNA ligase Pol I

RNase H DNA ligase Fen1

Okazaki fragment maturation

PCNA§ (one or three subunits)

Abbreviations: Fen1, flap endonuclease; MCM, mini-chromosome maintenance; PCNA, proliferating cell nuclear antigen; RFC, replication factor C; RPA, replication protein A; SSB, single-strand DNA-binding protein. * N.I., not identified, no homolog was detected by sequence alignment. † Creanarchaea have homologs of bacterial SSB and euryarchaea have homologs of eukaryal RPA. ‡ PolD is a euryarchaeal-specific polymerase. § The creanarchaeal PCNA ring is a heterotrimer while the euryarchaeal PCNA ring is a homotrimer.

The domain contains five β-strands that form an oligonucleotide-/oligosaccharide-binding (OB) fold and a β-finger motif. The domain is necessary and sufficient for hexamer/dodecamer formation (Kasiviswanathan et al., 2004). Based on the three-dimensional structure, it was proposed that the zinc-finger motif is required for dodecamer formation (Fletcher et al., 2003). Biochemical data, however, derived from mutant proteins devoid of Zn binding (Poplawski et al., 2001) or in which the entire domain B was removed (Kasiviswanathan et al., 2004) showed that the proteins retain their dodecameric or filamentous structure. In addition, domain C, which does not contain the zinc finger, was shown to form double hexamers in solution by itself (Kasiviswanathan et al., 2004). Furthermore, all archaeal MCM proteins contain a similar zinc-finger motif (Poplawski et al., 2001), yet most are hexameric and not dodecameric. The exact region(s) needed for dodecamer formation are currently unknown. The structure of MCM on DNA was studied with the enzyme from S. solfataricus using fluorescence resonance energy transfer (FRET) analysis. These studies revealed that, as one would expect, the C-terminal catalytic region faces the duplex DNA while the N-terminal part binds to the ssDNA portion behind the helicase following unwinding (McGeoch et al., 2005).

BIOCHEMICAL PROPERTIES OF MCM The biochemical properties of the archaeal MCM helicase have been extensively studied since the initial report on the enzyme from M. thermautotrophicus in 1999 (Chong et al., 2000; Kelman et al., 1999; Shechter et al., 2000). These studies elucidated many of the properties of the enzymes from different organisms (Carpentieri et al., 2002; Chong et al., 2000; De Felice et al., 2003; De Felice et al., 2004a; De Felice et al., 2004b; Fletcher et al., 2003; Fletcher et al., 2005; Grainge et al., 2003; Kasiviswanathan et al., 2004; Kelman et al., 1999; McGeoch et al., 2005; Pucci et al., 2004; Shechter et al., 2000). All MCM proteins possess ATP- and dATP-dependent 3′ → 5′ helicase activity with a processivity of greater than 500 bp. Although all can efficiently unwind a DNA

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substrate that contains only a 3′-ssDNA overhang region, the helicases demonstrated a more robust activity when provided with a forked DNA structure. As expected for a helicase, the proteins bind DNA. Mutational analysis of the M. thermautotrophicus MCM protein revealed that the zinc-finger motif participates in ss and dsDNA binding (Kasiviswanathan et al., 2004; Kasiviswanathan et al., 2006; Poplawski et al., 2001). Although the mutation reduced DNA binding, it did not abolish it. A mutation in the β-finger motif, on the other hand, completely abolished both ss and dsDNA binding (Fletcher et al., 2003; Kasiviswanathan et al., 2006). As expected from the inability to bind DNA, the mutant proteins cannot support helicase activity. S. solfataricus MCM contains two β-finger motifs (McGeoch et al., 2005). One is in the N-terminal region in a location similar to that of the M. thermautotrophicus enzyme. The other motif is located in the catalytic domain. Mutation of a single motif did not abolish DNA binding; only mutations in both β-fingers abolished DNA binding (McGeoch et al., 2005). However, both β-finger motifs are needed for efficient helicase activity. A mutation in the N-terminal motif substantially reduced helicase activity in comparison with the wild-type enzyme, while a mutation in the motif located at the catalytic domain or mutations in both regions completely abolished helicase activity (McGeoch et al., 2005). In addition to ss and dsDNA binding, studies with the MCM enzyme from A. fulgidus revealed that the enzyme binds preferentially to bubble structures compared with ss and dsDNA substrates (Grainge et al., 2003). The study also demonstrated cooperative binding between the hexamers on bubble substrates which was not observed with other substrates studied (Grainge et al., 2003). In addition to the ability of MCM to translocate along ssDNA, the enzymes from the archaeons M. thermautotrophicus and T. acidophilum were shown to translocate along duplex DNA as well (Kasiviswanathan et al., 2004; Kasiviswanathan et al., 2005; Kasiviswanathan et al., 2006; Shin et al., 2003b). The role of dsDNA translocation is currently unknown. It was proposed that this could play an important role during the initiation (and perhaps elongation) process (Laskey and Madine, 2003; Takahashi et al., 2005) (discussed subsequently) as replicative helicases from each of the three domains of life were shown to translocate along duplex DNA. The enzymes studied included the DnaB proteins from Thermus aquaticus and Escherichia coli (Kaplan and O’Donnell, 2002), and the MCM4,6,7 complexes from Schizosaccharomyces pombe and Saccharomyces cerevisiae (Kaplan et al., 2003; Shin et al., 2003b).

MECHANISM OF HELICASE ASSEMBLY AT THE ORIGIN Once a region of locally unwound DNA has been established by the OBP, helicase loading occurs before the DNA polymerase is recruited. The assembly of the helicase at the origin is considered to be the last step of the initiation process. The mechanism of helicase loading at the archaeal origin is currently unknown. However, the accumulating data on the biochemical properties of Cdc6 and MCM provide a possible model for the process. In addition to the biochemical properties described previously there is one additional observation that is important to formulate models on helicase loading. Besides specific Cdc6 binding to origin DNA, the archaeal Cdc6 proteins were shown to inhibit the MCM helicase (De Felice et al., 2003; De Felice et al., 2004a; Kasiviswanathan et al., 2005; Shin et al., 2003a). Using Cdc6 proteins with mutations in the AAA+ catalytic domain that rendered the enzyme incapable of ATP binding or hydrolysis it was found that ATP binding, but not hydrolysis, is required for the inhibition (Shin et al., 2003a). Studies using Cdc6 proteins devoid of DNA binding demonstrated that Cdc6–MCM interactions, but not Cdc6–DNA binding, are required for the inhibition of helicase activity (Kasiviswanathan et al., 2005). The inhibition of MCM helicase activity by Cdc6 is reminiscent of the observation made in E. coli in which the binding of the DnaC helicase loader to the DnaB helicase inhibits helicase

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G

D

H

A

I E

B J F

FIGURE 16.1 (See color insert following page 178.) Models for helicase assembly at the Methanothermobacter thermautotrophicus origin of replication.

activity. The DnaC interaction with DnaB depends on ATP binding to DnaC. Only upon proper assembly of the helicase at the origin is the ATP hydrolyzed, resulting in the severing of the interaction between DnaC and DnaB, which allows the helicase to initiate DNA unwinding. The similarity of Cdc6 to DnaC, together with other observations, can lead to various possible models for the process of helicase loading (Figure 16.1). Gaps in the available information regarding the archaeal initiation process were filled based on similarities to the bacterial and eukaryal replication systems. As described previously, the archaeal origins contain a number of inverted repeats (Figure 16.1, black arrows). Most archaea contain one or two Cdc6 homologs. In this model we assume two homologs, one that functions in origin recognition and one that functions as a helicase loader, but if the organism has only a single homolog the protein may posses both functions of origin recognition and helicase assembly. Upon binding of Cdc6 (orange) to the inverted repeats in the origin, the protein aggregates to form the initial replication bubble (Figure 16.1A) or a cruciform structure (Figure 16.1B). The cruciform structure was suggested as a possible form for the archaeal origin following Cdc6 binding (Kelman and Kelman, 2003). The other Cdc6 protein (purple), which functions as a helicase loader, then associates with MCM (green rings) and recruits it to the Cdc6–origin complex, forming a tertiary complex of the OBP, helicase loader, and helicase ternary complex (Figure 16.1C–F). Following MCM assembly at the origin, the Cdc6 proteins dissociate and release the helicase to initiate bidirectional DNA synthesis (Figure 16.1G–J). It is likely that ATP binding and/or hydrolysis plays a major role in the process because both Cdc6 proteins belong to the AAA+ family of ATPases, but since there are no data regarding the role of ATP this was not included in the model. It is not yet clear whether MCM translocates along ss or dsDNA while unwinding the chromosomes (Takahashi et al., 2005), but because the archaeal helicase can translocate on both ss and dsDNA substrates, both possibilities are depicted in the model (ssDNA translocation, Figure 16.1G; dsDNA translocation, Figure 16.1H–J). It is also not yet established whether the two helicase rings move along the DNA away from each other to form the two replication forks (Figure 16.1G and H) or that the protein remains stationary and the DNA is pulled through it (Figure 16.1I and Figure 16.1J).

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DNA REPLICATION AND CHROMATIN During chromosomal DNA replication, the replication form moves through chromatin and not naked DNA. Thus, the replication apparatus has to move along chromatin and presumably disassemble it to facilitate DNA synthesis. Most studies on archaeal DNA replication have been performed on naked DNA. Recently, however, studies of helicase activity on substrates which better resemble the chromosome structure in vivo have been performed. Archaeal genomes from the euryarchaea and nanoarchaea kingdom contain homologs of the eukaryotic histones as the major chromatin proteins (Reeve, 2003; Sandman and Reeve, 2005) forming nucleosome-like structures (Pereira and Reeve, 1998). While some crenarchaea do not have histones, their DNA is thought to be bound by another small basic protein, Alba (Sandman and Reeve, 2005). In addition to those proteins, the chromosomal DNA is also bound by transcription factor and inhibitors, RNA polymerase, and other DNA binding proteins. DNA helicases are located at the front of the replication machinery, and thus are the first to encounter the proteins that associate with the DNA. Recent studies with the M. thermautotrophicus MCM demonstrated that the helicase is capable of unwinding DNA substrates coated with various proteins, including histones and transcription factors (unpublished observations). Studies with the S. solfataricus MCM revealed that it is capable of unwinding substrates bound with Alba. It was found, however, that efficient unwinding could be observed only following Alba acetylation (Marsh et al., 2006). The acetylation may weaken the interaction of Alba with DNA and thus reduce the barrier for helicase movement.

CONCLUDING REMARKS In the last several years, studies on the mechanism of initiation of DNA replication in thermophilic archaea have led to important and interesting discoveries regarding this unique group of organisms. The study of the initiation process in thermophilic bacteria is not advanced. It is clear, however, that the process will be, at least in part, different than that of mesophilic bacteria. For example, in bacteria, the helicase is assembled at the origin by a helicase loader (DnaC or DnaI in gram-negative and gram-positive bacteria, respectively). Analysis of the complete genome of several thermophilic bacteria failed to identify a DnaC/DnaI homolog (Table 16.2). How is the DnaB helicase assembled at the origin in those organisms? Is there a functional homolog of the helicase loader with no sequence or structural similarity to the mesophilic bacteria? Can the helicase assemble itself? Hopefully, future studies will address these and other questions regarding the initiation process in thermophilic bacteria. Another important question that future studies will address is the regulation of origin firing and its coordination with other stages of the cell cycle. The identification of archaeal origins in the last several years provides a tool for such studies. It will also be interesting to see how origin use is coordinated and regulated in those organisms for which multiple origins have been identified. Several thermophilic bacterial and archaeal species contain multiple chromosomes. This is reminiscent of the situation in eukarya. It will be interesting to determine how the replication and segregation of these multiple chromosomes are coordinated and regulated. Is it similar to the mechanism in eukaryotes or via a microorganism-specific mechanism? Although in the last several years much has been learned about the archaeal helicase, less is known about the mechanism of helicase assembly at the origin. This is a key question if we would like to understand the regulation of DNA replication and thus it has to be addressed.

ACKNOWLEDGMENTS Research conducted in author’s laboratory was supported by a Research Scholar Grant from the American Cancer Society (RSG-04-050-01-GMC).

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REFERENCES Bell, S.P. (2002) The origin recognition complex: from simple origins to complex functions. Genes Dev, 16, 659–672. Bell, S.P. and Dutta, A. (2002) DNA replication in eukaryotic cells. Annu Rev Biochem, 71, 333–374. Bernander, R. (2000) Mapping of a chromosome replication origin in an archaeon [Viewpoint]. Trends Microbiol, 8, 535–537. Bernander, R. (2007) The cell cycle of Sulfolobus. Mol Microbiol 66, 557–562. Boulikas, T. (1996) Common structural features of replication origins in all life forms. J Cell Biochem, 60, 297–316. Bruck, I., Yuzhakov, A., Yurieva, O., Jeruzalmi, D., Skangalis, M., Kuriyan, J., and O’Donnell, M. (2002) Analysis of a multicomponent thermostable DNA polymerase III replicase from an extreme thermophile. J Biol Chem, 277, 17334–17348. Bullard, J.M., Williams, J.C., Acker, W.K., Jacobi, C., Janjic, N., and McHenry, C.S. (2002) DNA polymerase III holoenzyme from Thermus thermophilus identification, expression, purification of components, and use to reconstitute a processive replicase. J Biol Chem, 277, 13401–13408. Cann, I.K. and Ishino, Y. (1999) Archaeal DNA replication. Identifying the pieces to solve a puzzle. Genetics, 152, 1249–1267. Capaldi, S.A. and Berger, J.M. (2004) Biochemical characterization of Cdc6/Orc1 binding to the replication origin of the euryarchaeon Methanothermobacter thermoautotrophicus. Nucleic Acids Res, 32, 4821–4832. Carpentieri, F., De Felice, M., De Falco, M., Rossi, M., and Pisani, F.M. (2002) Physical and functional interaction between the MCM-like DNA helicase and the single-stranded DNA binding protein from the crenarchaeon Sulfolobus solfataricus. J Biol Chem, 277, 12118–12127. Chen, Y.J., Yu, X., Kasiviswanathan, R., Shin, J.H., Kelman, Z., and Egelman, E.H. (2005) Structural Polymorphism of Methanothermobacter thermautotrophicus MCM. J Mol Biol, 346, 389–394. Chong, J.P., Hayashi, M.K., Simon, M.N., Xu, R.M., and Stillman, B. (2000) A double-hexamer archaeal minichromosome maintenance protein is an ATP-dependent DNA helicase. Proc Natl Acad Sci U S A, 97, 1530–1535. De Felice, M., Esposito, L., Pucci, B., Carpentieri, F., De Falco, M., Rossi, M., and Pisani, F.M. (2003) Biochemical characterization of a CDC6-like protein from the crenarchaeon Sulfolobus solfataricus. J Biol Chem, 278, 46424–46431. De Felice, M., Esposito, L., Pucci, B., De Falco, M., Manco, G., Rossi, M., and Pisani, F.M. (2004a) Modular organization of a Cdc6-like protein from the crenarchaeon Sulfolobus solfataricus. Biochem J, 381, 645–653. De Felice, M., Esposito, L., Pucci, B., De Falco, M., Rossi, M., and Pisani, F.M. (2004b) A CDC6-like factor from the Archaea Sulfolobus solfataricus promotes binding of the mini-chromosome maintenance complex to DNA. J Biol Chem, 279, 43008–43012. Erzberger, J.P., Pirruccello, M.M., and Berger, J.M. (2002) The structure of bacterial DnaA: implications for general mechanisms underlying DNA replication initiation. EMBO J, 21, 4763–4773. Fletcher, R.J., Bishop, B.E., Leon, R.P., Sclafani, R.A., Ogata, C.M., and Chen, X.S. (2003) The structure and function of MCM from archaeal M. thermoautotrophicum. Nature Struct Biol, 10, 160–167. Fletcher, R.J., Shen, J., Gomez-Llorente, Y., Martin, C.S., Carazo, J.M., and Chen, X.S. (2005) Double hexamer disruption and biochemical activities of Methanobacterium thermoautotrophicum MCM. J Biol Chem, 280, 42405–42410. Forsburg, S.L. (2004) Eukaryotic MCM proteins: beyond replication initiation. Microbiol Mol Biol Rev, 68, 109–131. Forterre, P. and Elie, C. (1993) Chromosome structure, DNA topoisomerases, and DNA polymerases in archaebacteria (archaea). In Kates, M., Kushner, D., and Matheson, A.T. (eds.), The Biochemistry of archaea (archaebacteria). Elsevier, Amsterdam; New York, pp. 325–365. Giraldo, R. (2003) Common domains in the initiators of DNA replication in Bacteria, Archaea and Eukarya: combined structural, functional and phylogenetic perspectives. FEBS Microbiol Rev, 26, 533–554. Gomez-Llorente, Y., Fletcher, R.J., Chen, X.S., Carazo, J.M., and Martin, C.S. (2005) Polymorphism and Double Hexamer Structure in the Archaeal Minichromosome Maintenance (MCM) Helicase from Methanobacterium thermoautotrophicum. J Biol Chem, 280, 40909–40915. Grabowski, B. and Kelman, Z. (2001) Autophosphorylation of the archaeal Cdc6 homologues is regulated by DNA. J Bacteriol, 183, 5459–5464.

9214_C016.indd 275

11/5/2007 4:33:14 PM

276

Thermophiles: Biology and Technology at High Temperatures

Grabowski, B. and Kelman, Z. (2003) Archaeal DNA replication: Eukaryal Proteins in a Bacterial Context. Annu Rev Microbiol, 57, 487–516. Grainge, I., Scaife, S., and Wigley, D.B. (2003) Biochemical analysis of components of the pre-replication complex of Archaeoglobus fulgidus. Nucleic Acids Res, 31, 4888–4898. Hjort, K. and Bernander, R. (2001) Cell cycle regulation in the hyperthermophilic crenarchaeon Sulfolobus acidocaldarius. Mol Microbiol, 40, 225–234. Kaplan, D.L., Davey, M.J. and O’Donnell, M. (2003) Mcm4,6,7 Uses a “Pump in Ring” Mechanism to Unwind DNA by Steric Exclusion and Actively Translocate along a Duplex. J Biol Chem, 278, 49171–49182. Kaplan, D.L. and O’Donnell, M. (2002) DnaB drives DNA branch migration and dislodges proteins while encircling two DNA strands. Mol Cell, 10, 647–657. Kasiviswanathan, R., Shin, J.H., and Kelman, Z. (2005) Interactions between the archaeal Cdc6 and MCM proteins modulate their biochemical properties. Nucleic Acids Res, 33, 4940–4950. Kasiviswanathan, R., Shin, J.H., and Kelman, Z. (2006) DNA binding by the Methanothermobacter thermautotrophicus Cdc6 protein is inhibited by the minichromosome maintenance helicase. J Bacteriol, 188, 4577–4580. Kasiviswanathan, R., Shin, J.H., Melamud, E., and Kelman, Z. (2004) Biochemical Characterization of the Methanothermobacter thermautotrophicus Minichromosome Maintenance (MCM) Helicase N-terminal Domains. J Biol Chem, 279, 28358–28366. Kelman, L.M. and Kelman, Z. (2003) Archaea: an archetype for replication initiation studies? Mol Microbiol, 48, 605–615. Kelman, L.M. and Kelman, Z. (2004) Multiple origins of replication in archaea. Trends Microbiol, 12, 399–401. Kelman, Z. (2000a) DNA replication in the third domain (of life). Curr Prot Peptide Sci, 1, 139–154. Kelman, Z. (2000b) The replication origin of archaea is finally revealed. Trends Biochem Sci, 25, 521–523. Kelman, Z., Lee, J.K., and Hurwitz, J. (1999) The single minichromosome maintenance protein of Methanobacterium thermoautotrophicum ΔH contains DNA helicase activity. Proc Natl Acad Sci U S A, 96, 14783–14788. Laskey, R.A. and Madine, M.A. (2003) A rotary pumping model for helicase function of MCM proteins at a distance from replication forks. EMBO Rep, 4, 26–30. Liu, J., Smith, C.L., DeRyckere, D., DeAngelis, K., Martin, G.S., and Berger, J.M. (2000) Structure and function of Cdc6/Cdc18: implications for origin recognition and checkpoint control. Mol Cell, 6, 637–648. Lobry, J.R. (1996) Asymmetric substitution patterns in the two DNA strands of bacteria. Mol Biol Evol, 13, 660–665. Lopez, P., Philippe, H., Myllykallio, H., and Forterre, P. (1999) Identification of putative chromosomal origins of replication in Archaea [letter]. Mol Microbiol, 32, 883–886. Lundgren, M., Andersson, A., Chen, L., Nilsson, P., and Bernander, R. (2004) Three replication origins in Sulfolobus species: Synchronous initiation of chromosome replication and asynchronous termination. Proc Natl Acad Sci U S A, 101, 7046–7051. Lundgren, M. and Bernander, R. (2005) Archaeal cell cycle progress. Curr Opin Microbiol, 8, 662–668. Maisnier-Patin, S., Malandrin, L., Birkeland, N.K., and Bernander, R. (2002) Chromosome replication patterns in the hyperthermophilic euryarchaea Archaeoglobus fulgidus and Methanocaldococcus (Methanococcus) jannaschii. Mol Microbiol, 45, 1443–1450. Majernik, A.I., Lundgren, M., McDermott, P., Bernander, R., and Chong, J.P. (2005) DNA content and nucleoid distribution in Methanothermobacter thermautotrophicus. J Bacteriol, 187, 1856–1858. Marsh, V.L., McGeoch, A.T., and Bell, S.D. (2006) Influence of chromatin and single strand binding proteins on the activity of an archaeal MCM. J Mol Biol, 357, 1345–1350. Matsunaga, F., Forterre, P., Ishino, Y., and Myllykallio, H. (2001) In vivo interactions of archaeal Cdc6/Orc1 and minichromosome maintenance protein with the replication origin. Proc Natl Acad Sci U S A, 98, 11152–11157. Matsunaga, F., Norais, C., Forterre, P., and Myllykallio, H. (2003) Identification of short “eukaryotic” Okazaki fragments synthesized from a prokaryotic replication origin. EMBO Rep, 4, 154–158. McGeoch, A.T., Trakselis, M.A., Laskey, R.A., and Bell, S.D. (2005) Organization of the archaeal MCM complex on DNA and implications for the helicase mechanism. Nat Struct Mol Biol, 12, 756–762. Myllykallio, H. and Forterre, P. (2000) Mapping of a chromosome replication origin in an archaeon: Response [Response]. Trends in Microbiol, 8, 537–539.

9214_C016.indd 276

11/5/2007 4:33:14 PM

DNA Replication in Thermophiles

277

Myllykallio, H., Lopez, P., Lopez-Garcia, P., Heilig, R., Saurin, W., Zivanovic, Y., Philippe, H., and Forterre, P. (2000) Bacterial mode of replication with eukaryotic-like machinery in a hyperthermophilic archaeon. Science, 288, 2212–2215. Neuwald, A.F., Aravind, L., Spouge, J.L., and Koonin, E.V. (1999) AAA+: a class of chaperone-like ATPases associated with the assembly, operation, and disassembly of protein complexes. Genome Res, 9, 27–43. Ogura, T. and Wilkinson, A.J. (2001) AAA+ superfamily ATPases: common structure—diverse function. Genes Cells, 6, 575–597. Pape, T., Meka, H., Chen, S., Vicentini, G., van Heel, M., and Onesti, S. (2003) Hexameric ring structure of the full-length archaeal MCM protein complex. EMBO Rep, 4, 1079–1083. Pearson, C.E., Zorbas, H., Price, G.B., and Zannis-Hadjopoulos, M. (1996) Inverted repeats, stem-loops, and cruciforms: significance for initiation of DNA replication. J Cell Biochem, 63, 1–22. Pereira, S.L. and Reeve, J.N. (1998) Histones and nucleosomes in Archaea and Eukarya: a comparative analysis. Extremophiles, 2, 141–148. Perler, F.B., Kumar, S., and Kong, H. (1996) Thermostable DNA polymerases. Adv Protein Chem, 48, 377–435. Poplawski, A. and Bernander, R. (1997) Nucleoid structure and distribution in thermophilic Archaea. J Bacteriol, 179, 7625–7630. Poplawski, A., Grabowski, B., Long, S.E., and Kelman, Z. (2001) The zinc finger domain of the archaeal minichromosome maintenance protein is required for helicase activity. J Biol Chem, 276, 49371–49377. Pucci, B., De Felice, M., Rossi, M., Onesti, S., and Pisani, F.M. (2004) Amino acids of the Sulfolobus solfataricus mini-chromosome maintenance-like DNA helicase involved in DNA binding/remodeling. J Biol Chem, 279, 49222–49228. Reeve, J.N. (2003) Archaeal chromatin and transcription. Mol Microbiol, 48, 587–598. Robinson, N.P. and Bell, S.D. (2007) Extrachromosomal element capture and the evolution of multiple replication origins in archaeal chromosome. Proc Natl Acad Sci USA, 104, 5806–5811. Robinson, N.P., Dionne, I., Lundgren, M., Marsh, V.L., Bernander, R., and Bell, S.D. (2004) Identification of two origins of replication in the single chromosome of the archaeon Sulfolobus solfataricus. Cell, 116, 25–38. Sandman, K. and Reeve, J.N. (2005) Archaeal chromatin proteins: different structures but common function? Curr Opin Microbiol, 8, 656–661. Shechter, D.F., Ying, C.Y., and Gautier, J. (2000) The intrinsic DNA helicase activity of Methanobacterium thermoautotrophicum ΔH minichromosome maintenance protein. J Biol Chem, 275, 15049–15059. Shin, J.H., Grabowski, B., Kasiviswanathan, R., Bell, S.D., and Kelman, Z. (2003a) Regulation of minichromosome maintenance helicase activity by Cdc6. J Biol Chem, 278, 38059–38067. Shin, J.H., Jiang, Y., Grabowski, B., Hurwitz, J., and Kelman, Z. (2003b) Substrate requirements for duplex DNA translocation by the eukaryal and archaeal minichromosome maintenance helicases. J Biol Chem, 278, 49053–49062. Singleton, M.R., Morales, R., Grainge, I., Cook, N., Isupov, M.N., and Wigley, D.B. (2004) Conformational changes induced by nucleotide binding in Cdc6/ORC from Aeropyrum pernix. J Mol Biol, 343, 547–557. Takahashi, T.S., Wigley, D.B., and Walter, J.C. (2005) Pumps, paradoxes and ploughshares: mechanism of the MCM2-7 DNA helicase. Trends Biochem Sci, 30, 437–444. Yu, X., VanLoock, M.S., Poplawski, A., Kelman, Z., Xiang, T., Tye, B.K., and Egelman, E.H. (2002) The Methanobacterium thermoautotrophicum MCM protein can form heptameric rings. EMBO Rep, 3, 792–797. Zhang, R. and Zhang, C.T. (2003) Multiple replication origins of the archaeon Halobacterium species NRC-1. Biochem Biophys Res Commun, 302, 728–734. Zhang, R. and Zhang, C.T. (2004) Identification of replication origins in the genome of the methanogenic archaeaon, Methanocadococcus jannaschii. Extremophiles, 8, 253–258. Zhang, R. and Zhang, C.T. (2005) Identification of replication origins in archaeal genomes based on the Z-curve method. Archaea, 1, 335–346.

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DNA-Binding Proteins and DNA Topology Kathleen Sandman

CONTENTS Introduction ................................................................................................................................ Chromatin Proteins .................................................................................................................... Archaeal Histones ........................................................................................................... Alba ................................................................................................................................. Sul7 ................................................................................................................................. Sul10a ............................................................................................................................. Topoisomerases .......................................................................................................................... Conclusions ................................................................................................................................ References ..................................................................................................................................

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INTRODUCTION Extremophile microbiology has matured since the 1970s and 1980s when it was dominated by exploration of “ever-more-extreme” environments and classifying extremophiles, both as isolated organisms and through DNA sequences extracted from the environment. Prokaryotic genome sequencing has facilitated extensive comparison of related genes and gene products in extremophiles and nonextremophiles to complement molecular analyses of the processes necessary for life in extreme environments. Our understanding of prokaryotic chromosome structure has also matured from entangled DNA in a nucleoid to an organized nucleoprotein complex with three-dimensional structure and topology that respond to changes in environmental conditions. This chapter describes the architectural chromosomal (chromatin) proteins in thermophilic archaea from geothermal habitats, compares the DNA–protein complexes formed by these proteins, and discusses the contributions of both architectural proteins and enzymes to chromosome topology. In a recent comprehensive review, Charlier [1] discusses several factors that affect the stability of chromosomal DNA in thermophiles including intrinsic factors that maintain the chromosome as a circular, supercoiled molecule, and extrinsic factors such as high intracellular salt concentrations and long, linear polyamines. This chapter focuses on the roles of architectural DNA-binding proteins and topoisomerases and their influence on DNA topology and prokaryote genome stability. The dynamic complex now commonly referred to as chromatin (although this term originated in eukaryotes) includes the circular chromosome, any plasmids that may be present, and the associated small, basic, abundant architectural proteins that bind DNA with little sequence specificity. Although the term architectural evokes images of rigid assemblies, flexibility is critical for the prokaryotic chromosome where the need to maintain double-stranded character in a hot environment must be balanced with the need to facilitate local denaturation for replication, repair, and 279

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transcription [2]. The structure of chromatin at any time, then, is the outcome of a complex interplay between architectural proteins and enzymes responsible for these events. Chromatin proteins must compact the chromosome within a confined nucleoid, prevent DNA condensation in vivo at DNA concentrations approaching 100 mg/ml, constrain DNA supercoils, participate in gene expression and regulation, and stabilize the chromosome against thermal denaturation, yet each of these roles is not exclusive to chromatin proteins. Mutants lacking chromatin proteins have been isolated in several species, and the phenotypes of these mutants are numerous and subtle, confirming chromatin protein participation in many chromosomal transactions and underscoring their redundancy [3–5]. Bacteria rely almost universally on chromatin protein HU (COG0776) and the same pair of topoisomerases (topoisomerase I and gyrase) to generate chromosome topology. In contrast, archaea have an expanded repertoire of chromatin proteins and topoisomerases, with each species containing at least two different types of chromatin protein and many species with additional topoisomerases. Archaeal chromatin proteins have been recently reviewed [6], and this chapter will highlight the proteins in two archaeal species, Methanothermobacter thermautotrophicus (MT) and Sulfolobus solfataricus (SS). These organisms, in laboratory culture for decades, have become models for investigations of archaeal molecular biology. They represent two of the four phyla within the domain Archaea (Euryarchaea and Crenarchaea, respectively) (see Figure 9.1), and complete genome sequences are known. In both cases, chromosome architecture and topology have been investigated, and this chapter will address both the common challenges faced by organisms in geothermal environments and the different molecular solutions to those problems in these organisms. MT, originally named Methanobacterium thermoautotrophicum, has been cultured from a variety of thermal habitats, originally from a municipal waste treatment facility in Urbana, Illinois [7], but also from thermal springs in Yellowstone National Park [8]. A strict anaerobe, Methanothermobacter requires a reducing environment, temperatures between 45°C and 70°C, and a neutral pH. This methanogen grows autotrophically on two gaseous substrates, H2 and CO2, reducing the carbon in CO2 to methane (CH4). Sulfolobus species are typically found in acidic hot springs, as they derive energy from the oxidation of elemental sulfur to sulfuric acid. SS was isolated from volcanic hot springs near Naples, Italy [9] and grows aerobically, with a 75°C temperature optimum. MT and SS have circular chromosomes, of 1.75 and 2.99 Mbp, respectively, and no plasmids. Both organisms have in common the chromatin protein Alba, and topoisomerases III and VI, but MT encodes histones, while SS encodes an additional topoisomerase, reverse gyrase (RG), and chromatin proteins Sul7 and Sul10a. When examining the topological effects of chromatin proteins binding to DNA, local and global effects must be distinguished, that is, the effects of protein binding to a short (<100 bp) DNA fragment versus the effects of multiple copies of that protein on a large DNA molecule. The ability of the protein to bend DNA and/or distort the helix is typically evident in crystal structures, reflecting a local effect. Techniques for studying global effects in vitro include electron microscopy of protein– DNA complexes and assays to examine changes in plasmid topology upon chromatin protein binding. While local effects of protein binding on DNA can be readily extrapolated to the chromosome, global effects are more difficult to generalize for several reasons. First, large segments of the chromosome in vivo are rarely in complex with multiple copies of a single protein; second, topoisomerases also regulate global levels of supercoiling; and third, the bacterial chromosome is divided into independently supercoiled domains, and the archaeal chromosome is likely to be similarly partitioned.

CHROMATIN PROTEINS ARCHAEAL HISTONES One of the most abundant proteins in nature, histones (COG2036) organize the chromosomes of virtually all eukaryotes and many archaeal species, including MT. Histones are defined by their

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three-dimensional structure, the histone fold: a domain of three alpha helices separated by short beta-strand regions [10]. Archaeal histones comprise only the histone fold (~7.5 kDa), whereas eukaryotic histones (11–15 kDa) have embellished the histone fold with additional sequences both N-terminal and C-terminal to the fold (“tails”). Each of the alpha helices in a histone fold exhibits a hydrophobic face that is stabilized by association with a second histone fold, resulting in a globular histone dimer with a hydrophobic interior and a surface rich in the basic amino acids lysine and arginine. It is the spatial pattern of these positively charged surface residues in a histone dimer that specifies the path of the DNA as it wraps around the histones. An important difference between archaeal and eukaryotic histones lies in their selection of dimer partners: eukaryotic histones have strict associations (H2A•H2B and H3•H4) whereas archaeal histones can exhibit promiscuity. As all of the histones in an archaeon are very similar (see subsequently), they share an almost identical monomer–monomer interface, probably allowing all possible pairings. The basic unit of a histone-based chromosome is the nucleosome. Eukaryotic nucleosomes have a symmetric histone octamer structure with two copies each of the four core histones organized as a linear polymer of histone dimers: H2A•H2B/H4•H3/H3•H4/H2B•H2A [11]. Archaeal histones, on the other hand, are histone tetramers [12], analogous to the central (H3•H4)2 tetramer within the eukaryotic octamer, and histone hexamers have also been observed [13]. Based on the distribution of histone-encoding genes in archaeal genomes, and assuming that organisms that maintain histone-encoding genes express them this nucleosome-based format of chromosome structure is widespread in archaea. Most recently, Cˇubonˇová et al. [14] have expanded the known phylogenetic range of histones into the phylum Crenarchaeota with the discovery of four new histone sequences, three from the collection derived from whole-environmental sampling of the Sargasso Sea and one from Cenarchaeum symbiosum, a crenarchaeal symbiont of marine sponges. These four histone sequences are sufficiently similar to previously studied histones to be recognizable as archaeal histones, but sufficiently different to originate from distant relatives. The ability of crenarchaeal histones to generate archaeal nucleosomes in vitro was confirmed by assays to measure DNA wrapping (see subsequently). Archaeal genomes contain a variable number of histone-encoding genes, ranging from one to seven. Amino acid sequence comparisons of archaeal histones illustrate that within a specific lineage, the histones are most closely related to one another, implying duplication and sequence divergence within a lineage. While there is no correlation between the number of histone-encoding genes in a genome and the optimal growth temperature of an organism, it is noteworthy that all organisms with only a single histone gene are psychrophiles or mesophiles. In contrast, there does appear to be a correlation between the amount of histone protein present in an organism and its optimal growth temperature within the family Methanobacteriaceae. Histones have been extracted repeatedly from Methanothermus fervidus (Topt 83°C), MT (Topt 65°C), and Methanobacterium formicicum (Topt 37°C), and based on the amount of histone purified relative to measurements of total soluble protein, histones comprise ~4% of total soluble protein in M. fervidus [15], ~1% in MT [16], and significantly less than 1% in M. formicicum [17]. However, we have not observed variation in the amount of histones purified from MT grown at the high and low temperatures of its growth range. Histones comprise about 1% (±0.5%) of the total soluble protein in MT regardless of whether the growth temperature is 45°C, 55°C, or 70°C [16]. It is not known if higher histone levels in thermophiles are achieved through higher gene expression levels or lower protein turnover rates. Two additional factors contribute to the thermal stability of archaeal nucleosomes. In a biophysical study comparing the thermodynamic stability of archaeal histones from hyperthermophiles and mesophiles, M. fervidus histones were found to be intrinsically more stable at high temperatures than M. formicicum histones [18] and these effects have been attributed to structural features within the histone fold [19]. Higashibata et al. [20] demonstrated that polyamines, abundant in hyperthermophiles, not only stabilize naked DNA against thermal denaturation, but also confer additional compaction to DNA complexed with histones.

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MT encodes three archaeal histones, named here by their MT locus designation from the genome sequence, MT0254, MT0821, and MT1696 (formerly HMtB, HMtA1, and HMtA2, respectively [21]). Each has 67 residues in the mature protein, as the N-terminal methionine is proteolytically cleaved in the cases of MT0821 and MT1696, but not MT0254. Studies of native histones are complicated by the fact that the abundance of individual histones is regulated by growth phase [22], and studies of MT histones in particular are complicated by the fact that the three proteins are very similar: >75% amino acid sequence identity, molecular weights differing by only a few daltons, and isoelectric points in a narrow range from eight to nine. The DNA binding properties of each MT histone, then, have been studied by preparing recombinant histones in Escherichia coli, which are obligately homodimers in solution and homotetramers in archaeal nucleosomes. The unique organizing feature of histones is that they do not just bind and bend DNA, but provide a compact core to wrap DNA, forcing it into a tight circle, and histone assays exploit this property. The simplest assay for histone binding is an electrophoretic mobility shift assay (EMSA) of a linear DNA fragment in an agarose gel [23]. When wrapped and compacted by histones, the DNA mobility is increased (Figure 17.1). A drawback to this assay is that the DNA–histone interaction must be stable for the extended electrophoresis time (~16–20 h). Despite the high degree of similarity among the MT histones, they exhibit significant differences in DNA wrapping and compaction, and this is best, illustrated in comparisons of rMT0821 and rMT1696. In the EMSA, there are two characteristics of each histone that can be measured: the

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FIGURE 17.1 Agarose gel electrophoretic mobility shift assay (EMSA). Linear pBR322 DNA (50 ng) was incubated with increasing amounts of chromatin proteins and the complexes formed separated by electrophoresis through a 0.8% agarose gel. Increased mobility results from DNA compaction. Control lanes (–) had DNA alone. (a) Comparison of DNA compaction by rMT0821 and rMT1696. Lanes 2 through 7 have complexes formed by incubating the pBR322 DNA with 10, 20, 30, 40, 50, and 60 ng of rMT0821, and lanes 9 through 14 with 5, 10, 20, 30, 40, and 50 ng of rMT1696. (b) Electron micrographs show linear pBR322 compaction by archaeal histones [64]. (c) Comparison of DNA compaction by rHMfB [22] and rMT-AlbaR39. Lanes 2 through 5 contain complexes formed by pBR322 DNA and 10, 20, 40, and 100 ng of rHMfB and lanes 7 through 14 have complexes formed by incubation of pBR322 DNA with 100, 200, 250, 300, 350, 400, 450, and 500 ng of rMT-AlbaR39.

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degree of mobility shift, and the histone:DNA weight ratio at which the shift is saturated. Two EMSAs are illustrated in Figure 17.1, each covering the same range of histone:DNA weight ratios, and both aspects of DNA wrapping differ. The mobility of rMT0821–DNA complexes is significantly less than the mobility of rMT1696–DNA complexes, and rMT0821–DNA compaction is saturated at a much lower protein:DNA ratio than rMT1696-DNA. This assay measures the global compaction of a long DNA molecule, and differences reflect the stability of dimer–dimer interactions, a measure of DNA affinity [24]. rMT0821 and rMT1696 also differ in their DNA wrapping activity in the more precise global topology assay, illustrated in Figure 17.2. This assay measures an equilibrium topology resulting from the toroidal wrapping of DNA around nucleosomes in the presence of eukaryotic topoisomerase I. The products of the reaction are a set of plasmid topoisomers differing only in linking number, a topological property of closed circular DNA [25]. In this assay, increasing amounts of a histone result in the introduction of supercoils to the DNA, and those can be either positive or negative. High-percentage agarose gels separate topoisomers and permit identification of both the number of supercoils by visual inspection; the direction, positive or negative, is determined on twodimensional agarose gels. As previously documented for other archaeal histones [21,26], MT histones can wrap DNA in either positive or negative supercoils. This is best envisioned as resulting from a shift in the dimer–dimer interface of a DNA-bound histone tetramer that can permit either a clockwise or a

histone

(a)

de-prot

topoI

R control R

(b)

R

R

HMf R

rMT0821

+ 1

5

rMT1696

R

–

+

16

FIGURE 17.2 Topology assays. (a) Assembly of archaeal nucleosomes on relaxed circular pUC18 DNA (R) results in supercoiling of the unbound DNA. Incubation with eukaryotic topoisomerase I (topo I) relaxes these plectonemic supercoils, and de-proteinization (de-prot) results in plasmid topoisomers supercoiled in the direction and to the extent of the originally constrained supercoils. Topoisomers with different linking numbers can be separated by electrophoresis through 1.5% agarose gels and plasmids with the most supercoils, positive or negative, migrate fastest. (b) Comparison of pUC18 DNA supercoiling (500 ng) by assembly of rHMfB (HMf) [22], rMT0821, and rMT1696. Positive (+) and negative (–) supercoils were identified by twodimensional electrophoresis (data not shown). Lanes 2 through 4 contained topoisomers resulting from incubation with 100, 200, and 300 ng rHMfB, lanes 6 through 15 from incubation with 50, 100, 150, 200, 250, 300, 350, 400, 500, and 600 ng rMT0821, and lanes 17 through 23 from incubation with 50, 100, 150, 200, 250, 300, and 350 ng rMT1696.

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counterclockwise path for the DNA [27]. It has been proposed that this property of archaeal nucleosomes confers flexibility in the wake of the supercoiling changes that accompany moving polymerases. The addition of rMT0821 to relaxed DNA results in the introduction of both positive and negative supercoils, and as more protein is added, the distribution of topoisomers shifts from a mixture to exclusively positive supercoils. In contrast, addition of rMT1696 to relaxed DNA initially results in the introduction of negative supercoils, but as the amount of histone increases, the topoisomer population becomes less supercoiled overall and then positively supercoiled. The patterns observed in these two assays with rMT1696, that is, a significant mobility shift in the EMSA and the shift from negative to positive supercoiling in the topology assay, are typical of those routinely observed for histones derived from thermophiles, whereas the patterns observed with rMT0821 are more typical of a histone derived from a mesophile. MT is unique in this aspect, in that it contains examples of both types of histones, and this is perhaps a consequence of its optimal growth temperature (65°C), which falls too low for MT to be considered a hyperthermophile, yet too high for a mesophile.

ALBA Alba was originally isolated as an abundant component of nucleoid preparations from Sulfolobus species [28,29] and later independently as a chromatin protein reversibly modified by acetylation [30], hence its name Alba (Acetylation lowers binding affinity). Previously known as Sul10b and DBNP-B, this is the most widely distributed archaeal chromatin protein (COG1581), encoded in all archaeal genomes except the classes methanomicrobia and halobacteria [2]. Most archaea, including MT, encode a single Alba homolog of 10 to 10.5 kDa. Of the few species with two Alba homologs, some have two very similar Alba homologs that presumably result from a recent gene duplication event in that lineage, whereas in Sulfolobus species and Aeropyrum pernix one of the two Alba homologs, Alba2, is highly divergent, while the other, Alba1, is a more typical representative of the family [31]. Alba dimer structure has been solved by several research groups using x-ray crystallography and nuclear magnetic resonance (NMR) [32–36], and the protein exhibits a globular central body and two protruding β-hairpin arms, one from each monomer, with an appropriate geometry to penetrate the minor groove of DNA, extending over about 15 bp. Alba can pack tightly on the DNA and the binding site size has been measured at ~6 bp per Alba dimer [32], indicating an overlap of adjacent dimers. Although several research groups have observed that Alba is a dimer in solution, others have observed tetramers [34,37], and this may reflect differences in the behaviors of various Alba homologs. SS Alba exists as a mixture of (Alba1)2 homodimers and Alba1–Alba2 heterodimers [31]. With the ratios of Alba1:Alba2 measured at 19:1, virtually all of the Alba2 exists in a heterodimer form. Jelinska et al. [31] have suggested that the presence of a small amount of Alba2 may confer flexibility to Alba–DNA complexes as a way of disrupting extended homopolymers. Electron microscopy of Alba–DNA complexes shows protein filaments coating the DNA with no evidence of compaction [31,38]. This has been confirmed by the agarose gel EMSA to assay DNA compaction (Figure 17.1). The addition of increasing amounts of Alba to linear DNA fragments does not produce acceleration, but rather retardation of DNA mobility, reflecting the additional mass of the protein on the DNA but no compaction. Although Alba does not compact DNA, it nevertheless does appear to affect DNA topology [37]. In assays performed over a wide range of temperature, Alba was shown to act in a temperaturedependent manner to constrain negative supercoiling in nicked plasmids that were subsequently treated with ligase to trap the topology. Although this study reported an absence of supercoiling at 25°C, the temperature dependence was subsequently shown to reflect a structural transition in the Sulfolobus shibatae Alba (Ssh10b), a cis–trans isomerization of a Leu-Pro bond at rates that were dramatically increased at higher temperature [33].

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As its name suggests, SS Alba1 is a substrate for acetylation both at the N-terminus and residue Lys16 [30], but the consequences of acetylation in vivo are still in question. Alignment of the available Alba sequences reveals that residue 16 is Lys in a minority of sequences, although it is preserved in all Sulfolobus Alba1 sequences. In addition, although Lys16 is predicted to lie near the DNA-binding interface [32,34], measurements of the apparent dissociation constants of acetylated and recombinant (unacetylated) SS Alba with DNA have found either no difference [39] or only a three- to fourfold difference [31]. It has been suggested that rather than influence the Alba–DNA interface, the acetylated K16 might act at the level of protein oligomerization in Alba filaments [31]. Another Alba paradox relates to its ability to bind both DNA and RNA. Evidence has been presented in support of Alba as a chromatin protein that organizes DNA into filaments, but additional evidence implicates Alba in RNA binding [39,40]. While the three-dimensional structure of an Alba dimer is complementary to the DNA double helix, it may also be complementary to doublestranded RNA. Experiments supporting RNA binding have not identified any common structural features in the RNA substrate. Phylogenetic arguments hold that Alba may have originated as an RNA-binding domain, as its eukaryotic homologs play roles in RNA metabolism [41]. Although its contemporary role appears to be in chromatin organization, additional functions in RNA metabolism or stabilization cannot be discounted and have yet to be structurally modeled.

SUL7 Sul7 is the generic term given to a group of abundant, small (~7 kDa) proteins purified from the Sulfolobus nucleoid [28]. Their phylogenetic distribution is limited to Sulfolobus species, and the number of genes encoding Sul7 ranges from one to three per genome. Unlike histones and Alba, Sul7 is a monomer in solution and binds DNA as a monomer. Its three-dimensional structure has been solved by NMR and x-ray crystallography, alone and in complex with DNA, and shows a triple-stranded β-sheet lying over a double-stranded β-sheet with a C-terminal alpha helix [42–45]. The triple-stranded sheet is positively charged and also has an exposed hydrophobic surface featuring a prominent tryptophan residue, and this is the region of the protein that contacts the minor groove of DNA and intercalates the tryptophan into the double helix [46]. The binding site is only ~4 bp, and a severe kink is introduced in the DNA upon Sul7 binding. Sul7 proteins have been shown to constrain negative supercoils in topology assays performed at various temperatures [47–49], up to 80°C, and to relate local and global effects of Sul7 binding, Mai et al. [47] have calculated that a single negative supercoil is constrained in pUC18 (2.7 kb) upon binding of about 20 molecules of Sul7. Additional reported activities of Sul7 include DNA protection from thermal denaturation [29], renaturation of dsDNA at temperatures above their annealing temperatures [50], RNase [51] and as a protein chaperone [52].

SUL10A Like Sul7 proteins, Sul10a proteins were originally purified from the Sulfolobus nucleoid [28], and have therefore been designated chromatin proteins. They are widely but not universally distributed in archaea (COG3432) with one to two homologs per genome, and in SS their abundance peaks in exponential phase [53]. The structure of a Sul10a dimer has been solved [54] and shows two winged-helix (WH) domains separated by a 55-Å long two-stranded coiled coil. Each monomer contributes one strand of the coil, the dimerization region, and an intact WH domain, which contains a DNA-binding helix-turn-helix motif. In silico modeling this structure with B-form of DNA revealed that one or both partners would have to distort significantly if both WH domains were to simultaneously contact DNA [54]. Consistent with this, electron microscopy studies illustrated significant supercoiling of the DNA when bound with Sul10a [38].

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TOPOISOMERASES Global chromosome topology is a balance between forces that introduce supercoiling and forces that relax supercoiling. Moving polymerases, ATP-dependent topoisomerases, and DNA distortion by chromatin protein binding contribute to chromosome supercoiling, and those forces are adjusted locally by the action of adenosine triphosphate (ATP)-independent relaxing topoisomerases. The net level of global supercoiling is negative (underwound) in bacteria, and in archaea has been approximated from measurements of plasmids, where the supercoiling is negative for DNA isolated from mesophiles and relaxed for DNA isolated from thermophiles and hyperthermophiles [55]. Prokaryotes require at least two topoisomerases for viability, a type I and type II, and within archaea (including MT and SS) topoisomerases III and VI are the type I and II enzymes, respectively [56]. Both relax DNA: topo III relaxes negative supercoils while topo VI relaxes either positive or negative supercoils [57]. Hyperthermophiles (SS, but not MT) have a second type I topoisomerase, RG, an ATP-requiring enzyme that introduces positive supercoils. RG appears to have resulted from the fusion of a type I topoisomerase domain and a helicase domain and its precise mechanism has not yet been explained. The SS genome encodes two RG homologs, and any individual roles of the isozymes have not been explored. RG has attracted much attention, not only because of its unique enzymatic activity, but also because it is only the hyperthermophile-specific protein, found in both archaeal and bacterial hyperthermophiles, and it was hypothesized that its activity is required for chromosome stability at high temperatures. Interestingly, the first use of the newly developed genetic system in Thermococcus kodakaraensis (TK) was to test this hypothesis by deleting the gene that encodes RG (TK also has topoisomerases III and VI.) [58]. Not only could the mutant lacking RG be recovered readily, but its phenotype was subtle, as has been observed for mutants lacking chromatin proteins. The RG mutant exhibited a slower growth rate than the wild type, especially at temperatures approaching 90°C, and no other phenotypes were measured. Recent studies [59,60] have uncovered an additional renaturase activity for both topoisomerase III and RG, catalyzing the renaturation of single-stranded chromosomal regions. The importance of this enzymatic activity is underscored by the knowledge that DNA damage at high temperature is more rapid when the DNA is single-stranded [61], and therefore reannealing the chromosome after local denaturation is critical. Chen and Huang [59] measured a 100-fold difference in affinity of SS topoisomerase III for ssDNA over dsDNA. They suggested that the enzyme specifically recognizes ssDNA regions, cleaves one strand and passes the other through the break to accomplish its relaxation activity, and finally ligates, reanneals, and dissociates from the dsDNA product as a consequence of its lower affinity for that form of DNA. RG has also been demonstrated to have renaturase activity as a consequence of introducing positive supercoils. Hsieh and Plank [60] propose that RG introduces positive supercoils in an analogous fashion to topoisomerase III, by switching its binding preference from ssDNA to dsDNA as a consequence of ATP hydrolysis. These authors suggest that RG acts as a chromosome sentinel, binding single-stranded regions of the DNA through its helicase domain, and coupling the energy of ATP hydrolysis to strand passage in the topoisomerase I domain, resulting in a net effect of rewinding (i.e., the introduction of positive supercoils). In agreement with these renaturation activities of RG, recent studies have noted a physical association of RG and single-stranded binding protein [62], which may additionally target RG to single-stranded regions of the chromosome. RG has also been implicated in binding damaged DNA [61,63], although this may be a secondary effect of its binding to locally denatured chromosome segments.

CONCLUSIONS Microbes in a geothermal environment face the challenge of preserving the structural integrity of their chromosomal DNA while rendering it accessible to the enzymes that execute the reactions

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necessary for growth. Two mechanisms for protecting DNA are to cover it with chromatin proteins and to entwine the two strands to be topologically linked and overwound. The proteins responsible for these mechanisms, however, must be sufficiently flexible in their structure and activity to permit frequent, controlled local denaturation at origins of replication and promoters [2]. SS, a hyperthermophile, relies on Alba, Sul7, and Sul10a for DNA compaction and multiple topoisomerases to generate the appropriate DNA topology in its chromosome. Within its high temperature growth range, the additional DNA renaturase activities of RG and topoisomerase III contribute to chromosome maintenance. MT, a moderate thermophile, relies on Alba and the DNA wrapping activity of histones to compact the chromosome and constrain supercoils. As these organisms and many others have illustrated, there are multiple molecular mechanisms to permit life to thrive at high temperatures, and future extremophile research should prove exciting and enlightening.

REFERENCES 1. Charlier, D. and Droogmans, L., Microbial life at high temperature, the challenges, the strategies, Cell. Mol. Life Sci., 62, 2974, 2005. 2. Sandman, K. and Reeve, J.N., Archaeal chromatin proteins: different structures but common function? Curr. Opin. Micro., 8, 656, 2005. 3. Wada, M. et al., Construction and characterization of the deletion mutant of hupA and hupB genes in Escherichia coli, J. Mol. Biol., 204, 581, 1988. 4. Huisman, O. et al., Multiple defects in Escherichia coli mutants lacking HU protein, J. Bacteriol., 171, 3704, 1989. 5. Heinicke, I. et al., Mutational analysis of genes encoding chromatin proteins in the archaeon Methanococcus voltae indicates their involvement in the regulation of gene expression, Mol. Gen. Genomics, 272, 76, 2004. 6. Reeve, J.N. and Sandman, K., Chromatin and regulation, In Archaea, Garrett, R.A., and Klenk, J.-P., Eds., Blackwell Publishing, Copenhagen, 2007. 7. Zeikus, J.G. and Wolfe, R.S., Methanobacterium thermoautotrophicus, sp. n., an anaerobic, autotrophic extreme thermophile, J. Bacteriol., 109, 707, 1972. 8. Sandbeck, K.A. and Ward, D.M., Temperature adaptations in the terminal processes of anaerobic decomposition of Yellowstone National Park and Icelandic hot spring microbial mats, Appl. Env. Micro., 44, 844, 1982. 9. Zillig, W. et al., The Sulfolobus-Caldariella group: taxonomy on the basis of the structure of DNAdependent RNA polymerases, Arch. Micro., 125, 259, 1980. 10. Arents, G. and Moudrianakis, E.N., The histone fold: a ubiquitous architectural motif utilized in DNA compaction and protein dimerization, Proc. Natl. Acad. Sci. U S A, 92, 11170, 1995. 11. Luger, K. et al., Crystal structure of the nucleosome core particle at 2.8Å resolution, Nature, 389, 251, 1997. 12. Bailey, K. et al., Histone stoichiometry and DNA circularization in archaeal nucleosomes, Nucl. Acids Res., 27, 532, 1999. 13. Xie, Y. and Reeve, J.N., Transcription by an archaeal RNA polymerase is slowed but not blocked by an archaeal nucleosome, J. Bacteriol., 186, 3492, 2004. 14. Cˇubonˇová, L. et al., Histones in Crenarchaea, J. Bacteriol., 187, 5482, 2005. 15. Pereira, S.L. et al., Archaeal nucleosomes, Proc. Natl. Acad. Sci. U S A, 94, 12633, 1997. 16. Samson, R.Y., Sandman, K., and Reeve, J.N., unpublished data, 2006. 17. Darcy, T.J., Sandman, K., and Reeve, J.N., Methanobacterium formicicum, a mesophilic methanogen, contains three HFo histones, J. Bacteriol., 177, 858, 1995. 18. Li, W.-T. et al., Thermodynamic stability of archaeal histones, Biochemistry, 37, 10563, 1998. 19. Li, W.-T., Shriver, J.W., and Reeve, J.N., Mutational analysis of differences in thermostability between histones from mesophilic and hyperthermophilic Archaea, J. Bacteriol., 182, 812, 2000. 20. Higashibata, H. et al., Effect of polyamines of histone-induced DNA compaction of hyperthermophilic Archaea, J. Biosci. Bioeng., 89, 103, 2000. 21. Tabassum, R., Sandman, K.M., and Reeve, J.N., HMt, a histone-related protein from Methanobacterium thermoautotrophicum ΔH, J. Bacteriol., 174, 7890, 1992.

9214_C017.indd 287

11/5/2007 7:51:36 AM

288

Thermophiles: Biology and Technology at High Temperatures

22. Sandman, K. et al., Growth-phase-dependent synthesis of histones in the archaeon Methanothermus fervidus, Proc. Natl. Acad. Sci. U S A, 91, 12624, 1994. 23. Sandman, K. et al., HMf, a DNA binding protein isolated from the hyperthermophilic archaeon Methanothermus fervidus, is most closely related to histones, Proc. Natl. Acad. Sci. U S A, 87, 5788, 1990. 24. Marc, F. et al., Archaeal histone tetramerization determines DNA affinity and the direction of supercoiling, J. Biol. Chem., 277, 30879, 2002. 25. Bates, A.D. and Maxwell, A., DNA Topology, IRL Press, Oxford, 1993, Chap. 2. 26. Musgrave, D.R., Sandman, K.M., and Reeve, J.N., DNA binding by the archaeal histone HMf results in positive supercoiling, Proc. Natl. Acad. Sci. U S A, 88, 10397, 1991. 27. Sandman, K. and Reeve, J.N., Structure and functional relationships between archaeal and eukaryal histones and nucleosomes, Arch. Micro., 173, 165, 2000. 28. Dijk, J. and Reinhardt, R., The structure of DNA-binding proteins from eu- and archaebacteria, In Bacterial Chromatin, Gualerzi, C.O., and Pon, C.L., Eds., Springer-Verlag, Berlin, 1986, 185. 29. Reddy, T.R. and Suryanarayana, T., Novel histone-like DNA-binding proteins in the nucleoid from the acidothermophilic archaebacterium Sulfolobus acidocaldarius that protect DNA against thermal denaturation, Biochim. Biophys. Acta, 949, 87, 1988. 30. Bell, S.J. et al., The interaction of Alba, a conserved archaeal chromatin protein, with Sir2 and its regulation by acetylation, Science, 296, 148, 2002. 31. Jelinska, C. et al., Obligate heterodimerisation of the archaeal Alba2 protein with Alba1 provides a mechanism for control of DNA packaging, Structure, 13, 963, 2005. 32. Wardleworth, B.N. et al., Structure of Alba: an archaeal chromatin protein modulated by acetylation, EMBO J., 21, 4654, 2002. 33. Cui, Q. et al., Two conformations of archaeal Ssh10b, J. Biol. Chem., 278, 51015, 2003. 34. Zhao, K., Chai, X., and Marmorstein, R., Structure of a Sir2 substrate, Alba, reveals a mechanism for deacetylation-induced enhancement of DNA binding, J. Biol. Chem., 278, 26071, 2003. 35. Chou, C.-C. et al., Crystal structure of the hyperthermophilic archaeal DNA-binding protein Sso10b2 at a resolution of 1.85 angstroms, J. Bacteriol., 185, 4066, 2003. 36. Biyani, K. et al., Solution structure, stability, and nucleic acid binding of the hyperthermophile protein Sso10b2, Biochem., 44, 14217, 2005. 37. Xue, H. et al., An abundant DNA binding protein from the hyperthermophilic archaeon Sulfolobus shibatae affects DNA supercoiling in a temperature-dependent fashion, J. Bacteriol., 182, 3929, 2000. 38. Lurz, R. et al., Electron microscopic study of DNA complexes with proteins from the archaebacterium Sulfolobus acidocaldarius, EMBO J., 5, 3715, 1986. 39. Guo, R., Xue, H., and Huang, L., Ssh10b, a conserved thermophilic archaeal protein, binds RNA in vivo, Mol. Micro., 50, 1605, 2003. 40. Marsh, V.L., Peak-Chew, S.Y., and Bell, S.D., Sir2 and the acetyltransferase, Pat, regulate the archaeal chromatin protein, Alba, J. Biol. Chem., 280, 21122, 2005. 41. Aravind, L., Lakshminarayan, M.I., and Anantharamana, V., The two faces of Alba: the evolutionary connection between proteins participating in chromatin structure and RNA metabolism, Genome Biol., 4, R64.1, 2003. 42. Baumann, H. et al., Solution structure and DNA-binding properties of a thermostable protein from the archaeon Sulfolobus solfataricus, Struct. Biol., 1, 808, 1994. 43. Gao, Y.-G. et al., The crystal structure of the hyperthermophile chromosomal protein Sso7d bound to DNA, Nature Struct. Biol., 5, 782, 1998. 44. Agback, P. et al., Architecture of nonspecific protein-DNA interactions in the Sso7d-DNA complex, Nature Struct. Biol., 5, 579, 1998. 45. Krueger, J.K. et al., The solution structure of the Sac7d/DNA complex: a small-angle x-ray scattering study, Biochem., 38, 10247, 1999. 46. Baumann, H. et al., DNA-binding surface of the Sso7d protein from Sulfolobus solfataricus, J. Mol. Biol., 247, 840, 1995. 47. Mai, V.Q. et al., Small abundant DNA binding proteins from the thermoacidophilic archaeon Sulfolobus shibatae constrain negative DNA supercoils, J. Bacteriol., 180, 2560, 1998. 48. Lopez-Garcia, P. et al., In vitro DNA binding of the archaeal protein Sso7d induces negative supercoiling at temperatures typical for thermophilic growth, Nucl. Acids Res., 26, 2322, 1998.

9214_C017.indd 288

11/5/2007 7:51:36 AM

DNA-Binding Proteins and DNA Topology

289

49. Napoli, A. et al., DNA bending, compaction and negative supercoiling by the architectural protein Sso7d of Sulfolobus solfataricus, Nucl. Acids Res., 30, 2656, 2002. 50. Guagliardi, A. et al., Annealing of complementary DNA strands above the melting point of the duplex promoted by an archaeal protein, J. Mol. Biol., 267, 841, 1997. 51. Shehi, E. et al., The Sso7d DNA-binding protein from Sulfolobus solfataricus has ribonuclease activity, FEBS Lett., 497, 131, 2001. 52. Guagliardi, A., Mancusi, L., and Rossi, M., Reversion of protein aggregation mediated by Sso7d in cell extracts of Sulfolobus solfataricus, Biochem. J., 381, 249, 2004. 53. Edmondson, S.P. et al., Characterization of Sac10a, a hyperthermophile DNA-binding protein from Sulfolobus acidocaldarius, Biochemistry, 43, 13026, 2004. 54. Chen, L. et al., The hyperthermophile protein Sso10a is a dimer of winged helix DNA-binding domains linked by an antiparallel coiled coil rod, J. Mol. Biol., 341, 73, 2004. 55. Charbonnier, F. and Forterre, P., Comparison of plasmid DNA topology among mesophilic and thermophilic eubacteria and archaebacteria, J. Bacteriol., 176, 1251, 1994. 56. Serre, M.-C. and Duguet, M., Enzymes that cleave and religate DNA at high temperature: the same story with different actors, Prog. Nucl. Acids Res. Mol. Biol., 74, 37, 2003. 57. Gadelle, D. et al., Phylogenomics of type II DNA topoisomerases, BioEssays, 25, 232, 2003. 58. Atomi, H., Matsumi, R., and Imanaka, T., Reverse gyrase is not a prerequisite for hyperthermophilic life, J. Bacteriol., 186, 4829, 2004. 59. Chen, L. and Huang, L., Oligonucleotide cleavage and rejoining by topoisomerase III from the hyperthermophilic archaeon Sulfolobus solfataricus: temperature dependence and strand annealing-promoted DNA religation, Mol. Microbiol., 60, 783, 2006. 60. Hsieh, T.-S. and Plank, J.L., Reverse gyrase functions as a DNA renaturase, J. Biol. Chem., 281, 5640, 2006. 61. Kampmann, M. and Stock, D., Reverse gyrase has heat-protective DNA chaperone activity independent of supercoiling, Nucl. Acids Res., 32, 3537, 2004. 62. Napoli, A. et al., Functional interaction of reverse gyrase with single-strand binding protein of the archaeon Sulfolobus, Nucl. Acids Res., 33, 564, 2005. 63. Napoli, A. et al., Reverse gyrase recruitment of DNA after UV light irradiation in Sulfolobus solfataricus, J. Biol. Chem., 279, 33192, 2004. 64. Lurz, R., unpublished data, 2001.

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Structure and Evolution of the Thermus thermophilus Ribosome Steven T. Gregory and Albert E. Dahlberg

CONTENTS Thermus as a Model System ...................................................................................................... Structural Biology of the Thermus Protein Synthesis Machinery ............................................. Insights into Ribosome Function from Structural Studies of Thermus thermophilus Ribosomes ............................................................................. Genetics of T. thermophilus Ribosomes ......................................................................... Ribosome Synthesis in T. thermophilus ......................................................................... Evolutionary Adaptations of the Thermus Protein Synthetic Machinery to Thermal Extremes ......................................................................... Future Prospects ........................................................................................................................ Postscript .................................................................................................................................... References ..................................................................................................................................

291 293 294 295 297 297 301 301 302

The extremely thermophilic bacterium Thermus has been a major contributor to our understanding of the evolution of life at extreme temperatures and has provided fundamental insights into biology at the molecular level. The choice of Thermus as the focus of structural genomics efforts and the facile genetics of this organism have combined to make it the ideal model thermophile. One area of inquiry that clearly epitomizes the impact of this organism on biology is the investigation into the mechanism of protein synthesis. In this chapter, we describe some of the attributes of Thermus as a model organism, focusing on the genetics and structural biology of the ribosome and the mechanism by which it catalyzes the synthesis of proteins encoded by the genome.

THERMUS AS A MODEL SYSTEM Species belonging to the bacterial genus Thermus are extreme thermophiles found in terrestrial and submarine hot springs and from deep-sea thermal vents across the globe. They have also been isolated from a number of man-made habitats as well, making them among the most common and most abundant of thermophilic organisms (Williams and Sharp, 1995). The type species of the genus, Thermus aquaticus, was first isolated in 1969 from a hot spring in Yellowstone National Park in the United States by T. D. Brock (Brock and Freeze, 1969), and isolates of a closely related species, Thermus thermophilus, were later obtained from hot springs in Japan (Oshima and Imahori, 1971; Oshima and Imahori, 1974; Williams et al., 1995). Since then a number of other Thermus spp. have been identified in a variety of locales throughout the world (Williams and Sharp, 1995). 291

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Molecular sequence analyses indicate a distant phylogenetic relationship between Thermus and the radiation-resistant genus Deinococcus (Hensel et al., 1986). The Deinococcus–Thermus group, also referred to as the Deinococcus–Thermus phylum, is one of the most deeply branching bacterial lineages near the root of the universal phylogenetic tree (Chapter 1; Woese, 1987; Williams and Sharp, 1995), and does not show close affiliation to any other lineage. These conclusions are based on both protein (Griffiths and Gupta, 2004) and 16S rRNA (Williams and Sharp, 1995) sequences. Several other genera have been placed within this lineage, including Meiothermus (Nobre et al., 1996), Oceanithermus (Miroshnishenko et al., 2003a), Vulcanithermus (Miroshnichenko et al., 2003b), Marinithermus (Sako et al., 2003) and Truepera (Albuquerque et al., 2005). While Deincoccus is comprised of mesophilic (Brooks and Murray, 1981), slightly thermophilic (Ferreira et al., 1997), and psychrophilic (Hirsch et al., 2005) species, and the recently discovered Truepera (Albuquerque et al., 2005) is mesophilic, all other genera are moderate to extremely thermophilic. The thermophilic nature of so many members of this phylum would seem to suggest that thermophily is a primitive character of this group. However, a recent genomic comparison of T. thermophilus and Deinococcus radiodurans supports a mesophilic, or at most a moderately thermophilic, last common ancestor of Thermus and Deinococcus (Omelchenko et al., 2005), leaving this fundamental question open for debate. In any case, the relationship of Thermus to Deinococcus provides the potential for a comparative approach in dissecting the evolution of thermophily within a single phylogenetic group. T. thermophilus (and to a lesser extent the closely related species T. aquaticus) is one of the most comprehensively studied thermophilic organisms, for both historical and pragmatic reasons. As aerobes, Thermus spp. are easily isolated and cultivated in the laboratory (Brock and Freeze, 1969) in contrast to many other thermophiles which are strict anaerobes requiring elaborate media and culture conditions (for instance, see Blöchl et al., 1997). The relatively small genome size of T. thermophilus (a 1.9 Mbp chromosome plus a 0.2 Mbp megaplasmid pTT27; Henne et al., 2004; Masui et al., 2005) has also made this species attractive for sequencing efforts and structural genomics projects. Complete genome sequences have been determined for two T. thermophilus strains, HB27 (Henne et al., 2004) and HB8 (Masui et al., 2005), and systematic efforts are underway to solve the three-dimensional structure of each protein encoded in the HB8 genome (Yokoyama et al., 2000). One of the most attractive features of Thermus spp. is the potential to apply a genetic approach to questions about evolutionary adaptations to thermophily (see Chapter 12). T. thermophilus is naturally competent for transformation with DNA (Koyama et al., 1986). The development of genes encoding thermostable drug-resistance proteins including a kanamycin adenyltransferase (Hashimoto et al., 2001) and a bleomycin-binding protein (Brouns et al., 2005) facilitates the generation of gene knockouts and gene replacements (Hashimoto et al., 2001) and the combination of thermostable drug-resistance markers with cloned replication origins from naturally occurring Thermus plasmids has led to the construction of shuttle and expression vectors (de Grado et al., 1999; Kayser et al., 2001; Moreno et al., 2003; Takayama et al., 2004; Kobayashi et al., 2005). Linkage experiments indicate that transformation is a potentially powerful tool for genetically mapping tightly linked markers (Hoshino et al., 1994). Additional mechanisms of gene transfer may eventually be developed that would allow recombination mapping of markers separated by greater genetic distances. While phages specific for Thermus spp. have been identified (Sakaki and Oshima, 1975; Yu et al., 2006), generalized transduction has yet to be demonstrated. An Hfr-like transfer mechanism has been described for T. thermophilus HB8 (Ramírez-Arcos et al., 1998), although the origin of transfer for this Hfr element has not been identified and its use has been limited to genetic analysis of genes required for anaerobic growth. Further, it appears that this Hfr element may be unstable, and has been lost from the sample of HB8 used to determine the genome sequence (Brüggemann and Chen, 2006). Nevertheless, there exists the possibility that additional gene transfer methods will eventually be developed. The efficacy of genetic mapping by any approach will also be dependent on the accumulation of suitable mutations with selectable phenotypes, and these are currently limited in Thermus spp. (see Chapter 12).

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Proteins and RNAs from Thermus spp. have also proven to be exceptionally valuable for x-ray crystallographic analysis, presumably as a result of their intrinsic conformational stability. The contribution of Thermus to problems in structural biology cannot be overstated. As of May 2006, a search of the RCSB database (www.rcsb.org) using the keyword “Thermus thermophilus” produced 567 hits. Perhaps the most striking illustration of this organism’s impact on structural biology is in the number of solutions of large macromolecular complexes, such as the DNA polymerase I-DNA complex (Eom et al., 1996), the RNA polymerase holoenzyme-promoter DNA complex (Murakami et al., 2002), the type-V ATPase (Makiyo et al., 2005), the 30S ribosomal subunit (Schlüenzen et al., 2000; Wimberly et al., 2000), the 70S ribosome complexed with tRNAs and mRNA (Yusupov et al., 2001), and most recently, the 70S ribosome complexed with release factors RF-1 or RF-2 (Petry et al., 2005).

STRUCTURAL BIOLOGY OF THE THERMUS PROTEIN SYNTHESIS MACHINERY T. thermophilus HB8 has contributed more than any other organism to an atomic level view of the protein synthetic apparatus. Early crystallographic efforts produced crystals of 30S subunits (Yonath et al., 1988), 70S ribosomes (Trakhanov et al., 1989) and 50S subunits (Volkmann et al., 1990). These initial crystals diffracted only to low resolution (9.9 Å, 20 Å, and 9 Å, respectively) and it was only much later, when the technical difficulties of working with some of these crystals were overcome, that truly informative data began to emerge. Because of the seemingly insurmountable task of solving the structure of a complex as large and as asymmetric as the ribosome, a number of laboratories focused their efforts on solving the ribosome in pieces, from individual proteins to small protein-rRNA complexes, in many instances using T. thermophilus or T. aquaticus as sources for crystallizable material. These efforts proved quite fruitful, as x-ray and NMR structures of individual components of the protein synthetic apparatus accumulated. These structures were crucial for the interpretation of electron density maps of intact ribosomes that would later be solved by x-ray crystallography. Structures of Thermus protein synthesis factors were solved, including elongation factor EF-Tu (Berchtold et al., 1993; Kjeldgaard et al., 1993), the EF-Tu-tRNA-GTP ternary complex (Nissen et al., 1995), elongation factor EF-G (Czworkowski et al., 1994), the EF-Tu·EF-Ts complex (Wang et al., 1997), ribosome release factor (RRF) (Toyoda et al., 2000) and elongation factor EF-P (Hanawa-Suetsugu et al., 2004). Single Thermus ribosomal protein structures were also solved, including S6 (Lindahl et al., 1994), L1 (Nikonov, 1996), S7 (Wimberly et al., 1997), S8 (Nevskaya et al., 1998), both wild-type (Unge et al., 1998) and mutant (Davydova et al., 2002) L22, L30 (Fedorov et al., 1999), S19 (Helgstrand et al., 1999), S16 (Allard et al., 2000), L36 (Hard et al., 2000), L18 (Woestenenk et al., 2002), L23 (Ohman et al., 2003), L16 (Nishimura et al., 2004) and L27 (Wang et al., 2004), and the S15-rRNA complex (Nikulin et al., 2000). A complex of proteins S6, S15, S18 and their 16S rRNA binding site was solved by crystallography, revealing a subunit assembly intermediate (Agalarov et al., 2000). At the same time, longstanding efforts to solve structures using the crystals of the intact T. thermophilus 30S subunit and the entire 70S ribosome-tRNA-mRNA complex proved fruitful. 5.5 Å resolution 30S subunit (Clemons et al., 1999) and 7.8 Å 70S ribosome (Cate et al., 1999) structures were published in 1999. These were followed by two 30S subunit structures at 3 Å resolution (Schlüenzen et al., 2000; Wimberly et al., 2000). A 5.5 Å structure of the entire 70S ribosome complexed with tRNAs was produced the next year (Yusupov et al., 2001). This period marked the beginning of a flurry of crystallographic activity, with crystal structures of the T. thermophilus 30S subunit or 70S ribosome complexed with substrates or accessory protein factors appearing. These included the 30S subunit complexed with either initiation factors IF-1 (Carter et al., 2001) or IF3 (Pioletti et al., 2001), the 70S ribosome with a translational operator sequence of mRNA (Jenner et al., 2005), and the 70S ribosome with release factors RF-1 and RF-2 and a termination codon (Petry et al., 2005). An all-atom model of the T. thermophilus 70S ribosome was also developed using the available ribosome crystallographic data (Tung and Sanbonmatsu, 2004).

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High-resolution crystal structures of T. thermophilus ribosomes can also facilitate the interpretation of low-resolution cryo-electron-microscopic (cryo-EM) reconstructions of homologous complexes, including the T. thermophilus 30S subunit-IF3 complex (McCutcheon et al., 1999), the T. thermophilus 30S subunit-Era complex (Sharma et al., 2005), and the T. thermophilus 70S ribosome-IF2 complex (Myasnikov et al., 2005). In the latter two studies the cryo-EM contours of the homologous complexes were fitted to the T. thermophilus crystal structures. These results show the utility of using T. thermophilus complexes for cryo-EM studies as well as crystallographic efforts.

INSIGHTS INTO RIBOSOME FUNCTION FROM STRUCTURAL STUDIES OF THERMUS THERMOPHILUS RIBOSOMES The goal of any structural study is, of course, to produce insights into function, and structural studies of the ribosome are no exception. Perhaps the greatest contribution of the multiple structures of T. thermophilus ribosomes is that they provide a structural framework for the interpretation of several decades of genetic and biochemical data, allowing insights into ribosome function that were previously hampered by the lack of a three-dimensional context. Some interpretations were immediately obvious upon inspection of the structures, for instance, by the placement of sites of mutations or chemical footprints into the structure. They will of course also allow a more complete interpretation of the results of any future genetic or biochemical experiments. On the gross scale, even the low-resolution structures produced tantalizing previews of the structural bonanza in store for molecular biologists. The 30S subunit is comprised of three distinct structural domains, referred to as the head, body, and platform (Wimberly et al., 2000), an overall organization that was presaged in the 16S rRNA secondary structure models derived by comparative sequence analysis some two decades earlier (Cannone et al., 2002) and from electron microscopic reconstructions in the 1970s and 1980s (Oakes et al., 1986). The structural relevance of these domains is reflected in the ability of the three 16S rRNA secondary structure domains to independently assemble into stable ribonucleoprotein particles (Agalarov et al., 1999). One interpretation of this organization is that it in part reflects the necessity for individual domains to move relative to one another, and to facilitate conformational changes attributed to the 30S subunit during different stages of protein synthesis. The most stunning insights of course resulted from the high-resolution structures. The Ramakrishnan laboratory has developed a detailed model for the decoding process that invokes an open-closed conformational transition of the 30S subunit upon cognate tRNA binding (Ogle et al., 2002). These crystallographic experiments reveal a series of interactions that are formed and broken sequentially. These include interactions between ribosomal proteins S4 and S5 on the backside of the 30S subunit, and interactions between ribosomal protein S12 and 16S rRNA on the side of the subunit facing the 50S subunit. Global domain movements of the head, body, and platform accompany this transition. These studies also revealed the stereochemical details of the mechanism of codon–anticodon recognition by specific base–base hydrogen bonding interactions between residues in 16S rRNA and the RNA mini-helix formed by the codon–anticodon interaction (Ogle and Ramakrishnan, 2005). Further, cocrystal structures of the 30S subunit bound to antibiotic inhibitors of protein synthesis (Carter et al. 2000; Brodersen et al., 2000; Pioletti et al., 2001) provide a wealth of information about antibiotic modes of action and the functions they are known to affect. In particular, the binding of aminoglycosides, known to antagonize the accuracy of mRNA decoding, act by facilitating the transition of the 30S subunit to the closed conformation. Further, the sites of mutations affecting translational accuracy in ribosomal proteins S4, S5, and S12 are all situated to affect the open-closed transition in a way that is predictable from the crystallographic data and serve to correlate a body of genetic and biochemical results with the structural details (Ogle and Ramakrishnan, 2005).

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Structures of the 70S ribosome, albeit at lower resolution, showed with greater clarity than previously achieved by other methods, the intersubunit bridges and the binding modes of intact tRNAs (Yusupov et al., 2001) and the interactions between mRNA and the 70S ribosome (Yusupova et al., 2001). Intersubunit interactions are known to be important not only for 70S ribosome formation, but for intersubunit movements, which have been shown by cryo-EM to occur during the translocation step of protein synthesis (Frank and Agrawal, 2000). A reasonable hypothesis is that such movements that accompany translocation are also important for it’s execution, and it is expected that future structures of functional complexes representing various stages of the elongation cycle will reveal rearrangements of many of the intersubunit interactions.

GENETICS OF T. THERMOPHILUS RIBOSOMES Ribosomal genes of Thermus spp. differ from those of more familiar bacterial species in some interesting ways. Most bacteria have rrn operons expressing single precursor transcripts composed of 16S rRNA-tRNA-23S rRNA-5S rRNA, with the individual mature rRNAs being processed from this nascent transcript. In contrast, rRNA genes of Thermus spp. have the highly unusual arrangement of two independently-transcribed 16S rRNA genes, rrsA and rrsB, and two operons, rrlArrfA-glyT and rrlB-rrfB-glyT, encoding 23S rRNA-5S rRNA-tRNAGly precursors (Hartmann et al., 1987; Hartmann and Erdmann, 1989; Henne et al., 2004; Masui et al., 2005). There is, correspondingly, no spacer tRNA gene as there is no rrs-rrl intergenic spacer. The four independently transcribed rRNA loci are scattered about the T. thermophilus chromosome (Henne et al., 2004; Masui et al., 2005). Such deviation from the canonical arrangement, though quite rare, has been observed in the bacteria Mycoplasma gallisepticum (Chen and Finch, 1989), Pirellula marina (Liesack and Stackebrandt, 1989), Borrelia burgdorferi (Fukunaga et al., 1992), Buchnera aphidicola (Munson et al., 1993; Rouhbakhsh and Baumann, 1995) and Wolbachia pipientis (Bensaadi-Merchermek et al., 1995), and in the archaeon Thermoplasma acidophilum (Ree and Zimmermann, 1990). The organization of ribosomal protein genes resembles that of other bacteria. All ribosomal protein genes are present in single copy (Henne et al., 2004; Masui et al., 2005). However, there is a fusion of the str, S10, and spc operons into a single, large transcription unit (Pfeiffer et al., 1995; Vysotskaya, 1997). Thermus species differ from Escherichia coli in that they lack ribosomal protein S21 (Tsiboli et al., 1994), and carry an additional ribosomal protein called Thx encoded by the thx gene, immediately following rpsT in a two-gene operon (Tsiboli et al., 1994; Leontiadou et al., 2001). The only bacterium in which this protein has been shown to exist is T. thermophilus, although thx homologs have been identified in plant chloroplasts (Schmidt et al., 1993; Yamaguchi and Subramanian, 2003), suggesting common ancestry between Thermus spp. and chloroplasts, lateral gene transfer, or convergent evolution. The cis-acting mRNA elements for translation in T. thermophilus resemble those of other bacteria. There are putative Shine–Dalgarno sequences in mRNAs upstream of AUG translation start sites, and these sequences show complementarity to the putative anti-Shine–Dalgarno sequence in 16S rRNA. Another feature commonly found in T. thermophilus genes is the overlapping of open reading frames. Comparisons among completed bacterial genome sequences reveal that T. thermophilus has one of the smallest median intergenic spacing, with only nine nucleotides in contrast to 85 nucleotides for the mesophile E. coli (Giovannoni et al., 2005). Such close intergenic spacing and frequent overlaps occur among ribosomal protein genes; for instance, the UGA of rpsH overlaps the AUG of rplF (UGAUG). The most extensive overlap of ORFs in the str operon is the junction between rpsE and rpmD, which has the sequence 5′-CCCAUGCCCAGGCUCAAGGUUAAGCU-3′ where the AUG of rpmD and the termination codon of rpsE are underlined. Overlap of open reading frames is taken to the extreme in the case of rpmH and rnpA, coding for ribosomal protein L34 and

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RNase P protein, respectively (Feltens et al., 2003). In this situation, the entire rpmH coding sequence is contained within the rnpA coding sequence, but in a different reading frame. The start of this overlapping sequence is as follows: 5′-GGAGGGAGAUGGAUGAAAGGA-3′ with the AUG of rnpA followed by the AUG of rpmH. The evolutionary adaptive value of this degree of overlapping of open reading frames is unclear, although many of the other organisms with similar degrees of intergenic contraction are also thermophiles (Giovannoni et al., 2005). Conceivably, overlap of start and stop codons eliminates the requirement for subunit dissociation and reassociation during the reinitiation of translation, which might be more problematic at higher temperatures. Despite the significant potential for the application of a genetic approach to examining ribosomes of T. thermophilus, only recently have such efforts been undertaken. In one study, the rpsQ gene encoding ribosomal protein S17 was deleted and replaced with a thermostable kanamycinresistance gene (Simitsopoulou et al., 1999). This mutant, as expected from the position of S17 in the 30S subunit assembly pathway, is viable, but exhibits a subunit assembly defect. Interestingly, this mutant is also temperature sensitive for growth. Perhaps the more significant aspect of this study was the demonstration of the feasibility of the gene knockout approach. More recent was the description of a number of antibiotic-resistant mutants of T. thermophilus with mutations in ribosomal protein and rRNA genes. The first among these were streptomycinresistant, streptomycin-dependent, and streptomycin-pseudodependent mutants containing single amino acid substitutions in ribosomal protein S12 (Gregory et al., 2001a). Almost all of these mutations had been previously described in other species, particularly E. coli, demonstrating that similar phenotypic selections could produce predictable mutations. The very first rRNA mutation, A2058G, was identified in one of the two 23S rRNA genes (Gregory et al., 2001b), indicating that it was possible to isolate dominant rRNA mutations despite the presence of two copies of each rRNA gene on the T. thermophilus chromosome. Mutants resistant to the translocation inhibitor thiostrepton were subsequently isolated and were shown to have mutations in either rplK encoding ribosomal protein L11 or in the rrl genes encoding 23S rRNA, all within the known thiostrepton binding site (Cameron et al., 2004). In this case, 23S rRNA mutants were shown to have the identical mutations in both 23S rRNA gene copies, suggesting that the mutations arose in one gene and were then transferred to the second copy by gene conversion, and indicating that mutants with homogeneous ribosome populations could in fact be isolated. A subsequent study described a large number of mutants containing base substitutions in 16S or 23S rRNA genes, with mutations arising in both rRNA gene copies, again suggesting the occurrence of an efficient gene conversion process (Gregory et al., 2005). Aminoglycoside antibiotics and capreomycin were used in selections to produce mutations in the decoding center of the 30S subunit, while chloramphenicol, tylosin, and sparsomycin were used to select for mutations in the peptidyltransferase active site. These experiments not only provided insight into mechanisms of antibiotic resistance, but also generated a number of useful genetic markers for mapping novel mutations. In addition to selecting spontaneous mutants, it is also possible to perform site-directed mutagenesis using gene replacement (Carr et al., 2005; Carr et al., 2006). Streptomycin dependence due to mutations in rpsL, the gene encoding ribosomal protein S12, is a conditional lethal phenotype, providing a highly effective counter-selection. Transformation of a streptomycin-dependent mutant with an rpsL gene carried on a nonreplicating plasmid and selection for streptomycin independence results in the efficient replacement of the streptomycin-dependent allele with the plasmid-encoded allele. This allows the introduction of any amino acid substitution, including those requiring multiple base substitutions, without prior knowledge of that substitution’s phenotype. In this way, a systematic dissection of the phenotypic effects of various amino acid substitutions at one position on streptomycin dependence was performed (Carr et al., 2005). Similarly, the influence of various

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amino acid substitutions on a post-translational methylthiolation of a conserved aspartic acid residue in S12 was determined (Carr et al., 2006). The results from this study indicate that the phenotypes of the various rpsL alleles are not due simply to the loss of this modification, but are a direct consequence of the amino acid substitution. Deletion of one of the two 16S rRNA genes has allowed the identification of streptomycin resistance and streptomycin dependence mutations in the remaining 16S rRNA gene, and should allow analyses of 16S rRNA similar to those described for rpsL (Gregory et al., unpublished results). Having a large collection of mutations with selectable phenotypes in rRNA as well as ribosomal protein genes allows the mapping of novel mutations arising in other types of selections (e.g., temperature sensitive alleles). It is also now possible to use matrix-assisted laser desorption ionization mass spectrometry (MALDI/TOF MS) to identify amino acid substitutions or changes in posttranslational modification of ribosomal proteins (Cameron et al., 2004; Carr et al., 2005; Suh et al., 2005). This is a rapid way to identify mutations in ribosomal protein genes in the absence of convenient selectable genetic markers and can serve to complement the genetic mapping approach.

RIBOSOME SYNTHESIS IN T. THERMOPHILUS The rRNA genes of T. thermophilus contain the canonical transcription antitermination signals BoxA and Box C (Hartmann and Erdmann, 1989) and genes encoding the antitermination factors NusA (Vornlocher et al., 1997) and NusG (Heinrich et al., 1992) have been identified (Henne et al., 2004; Masui et al., 2005). While these observations suggest the existence of an antitermination system similar to that of the more familiar E. coli, no actual work on the antitermination mechanism of Thermus spp. has been conducted. Studies with E. coli have shown that many ribosomal protein operons are controlled by feedback repression via the binding of specific ribosomal proteins to their own mRNA (Nomura et al., 1980). Little work has been done to examine feedback regulation of ribosomal protein operon expression in Thermus spp., but in one study where feedback repression by ribosomal protein S15 was reported, it was found that the mRNA binding site for S15 adopts a different structure from that of it’s E. coli homolog (Serganov et al., 2003). Thus, while we can expect some common themes in gene regulation, there undoubtedly will be many unexpected nuances to this phylogenetically distant group, some of which may reflect adaptations to high-temperature existence. The assembly of the T. thermophilus 30S subunit from its constituent 16S rRNA and ribosomal proteins has been examined in vitro. The three morphological features of the 30S subunit can assemble independently from the three individual 16S rRNA secondary structure domains and their constituent proteins (Agaralov et al., 1999). Using computational approaches, the assembly pathway for the T. thermophilus 30S subunit has been explored (Hamacher et al., 2006; Trylska et al, 2005). One prediction is the placement of Thx in the assembly map as a late binding protein. As analysis of T. thermophilus ribosome assembly is still at an early stage, specific adaptations that facilitate assembly at high temperature have yet to be identified.

EVOLUTIONARY ADAPTATIONS OF THE THERMUS PROTEIN SYNTHETIC MACHINERY TO THERMAL EXTREMES Substantial effort has been invested in elucidating the structural basis of thermostabilization of proteins from thermophiles, revealing that a variety of factors, including greater hydrophobic packing in protein interiors, increased numbers of ion pairs and formation of disulfide bonds in intracellular proteins, can influence protein stability (see Chapters 2 and 3). Early work demonstrated that ribosomes from T. aquaticus are highly thermostable with a melting temperature closely correlating with the maximum growth temperature of this species (Zeikus et al., 1970). Since these initial observations, however, relatively little has been learned about the mechanism of ribosome thermostabilization, even with the advent of crystal structures of the 30S subunit (Schlüenzen et al., 2000;

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Wimberly et al., 2000) and the 70S ribosome (Petry et al., 2005; Yusupov et al., 2001). This relatively minimal contribution to this question from structural biology has been primarily due to the absence of comparable structures from mesophilic or psychrophilic sources, although the recent solution of the E. coli 70S ribosome structure at 3.5 Å resolution (Schuwirth et al., 2005) should now permit a comparative approach to address this question. Given that the ribosome is composed largely of RNA, it seems reasonable to expect that some property of RNA structure will be a major determinant of stability. However, little is known about factors that determine the stability of very large RNA molecules. RNA folding is driven principally by the two forces of hydrogen bonding and base stacking rather than by hydrophobic forces as is the case for proteins. Sufficiently large RNA molecules can also adopt tertiary structure, and it is through such interactions that additional stability can be acquired. A number of well known tertiary structure motifs, including the A-minor interaction and ribose zipper, have been observed in ribosome crystal structures (Tamura and Holbrook, 2002). Examination of a thermostable ribozyme suggests that, as with proteins, relatively minor tertiary structure changes can create substantial thermostabilization (Fang et al., 2001). More recently, comparisons of crystal structures of RNase P RNAs from T. thermophilus and E. coli (Krasilnikov et al., 2004; Baird et al., 2006) indicate the importance of specific RNA tertiary interactions in RNA thermostabilization. Whatever structural adaptations have been acquired by rRNAs, they do not appear to substantially alter at least some of their conserved functional aspects, as indicated by the ability of T. thermophilus and E. coli ribosomal subunits to interact sufficiently well with one another so as to synthesize proteins in vitro (Thompson and Dahlberg, 2004). It is the current consensus view that binding of ribosomal proteins evolved to aid in folding nascent rRNA molecules (Noller, 2004). It seems highly plausible that adaptive changes in ribosomal protein-rRNA interactions could contribute substantially to ribosome stability, and there is some experimental evidence that such interactions are more stable in thermophiles. A remarkable set of experiments performed in the early 1990s demonstrated that peptide bond formation catalyzed by 50S subunits from T. aquaticus was highly resistant to protein extraction (Noller et al., 1992), whereas 50S subunits from the mesophile E. coli were shown to lose activity with such treatments. Subsequently, this resistance was shown to result from the robust binding of T. aquaticus ribosomal proteins L2, L3, L13, L15, L17, L18, L21, and L22 to 23S rRNA (Khaitovich et al., 1999a). Greater stability of ribosomal protein-RNA interactions may also explain in part the observation that functional T. aquaticus 50S subunits can be reconstituted from in vitro-transcribed rRNA and ribosomal proteins, whereas E. coli subunits require natural, modified rRNA (Khaitovich, 1999b). A comparative study of ribosomal protein S8 binding to it’s rRNA binding site (Gruber et al., 2003) showed that the S8-rRNA interaction in T. thermophilus and Methanococcus spp. possessed a 10-fold and 100-fold greater affinity, respectively, than the equivalent complex from E. coli. These results support thermal stability of Thermus ribosomes as being in part due to tighter binding of ribosomal proteins to rRNA. The T. thermophilus 30S subunit crystal structures reveal some features of ribosomal proteins that might be expected to positively influence structural stability at high temperatures (Brodersen et al., 2001; Schlüenzen et al., 2000; Wimberly et al., 2000). Ribosomal protein S17 of T. thermophilus consists of two domains, a β-barrel domain adopting the oligonucleotide-oligosaccharide-binding fold, and a C-terminal extension consisting of an α-helix and a 12-amino-acid tail. The α-helix and tail contact multiple 16S rRNA helices, and in all likelihood help to stabilize the RNA packing in this region (Figure 18.1). This notion is consistent with the 30S subunit assembly defect and temperature-sensitive phenotype associated with the deletion of the T. thermophilus gene encoding S17, as discussed earlier (Simitsopoulou et al., 1999). The adjunct α-helical extension is absent from S17 of most mesophiles including E. coli and Bacillus subtilis, and from psychrophiles, such as Colwellia psychrerythrea and Polaromonas napthalenivorans, although an extension is found in the mesophile Blastopirellula marina. C-terminal extensions are present in thermophiles such as Thermotoga maritima, Aquifex aeolicus, and Carboxydothermus

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S17

Thx

S17 C-terminal alignment Thermus thermophilus Thermotoga maritima Aquifex aeolicus Carboxydothermus hydrogenoform Deinococcus geothermalis Deinococcus radiodurans Blastopirellula marina Escherichia coli Bacillus subtilis Colwellia psychrerythraea Polaromonas naphthalenivorans

PISKRKRFRVLRLVESGRMDLVEKYLIRRQNYESLSKRGGKA PLSKTKRWRVVRIIQRFEPERVVKEKEDIQEEIEAVEGKGGVES PLSKTKRWVVVKILQRARRPEEEIQKQQEGQEQ PLSKEKRWRVVEIIERGKVLGEEENLETIEG PISKTKTWKVTRLIERPRGIETTAVETEGGNA PISKTKTWKVTKLIERPRGIETTLAETEVAGGEA PLSKTKRWALVRIVEKSREVDVAALKAARDQAAENLASES PLSKTKSWTLVRVVEKAVL PLSATKRFRLVEVVEEAVII PISKSKNWKLVDVITKA PISKTKNWVVTRLVQKAEIV

FIGURE 18.1 (See color insert following page 178.) Two examples of structural features potentially contributing to thermal stability of the T. thermophilus ribosome, as observed in the high-resolution crystal structure of the 30S subunit (Wimberly et al., 2000). Ribosomal protein S17 (left) has a β-barrel domain (blue) and a C-terminal α-helix and extended tail (red) that contacts multiple 16S rRNA helices. This extension is generally absent from S17 of mesophiles and psychrophiles. Even among thermophiles, the sequence of the extension is highly variable. Ribosomal protein Thx (right, in green), unique to T. thermophilus (and some plant chloroplasts), contacts multiple secondary structure elements in the head of the 30S subunit. Structures were rendered using MacPyMol 0.99 (DeLano, 2002) and PDB file 1J5E.pdb (Wimberly et al., 2000). An alignment (bottom) of selected S17 protein sequences obtained from NCBI (http://www.ncbi.nlm.nih.gov). Complete S17 protein sequences were aligned with Clustal W (Thompson et al., 1994), although for clarity only the C-terminal residues are shown. The portion of the T. thermophilus S17 sequence corresponding to the C-terminal α-helix and tail are highlighted in red. The first five sequences belong to thermophilic species, while the next four belong to mesophiles, and the last two belong to psychrophiles. Protein identification numbers are as follows: Thermus thermophilus HB8, gi|55981652; Thermotoga maritima MSB8, gi|4982055; Aquifex aeolicus VF5, gi|2982774; Carboxydothermus hydrogenoformans Z-2901, gi|94730505; Deinococcus geothermalis DSM 11300, gi|94985958; Deinococcus radiodurans R1, gi|6457994; Blastopirellula marina DSM 3645, gi|87306529; Escherichia coli W3110, gi|85676730; Bacillus subtilis subsp. subtilis str. 168, gi|16077193; Colwellia psychrerythraea, 34H gi|71147715; Polaromonas naphthalenivorans CJ2, gi|84711025.

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hydrogenoformans. Both Deinococcus geothermalis and D. radiodurans have extensions, perhaps reflecting their phylogenetic affiliation with Thermus. Interestingly, the terminal extensions found in thermophiles do not share any sequence similarity, suggesting that they probably have arisen independently as different solutions to the same problem. One feature that seems to be specific to ribosomes from Thermus spp. is the presence of ribosomal protein Thx, mentioned earlier in this chapter (Choli et al., 1993). Thx is a small, 26 amino acid α-helical polypeptide located in the head of the 30S ribosomal subunit (Wimberly et al., 2000). It contacts several rRNA helices (Figure 18.1), consistent with a putative role in enhancing structural stability. However, no direct evidence of such a role has yet been reported. Another potentially stabilizing structural feature are Zn-binding motifs in ribosomal proteins S4 and S14 observed in the T. thermophilus 30S subunit structure (Brodersen et al., 2001) and in the NMR structure of T. thermophilus ribosomal protein L36 (Hard et al., 2000). One curious observation is the existence in 50S subunits of T. thermophilus and T. aquaticus of a heptameric ribosomal protein stalk complex (L12)6/L10 rather than the more familiar pentameric (L12)4/L10 complex found in mesophiles, such as E. coli and B. subtilis (Ilag et al., 2005). This stalk region, which is highly conserved and interacts with a number of protein synthesis factors, is the most flexible component of the 50S subunit. The heptameric arrangement is not a phylogenetic peculiarity since it was also found in ribosomes of the hyperthermophile T. maritima. The fact that a complex of conserved proteins at a critical functional site of the ribosome has a variable organization is surprising. How this altered protein stoichiometry is related to a thermophilic existence is as yet unclear, although it is tempting to speculate that it may relate in some way to stabilizing ribosome–factor interactions at high temperatures. What is also unknown at this time is whether this stoichiometry is modulated at different growth temperatures. Post-transcriptional modifications of T. thermophilus tRNAs have been described and evidence suggests that they enhance thermal stability. Levels of 2′-O-methylguanosine (Kumagi et al., 1980) and of 2-thioribothymidine (Shigi et al., 2006b) increase with growth temperature. In both cases, the increase in modification was inferred or demonstrated to result from higher enzyme activity at elevated temperatures. The importance of tRNA modifications in some instances has been demonstrated, as inactivation of genes responsible for the biosynthesis of N1-methyladenosine at position 58 (m1A58) (Droogmans et al., 2003) or 2-thioribothymidine at position 54 (s2T54) (Shigi et al., 2006a) causes temperature-sensitive phenotypes. The identification and mapping of post-transcriptional modifications of T. thermophilus 16S rRNA (Guymon et al., 2006) and 23S rRNA (Mengel-Jørgensen et al., 2006) have recently been accomplished. Surprisingly, the pattern of T. thermophilus 16S rRNA modification is very similar to that of 16S rRNA of the mesophile E. coli. Of the 14 modifications found in T. thermophilus and 11 found in E. coli, 8 are common between the two species. The modifications of T. thermophilus 23S rRNA also do not differ dramatically from those of E. coli. If rRNA modifications contribute to thermal stability of the T. thermophilus ribosome, the limited number of genes potentially involved makes this hypothesis practically testable by knockout mutagenesis of genes encoding putative modification enzymes. Two of the 16S rRNA modifications found in T. thermophilus, tandem pseudouridines adjacent to the anti-Shine–Dalgarno sequence in the very 3′ terminus of the 16S rRNA molecule, are unlikely to contribute to thermal stability of the subunit, although potential stabilization of interaction with mRNA cannot been excluded. The implication from these two studies is that, in contrast with tRNAs, post-transcriptional modifications of rRNA are unlikely to be the major determinant of global thermal stability. This interpretation is bolstered by the aforementioned in vitro reconstitution of T. aquaticus 50S subunits using unmodified rRNA (Khaitovich et al., 1999b). Of course, this interpretation does not exclude a role for modifications in the stabilization of specific local structures, or in the fine-tuning of specific interactions such as substrate binding and selection during ribosome function. Indeed, most of the modified bases in rRNA are located in critical functional sites, including sites of contact with tRNAs (Guymon et al., 2006; Mengel-Jørgensen et al., 2006).

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Similarly, post-translational modifications of ribosomal proteins have also been considered as a potential contributor to ribosome stability, although few studies have addressed this question. In one report, the ribosomal protein L11 methyltransferase, encoded by the prmA gene, was found to incorporate additional methyl groups relative to the E. coli homolog (Cameron et al., 2004). However, as in E. coli, deletion of the prmA gene was found to have no detectable physiological effect, arguing against a significant role for these modifications in thermal adaptation. Finally, there are extrinsic factors that can aid in RNA stability. T. thermophilus contains a number of polyamines, including novel ones, that can bind and add thermal stability to nucleic acids, including tRNA (Terui et al., 2005) and presumably rRNA as well. In fact, early efforts to develop cell-free protein synthesis from T. thermophilus found that polyamines were an essential component for synthesis at high temperature, although it is not clear if polyamines acted by stabilizing individual components, or by stabilizing interactions between components in the system (OhnoIwashita et al., 1975). While the contribution of individual polyamines to ribosome thermal stability has yet to be systematically examined, the available evidence indicates that they will be found to be of some importance. Thus, at present, our understanding of the evolutionary adaptations acquired by thermophilic ribosomes is at a very early stage, and few generalizations can currently be drawn. However, given the wealth of structural information, and the potential for applying a genetic approach to this problem, it seems likely that some insights will be gathered in the not too distant future.

FUTURE PROSPECTS It is clear that T. thermophilus has made a significant contribution to our understanding of protein synthesis mechanisms, particularly from a structural perspective, and it seems certain that it will continue to do so in the future, even with the advent of crystal structures of ribosomes from the more familiar E. coli (Schuwirth et al., 2005). If history is any indication, future crystallographic efforts will continue to take advantage of the thermal stability of T. thermophilus ribosomes, and with further development of the genetics of ribosomes from this species, structures of mutant ribosomes will begin to appear. It is also plausible that the ability to combine genetic strategies and biophysical techniques will ultimately make T. thermophilus the preferred system for studying ribosome structure and function. One area where T. thermophilus may ultimately make its greatest contribution will be in furthering our understanding of the mechanism by which ribosomes evolve to adapt to functioning under extreme conditions. While direct comparisons of crystal structures of ribosomes from T. thermophilus and E. coli will suggest possible structural adaptations, an experimental approach using genetics and biochemistry will be required to test hypotheses posed by observations from structural biology. Understanding adaptation of ribosomes to thermophily will also have impacts beyond the study of ribosomes. Indeed, the particular adaptations of ribosomes to thermal stability should serve as a general paradigm for the description of the forces and mechanisms driving the evolution of large macromolecular complexes.

POSTSCRIPT Since the preparation of this chapter, there have been several significant developments in the structural biology of Thermus ribosomes. Perhaps the most noteworthy is the 2.8 Å crystal structure of the 70S ribosome complexed with tRNAs (Selmer et al., 2006). A lower resolution (3.7 Å) 70S structure was also produced (Korostelev et al., 2006). A 3.3 Å resolution structure of the T. thermophilus 30S subunit bound with mRNA revealed the Shine-Dalgarno interaction (Kaminishi et al., 2007). Cryo-electron microscopy showed greater details of the T. thermophilus 70S ribosome

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complexed with EF-G (Connell et al., 2007). Finally, co-crystal structures of ribosomal protein L11 complexed with its methyltransferase PrmA revealed how this modification enzyme recognizes multiple sites on its substrate protein (Demirci et al., 2007).

REFERENCES Agalarov, S. C., Selivanova, O. M., Zheleznyakova, E. N., Zheleznaya, L. A., Matvienko, N. I., and Spirin, A. S. (1999) Independent in vitro assembly of all three major morphological parts of the 30S ribosomal subunit of Thermus thermophilus. Eur. J. Biochem. 266:533–537. Agalarov, S. C., Sridhar Prasad, G., Funke, P. M., Stout, C. D., and Williamson, J. R. (2000) Structure of the S15, S6, S18-rRNA complex: assembly of the 30S ribosome central domain. Science 288:107–113. Albuquerque, L., Simões, C., Nobre, M. F., Pino, N. M., Battista, J. R., Silva, M. T., Rainey, F. A., and da Costa, M. S. (2005) Truepera radiovictrix gen. nov., sp. nov., a new radiation resistant species and the proposal of Trueperaceae fam. nov. FEMS Microbiol. Lett. 247:161–169. Allard, P., Rak, A. V., Wimberly, B. T., Clemons, W. M. Jr., Kalinin, A., Helgstrand, M., Garber, M. B., Ramakrishnan, V., and Hard, T. (2000) Another piece of the ribosome: solution structure of S16 and its location in the 30S subunit. Structure Fold. Des. 8:875–882. Baird, N. J., Srividya, N., Krasilnikov, A. S., Mondragón, A., Sosnick, T. R., and Pan, T. (2006) Structural basis for altering the stability of homologous RNAs from a mesophilic and a thermophilic bacterium. RNA 12:598–606. Bensaadi-Merchermek, N., Salvado, J. C., Cagnon, C., Karama, S., and Mouchés, C. (1995) Characterization of the unlinked 16S rDNA and 23S-5S rRNA operon of Wolbachia pipientis, a prokaryotic parasite of insect gonads. Gene 165:81–86. Berchtold, H., Reshetnikova, L., Reiser, C. O., Schirmer, N. K., Sprinzl, M., and Hilgenfeld, R. (1993) Crystal structure of active elongation factor Tu reveals major domain rearrangements. Nature 365:126–132. Blöchl, E., Rachel, R., Burggraf, S., Hafenbradl, D., Jannasch, H. W., and Stetter, K. O. (1997) Pyrolobus fumarii, gen. and sp. nov., represents a novel group of archaea, extending the upper temperature limit for life to 113°C. Extremophiles 1:14–21. Brock, T. D. and Freeze, H. (1969) Thermus aquaticus gen. n. and sp. n., a non-sporulating extreme thermophile. J. Bacteriol. 98:289–297. Brodersen, D. E., Clemons W. M., Jr., Carter, A. P., Wimberly, B. T., and Ramakrishnan, V. (2001) Crystal structure of the 30S ribosomal subunit from Thermus thermophilus: structure of the proteins and their interaction with 16S RNA. J. Mol. Biol. 316:725–768. Brodersen, D. E., Clemons, W. M. Jr., Carter, A. P., Morgan-Warren, R. J., Wimberly, B. T., Ramakrishnan, V. (2000) The structural basis for the action of the antibiotics tetracycline, pactamycin, and hygromycin B on the 30S ribosomal subunit. Cell 103:1143–1154. Brooks, B. W. and Murray, R. G. E. (1981) Nomenclature for ‘Micrococcus radiodurans’ and other radiationresistant cocci: Deinococcaceae fam. nov. and Deinococcus gen. nov., including five species. Int. J. Syst. Bacteriol. 31:353–360. Brouns, S. J. J., Wu, H., Akerboom, J., Turnbull, A. P., de Vos, W. M., van der Oost, J. (2005) Engineering a selectable marker for hyperthermophiles. J. Biol. Chem. 280:11422–11431. Brüggemann, H. and Chen, C. (2006) Comparative genomics of Thermus thermophilus: Plasticity of the megaplasmid and its contribution to a thermophilic lifestyle. J. Biotechnol. 124:654–661. Cameron, D. M., Gregory, S. T., Thompson, J., Suh, M. J., Limbach, P. A., and Dahlberg, A. E. (2004) Thermus thermophilus L11 methyltransferase, PrmA, is dispensable for growth and preferentially modifies free ribosomal protein L11 prior to ribosome assembly. J. Bacteriol. 186:5819–5825. Cameron, D., Thompson, J., Gregory, S. T., March, P. E., and Dahlberg, A. E. (2004) Thiostrepton-resistant mutants of Thermus thermophilus. Nucleic Acids Res. 32:3220–3227. Cannone, J. J., Subramanian, S., Schnare, M. N., Collett, J. R., D’Souza, L. M., Du, Y., Feng, B., Lin, N., Madabusi, L. V., Muller, K. M., Pande, N., Shang, Z., Yu, N., and Gutell, R. R. (2002) The Comparative RNA Web (CRW) Site: an online database of comparative sequence and structure information for ribosomal, intron, and other RNAs. BioMed Central Bioinformatics 3:2. Carr, J. F., Gregory, S. T., and Dahlberg, A. E. (2005) Severity of the streptomycin resistance and streptomycin dependence phenotypes of ribosomal protein S12 of Thermus thermophilus depends on the identity of highly conserved amino acid residues. J. Bacteriol. 187:3548–3550.

9214_C018.indd 302

11/5/2007 4:34:08 PM

Structure and Evolution of the Thermus thermophilus Ribosome

303

Carr, J. F., Hamburg, D. M., Gregory, S. T., Limbach, P. A., and Dahlberg, A. E. (2006) Effects of streptomycin resistance mutations on posttranslational modification of ribosomal protein S12. J. Bacteriol. 188:2020–2023. Carter, A. P., Clemons, W. M. Jr., Brodersen, D. E., Morgan-Warren, R. J., Hartsch, T., Wimberly, B. T., and Ramakrishnan, V. (2001) Crystal structure of an initiation factor bound to the 30S ribosomal subunit. Science 291:498–501. Carter, A. P., W. M. Clemons, D. E. Brodersen, R. J. Morgan-Warren, B. T. Wimberly, and V. Ramakrishnan. (2000) Functional insights from the structure of the 30S ribosomal subunit and its interactions with antibiotics. Nature 407:340–348. Cate, J. H., Yusupov, M. M., Yusupova, G. Z., Earnest, T. N., and Noller, H. F. (1999) X-ray crystal structures of 70S ribosome functional complexes. Science 285:2095–2104. Chen, X. and Finch, L. R. (1989) Novel arrangement of rRNA genes in Mycoplasma gallisepticum: separation of the 16S gene of one set from the 23S and 5S genes. J. Bacteriol. 171:2876–2878. Choli, T., F. Franceschi, A. Yonath, and B. Wittmann-Liebold. 1993. Isolation and characterization of a new ribosomal protein from the thermophilic eubacteria, Thermus thermophilus, T. aquaticus and T. flavus. Biol. Chem. 374:377–383. Clemons, W. M. Jr., May, J. L., Wimberly, B. T., McCutcheon, J. P., Capel, M. S., and Ramakrishnan, V. (1999) Structure of a bacterial 30S ribosomal subunit at 5.5 Å resolution. Nature 400:833–840. Connell, S. R., Takemoto, C., Wilson, D. N., Wang, H., Murayama, K., Terada, T., Shirouzu, M., Rost, M., Schuler, M., Giesebrecht, J., Dabrowski, M., Mielke, T., Fucini, P., Yokoyama, S., and Spahn, C. M. (2007) Structural basis for interaction of the ribosome with the switch regions of GTP-bound elongation factors. Mol. Cell. 25:751–764. Czworkowski, J., Wang, J., Steitz, T. A., and Moore, P. B. (1994) The crystal structure of elongation factor G complexed with GDP, at 2.7 Å resolution. EMBO J. 13:3661–3668. Davydova, N., Streltsov, V., Wilce, M., Liljas, A., and Garber, M. (2002) L22 ribosomal protein and effect of its mutation on ribosome resistance to erythromycin. J. Mol. Biol. 322:635–644. Demirci, H., Gregory, S. T., Dahlberg, A. E., and Jogl, G. (2007) Recognition of ribosomal protein L11 by the protein trimethyltransferase PrmA. EMBO J. 26:567–577. de Grado, M., Castán, P., and Berenguer, J. (1999) A high-transformation-efficiency cloning vector for Thermus thermophilus. Plasmid 42: 241–245. DeLano, W. L. (2002) The PyMol Molecular Graphics System. DeLano Scientific, San Carlos, CA. Droogmans, L., Roovers, M., Bujnicki, J. M., Tricot, C., Hartsch, T., Stalon, V., and Grosjean, H. (2003) Cloning and characterization of tRNA (m1A58) methyltransferase (TrmI) from Thermus thermophilus HB27, a protein required for cell growth at extreme temperatures. Nucleic Acids Res. 31:2148–2156. Eom, S. H., Wang, J., and Steitz, T. A. (1996) Structure of Taq ploymerase with DNA at the polymerase active site. Nature 382:278–281. Fang, X.-W., Golden, B. L., Littrell, K., Shelton, V., Thiyagarajan, P., Pan, T., and Sosnick. T. R., (2001) The thermodynamic origin of the stability of a thermophilic ribozyme. Proc. Natl. Acad. Sci. USA 98:4355–4360. Fedorov, R., Nevskaya, N., Khairullina, A., Tishchenko, S., Mikhailov, A., Garber, M., and Nikonov, S. (1999) Structure of ribosomal protein L30 from Thermus thermophilus at 1.9 Å resolution: conformational flexibility of the molecule. Acta Crystallogr. D Biol. Crystallogr. 55:1827–1833. Feltens, R., Gossringer, M., Willkomm, D. K., Urlaub, H., and Hartmann, R. K. (2003) An unusual mechanism of bacterial gene expression revealed for the RNase P protein of Thermus strains. Proc. Natl. Acad. Sci. USA. 100:5724–5729. Ferreira, A. C., Nobre, M. F., Rainey, F. A., Silva, M. T., Wait, R., Burghardt, J., Chung, A. P., and da Costa, M. S. (1997) Deinococcus geothermalis sp. nov. and Deinococcus murrayi sp. nov., two extremely radiation-resistant and slightly thermophilic species from hot springs. Int. J. Syst. Bacteriol. 47: 939–947. Frank, J. and Agrawal, R. K. (2000) A ratchet-like inter-subunit reorganization of the ribosome during translocation. Nature 406:318–322. Fukunaga, M., Yanagihara, Y., and Sohnaka, M. (1992) The 23S/5S ribosomal RNA genes (rrl/rrf) are separate from the 16S ribosomal RNA gene (rrs) in Borrelia burgdorferi, the aetiological agent of Lyme disease. J. Gen. Microbiol. 138:871–877. Giovannoni, S. J., Tripp, H. J., Givan, S., Podar, M., Vergin, K. L., Baptista, D., Bibbs, L., Eads, J., Richardson, T. H., Noordewier, M., Rappé, M. S., Short, J. M., Carrington, J. C., and Mathur, E. J. (2005) Genome streamlining in a cosmopolitan oceanic bacterium. Science 309:1242–1245.

9214_C018.indd 303

11/5/2007 4:34:09 PM

304

Thermophiles: Biology and Technology at High Temperatures

Gregory, S. T., Carr, J. F., Rodriguez-Correa, D., and Dahlberg, A. E. (2005) Mutational analysis of 16S and 23S rRNA genes of Thermus thermophilus. J. Bacteriol. 187:4804–4812. Gregory, S. T., Cate, J. H. D., and Dahlberg, A. E. (2001a) Streptomycin-resistant and streptomycin-dependent mutants of the extreme thermophile Thermus thermophilus. J. Mol. Biol. 309:333–338. Gregory, S. T., Cate, J. H. D., and Dahlberg, A. E. (2001b) Spontaneous erythromycin resistance mutations in a 23S rRNA gene, rrlA, of the extreme thermophile Thermus thermophilus IB-21. J. Bacteriol. 183:4382–4385. Griffiths, E. and Gupta, R. S. (2004) Distinctive protein signatures provide molecular markers and evidence for the monophyletic nature of the Deinococcus-Thermus phylum. J. Bacteriol. 186:3097–3107. Gruber, T., Köhrer, C., Lung, B., Shcherbakov, D., and Piendl, W. (2003) Affinity of ribosomal protein S8 from mesophilic and (hyper)thermophilic archaea and bacteria for 16S rRNA correlates with the growth temperatures of the organisms. FEBS Lett. 549:123–128. Guymon, R., Pomerantz, S. C., Crain, P. F., and McCloskey, J. A. (2006) Influence of phylogeny on posttranscriptional modification of rRNA in thermophilic prokaryotes: the complete modification map of 16S rRNA of Thermus thermophilus. Biochemistry 45:4888–4889. Hamacher, K., Trylska, J., and McCammon, J. A. (2006) Dependency map of proteins in the small ribosomal subunit. PLoS Comp. Biol. 2:80–87. Hanawa-Suetsugu, K., Sekine, S., Sakai, H., Hori-Takemoto, C., Terada, T., Unzai, S., Tame, J. R., Kuramitsu, S., Shirouzu, M., and Yokoyama, S. (2004) Crystal structure of elongation factor P from Thermus thermophilus HB8. Proc. Natl. Acad. Sci. USA 101:9595–9600. Hard, T., Rak, A., Allard, P., Kloo, L., and Garber, M. (2000) The solution structure of ribosomal protein L36 from Thermus thermophilus reveals a zinc-ribbon-like fold. J. Mol. Biol. 296:169–180. Hartmann, R. K. and Erdmann, V. A. (1989) Thermus thermophilus 16S rRNA is transcribed from an isolated transcription unit. J. Bacteriol. 171:2933–2941. Hartmann, R. K., Ulbrich, N., and Erdmann, V. A. (1987) An unusual rRNA operon constellation: in Thermus thermophilus HB8 the 23S/5S rRNA operon is a separate entity from the 16S rRNA operon. Biochimie 69:1097–1104. Hashimoto, Y., Yano, T., Kuramitsu, S., and Kagamiyama, H. (2001) Disruption of Thermus thermophilus genes by homologous recombination using a thermostable kanamycin-resistant marker. FEBS Lett. 506:231–234. Heinrich, T., Schroder, W., Erdmann, V. A., and Hartmann, R. K. (1992) Identification of the gene encoding transcription factor NusG of Thermus thermophilus. J. Bacteriol. 174:7859–7863. Helgstrand, M., Rak, A. V., Allard, P., Davydova, N., Garber, M. B., and Hard, T. (1999) Solution structure of the ribosomal protein S19 from Thermus thermophilus. J. Mol. Biol. 292:1071–1081. Henne, A., Bruggemann, H., Raasch, C., Wiezer, A., Hartsch, T., Liesegang, H., Johann, A., Lienard, T., Gohl, O., Martinez-Arias, R., Jacobi, C., Starkuviene, V., Schlenczeck, S., Dencker, S., Huber, R., Klenk, H. P., Kramer, W., Merkl, R., Gottschalk, G., and Fritz, H. J. (2004) The genome sequence of the extreme thermophile Thermus thermophilus. Nature Biotechnol. 22:547–553. Hensel, R., Demharter, W., Kandler, O., Kroppenstedt, R. M., and Stackebrandt, E. (1986) Chemotaxonomic and molecular-genetic studies of the genus Thermus: evidence for a phylogenetic relationship of Thermus aquaticus and Thermus ruber to the genus Deinococcus. Int. J. Systemat. Bacteriol. 36:444–453. Hirsch, P., Gallikowski, C. A., Siebert, J., Peissl, K., Kroppenstedt, R., Schumann, P., Stackebrandt, E., and Anderson, R. (2004) Deinococcus frigens sp. nov., Deinococcus saxicola sp. nov., and Deinococcus marmoris sp. nov., low temperature and draught-tolerating, UV-resistant bacteria from continental Antarctica. System. Appl. Microbiol. 27:636–645. Hoshino, T., Hidaka, Y. Y., Kosuge, T., Hasegawa, M., Saito, T., and Nakahara, T. (1994) Transformation as a tool for genetic fine mapping of closely located mutations in Thermus thermophilus. FEMS Microbiol. Lett. 117: 175–180. Ilag, L. L., Videler, H., McKay, A. R., Sobott, F., Fucini, P., Nierhaus, K. H., and Robinson, C. V. (2005) Heptameric (L12)6/L10 rather than canonical pentameric complexes are found by tandem MS of intact ribosomes from thermophilic bacteria. Proc. Natl. Acad. Sci. USA 102:8192–8197. Jenner, L., Romby, P., Rees, B., Schulze-Briese, C., Springer, M., Ehresmann, C., Ehresmann, B., Moras, D., Yusupova, G., and Yusupov, M. (2005) Translational operator of mRNA on the ribosome: how repressor proteins exclude ribosome binding. Science 308:120–123. Kaminishi, T., Wilson, D. N., Takemoto, C., Harms, J. M., Kawazoe, M., Schlüenzen, F., Hanawa-Suetsugu, K., Shirouzu, M., Fucini, P., and Yokoyama, S. (2007) A snapshot of the 30S ribosomal subunit capturing mRNA via the Shine-Dalgarno interaction. Structure 15:289–297.

9214_C018.indd 304

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Structure and Evolution of the Thermus thermophilus Ribosome

305

Kayser, K. J., Kwak, J. H., Park, H. S., and Kilbane, J. J. 2nd. (2001) Inducible and constitutive expression using new plasmid and integrative expression vectors for Thermus sp. Lett. Appl. Microbiol. 32:412–418. Khaitovich, P., Mankin, A. S., Green, R., Lancaster, L., and Noller, H. F. (1999) Characterization of functionally active subribosomal particles from Thermus aquaticus. Proc. Natl. Acad. Sci. USA 96:85–90. Khaitovich, P., Tenson, T., Kloss, P., and Mankin, A. S. (1999) Reconstitution of functionally active Thermus aquaticus large ribosomal subunits with in vitro-transcribed rRNA. Biochemistry 38:1780–1788. Kjeldgaard, M., Nissen, P., Thirup, S., and Nyborg, J. (1993) The crystal structure of elongation factor EF-Tu from Thermus aquaticus in the GTP conformation. Structure 1:35–50. Kobayashi, H., Kuwae, A., Maseda, H., Nakamura, A., and Hoshino, T. (2005) Isolation of a low-molecularweight, multicopy plasmid, pNHK101, from Thermus sp. TK10 and its use as an expression vector for T. thermophilus HB27. Plasmid 54:70–79. Korostelev, A., Trakhanov, S., Laurberg, M., and Noller, H. F. (2006) Crystal structure of a 70S ribosometRNA complex reveals functional interactions and rearrangements.Cell. 126:1065-1077. Koyama, Y., Hoshino, T., Tomizuka, N., and Furukawa, K. (1986) Genetic transformation of the extreme thermophile Thermus thermophilus and of other Thermus spp. J. Bacteriol. 166: 338–340. Krasilnikov, A. S., Xiao, Y., Pan, T., and Mondragon, A. (2004) Basis for structural diversity in homologous RNAs. Science 306:104–107. Kumagai, I., Watanabe, K., and Oshima, T. (1980) Thermally induced biosynthesis of 2′-O-methylguanosine in tRNA from an extreme thermophile, Thermus thermophilus HB27. Proc. Natl. Acad. Sci. USA 77:1922–1926. Leontiadou, F., Triantafillidou, D., and Choli-Papadapalou, T. (2001) On the characterization of the putative S20–Thx operon of Thermus thermophilus. Biol. Chem. 382:1001–1006. Liesack, W. and Stackebrandt, E. (1989) Evidence for unlinked rrn operons in the Planctomycete Pirellula marina. J. Bacteriol. 171:5025–5030. Lindahl, M., Svensson, L. A., Liljas, A., Sedelnikova, S. E., Eliseikina, I. A., Fomenkova, N. P., Nevskaya, N., Nikonov, S. V., Garber, M. B., Muranova, T. A., Rykonova, A. I., and Amons, R. (1994) Crystal structure of the ribosomal protein S6 from Thermus thermophilus. EMBO J. 13:1249–1254. Makyio, H., Iino, R., Ikeda, C., Imamura, H., Tamakoshi, M., Iwata, M., Stock, D., Bernal, R. A., Carpenter, E. P., Yoshida, M., Yokoyama, K., and Iwata, S. (2005) Structure of a central stalk subunit F of prokaryotic V-type ATPase/synthase from Thermus thermophilus. EMBO J. 24:3974–3983. Masui, R., Kurokawa, K., Nakagawa, N., Tokunaga, F., Koyama, Y., Shibata, T., Oshima, T., Yokoyama, S., Yasunaga, T., and Kuramitsu, S. NCBI. (2005) Thermus thermophilus HB8, complete genome. http://www.ncbi.nlm.nih.gov/genomes/framik.cgi?db=Genome&gi=530. McCutcheon, J. P., Agrawal, R. K., Philips, S. M., Grassucci, R. A., Gerchman, S. E., Clemons, W. M. Jr., Ramakrishnan, V., and Frank, J. (1999) Location of translational initiation factor IF3 on the small ribosomal subunit. Proc. Natl. Acad. Sci. USA 96:4301–4306. Mengel-Jørgensen, J., Jensen, S. S., Rasmussen, A., Poehlsgaard, J., Iversen, J. J., and Kirpekar, F. (2006) Modifications in Thermus thermophilus 23S ribosomal RNA are centred in regions of RNA–RNA contact. J. Biol. Chem. 281:22108–22117. Miroshnichenko, M. L., L’Haridon, S. L., Jeanthon, C., Antipov, A. N., Kostrikina, N. A., Tindall, B. J., Schumann, P., Spring, S., Stackebrandt, E., and Bonch-Osmolovskaya, E. A. (2003) Oceanithermus profundus gen. nov., sp. nov., a thermophilic, microaerophilic, facultatively chemolithotrophic bacterium from a deep-sea hydrothermal vent. Int. J. Syst. Evol. Microbiol. 53:747–752. Miroshnichenko, M. L., L’Haridon, S., Nercessian, O., Antipov, A. N., Kostrikina, N. A., Tindall, B. J., Schumann, P., Spring, S., Stackebrandt, E., Bonch-Osmolovskaya, E. A., and Jeanthon, C. (2003) Vulcanithermus mediatlanticus gen. nov., sp. nov., a novel member of the family Thermaceae from a deep-sea hot vent. Int. J. Syst. Evol. Microbiol. 53:1143–1148. Moreno, R., Zafra, O., Cava, F., and Berenguer, J. (2003) Development of a gene expression vector for Thermus thermophilus based on the promoter of the respiratory nitrate reductase. Plasmid 49:2–8. Munson, M. A., Baumann, L., and Baumann, P. (1993) Buchnera aphidicola (a prokaryotic endosymbiont of aphids) contains a putative 16S rRNA operon unlinked to the 23S rRNA-encoding gene: sequence determination, and promoter and terminator analysis. Gene 137:171–178. Murakami, K. S., Masuda, S., Campbell, E. A., Muzzin, O., and Darst, S. A. (2002) Structural basis of transcription initiation: an RNA polymerase holoenzyme-DNA complex. Science 296:1285–1290.

9214_C018.indd 305

11/5/2007 4:34:09 PM

306

Thermophiles: Biology and Technology at High Temperatures

Myasnikov, A. G., Marzi, S., Simonetti, A., Giuliodori, A. M., Gualerzi, C. O., Yusupova, G., Yusupov, M., and Klaholz, B. P. (2005) Conformational transition of initiation factor 2 from the GTP- to GDPbound state visualized on the ribosome. Nat. Struct. Mol. Biol. 12:1145–1149. Nevskaya, N., Tishchenko, S., Nikulin, A., al-Karadaghi, S., Liljas, A., Ehresmann, B., Ehresmann, C., Garber, M., and Nikonov, S. (1998) Crystal structure of ribosomal protein S8 from Thermus thermophilus reveals a high degree of structural conservation of a specific RNA binding site. J. Mol. Biol. 279:233–244. Nikonov, S., Nevskaya, N., Eliseikina, I., Fomenkova, N., Nikulin, A., Ossina, N., Garber, M., Jonsson, B. H., Briand, C., Al-Karadaghi, S., Svensson, A., Ævarsson, A., and Liljas, A. (1996) Crystal structure of the RNA binding ribosomal protein L1 from Thermus thermophilus. EMBO J. 15:1350–1359. Nikulin, A., Serganov, A., Ennifar, E., Tishchenko, S., Nevskaya, N., Shepard, W., Portier, C., Garber, M., Ehresmann, B., Ehresmann, C., Nikonov, S., and Dumas, P. (2000) Crystal structure of the S15-rRNA complex. Nat. Struct. Biol. 7:273–277. Nishimura, M., Yoshida, T., Shirouzu, M., Terada, T., Kuramitsu, S., Yokoyama, S., Ohkubo, T., and Kobayashi, Y. (2004) Solution structure of ribosomal protein L16 from Thermus thermophilus HB8. J. Mol. Biol. 344:1369–1383. Nissen, P., Kjeldgaard, M., Thirup, S., Polekhina, G., Reshetnikova, L., Clark, B. F., and Nyborg, J. (1995) Crystal structure of the ternary complex of Phe-tRNA Phe, EF-Tu, and a GTP analog. Science 270:1464–1472. Nobre, M. F., Truper, H. G., and da Costa, M.S. (1996) Transfer of Thermus ruber (Loginova et al. 1984), Thermus silvanus (Tenreiro et al. 1995), and Thermus chliarophilus (Tenreiro et al.1995) to Meiothermus gen. nov. as Meiothermus ruber comb. nov., Meiothermus silvanus comb. nov., and Meiothermus chliarophilus comb. nov., respectively, and emendation of the genus Thermus. Int. J. Syst. Bacteriol. 46:604–606. Noller, H. F. (2004) The driving force for molecular evolution of translation. RNA 10:1833–1837. Noller, H. F., Hoffarth, V., and Zimniak, L. (1992) Unusual resistance of peptidyl transferase to protein extraction procedures. Science 256:1416–1419. Nomura, M., Yates, J. L., Dean, D., and Post, L. E. (1980) Feedback regulation of ribosomal protein gene expression in Escherichia coli: structural homology of ribosomal RNA and ribosomal protein mRNA. Proc. Natl. Acad. Sci. USA 77:7084–7088. Oakes, M., Henderson, E., Scheinman, A., Clark, M., and Lake, J. A. (1986) Ribosome structure, function and evolution: mapping ribosomal RNA, proteins, and functional sites in three dimensions. In Structure, function, and genetics of ribosomes. B. Hardesty and G. Kramer, eds., Springer-Verlag, New York., pp. 47–67. Ogle, J. M. and Ramakrishnan, V. (2005) Structural insights into translational fidelity. Annu. Rev. Biochem. 74:129–177. Ogle, J. M., Murphy F. V., IV, Tarrry, M. J., and Ramakrishnan, V. (2002) Selection of tRNA by the ribosome requires a transition from an open to a closed form. Cell 111:721–732. Ohman, A., Rak, A., Dontsova, M., Garber, M. B., and Hard, T. (2003) NMR structure of the ribosomal protein L23 from Thermus thermophilus. J. Biomol. NMR 26:131–137. Ohno-Iwashita, Y., Oshima, T., and Imahori, K. (1975) In vitro protein synthesis at elevated temperature by an extract of an extreme thermophile: Effects of polyamines on the polyuridylic acid-directed reaction. Arch. Biochem. Biophys. 171:490–499. Omelchenko, M. V., Wolf, Y. I., Gaidamakova, E. K., Matrosova, V. Y., Vasilenko, A., Zhai, M., Daly, M. J., Koonin, E. V., and Makarova, K. S. (2005) Comparative genomics of Thermus thermophilus and Deinococcus radiodurans: divergent routes of adaptation to thermophily and radiation resistance. BioMed Central Evol. Biol. 5:57. Oshima, T. and Imahori, K. (1971) Isolation of an extreme thermophile and thermostability of its transfer ribonucleic acid and ribosomes. J. Gen. Appl. Microbiol. 17:513–517. Oshima, T. and Imahori, K. (1974) Description of Thermus thermophilus (Yoshida and Oshima), comb. nov., a nonsporulating thermophilic bacterium from a Japanese thermal spa. Int. J. Syst. Bacteriol. 24:102–112. Petry, S., Brodersen, D. E., Murphy, F. V. 4th, Dunham, C. M., Selmer, M., Tarry, M. J., Kelley, A. C., and Ramakrishnan, V. (2005) Crystal structures of the ribosome in complex with release factors RF1 and RF2 bound to a cognate stop codon. Cell 123:1255–1266. Pfeiffer, T., Jorcke, D., Feltens, R., and Hartmann, R. K. (1995) Direct linkage of str-, S10- and spc-related gene clusters in Thermus thermophilus HB8, and sequences of ribosomal proteins L4 and S10. Gene 167:141–145.

9214_C018.indd 306

11/5/2007 4:34:09 PM

Structure and Evolution of the Thermus thermophilus Ribosome

307

Pioletti, M., Schlünzen, F., Harms, J., Zarivach, R., Glühmann, M., Avila, H., Bashan, A., Bartels, H., Auerbach, T., Jacobi, C., Hartsch, T., Yonath, A., and Franceschi, F. (2001) Crystal structures of complexes of the small ribosomal subunit with tetracycline, edeine and IF3. EMBO J. 20:1829–1839. Ramírez-Arcos, S., Fernández-Herrero, L. A., Marín, I., and Berenguer, J. (1998) Anaerobic growth, a property horizontally transferred by an Hfr-like mechanism among extreme thermophiles. J. Bacteriol. 180:3137–3143. Ree, H. K. and Zimmermann, R. A. (1990) Organization and expression of the 16S, 23S and 5S ribosomal RNA genes from the archaebacterium Thermoplasma acidophilum. Nucleic Acids Res. 18:4471–4478. Sakaki, Y. and Oshima, T. (1975) Isolation and characterization of a bacteriophage infectious to an extreme thermophile, Thermus thermophilus HB8. J. Virol. 15:1449–1453. Sako, Y., Nakagawa, S., Takai, K., and Horikoshi, K. (2003) Marinithermus hydrothermalis gen. nov., sp. nov., a strictly aerobic, thermophilic bacterium from a deep-sea hydrothermal vent chimney. Int. J. Syst. Evol. Microbiol. 53:59–65. Schlüenzen, F., Tocilj, A., Zarivach, R., Harms, J., Gluehmann, M., Janell, D., Bashan, A., Bartels, H., Agmon, I., Franceschi, F., and Yonath, A. (2000) Structure of functionally activated small ribosomal subunit at 3.3 Å resolution. Cell 102:615–623. Schmidt, J., Srinivasa, B., Weglohner, W., and Subramanian, A. R. (1993) A small novel chloroplast ribosomal protein (S31) that has no apparent counterpart in the E. coli ribosome. Biochem. Mol. Biol. Int. 29:25–31. Schuwirth, B. S., Borovinskaya, M. A., Hau, C. W., Zhang, W., Vila-Sanjurjo, A., Holton, J. M., and Cate, J. H. D. (2005) Structures of the bacterial ribosome at 3.5 Å resolution. Science 310:827–34. Selmer, M., Dunham, C. M., Murphy, F. V. 4th, Weixlbaumer, A., Petry, S., Kelley, A. C., Weir, J. R., and Ramakrishnan, V. (2006) Structure of the 70S ribosome complexed with mRNA and tRNA. Science 313:1935–1942. Serganov, A., Polonskaia, A., Ehresmann, B., Ehresmann, C., and Patel, D. J. (2003) Ribosomal protein S15 represses its own translation via adaptation of an rRNA-like fold within its mRNA. EMBO J. 22:1898–908. Sharma, M. R., Barat, C., Wilson, D. N., Booth, T. M., Kawazoe, M., Hori-Takemoto, C., Shirouzu, M., Yokoyama, S., Fucini, P., and Agrawal, R. K. (2005) Interaction of Era with the 30S ribosomal subunit implications for 30S subunit assembly. Mol. Cell 8:319–329. Shigi, N., Sakaguchi, Y., Suzuki, T., and Watanabe, K. (2006) Identification of two tRNA thiolation genes required for cell growth at extremely high temperatures. J. Biol. Chem. 281:14296–14306. Shigi, N., Suzuki, T., Terada, T., Shirouzu, M., Yokoyama, S., and Watanabe, K. (2006) Temperaturedependent biosynthesis of 2-thioribothymidine of Thermus thermophilus tRNA. J. Biol. Chem. 281:2104–2113. Simitsopoulou, M., Avila, H., and Franceschi, F. (1999) Ribosomal gene disruption in the extreme thermophile Thermus thermophilus HB8. Generation of a mutant lacking ribosomal protein S17. Eur. J. Biochem. 266:524–532. Suh, M.-J., Hamburg, D.-M., Gregory, S. T., Dahlberg, A. E., and Limbach, P. A. (2005) Extending protein identifications to unsequenced bacterial strains using matrix-assisted laser desorption/ionization (MALDI) mass spectrometry. Proteomics 5:4818–4831. Takayama, G., Kosuge, T., Maseda, H., Nakamura, A., and Hoshino, T. (2004) Nucleotide sequence of the cryptic plasmid pTT8 from Thermus thermophilus HB8 and isolation and characterization of its highcopy-number mutant. Plasmid 51:227–237. Tamura, M. and Holbrook, S. R. (2002) Sequence and structural conservation in RNA ribose zippers. J. Mol. Biol. 320:455–474. Terui, Y., Ohnuma, M., Hiraga, K., Kawashima, E., and Oshima, T. (2005) Stabilization of nucleic acids by unusual polyamines produced by an extreme thermophile, Thermus thermophilus. Biochem. J. 388:427–433. Thompson, J. D., Higgins, D. G., and Gibson, T. J. (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. 22:4673–4680. Thompson, J. and Dahlberg, A. E. (2004) Testing the conservation of the translational machinery over evolution in diverse environments: assaying Thermus thermophilus ribosomes and initiation factors in a coupled transcription-translation system from Escherichia coli. Nucleic Acids Res. 32:5954–5961.

9214_C018.indd 307

11/5/2007 4:34:10 PM

308

Thermophiles: Biology and Technology at High Temperatures

Toyoda, T., Tin, O. F., Ito, K., Fujiwara, T., Kumasaka, T., Yamamoto, M., Garber, M. B., and Nakamura, Y. (2000) Crystal structure combined with genetic analysis of the Thermus thermophilus ribosome recycling factor shows that a flexible hinge may act as a functional switch. RNA 6:1432–1444. Trakhanov, S., Yusupov, M., Shirokov, V., Garber, M., Mitschler, A., Ruff, M., Thierry, J. C., and Moras, D. (1989) Preliminary X-ray investigation of 70 S ribosome crystals from Thermus thermophilus. J. Mol. Biol. 209:327–328. Trylska, J., McCammon, J. A., and Brooks, C. L. III, (2005) Exploring assembly energetics of the 30S ribosomal subunit using an implicit solvent approach. J. Am. Chem. Soc. 127:11125–11133. Tsiboli, P., Herfurth, E., and Choli, T. (1994) Purification and characterization of the 30S ribosomal proteins from the bacterium Thermus thermophilus. Eur. J. Biochem. 226:169–177. Tung, C.-S. and Sanbonmatsu, K. Y. (2004) Atomic model of the Thermus thermophilus 70S ribosome developed in silico. Biophys. J. 87:2714–2722. Unge, J., van den Berg, A., Al-Kharadaghi, S., Nikulin, A., Nikonov, S., Davydova, N., Nevskaya, N., Garber, M, and Liljas, A. (1998) The crystal structure of ribosomal protein L22 from Thermus thermophilus: insights into the mechanism of erythromycin resistance. Structure 6:1577–1586. Volkmann, N., Hottentrager, S., Hansen, H. A., Zayzsev-Bashan, A., Sharon, R., Berkovitch-Yellin, Z., Yonath, A., and Wittmann, H. G. (1990) Characterization and preliminary crystallographic studies on large ribosomal subunits from Thermus thermophilus. J. Mol. Biol. 216:239–241. Vornlocher, H. P., Kreutzer, R., and Sprinzl, M. (1997) Organization of the Thermus thermophilus nusA/infB operon and overexpression of the infB gene in Escherichia coli. Biochimie 79:195–203. Wang, H., Takemoto, C. H., Murayama, K., Sakai, H., Tatsuguchi, A., Terada, T., Shirouzu, M., Kuramitsu, S., and Yokoyama, S. (2004) Crystal structure of ribosomal protein L27 from Thermus thermophilus HB8. Protein Sci. 13:2806–2810. Wang, Y., Jiang, Y., Meyering-Voss, M., Sprinzl, M., and Sigler, P. B. (1997) Crystal structure of the EF-Tu.EFTs complex from Thermus thermophilus. Nat. Struct. Biol. 4:650–656. Williams, R. A., Smith, K. E., Welch, S. G., Micallef, J., and Sharp, R. J. (1995) DNA relatedness of Thermus strains, description of Thermus brockianus sp. nov., and proposal to reestablish Thermus thermophilus (Oshima and Imahori). Int. J. Syst. Bacteriol. 45:495–499. Williams, R. and R. Sharp. (1995) The taxonomy and identification of Thermus. In Thermus Species. R. Sharp and R. Williams, eds., Plenum Press, New York., pp. 1–42. Wimberly, B. T., Brodersen, D. E., Clemons W. M., Jr., Morgan-Warren, R. J., Carter, A. P., Vonrhein, C., Hartsch, T., and Ramakrishnan, V. (2000) Structure of the 30S ribosomal subunit. Nature 407:327–339. Wimberly, B. T., White, S. W., and Ramakrishnan, V. (1997) The structure of ribosomal protein S7 at 1.9 Å resolution reveals a beta-hairpin motif that binds double-stranded nucleic acids. Structure 5:1187–1198. Woese, C. R. (1987) Bacterial evolution. Microbiol. Rev. 51:221–271. Woestenenk, E. A., Gongadze, G. M., Shcherbakov, D. V., Rak, A. V., Garber, M. B., Hard, T., and Berglund, H. (2002) The solution structure of ribosomal protein L18 from Thermus thermophilus reveals a conserved RNA-binding fold. Biochem. J. 363(Pt 3):553–561. Yamaguchi, K. and Subramanian, A. R. (2003) Proteomic identification of all plastid-specific ribosomal proteins in higher plant chloroplast 30S ribosomal subunit. PSRP-2 (U1A-type domains), PSRP-3/(ycf65 homologue) and PSRP-4 (Thx homologue). Eur. J. Biochem. 270:190–205. Yokoyama, S., Hirota, H., Kigawa, T., Yabuki, T., Shirouzu, M., Terada, T., Ito, Y., Matsuo, Y., Kuroda, Y., Nishimura, Y., Kyogoku, Y., Miki, K., Masui, R., and Kuramitsu, S. (2000) Structural genomics projects in Japan. Nature Struct. Biol. Suppl. 943–945. Yonath, A., Glotz, C., Gewitz, H. S., Bartels, K. S., von Bohlen, K., Makowski, I., and Wittmann, H. G. (1988) Characterization of crystals of small ribosomal subunits. J. Mol. Biol. 203:831–834. Yu, M. X., Slater, M. R., and Ackermann, H. W. (2006) Isolation and characterization of Thermus bacteriophages. Arch. Virol. 151:663–679. Yusupov, M. M., Yusupova, G. Zh., Baucom, A., Lieberman, K., Earnest, T. N., Cate, J. H., and Noller, H. F. (2001) Crystal structure of the ribosome at 5.5 A resolution. Science 292:883–896. Yusupova, G. Zh., Yusupov, M. M., Cate, J. H. D., and Noller, H. F. (2001) The path of messenger RNA through the ribosome. Cell 106:233–241. Zeikus, J. G., Taylor, M. W., and Brock, T. D. (1970) Thermal stability of ribosomes and RNA from Thermus aquaticus. Biochim. Biophys. Acta 204:512–520.

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Protein Phosphorylation at 80°C and Above Peter J. Kennelly

CONTENTS Ubiquitous Regulatory Mechanism ........................................................................................... Chemistry of Protein Phosphorylation ...................................................................................... Phosphoacceptor Amino Acids ...................................................................................... Mechanisms of Phosphoprotein Modulation .................................................................. Thermodynamics of Protein Phosphorylation ................................................................ Phosphoproteins Can Be Dephosphorylated In Vivo ........................................... Influence of Thermodynamics on Cascade Architecture .................................... Distribution of Protein Kinase and Phosphatase Families among Hyperthermophiles ............ Histidine Kinases and Response Regulators .................................................................. Protein-Histidine and Protein-Aspartate Phosphatases .................................................. Protein-Serine/Threonine/Tyrosine Kinases .................................................................. “Eukaryote-Like” Protein Kinases ...................................................................... Other Protein-Serine/Threonine/Tyrosine Kinases ............................................. Protein-Serine/Threonine/Tyrosine Phosphatases ......................................................... Characteristics of Protein Kinases and Protein Phosphatases from Hyperthermpophiles ....... Histidine Kinases ............................................................................................................ HpkA from Thermatoga maritima ...................................................................... CheA from Thermatoga maritima ....................................................................... HK853 from Thermatoga maritima .................................................................... Eukaryote-Like Protein Kinases .................................................................................... Rio1 and Rio2 from Archaeoglobus fulgidus ...................................................... SsoPK2 and SsoPK3 from Sulfolobus solfataricus ............................................. Ph0512 from Pyrococcus horikoshii ................................................................... Protein Kinases of Undetermined Sequence .................................................................. Protein Phosphatases ...................................................................................................... Protein-Serine/Threonine Phosphatases .............................................................. Protein-Tyrosine Phosphatases ............................................................................ Phosphoproteins from Hyperthermophiles ................................................................................ Two-Component System Phosphoproteins ..................................................................... CheY from Thermotoga maritima ....................................................................... DrrA, DrrB, and DrrD from Thermatoga maritima ........................................... NtrC1 from Aquifex aeolicus ............................................................................... Phosphoserine- and Phosphothreonine-Containing Proteins ......................................... Initiation Factor 2α from Pyrococcus horikoshii ................................................ Cdc6 from Sulfolobus acidocaldarius .................................................................

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Phosphohexomutase from Sulfolobus solfataricus .............................................. SsoPK3 from Sulfolobus solfataricus .................................................................. Phosphotyrosine-Containing Proteins ............................................................................ Phosphoproteins of Undetermined Phosphoamino Acid Content .................................. Glycogen Synthase from Sulfolobus acidocaldarius .......................................... d-Gluconate Dehydratase from Sulfolobus solfataricus ..................................... Chaperonin-Associated Aminopeptidase from Sulfolobus solfataricus ............. Physiological Role of Protein Phosphorylation in Hyperthermophiles ..................................... Influence of Temperature ................................................................................................ Protein-Serine/Threonine/Tyrosine Phosphorylation .................................................... Two-Component System ................................................................................................. Conclusion .................................................................................................................................. Acknowledgments ...................................................................................................................... References ..................................................................................................................................

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The phosphorylation and dephosphorylation of proteins represents one of nature’s most prolific mechanisms for regulating cellular functions. Does protein-phosphorylation play a prominent role in the lives of hyperthermophilic organisms? What proteins are targeted for modification by phosphorylation in these extremophiles? The chapter below will try to summarize our current state of knowledge regarding the phosphoproteins, protein kinases, and protein phosphatases found in hyperthermophiles. This information will then be used to consider how extreme temperatures may have shaped the development of their protein phosphorylation cascades.

UBIQUITOUS REGULATORY MECHANISM The regulation of protein function is fundamental to the adaptation and survival of living organisms [1]. The functional plasticity of select proteins represents the key to directing and coordinating central metabolism; to sensing and reacting to environmental changes; to dynamically reconfiguring the proteome through control of gene expression as well as the degradation of proteins and the mRNAs that encode them; and to identifying the appropriate moment to commit to global, resourceintensive processes, such as cell division and cell differentiation. The two predominant mechanisms by which the functional properties of extant proteins are modulated to meet the moment-to-moment needs of the cell are (i) the binding of dissociable allosteric ligands and (ii) the covalent modification of protein structure. Most allosteric ligands can be grouped into one of a handful of general categories. Feedback metabolites regulate the expression and/or activity of enzymes critical to their synthesis. Indicator metabolites act pleiotropically to adjust metabolic and other functions in response to changes in global cellular parameters, such as energy state, nutrient availability, redox state, and so on. Second messengers, such as Ca2+ and cAMP are specialized allosteric ligands produced or released in response to the activation of a cell receptor. Unlike feedback or indicator metabolites, they participate in no other cellular processes. Similarly, the broad spectrum of covalent modifications that occur in nature can be grouped into a few functional categories. Post-translational processing events transform a nascent polypeptide into a mature, functionally competent protein. Common post-translational modifications include the removal, via partial proteolysis, of segments of the polypeptide chain that serve a transient purpose, such as a signal peptide; the addition of nonprotein structural moieties, such as oligosaccharides and lipid or glycolipid anchors; or the incorporation of prosthetic groups, such as metals or pyridoxal, that form part of its catalytic machinery. Post-translational processing events are characterized by their close temporal association with the synthesis of the affected polypeptide, their

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universality to all polypeptides of a given type, and their constancy over the lifetime of the protein. Regulatory modifications are characterized by their dynamic nature and their intimate association with specific intra- or extra-cellular stimuli. Unlike post-translational processing events, they vary in their timing, persistence, and/or extent. Among the numerous covalent modifications employed for the regulation of protein function, none can match phosphorylation in the number and diversity of its protein targets [2]. Open reading frames (ORF) encoding potential protein kinases and protein phosphatases have been identified in all but a minute handful of the hundreds of genomes sequenced to date [3–9]. Even more striking than its gross numerical dominance, however, is the remarkable versatility of protein phosphorylation. It exhibits no appreciable limits with regard to either the type of protein—enzymes, chromatin proteins, cytoskeletal proteins, motor proteins, transcription factors, translation factors, receptors, and so on—or functional property— specific activity, subcellular location, oligomerization, substrate selectivity, stability, susceptibility to other covalent modification events, and so on—that can be modulated by this mechanism. Its prolific nature can be attributed, in large part, to the special combination of biophysical properties inherent in the phosphoryl group.

CHEMISTRY OF PROTEIN PHOSPHORYLATION PHOSPHOACCEPTOR AMINO ACIDS The covalent modification of proteins by phosphorylation takes place on a variety of amino acid side chains (Figure 19.1). In eukaryotic organisms, phosphorylation predominantly targets the side chain hydroxyl groups of serine, threonine, and tyrosine—forming phosphoesters. In most Bacteria, as well as some members of the Archaea and Eucarya, phosphorylation also takes place on N-1 or, more frequently, N-3 of the imidazole side chain of histidine and on the β-carboxylate of aspartic acid. Curiously, the γ-carboxylate group of glutamic acid is rarely—if ever—modified by phosphorylation in vivo [10]. Although the existence of phospholysine- and phosphoarginine-containing proteins was reported several decades ago [11], little is known concerning either the identities of these proteins or their cognate protein-lysine or protein-arginine kinases and phosphatases [12,13]. The phosphothioester phosphocysteine is transiently formed as a catalytic intermediate in many protein-tyrosine phosphatases (PTP) and their derivatives [14], as well as enzyme II of the phosphoenolpyruvate:sugar phosphotransferase system [15]. However, no reports implicating phosphocysteine as either a structural or regulatory modification have appeared in the literature.

MECHANISMS OF PHOSPHOPROTEIN MODULATION The biophysical properties of the phosphoryl group render it a potent agent for perturbing the structural and functional characteristics of target proteins [16–18]. At intracellular pH, the phosphoryl group generally exhibits (i) a high-negative charge of −1.5 to −2.0, (ii) a large capacity for hydrogen bonding, and (iii) a marked propensity to form salt bridges with the protonated side chains of arginine [19,20] and lysine [21] residues. While protein phosphorylation’s stereotypic mode of action is the inducement of some general conformational change in target proteins (Figure 19.2a) [22–28], the addition of a phosphoryl group can also influence function by more spatially discrete mechanisms. The phosphorylation of phenylalanine hydroxylase enhances catalytic activity by shifting the position of the enzyme’s N-terminal tail to render the active site more accessible to substrates, but leaves the conformation of the core catalytic domain largely undisturbed (Figure 19.2b) [29]. In isocitrate dehydrogenase, the addition of a phosphoryl group to a serine residing at the mouth of the active site impedes catalysis by erecting an electrosteric barrier against the binding of isocitrate, a tricarboxylic acid (Figure 19.2c) [30–32]. When the serine residue adjacent to the catalytically essential histidine residue in 3-hydroxy-3-methylglutaryl coenzyme A reductase becomes phosphorylated, on the other hand, a new hydrogen bond forms between the protonated imidazole ring

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Thermophiles: Biology and Technology at High Temperatures OH P

HO

H2N

O

HO

P

CH2

CH H2N

COOH

C H

Phosphoserine (P-Ser)

Phosphothreonine (P-Thr)

CH2 H2N

HO P

O

Phosphotyrosine (P-Tyr)

O

CH2 COOH

H 3- Phosphohistidine (P-His)

FIGURE 19.1

O

C

CH2 C

COOH

O

OH

H2N

P

C H

OH OH N

O

CH3

H

N

P O

O

COOH

HO

O

O

C

OH

OH

H2N

C

COOH

H Phosphoaspartic acid (P-Asp)

Structures of the most frequently encountered phosphoamino acids.

and the anionic phosphoryl group (Figure 19.2d) [33]. As a consequence, the histidine residue is no longer able to protonate the thiolate moiety of coenzyme A, one of the reaction products— precipitously decreasing the enzyme’s catalytic efficiency. Phosphorylation serves as a trigger for the assembly of multiprotein complexes. One simple mechanism for accomplishing this is to utilize the phosphoryl group as an integral component of the protein–protein docking site itself (Figure 19.2e). The list of phospho-specific protein binding modules includes SH2 and PTB domains, which recognize phosphotyrosine-containing motifs [34], and WW, FHA, and 14-3-3 domains, which recognize phosphoserine/phosphothreonine-containing motifs [35–37]. Conversely, phosphorylation of some interaction sites may abrogate their protein binding properties, preventing the formation or facilitating the dissolution of a polypeptide complex [38–40]. In proteins that contain multiple sites of phosphorylation, interactions between sites can be employed to elicit more complex patterns of regulation (Figure 19.2f). For example, in glycogen synthase the phosphorylation of a quartet of clustered phosphorylation sites by glycogen synthase kinase-3 inhibits catalytic activity [41]. However, prior phosphorylation of a nearby site by casein kinase II is a pre-requisite for phosphorylation and inactivation by glycogen synthase kinase-3. Phosphorylation by casein kinase II in and of itself exerts no effect on activity. Its role is simply to enable phosphorylation by glycogen synthase kinase-3, a phenomenon called hierch(ic)al phosphorylation [42]. A reciprocal hierarchy exists in hormone sensitive lipase, where phosphorylation of Ser-565 by the AMP-activated protein kinase blocks the phosphorylation of Ser-563 and consequent activation of the enzyme by the cAMP-dependent protein kinase [43]. As the number of interactive

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FIGURE 19.2 Mechanisms by which phosphorylation alters function. Shown is a set of schematic representations illustrating the basic features of various mechanisms by which covalent phosphorylation influences the structural and functional properties of proteins. Proteins are represented by shaded ovals and polygons. Indentations represent enzyme active sites (a–c, f, g) and/or ligand binding sites (e, g). The filled diamond represents a dissociable allosteric ligand. Pi is used to indicate phosphoryl groups. Solid lines indicate covalent bonds, dotted lines indicate hydrogen bonds.

sites of phosphorylation increases, so does the potential to produce more sophisticated input–output algorithms. For example, cooperation among the thirteen sites of phosphorylation in transcription factor NFAT1 creates a sharp activation threshold that functions as an all-or-none trigger [44]. By contrast, the multiple phosphorylation sites in the transcriptional activator Ets-1 interact in an additive fashion to alter its DNA-binding affinity in a graduated manner [45]. Protein phosphorylation also can interact with covalent modification processes. In c-Myc, the glycosylation of Thr-58 is blocked by prior phosphorylation of this same residue (and vice-versa), the latter of which is, in turn, dependent upon the phosphorylation of Ser-62 [46]. Conversely, the

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bacterial pathogen Yersinia pestis prevents the activation of one of the host organisms key signal transduction proteins, mitogen-activated protein kinase, by acetylating key serine and threonine residues within its activation loop [47]. Acetylation preempts the phosphorylation of these residues by an upstream protein kinase. The principle of hierarchal regulation can also be extended to embrace the interplay between sites of phosphorylation and the binding of allosteric ligands. Only the phosphorylated form of cytochrome c oxidase, for example, is sensitive to allosteric inhibition by ATP [48], while phosphorylation of C4 phosphoenol pyruvate carboxylase decreases its responsiveness to glucose 6-phosphate by increasing Kact seven-fold and decreasing the Vmax of the activated enzyme by fivefold [49]. Phosphorylation of mammalian carbamoyl phosphate synthetase by a mitogen-activated protein kinase decreases the former’s sensitivity to feedback inhibition by UTP while concomitantly sensitizing it to feed forward activation by phosphoribosylpyrophosphate [50]. Hierarchal relationships between phosphorylation sites and between sites of protein phosphorylation and other covalent modifications and ligand binding, when coupled with the potential interplay between the protein kinase(s) and protein phosphatase(s) that target a given phosphorylation site, can transform proteins into sophisticated computer chips capable of processing a multiplicity of signal inputs [2].

THERMODYNAMICS OF PROTEIN PHOSPHORYLATION Phosphoproteins Can Be Dephosphorylated In Vivo Protein phosphorylation derives many of its capabilities from a combination of two seemingly antagonistic physicochemical properties—reactivity and stability [51]. The “high energy” of organophosphates renders them inherently chemically reactive, that is, thermodynamically unstable. They are thus capable of performing thermodynamic work, as reflected in the selection of ATP as the primary cellular energy carrier. However, the relative kinetic stability of most organophosphates at intracellular pH enables a cell to dictate when, where, and how this “latent” reactivity is harnessed via the selective intervention of enzyme catalysts. One potential fate for a protein-bound phosphoryl group is transfer to the weak, but highly abundant nucleophile, water—a process catalyzed by protein phosphatases. The resultant ability to controllably and repeatedly interconvert a protein between its phosphorylated and dephosphorylated forms permits an organism to repeatedly switch protein functions on and off “on demand.” Moreover, the “reversibility” of protein phosphorylation can employed to transform even the simplest phosphoprotein into a decision node for processing sensory information, as the protein kinase(s) and protein phosphatase(s) that act upon it often are controlled through distinct sensor-response cascades. Under these circumstances, the multiple signals propagated through these enzymes will converge upon and be resolved as the net stoichiometry and persistence of phosphorylation of the target phosphoprotein [2]. Influence of Thermodynamics on Cascade Architecture Differences in the physicochemical properties of the various phosphoamino acids influence the manner in which signal transduction cascades are constructed. The standard free energy of hydrolysis of the acid anhydride bond in phosphoaspartate (P-Asp) is comparable, −10.3 kcal/mole, to that of the phosphoramide bond in phosphohistidine (P-His) and somewhat exceeds that of ATP [52]. Hence, the energetic barrier for transfer of phosphate groups from histidine to aspartate, and viceversa, is relatively small. Many two-component signal cascades (see section “Histidine Kinases and Response Regulators”) exploit this property to construct multistep His-Asp phosphorelays whose length and complexity exceed that of the archetypical two-component cascade [53–56] and/or to terminate signal transmission via retrotransfer of phosphate when receptor stimulation ceases [57]. At ≈−3 kcal/mole, the phosphoesters of serine, threonine, and tyrosine are not nearly as energy-rich as P-Asp and P-His [52]. One consequence of their lower group transfer potential is that phosphoryl

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groups bound to hydroxyl amino acids rarely undergo intra- and inter- protein transfer in vivo. Thus, every phosphorylation event in a multi-step serine/threonine/tyrosine phosphorylation cascade consumes one molecule of ATP. By contrast, once it has been “primed” using a single molecule of ATP, a His-Asp phosphorelay can pass a single phosphoryl group all the way down the cascade via a series interdomain and interprotein transfers [53–55]. The apparent economic disadvantage of phosphoester-based cascades is likely more than offset by their greater inherent fidelity. As the capability and complexity of signal transduction cascades is increased via the incorporation of additional steps, branches, and hierarchally phosphorylated proteins; the opportunities for “short circuits” arising from anomalous phosphotransfer events rapidly multiply [58]. As phosphoesters appear to be considerably less prone to adventitious phosphotransfer, they offer a clear advantage over phosphoramides and mixed acid anhydrides for the construction of architecturally complex signal transduction networks.

DISTRIBUTION OF PROTEIN KINASE AND PHOSPHATASE FAMILIES AMONG HYPERTHERMOPHILES HISTIDINE KINASES AND RESPONSE REGULATORS Histidine kinase and response regulator domains constitute the core building blocks of a conserved signal transmission module known as the two-component system [56,59,60]. Histidine kinases phosphorylate themselves on a conserved histidine residue using ATP as phosphodonor substrate. The autophosphorylated histidine kinase then serves as a specialized, regulated phosphodonor substrate for the intrinsic phosphotransferase activity of its partner response regulator domain(s) [61]. The response regulator autophosphorylates on a conserved aspartic acid residue. In the basic twocomponent systems, autophosphorylation of the response regulator domain triggers a change in its conformation [62] that influences the functional properties of an associated domain or protein, such as a flagellar motor protein, a methylesterase domain, or a DNA-binding domain. However, some response regulator domains also catalyze the transfer of the phosphoryl group from the conserved aspartic acid residue to exogenous proteins, such as histidine kinases (retrotransfer) or, in multi-step variants known as His-Asp phosphorelays, to a histidine residue on a third conserved module called an Hpt domain [55]. Although theoretically possible, no examples of transfer to a serine, threonine, or tyrosine have emerged. The genomes of the three hyperthermophilic or near hyperthermophilic, that is, Thermoanaerobacter tengcongensis (T optimum 75°C [63]), bacteria listed in Table 19.1 each encode known or potential histidine kinases and response regulators, as do 4 of the 14 archaeal hyperthermophiles. The striking disparity in the proportion of bacterial versus archaeal hyperthermophiles that harbor two-component systems is attributable, in large measure, to the evolutionary history of this signal transduction paradigm. The two-component system originated and has flourished in the Bacteria [5,6]. ORFs encoding deduced histidine kinases and response regulators have been identified in 85% to 90% of all bacterial genomes [1,64], where they account for ≈1% of coded proteins [65]. The Archaea and Eucarya acquired the two-component system via horizontal gene transfer [5,6]. Consequently, both the pervasiveness and multiplicity of two-component ORFs is considerably lower among the members of the archaeal and eucaryal domains [3,66,67].

PROTEIN-HISTIDINE AND PROTEIN-ASPARTATE PHOSPHATASES Dephosphorylation of phosphorylated response regulator can take place via several mechanisms: (i) hydrolysis by an autophosphatase activity intrinsic to the response regulator protein, (ii) retrotransfer to a cognate histidine kinase, or (iii) intervention of an exogenous protein-aspartate phosphatase [56]. Of the three, the most common appears to be autodephosphorylation. In some instances, this process is regulated by a protein cofactor such as CheZ. Binding of CheZ to CheY completes the active site

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TABLE 19.1 Distribution of ORFs Encoding Known or Potential Protein Kinases or Protein Phosphatases in Hyperthermophilic Organisms T°C

HK

SixA

RR

CheC/FliY

ePK

PPP

PPM

cPTP

LMW PTP

Archaea Aeropyrum pernix

95

−

−

−

−

+

+

−

−

−

Archaeoglobus fulgidus

83

+

−

+

+

+

+

−

−

+

Ignicoccus

90

−

−

−

−

+

+

−

−

−

Methanococcus jannaschii

85

−

−

−

−

+

−

−

+

−

Methanopyrus kandleri

98

−

−

−

−

+

+

−

−

−

Nanoarchaeum equitans

90

−

−

−

−

−

−

−

−

−

Pyrobaculum aerophilum

100

−

+

−

−

+

+

−

+

−

96

+

−

+

+

+

+

−

+

−

100

−

−

−

−

+

−

−

+

−

Pyrococcus horikoshii

96

+

−

+

+

+

−

−

+

−

Sulfolobus acidocaldarius

80

−

−

−

−

+

+

−

+

−

Sulfolobus solfataricus

80

−

+

−

−

+

+

−

+

−

Sulfolobus tokodaii

80

−

+

−

−

+

+

−

+

−

102

+

−

+

+

+

+

−

+

−

93

+

−

+

+

+

−

+

−

−

75

+

−

+

+

+

−

+

−

+

90

+

−

+

+

+

+

−

−

−

ORGANISM

Pyrococcus abyssi Pyrococcus furiosus

Thermococcus kodakaraensis Bacteria Aquifex aeolicus Thermoanaerobacter tengcongensis Thermotoga maritima

Note: Plus and minus signs are used to indicate whether one or more ORFs encoding a protein that displays similarity to the members of the protein family listed at the head of each column is (+) or is not (−) present in the genome of the organism listed. Abbreviations: CheC/FliY, CheC/FliY protein-aspartate phosphatase; cPTP, conventional proteintyrosine phosphatase; ePK, eukaryote-like protein-serine/threonine/tyrosine kinase; HK, histidine kinase; LMWPTP, low molecular weight protein-tyrosine phosphatase; PPM, PPM family protein-serine/threonine phosphatase; PPP, PPP-family protein-serine/threonine phosphatase; RR, response regulator; SixA, SixA phosphohistidine phosphatase; T, approximate optimum growth temperature.

of the latter by providing a glutamine side chain that facilitates hydrolysis by orientating the water molecule that attacks and displaces the phosphoryl group from the phosphoaspartate residue [68]. No homologs of CheZ have been identified in the hyperthermophiles listed in Table 19.1, however. When phosphate is removed via retrotransfer to a histidine kinase, completion of the hydrolytic process can take place by one of several mechanisms [69]. Some histidine kinases are bifunctional, that is, they are capable of catalyzing their own dephosphorylation [56,70]. Alternatively, the histidine kinase might be dephosphorylated through the action of an exogenous protein-histidine phosphatase [70]. Lastly, if the two-component cascade is divided into branches, the histidine kinase can transfer the phosphoryl group to a second response regulator protein. The alternate response regulator may possess high autophosphatase activity or be susceptible to attack by an exogenous protein-aspartate phosphatase [71–73]. Two families of potential protein-aspartate phosphatases have been identified thus far: the response regulator aspartyl-phosphate (RapP) phosphatases [74] and CheC/FliY [69]. Among the

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hyperthermophiles, only T. tencogenesis encodes a homolog of the Rap phosphatases. By contrast, homologs of CheC/FliY were present in every hyperthermophile whose genome contained ORFs encoding deduced histidine kinases and response regulators (Table 19.1). Significantly, no homologs of Chec/FliY were evident in those hyperthermophiles lacking potential two-component systems. By contrast, the prospective protein-histidine phosphatase SixA [75] was present in only three hyperthermophiles—the archaeons P. aerophilum, S. solfataricus, and S. tokadaii (Table 19.1)— none of which encoded potential histidine kinases.

PROTEIN-SERINE/THREONINE/TYROSINE KINASES “Eukaryote-Like” Protein Kinases The term “eukaryote-like” protein kinase, or ePK, refers to nature’s most prolific, and indeed overwhelmingly dominant, family of protein-serine/threonine/tyrosine kinases [8,76] for which the catalytic domain of the cAMP-dependent protein kinase serves as prototype [77]. Prototypical ePKs are characterized by a set of twelve conserved sequence motifs, subdomains I-V, Via, VIb, and VIIXI that are distributed among two functional lobes: an N-terminal, β sheet-rich nucleotide binding domain and a C-terminal, helix-rich, protein substrate binding domain [75]. Subdomain VIb, which lies at the interface between these domains, contains several residues that participate directly in catalysis. In addition to the prototypic or conventional ePKs, several subfamilies have been discovered that deviate from the established ePK paradigm, among them the RIO kinases and the piD261 protein kinases [7,79,80]. These “atypical” ePKs generally are characterized by the absence of the basic amino acid residue in subdomain VIb that is characteristic of conventional ePKs [81] and a compressed or truncated C-terminal protein binding lobe that may lack obvious equivalents for any or all of subdomains IX-XI [80,82]. Ironically, the conventional ePKs appear to be descended from one of the atypical ePKs [7,83]. Prospective members of the extended ePK superfamily were identified in the genome of nearly every hyperthermophile examined (Table 19.1). The sole exception, Nanoarchaeum equitans, is an outlier in terms of both its obligatorily parasitic lifestyle and minute genome. It may be noteworthy that the genome of this nanoarchaeon’s host, the archaeon Ignicoccus [84], does encode deduced ePKs. While conventional ePKs predominate in bacterial organisms [7,9], including the hyperthermophilic ones, atypical protein kinases dominate among the Archaea [1]. To date, only a small handful of archaeons, among them the hyperthermophiles of the genus Sulfolobus, harbor deduced proteins resembling conventional ePKs [1]. These same archaeons also contain atypical ePKs. It remains to be determined whether the conventional ePKs in the Archaea evolved independently of their counterparts in the Eucarya, or were acquired by horizontal gene transfer [7,9]. Other Protein-Serine/Threonine/Tyrosine Kinases Despite the overwhelming dominance of ePK superfamily, other sources of protein-serine/ threonine/tyrosine kinase activity exist. These include the alpha kinases of the Eucarya [85]; AceK, the isocitrate dehydrogenase kinase/phosphatase found in enteric bacteria [86]; the HPr kinase/ phosphatase that (de)phosphorylates the histidine rich protein of the sugar phosphotransferase system in Gram-positive bacteria and proteobacteria [87,88]; and several serine/threonine/tyrosinespecific derivatives of two component histidine kinases such as the Rsb and Spo kinases and mitochondrial protein kinases [6]. Among the hyperthermophiles listed in Table 19.1, only T. tencongensis possesses potential alternative protein-serine/threonine kinases, specifically a prospective HPr kinase/phosphatase as well as homologs of the Rsb kinases of Bacillus subtilis [89].

PROTEIN-SERINE/THREONINE/TYROSINE PHOSPHATASES In sharp contrast to the dominance of the extended ePK family as a source of protein-serine/ threonine/tyrosine kinase activity, nature employs four major families of protein phosphatases (and

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at least three minor ones) to dephosphorylate protein-bound phosphoserine, phosphothreonine, and phosphotyrosine [14,90]. The four major families are the PPP and PPM protein-serine/threonine phosphatases [91], the conventional protein-tyrosine phosphatases (cPTP) [92], and the low molecular weight protein-tyrosine phosphatases (LMW PTP) [93]. The PPP and cPTP families include some variants that are dual-specific, that is, they hydrolyze both the phosphoester of the aromatic alcohol tyrosine, as well as phosphoesters of the aliphatic alcohols serine and threonine. Unlike the Eucarya, wherein the four families are ubiquitous [93–96], the genomes of prokaryotic organisms are populated by heterogeneous and seemingly random combinations of these protein phosphatase paradigms [4,90]. Among the hyperthermophilic prokaryotes, some clear trends are evident, however. Discounting the parasite/symbiont N. equitans, which lacks any recognizable sources of protein kinase or protein phosphatase activity, 10 of 13 archaeal genomes—77%—encode known or potential PPPs. A nearly equivalent number, nine—69%—possess deduced cPTPs (Table 19.1). Intriguingly, every one of the 13 possessed a PPP and/or a cPTP. No potential PPMs were identified among the archaeal hyperthermophiles. Potential LMW cPTPs were nearly as scarce, being present in only one or perhaps two archaeal hyperthermophiles (Table 19.1). Although the sample size was small, the situation with respect to the hyperthermophilic Bacteria is roughly the inverse of that in the Archaea. Two of three, that is, A. aeolicus and T. tencongensis, encode both potential PPM family protein phosphatases and potential LMW PTPs. T. maritima, on the other hand, contained only a deduced contains PPP while none harbored ORFs for the possible LMW cPTPs that were so prevalent among the archaeal hyperthermophiles.

CHARACTERISTICS OF PROTEIN KINASES AND PROTEIN PHOSPHATASES FROM HYPERTHERMPOPHILES HISTIDINE KINASES HpkA from Thermatoga maritima The gene encoding HpkA was cloned by PCR using degenerate primers modeled after conserved sequences in previously identified histidine kinases [97]. The first 20 amino acids of the 412residue polypeptide are predicted to form a transmembrane segment, implying that this histidine kinase partners with a membrane receptor protein. Recombinant versions of the HpkA autophosphorylate when incubated with ATP. Phosphorylated HpkA serves as a phosphodonor substrate in vitro for the endogenous response regulator protein DrrA, whose coding gene is located immediately upstream of that for HpkA [97,98]. The pH stability profile of autophosphorylated HpkA was consistent with modification of a histidine residue [99]. CheA from Thermatoga maritima The CheA histidine kinase from T. maritima has been the object of extensive structural studies [100–105]. Its domain composition and topography resembles that of the histidine kinases that transmit signals from chemo-, photo-, and aero-tactic receptor proteins to the flagellar motor proteins that control swimming behavior. Recombinantly produced versions of CheA from T. maritima form homodimers in solution [103]. CheA binds a homolog of CheW, a small adapter protein that likely couples it to the receptor(s) that modulate the former’s histidine kinase activity [106]. X-ray crystallographic analysis revealed that histidine phosphotransferase domain of CheA alternates between two distinct conformations, suggesting that this thermophilic protein transduces signals— at least in part—via a conformational switching mechanism [104]. Intact CheA from T. maritima catalyzes its own phosphorylation, while a truncated fragment consisting of the ATP-binding and catalytic subdomains exhibits phosphotransferase activity toward exogenous P1 domains that contain the predicted phosphoacceptor histidine residue [105].

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HK853 from Thermatoga maritima The amino acid sequence of HK853, the protein product of ORF TM0853 in the genome of T. maritima, exhibits the domain organization typical of a receptor kinase, that is, an extracellular/ periplasmic domain linked to an intracellular catalytic domain via a transmembrane helix. HK583’s closest sequence homologs are the EnvZ and PhoQ histidine kinases that modulate gene expression in response to environmental factors, such as osmolarity and phosphate concentration [107]. The ligand for the presumed extracellular receptor domain of HK853 remains to be identified, although this region exhibits weak homology to the GAF domains implicated in cGMP or photopigment binding [3]. Recombinant HK853 is a dimer that catalyzes its ATP-dependent autophosphorylation. Intriguingly, HK853 exhibits functional competency in a biological context. Expression of the T. maritima protein in an E. coli strain lacking EnvZ restored expression of a reporter gene whose expression is modulated by the latter’s cognate response regulator, OmpR [107].

EUKARYOTE-LIKE PROTEIN KINASES Rio1 and Rio2 from Archaeoglobus fulgidus The genome of the hyperthermophilic archaeon A. fulgidus codes for two members of the RIO family of atypical ePKs, Rio1 and Rio2 [108,109]. Recombinantly expressed forms of both proteins exhibit protein-serine kinase activity toward exogenous substrates, such as histone H1 and myelin basic protein. In addition, both Rio1 and Rio2 autophosphorylate on a conserved serine residue located within a flexible loop that protrudes from the amino-terminal nucleotide-binding lobe [109,110]. In contrast to many conventional ePKs, autophosphorylation does not appear to be essential for activation of these atypical ePKs, as a form of Rio1 in which the phosphoacceptor serine residue had been replaced by alanine phosphorylated exogenous protein substrates at rates comparable to wild type [109]. It must be noted, however, that these assays were—because of the mammalian origins of the substrate proteins utilized—performed at a temperature well below that which Rio1 normally operates, that is, 37°C versus 80°C+. While the physiologic substrates of Rio1 and Rio2 have yet to be determined, it has been reported that their yeast homologs are essential for ribosome biosynthesis and cell cycle progression [111,112]. Intriguingly, the amino terminus of Rio2 (but not Rio1) from A. fulgidus contains a winged-helix motif characteristic of many DNA-binding proteins [112]. SsoPK2 and SsoPK3 from Sulfolobus solfataricus SsoPK2 is an atypical ePK encoded by ORF sso2387 in the genome of S. solfataricus [113]. The catalytic domain of the protein resides in the C-terminal portion of the 583-residue polypeptide. The function of the amino terminal portion of the protein is unknown. The N-terminal region is devoid of readily identifiable sequence motifs. Despite the fact that the catalytic domain is significantly truncated relative to those of conventional ePKs, essentially terminating after putative subdomain VIII, the recombinant protein catalyzes both its own phosphorylation as well as that of several exogenous proteins at low, but measurable rates [113]. However, it remains to be determined whether proteins represent the physiological target of the enzyme’s phosphotransferase activity or whether indeed SsoPK2 functions as a phosphotransferase in vivo, as it has recently been annotated as an ATPase implicated in protein secretory events [114]. SsoPK2 utilizes ATP as phosphodonor substrate, and exclusively targeted serine residues in vitro [113]. The sites of autophosphorylation in SsoPK2 were mapped to distinct two regions of the protein via mass peptide profiling. One set resides in the central region beyond the estimated amino terminal boundary of the presumed catalytic domain. The second set occupies the region between putative subdomains VII and VIII within the catalytic domain [113]. This area roughly corresponds to the activation loop found in conventional ePKs. In the majority of the latter, phosphorylation of this loop, either autocatalytically or through the intervention of an exogenous protein kinase, is required

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to attain full catalytic competency [115]. Intriguingly, autophosphorylation only occurred at temperatures >65°C, indicating that the protein undergoes a temperature-dependent conformational transition between 37°C and 65°C [113]. Expression of SsoPK2 could be detected only in cultures that were grown on rich media containing sucrose, tryptone, and yeast extract [114]. SsoPK3 was first identified as a phosphothreonine-containing protein that became radiolabeled when partially-purified detergent extracts of the membrane fraction of S. solfataricus were incubated with 32P-lablled purine nucleotides (see section “SsoPK3 from Sulfolobus solfataricus”). The protein product of ORF sso0469, SsoPK3, is an atypical ePK. The 582-residue polypeptide contains plausible candidates for all of the subdomains, that is, I–XI, characteristic of the extended ePK family [116]. Certain of these assignments were verified via mutagenic alteration of residues predicted to be essential for catalysis, which produced enzyme forms displaying little or no catalytic activity. The first 180 residues of the protein share no obvious homology with other proteins or motifs. The extreme C-terminus, on the other hand, contains a putative leucine zipper, a motif commonly involved in protein–protein interactions. SsoPK3 was specific for ATP as phosphodonor substrate, but exhibited no propensity to autophosphorylate. Phosphotransfer to exogenous proteins, such as casein, myelin basic protein, and bovine serum albumin targeted serine and, less frequently, threonine residues [116]. Ph0512 from Pyrococcus horikoshii The protein product of ORF Ph0512 was one of two identified from the genome of P. horikoshii as potential homologs of an RNA-dependent ePK from the Eucarya known as PKR [117]. Recombinantly expressed Ph0512 phosphorylated aIF2α, the archaeal counterpart of the physiological substrate of PKR in Eucarya (see section “Initiation factor 2α from Pyrococcus horikoshii”). Both PH0512 and PKR phosphorylated aIF2α predominantly on Ser-48 [117].

PROTEIN KINASES OF UNDETERMINED SEQUENCE Detergent extracts of the membrane fraction of S. solfataricus contain a protein kinase activity that is, unusual in several respects. First, the polypeptide source of this activity, SsoPK1, can be renatured and its catalytic capabilities assayed following polyacrylamide electrophoresis in the presence of sodium dodecyl sulphate [118]. Second, it displayed a marked preference for modifying threonine residues. While some protein-serine kinase activity could be detected, the Vmax for phosphorylation of a threonine-containing peptide was 20-fold lower than that of an identical peptide in which the threonine had been replaced by serine [119]. Third, while ATP was the most efficient phosphodonor substrate, the enzyme also could utilize other purine nucleotides, for example, GTP, ADP, or GDP— in order of decreasing preference, as phosphodonor substrate in vitro [118,119]. SsoPK1 catalyzed the phosphorylation of several exogenous, physiologically irrelevant, protein, and peptide substrates in vitro, such as histone H4 and casein [119]. By taking advantage of its unusually broad phosphodonor specificity, a second membrane-associated protein kinase, SsoPK3, was identified as a potential endogenous phosphoprotein substrate for SsoPK1 (see section “SsoPK3 from Sulfolobus solfataricus”). SsoPK1 is subject to two types of covalent modifications, autophosphorylation, and glycosylation [120]. As modification by glycosylation is associated with proteins or domains that reside on the outer surface of the plasma membrane or beyond, it appears likely that SsoPK1 adopts a receptor-like transmembrane topology in vivo. The sequence of this protein kinase has yet to be determined.

PROTEIN PHOSPHATASES Protein-Serine/Threonine Phosphatases PP1-Arch1 from S. solfataricus [121,122] and Py-PP1 from P. abyssi [123] are members of the PPPfamily of protein phosphatases. The gross functional characteristics of this pair of archaeal PPPs

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were quite comparable. Both consist of a single core catalytic domain that possesses no obvious regulatory, membrane binding, or other auxiliary domains [122,123]. Thus, it is unclear how or even whether their activity is controlled in vivo. Both archaeal PPPs appear to be serine/threonine specific, like their eukaryotic homologs [121,123]. However, unlike their mammalian counterparts, all of which are metalloenzymes, the affinity of these hyperthemophile-derived protein phosphatases for divalent metal ion cofactors is relatively weak [90]. Hence, catalysis requires the presence of exogenous metal ions, of which Mn+2 appears to be the most efficacious. Protein-Tyrosine Phosphatases Tk-PTP from T. kodakaraensis is a member of the conventional family of PTPs. A recombinant version of the protein hydrolyzed both free phosphotyrosine and free phosphoserine in vitro, and did so at comparable rates [124]. However, it was not determined whether Tk-PTP acts upon phosphoprotein substrates, although a substrate-trapping version of the protein bound three phosphotyrosine-containing proteins when used as an affinity purification agent (see section “Phosphotyrosine-Containing Proteins”). While it is common for PTPs to hydrolyze free phosphotyrosine, it is extremely rare that any protein phosphatase—even those with protein-serine phosphatase activity—will dephosphorylate free phosphoserine at a measurable rate [125]. Further evidence that the catalytic capabilities of this thermophilic cPTP differ from those of its mesophilic equivalents was provided by the analysis of mutagenically altered versions of the enzyme [124]. The conserved amino acid residues that play crucial roles in the catalytic mechanism of well-characterized cPTPs include (i) the active site nucleophile—a cysteine, (ii) a catalytic acid/base—an aspartate, and (iii) a substrate recognition and transition state stabilization group—an arginine—separated by five residues from the active site cysteine, for example, CX5R [126]. Mutagenic alteration of either the predicted active site nucleophile, Cys-93, or the conserved arginine, Arg-99, in the active site loop reduced catalytic efficiency 20-fold or more [124]. However, substitution of the predicted catalytic acid/base, Asp66, by alanine actually improved the catalytic performance of Tk-PTP slightly at 80°C. This behavior was not peculiar to hyperthermophilic conditions under which the enzyme normally operates, as the relative kcat/K m value of the mutagenically altered phosphatase remained comparable to that of the wild-type when kinetic measurements were performed at 25°C [124]. It would therefore appear that the catalytic mechanism of Tk-PTP has diverged somewhat from its better-known counterparts.

PHOSPHOPROTEINS FROM HYPERTHERMOPHILES TWO-COMPONENT SYSTEM PHOSPHOPROTEINS CheY from Thermotoga maritima CheY refers to a family of two-component response regulator proteins that bind to and modulate the action of flagellar motor proteins following autophosphorylation on a conserved aspartic acid residue [57,127]. Autophosphorylation is regulated by controlling the availability of CheY’s phosphodonor substrate, the autophosphorylated CheA histidine kinase, via the latter’s association with receptors responsive to chemotactic, phototactic, or aerotactic signals. A gene encoding a homolog of CheY from the hyperthermophilic bacterium T. maritima has been cloned and its protein product expressed [128]. The recombinant protein catalyzed its own phosphorylation in the absence of a cognate histidine kinase using acetyl-phosphate as phosphodonor substrate. It has been postulated that the ability of certain response regulators to utilize acetyl phosphate as an alternate phosphodonor may represent mechanism for responding to changes in metabolic status via an internal “acetate switch” [129]. Based upon an analysis of the X-ray crystal structure of CheY from T. maritima, it has been suggested that phosphorylation of the conserved aspartate residue, Asp-54, might affect protein function through the formation of a hydrogen bond with Ser-82 [130]. The resulting rotation

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of the side chain of Ser-82 about the Cα-Cβ bond would shield Phe-101 from solvent. The resulting changes in the surface characteristics of CheY presumably unmask a latent site for binding flagellar motor proteins. DrrA, DrrB, and DrrD from Thermatoga maritima DrrA, DrrB, and DrrD from T. maritima are members of the OmpR/PhoB subfamily of response regulators [131]. Each Drr protein contains two major structural domains: an N-terminal response regulator domain and a C-terminal winged-helix domain that is predicted to bind DNA. Wellcharacterized members of the OmpR/PhoB subfamily modulate the transcription of specific genes in response to environmental signals. The thermostability of response regulators from T. maritima has rendered them attractive subjects for x-ray crystallography, Hence, three-dimensional structures for the dephosphorylated forms of both DrrB and DrrD have been determined [131,132]. DrrA can be phosphorylated in vitro by the two-component histidine kinase HpkA, the product of the ORF lying directly adjacent to that encoding DrrA [97]. As has been observed for several other response regulators, DrrA catalyzes its autodephosphorylation in vitro [99]. This hydrolytic activity requires a divalent metal ion cofactor such as Mg+2. Phosphorylation of DrrB modulates its quaternary structure in vitro. Recombinantly produced, unphosphorylated DrrB is a monomer in solution. However, incubation with phosphoramidate, an artificial phosphodonor substrate for intrinsic autophosphorylation activity of many response regulators, caused the protein to dimerize [132]. As dimerization appears to be a prerequisite for DNA binding by the members of the OmpR/ PhoB family of response regulators [133], phosphorylation of DrrB may be necessary, and perhaps sufficient, to trigger its association with DNA in vivo. NtrC1 from Aquifex aeolicus The annotation of the gene product of ORF aq1117 from A. aeolicus as NtrC1 reflects its high degree of sequence similarity, 60%, with NtrC from E. coli [134]. In E. coli, NtrC regulates gene transcription by activating the σ54 RNA polymerase [135]. NtrC1 is organized into three major structural domains: a C-terminal DNA-binding domain, a central AAA+ ATPase domain, and an N-terminal response regulator domain [134]. Dephosphorylated NtrC1, a homodimer, exhibits little or no propensity to stimulate transcription of a σ54-dependent reporter gene in E. coli. However, constructs lacking the response regulator domain were active in these same assays, implying that interactions between the response regulator domains impose some constraint upon the other domains that is relieved by phosphorylation of the former [136]. Structural studies have confi rmed that phosphorylation of the response regulator domains in NtrC1 induces a reconfiguration of its interdomain contacts, as predicted by this model [134,136].

PHOSPHOSERINE- AND PHOSPHOTHREONINE-CONTAINING PROTEINS Initiation Factor 2α from Pyrococcus horikoshii The archaeal translational initiation factor 2 complex (aIF2) exhibits a high degree of structural [137] and functional [138] similarity to its eucrayal counterpart, eIF2. In the Eucarya, phosphorylation of a conserved serine residue in the α-subunit of this complex inhibits protein synthesis. Intriguingly, incubation of aIF2α from Pyrococcus horikoshii with one of the protein kinases known to phosphorylate eIF2α in mammals, an RNA-dependent protein kinase known as PKR, resulted in the phosphorylation of the aIF2α on Ser-48 [117]. Moreover, Ser-48 falls within one residue of aligning directly with the phosphorylated serine in eIF2α. A serine is also found at this position in the homologs of aIF2α encoded by other archaeons, including the hyperthermophiles. Since a protein kinase endogenous to P. horikoshii, Ph0512, phosphorylates aIF2α on Ser-48 in vitro (see section “Ph0512 from Pyrococcus horikoshii”), it is probable that archaeal hyperthermophiles

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employ protein phosphorylation to regulate protein translation in a similar manner to members of the Eucarya. Cdc6 from Sulfolobus acidocaldarius Given that the fundamental elements of cellular information transfer processes, such as transcription and translation, are fairly well conserved between the Archaea and Eucarya, it was somewhat surprising that few obvious homologs of eucaryal cell division cycle (Cdc) proteins could be identified among archaeal genomes [139]. One notable exception was Cdc6 [140]. In the Eucarya, Cdc6 plays an essential role in the initiation of DNA replication. Members of the Cdc6 family bind ATP, presumably to provide the energy to drive their activity. Incubation of Cdc6 from the hyperthermophilic archaeaon P. aerophilum with [γ-32P]ATP resulted in the formation of phosphoserine [141]. A Cdc6 homolog from S. solfataricus behaved in a similar fashion, although the nature of the phosphoryl-protein bond was not determined [142]. Phosphorylation of both proteins was self-catalyzed, as it required the presence of an intact Walker A nucleotide binding motif [141,142]. The physiological role of Cdc6 autophosphorylation in hyperthermophilic archaeons remains to be determined. Intriguingly, autophosphorylation of Cdc6 from P. aerophilum could be inhibited by both single- and double-stranded DNA [141]. While autophosphorylation of Cdc6 from S. solfataricus was unaffected by the presence of DNA [142], a homolog from the moderately thermophilic archaeon Methanobacterium thermoautotrophicum also was inhibited by DNA [143]. In addition, the latter protein was stimulated by association with the minichromosome maintenance complex [143]. The behavior of these DNA-sensitive versions of Cdc6 suggests that autophosphorylation is a regulated, and hence functionally consequential, event; and not simply an innocuous side reaction, as is apparently the case for the autophosphorylation of bacterial nucleoside diphosphate kinases [144]. Phosphohexomutase from Sulfolobus solfataricus Phosphohexomutases catalyze the net transfer of a phosphate group between hydroxyl groups on their phosphosugar substrates. Catalysis proceeds via formation of a phosphoenzyme intermediate involving a serine residue within the active site [145]. Since sequence comparisons implicated Ser-59 as the catalytically essential serine residue in the deduced phosphohexomutase from S. solfataricus, it was somewhat surprising when mass peptide profiling of protein extracts from this archaeon indicated that Ser-309 was phosphorylated in vivo [146]. When Ser-59 was found to be essential for catalysis while Ser-309 was not, the x-ray structure of a bacterial homolog was used as a scaffold to extrapolate the position of the latter. The model indicated that Ser-309 lies in the vicinity of the active site [146], suggesting that a phosphoryl group located at this point would act as an electrosteric barrier against the binding of phosphosugar substrates in a manner analogous to the phosphorylation of isocitrate dehydrogenase (see section “Mechanisms of Phosphoprotein Modulation”). This inference was supported by mutagenic alterations of Ser-309. Substitution of Ser-309 with a potentially negatively charged aspartic acid residue resulted in an enzyme whose Vmax was only 4% that of wild-type, while neutral residues, such as Ala and Gln only marginally reduced catalytic efficiency [146]. SsoPK3 from Sulfolobus solfataricus Membrane extracts of S. solfataricus contain a protein kinase, SsoPK1, that displays some unique characteristics in vitro, for example, a marked preference for modifying threonine over serine residues and the ability to utilize purine nucleotide tri- and di-phosphates as phosphodonor substrates (see section “Protein Kinases of Undetermined Sequence”). When detergent extracts enriched for SsoPK1 were incubated with radiolabeled nucleotides, a second protein-serine kinase SsoPK3 was phosphorylated [116]. Several observations indicated that SsoPK3 was phosphorylated by SsoPK1

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and not via an autocatalytic process [116]. First, recombinantly expressed SsoPK3 displayed no propensity to autophosphorylate in vitro. Second, phosphorylation took place on threonine, which is the preferred target for SsoPK1, while SsoPK3 appears to prefer serine residues. Third, whereas recombinant SsoPK3 was ATP-specific, phosphorylation could be detected in extracts incubated with [γ-32P]GTP or [β-32P]GDP, which can be used as phosphodonor substrate by SsoPK1 in vitro. While the physiological function, if any, of this phosphorylation event remains to be determined, the phosphorylation of one protein kinase by another is an extremely common feature of the multistep signal transduction cascades in the Eucarya [147].

PHOSPHOTYROSINE-CONTAINING PROTEINS Historically, the occurrence and role of protein-tyrosine phosphorylation in “lower” organisms has proven controversial [148]. However, in the past several years, a compelling body of evidence has accumulated indicating that many prokaryotic organisms harbor protein-tyrosine kinases, PTPs, and phosphotyrosine-containing proteins [149]. Thus far, the only hyperthermophile in which protein-tyrosine phosphorylation has been investigated in any depth is the archaeon Thermococcus kodakaraensis [124]. Three phosphotyrosine-containing proteins were isolated from extracts of T. kodakaraensis using a specially engineered form of the endogenous PTP, Tk-PTP (see section “Protein-Tyrosine Phosphatases”), referred to as a “substrate-trapping mutant.” This approach capitalizes on the observation that many conventional PTPs retain the ability to bind cognate phosphoprotein substrates with high affinity after certain catalytically essential amino acid residues are altered via site-directed mutagenesis [150]. When extracts from T. kodakaraensis were fractionated on a column of inactive Tk-PTP, three proteins adhered thereto that also displayed immunoreactivity toward antibodies against phosphotyrosine. Amino terminal sequence analysis revealed them to be (i) a putative phosphomannomutase, (ii) the deduced β-subunit of phenylalanine t-RNA synthetase, and (iii) the unidentified protein product of an ORF within the RNA terminal phosphate cyclase operon [124]. With regard to the first of these, it is perhaps noteworthy that the phosphohexosemutase from S. solfataricus is a phosphoprotein, albeit a phosphoserine-containing one (see section “Phosphohexomutase from Sulfolobus solfataricus”). Curiously, it was not reported whether Tk-PTP catalyzed the dephosphorylation of these proteins in vitro.

PHOSPHOPROTEINS OF UNDETERMINED PHOSPHOAMINO ACID CONTENT Glycogen Synthase from Sulfolobus acidocaldarius Analysis of the polypeptides associated with glycogen particles isolated from the archaeon S. acidocaldarius revealed the presence of an M r ≈ 60 kDa polypeptide whose sequence resembled that of prototypic glycogen synthases [151]. Analysis of a recombinant version of the protein indicated that it did indeed possess glycosyl transferase activity. Following isolation from cultures of S. acidocaldarius that had been grown in the presence of 32P-labeled orthophosphate, three distinct forms of the polypeptide were resolved by two-dimensional electrophoresis that differed in pI, but not Mr. The two more acidic of these were labeled with 32P, as would be expected if they represented differentially phosphorylated forms of a single polypeptide [151]. The radiolabel remained associated with the protein when the latter was incubated at pH 2.5, indicating that the phosphoryl group was likely bound through an ester linkage. D-Gluconate

Dehydratase from Sulfolobus solfataricus

d-Gluconate dehydratase catalyzes the second step of the modified, semi-phosphorylative Entner-Doudooroff pathway in hyperthermophilic Archaea [152]. When isolated from cultures of S. solfataricus that had been incubated in the presence of 32P-orthophosphate, d-gluconate

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dehydratase was radiolabeled [153]. While the chemical nature of the protein-phosphoryl linkage was not determined, d-gluconate dehydratase lost activity when treated with either of two broad-spectrum phosphatases, potato acid phosphatase or bacterial alkaline phosphatase [153], implying that the enzyme was activated by phosphorylation. Chaperonin-Associated Aminopeptidase from Sulfolobus solfataricus Incubation of cell lysates of S. solfataricus with [γ-32P]ATP resulted in the radiolabeling of a protein with an Mr of ≈ 60 kDa that coimmunoprecipitated with the chaperonin complex [154]. Sequence analysis and assays of enzyme activity indicated that the phosphoprotein was an aminopeptidase. Phosphorylation does not appear to modulate the catalytic efficiency of the enzyme as neither preincubation of cell lysates with ATP nor with an unidentified phosphatase influenced the level of aminopeptidase activity detected in immunoprecipitates [154]. The nature of the phosphoacceptor amino acid was not determined.

PHYSIOLOGICAL ROLE OF PROTEIN PHOSPHORYLATION IN HYPERTHERMOPHILES INFLUENCE OF TEMPERATURE The conspicuous electrostatic properties of the phosphoryl group render it an effective tool for manipulating protein structure and function at elevated temperatures [155,156]. It therefore is not surprising that, with the exception of the parasitic symbiont N. equitans, all of the hyperthermophiles listed in Table 19.1 encode known or potential protein kinases and protein phosphatases within their genomes. Moreover, even N. equitans benefits from the contributions of protein kinases and protein phosphatases by virtue of its intimate, almost organelle-like, relationship with its obligate host, Ignicoccus. The distribution of ePKs, histidine kinases, and response regulators amongst the hyperthermophiles suggests that elevated temperatures influence the nature of cellular protein phosphorylation events. Representatives of both the ePK family of protein-serine/threonine/tyrosine kinases and the histidine kinases and response regulators of the two-component system are found among the hyperthermophiles. However, while every organism surveyed, with the previously noted exception of N. equitans, contains one or more ORFs encoding potential ePKs, less than half harbored ORFs for potential two-component histidine kinases and response regulators. Moreover, the total number of two-component modules in a given hyperthermophile (see section “Two-Component System”) tended to fall significantly below the 20 to 50 or more harbored by a typical free-living prokaryote [3,64,66]. The apparent preference for protein-serine/threonine/tyrosine phosphorylation among the hyperthermophiles may simply reflect the large proportion of archaeal organisms within this group. The ePK family of protein kinases, which target side chain hydroxyl groups, is indigenous to the members of the domain [7], whereas two-component histidine kinases and response regulators were imported from the Bacteria, where they originated [5,6]. However, when one compares the stabilities of phosphoesters, phosphoramides, and acid anhydrides at elevated temperatures, it is difficult to discount the impact of thermophily on the origins and distribution of various protein phosphorylation paradigms. Even within the Archaea, a rough correlation is evident between growth temperature and the presence and multiplicity of deduced two-component cascades [66]. Of 13 archaeons containing histidine kinases and response regulators, only four are hyperthermophiles and only one of them, A. fulgidus, possesses sufficient components to construct multiple two-component cascades. Eight of the remaining nine nonhyperthemophiles, on the other hand, encode a variety of histidine kinases and response regulators. By contrast, hyperthermophiles account for 10 of the 15 archaeons that lack two-component histidine kinases and response regulators.

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As temperatures increase from 25–37°C to 80°C and beyond, the half-lives of phosphoramides and mixed acid anhydrides in neutral, aqueous solution are reduced from a few hours to a few minutes or less [157–163]. While phosphoesters suffer a commensurate relative decline in stability as temperatures increase, they still persist for relatively long periods—hours to days—as compared to the time frames on which most molecular processes take place in the cell [157,164]. The transience of P-His and P-Asp at hyperthermophilic temperatures render their application to the regulation of many cellular processes problematic. While it is true that the lifetime of a given phosphoamino acid can be influenced by the protein of which it is a component [99], (de)stabilization must be purchased at the expense of structural constraints that must narrow the spectrum of molecular processes amenable to this mode of regulation.

PROTEIN-SERINE/THREONINE/TYROSINE PHOSPHORYLATION A clear dichotomy is evident in our body of knowledge regarding protein phosphorylation in hyperthermophiles. Studies on two-component mediated signal transduction have been confined almost exclusively to bacterial organisms. The archaeal hyperthermophiles enjoy a similar monopoly as venues for the study of protein-serine/threonine/tyrosine phosphorylation. In hyperthermophilic archaeons, evidence accumulated to date implicates at least three cellular processes as targets for regulation via the phosphorylation of serine, threonine, and/or tyrosine residues. The first is carbohydrate metabolism. Phosphoproteins have been identified that participate in glycolysis (d-gluconate dehydratase, see section “d-Gluconate Dehydratase from Sulfolobus solfataricus”), carbohydrate storage (glycogen synthase, see section “Glycogen Synthase from Sulfolobus acidocaldarius”), and oligosaccharide synthesis (phosphohexomutase, see section “Phosphohexomutase from Sulfolobus solfataricus” and “Phosphotyrosine-Containing Proteins). While the nature of the phosphoamino acids in d-gluconate dehydratase and glycogen synthase have yet to be determined, given that the organism from which they were isolated, S. solfataricus, possesses only ePKs (Table 19.1), they presumably are modified on one of the hydroxyl amino acids. The second is DNA replication, as evidenced by the reports that Cdc6 is phosphorylated in three archaeons, including two hyperthermophiles (see section “Cdc6 from Sulfolobus acidocaldarius”). Third is protein synthesis. The parallels between the sites phosphorylated in eIF2α and aIF2α strongly suggest that phosphorylation of the latter is of regulatory significance (see section “Initiation Factor 2α from Pyrococcus horikoshii”). The identification of the deduced β-subunit of phenylalanine t-RNA synthetase as a tyrosine-phosphorylated protein in T. kodakaraensis [124] also is consistent with a role for phosphorylation in translation.

TWO-COMPONENT SYSTEM In the seven hyperthermophiles that encode potential two-component signal transduction cascades, the number of such modules exhibits a strikingly bimodal distribution. A. fulgidus, T. maritima, and T. tencongensis each possess 19 or more total histidine kinase and response regulator domains, while A. aoelicus, P. abyssi, P. horikoshii, and T. kodakaraensis contain three to eight [3,64,66]. A common leitmotif among these organisms was chemotaxis. Six of this set of seven hyperthemophiles possesses ORFs encoding one or more CheA-like histidine kinases and CheY-like response regulators—the exception being the bacterium A. aeolicus [64]. The conservation of the Che system in hyperthermophiles may be attributable, at least in part, to the extremely rapid and dynamic nature of chemo-, aero-, and photo- tactic sensor-response processes. In A. aeolicus, the domain architectures of its deduced response regulator proteins suggest that the two-component cascade is employed exclusively to regulate gene expression [64]. Similarly, many of the “nonchemotactic” response regulators in T. tencongensis and T. maritima are fused with DNA binding domains. In A. fulgidus, on the other hand, all of the predicted response regulators are CheY- or CheB-like [66], suggesting that this organism has developed a particularly extensive sensor-response network to guide its motions.

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CONCLUSION At fi rst glance, it would appear that hyperthermophiles employ protein phosphorylationdephosphorylation in ways similar to those of their mesophilic counterparts. However, as life under hyperthermophilic conditions magnifies the physicochemical strengths and limitations of the various molecular mechanisms, much remains to be learned from these organisms regarding the chemistry and evolution of this important molecular regulatory mechanism.

ACKNOWLEDGMENTS The author gratefully acknowledges the support of NSF grant MCB-0315122.

REFERENCES 1. Kennelly, P.J., Archaeal protein kinases and protein phosphatases—Insights from genomics and biochemistry, Biochem. J., 370, 373, 2003. 2. Wurgler-Murphy, S.M., King, D.M. and Kennelly, P.J., The Phosphorylation Site Database: A guide to the serine-, threonine-, and/or tyrosine-phosphorylated proteins in prokaryotic organisms, Proteomics, 4, 1562, 2004. 3. Galperin, M.Y., A census of membrane-bound and intracellular signal transduction proteins in Bacteria: Bacterial IQ, extroverts and introverts. BMC Microbiol., 5, 35, 2005. 4. Kennelly, P.J., Protein kinases and protein phosphatases in prokaryotes: A genomic perspective, FEMS Microbiol. Lett., 206, 1, 2002. 5. Kim, D.-J. and Forst, S., Genomic analysis of the histidine kinase superfamily in bacteria and archaea, Microbiology, 147, 1197, 2001. 6. Koretke, K.K. et al., Evolution of the two-component regulatory system, Mol. Biol. Evol., 17, 1956, 2000. 7. Leonard, C.J., Aravind, L. and Koonin, E.V., Novel families of putative protein kinases in Bacteria and Archaea: Evolution of the ‘eukaryotic’ protein kinase superfamily, Genome Res., 8, 1038, 1998. 8. Manning, G. et al., Evolution of protein kinase signaling from yeast to man, Trends Biochem. Sci., 27, 514, 2002. 9. Ponting, C.P. et al., Eukaryotic signaling domain homologues in Archaea and Bacteria. Ancient ancestry and horizontal gene transfer. J. Mol. Biol., 289, 729, 1999. 10. Trumbore, M.W. et al., Prothymosin α in vivo contains phosphorylated glutamic acid residues, J. Biol. Chem., 272, 26394, 1997. 11. Walinder, O., Zetterquist, O. and Engstrom, L., Purification of a bovine liver protein rapidly phosphorylated by adenosine triphosphate. Isolation of 1-32P-phosphohistidine, 3-32P-phosphohistidine, and N-ε32P-phospholysine from 32P-labeled protein, J. Biol. Chem., 243, 2793, 1968. 12. Chen, C.-C. et al., Occurrence and distribution of acid-labile histone phosphates in regenerating rat liver, Biochemistry, 13, 3785, 1974. 13. Wakim, B.T. and Aswad, G.D., Ca2+ -calmdolulin-dependent phosphorylation of arginine in histone 3 by a nuclear protein kinase from mouse leukemia cells, J. Biol. Chem., 269, 2722, 1994. 14. Jackson, M.D. and Denu, J.M., Molecular reactions of protein phosphatases—Insights from structure and chemistry, Chem. Rev., 101, 2313, 2001. 15. Legler, P.M. et al., Three-dimensional solution structure of the cytoplasmic B domain of the mannitol transporter IIMannitol of the Escherichia coli phosphootransferase system, J. Biol. Chem., 279, 39115, 2004. 16. Appleby, J.L. and Bourret R.B., Activation of CheY mutant D57N by phosphorylation at an alternative site, Ser-56, Mol. Microbiol., 34, 915, 1999. 17. Signarvic, R.S. and DeGrado, W.F., De novo design of a molecular switch: Phosphorylation-dependent association of designed peptides, J. Mol. Biol., 334, 1, 2003. 18. Waud, J.P. et al., Engineering the C-terminus of firefly luciferase as an indicator of covalent modification of proteins, Biochim. Biophys. Acta, 1292, 89, 1996. 19. Johnson, L.N. and Barford, D., Electrostatic effects in the control of glycogen phosphorylase by phosphorylation, Prot. Sci., 3, 1726, 1994. 20. Johnson, L.N. and Lewis, R.J., Structural basis for control by phosphorylation, Chem. Rev., 101, 2209, 2001.

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328

Thermophiles: Biology and Technology at High Temperatures

21. Errington, N. and Doig, A.J., A phosphoserine-lysine salt bridge within an α-helical peptide, the strongest α-Helix side-chain interaction measured to date, Biochemistry, 44, 7553, 2005. 22. Fernando, P., Megeney, L.A. and Heikkila, J.J., Phosphorylation-dependent structural alterations in the small hsp30 chaperone are associated with cellular recovery, Exp. Cell Res., 286, 175, 2003. 23. Ganguly, C. et al., Limited proteolysis reveals a structural difference in the globular head domains of dephosphorylated and phosphorylated Acanthamoeba myosin II, J. Biol. Chem., 267, 20905, 1992. 24. Johnson, L.N. and Barford, D., The effects of phosphorylation on the structure and function of proteins, Annu. Rev. Biophys. Biomol. Struct., 22, 1999, 1993. 25. Johnson, L.N. and Lewis, R.J., Structural basis for control by phosphorylation, Chem. Rev., 101, 2209, 2001. 26. Metcalfe, E.E., Traaseth, N.J. and Veglia, G., Serine 16 phosphorylation induces an order-to-disorder transition in monomeric phospholamban, Biochemistry, 44, 4386, 2005. 27. Wittekind, M. et al., Common structural changes accompany the functional inactivation of HPr by seryl phosphorylation or by serine to aspartate subsitution, Biochemistry, 28, 9908, 1989. 28. Zhang, J. and Corden, J.L., Phosphorylation causes a conformational change in the carboxylterminal domain of the mouse RNA polymerase II largest subunit, J. Biol. Chem., 266, 2297, 1991. 29. Miranda, F.F. et al., Phosphorylation and mutations of Ser16 in human phenylalanine hydroxylase, J. Biol. Chem., 277, 40937, 2002. 30. Dean, A.M. and Koshland, D.E. Jr., Electrostatic and steric contributions to regulation at the active site of isocitrate dehydrogenase, Science, 249, 1044, 1990. 31. Hurley, J.H. et al., Regulation of an enzyme by phosphorylation at the active site, Science, 249, 1012, 1990. 32. Hurley, J.H. et al., Regulation of isocitrate dehydrogenase by phosphorylation involves no long-range conformational change in the free enzyme, J. Biol. Chem., 265, 3602, 1990. 33. Omkumar, R.V. and Rodwell, V.W., Phosphorylation of Ser871 impairs the function of His865 of Syrian hamster 3-hydroxy-3-methylglutaryl-CoA reductase, J. Biol. Chem., 269, 16862, 1994. 34. Pawson, T. and Nash, P. Assembly of cell regulatory systems through protein interaction domains, Science, 300, 445, 2003. 35. Dodge, K.L. et al., mAKAP assembles a protein kinase A/PDE4 phosphodiesterase cAMP signaling module, EMBO J., 20, 1921, 2001. 36. Yaffe, M.B. and Elia, A.E., Phosphoserine/thereonine-binding domains, Cell Biol., 13, 131, 2001. 37. Zor, T. et al., Roles of phosphorylation and helix propensity in the binding of the KIX domain of CREBbinding protein by constitutive (c-Myb) and inducible (CREB) activators, J. Biol. Chem., 277, 42241, 2002. 38. Daujat, S. et al., HP1 binds specifically to Lys26 -methylated histone H1.4, whereas simultaneous Ser27 phosphorylation blocks HP1 binding, J. Biol. Chem., 280, 38090, 2005. 39. Heist, E.K., Srinivasan, M. and Schulman, H., Phosphorylation at the nuclear localization signal of Ca2+/ calmodulin-dependent protein kinase II blocks its nuclear targeting, J. Biol. Chem., 273, 19763, 1998. 40. Hutchins, J.R.A., et al., Phosphorylation regulates the dynamic interaction of RCC1 with chromosomes during mitosis, Curr. Biol., 14, 1099, 2004. 41. Wang, Y. and Roach, P.J., Inactivation of rabbit muscle glycogen synthase by glycogen synthase kinase3, J. Biol. Chem., 268, 23876, 1993. 42. Roach, P.J., Multisite and hierarchal protein phosphorylation, J. Biol. Chem., 266, 14139, 1991. 43. Yeaman, S.J., Hormone-sensitive lipase—A multipurpose enzyme in lipid metabolism, Biochim. Biophys. Acta, 1052, 128, 1990. 44. Salazar, C. and Höfer, T., Allosteric regulation of the transcription factor NFAT1 by multiple phosphorylation sites: A mathematical analysis, J. Mol. Biol., 327, 31, 2003. 45. Pufall, M.A. et al., Variable control of Ets-1 DNA binding by multiple phosphates in an unstructured region, Science, 309, 142, 2005. 46. Kamemura, K. et al., Dynamic interplay between O-glycosylation and O-phosphorylation of nucleocytoplasmic proteins: Alternative glycosylation/phoosphorylation of THR-58, a known mutational hot spot of c-Myc in lymphomas, is regulated by mitogens, J. Biol. Chem., 277, 19229, 2002. 47. Mukherjee, S. et al., Yersinia YopJ acetylates and inhibits kinase activation by blocking phosphorylation, Science, 312, 1211, 2006. 48. Bender, E. and Kadenbach, B., The allosteric ATP-inhibition of cytochrome c oxidase activity is reversibly switched on by cAMP-dependent phosphorylation, FEBS Lett., 466, 130, 2000.

9214_C019.indd 328

11/5/2007 9:08:52 PM

Protein Phosphorylation at 80°C and Above

329

49. Duff, S.M.G. et al., Kinetic analysis of the non-phosphorylated, in vitro phosphorylated, and phosphorylation-site-mutant (Asp8) forms of intact recombinant C4 phosphoenolpyruvate carboxylase from sorghum, Eur. J. Biochem., 228, 92, 1995. 50. Graves, L.M. et al., Regulation of carbamoyl phosphate synthetase by MAP kinase, Nature, 403, 328, 2000. 51. Westheimer, F.H., Why nature chose phosphates, Science, 235, 1173, 1987. 52. Stock, J.B., Stock, A.M. and Mottonen, J.M., Signal transduction in bacteria, Nature, 344, 395, 1990. 53. Appleby, J.L., Parkinson, J.S. and Bourret, R.B., Signal transduction via the multi-step phosphorelay: Not necessarily a road less traveled, Cell, 86, 845, 1996. 54. Hoch, J.A., Two-component and phosphorelay signal transduction, Curr. Opin. Microbiol., 3, 165, 2000. 55. Perraud, A.L., Weiss, V. and Gross, R., Signaling pathways in two-component phosphorelay systems. Trends Microbiol., 7, 115, 1999. 56. Stock, A.M., Robinson, V.L. and Goudreau, P.N., Two-component signal transduction, Annu. Rev. Biochem., 69, 183, 2000. 57. Szurmant, H. and Ordal, G.W., Diversity of chemotaxis mechanisms among the Bacteria and Archaea, Microbiol. Mol. Biol. Rev., 68, 301, 2004. 58. Porter, S.L. and Armitage, J.P., Chemotaxis in Rhodobacter sphaeroides requires an atypical histidine protein kinase, J. Biol. Chem., 279, 54573, 2004. 59. Bourret, R.B. and Stock, A.M., Molecular information processing: Lessons from bacterial chemotaxis, J. Biol. Chem., 277, 9625, 2002. 60. Fabret, C., Feher, V.A. and Hoch, J.A., Two-component signal transduction in Bacillus subtilis: How one organism sees its world, J. Bacteriol., 181, 1975, 1999. 61. Grebe, T.W. and Stock, J.B., The histidine protein kinase superfamily, Adv. Microbial Physiol., 41, 139, 1999. 62. Stock, J. and De Re, S., Signal transduction: Response regulators on and off, Curr. Biol., 10, R420, 2000. 63. Xue, Y. et al., Thermoanaerobacter tengcongensis sp. Nov., a novel anaerobic, thermophilic bacterium isolated from a hot spring in Tencong, China, Int. J. Syst. Evol. Microbiol., 51, 1335, 2001. 64. Ashby, M.K., Survey of the number of two-component response regulator genes in the complete and annotated genome sequences of prokaryotes, FEMS Microbiol. Lett., 231, 277, 2004. 65. West, A.H. and Stock, A.M., Histidine kinases and response regulators in two-component signaling systems, Trends Biochem. Sci., 26, 369, 2001. 66. Ashby, M.K., Distribution, structure, and diversity of “bacterial” genes encoding two-component proteins in the Euryarchaeota, Archaea, 2, 11, 2006. 67. Chang, C. and Stewart, R. C., The two-component system, Plant Physiol., 117, 723, 1998. 68. Zhao, R. et al., Structure and catalytic mechanism of E. coli chemotaxis phosphatase CheZ, Nat. Struct. Biol., 9, 570, 2002. 69. Szurmant, H., Muff, T.J. and Ordal, G.W., Bacillus subtilis CheC and FliY are members of a novel clas of CheY-P-hydrolyzing enzymes in the chemotactic signal transduction cascade. J. Biol. Chem., 279, 21787, 2004. 70. Klumppp, S. and Krieglstein, J., Phosphorylation and dephosphorylation of histidine residues in proteins, Eur. J. Biochem., 269, 1067, 2002. 71. Jimenez-Pearson, M.-A. et al., Phosphate flow in the chemotactic response regulator system of Helicobacter pylori, Microbiology, 151, 3299, 2005. 72. Porter, S.L. and Armitage, J.P., Phosphotransfer in Rhodobacter sphaeroides chemotaxis, J. Mol. Biol., 324, 35, 2002. 73. Sourjic, V. and Schmitt, R., Phosphotransfer between CheA, CheY1, and CheY2 in the chemotaxis signal transduction chain of Rhizobium meliloti, Biochemistry, 37, 2327, 1998. 74. Reizer, J. et al., Characterization of a family of bacterial response regulator aspartyl phosphate (RAP) phosphatases, Microb. Comp. Genomics, 2, 103, 1997. 75. Matsubara, M. and Mizuno, T., The SixA phospho-histidine phosphatase modulates the ArcB phosphorelay signal transduction in Escherichia coli, FEBS Lett., 470, 118, 2000. 76. Manning, G., et al., The protein kinase complement of the human kinase, Science, 298, 1912, 2002. 77. Taylor, S.S. et al., A template for the protein kinase family, Trends Biochem. Sci., 18, 84, 1993. 78. Hanks, S.K. and Hunter, T., The eukaryotic protein kinase superfamily: Kinase (catalytic) domain structure and classification, FASEB J., 9, 576, 1995. 79. Angermmayr, M. and Bandlow, W., RIO1, an extraordinary novel protein kinase, FEBS Lett., 524, 31, 2002.

9214_C019.indd 329

11/5/2007 9:08:52 PM

330

Thermophiles: Biology and Technology at High Temperatures

80. Facchin, S. et al., Structure-function analysis of yeast piD261/Bud32, an atypical protein kinase essential for normal cell life, Biochem. J., 364, 457, 2002. 81. Hanks, S.K. and Lindberg, R.A., Use of degenerate oligonucleotide probes to identify clones that encode protein kinases, Meth. Enzymol, 2000, 525, 1991. 82. LaRonde-LeBlanc, N. and Wlodawer, A., A family portrait of the RIO kinases, J. Biol. Chem., 280, 37297, 2005. 83. Facchin, S. et al., Acidophilic nature of yeast PID261/BUD32, a putative ancestor of eukaryotic protein kinases, Biochem. Biophys. Res. Commun., 296, 1366, 2002. 84. Waters, E. et al., The genome of Nanoarchaeum equitans: Insights into early archaeal evolution and derived parasitism, Proc. Natl. Acad. Sci. U.S.A., 100, 12984, 2003. 85. Drennan, D. and Ryazanov, A.G., Alpha-kinases: Analysis of the family and comparison with conventional protein kinases, Prog. Biophys. Mol. Biol., 85, 1, 2004. 86. LaPorte, D.C., The isocitrate dehydrogenase phosphorylation cycle: Regulation and enzymology, J. Cell. Biochem., 51, 14, 1993. 87. Poncet, S. et al., HPr kinase/phosphatase, a Walker motif A-containing bifunctional sensor enzyme controlling catabolite repression in Gram-positive bacteria, Biochim. Biophys. Acta, 1697, 123, 2004. 88. Stonstrom, A. et al., Bioinformatic analyses of bacterial HPr kinase/phosphatase homologues, Res. Microbiol., 156, 443, 2005. 89. Bao, Q. et al., A complete sequence of the T. tencongensis genome, Genome Res., 12, 689, 2002. 90. Kennelly, P.J., Protein phosphatases: A phylogenetic perspective, Chem. Rev., 101, 2291, 2001. 91. Barford, D., Molecular mechanisms of the protein serine/threonine phosphatases, Trends Biochem. Sci., 21, 407, 1996. 92. Barford, D., Jia, Z. and Tonks, N.K., Protein tyrosine phosphatases take off, Nat. Struct. Biol., 2, 1043, 1995. 93. Ramponi, G. and Stefani, M., Structure and function of the low M r phosphotyrosine protein phosphatases, Biochim. Biophys. Acta, 1341, 137, 1997. 94. Andersen, J.K. et al., Structural and evolutionary relationships among protein tyrosine phosphatase domains, Mol. Cell. Biol., 21, 7117, 2001. 95. Barton, G.J., Cohen, P.T.W. and Barford, D., Conservation analysis and structure prediction of protein serine/threonine phosphatases. Sequence similarity with diadenosine tetraphosphatase from Escherichia coli suggests homology to protein phosphatases, Eur. J. Biochem., 220, 225, 1994. 96. Bork, P. et al., The protein phosphatase 2C (PP2C) superfamily: Detection of bacterial homologues, Prot. Sci., 5, 1421, 1996. 97. Lee, P. and Stock, A.M. Characterization of the genes and proteins of a two-component system from the hyperthermophilic bacterium Thermotoga maritima, J. Bacteriol., 178, 5579, 1996. 98. Foster, J.E. et al., Kinetic and mechanistic analyses of new classes of inhibitors of two-component signal transduction systems using a coupled assay containing HpkA-DrrA from Thermotoga maritime, Microbiology, 150, 885, 2004. 99. Goudreau, P.N., Lee, P. and Stock, A.M., Stabilization of the phospho-aspartyl residue in a twocomponent signal transduction system in Thermotoga maritima, Biochemistry, 37, 14575, 1998. 100. Bilwes, A.M. et al., Structure of CheA, a signal-transducing histidine kinase, Cell, 96, 131, 1999. 101. Bilwes, A.M. et al., Nucleotide binding by the histidine kinase CheA, Nat. Struct. Biol., 8, 353, 2001. 102. Park, S. et al., In different organisms, the mode of interaction between two signaling proteins is not necessarily conserved, Proc. Natl. Acad. Sci. U.S.A., 101, 11646, 2004. 103. Park, S. et al., Subunit exchange by CheA histidine kinases from the mesophile Escherichia coli and the thermophile Thermotoga maritima, Biochemistry, 43, 2228, 2004. 104. Quezada, C.M. et al., Helical shifts generate two distinct conformers in the atomic resolution structure of the CheA phosphotransferase domain from Thermotoga maritima, J. Mol. Biol., 341, 1283, 2004. 105. Quezada, C.M. et al., Structural and chemical requirements for histidine phosphorylation by the chemotaxis kinase CheA, J. Biol. Chem., 280, 30581, 2005. 106. Griswold, I.J., The solution structure and interactions of CheW from Thermotoga maritima, Nat. Struc. Biol., 9, 121, 2002. 107. Marina, A., Waldberger, C.D. and Hendrickson, W.A., Structure of the entire cytoplasmic portion of a sensor histidine-kinase protein, EMBO J., 24, 4247, 2005. 108. LaRonde-LaBlanc, N. and Wlodawer, A., Crystal structure of A. fulgidus Rio2 defines a new family of serine protein kinases, Structure, 12, 1585, 2004.

9214_C019.indd 330

11/5/2007 9:08:52 PM

Protein Phosphorylation at 80°C and Above

331

109. LaRonde-LeBlanc, N. et al., Structure and activity of the atypical serine kinase Rio1, FEBS J., 272, 3698, 2005. 110. LaRonde-LeBlanc, N. et al., Autophosphorylation of Archaeoglobus fulgidus Rio2 and crystal structures of its nucleotide-metal ion complexes, FEBS J., 272, 2800, 2005. 111. Angermayr, M., Roidl, A. and Brandlow, W., Yeast Rio1p is the founding member of a novel subfamily of protein serine kinases involved in control of cell cycle progression, Mol. Microbiol., 44, 309, 2002. 112. LaRonde-LeBlanc, N. and Wlodawer, A., The RIO kinases: An atypical protein kinase family required for ribosome biogenesis and cell cycle progression, Biochim. Biophys. Acta, 1754, 14, 2005. 113. Lower, B.H. and Kennelly, P.J., Open reading frame sso2387 from the archaeon Sulfolobus solfataricus encodes a polypeptide with protein-serine kinase activity, J. Bacteriol., 185, 3436, 2003. 114. Alber, S.-V. and Driessens, A.J.M., Analysis of ATPases of putative secretion operons in the thermophilic archaeon Sulfolobus solfataricus, Microbiology, 151, 763, 2005. 115. Johnson, L.N., Noble, M.E.E. and Owen, D.J., Active and inactive protein kinases: Structural basis for regulation, Cell, 85, 149, 1996. 116. Lower, B.H., Potters, M.B. and Kennelly, P.J., A phosphoprotein from the archaeon Sulfolobus solfataricus with protein-serine kinase activity, J. Bacteriol., 186, 463, 2004. 117. Tahara, M. et al., In vitro phosphorylation of initiation factor 2α (aIF2α) from hyperthermophilic archaeon Pyrococcus horikoshii OT3, J. Biochem. (Tokyo), 135, 479, 2004. 118. Bischoff, K.M. and Kennelly, P.J., “In-gel” assay for identifying alternative nucleotide substrates for protein kinases. Anal. Biochem., 271, 199, 1999. 119. Lower, B.H., Bischoff, K.M. and Kennelly, P.J., The archaeon Sulfolobus solfataricus contains a membrane-associated protein kinase that preferentially phosphorylates threonine residues, J. Bacteriol., 182, 3452, 2000. 120. Lower, B.H. and Kennelly, P.J., The membrane-associated protein-serine/threonine kinase from Sulfolobus solfataricus is a glycoprotein, J. Bacteriol., 184, 2614, 2002. 121. Kennelly, P.J. et al., Identification of a serine/threonine-specific protein phosphatase from the archaebacterium Sulfolobus solfataricus, J. Biol. Chem., 268, 6505, 1993. 122. Leng, J. et al., Isolation and cloning of a protein-serine/threonine phosphatase from an archaeon, J. Bacteriol., 177, 6510, 1995. 123. Mai, B. et al., Molecular cloning and functional expression of a protein-serine/threonine phosphatase from the hyperthermophilic archaeon Pyrodictium abyssi TAG11, J. Bacteriol., 180, 4030, 1998. 124. Jeon, S.J. et al., Tk-PTP, a protein-tyrosine phosphatase from the hyperthermophilic archaeon Thermococcus kodakaraensis KOD1: Enzymatic characteristics and identification of its substrate proteins, Biochem. Biophys. Res. Commun., 295, 508, 2002. 125. Savle, P.S. et al., N-(Cyclohexanecarboxyl)-O-phospho-L-serine, a minimal substrate for the dualspecificity protein phosphatase IphP, Arch. Biochem. Biophys., 376, 439, 2000. 126. Zhang, Z.-Y., Protein-tyrosine phosphatases: Biological function, structural characteristics, and mechanism of catalysis, Crit. Rev. Biochem. Mol. Biol., 33, 1, 1998. 127. Baker, M.D., Wolanin, P.M. and Stock, J.B., Signal transduction in bacterial chemotaxis. BioEssays, 28, 9, 2005. 128. Swanson, R.V., Sanna, M.S. and Simon, M.I., Thermostable chemotaxis proteins from the hyperthermophilic bacterium Thermotoga maritima, J. Bacteriol., 178, 484, 1996. 129. Wolfe, A.J., The acetate switch, Microbiol. Mol. Biol. Rev., 69, 12, 2005. 130. Usher, K.C. et al., Crystal structures of CheY from Thermotoga maritima do not support conventional explanations for the structural basis of enhanced thermostability, Protein Sci., 7, 403, 1998. 131. Buckler, D.R., Zhou, Y. and Stock, A.M., Evidence of intradomain and interdomain flexibility in an OmpR/RhoB homolog from Thermotoga maritima, Structure, 10, 153, 2002. 132. Robinson, V.L., Wu, T. and Stock, A.M., Structural analysis of the domain interface in DrrB, a response regulator of the OmpR/PhoB subfamily, J. Bacteriol., 185, 4186, 2003. 133. Blanco, A.G. et al., Tandem DNA recognition by PhoB, a two-component signal transduction transcriptional activator, Structure, 10, 701, 2002. 134. Lee, S. et al., Regulation of the transcriptional activator NtrC1: Structural studies of the regulatory and AAA+ ATPase domains, Genes Develop., 17, 2552, 2003. 135. Hirschman, J. et al., Products of nitrogen regulatory genes ntrA and ntrC of enteric bacteria activate glnA transcription in vitro: Evidence that the ntrA product is a σ factor, Proc. Natl. Acad. Sci. U.S.A., 82, 7525, 1985.

9214_C019.indd 331

11/5/2007 9:08:53 PM

332

Thermophiles: Biology and Technology at High Temperatures

136. Doucleff, M. et al., Negative regulation of AAA+ ATPase assembly by two component receiver domains: A transcription activation mechanism that is conserved in mesophilic and extremely hyperthermophilic bacteria, J. Mol. Biol., 353, 242, 2005. 137. Yatime, L. et al., Structure—Function relationships of the intact aIF2α subunit from the archaeon Pryococcus abyssi, Biochemistry, 44, 8749, 2005. 138. Pedulla, N. et al., The archaeal elF2 homologue: functional properties of an ancient translation initiation factor, Nucl. Acids Res., 33, 1804, 2005. 139. Lundgren, M. and Bernander, R., Archaeal cell cycle progress, Microbiology, 8, 662, 2005. 140. Grabowski, B. and Kelman, Z., Archaeal DNA replicaton: Eukaryal proteins in a bacterial context, Annu. Rev. Microbiol., 57, 487, 2003. 141. Grabowski, B. and Kelman, Z., Autophosphorylation of archaeal cdc6 homologues is regulated by DNA, J. Bacteriol., 183, 5459, 2001. 142. De Felice, M.L. et al., Biochemical characterization of a CDC6-like protein from the crenarchaeon Sulfolobus solfataricus. J. Biol. Chem., 278, 46424, 2003. 143. Kasiviswanathan, R., Shin, J. and Kelman, Z., Interactions between the archaeal Cdc6 and MCM proteins modulate their biochemical properties, Nucl. Acids Res., 33, 4940, 2005. 144. Almaula, N. et al., Nucleoside diphosphate kinase from Escherichia coli, J. Bacteriol, 177, 2524, 1995. 145. Regni, C., Schramm, A.M. and Beamer, L.J., The reaction of phosphohexomutase from Pseudomonas aeruginosa. Structural insight into a simple processive enzyme, J. Biol. Chem., 281, 15564, 2006. 146. Ray, W.K. et al., A phosphohexomutase from the archaeon Sulfolobus solfataricus is covalently modified by phosphorylation on serine, J. Bacteriol., 187, 4270, 2005. 147. Graves, J.D. and Krebs, E.G., Protein phosphorylation and signal transduction, Pharmacol. Ther., 82, 111, 1999. 148. Kennelly, P.J. and Potts, M., Fancy meeting you here! A fresh look at ‘prokaryotic’ protein phosphorylation, J. Bacteriol., 178, 4759, 1996. 149. Cozzone, A.J. et al., Protein phosphorylation on tyrosine in Bacteria, Arch Microbiol, 181, 171, 2004. 150. Flint, A.J. et al., Development of “substrate-trapping” mutants to identify physiological substrates of protein tyrosine phosphatases, Proc. Natl. Acad. Sci., 94, 1680, 1997. 151. Cardona, S. et al., The glycogen-bound polyphosphate kinase from Sulfolobus solfataricus is actually a glycogen synthase, Appl. Environ. Microbiol., 67, 4773, 2001. 152. Ahmed, H. et al., The semi-phosphorylative Entner-Doudoroff pathway in hyperthermophilic archaea: A re-evaluation, Biochem. J., 390, 529, 2005. 153. Kim, S. and Lee, S.B., Identification and characterization of Sulfolobus solfataricus D-gluconate dehydratase: A key enzyme in the non-phosphorylated Entner-Doudoroff pathway, Biochem. J., 387, 271, 2005. 154. Condo, I., A novel aminopeptidase associated with the 60 kDa chaperonin in the thermophilic archaeon Sulfolobus solfataricus, Mol. Microbiol., 29, 775, 1998. 155. Kumar, S. and Nussinov, R., Close-range electrostatic interactions in proteins, ChemBioChem, 3, 604, 2002. 156. Pain, R.H., Temperature and macromolecular structure and function, Symp. Soc. Exp. Biol., 41, 21, 1987. 157. Daniel, R.M. and Cowan, D.A., Biomolecular stability and life at high temperatures, Cell. Molec. Life Sci., 57, 250, 2000. 158. Gross, J. et al., Energy coupling in Salmonella typhimurium nicotinic acid phosphoribosyltransferase: Identification of His-219 as site of phosphorylation, Biochemistry, 35, 3917, 1996. 159. Hultquist, D.E., Moyer, R.W. and Boyer, P.D., The preparation and characterization of 1-phosphohistidine and 3-phosphohistidine, Biochemistry, 5, 322, 1966. 160. McCleary, W.R. and Zusman, D.R., Purification and characterization of the Myxococcus xanthus FrzE protein shows that it has autophosphorylation activity, J. Bacteriol., 172, 6661, 1990. 161. Post, R.L. and Kume, S., Evidence for an aspartyl phosphate residue at the active site of sodium and potassium ion transport adenosine triphosphatase, J. Biol. Chem., 248, 6993, 1973. 162. Sheeler, N.L., MacMillan, S.V. and Nodwell, J.R., Biochemical activities of the absA two-component system of Streptomyces coelicolor, J. Bacteriol., 187, 687, 2005. 163. Weiss, V. and Magasanik, B., Phosphorylation of nitrogen regulator I (NRI) of Escherichia coli, Proc. Natl. Acad. Sci. U.S.A., 85, 8919, 1988. 164. Demidov, V.V., Radiation and thermal stabilities of adenine nucleotides, Adv. Space Res., 15, 131, 1995.

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Archaeal 20S Proteasome: A Simple and Thermostable Model System for the Core Particle Joshua K. Michel and Robert M. Kelly

CONTENTS Introduction ................................................................................................................................ 20S Proteasome Structure ......................................................................................................... 20S Proteasome Assembly ......................................................................................................... Role of α Subunits ..................................................................................................................... Role of β Subunits ...................................................................................................................... Proteasome Biocatalysis ............................................................................................................ Concluding Remarks.................................................................................................................... Acknowledgments ...................................................................................................................... References ..................................................................................................................................

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INTRODUCTION The 20S proteasome (also referred to as the core particle, CP) is a cylindrically shaped protease ubiquitous to both Archaea and Eukarya.1,2 The 20S proteasome has also been found in the actinomycetes Rhodococcus erythropoli,3 Myobacterium tuberculosis,4 Streptomyces coelicolar,5 and Frankia.6 The lack of proteasome genes encoded in other bacterial genomes suggests that the actinomycetes acquired the proteasome by lateral gene transfer.7 Most bacteria contain a related complex, ClpQY (or HslVU), that shares a similar catalytic mechanism to the proteasome and apparently plays a similar functional role.8 The eukaryotic and archaeal 20S proteasomes share a similar overall size and structure, but differ in complexity—the archaeal proteasome is based on fewer unique proteins (which may be processed prior to being incorporated into the macromolecular complex as subunits). As such, the 20S archaeal proteasome serves as a simpler model system for examining the significance of subunit composition on biochemical, biophysical, and functional properties of the multimeric protease. In addition, the archaeal proteasome, presumably related to the ancestral eukaryotic proteasome precursor, may provide insight into how eukaryotic proteasomes developed into the complex proteases that now exist.

20S PROTEASOME STRUCTURE The 20S proteasome from the thermophilic archaeon Thermoplasma acidophilum was the first archaeal proteasome structure resolved and subsequently has been found to be closely related to all 333

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FIGURE 20.1 Assembly of the proteasome involves formation of ~300 kDa half-proteasome complexes (which are inactive and contain α and β pro-subunits) prior to cleavage of the β protein pro-peptide region. Combination of two half-proteasomes results in an active 600 to 700 kDa enzyme capable of degrading peptides and unfolded proteins.

archaeal proteasomes examined to date.2,9–13 Like its eukaryotic counterpart, the 20S archaeal proteasome is composed of four stacked heptametric rings that form a barrel-like structure with a hollow channel extending down the center.14 The hollow channel is comprised of three cavities consisting of two antechambers formed by the α rings and a central channel formed by the β rings.9 To prevent undesirable protein degradation in the cytosol, active sites are compartmentalized within the interior channel of the 20S proteasome. The overall length of the cylindrical enzyme is 148 Å with maximum and minimum diameters of 113 Å and 75 Å, respectively.9 Each homomeric ring is composed entirely of either α or β subunits, arranged in the order α7β7β7α7, as shown in Figure 20.1. The catalytic core consists of 6 to 14 active sites [typically involving a Threonine (Thr) residue] located on the N-terminal regions of the β subunits.15,16 Access to the central chamber of the archeal 20S proteasome is facilitated through 13 Å openings on either end of the cylinder.17 Comparatively, the eukarotic 20S proteasome core is inaccessible through the ends, except by major rearrangement of the α subunit N-termini that creates an opening of 10 Å.15 Unlike eukaryotic genomes that have been found to encode up to 23 unique proteasome α/β subunit proteins, the T. acidophilum proteasome is comprised of only two different subunits (one α and one β) with molecular weights of 20 and 35 kDa, respectively.17,18 This simple basis for 20S proteasome structure is typical among all archaea, whose genomes encode between two and four different α/β subunit proteins.19–21

20S PROTEASOME ASSEMBLY The creation of the 20S proteasome is a complex process in which α proteins self-assemble into rings that then act as scaffolds for the subsequent β ring formation.22,23 The result is a halfproteasome complex, with an approximate size of 300 kDa. These ring-dimers in turn associate and cleave the β pro-pepide to form the complete and functional 600 to 700 kDa enzyme.24 As one of several measures to prevent to unwanted proteolysis, the β proteins cannot form ring structures in the absence of α proteins.25 For Archaeoglobus fulgidus, there is no major conformational change of contact areas after formation of the α–β complex, suggesting that these regions are complementary prior to assembly.26 This assembly process appears to be conserved across all archaea, but as structural complexity increases (more than one α and/or β subunit type) so do the assembly mechanisms.24 For many thermophilic archaea, in vitro combination of the α and β proteins leads to spontaneous self-directing assembly of the 20S proteasome, without need for chaperones or other accessory proteins.20,27 However, this process can be extremely inefficient, such as is the case for the Methanosarcina thermophila 20S proteasome in which only 50% of the β proteins were processed

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and incorporated into a fully active 20S CP.25 Likewise, the assembly of the Pyrococcus furiosus 20S proteasome at physiologically relevant temperatures yielded a substantial amount of unincorporated β proteins.28 The recombinant 20S proteasome from Methanococcus jannaschii required denaturation of α and β subunits with urea, followed by their combination at high temperature and subsequent refolding by removal of the denaturant using dialysis. Without unfolding-refolding of the M. jannaschii proteasome, an active enzyme was produced, but with a markedly decreased optimal temperature (95°C) compared to the native version (119°C).29 Presumably, the native intracellular assembly environment increases the folding efficiency of 20S proteasomes. As a safeguard to unwanted proteolysis prior to assembly, β proteins are expressed as precursors that undergo self-processing (removal of ~10 N-terminal amino acid residues to expose the active site Thr) during assembly into the active structure.25,26,30,31 The precursor N-terminal region in yeast is essential for proper incorporation of the β subunit into the proteasome32, thus acting as an intramolecular chaperone. Conversely, β proteasome incorporation in T. acidophilum, showed no dependence on the pro-peptide region,22 indicating that the archaeal β pro-peptide functions solely as a temporary inhibitor of proteolysis. The pro-peptide also acts to protect the active site; premature processing of the β protein results in acetylation of Thr residues, yielding an inactive enzyme.32 Cleavage of the pro-peptide occurs during combination of dimer rings into a fully functional 20S proteasomes.33 The processing of the β-pro-peptide appears to be intramolecular and autocatalytic in nature, while the α protein is not subject to any cleavage.31 The N-terminal region of the α subunit contains an α-helix that is required for proper ring formation.22 While the α proteins do not contain a pro-peptide sequence, it has been suggested that the archaeal α subunits may be subjected to post-translational modification, such as phosphorylation.34,35 The influence and purpose of α protein modifications on the 20S proteasome’s interaction with regulatory proteins such as PAN (proteasome-activating nucleotidase) have yet to be fully elucidated.

ROLE OF α SUBUNITS Access to the 20S proteasome interior is regulated by the α subunits, which in eukaryotes form a stable plug on each end of the cylinder.15 In yeast, extension of the N-termini from α1,2,3,6,7 prevents access of proteins to the center chamber; multiple hydrogen bonds are formed between the overlapping α subunit N-termini, contributing significantly to the stability of the “gates.”15,36 Opening of the 20S eukaryotic proteasome axial channel is facilitated by attachment of the 19S regulatory component.37–39 Combination of the 19S and 20S proteasomes, into the 26S complex, creates an enzymatic machine cable of selectively degrading ubiquitin-tagged proteins.40–43 Conversely, 20S proteasome structures from T. thermophila and A. fulgidus show disorder among the α N-termini with the presence of only one hydrogen bond between the Asp9 and Tyr8 residues of adjacent α subunits.9,17,26 Lack of α subunit interaction contributes to N-terminal flexibility of archaeal proteasomes, which allows entry of unfolded proteins and peptides without need for the ATPdependent protein PAN.44,45 Hyperthermophilic archaea encode both single and multiple homologs of α and β proteasome components, as shown in Table 20.1. The rarity of archaeal genomes encoding multiple α proteins supports genome sequence data suggesting β differentiation evolutionarily predated that of the α-subunits.46 One example of α differentiation is found in the native proteasome from Methanosarcina thermophila, which contain multiple α subunits differing in length by four amino acids, but encoded by the same gene.47 The difference in α subunit length has been attributed to three potential translation start sites within the α gene.25 Alternatively, multiple native 20S proteasome sub-types have also been found in the haloarchaea; Haloferax volcanii produces at least two distinct proteasome sub-types resulting from two unique α genes.48 Immunoanalysis of the H. volcanii proteasome shows the α1 subunit represented 60% of incorporated α protein, while α2 comprised the other 40%.48 However, α2 has exhibited a seven-fold transcriptional increase during the transition from exponential growth to stationary phase,49 indicating the involvement of post-translational mechanisms

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95 83 50 50–53 45 85 98 50 65–70 45 90 60 100 96 100 98 75 87 80 59 60 95

No ORF annotated as PAN, but possible homolog detected. Note: ND - No PAN homolog detected Source: Adapted from Madding, L.S. et al., J. Bacteriol., 189, 583, 2007.

*

Aeropyrum pernix Archaeoglobus fulgidus Halobacterium sp. NRC-1 Haloarcula marismortui 43049 Haloferax volcanii Methanocaldococcus jannaschii Methanopyrus kandleri AV19 Methanosarcina thermophila Methanotherm. thermautotrophicus Natronomonas pharaonis 2160 Nanoarchaeum equitans Kin4-M Picrophilus torridus DSM 9790 Pyrobaculum aerophilum Pyrococcus abyssi Pyrococcus furiosus Pyrococcus horikoshii Sulfolobus acidocaldarius Sulfolobus solfataricus Sulfolobus tokodaii Thermoplasma acidophilum Thermoplasma volcanium Thermococcus kodakarensis

(ºC)

AAV46124

α protein APE1449 AF0490 VNG0166G AAV46668 T48679 MJ0591 MK0385 MTU30483 MTH686 NP3738A AAR39362 PTO0804 PAE2215 PAB0417 PF1571 PH1553 AAY80005 SSO0738 ST0446 TA1288 TVN0304 TK1637

TABLE 20.1 Proteasomes from Thermophilic and Hyperthermophilic Archaea APE0521 AF0481 VNG0880G AAV45476 T48677 MJ1237 MK1228 MTU22157 MTH1202 NP3472A AAR39057 PTO0686 PAE3595 PAB1867 PF1404 PH1402 AAY80046 SSO0766 ST0477 TA0612 TVN0663 TK1429 TK2207

PAE0807 PAB2199 PF0159 PH0245 AAY80272 SSO0278 ST0324

AAV46667

APE0507

β protein APE2012 AF1976 VNG2000G AAV47895 AAV38127 MJ1176 MK0878 ND MTH728 NP1524A AAR39040 PTO0456* PAE0696* PAB2233 PF0115 PH0201 AAO73475 SSO0271 ST0330 TA0840* TVN0947* TK2252

PAN

NP5038A

VNG0510G AAV48212 AAV38126

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to regulate subunit levels.49 While noted for a limited number of archaeal proteins,50–52 α1 and α2 proteins have been found subject to N-terminal acetylation.34 Alternatively, H. volcanii may alter proteasome composition to better adapt to certain growth phase conditions. With the limited data available for multiple α proteins in Archaea, the impact of different α subunits on structure and activity is largely unknown. In eukaryotic organisms, the 20S proteasome is distributed throughout the cellular space,53 but during mitosis there is an increased movement of proteasomes to the nucleus.54 In solid tumor cells, cellular stress (e.g., hypoxia and starvation) causes nuclear localization of the proteasome, which subsequently induces drug resistance.55 Nuclear localization signals (NLS), found on the α subunits, facilitate intracellular proteasome transport; a highly similar sequence to the NLS is also found on the T. acidophilum α-subunit.56 The production of a recombinant version of this archaeal proteasome in mouse cells results in active import of the T. acidophilum proteasome into the nucleus.57 Since T. acidophilum does not contain a nucleus, this NLS-similar sequence may represent an alternative signal that later developed into the NLS of eukaryotic organisms. Observations that T. acidophilum cell-lysate facilitates the nuclear import of both human and Thermoplasma proteasome58 supports the use of the archaeal system as a model for NLS-mediated relocation.

ROLE OF β SUBUNITS Eukaryotic organisms ubiquitously encode a highly diversified set of subunit proteins: the yeast proteasomes consist of seven α and seven β subunits, while mammalian cells encode for seven α and ten β proteins.10 The 14 different subunits encoded by yeast are essential for cellular survival under standard growth conditions.32,59,60 Other efforts with eukaryotic organisms demonstrated significant cellular changes from loss of proteasome subunit function.61,62 The presence of three γ−inducible, mammalian proteasome β subunits has been linked to production of alternative degradation products for antigenic presentation on cellular surfaces.63 Similarly, Drosophila melanogaster produces six male-specific 20S proteasome subunit isoforms only during late spermatogenesis.64 Functional requirement for proteasome activity varies by organism, and may depend on the availability of complementary proteases to compensate for proteasome loss.21 Mutation of the 20S proteasome in Mycobacterium smegmatis showed no effect on cellular growth or degradation of peptides, but in Mycobacterium tuberculosis the proteasome was required for growth during oxidative or nitrosactive stress.4,65 For eukaryotes, the large number of proteases encoded in their genomes makes determination of proteasome-specific functions difficult. Native versions of the archaeal proteasome have been isolated and characterized from P. furiosus,66 M. jannaschii,67 M. thermophila,47 H. volcanii,35 and Haloarcula marismortui.68 Inhibition of the entire T. acidophilum 20S proteasome impacted cellular survival during heat stress.69 Proteasome inhibition studies carried out in H. volcanii demonstrated that the lack of an active 20S proteasome led to a 30% decrease in growth rate under otherwise optimal conditions.70 The necessity of proteasome function during stress conditions most likely stems from this protease’s ability to degrade large unfolded, misfolded, and damaged proteins. The genomes of M. thermophila, H. volcanii, M. jannaschii and A. fulgidus encode only single versions of α and β proteins.71 On the other hand, the genomes of Sulfolobus solfataricus and P. furiosus contain a single α protein gene and two distinct β protein genes.21 The purpose and function of these additional β subunit homologs in archaeal proteasomes is not known, although this simple model could offer insight into the maturation of the diverse set of homologous β subunits found in eukaryotes. Transcriptional response studies on hyperthermophilic archaea containing proteasomes comprised of single α and β subunits show a consistent tendency for higher transcription of β during stress conditions. The A. fulgidus proteasome exhibited a 2.7-fold decrease in transcription of the α gene after heat shock, while transcripts for β exhibited a slight increase.72 M. jannaschii showed no significant change in transcription of 20S proteasome genes in response to a 10°C temperature increase.73 But, M. jannaschii did respond to a 20°C decease in temperature with a 2.3-fold

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increase in β transcription, while α remained unchanged.73 The increased M. jannaschii β transcription during cold shock may be related to the difficulty of proteasome formation at sub-optimal conditions.29,74 In addition, the α protein of M. jannaschii is unique because of the conservation of amino acid residues associated with the active site of β, which may result in transcriptional trends differing from other archaea. These conserved residues in the M. jannaschii α protein are likely a remnant from times before differentiation of α and β. Unlike the case in eukaryotes, in which some multiple versions of α and β proteins have been observed to share 90% identical protein sequences,75 the two P. furiosus β proteins are only 48% identical at the amino acid level.28 This suggests that the β subunits can play distinct roles in P. furiosus CP structure and function. Transcriptional analysis of the three subunits from P. furiosus showed that, under heat shock conditions, the α gene was down-regulated 2.0-fold, β1 increased 2-fold, and β2 remained relatively constant. The decreased transcription of the P. furiosus α subunit is consistent with results for heat shocked A. fulgidus,72 and may relate to the inherent thermal stability of α proteins compared with β. The P. furiosus β1 and β2 proteins exhibit melting temperatures of 104.4°C and 93.1°C respectively28, while the α protein is estimated to melt at approximately 135°C.76 For P. furiosus, active recombinant enzymes can be formed from combinations of α and β2, as well as α+β1+β2.28 When these enzymes were assembled at 90°C, the version with β1 demonstrated a slightly higher activity. However, versions containing α+β1+β2 assembled at 105°C demonstrated significantly higher activity and thermostability. Two-dimensional gel electrophoresis showed a greater amount of β1, compared to β2, incorporated into the recombinant P. furiosus proteasome at higher assembly temperatures.28 Native proteasomes isolated from P. furiosus cells grown under heat shock conditions demonstrated a ratio of β1 to β2 similar to the ratio noted for the recombinant enzymes assembled at higher temperatures; native proteasomes from nonstressed cells contained a lower amount of β1, as shown in Figure 20.2. This suggests that the β1 subunit, while not essential for catalytic activity, plays a role in stabilizing and activating the P. furiosus CP assembly, particularly at supraoptimal temperatures, such as those encountered during thermal stress events. Furthermore, the response of P. furiosus to produce proteasomes with higher β1 content was controlled at both the transcriptional level and during enzyme assembly. For the hyperthermophilic archaea, it is not clear whether CPs with 2 β subunits create any special physiological or ecological advantages during thermal stress response. Perhaps hyperthermophiles whose genomes encode two β proteins experience frequent temperature excursions in their natural habitats. Recent results from our laboratory showed that when S. solfataricus was shifted from optimal (80°C) to supraoptimal (90°C) temperatures, the transcription of ORFs encoding CP proteins (α, β1, β2) were unaffected throughout the 60 min period following the temperature shift.77 However, it was also noted that the transcriptional level of CP proteins in S. solfataricus were much higher than in P. furiosus under both normal and stressed conditions. In any case, these data suggest that the impact of CP β subunit content on function at suboptimal, optimal, and supraoptimal temperatures merits further examination. The relative conservation of proteasome constituent proteins across all domains of life is a primary reason that archaeal proteasomes can serve as model systems for investigating structure and function issues of this complex protease. In fact, recombinant 20S proteasome proteins form active proteases when combined with pro-subunits from different organisms. The possibility to form a hybrid proteasome, consisting of subunits from different organisms was first noted for Aeropyrum pernix and A. fulgidus.26 The α protein from A. fulgidus has recently demonstrated the ability to form active proteasome CPs when combined with either P. furiosus β1, β2, or the combination of both β1 and β2.76 In addition to the creation of a hybrid CP, the formation of an active enzyme by A. fulgidus α + P. furiosus β1 was interesting since P. furiosus β1 does not from an active CP when combined with P. furiosus α.28 While intra-domain hybrid 20S proteasomes are fully assembled and active, the creation of inter-domain hybrids has yet to be demonstrated. Formation of such hybrids might be biotechnologically relevant in light of reports demonstrating the in vivo degradation of aggregation-prone proteins by a mesophilic archaeal proteasome expressed in mammalian cells.78

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FIGURE 20.2 The β1/β2 ratio for the recombinant P. furiosus proteasomes assembled at temperatures between 80ºC and 105ºC, as well as for the native proteasomes isolated from P. furiosus grown at 80ºC and 90ºC. Above each temperature bar, schematic representations of the 20S proteasome assembly are shown accounting for the altered β1 content.

PROTEASOME BIOCATALYSIS The 20S proteasome is a member of the T1 peptidase family71 characterized by an N-terminal Thr nucleophile.79 While initial observations noted that a Ser residue could be substituted in place of Thr without modifying hydrolysis of LLVY-Amc,80 it was subsequently shown that the cleavage pattern was impacted as a result of the residue change.81 The hydroxyl group of Thr apparently initiates hydrolysis of the peptide bond, followed by nucleophilic attack from a water molecule resulting in peptide bond cleavage.81,82 Even though the substrate specificity of 20S proteasomes may vary, the N-terminal Thr residue is conserved in all known active β subunits.1,19,20,27,43 Individual β proteins comprising eukaryotic proteasome are associated with certain substrate specificities.32 Yeast proteasome subunits β1, β2, and β5 are linked to chymotrypsin-like, trypsin-like, and post-glutamyl peptide hydrolase-like activities, respectively.15,59,83,84 The β1 and β2 active sites prefer cleavage after acidic and basic residues on the substrate protein or peptide.85,86 However, while the three active sites differ in their cleavage preference, all contribute significantly to protein

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degradation.87 The limited number of active sites (i.e., 3) in eukaryotic proteasomes differs significantly from the 14 active sites present in the archaeal CP that exhibit only chymotrypsin-like activity. The increased number of active sites present in the archaeal proteasome has yet to be connected to a physiological role or advantage. In part, this is due to a lack of understanding of which factors influence activity and substrate specificity. A prime example is the M. jannaschii proteasome that consists of two unique subunits (α + β) with 14 active sites distributed in β chamber. The M. jannaschii native proteasome exhibits an optimal activity at 119°C,67 although recombinant versions showed optimal temperatures of 95°C and 110°C, depending on assembly conditions.74 The recombinant M. jannaschii 20S proteasome with an optimal temperature closest to the native organism was subjected to chemical denaturation followed by refolding at elevated temperatures, thus demonstrating the importance of assembly conditions on enzyme function. In addition, recombinant M. jannaschii proteasomes have demonstrated decreased enzymatic activity in response to high-osmotic pressure, while the native enzyme exhibited increased protease activity under these conditions.29 Most studies on native proteasomes have focused on structure, size, and substrate preferences. Table 20.2 summarizes the biochemical data available for several of the native archaeal proteasomes characterized to date; note that detailed kinetics are typically carried out only on recombinant versions such that information from such studies may not reflect in vivo properties (Table 20.3). The range of activities presented in Table 20.2 may indicate diversity within the archaeal 20S proteasome family; however, the complete picture remains unclear without detailed kinetics and cleavage patterns for the native enzymes. Inconsistencies in extents of protein purification make direct biocatalytic comparisons between specific proteasomes intractable. Differences between protocols and purification levels for native and recombinant proteasomes represent an obstacle to the more widespread use of archaeal 20S proteasomes as model systems. The impact of multiple α or β proteins on archaeal proteasome biocatalysis has not been studied to any extent as yet. The 20S proteasome from P. furiosus has been examined along these lines and the presence of two alternative β pro-subunits impacted biocatalytic and biophysical properties.66 For the P. furiosus CP, both the β1 and β2 proteins contain the conserved active site Thr residue,

TABLE 20.2 Kinetic and Physical Properties of Selected Native Proteasomes from High-Temperature Microorganisms Organism

Methanococcus jannaschii

Methanosarcina thermophila

Preferred Substrate Sp.Activity (nmol/min mg)

Cbz-AAL-βNa

Cbz-LLE-βNa

Suc-AAF-Amc

Suc-LLVY-Amc

Suc-VKM-Amc

116000 (95ºC)*

115.0 (65ºC)*

340.0 (60ºC)*

0.79

Vm = 2200

pHoptimal Toptimal (ºC) References

7.5-7.8 119 67

NR NR 25,47

Haloferax volcanii

7.0–9.3 75 35

Thermoplasma acidophilum

Pyrococcus furiosus

(pmol/min μg)

NR NR 17

kcat/Km = 3.64 (s−1 mM−1) 6.5 95 66

*

Represents assay temperature for reported specific activity. Abbreviations: Amc, 7-amino-4-methylcoumarin; βNa, β-naphthylamine; NR, Not reported.

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NR 75 LLE-βNa LLVY-Amc AAF-Amc 25

0.018 119 LLE-βNa LLVY-Amc AAF-Amc 29,74 35

NR NR LLVY-Amc

NR

NR

NR

NR

Haloferax volcanii

Abbreviations: Amc, 7-amino-4-methylcoumarin; βNa, β-naphthylamine; NR, Not reported.

References

NR

NR

kcat/Km (s−1 mM−1) ki (115ºC) Toptimal (ºC) Substrate Preference

NR

NR

kcat (s−1)

μg)

NR NR

NR NR

Km (μM)

Vm (pmol/min

Methanosarcina thermophila

Methanococcus jannaschii

Organisms

80,81

NR 60 GGL-Amc LLVY-Amc AAF-Amc

7.0

0.03

250

30.0

Thermoplasma acidophilum

28

0.15 >100.0 VKM-Amc AAF-Amc LLVY-Amc

22.1

1.0

1159

45.2

Pyrococcus furiosus (90ºC Assembly)

TABLE 20.3 Kinetic and Physical Properties of Selected Recombinant Proteasomes from High-Temperature Microorganisms

0.025 >100.0 VKM-Amc LLVY-Amc AAF-Amc

51.5

1.9

2194

36.4

Pyrococcus furiosus (105ºC Assembly)

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implying that 14 active sites are present in this enzyme. However, the increased incorporation of β1 at 105°C assembly (Table 20.2) altered the substrate preference by increasing the degradation of LLVY-MCA compared to AAF-MCA.28 Alternatively, a preference was shown for AAF-MCA by the P. furiosus proteasome assembled at 90°C (lower ratio of β1). Furthermore, increased incorporation of β1 into the CP yielded an enzyme with almost twice the Vmax of the version with less β1.28 These results support a role for β1 in both biocatalysis as well as structural integrity at elevated temperatures. It is worth noting that the kinetic properties of the recombinant P. furiosus 20S proteasome were consistent with previously reported values for the native CP in terms of both Vmax (2200 pmol/min-μg) and K m (0.46 mM).66

CONCLUDING REMARKS Archaeal 20S proteasomes have the potential to serve as simpler model systems for core particles from eukaryotic sources, especially in determining the role of α/β subunit composition in biochemical and physiological function. This is primarily due to the limited number of β subunits found within a single archaeal organism’s proteasome and the fact that archaeal proteasomes are apparently functional in eukarya. As such, the mechanisms of assembly, biochemical and biophysical characteristics, and cellular roles for archaeal 20S proteasomes provide a basis for examining aspects of the complex heteromultimeric proteasomes in eukaryotic systems. Additionally, the ease of recombinant production, assembly, and purification of thermophilic and hyperthermophilic proteasomes makes them attractive model systems from a logistical perspective. One of the current disadvantages of using hyperthermophilic proteasomes as model systems is the lack of data regarding the enzyme’s in vivo function. However, as genetic systems become more widely available for archaea, such as in the hyperthermophilic organisms Thermococcus KOD and Sulfolobus spp (see Chapters 11 and 13), the contribution of the 20S proteasome to cellular function can be scrutinized.88–90 Furthermore, intriguing possibilities using archaeal proteasomes for novel therapeutic strategies exist and merit further examination in their own right.78

ACKNOWLEDGMENTS This work was supported in part by grants from the Biotechnology Program of the U.S. National Science Foundation and the Energy Biosciences Program of the U.S. Department of Energy. JK Michel acknowledges support from an NIH Biotechnology Traineeship.

REFERENCES 1. Bochtler, M. et al., The proteasome, Annu. Rev. Biophys. Biomol. Struct., 28, 295, 1999. 2. Barber, R.D. and Ferry, J.G., Archaeal proteasomes, Methods Enzymol., 330, 413, 2001. 3. Tamura, T. et al., The first characterization of a eubacterial proteasome—The 2os complex of Rhodococcus, Curr. Biol., 5, 766, 1995. 4. Darwin, K.H. et al., The proteasome of Mycobacterium tuberculosis is required for resistance to nitric oxide, Science, 302, 1963, 2003. 5. Nagy, I. et al., The 20S proteasome of Streptomyces coelicolor, J. Bacteriol., 180, 5448, 1998. 6. Benoist, P. et al., High-molecular-mass multicatalytic proteinase complexes produced by the nitrogenfixing Actinomycete frankia Strain Br, J. Bacteriol., 174, 1495, 1992. 7. Lupas, A. et al., Self-compartmentalizing proteases, Trends Biochem. Sci., 22, 399, 1997. 8. Bochtler, M. et al., Crystal structure of heat shock locus V (HslV) from Escherichia coli, Proc. Natl. Acad. Sci. U. S. A., 94, 6070, 1997. 9. Lowe, J. et al., Crystal structure of the 20S proteasome from the archaeon T. acidophilum at 3.4 A resolution, Science, 268, 533, 1995.

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10. Groll, M. and Huber, R. in Ubiquitin And Protein Degradation, Part A 329 (2005). 11. Groll, M. et al., A gated channel into the proteasome core particle, Nat. Struct. Biol., 7, 1062, 2000. 12. De Castro, R.E. et al., Haloarchaeal proteases and proteolytic systems, FEMS Microbiol. Rev., 30, 17, 2006. 13. Puhler, G. et al., Proteasomes—multisubunit proteinases common to Thermoplasma and Eukaryotes, Syst. Appl. Microbiol., 16, 734, 1994. 14. Rechsteiner, M., Hoffman, L. and Dubiel, W., The Multicatalytic And 26S Proteases, J. Biol. Chem., 268, 6065, 1993. 15. Groll, M. et al., Structure of 20S proteasome from yeast at 2.4 angstrom resolution, Nature, 386, 463, 1997. 16. Hegerl, R. et al., The 3-dimensional structure of proteasomes from Thermoplasma acidophilum as determined by electron-microscopy using random conical tilting, FEBS Lett., 283, 117, 1991. 17. Zwickl, P. et al., Primary structure of the Thermoplasma proteasome and its implications for the structure, function, and evolution of the multicatalytic proteinase, Biochemistry (Mosc), 31, 964, 1992. 18. Puhler, G. et al., Subunit stoichiometry and 3-dimensional arrangement in proteasomes from Thermoplasma acidophilum, EMBO J., 11, 1607, 1992. 19. Groll, M. and Clausen, T., Molecular shredders: how proteasomes fulfill their role, Curr. Opin. Struct. Biol., 13, 665, 2003. 20. Maupin-Furlow, J.A. et al., Proteasomes: perspectives from the Archaea, Front. Biosci., 9, 1743, 2004. 21. Ward, D.E. et al., Proteolysis in hyperthermophilic microorganisms, Archaea, 1, 63, 2002. 22. Zwickl, P., Kleinz, J. and Baumeister, W., Critical elements in proteasome assembly, Nat. Struct. Biol., 1, 765, 1994. 23. Baumeister, W. et al., The proteasome: paradigm of a self-compartmentalizing protease, Cell, 92, 367, 1998. 24. Zuhl, F. et al., Dissecting the assembly pathway of the 20S proteasome, FEBS Lett., 418, 189, 1997. 25. Maupin-Furlow, J.A., Aldrich, H.C. and Ferry, J.G., Biochemical characterization of the 20S proteasome from the methanoarchaeon Methanosarcina thermophila, J. Bacteriol., 180, 1480, 1998. 26. Groll, M. et al., Investigations on the maturation and regulation of archaebacterial proteasomes, J. Mol. Biol., 327, 75, 2003. 27. Maupin-Furlow, J.A. et al., Archaeal proteasomes: proteolytic nanocompartments of the cell, Adv. Appl. Microbiol., 50, 279, 2001. 28. Madding, L.S. et al., Role of β1 subunit in function and stability of the 20S proteasome from the hyperthermophilic archaeon Pyrococcus furiosus, J. Bacteriol., 189, 583, 2007. 29. Frankenberg, R.J., Andersson, M. and Clark, D.S., Effect of temperature and pressure on the proteolytic specificity of the recombinant 20S proteasome from Methanococcus jannaschii, Extremophiles, 7, 353, 2003. 30. Chen, P. and Hochstrasser, M., Autocatalytic subunit processing couples active site formation in the 20S proteasome to completion of assembly, Cell, 86, 961, 1996. 31. Seemuller, E., Lupas, A. and Baumeister, W., Autocatalytic processing of the 20S proteasome, Nature, 382, 468, 1996. 32. Jager, S. et al., Proteasome beta-type subunits: Unequal roles of propeptides in core particle maturation and a hierarchy of active site function, J. Mol. Biol., 291, 997, 1999. 33. Nandi, D. et al., Intermediates in the formation of mouse 20S proteasomes: Implications for the assembly of precursor beta subunits, EMBO J., 16, 5363, 1997. 34. Humbard, M.A., Stevens, S.M. and Maupin-Furlow, J.A., Posttranslational modification of the 20S proteasomal proteins of the archaeon Haloferax volcanii, J. Bacteriol., 188, 7521, 2006. 35. Wilson, H.L., Aldrich, H.C. and Maupin-Furlow, J., Halophilic 20S proteasomes of the archaeon Haloferax volcanii: purification, characterization, and gene sequence analysis, J. Bacteriol., 181, 5814, 1999. 36. Whitby, F.G. et al., Structural basis for the activation of 20S proteasomes by 11S regulators, Nature, 408, 115, 2000. 37. Smith, D.M., Benaroudj, N. and Goldberg, A., Proteasomes and their associated ATPases: A destructive combination, J. Struct. Biol., 156, 72, 2006.

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38. Dennis, A.P. and O’malley, B.W., Rush hour at the promoter: How the ubiquitin-proteasome pathway polices the traffic flow of nuclear receptor-dependent transcription, J. Steroid Biochem. Mol. Biol., 93, 139, 2005. 39. Groll, M. and Huber, R., Substrate access and processing by the 20S proteasome core particle, Int. J. Biochem. Cell Biol., 35, 606, 2003. 40. Glickman, M.H. and Adir, N., The proteasome and the delicate balance between destruction and rescue, Plos Biology, 2, 25, 2004. 41. Glickman, M.H. and Ciechanover, A., The ubiquitin-proteasome proteolytic pathway: destruction for the sake of construction, Physiol. Rev., 82, 373, 2002. 42. Pickart, C.M. and Fushman, D., Polyubiquitin chains: polymeric protein signals, Curr. Opin. Chem. Biol., 8, 610, 2004. 43. Wolf, D.H. and Hilt, W., The proteasome: a proteolytic nanomachine of cell regulation and waste disposal, Biochimica Et Biophysica Acta-Molecular Cell Research, 1695, 19, 2004. 44. Medalia, N. et al., Functional and structural characterization of the Methanosarcina mazei proteasome and PAN complexes, J. Struct. Biol., 156, 84, 2006. 45. Forster, A. et al., The 1.9 A structure of a proteasome-11S activator complex and implications for proteasome-PAN/PA700 interactions, Mol. Cell, 18, 589, 2005. 46. Gille, C. et al., A comprehensive view on proteasomal sequences: implications for the evolution of the proteasome, J. Mol. Biol., 326, 1448, 2003. 47. Maupin-Furlow, J.A. and Ferry, J.G., A proteasome from the methanogenic Archaeon Methanosarcina thermophila, J. Biol. Chem., 270, 28617, 1995. 48. Kaczowka, S.J. and Maupin-Furlow, J.A., Subunit topology of two 20S proteasomes from Haloferax volcanii, J. Bacteriol., 185, 165, 2003. 49. Reuter, C.J., Kaczowka, S.J. and Maupin-Furlow, J.A., Differential regulation of the PanA and PanB proteasome-activating nucleotidase and 20S proteasomal proteins of the haloarchaeon Haloferax volcanii, J. Bacteriol., 186, 7763, 2004. 50. Mattar, S. et al., The primary structure of Halocyanin, an archaeal blue copper protein, predicts a lipid anchor for membrane fixation, J. Biol. Chem., 269, 14939, 1994. 51. Klussmann, S. et al., N-terminal modification and amino acid sequence of the ribosomal protein Hmas7 From Haloarcula marismortui and homology studies to other ribosomal proteins, Biol. Chem. Hoppe. Seyler, 374, 305, 1993. 52. Maras, B. et al., The protein sequence of glutamate dehydrogenase From Sulfolobus solfataricus, A thermoacidophilic Archaebacterium—is the presence of N-epsilon-methyllysine related to thermostability, Eur. J. Biochem., 203, 81, 1992. 53. Reits, E.A.J. et al., Dynamics of proteasome distribution in living cells, EMBO J., 16, 6087, 1997. 54. Kruger, E., Kloetzel, P.M. and Enenkel, C., 20S proteasome biogenesis, Biochimie, 83, 289, 2001. 55. Ogiso, Y., Tomida, A. and Tsuruo, T., Nuclear localization of proteasomes participates in stressinducible resistance of solid tumor cells to topoisomerase II-directed drugs, Cancer Res., 62, 5008, 2002. 56. Nederlof, P.M., Wang, H.R. and Baumeister, W., Nuclear localization signals of human and Thermoplasma proteasomal alpha subunits are functional in vitro, Proc. Natl. Acad. Sci. U. S. A., 92, 12060, 1995. 57. Wang, H.R. et al., Import of human and Thermoplasma 20S proteasomes into nuclei of HeLa cells requires functional NLS sequences, Eur. J. Cell Biol., 73, 105, 1997. 58. Mayr, J. et al., The import pathway of human and Thermoplasma 20S proteasomes into HeLa cell nuclei is different from that of classical NLS-bearing proteins, Biol. Chem., 380, 1183, 1999. 59. Heinemeyer, W. et al., The active sites of the eukaryotic 20S proteasome and their involvement in subunit precursor processing, J. Biol. Chem., 272, 25200, 1997. 60. Heinemeyer, W., Ramos, P.C. and Dohmen, R.J., Ubiquitin-proteasome system—The ultimate nanoscale mincer: assembly, structure and active sites of the 20S proteasome core, Cell. Mol. Life Sci., 61, 1562, 2004. 61. Finley, D., Ozkaynak, E. and Varshavsky, A., The Yeast polyubiquitin gene is essential for resistance to high temperatures, starvation, and other stresses, Cell, 48, 1035, 1987.

9214_C020.indd 344

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Archaeal 20S Proteasome: A Simple and Thermostable Model System

345

62. Ciechanover, A., Finley, D. and Varshavsky, A., Ubiquitin dependence of selective protein-degradation demonstrated in the mammalian-cell cycle mutant Ts85, Cell, 37, 57, 1984. 63. Van Den Eynde, B.J. and Morel, S., Differential processing of class-I-restricted epitopes by the standard proteasome and the immunoproteasome, Curr. Opin. Immunol., 13, 147, 2001. 64. Ma, J., Katz, E. and Belote, J.M., Expression of proteasome subunit isoforms during spermatogenesis in Drosophila melanogaster, Insect Mol. Biol., 11, 627, 2002. 65. Knipfer, N. et al., Species variation in ATP-dependent protein degradation: protease profiles differ between mycobacteria and protease functions differ between Mycobacterium smegmatis and Escherichia coli, Gene, 231, 95, 1999. 66. Bauer, M.W., Halio, S.B. and Kelly, R.M., Purification and characterization of a proteasome from the hyperthermophilic archaeon, Pyrococcus furiosus, Appl. Environ. Microbiol., 63, 116, 1997. 67. Michels, P.C. and Clark, D.S., Pressure-enhanced activity and stability of a hyperthermophilic protease from a deep-sea methanogen, Appl. Environ. Microbiol., 63, 3985, 1997. 68. Franzetti, B. et al., Characterization of the proteasome from the extremely halophilic archaeon Haloarcula marismortui, Archaea, 1, 53, 2002. 69. Ruepp, A. et al., Proteasome function is dispensable under normal but not under heat shock conditions in Thermoplasma acidophilum, FEBS Lett., 425, 87, 1998. 70. Reuter, C.J. and Maupin-Furlow, J.A., Analysis of proteasome-dependent proteolysis in Haloferax volcanii cells, using short-lived green fluorescent proteins, Appl. Environ. Microbiol., 70, 7530, 2004. 71. Rawlings, N.D., Morton, F.R. and Barrett, A.J., MEROPS: the peptidase database, Nucleic Acids Res., 34, D270, 2006. 72. Rohlin, L. et al., Heat shock response of Archaeoglobus fulgidus, J. Bacteriol., 187, 6046, 2005. 73. Boonyaratanakornkit, B.B. et al., Transcriptional profiling of the hyperthermophilic methanarchaeon Methanococcus jannaschii in response to lethal heat and non-lethal cold shock, Environ. Microbiol., 7, 789, 2005. 74. Frankenberg, R.J. et al., Chemical denaturation and elevated folding temperatures are required for wild-type activity and stability of recombinant Methanococcus jannaschii 20S proteasome, Protein Sci., 10, 1887, 2001. 75. Fu, H. et al., Molecular organization of the 20S proteasome gene family from Arabidopsis thaliana, Genetics, 149, 677, 1998. 76. Michel, J.K. and Kelly, R.M., Unpublished Data. 77. Tachdjian, S. and Kelly, R.M., Dynamic metabolic adjustments and genome plasticity are implicated in the heat shock response of the extremely thermoacidophilic Archaeon Sulfolobus solfataricus, J. Bacteriol., 188, 4553, 2006. 78. Yamada, S.I. et al., Archaeal proteasomes effectively degrade aggregation-prone proteins and reduce cellular toxicities in mammalian cells, J. Biol. Chem., 2006. 79. Brannigan, J.A. et al., A protein catalytic framework with An N-terminal nucleophile Is capable of self-activation, Nature, 378, 416, 1995. 80. Seemuller, E. et al., Proteasome from Thermoplasma acidophilum—A Threonine protease, Science, 268, 579, 1995. 81. Kisselev, A.F., Songyang, Z. and Goldberg, A.L., Why does threonine, and not serine, function as the active site nucleophile in proteasomes? J. Biol. Chem., 275, 14831, 2000. 82. Zwickl, P. et al. in Protein Folding In The Cell 187–222 (2002). 83. Arendt, C.S. and Hochstrasser, M., Identification of the yeast 20S proteasome catalytic centers and subunit interactions required for active-site formation, Proc. Natl. Acad. Sci. U. S. A., 94, 7156, 1997. 84. Hochstrasser, M., Ubiquitin-dependent protein degradation, Annu. Rev. Genet., 30, 405, 1996. 85. Dick, T.P. et al., Contribution of proteasomal beta-subunits to the cleavage of peptide substrates analyzed with yeast mutants, J. Biol. Chem., 273, 25637, 1998. 86. Nussbaum, A.K. et al., Cleavage motifs of the yeast 20S proteasome beta subunits deduced from digests of enolase 1, Proc. Natl. Acad. Sci. U. S. A., 95, 12504, 1998. 87. Kisselev, A.F., Callard, A. and Goldberg, A.L., Importance of the different proteolytic sites of the proteasome and the efficacy of inhibitors varies with the protein substrate, J. Biol. Chem., 281, 8582, 2006.

9214_C020.indd 345

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346

Thermophiles: Biology and Technology at High Temperatures

88. Rother, M. and Metcalf, W.W., Genetic technologies for Archaea, Curr. Opin. Microbiol., 8, 745, 2005. 89. Allers, T. and Ngo, H.P., Genetic analysis of homologous recombination in Archaea: Haloferax volcanii as a model organism, Biochem. Soc. Trans., 31, 706, 2003. 90. Bitan-Banin, G., Ortenberg, R. and Mevarech, M., Development of a gene knockout system for the halophilic archaeon Haloferax volcanii by use of the pyrE gene, J. Bacteriol., 185, 772, 2003.

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Index A AAA+ ATPase, 57 ABC transporter, 41, 46 Acetyl-CoA, 101, 102, 105, 106, 109 Acetyl-CoA synthetase (ADP forming), 106, 109 Acidianus ambivalens, 142, 193, 194, 198, 225, 227, 230, 232 Acidanus spp., 197 Acyclic tetraether phosphatidylcholine (a-TEPC), 77, 84 Adenylate kinase, 29 Aeropyrum pernix K1, 11, 43, 44, 49, 57, 60, 129, 131–135, 163, 184, 267, 284, 336, 338 Alcohol dehydrogenase (ADH), 12, 29, 62, 133, 135, 151, 190, 196 Aldolase, 101, 103, 104 Alpha-amylase, 26, 27, 33, 34, 120, 124 Alpha-amylase, gene, 200, 221 Alpha-galactosidase, 31 Alpha-glucosidases, 125 Aminotransferase, 30 Ammonification, 162 Ampullaviridae, 230 Amyloid fibrils, 13, 14, 16, 19 Amylomaltases, 127 Anaerobic dissimilatory nitrate reduction, 162 Antibiotic resistance gene, hygromycin, 197 Aquifex aeolicus, 42, 300, 301 Aquifex pyrophilus, 162 Aquifex spp., 11 Arabinose isomerase, 134, 158 Arabinose isomerases, 134, 137 Archaeal histones, 283 Archaeal nucleosomes, 281 Archaeoglobus fulgidus, 43, 57, 219 Archaeoglobus fulgidus FEN-1, 242 cleavage specificity, 242 domain structure, 246 XPB homolog, 245 XPD proposed mechanism, 247 Archaeoglobus fulgidus protein kinases Rio1 and Rio2, 319 Rio 2 winged-helix motif, 320

Archaeoglobus profundus, 196 Archaeols, 74 Aspartate aminotransferase ATPase domains, Rad51 homologs, 253 Autophosphatase activity, 316 Auxotrophy, 214

B Bacillus azotoformans, 162, 166, 167 Bacillus halodenitrificans, 162 Beta-amylases, 124 Beta-Glycosidase, 5, 192 Bipolar tetraether lipids, 78 Biomining, 145 Bleomycin binding protein, 293 Blue copper proteins, 167 BRCA2 mutations BRC repeats, 255 cancer risk, 254 interactions with Rad51, 254

C Caldarchaeol, 74 Calditoglycerocaldarchaeol, 75 cAMP-dependent protein kinase, 318 Capreomycin, 297 Carbonic anhydrase, 35 Carboxydothermus hydrogenoformans Z2901, 300, 301 Carboxyfluorescein fluorescence (5,6-CF), 82 Carboxylesterase, from Pyrobaculum calidifontis, 132 Cdc6 from Sulfolobus acidocaldarius, phosphorylation, 323 Cenarchaeum symbiosum, 281 Channels, 44 Chaperonin, 59, Table 5.2 Chaperonin-Associated Aminopeptidase from Sulfolobus solfataricus, 325 CheA from Thermatoga maritime, 318 Chemoautotroph, 5 Chitinase, 129

347

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348 Chlorate reduction by nitrate reductase, 169 Chromatin architectural proteins, 279, 280 Chromatin proteins, 279, 280, 287 acetylation, 285 “Alba,” 284, 287 HU, 280 Sul7, 285, 287 Sul10A, 285, 287 Chromatin, 280 Chromosome stability, 286 Chromosome structure, 279 CIRCE elements, 58 Circular dichroism (CD) spectroscopy, 19 ClpA, 56 Cold shock protein (CSP), 27, 28 Colwellia psychroerythraea 34H, 300 Compatible solutes, 9, 10 Conjugation, 191–192, 195 bacterial conjugation proteins, 191 cell pairing, 192 Copper, role in nitrite reduction, 169 Corresponding states, 28, 29, 30, 31 Crenarchaeota, 162, 281 Cryo-electron microscopic (cryo-EM) reconstruction, 294, 302 Cyclopentane rings, in archaeal lipids, 75 Cytochromes, in denitrification, 164, 165

Index Directed evolution, 30 DNA compaction, 283, 284 double-stranded, 286 gyrase, 280 local denaturation, 287 N-glycosylases, 208 single-stranded, 286 supercoiling, 280, 284–286 topoisomerases, 219, 279, 280, 286, 344 topology, 283 wrapping, 281, 282, 287 DNA ligases, 244 active site, 244 catalytic cycle, 244 from Sulfolobus solfataricus, 244 DNA Polymerase I-DNA complex, in Thermus spp., 294 DNA polymerases, fidelity and processivity, 240 DNA recombination, 240 DNA repair, 240 pathways, 240 double-strand breaks, 247–249 DNA replication, 240 role of helicases, 240 DNA synthesis, initiation, 240 DNA topoisomerases, 141, 142, 280, 286 DNA, 5′-flap structure , 241 DNA-histone interaction, 282

D D-xylose ketol-isomerase, 134 Deinococcus geothermalis, 300 Deinococcus radiodurans, 292, 300 Deletion, 192 Denitrification, 161–171 enzyme localization, 161 in acidic environments, 163 thermophilic bacteria and archaea, 161 Denitrification enzymes, cellular localization, 166, 168 cofactors, 168 diversity, 171 membrane-bound, 167 Denitrifiers, phylogeny, 162 Desolvation penalty, 56 Desulfurococcus mobilis, 198 Di-myo-inositol phosphate (DIP), 10, 11 Diacylphophatidylcholine, 84 Diether lipids, 74 Differential scanning calorimetry Lipids, 80,81

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E Electron transport, in denitrification, 166 Electrophoretic mobility shift assay (EMSA), 282, 284 Electroporation, 197 Elongation Factor P (EF-P), of Thermus spp., 294 Elongation Factor Tu (EF-Tu) of Thermus spp. Ef-Tu structure, 294 EF-Tu-Ef-Ts complex, 294 EF-Tu-tRNA-GTP ternary complex, 294 Embden-Meyerhof pathway, modified In hyperthermophiles, 103–106 In Sulfolobus spp., 220 Endoglucanases, 115, 116 Enolase, 103, 105, 107, 110, 345 Enthalpy, 34 Escherichia coli RecA protein, 251 Esterases, 132 Euryarchaeota, 102, 104, 109, 142 Exopolygalacturonase, 128 Extremophiles, 3, 73, 114, 120, 146, 150, 163, 279, 310

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349

Index

F 5-Fluoro-orotate (FOA), 6, 214–216 FEN-1 (flap endonuclease 1), 241, 244 conformational changes upon substrate binding, 242 sequence-independent DNA recognition, 242 Ferroglobus placidus, 162 Fluorescence recovery after photobleaching (FRAP), 87 Formyltransferase, of Methanopyrus kandleri, 12 Fourier-tranform infrared (FTIR) spectroscopy, 14 Fructose-1,6-bisphosphatase, 219 Fructose-6-Phosphate phosphorylation, 104

G Gene disruption, 214–219 Genetic markers, 190 Genetic systems, archaea, 190 Genetic transformation, efficiency, 217 Genome sequences, 163 Geobacillus stearothermophilus, 134 Geothermal environments, 280 Global genome DNA repair, 245 Global nitrogen cycle, 162 Glucoamylase, 124, 125 Gluconeogenesis, 100, 104, 109, 110, 219 Glucopyranose, 76 Glucose isomerase, 134 Glucose phosphorylation, 103 Glutamate dehydrogenase (GDH), 12 Glyceraldehyde-3-phosphate dehydrogenase, 21 Glycerol-dialkyl-glycerol-tetraether (GDGT) lipids, 74–81 Glycerol-dialkyl-nonitol-tetraether (GDNT) lipids, 74–86 GroEL/GroES, 59

Heme proteins, in denitrification pathways, 165 Heme-copper oxidases, 170 Hfr element, in Thermus thermophilus, 293 Histidine kinases, 315, 316 Histone fold, 281 Histone pairing, 281 Histones, 280–282 HK853 histidine kinase from Thermatoga maritime, 319 Hofmeister ions, 16 Holliday junction, 255 branch migration, 255, 257 Homologous recombination, 215, 218, 249, 255 archaeal proteins, 250 HpkA histidine kinase from Thermatoga maritime, 318 HPr kinase/phosphatase of Bacillus subtilis, 318 HSP60, 56 HtpX, 57 hybrid proteasome, 338 Hydrogen exchange, 26, 27 Hyperthermophiles, 213, 221 Hyperthermophile-specific, 286

I ibpA, ibpB, 57 Iceland, 192 Ignicoccus spp., 317 Indole glycerol phosphate synthetase (IGPS), 33 Inducible promoters, 197 Intron, 198 Ion-pair networks, ion clusters, 14, 55 Iron-sulfur clusters, in nitrate reductases, 167, 169 Isopropylmalate dehydrogenase, 27 Italy, 195

K

H

Kanamycin adenyltransferase, 31, 293

3-Hydroxy-3-methylglutaryl coenzyme A (HMG-CoA) reductase, 217 Haloarcula valismortui, 162 Haloarcula marismortui, 162, 165–170 Haloferax denitrificans, 162, 167 Haloferax mediterranei, 162, 169 Halomonas sp., 162 Heat shock proteins (HSPs), 56, 57–64 Helical protrusion, 60 Helicases XPB and XPD, 245

L

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lacS gene, 192, 220, 221 Lactate dehydrogenase, 12 Lactate oxidase, 31 Laurdan fluorescence, 84 Ligase chain reaction, 142 Lipase, 30, 132–133 Liposomes 78, 84, 85 Lipothrixviridae, 227

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350

M Macrocyclic-Tetraetherphosphatidylcholine (m-TEPC), 86 Malate dehydrogenase, 12 Mannosylglycerate, 10, 11, 12 Marker genes, 215, 216 Meiothermus, 293 Membrane anchor proteins, nitrate reductases, 167 Membrane fusion, 83 Membrane potential, in nitrate reduction, 169 Metallosphaera sedula TA-2, 75 Methanobacterium formicicum, 281 Methanocaldococcus jannaschii, 41, 219, 220 FEN-1, 242 Methanopyrus kandleri, 43 Lipids, 74 Methanosarcina acetivorans, 59 Methanosarcina thermophila, 46 Methanosarcinacea, 59 Methanospirillum hungatei, 79, 88 Methanothermobacter thermoautotrophicum, 280–282 Methanothermus fervidus, 281 Molecular genetic tools, 190 Molecular motor, DNA packaging, 230 Molecular self-assembly, 225 Molybdopterin cofactor, 167, 169 Monopolar diether liposomes, 83 Morphological transformation, 225 Mre11, Rad50 protein complex, 247 biochemical activities, 247 structural domains, 247

N N-acylamino acid racemase N-methyl-N′-nitro-N-nitrosoguanidine, 142 N2O reductase (Nos) genes, 165 Nanoarchaeum equitans Kin4-M, 43, 56 narM gene, 164 Nascent Associated Complex (NAC), 65, 66 Neutron scattering 14 New Zealand, 195 Nitrate reductase, 161, 162 Nitrate reductase (Nar) genes, 163–165 transcription, 165 Nitrate reductase, catalytic cycle, 168, 169 Nitrate reductase subunits, 163, 167 Nitrate reduction, 100, 102, 166 Nitric oxide reductases, 167, 170 modified hemes, 170 substrate inhibition, 170

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Index Nitrile-degrading enzymes, 138 Nitrite reductase (Nir) genes, 165 Nitrite reductases, 169 Nitrous oxide reductases, 170 copper centers, 170 NO detoxification, 170 NO reductase (Nor) genes, 165 Nonlamellar lipid assemblies, 73 Nuclear localization signals (NLS), 337 Nuclear Overhauser effect, 28 Nucleocapsid, 231 Nucleoid, 280 Nucleoprotein complex, archaeal viruses, 229 Nucleoprotein filaments, 249, 251 Nucleosome, 281 Nucleotide excision repair (NER), 245

O Oceanithermus spp., 292 Okazaki fragment maturation, 243 open kinked DNA, 243 rotary-handoff mechanism, 243 Ornithine carbamoyltransferase, 33 Osmoadaptation, 10

P 6-phospho-3-hexuloisomerase (PHI), 220 Para-nitrobenzyl esterase, 31, 32 Paracoccus sp., 162 PCNA (proliferating cell nuclear antigen), 241, 242 homologs, interactions with replication proteins, 244 Pectinase, 128 Pentose-phosphate pathway, 220 Peptidyl-prolyl cis-trans isomerases, 144 Permease, 41 Persephonella marina, 11 Perylene fluorescence, 80 petB gene, 164 Petrotoga miotherma, 11 PGK (3-phosphoglycerate kinase), 26 Phase transitions (archaeal lipids), 41, 79 phi29 phage, 230 Phosphoacceptor amino acids, 311 Phosphoaspartate (P-Asp), 315 Phosphoenolpyruvate (PEP), 102, 105 Phosphoenolpyruvate (PEP) dikinase, 105 Phosphoenolpyruvate (PEP) synthase, 220 Phosphoenolpyruvate (PEP)-dependent phosphotransferase, 101

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351

Index Phosphoglucose isomerase, 101, 104 Phosphoglucose isomerase (PGI) (Cupin fold), 104 Phosphoglucose/phosphomannose isomerase (PGI/PMI), 103 Phosphoglycerate kinase, 105 Phosphoglycerate mutase (PGM), 105 Phosphohexomutase from Sulfolobus solfataricus, 323 Phosphoproteins from hyperthermophiles, 321 Phosphoribosyl anthranilate isomerase (PRAI), 38 Phosphorylative glyceraldehyde 3-phosphate dehydrogenase (GAPDH), 110 Phosphothioester phosphocysteine intermediate in protein phosphorylation, 312 Phr, Pyrococcus heat-shock regulator, 57 Picrophilus oshimae, 196 Picrophilus torridus, 43, 55 Plasmid archaeal, 190 conjugative, 190–191 cryptic, 193–195 from Pyrococcus and Thermococcus, 196 in Sulfolobus, 190, 193, 195, 198 in thermophilic archaea, 198 in thermophilic archaea, copy number, 192 in Thermotoga, 190 integrated forms, 193 non-conjugative, 195 open reading frames, 192 origin of replication, 192 pBR322 Rom/Rop gene, 196 pNOB8, pING families, 192 pRN family, 193, 195 protein, CopG homolog, 195 pSOG family, 192 pSSVx family, 193 replication, 195 replication proteins, 192 transfer, 192 transformation, 196 vectors for thermophilic archaea, 196 Polaromonas naphthalenivorans CJ2, 300 Polyisoprenoid chains, 74 Polymerase chain reaction (PCR), 4 Polymerization motif, Rad51 homologs, 253 Prefoldin, 61, 62, 63 PrmA methyltransferase, 302 Prokaryotic chromosome, 279 Proteases, 129–132 Proteasome, 20S, 333 Protein, thermal stability, 239 Protein complexes, kinetic trapping, 239

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Protein interactions, reversible, 240 Protein kinase families, 315 Protein-Serine/Threonine Phosphatases of Archaea, 320 Protein-Tyrosine Phosphatases of Archaea, 321 Proton motive force (PMF), 40 Proton-motive force, 161 PTS system, 42 Pullulanases, 126 pyrF gene, 215, 216 Pyrimidine biosynthesis pathway, 214 Pyrobaculum aerophilum, 43, 100 Pyrobaculum Spherical Virus (PSV), 230 Pyrococcus abyssi, 57,190, 195, 197 Pyrococcus furiosus, 43, 196, 219 Pyrococcus furiosus DNA-repair proteins FEN-1241 Mre11, nuclease activity, 248 Rad50 protein, 249 ATPase domains, dimerization, 249 conformational cycle, 249 Rad51 homolog, 251–253 Pyrococcus horikoshii, 43 protein kinase Ph0512, 320 Pyrolobus fumarii, 5 Pyruvate dehydrogenase, 101 Pyruvate ferredoxin oxidoreductase (POR), 101 Pyruvate formate lyase (PFL), 101 Pyruvate kinase, 104, 220 Pyruvate phosphate dikinase, 105

Q Quinols, 167

R Rad50 protein, coiled-coil structure, 248 Rad51 nucleoprotein filament, 253, 254 Rad51 protein, 249 single-strand DNA vs. double-strand DNA binding, 254 structural analysis, 254 Rad51/RadA homologs, 251 comparison to RecA proteins, 251 Rational design, 25, 30 Recombination, 192 Regulator aspartyl-phosphate (RapP) phosphatases, 312 Replication, 192 Reverse DNA gyrase, 218, 219, 222, 286, 287 Rhodococcus erythropoli, 333

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352 Rhodothermus marinus, 11, 115–118, 121, 123, 124, 129–140, 144 Ribonuclease A, 13 Ribonuclease H1, 28, 33 Ribosomal stalk complex, 301 Ribosome release factor (RRF), of Thermus thermophilus, 294 Ribosome, 30S of Thermus thermophilus, 294–300 Complexed with mRNA, 303 Ribosome, 70S of Thermus thermophilus, 294–300 Complexed with tRNAs, 303 Ribulose-5-phosphate, 220 Rieske-type iron-sulfur protein, 164 RNA binding, 285 ROSE, Repression of heat shock gene expression, 57 Rosettasome rrs-rrl intergenic spacer, 296 Rubredoxin, 18 Rudiviridae, 228 RuvA protein, 255 RuvA-RuvB complex, 256, 257 structural features, 256 RuvB protein, 255 from Thermotoga maritima, 255 from Thermus thermophilus, 255

S S-layers, 232 Saccharomyces cerevisiae, 214, 241 Salt bridge, 14 Secondary transporters, 42, 45 Selection for pyrE, pyrF mutants, 214 Self-assembly, 232 Shine-Dalgarno sequence, 296, 300, 301 Shuttle vectors, S. solfataricus/E. coli, 197 Simvastatin, 218 Single-stranded binding protein, 286 Small angle X-ray diffraction of lipids, 79, 80 Small Heat Shock Protein, sHSP, 57 Sparsomycin, 297 Staphylococcal nuclease (SNase), 13 Starch processing enzymes, 120–127 Stealth liposomes, 88 Strain, 121, 5 Streptococcus thermophilus, 43 Streptomycin-resistant, -dependent and -pseudodependent mutants, of Thermus 297

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Index Structural flexibility, of proteins, 26–28 Structure-specific nucleases, 241 STVIIE, 14 Substrate binding protein, 47 Sulfolobus acidocaldarius, 43, 190, 192, 196, 198, 218 Sulfolobus “islandicus,” 192 Sulfolobus shibatae, 134, 197, 284 Sulfolobus solfataricus, 43, 57, 190–197, 213, 220, 280, 319 d-Gluconate Dehydratase, 325, 327 protein kinases SsoPK2 and SsoPK3, 319 RadA protein, 253 Sulfolobus spp., 43, 284 Sulfolobus tokodaii, 193 Supercoils, 283 Symbiobacterium thermophilum, 56 Synechococcus vulcanus 57, 58

T 2-Thioribothymidine, 301 Targeted gene disruption, 221 Tetraether lipids, 41, 74 Thermal stability, 30 Thermo-inducible conformational changes, 226 Thermoacidophiles, 55 Thermoanaerobacter italicus, 128 Thermoanaerobacter mathranii, 134 Thermoanaerobacter tengcongensis, 134, 317, 326, Thermoanaerobacterium saccharolyticum B6A-RI, 134 Thermoanaerobacterium strain JW/SL-YS 489, 134 Thermococcus fumicolans, 142 Thermococcus kodakarensis, 43 Thermococcus KS-1, 58 Thermococcus litoralis, 12 Thermococcus marinus, 142 Thermococcus radiotolerans, 142 Thermophilic denitrifiers, 171 Thermoplasma acidophilum, 43, 88, 60, 296 Thermoplasma sp., DNA repair, 197 Tetraether lipids, 41, 75 Thermoproteus tenax Spherical Virus (TTSV), 231 Thermosolutes, 16, 17 Thermosome, 62, 67 Thermotoga maritima, 27, 42, 196, 300 Thermotoga neopolitana, 134 Thermus aquaticus, 291–301 Thermus flavus AT62, 134 Thermus scotoductus, 142 Thermus thermophilus, 163

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353

Index Thermus thermophilus HB8, 31, 293–302 Thioalkalivibrio spp., 162 Thx protein, 296 Tk-PTP of Thermococcus KOD, 325 Tobacco Mosaic virus, 229 Topoisomerase III, 286 Topoisomerase Type I, 142, 208 Transcription-coupled DNA repair, 245 Transformation efficiency, 198 Transport, of nitrate and nitrite, 166 Triosephosphate isomerase, 30, 101, 103, 104, 107, 110 trpE gene, 215, 216 Truepera spp., 293 Tryptophan auxotrophy, 215 Tryptophan fluorescence quenching, 14 Tryptophan phosphorescence lifetime, 32 Tylosin, 297 Type IV pilin-like signal sequence, 48

U Ubiquitin Associated (UBA) domain, 64 Uracil, 214 Uracil auxotrophs, 215 Uvr ABC exinuclease, 183

V V-type ATPase, 294 Vector-host sytems, 198 Vectors, 197

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Venus fly trap, 47 Virion adsorption, 228 as helical scaffold, 229 functionalization, 229 stability, 229 structure, 226, 231 Viruses from thermophilic archaea, 190, 197 Acidianus bottle-shaped virus, 230 Acidianus filamentous virus (AFV), 227 Acidianus rod-shaped virus (ARV), 228 Acidianus two-tailed virus (ATV), 225 Acidianus virus, claw-like appendage, 228 enveloped, 230, 231 extracellular virion mophogenesis, 226 fuselloviruses, 197, 198 nanotechnology, 225 sequence conservation, 198 SSV family, 190, 193, 197, 231, 232 Sulfolobus islandicus filamentous virus (SIFV), 227 Sulfolobus islandicus rod-shaped virus (SIRV), 228 Sulfolobus shibatae 57 Sulfolobus Turreted Icosahedral Virus (STIV), 230 tail fibers, 231 thermostable, acid-stable, 233 Virus-like particle, 232 Vulcanithermus spp., 293

Z Zipper Virus-like Particles (ZVLP), 232

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FIGURE 5.3 The crystal structure of homodimeric nascent polypeptide associated complex (NAC) from Methanobacterium thermoautotrophicum (from PDB: 1TR8). NAC and ubiquitin-associated domain (UBA) domains are labeled. The figure was drawn with the three-dimensional molecular viewer in the VectorNTI 10.0 package. FIGURE 15.1 Archaeal protein structures prompted the proposal of a rotary-handoff mechanism mediated by proliferating cell nuclear antigen (PCNA) for sequential transition of DNA from DNA polymerase, to flap endonuclease (FEN)-1, and to ligase in Okazaki fragment maturation. SsPCNA (PDB code, 2HII) is presented in three colored surfaces (red, green, and blue). The backbone of DNA strands are in lines. To start DNA synthesis, PCNA is loaded to the 3′-end of a primer by the clamp loader (not shown). Binding of DNA polymerase (gray) to PCNA1 bends the template strand for DNA synthesis. When this complex meets the 5′-end of the adjacent Okazaki fragment, it displaces a short fragment to create a double-flap structure and hands the DNA over to FEN-1 (cyan) bound to PCNA2. FEN-1 cleaves the flapped 5′-fragment and hands over the nicked DNA to ligase (yellow) bound to PCNA3. DNA ligase then covalently connects the two Okazaki fragments together. This mechanism requires the kinked DNA rotates around the three PCNA subunits to interact with different enzymes at different stages of reactions. DNA polymerase, FEN-1, and ligase can bind to PCNA simultaneously as described for SsPCNA. The interactions of DNA with different enzymes are therefore regulated by the flexible interactions between distinct PCNA subunit and each enzyme through conformational changes. In other systems, these three enzymes may bind sequentially to PCNA to fulfill their distinct role during the process. The structure of AfFEN-1:DNA complex (PDB code, 1RXW) is presented in ribbon diagram with the helical clamp highlighted in magenta and DNA in sticks. The structure of SsLig (PDB code, 2HIV) is also presented in ribbon diagram with three domains colored differently. In addition, the structure of human ligase 1:DNA complex (PDB code, 1X9N) is presented in ribbon diagram with dsDNA in gray, and a bound adenosine monophosphate in the sphere at the active center.

FIGURE 15.3 Archaeal protein structures revealed the architecture of the Mre11/Rad50 complex. Center: the Mre11/Rad50 complex assembly formed by heterotetramerization of Mre11/Rad50 (M2R2). Larger complexes 2X(M2R2) observed by negative stain electron microscopy through M2R2 intercomplex hook–hook interactions. Archael structures of the Mre11/Rad50 subcomplexes are highlighted by boxed regions: (a) structure of the Rad50 Zn-hook domain. CXXC motifs coordinate Zn2+ ions to bridge the apices of the Rad50 coiled coils and facilitate long-range DNA tethering; (b) structure of the Mre11 phosphoesterase domain bound Mn2+ and a 5′-adenosine monophosphate (AMP) nucleotide reaction product; (c) structures of ATP bound (top) and apo-Rad50 minimal ATPase domain. Nucleotide binding-induced dimerization of Rad50 ATPase halves within the M2R2 DNA-binding head. Hydrolysis causes dimeric ATPase release and a dramatic conformational twisting to the ATPase-N domain relative to ATPase-C domain.

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FIGURE 16.1 Models for helicase assembly at the Methanothermobacter thermautotrophicus origin of replication.

FIGURE 18.1 Two examples of structural features potentially contributing to thermal stability of the T. thermophilus ribosome, as observed in the high-resolution crystal structure of the 30S subunit [Wimberly et al., 2000]. Ribosomal protein S17 (left) has a β-barrel domain (blue) and a C-terminal α-helix and extended tail (red) that contacts multiple 16S rRNA helices. This extension is generally absent from S17 of mesophiles and psychrophiles. Even among thermophiles, the sequence of the extension is highly variable. Ribosomal protein Thx (right, in green), unique to T. thermophilus (and some plant chloroplasts), contacts multiple secondary structure elements in the head of the 30S subunit. Structures were rendered using MacPyMol 0.99 [DeLano, 2002] and PDB file 1J5E.pdb [Wimberly et al., 2000]. An alignment (bottom) of selected S17 protein sequences obtained from NCBI (http://www.ncbi.nlm.nih.gov). Complete S17 protein sequences were aligned with Clustal W [Thompson et al., 1994], although for clarity only the C-terminal residues are shown. The portion of the T. thermophilus S17 sequence corresponding to the C-terminal α-helix and tail are highlighted in red. The first five sequences belong to thermophilic species, while the next four belong to mesophiles, and the last two belong to psychrophiles. Protein identification numbers are as follows: T. thermophilus HB8, gi|55981652; Thermotoga maritima MSB8, gi|4982055; Aquifex aeolicus VF5, gi|2982774; Carboxydothermus hydrogenoformans Z-2901, gi|94730505; Deinococcus geothermalis DSM 11300, gi|94985958; Deinococcus radiodurans R1, gi|6457994; Blastopirellula marina DSM 3645, gi|87306529; Escherichia coli W3110, gi|85676730; Bacillus subtilis subsp. subtilis str. 168, gi|16077193; Colwellia psychrerythraea, 34H gi|71147715; Polaromonas naphthalenivorans CJ2, gi|84711025.

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