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.

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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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27

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 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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Thermophilic Protein-Folding Systems

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.

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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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79

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 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 (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

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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.

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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

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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Thermophiles: Biology and Technology at High Temperatures

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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125

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

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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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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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Thermophiles: Biology and Technology at High Temperatures

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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Thermophiles: Biology and Technology at High Temperatures

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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148

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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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Thermophiles: Biology and Technology at High Temperatures

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.

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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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Thermophiles: Biology and Technology at High Temperatures (a)

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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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).

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

179 180 180 181 182 183 184 184 185

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

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tio ele

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pING1/p KEF

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4

/VirD

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ase plrA integ r

Tr

bE

NG

/V

de

irB

4

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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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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.

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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.

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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.

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

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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 (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.

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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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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

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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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