ADVANCES IN PROTEIN CHEMISTRY Volume 69 DNA Repair and Replication
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ADVANCES IN PROTEIN CHEMISTRY EDITED BY FREDERIC M. RICHARDS
DAVID S. EISENBERG
Department of Molecular Biophysics and Biochemistry, Yale University New Haven, Connecticut
Department of Chemistry and Biochemistry Center for Genomics and Proteomics University of California, Los Angeles Los Angeles, California
JOHN KURIYAN Department of Molecular and Cellular Biology University of California, Berkeley Berkeley, California
VOLUME 69
DNA Repair and Replication EDITED BY Wei Yang Section Chief of Structure and Mechanism, Laboratory of Molecular Biology, NIDDK, NIH, Bethesda, Maryland
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CONTENTS PREFACE . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
ix
Base Excision Repair J. Christopher Fromme and Gregory L. Verdine
I. II. III. IV. V. VI.
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Types of Damage Repaired by BER . . . . . . . . . . . . . . . . . . . . . . DNA Glycosylases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Downstream BER Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mammalian BER . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Roles of BER Enzymes in Other Processes . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
1 5 7 18 23 27 28
Nucleotide Excision Repair in E. Coli and Man Aziz Sancar and Joyce T. Reardon
I. II. III. IV. V. VI.
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Damage Recognition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mechanism of Excision Repair . . . . . . . . . . . . . . . . . . . . . . . . . . . Transcription-Coupled Repair . . . . . . . . . . . . . . . . . . . . . . . . . . . Repair of Chromatin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusion. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
43 45 48 56 59 63 65
Photolyase and Cryptochrome Blue-Light Photoreceptors Aziz Sancar
I. II. III. IV. V. VI.
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Phylogenetics. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Structure of Photolyase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Reaction Mechanism of Photolyase . . . . . . . . . . . . . . . . . . . . . . . (6–4) Photolyase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cryptochrome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . v
73 74 77 79 86 90
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VII. Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
96 96
Coordination of Repair, Checkpoint, and Cell Death Responses to DNA Damage Jean Y. J. Wang and Sarah K. Cho
I. Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Overview of Biological Responses to DNA Damage. . . . . . . . . . III. Molecular Components for the Initiation of DNA Damage Responses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Apoptotic Effectors in DNA Damage Response . . . . . . . . . . . . . V. DNA Repair Proteins in Damage Signaling . . . . . . . . . . . . . . . . VI. Alternative Models for the Temporal Coordination of DNA Damage Responses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VII. Future Prospects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
101 104 109 115 120 123 127 128
Functions of DNA Polymerases Katarzyna Bebenek and Thomas A. Kunkel
I. II. III. IV. V. VI. VII. VIII. IX. X. XI. XII. XIII. XIV.
DNA Polymerase Families . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Structures and Compositions of DNA Polymerases . . . . . . . . . . Functions of DNA Polymerases . . . . . . . . . . . . . . . . . . . . . . . . . . . Polymerases for DNA Repair . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Polymerases for Replicating Undamaged DNA . . . . . . . . . . . . . Polymerases for Sister Chromatid Cohesion. . . . . . . . . . . . . . . . Mitochondrial DNA Replication . . . . . . . . . . . . . . . . . . . . . . . . . . Polymerases for Replicating Damaged DNA . . . . . . . . . . . . . . . . Polymerases and Cell-Cycle Checkpoints. . . . . . . . . . . . . . . . . . . Polymerases for Replication Restart and Homologous Recombination . . . . . . . . . . . . . . . . . . . . . . . . . . . . Polymerases for DNA Mismatch Repair . . . . . . . . . . . . . . . . . . . . Polymerases in the Development of the Immune System . . . . Biological Consequences of Polymerase Dysfunction . . . . . . . . Closing Comments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
137 139 141 141 150 151 152 152 155 155 156 156 157 158 159
CONTENTS
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Cellular Functions of DNA Polymerase and Rev1 Protein Christopher W. Lawrence
I. II. III. IV.
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Enzymological Studies With Pol and Rev1p . . . . . . . . . . . . . . . Genetic Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Processes Other than General Translesion Replication that Employ Pol and Rev1p. . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Regulation of Pol and Rev1p and Interactions with other Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VI. Conclusions and Speculations . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
167 172 178 182 186 190 195
DNA Polymerases and Alexandra Vaisman, Alan R. Lehmann, and Roger Woodgate
I. II. III. IV. V. VI. VII. VIII.
Historical Perspective. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Identification of RAD30 and its Orthologs . . . . . . . . . . . . . . . . Biochemical Properties of Pol and Pol . . . . . . . . . . . . . . . . . . Translesion Synthesis by Pol and Pol. . . . . . . . . . . . . . . . . . . . Structure of the Catalytic Core of S. cerevisiae Pol . . . . . . . . . . Regulation and Localization of Pol and Pol . . . . . . . . . . . . . . Mutations in Pol in XP Variants. . . . . . . . . . . . . . . . . . . . . . . . . Pols and and the Polymerase Switch: Interactions with PCNA and Rev1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IX. Protection from Cellular Effects of DNA Damage . . . . . . . . . . X. Roles of Pol and Pol in Somatic Hypermutation . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
205 206 207 210 212 215 217 219 220 221 222
Properties and Functions of Escherichia Coli : Pol IV and Pol V Robert P. Fuchs, Shingo Fujii, and JØro^ me Wagner
I. DNA Pol IV, the dinB Gene Product . . . . . . . . . . . . . . . . . . . . . . II. DNA Polymerase V, the umuDC Gene Product . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
230 248 257
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Mammalian Pol : Regulation of Its Expression and Lesion Substrates Haruo Ohmori, Eiji Ohashi, and Tomoo Ogi
I. Structures of the Genes and Proteins . . . . . . . . . . . . . . . . . . . . . II. Enzymatic Properties of Pol . . . . . . . . . . . . . . . . . . . . . . . . . . . . III. Possible Mechanisms of TLS by Pol In Vivo . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
265 271 274 275
DNA Postreplication Repair Modulated by Ubiquitination and Sumoylation Landon Pastushok and Wei Xiao
I. II. III. IV. V. VI. VII. VIII. IX.
Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . DNA Postreplication Repair Prokaryotes. . . . . . . . . . . . . . . . . . . DNA Postreplication Repair in Eukaryotes . . . . . . . . . . . . . . . . . Ubiquitination . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Protein Conjugation in PRR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Postreplication Repair via Covalent Modifications of PCNA . . Functional Conservation of Eukaryotic Postreplication Repair Future Directions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
279 280 281 283 286 292 295 297 300 301
Somatic Hypermutation: A Mutational Panacea Brigette Tippin, Phuong Pham, Ronda Bransteitter, and Myron F. Goodman
I. II. III. IV.
Generation of Antibody Diversity . . . . . . . . . . . . . . . . . . . . . . . . . Somatic Hypermutation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Apobec Protein Family . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Biochemical Perspective. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
307 313 323 326 327
AUTHOR INDEX. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
337 369
SUBJECT INDEX . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
PREFACE The topic of DNA repair, now firmly embedded in the larger field of biological responses to DNA damage, is in an extraordinarily dynamic state at this time. In particular, productive inroads are being made into the central issue of how cells sense the presence of DNA damage and initiate specific signals in response to genomic insult of one type or another. The term ‘‘genomic insult’’ is deliberately used in deference to the more familiar term ‘‘DNA damage’’ because it is becoming increasingly apparent that the primary event that initiates biological responses to genomic insult may be arrested DNA replication rather than DNA damage itself. This volume, entitled DNA Repair and Replication, is thus timely and appropriately focused. The book comprises eleven chapters contributed by experts in the field. It opens with discussions of the primary modes of the repair of DNA base damage in the strict biochemical sense, namely, base excision repair (BER), enzymatic photoreactivation (EPR) of both cyclobutane pyrimidine dimers and [6–4] photoproducts, nucleotide excision repair (NER), mismatch repair (MMR), and the repair of DNA double-strand breaks (DSBR). BER comprises a fundamental group of biochemical reactions that notably processes many types of spontaneous base damage to DNA, especially the damage generated by the pervasive reactive oxygen species that continually swamp the intracellular milieu. Fromme and Verdine provide a comprehensive discussion of these reactions, with an emphasis on BER in mammalian cells. They also discuss the possible novel roles of BER proteins in sensing damage and initiating checkpoint responses. The chapter on NER contributed by Aziz Sancar and Joyce Reardon provide a historical introduction to the biological response to DNA damage that is evoked primarily by environmental reagents reactive with DNA and is now known to comprise both transcription-coupled and transcription-independent pathways. Its links to human hereditary diseases are also discussed, as is the important and still murky area of chromatin remodeling during NER. This phenomenon has long been intuitively considered indispensable for the process, since the NER machinery is physically massive once fully assembled, and it is not obvious how it could access sites of base damage without some sort of chromatin modification. Aziz Sancar has provided a comprehensive survey of EPR including the phylogenetic relationship between DNA photolyases and other light-absorbing proteins, notably those involved in circadian rhythm. Unfortunately, this volume was not able to include chapters on other ix
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PREFACE
aspects of DNA repair, such as mismatch repair and the repair of DNA strand breaks. Jean Wang and Sarah Cho have effectively tackled the formidable task of attempting to integrate the coordination of DNA damage sensing and the elaboration of checkpoint signals, a rapidly evolving and complex area of contemporary research in biological responsiveness to genomic insult. Sustained arrested DNA replication is lethal to cells. Both prokaryotic and eukaryotic cells react to this threat with a series of diverse biological responses that are designed to relieve the arrest while tolerating the presence of the offending damage. Such damage can presumably be repaired once the replicative crisis is resolved, although it remains unclear why lesions at sites of replicative arrest are apparently refractory to repair. Some of these responses promote the resumption of high-fidelity semiconservative DNA synthesis without incurring mutations. Others, presumably employed as a last-ditch response to arrested replication, support DNA synthesis across sites of template DNA damage with a high probability of mutation. Indeed, this phenomenon may constitute a primary source of spontaneous mutagenesis in many organisms. Gratifyingly, half of this volume is dedicated to a discussion of these so-called DNA damage–tolerance mechanisms, for much remains to be learned about how cells determine which of the multiple responses to arrested DNA replication to activate, and in what order. Our understanding of error-prone (mutagenic) responses to arrested DNA replication has experienced a flowering in the last 5 years because of the discovery that prokaryotes, and especially eukaryotes, are endowed with multiple specialized DNA polymerases that are able to effect extension of DNA primer strands across sites of damage that cause the arrest of high-fidelity DNA synthesis. It is now well appreciated that the fundamental property of these specialized DNA polymerases that promotes their ability to support so-called translesion DNA synthesis is a dramatically reduced fidelity for nucleotide incorporation. This property necessarily carries the risk of mutational catastrophe if such enzymes gain access to stretches of undamaged DNA, and much remains to be learned about how the regulation of such access is controlled in cells. A series of chapters by Katarzyna Bebenek and Thomas Kunkel; Christopher Lawrence; Alexandra Vaisman, Alan Lehmann, and Roger Woodgate; Robert Fuchs, Shingo Fujii and Je´roˆme Wagner; Haruo Ohmori, Eiji Ohashi, and Tomoo Ogi present current information on the plethora of specialized polymerases in prokaryotes and eukaryotes. These chapters include intriguing hints how critical switching events between high-fidelity and low-fidelity polymerases may transpire during translesion DNA synthesis.
PREFACE
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Error-free mechanisms for DNA damage tolerance have been variously dubbed postreplication repair, postreplicative recombinational repair, repair by gap-filling, and replication fork regression, to name a few terms. Landon Pastushok and Wei Xiao have provided a useful summary of the important regulatory roles of specific types of posttranslational modification of certain proteins, notably monoubiquitination and sumoylation, in these biological responses, with an appropriate focus on the yeast Saccharomyces cerevisiae. The enormous utility of the genetic versatility of this lower eukaryotic is once again becoming a boon to the field. In recent years, studies from several laboratories have generated suggestive solutions to the long-standing mystery of how somatic hypermutation transpires in immunoglobulin genes during antibody maturation. Intriguingly, various familiar players on the DNA repair stage appear to be critical, including the specialized DNA polymerases just mentioned. In this volume, Myron Goodman and his colleagues have provided a cogent summary of class switch recombination and somatic hypermutation that incorporates the known role of activation-induced cytosine deaminase (AID), as well as the possible roles of error-prone DNA polymerases, base excision repair, and mismatch repair. The field of biological responses to DNA damage is grossly underserved with contemporary texts that address current problems and perspectives. Wei Yang and her publisher, Elsevier, are to be congratulated on their timely elaboration of this comprehensive volume. This collection of reviews will serve well not only the DNA repair community, but also students of DNA metabolism and cell regulation in general. Errol C. Friedberg Laboratory of Molecular Pathology Department of Pathology University of Texas Southwestern Medical Center Dallas, TX 75390–9072
BASE EXCISION REPAIR By J. CHRISTOPHER FROMME* AND GREGORY L. VERDINE*,À *Department of Molecular and Cellular Biology, Harvard University, Cambridge, Massachusetts, 02138 ÀDepartment of Chemistry and Chemical Biology, Harvard University, Cambridge, Massachusetts, 02138
I. Introduction . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . II. Types of Damage Repaired by BER . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . III. DNA Glycosylases. . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Mechanistic Classes . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. Distinct Structural Classes. .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . C. Damage Recognition and Searching . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . D. Catalysis . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . IV. Downstream BER Enzymes . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. AP Endonucleases. . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. Short-Patch versus Long-Patch Repair . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . V. Mammalian BER . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Additional Components . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. Knockout Mice and the Role of BER Proteins in Human Disease . . . .. . . . . . VI. Roles of BER Enzymes in Other Processes . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. BER-Like Enzymes in Plant Development . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. Thymine DNA Glycosylase .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . .
1 5 7 7 9 10 16 18 18 19 23 23 24 27 27 28 28
I. Introduction When Marshall McLuhan coined the phrase ‘‘the medium is the message,’’ he could not have imagined just how perfectly his shrewd commentary on pop culture would summarize the entire molecular underpinnings of genetics. The medium of heredity—the covalent structure of the nucleobases in DNA and RNA—directly provides the information content for all genetic transactions: Change the covalent structure, change the biologic meaning. It thus comes as no surprise that living systems expend considerable energy in an ongoing struggle against spontaneous genetic change. Uncovering the origins of such change and the nature of the defense against it is of broad significance for understanding issues ranging from evolution to carcinogenesis. The most wide-ranging threat to genetic integrity is posed by the attack of environmental agents on DNA nucleobases, resulting in their spontaneous covalent modification (Lindahl, 1993). Nucleobases are subject to the attack of oxidants, alkylating agents, ultraviolet light, and other forms of electromagnetic radiation; even the solvent water that endows DNA with 1 ADVANCES IN PROTEIN CHEMISTRY, Vol. 69
Copyright 2004, Elsevier Inc. All rights reserved. 0065-3233/04 $35.00
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FROMME AND VERDINE
its miraculous biologic properties can cause hydrolytic damage to DNA bases. Nearly all of the damaged bases resulting from chemical modification, often referred to as ‘‘lesions,’’ interfere with the template function of DNA, thereby giving rise to point mutations (Lindahl, 1993) or aberrant transcripts (Bregeon et al., 2003). In certain cases, the lesions possess unstable glycosidic bonds and therefore undergo further spontaneous conversion to abasic sites (Stivers and Jiang, 2003), which are themselves highly genotoxic (Loeb and Preston, 1986). The negative selective pressure imposed by lesion nucleobases has driven the evolution of repair pathways dedicated to recognition and removal of base lesions, followed by restoration of the original DNA sequence(Lindahl, 1993). The major pathway for this type of repair (Fig. 1) is initiated by the excision of a damaged base and is therefore known as base excision DNA repair (BER). Another repair pathway, nucleotide excision repair (NER), can also repair some lesion nucleobases that severely distort the DNA helix. The NER pathway is discussed in Chapter 2. The enzymes responsible for catalyzing the initiation of BER are designated as DNA glycosylases, because they catalyze cleavage of the glycosidic bond linking a lesion nucleobase to the DNA backbone. DNA glycosylases are specialists, each recognizing one lesion, or at most a small subset of lesions; most organisms express multiple enzymes to gain broad protection from various kinds of routinely encountered genetic insults. Locating these damaged bases embedded in a more than million-fold excess of normal DNA represents a formidable biologic version of the ‘‘needle in the haystack’’ problem. DNA glycosylases channel the chemically diverse universe of lesion bases into a small number of products suitable for processing by downstream enzymes in the BER pathway. DNA glycosylases fall into two mechanistic classes: monofunctional DNA glycosylases displace the lesion base using a molecule of solvent water, whereas bifunctional DNA glycosylases (glycosylase/lyases) displace the lesion base by using a nucleophilic active site residue on the protein, which is always an amine (Fig. 2). The covalently linked enzyme/DNA intermediate thus formed undergoes a cascade of further reactions, ultimately resulting in cleavage of the DNA backbone on the 30 -side of the lesion base. After a DNA glycosylase excises a lesion base, the resulting product is further processed by an AP (apurinic/apyrimidinic) endonuclease, which nicks the DNA backbone on the 50 side of the lesion base. When an AP endonuclease acts on an abasic site generated by a monofunctional DNA glycosylase, the product is a 50 -deoxyribosephosphate (50 -dRP) group (Fig. 3). This species is usually removed by DNA polymerase , the repair polymerase. However, an alternative pathway is sometimes used in which DNA polymerase /", together with PCNA and FEN-1, displaces the strand
BASE EXCISION REPAIR
3
Fig. 1. Base excision repair pathway for repair of DNA base damage. Base damage is indicated by the color red and an asterisk. The key enzymatic activities are indicated at each step, but some steps require proteins not indicated. ‘‘Pol’’ is an abbreviation for ‘‘polymerase.’’ (See Color Insert.)
on the 30 side of the nick. This ‘‘long-patch’’ repair process is thought to be important for replication-associated repair of base damage. In contrast, when AP endonuclease acts on the product generated by a bifunctional glycosylase, the entire lesion nucleoside is removed, and repair polymerization can proceed directly. Finally, DNA ligase covalently reseals the DNA backbone.
4 FROMME AND VERDINE
Fig. 2. Mechanistic scheme of the reactions catalyzed by DNA glycosylases. DNA glycosylases must first recognize and bind to nucleobase damage (1). The monofunctional DNA glycosylases, such as uracil DNA glycolase and MutY, then catalyze one-step removal of the lesion base to generate an abasic site (4). Bifunctional DNA glycosylases, such as hOGG1, MutM, and EndoIII, catalyze additional reactions via attachment of an active-site nucleophilic moiety (Nu) (2), resulting in 30 - (3, in the case of glycosylases with a primary amine nucleophile) or 30 - and 50 - (6, in the case of glycosylases with a secondary amine nucleophile) nicking of the DNA strand. A characteristic of the Schiff base intermediate (2) is that it can be intercepted by the reducing agent sodium borohydride (NaBH4) to generate a stable covalent enzyme-DNA complex (5). This figure is reprinted with permission (Fromme et al., 2004b).
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Fig. 3. Reactions catalyzed by apurinic/apyrimidinic (AP) endonucleases. An AP endonuclease will generate a 50 -dRP group and a nicked strand when acting on an abasic site. Alternatively, a single-nucleotide gap will be created if the substrate is the -elimination product of a bifunctional DNA glycosylase.
II. Types of Damage Repaired by BER As mentioned above, a wide variety of damaged DNA bases result from the attack of various environmental agents (Table I). The enormous molar excess of water (55 M) over the genome provides driving force for water to cause hydrolytic deamination of amine-containing DNA bases. The
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Table I Substrates of Base Excision Repair Source of damage
Example lesion
Responsible DNA glycosylases
Water/alkaline Oxidation Alkylation Ultraviolet light
Uracil 8-oxoguanine 3-methyladenine Thymine dimer
UDG, MUG, SMUG OGG1, MutM MagI, AlkA, MagIII, AAG T4 endonuclease V
predominant target of this reaction is cytosine, which undergoes conversion to uracil (Lindahl, 1993). Uracil residues in DNA are an efficient substrate for BER catalyzed by uracil DNA glycosylases, of which several distinct forms are known. Interestingly, hydrolytic deamination of 5-methylcytosine generates the normal base thymine, but this is recognized as a BER substrate by its being paired opposite G instead of A. Hydrolytic deamination of adenine generates hypoxanthine, another BER substrate (Stivers and Jiang, 2003). The by-products of cellular respiration include several ferociously reactive oxygen species (ROS) such as hydroxyl radical, superoxide radical, and hydrogen peroxide. ROS are also found at high levels in cigarette smoke. ROS can react with many components of the cell, including DNA bases. Of the four canonical DNA bases, guanine is the most sensitive to oxidation (Burrows and Muller, 1998). The oxidation product, 8-oxoguanine (oxoG), is quite stable in DNA (Cullis et al., 1996) and is a potent mutagen owing to its overwhelming preference for mispairing with adenine during DNA replication (Grollman and Moriya, 1993; Shibutani et al., 1991). BER is the primary pathway for repair of oxoG lesions. Other known oxidative lesions include thymine glycol, 5-hydroxycytosine, and formamidopyrimidine (fapy) (Lu et al., 2001). Many exogenous mutagens are alkylation agents that express their toxicity through covalent attachment to DNA bases. Alkylation of a DNA base can lead to mispairing or result in a lesion base with a particularly weak glycosidic bond, increasing the likelihood of depurination. There are several known alkylation-damage products, including 7-methylguanine and 3-methyladenine, which are repaired by BER (Wyatt et al., 1999). Some alkylated bases are repaired by systems other than BER; for instance, the alkylation product O6-methylguanine is repaired by direct reversion (Foote et al., 1980; Myrnes et al., 1982; Olsson and Lindahl, 1980). Recently, a novel dioxygenase mechanism for direct reversion of alkylated bases has been observed in catalysis by AlkB (Sedgwick, 2004). The most well-known form of ultraviolet light–induced base damage is the pyrimidine dimer. Although the hallmark repair system for this type of
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DNA lesion is NER, the BER response is also known to process cyclobutane thymine dimers (Seawell et al., 1980).
III. DNA Glycosylases A. Mechanistic Classes There are two distinct mechanistic classes of DNA glycosylases. The monofunctional glycosylases remove lesion bases from the DNA backbone by cleavage of the glycosidic bond, displacing the lesion base using hydroxide derived from water. The product of this reaction is an abasic site (Dodson et al., 1994; Sun et al., 1995) (Fig. 2, structure 4), which is the primary substrate for AP endonuclease (Mol et al., 2000a; Wilson and Barsky, 2001). Bifunctional glycosylases remove lesion bases via displacement with a nucleophilic active-site residue (Dodson et al., 1994; Sun et al., 1995; Weiss and Grossman, 1987). Dependent on the structural family of bifunctional glycosylase, the nucleophile is the primary amine of an internal lysine residue (Dodson et al., 1994; Kuo et al., 1992; Nash et al., 1997; Sun et al., 1995; Thayer et al., 1995), the secondary amine of an Nterminal proline residue (Tchou and Grollman, 1995; Zharkov et al., 1997), or the primary amine of a N-terminal threonine residue (Dodson et al., 1993). Bifunctional glycosylases, by virtue of the covalent attachment they form with the lesion nucleoside (Fig. 2, structure 2), catalyze additional transformation of the substrate DNA, resulting in a nicked phosphate backbone (Dodson et al., 1994). They also perform this strand– nicking reaction (sometimes referred to as ‘‘AP lyase’’ activity) on abasic site substrates (McCullough et al., 2001). Depending on the nature of the active-site amine, the product of strand nicking may be singly (glycosylases with a primary amine nucleophile) or doubly (glycosylases with a secondary amine nucleophile) nicked (Fig. 2, structures 3 and 6). The strand nicking occurs via a -elimination (Bailly and Verly, 1987; Kow and Wallace, 1987; Mazumder et al., 1991) (singly nicked) or , -elimination (doubly nicked) mechanism (Bhagwat and Gerlt, 1996). An experimentally useful feature of the bifunctional mechanism is the capability of reducing agent (i.e., NaBH4) to intercept the covalent enzyme-DNA intermediate (Dodson et al., 1994; Nash et al., 1996; Sun et al., 1995). The result of reduction is a covalent single bond between the glycosylase and substrate DNA that is quite stable (Fig. 2, structure 5). This ‘‘borohydride trapping’’ procedure has been useful in the isolation (Bruner et al., 1998; Nash et al., 1996; Piersen et al., 1995) and characterization (Girard et al., 1997; Ikeda et al., 1998; Nash et al., 1997; Sidorkina
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Fig. 4. Structures of DNA glycosylase-DNA complexes, representing the five known structural classes of DNA glycosylase. In each panel DNA is shown as gold sticks with the
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and Laval, 2000; Sun et al., 1995; Zharkov et al., 1997) of numerous bifunctional DNA glycosylases. DNA glycosylase catalytic mechanisms are discussed in further detail below.
B. Distinct Structural Classes There are several different structural classes of DNA glycosylases (Fig. 4), but the boundaries of these classes do not strictly coincide with those of the mechanistic classes. The UDG structural family includes the DNA glycosylases (UDG or UNG), MUG, TDG, and SMUG, all monofunctional glycosylases. These proteins contain a single domain and recognize uracil or thymine mismatches in DNA. The MutM structural family includes the two-domain bifunctional glycosylases MutM (also known as 8-oxoguanine glycosylase or Fpg) and endonuclease VIII (also known as Nei), which repair various types of oxidative damage. Until recently it was believed that MutM family members were found only in bacteria. This view changed with the relatively recent discovery of several mammalian homologs, referred to as the NEIL (Nei-like) proteins (Bandaru et al., 2002; Hazra et al., 2003; Morland et al., 2002; Takao et al., 2002a). AAG (also known as ANPG), the only known member of its own structural class, is a single-domain eukaryotic monofunctional glycosylase that repairs alkylation damage (Engelward et al., 1993). Endonuclease V from the T4 virus, which also has no known structural relatives, is a small (16-kD) bifunctional glycosylase that repairs ultraviolet-induced thymine dimers (Dodson et al., 1993). The largest and most functionally diverse structural family of DNA glycosylases is the HhH-GPD superfamily (Nash et al., 1996; Thayer et al., 1995). The family includes both monofunctional and bifunctional DNA glycosylases, and members recognize a variety of lesions arising from oxidative, alkylation, and hydrolytic damage. Some members of both the MutM and HhH-GPD families contain structural metal ions that play no direct role in catalysis.
damage base(s) in red, and glycosylases are shown in ribbon format. (A) T4 endonuclease V bound to DNA containing a thymine dimer (Vassylyev et al., 1995). (B) Uracil DNA glycosylase bound to pseudouridine-containing DNA (Parikh et al., 2000). (C) Human 8-oxoguanine glycosylase 1 bound to DNA containing 8-oxoguanine (Bruner et al., 2000). (D) Human alkyladenine glycosylase bound to DNA containing ethenoadenine (Lau et al., 2000). (E) MutM (bacterial 8-oxoguanine glycosylase) bound to DNA containing 8-oxoguanine (Fromme and Verdine, 2003a). (See Color Insert.)
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C. Damage Recognition and Searching One of the major challenges facing DNA glycosylases is locating damaged bases embedded amid an overwhelmingly larger number of normal bases. It is undesirable for DNA glycosylases to act on undamaged DNA, if only because of the potential toxicity of their abasic-site products (discussed later). To understand how glycosylases search for damage, we must first understand the basis for recognition of damage. A particularly informative avenue for revealing the atomic basis of damage selection is structural biology. There has been a recent windfall of structural data addressing the issue of damage recognition by DNA glycosylases (Fromme et al., 2004b). The structure of a recognition complex, in which a DNA glycosylase is bound to duplex DNA containing a lesion base, can be especially informative. The caveat associated with any recognition complex is that some perturbation is necessary to obtain the structure, either a mutation of the enzyme or the use of a noncleavable substrate analog, so that one is never viewing a true precatalytic state. However, if the perturbations are subtle enough, it can be assumed that the structure obtained is a reasonable facsimile of the true natural state.
1. Damage Recognition Lesion base-containing recognition complexes for many DNA glycosylases have now been structurally characterized (Table II). These include uracil-DNA glycosylase (UDG), representing the first structure of a eukaryotic DNA glycosylase-DNA complex (Parikh et al., 1998; Slupphaug et al., 1996) and MUG (Barrett et al., 1998, 1999), both members of the same structural family. These enzymes remove uracil from DNA, and MUG will also remove thymine when paired opposite guanine. MUG recognizes its substrate primarily by interacting with the guanine paired opposite the substrate base, and it exhibits little selectivity within the active site. UDG does not make any specific contacts with the guanine base and only makes direct contact with uracil in the product complex. The authors suggest that UDG substrate discrimination is mainly a result of the instability of U:G pairs. Extensive work has been performed on the structural biology of UDG, often with the goal of elucidating the catalytic mechanism (see below). Structures of the uracil glycosylase SMUG1 in complex with DNA have recently been obtained (Wibley et al., 2003). Unfortunately, it was not possible to obtain a true recognition complex with SMUG1, so the details of uracil recognition by that enzyme remain to be determined. Recognition complexes are also available for T4 endonuclease V (Vassylyev et al., 1995), hAAG (Lau et al., 2000), MutM (Fromme and Verdine, 2003a), and the HhH-GPD glycosylases endonuclease III ( J. C. Fromme
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Table II Structurally Characterized DNA Glycosylase-DNA Complexes Glycosylase
Mechanistic class
Structural class
AAG
Monofunctional
AAG
AlkA EndoIII
Monofunctional Bifunctional
HhH-GPD HhH-GPD
EndoVIII OGG1
Bifunctional Bifunctional
MutM HhH-GPD
MUG
Monofunctional
UDG
MutM
Bifunctional
MutM
MutY
Monofunctional
HhH-GPD
SMUG1 T4 EndoV UDG
Monofunctional Bifunctional Monofunctional
UDG T4 EndoV UDG
DNA-bound structures available Recognition, abasic analog, and mutant complexes Abasic analog complex Recognition and trapped intermediate complexes Trapped intermediate complex Recognition, trapped intermediate, and mutant complexes Product, substrate analog, and nonspecific complexes Recognition, trapped intermediate, product, and abasic analog complexes Recognition and product-like complexes Product and nonspecific complexes Recognition complex Product, substrate analog, and mutant complexes
and G. L. Verdine, unpublished), hOGG1 (Bruner et al., 2000), and MutY (Fromme et al., 2004a). T4 endonuclease V, the first DNA glycosylase for which the structure of a lesion-containing DNA complex was determined, is unique among DNA glycosylases by virtue of its intrahelical recognition and removal mode. Whereas all other DNA glycosylases extrude their substrate base into an extrahelical active site pocket, T4 endonuclease V instead extrudes one of the two adenine bases paired opposite the thymine-dimer substrate to gain access to the lesion. On the basis of a structural study, AAG was thought to select for its diverse array of alkylated substrates in DNA by sensing their electron deficiency via stacking with aromatic residues (Lau et al., 2000). More recently, it has been suggested that AAG recognizes its substrates, including the deamination product hypoxanthine, principally by excluding normal bases (O’Brien and Ellenberger, 2003). AAG has little discrimination for the identity of the base paired opposite a lesion, and consequently makes sparse contact with this base. The two distinct oxoG-glycosylases, MutM (found in prokaryotes) and OGG1 (found in eukaryotes), recognize the oxoG:C pair in DNA somewhat differently (Fig. 5). OGG1 binds to oxoG in the anti glycosidic
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Fig. 5. Comparison of 8-oxoguanine binding to the active sites of two different 8-oxoguanine glycosylases. The oxoG base is shown in red, DNA in gold, and enzyme residues in cyan. (A) hOGG1 active site (Bruner et al., 2000). (B) MutM active site (Fromme and Verdine, 2003a). Both panels used with permission. (See Color Insert.)
conformation, makes contact with the Watson–Crick face of oxoG using a glutamine sidechain, and recognizes the oxidation product by hydrogen bonding to the N7 proton of oxoG with the main-chain carbonyl oxygen of a glycine residue (Bruner et al., 2000). MutM binds to oxoG in the syn glycosidic conformation, makes contacts with the Watson–Crick face using
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threonine and glutamate sidechains, and recognizes the oxidation status of oxoG via hydrogen bonding to the N7 position with the main-chain carbonyl oxygen of a serine residue (Fromme and Verdine, 2003a). The strategy for cytosine recognition differs between the two enzymes, with OGG1 employing two arginine residues and MutM using a single arginine residue to contact the Watson–Crick face of cytosine. Substrates of the structurally related enzymes endonuclease III and MutY share one aspect in common: the base paired opposite the lesion has a guanine-like Watson–Crick face. In the case of endonuclease III, the preferred substrates are oxidized pyrimidines when paired opposite guanine (though lesions paired opposite adenine are also processed). In the case of MutY, the ‘‘lesion’’ substrate is adenine when paired opposite oxoG. Both enzymes recognize the guanine-like Watson–Crick face of the opposite base using two main-chain carbonyl oxygens (Fromme and Verdine, 2003b; Fromme et al., 2004a). However, MutY possesses an additional C-terminal domain that has been demonstrated to be essential for proper functioning of MutY (Chmiel et al., 2001; Li et al., 2000; Noll et al., 1999). Interestingly, this domain shares low structural homology with the MutT enzyme, an oxo-dGTP triphosphatase. Using data from a fluorescence-based study, it was initially proposed that MutY would extrude not only the substrate adenine but also the partner oxoG from the DNA helix (Bernards et al., 2002). This ‘‘double-flipping’’ mechanism was appealing for several reasons, but the experimental evidence used to reach this conclusion was unable to distinguish between the effects of DNA bending and base extrusion. In fact, the crystal structure of MutY bound to duplex DNA containing an oxoG:A pair reveals that the oxoG remains intrahelical in the recognition complex (Fromme et al., 2004a) (Fig. 6). An interesting aspect of the MutY oxoG recognition mode is that oxoG is bound in the anti conformation, in contrast to the syn conformation oxoG normally adopts when paired with adenine in DNA (Kouchakdjian et al., 1991; McAuley-Hecht et al., 1994). The anti conformation forces the substrate adenine into the active site, and the syn conformation may serve as a homing mechanism for MutY while searching for damage (see following). Recognition of adenine by MutY seems to occur mainly by exclusion of cytosine, the other possible partner to oxoG. Endonuclease III can process a variety of lesions, which is borne out in the structure of a dihydrouracilcontaining recognition complex, evidenced by a lack of direct interactions between the enzyme and substrate base ( J. C. Fromme and G. L. Verdine, unpublished results). Structures that lack a bound DNA duplex, but contain a substrate lesion in the free-base form, can also be informative. The nuclear magnetic resonance structures of 3-methyladenine glycosylase I (known as MagI or
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Fig. 6. Structure of MutY bound to DNA containing an A:oxoG pair. The DNA is shown in gold, with oxoG in magenta and the substrate adenine in purple. MutY extrudes the adenine base from the duplex, but the oxoG base remains intrahelical. The [4Fe-4S] cluster is shown as yellow and orange spheres. Reprinted with permission (Fromme et al., 2004a). (See Color Insert.)
Tag) with bound substrate analogs and accompanying biochemistry show that MagI recognizes its substrate bases primarily through van der Waals interactions and hydrogen bonds with the Watson–Crick and major-groove face of the lesions (Cao et al., 2003). Crystal structures of 3-methyladenine glycosylase III (MagIII) bound to substrate analogs, together with kinetic analysis, demonstrate that MagIII recognizes its substrate via aromatic stacking interactions and steric exclusion (Eichman et al., 2003). Other crystal structures that hint at substrate recognition mechanism— despite the absence of a lesion base—include the complex between the bacterial 3-methyladenine glycosylase AlkA (MagII) and DNA containing an abasic site analog (Hollis et al., 2000) and a trapped-intermediate complex of endonuclease VIII (Zharkov et al., 2002). Both structures establish the overall DNA binding mode but lack a concrete depiction of substrate base recognition. Several different DNA glycosylases are known to act on T:G mismatches in DNA. The primary source of these mismatches is deamination of 5-methylcytosine (Duncan and Miller, 1980; Sved and Bird, 1990). 5-Methylcytosine is found in eukaryotes as an epigenetic modifier of chromatin structure and transcription, and in prokaryotes as the basis for self-protection from endogenous restriction enzymes. The human
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MBD4 protein is of particular interest among these evolutionarily widespread proteins. It possesses two domains, a methyl-CpG binding domain and a T:G glycosylase domain. The thymine glycosylase activity of the second domain is sequence specific, preferring substrates with a methylated CpG site paired opposite the substrate thymine base (Hendrich et al., 1999). The crystal structure of the glycosylase domain has been determined in the absence of substrate (Wu et al., 2003), confirming its place within the HhH-GPD superfamily. The bacterial thymine DNA glycosylase Mig is also a member of the HhH-GPD superfamily. The crystal structure of Mig has been determined (Mol et al., 2002) but also lacks substrate. Both of these enzymes have the potential to teach us new substrate recognition mechanisms, but we are awaiting structures obtained in the presence of lesion-containing DNA.
2. Searching for Damage There are several competing theories concerning the nature of the damage search process. The most basic mechanism involves a simple three-dimensional diffusion search, wherein encounters between the enzyme and DNA are random and transient, unless damage is located during the encounter. Based on the steady-state frequency of oxoG-lesions in the genome—about two oxoG residues per million guanines (ESCODD, 2002) or 1500 in a human cell—this method seems unlikely. A more expedient mechanism invokes a one-dimensional search process, in which the DNA glycosylase slides along the DNA duplex, thus greatly improving the likelihood of finding damage. Within this latter mechanism, there is room for variant methods of detecting DNA damage. It has been suggested that DNA glycosylases that act on non-helix-destabilizing lesions must extrude every base they encounter to recognize damaged bases (Verdine and Bruner, 1997). In contrast, DNA glycosylases that act on helix-destabilizing lesions may home in on damaged bases indirectly, simply by binding preferentially to deformable sites in DNA. The structure of a MutY-DNA complex indicates a possible avenue for direct detection (Fromme et al., 2004a). The syn configuration oxoG adopts when paired with adenine projects the N1, N2, and O6 atoms of oxoG out of the major groove. This protuberance interrupts an otherwise relatively smooth major groove surface, and it is possible that the projection is directly detected by MutY. This mechanism is satisfying in part because it removes the possibility of MutY stalling at oxoG:C pairs, in which oxoG adopts the anti conformation. Using duplex DNA with multiple substrates separated by defined distances, ‘‘processivity’’ has been demonstrated for several DNA glycosylases
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(Francis and David, 2003; Higley and Lloyd, 1993). However, it should be noted that these studies do not use statistical arguments to distinguish between true processivity and proximity bias. A recent study on MutY demonstrated that its [4Fe-4S] cluster is redox-active when the enzyme is DNA-bound (Boon et al., 2003), whereas it was previously believed to be redox-inert. Furthermore, the redox state of the metal cluster modulated the affinity of MutY for DNA. The authors of this study suggest the cluster may play a role in damage searching, insofar as oxidative lesions prevent transmission of electrons through DNA. MYH is known to associate with PCNA, a protein complex essential for DNA replication (Chang and Lu, 2002; Parker et al., 2001), and it has been shown to associate with replication forks (Boldogh et al., 2001). This colocalization may simplify MutY damage searching. In eukaryotic organisms, DNA exists in a chromatin-bound state, with accessibility related to transcriptional status. It seems likely that the chromatin state will influence the ability of BER enzymes to search for and repair damage. Indeed a recent study has shown that nucleosomebound DNA can be acted on by UDG and AP endonuclease, but not by DNA polymerase (Beard et al., 2003). These results indicate that chromatin remodeling is necessary for completion of the BER process, but not for the recognition of damage. However, in another study, polymerase was active upon nucleosome-bound substrates (Nilsen et al., 2002), though it is possible that the nucleosomal structures were more loosely packed in this study. Further efforts are necessary to establish the unique requirements of BER on a chromatin substrate.
D. Catalysis DNA glycosylases are interesting subjects for enzymological studies, if only because their substrate is the genome. Bifunctional glycosylases are especially interesting from a mechanistic standpoint because they catalyze several sequential reactions within a single active site. The base removal, or excision, step is shared by both mechanistic classes, and it is likely catalyzed similarly by both. Catalytic studies of DNA glycosylases are frequently complicated by several issues. One problem is that many glycosylases are severely end-product inhibited, rendering multiple-turnover kinetic studies uninformative. A second problem is that substrate binding occurs in several time-consuming steps, as a result of the need for gross structural rearrangement of the DNA duplex ( Jiang and Stivers, 2002). Thus, the rates of Michaelis complex formation may be slower than those of the chemical steps, making it difficult to ascribe rate constants to specific events.
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The mechanism of catalysis by UDG, including events from the initial DNA binding process through nucleoside extrusion and uracil excision, has been studied extensively by Tainer, Stivers, and their colleagues (Parikh et al., 2000; Stivers and Drohat, 2001). Some of the highlights of this research include the importance of a leucine residue for extrusion of the uridine nucleoside from the helix, which can be mimicked by replacement of the opposite guanine with a bulky pyrene residue ( Jiang et al., 2002). Extrusion of uracil from the DNA duplex is the rate-limiting step in catalysis by UDG ( Jiang and Stivers, 2002). Other studies have demonstrated that uracil excision by UDG follows an SN1-type pathway (Dinner et al., 2001; Werner and Stivers, 2000). The active site of UDG lowers the pKa of the uracil leaving group by 3.4 units to facilitate catalysis (Drohat and Stivers, 2000). Monofunctional and bifunctional glycosylases alike use aspartate or glutamate sidechains to stabilize the partial positive charge that is proposed to develop either on O40 or C10 in the transition state of the excision reaction. An additional driving force for excision of uracil is thought to derive from nearby phosphates in the DNA backbone (Dinner et al., 2001; Jiang et al., 2003). Alkylated lesion bases have especially labile glycosidic bonds, and therefore require less ‘‘powerful’’ catalysis by DNA glycosylases. Accordingly, MagIII performs catalysis on alkylated substrates even when the conserved catalytic aspartate is mutated (Eichman et al., 2003). However, the alkylation damage-specific glycosylase AlkA has been shown to be a sufficiently powerful catalyst to remove normal bases from DNA (Berdal et al., 1998), and AAG exhibits a 108-fold catalytic rate enhancement on the substrate hypoxanthine (O’Brien and Ellenberger, 2003). Monofunctional glycosylases replace the leaving group lesion base with a water molecule. In contrast, bifunctional glycosylases substitute a nucleophilic sidechain. The moieties used are an N-terminal proline in MutM family members, an N-terminal threonine in T4 endonuclease V, and an internal lysine in HhH-GPD glycosylases. The formation of iminium intermediates, available only to bifunctional glycosylases, is what allows for the subsequent steps of strand nicking through -elimination. As mentioned above, the primary bifunctional intermediate can be ‘‘trapped’’ with a reducing agent. There are now crystal structures of trapped intermediates available from several glycosylases (Fromme et al., 2004b). One such structure led to the discovery that hOGG1 uses the product of base excision, the oxoG lesion base, as a cofactor to assist in catalysis of -elimination (Fromme et al., 2003). The catalytic mechanisms of DNA glycosylases, with special emphasis on monofunctional glycosylases, is the subject of a recent review (Stivers and Jiang, 2003).
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IV. Downstream BER Enzymes A. AP Endonucleases The products of DNA glycosylase activity are quite similar despite the varied nature of glycosylase substrates. All monofunctional DNA glycosylases produce the same species: the abasic site. The abasic site produced enzymatically is chemically identical to the abasic product of spontaneous base loss from DNA. The abasic site exhibits considerable genotoxicity. Not only do abasic sites lack the coding information necessary for template-directed DNA synthesis but they can lead to stalled replication forks (Higuchi et al., 2003; Loeb and Preston, 1986) and transcription bubbles (Yu et al., 2003); events that are potentially mutagenic or even lethal. They have also been shown to trap topoisomerase I through a covalent interaction (Pourquier et al., 1997). Two studies by Guillet and Boiteux indicate that, at least in yeast, the single major source of abasic sites in the genome is from the action of UDG on uracil, and that these abasic sites are lethal in the absence of AP endonuclease activity (Guillet and Boiteux, 2002, 2003). Importantly, lethality was suppressed by deletion of UDG, but not by any other DNA glycosylase examined. AP endonucleases cleave the DNA strand on the 50 side of abasic sites, resulting in a nick having 30 -OH and 50 -deoxyribosephosphate (dRP) groups. In addition, AP endonucleases will process the -elimination products of bifunctional DNA glycosylases by completely removing the processed lesion nucleoside, leaving behind a free 30 -OH. AP endonucleases also possess 30 -phosphoesterase activity, enabling them to remove the 30 -phosphate group remaining after the ,-elimination activity of some bifunctional glycosylases (Fig. 3). The bacterium Escherichia coli possesses two different AP endonucleases, exonuclease III and endonuclease IV (Ljungquist and Lindahl, 1977). These enzymes define the two known structural classes of AP endonuclease. The predominant enzyme in E. coli is exonuclease III, which has 30 ! 50 exonuclease activity in addition to the characteristic AP endonuclease activities detailed above (Rogers and Weiss, 1980; Weiss, 1976). Endonuclease IV, whose expression in E. coli is induced in response to oxidative stress (Chan and Weiss, 1987), is homologous to the major AP endonuclease found in baker’s yeast, APN1 (Popoff et al., 1990). The major human AP endonuclease, APE1 (also known as Ref-1 or HAP1), is homologous to exonuclease III (Demple et al., 1991). APE1 is multifunctional, possessing redox-dependent transcriptional activation (Xanthoudakis et al., 1992) and acetylation-dependent, redox-independent transcriptional repression activities (Bhakat et al., 2003; Okazaki et al., 1994) in addition to AP
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endonuclease activities (Fritz et al., 2003). It has also been suggested that APE1 is the main proofreading enzyme for mistakes made by DNA polymerase , based on its ability to remove mismatched bases at 30 termini (Chou and Cheng, 2002). The additional roles of APE1 will be discussed in greater detail later. Because AP endonuclease acts immediately downstream of the DNA glycosylases, the DNA repair community has actively sought evidence of interactions between the two classes of enzyme (Wilson and Kunkel, 2000). Human APE1 has been shown to stimulate the activity of the human DNA glycosylases UDG (Parikh et al., 1998), TDG (Waters et al., 1999), hOGG1 (Vidal et al., 2001b), and NTH1 (Marenstein et al., 2003), most likely through competitive substrate binding, but perhaps through a more active mechanism. The only demonstration to date of a direct physical interaction between AP endonuclease and a DNA glycosylase is between the human adenine glycosylase MYH and APE1 (Parker et al., 2001). The structural biology of AP endonucleases has been well established through the work of Tainer and colleagues (Mol et al., 2000a), with DNAbound structures available from structurally distinct human APE1 (Mol et al., 2000b) and E. coli endonuclease IV (Hosfield et al., 1999). The DNA cocrystal structures reveal how these enzymes use different folds to achieve similar results (Fig. 7A and B). Both enzymes extrude the substrate abasic site from the DNA helix in a similar manner to the DNA glycosylases, bending the duplex in the process and inserting protein residues into the minor groove. Both enzymes use bound metal ion(s) to catalyze strandnicking via hydrolysis, and each structure led to a plausible proposal for the enzymatic mechanism. Notably, endonuclease IV appears to use three zinc ions for catalysis. A somewhat surprising aspect of the structure of the APE1-DNA complex is the location of the two conserved cysteine residues responsible for the redox activity of APE1. These residues—Cys65 and Cys93 (Walker et al., 1993)—are not exposed to solvent but, instead, are buried within the protein. This indicates that APE1 might undergo a significant conformational change to carry out its redox activity (Fig. 7C).
B. Short-Patch versus Long-Patch Repair After the action of AP endonuclease on an abasic site, DNA repair polymerase (polymerase in eukaryotes or polymerase I in bacteria) can remove the dRP group (so-called ‘‘dRPase activity’’) (Allinson et al., 2001). This reaction is catalyzed by the N-terminal domain of polymerase via an imine intermediate, analogous to the AP lyase activity of bifunctional glycosylases (Piersen et al., 1996; Prasad et al., 1998). If this step
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Fig. 7. The two structurally distinct AP endonucleases. (A) Human APE1, in blue, bound to DNA containing a substrate abasic site, in red (Mol et al., 2000b). (B) Endonuclease IV, in red, bound to its product DNA (Hosfield et al., 1999). The abasic moiety of the dRP group is colored blue. (C) View of the APE1/DNA complex highlighting the positions of the two cysteines involved in the redox activity of this enzyme. The cysteine residues are yellow, seen buried within the interior of the protein. (See Color Insert.)
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transpires, the product is chemically identical to the gap resulting from AP endonuclease activity on a bifunctional elimination product. The one nucleotide gap is filled in by the polymerase domain of DNA polymerase , and the strand is resealed by a DNA ligase III/XRCC1 complex (Cappelli et al., 1997). DNA ligase I has also been implicated as the functional ligase acting at this stage, based on its association with polymerase (Prasad et al., 1996) and competence within an in vitro BER assay (Nicholl et al., 1997). The final steps described above are referred to as ‘‘short-patch’’ base excision repair, because of the single nucleotide polymerization needed to complete the repair process (Fig. 1, bottom right). An alternative repair process (Frosina et al., 1996; Matsumoto et al., 1994), termed ‘‘long-patch’’ base excision repair (Fig. 1, bottom left), occurs when DNA polymerase /" initiates polymerization from the free 30 -OH adjacent to the dRP group resulting from AP endonuclease activity. The polymerization incorporates between 2 and 15 nucleotides, displacing the strand containing the 50 -dRP (called the ‘‘flap’’) and requiring FEN-1, a nuclease that removes this flap (Klungland and Lindahl, 1997). Longpatch repair has been reconstituted in vitro (Matsumoto et al., 1999; Pascucci et al., 1999) and was shown to require both PCNA and the nuclease FEN-1. If flap removal produces a gap of fewer than 6 nucleotides, polymerase can be a key player in long-patch repair (Fortini et al., 1998; Wilson, 1998). An in vitro reconstitution of long-patch repair was achieved using polymerase and demonstrated that PARP-1 (poly ADP-ribose polymerase-1) stimulated the long-patch activity of polymerase (Prasad et al., 2001). However, another report indicated that PARP-1 actually slows down the repair reaction (Allinson et al., 2003). PARP-1 binds to single-strand nicks in DNA (Benjamin and Gill, 1980) and is believed to play a role in damage repair by scanning the genome for these nicks. PARP-1 was shown to interact with the product of AP endonuclease activity in an ultraviolet cross-linking study (Lavrik et al., 2001). The role of PARP in BER is discussed further later. Recent structural studies of complexes between FEN-1, PCNA, and DNA reveal the basis for the flap specificity of FEN-1 (Chapados et al., 2004). FEN-1 binds to a kinked structure of the DNA duplex made possible by the discontinuity of the nicked strand. The authors use the structure of a FEN1 peptide bound to PCNA to propose a model of how PCNA orients FEN-1 for interaction with DNA at a replication site (Fig. 8). Furthermore, it is proposed that FEN-1 and the next downstream enzyme, DNA ligase, both use similar features of nicked-DNA for substrate recognition while bound to PCNA. Physical interactions between APE1 and FEN-1 and APE1 and PCNA have been demonstrated in copurification experiments (Dianova et al.,
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Fig. 8. Modeled structure of a FEN-1/PCNA/DNA complex. This model was generated using coordinates from structures of a FEN-1/DNA complex and a complex between PCNA and a peptide fragment of FEN-1 (Chapados et al., 2004). PCNA is shown in green, FEN-1 in blue, and DNA in gold. The DNA is presumed to extend downward from its point of contact with FEN-1. (See Color Insert.)
2001). Together with the known interactions between FEN-1 and PCNA (Wu et al., 1996), these interactions indicate that long-patch repair is a tightly coordinated event, using interactions between enzymes and with PCNA as a scaffold. It has been noted that 50 -dRPs are more likely to provoke long-patch repair than are single-nucleotide gaps (Klungland et al., 1999a). 50 -dRPs arise after the sequential actions of a monofunctional glycosylase and AP endonuclease, whereas single-nucleotide gaps arise from the actions of a bifunctional glycosylase and AP endonuclease. Furthermore, bifunctional glycosylases (OGG1, MutM, EndoIII, and EndoVIII) seem to be restricted to oxidatively damaged substrates. What is unique about these substrates that requires short-patch repair instead of long-patch repair? Lindahl and colleagues have proposed that ionizing radiation, a major source of
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oxidative lesions via generation of free radicals, tends to generate multiple lesions in a concentrated area; long-patch repair might lead to a disastrous double-strand break if two lesions lay on opposite strands within a few basepairs of each other (Klungland et al., 1999a). Therefore, by using bifunctional glycosylases to initiate repair at oxidative lesions, cells can avoid the potentially harmful long-patch repair process.
V. Mammalian BER A. Additional Components BER in mammalian cells is more complex than in simpler eukaryotes (Izumi et al., 2003; Memisoglu and Samson, 2000), involving interactions with additional proteins like XRCC1 (Thompson and West, 2000) and p53 (Offer et al., 2001b) that coordinate repair and alert the cell to the presence of damage. APE1 is suspected to be the key BER protein involved in these extra layers of complexity (Fritz et al., 2003). The redox activity of APE1 is known to regulate the DNA-binding affinity of several transcription factors. The first example of such an interaction was with the transcription factors Fos and Jun (Xanthoudakis et al., 1992), which heterodimerize to form AP-1, an oxidation-sensitive transcription factor. APE1, using two specific cysteine residues, reduces AP-1 in a thioredoxindependent manner (Wei et al., 2000). The yeast version of APE1 does not contain the conserved cysteine residues critical for redox function, lending credence to the view that this activity occurs only in higher eukaryotes (Fritz et al., 2003). APE1 also serves as a redox factor for p53 (Gaiddon et al., 1999; Jayaraman et al., 1997), NF-B (Mitomo et al., 1994), HIF-1 (Huang et al., 1996), and other transcription factors (Flaherty et al., 2001). The redox activity of APE1 has been shown to be stimulated by phosphorylation by casein kinase II and protein kinase C both in vitro and in vivo (Fritz and Kaina, 1999; Hsieh et al., 2001), indicating that there may be a mechanism whereby damage sensed by APE1 leads to its phosphorylation and subsequent activation. This hypothesis is especially appealing considering the roles of casein kinase II and protein kinase C in the DNA damage response (Ghavidel and Schultz, 2001; Yoshida et al., 2003). PARP-1 is a signaling enzyme that transfers ADP-ribose groups from NADþ to itself and other nuclear proteins in response to detecting DNA damage. Poly(ADP)ribosylation of histones has been shown to loosen chromatin (de Murcia et al., 1986), indicating that damage detection by PARP-1 leads to increased access to the site of damage for other repair
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proteins. PARP-1 interacts with DNA polymerase in GST-pulldown assays (Dantzer et al., 2000), and a knockout mouse model demonstrated the key role this enzyme plays in BER (Dantzer et al., 1999), as demonstrated by the inadequate repair of abasic sites by PARP-1-deficient cell extracts. Interestingly, the interaction between PARP-1 and polymerase does not depend on poly(ADP)ribosylation but does depend on the presence of damaged DNA, whereas the interaction between PARP-1 and XRCC1 is dependent on poly(ADP)ribosylation but independent of DNA (Dantzer et al., 2000; Masson et al., 1998). XRCC1 (x-ray repair cross-complementing protein 1) is a nonenzymatic scaffolding protein that binds to other key BER enzymes, APE1 (Vidal et al., 2001a), DNA polymerase (Caldecott et al., 1996; Kubota et al., 1996), and DNA ligase III (Caldecott et al., 1994). More recently, XRCC1 has been shown to interact with hOGG1 in both GST-pulldown and yeast two-hybrid assays (Marsin et al., 2003). In addition, XRCC1 stimulates the repair activity of hOGG1 on DNA containing oxoG, similar to stimulation by APE1. The tumor suppressor p53 plays a more direct role in BER in addition to transcriptional regulation. p53 stimulates the rate of an in vitro BER system by directly interacting with DNA polymerase and stabilizing the interaction between the polymerase and abasic DNA (Zhou et al., 2001). The in vivo relevance of this finding was validated when mutations in the transactivating domain of p53 failed to inactivate BER in mammalian cells, whereas mutations elsewhere in p53 diminished BER (Offer et al., 2001a). Another more recent study indicates that p53 can potentiate the formation of abasic sites by repressing transcription of 3-methyladenine DNA glycosylase in response to genotoxic stress (Zurer et al., 2004). This mechanism might alleviate the mutational burden by ensuring that the level of abasic site formation does not outpace the level of APE1. XPG is an endonuclease associated with the NER pathway in eukaryotes. The repair of oxidized lesions in vitro has been shown to be accelerated by XPG, through interactions with the DNA glycosylase hNTH1 that increase the affinity of hNTH1 for DNA (Klungland et al., 1999a).
B. Knockout Mice and the Role of BER Proteins in Human Disease Given the importance of DNA glycosylases for BER, the DNA repair community anxiously awaited the results of mouse knockout studies. The initial results were generally greeted with disappointment, as animals lacking particular DNA glycosylases seemed phenotypically normal. Early studies of AAG (also known as ANPG)-knockout mice demonstrated an increased sensitivity to methylating agents in cells derived from knockout
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embryos, but the mice themselves exhibited no apparent phenotype (Engelward et al., 1997). Parenthetically, the knockout studies were useful in defining the in vivo substrate range for this glycosylase (Engelward et al., 1997; Hang et al., 1997). The generation of UDG (also known as UNG) knockout mice (Nilsen et al., 2000) also revealed these animals to be outwardly normal despite an increase in the steady-state genomic level of uracil. Cell extracts from knockout animal tissue demonstrated a persistent uracil glycosylase activity, later definitively attributed to SMUG1 (Nilsen et al., 2001). These two studies established the role of UDG during DNA replication and that of SMUG1 as a constitutively expressed enzyme found only in higher eukaryotes. A more recent study revealed that UDG knockout mice eventually develop B cell lymphomas, highlighting the increasingly recognized importance of UDG in the immune system (Nilsen et al., 2003). The role of UDG in somatic hypermutation is discussed in Chapter 11. OGG1 knockout mice appeared phenotypically normal but displayed increased levels of oxoG in their genomes (Klungland et al., 1999b; Minowa et al., 2000). Further studies showed that cells from knockout mice were more susceptible to exogenous mutagens both in vitro and in vivo (Arai et al., 2002, 2003). Knockout mouse embryo fibroblasts were used to show that oxoG can be repaired independent of OGG1 in a transcription-coupled repair process (Le Page et al., 2000). A recent study has shown that OGG1 knockout mice exhibit a fivefold increase in lung tumors at age 18 months (Sakumi et al., 2003), indicating that the resulting mutational burden eventually manifests itself via carcinogenesis. Curiously, in this same study it was shown that disruption of MTH1, which is responsible for sanitizing the nucleotide precursor pool of oxo-dGTP by catalyzing its hydrolysis to oxo-dGMP and pyrophosphate, suppressed the tumorigenic phenotype of OGG1 deletion. MBD4 knockout mice are viable but display increased mutation rates at CpG sites (Millar et al., 2002). When these knockout mice where crossed with mice predisposed to colon cancer, the tumorigenic potential of the MBD4 deletion was revealed. A further study highlighted the important role of MBD4 in triggering apoptosis, as a reduced apoptotic response was observed in cells from knockout mice (Sansom et al., 2003). NTH1 knockout mice, though otherwise viable, were crucial to the discovery of novel mammalian homologs of Nei, the NEIL proteins (Elder and Dianov, 2002; Ocampo et al., 2002; Takao et al., 2002b). The NEIL family currently consists of three family members. Surprisingly, NEIL3 contains a C-terminal domain that is homologous to both topoisomerase III and APE2, an AP endonuclease-like protein with unknown function (Takao et al., 2002a).
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It was initially assumed that functional redundancy obscured any obvious phenotypic effects in animals genetically engineered to lack certain DNA glycosylases (Parsons and Elder, 2003). However, the recent results with OGG1 and UDG knockout animals indicate that deficiency of a single DNA glycosylase in some cases is sufficient to evoke carcinogenesis, but this phenotype takes time to develop. Although no MutY knockout animal is available yet, there is now compelling genetic evidence that this DNA glycosylase acts as a tumor suppressor in humans. The human homolog of MutY (adenine DNA glycosylase), hMYH, has been persuasively linked to colon cancer. The first observation of this fact came from a study of familial colon cancers not associated with inherited deficiencies in the usual suspect, APC (Al-Tassan et al., 2002). This and subsequent studies confirmed the prevalence of two specific MYH polymorphisms in such cancers and showed that these polymorphisms are associated with tumorigenesis in a number of unrelated patients (Cheadle and Sampson, 2003; Jones et al., 2002; Sampson et al., 2003; Sieber et al., 2003). The effects of the polymorphisms on enzyme function were shown using the equivalent mutations in the E. coli homolog (Chmiel et al., 2003) and seem to stem mainly from reduced activity of the enzyme. Furthermore, the reduction in activity was exacerbated at known ‘‘hotspot’’ sequences (Al-Tassan et al., 2002) mutated in the somatic APC genes of patients inheriting defective MYH. APE1 is overexpressed in many cancers (Holmes et al., 2002; Kakolyris et al., 1997; Moore et al., 2000; Robertson et al., 2001) and is essential for development (Xanthoudakis et al., 1996). A recent study found that granzyme A cleaves APE1 as part of the cytotoxic T cell–initiated cascade leading to cell death; overexpression of a noncleavable version of APE1 resulted in cells that were less sensitive to granzyme A-mediated cell death (Fan et al., 2003). Granzyme A activity was seen to affect both the AP endonuclease and redox functions of APE1, though redox activation of transcription factors was suggested to be more critical for death avoidance. The authors of this study propose that inactivation of APE1 is critical for the granzyme A-mediated cell death pathway because the transcription factors activated by APE1 inhibit apoptosis during DNA repair. Others have suggested that the DNA repair activity of APE1 is responsible for resistance to mutation-inducing cancer therapies (Fritz et al., 2003). A similar finding with PARP-1 provides evidence for the role of this enzyme in regulation of cell fate. PARP-1 is degraded by caspase 3 during apoptosis (Lazebnik et al., 1994; Nicholson et al., 1995), indicating that it normally provides resistance to apoptosis. A complementary finding is that PARP-1 poly(ADP)ribosylates the Ca2þ/Mg2þ endonuclease, suppressing the action of this proapoptotic enzyme (Yakovlev et al., 2000).
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Imbalances in BER are known to induce mutagenesis and to be associated with cancer. This phenomenon has been illustrated by overexpression of DNA glycosylases (Coquerelle et al., 1995; Glassner et al., 1998) or polymerase (Bergoglio et al., 2002; Canitrot et al., 1998). The increase in mutagenesis in the former case has been attributed to accumulation of abasic sites. Overexpression of polymerase is thought to lead to mutagenesis because of the poor fidelity of this polymerase.
VI. Roles of BER Enzymes in Other Processes A. BER-Like Enzymes in Plant Development Accumulating evidence indicates a critical role for proteins with DNA glycosylase activity in seed development of the plant Arabidopsis thaliana. The first evidence of this phenomenon came from a study on the imprinting of the MEA gene. This gene is normally silenced by methylation, but the maternal allele is activated in the endosperm of seeds. The activation was shown to accompany demethylation of the maternal allele, and the surprising cause of demethylation was shown to be the DNA glycosylase DEMETER (Choi et al., 2002). Subsequent studies confirmed this finding while identifying MET1 as the methyltransferase maintaining MEA in the repressed state (Xiao et al., 2003) and found that the FWA gene was regulated in an identical manner, also requiring DEMETER for activation (Kinoshita et al., 2004). Another Arabidopsis protein related to DEMETER, ROS1, is a DNA glycosylase that functions in a similar manner. Furthermore, a purified MBP-ROS1 fusion protein was shown to nick (presumably through bifunctional glycosylase activity) plasmid DNA containing methylated CpG sites (Gong et al., 2002). The most intriguing question surrounding this regulatory system is the exact mechanism whereby the activities of DEMETER and ROS1 lead to cytosine demethylation at CpG sites. The authors of these studies have proposed competing mechanisms, invoking either an intrinsic 5-methylcytosine DNA glycosylase activity within DEMETER and ROS1 or an indirect pathway in which these proteins create single-stranded nicks near methylated CpG sites that somehow lead to replacement with unmethylated nucleotides. In fact, no mechanism for chromosomal demethylation has been conclusively established, and reports of 5-methylcytosine glycosylase activity ( Jost et al., 2001; Zhu et al., 2000) have not been independently confirmed. It is nonetheless intriguing that DNA glycosylases appear to be key players in epigenetic demethylation, and we anxiously await further clarification of the details.
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B. Thymine DNA Glycosylase Thymine DNA glycosylase (TDG) is unique among the DNA glycosylases with regard to the number and diversity of its protein partners. As discussed above, the function of TDG is critical for removal of T:G mismatches arising via deamination of 5-methylcytosine. TDG has been shown to be sumoylated at a single-lysine residue (Hardeland et al., 2002). Sumoylation increases the turnover rate of TDG while suppressing the activating effect of APE1 by reducing the affinity of TDG for DNA. This type of regulation is thus far unique to TDG among DNA glycosylase. TDG is known to interact with the transcriptional coactivators CBP/p300 (Tini et al., 2002). This interaction stimulates transcription in cells and leads to acetylation of TDG. The significance of increased transcription may be that the loosened chromatin structure of a transcriptionally active region allows DNA repair proteins easier access to the damaged site. Acetylated TDG no longer binds CBP and no longer recruits APE1 to damaged sites, indicating that acetylation serves as another level of regulation. TDG has also recently been reported to function as a coactivator for the nuclear estrogen receptor alpha (Chen et al., 2003), thus forging yet another link between BER and other critical aspects of cellular function.
Acknowledgments We are grateful to Anirban Banerjee for helpful discussions. J.C.F. is sponsored by a MerckWiley Fellowship.
References Allinson, S. L., Dianova, I. I., and Dianov, G. L. (2001). DNA polymerase beta is the major dRP lyase involved in repair of oxidative base lesions in DNA by mammalian cell extracts. EMBO J. 20, 6919–6926. Allinson, S. L., Dianova, I. I., and Dianov, G. L. (2003). Poly(ADP-ribose) polymerase in base excision repair: Always engaged, but not essential for DNA damage processing. Acta Biochim. Pol. 50, 169–179. Al-Tassan, N., Chmiel, N. H., Maynard, J., Fleming, N., Livingston, A. L., Williams, G. T., Hodges, A. K., Davies, D. R., David, S. S., Sampson, J. R., and Cheadle, J. P. (2002). Inherited variants of MYH associated with somatic G:C–>T:A mutations in colorectal tumors. Nat. Genet. 30, 227–232. Arai, T., Kelly, V. P., Komoro, K., Minowa, O., Noda, T., and Nishimura, S. (2003). Cell proliferation in liver of Mmh/Ogg1-deficient mice enhances mutation frequency because of the presence of 8-hydroxyguanine in DNA. Cancer Res. 63, 4287–4292. Arai, T., Kelly, V. P., Minowa, O., Noda, T., and Nishimura, S. (2002). High accumulation of oxidative DNA damage, 8-hydroxyguanine, in Mmh/Ogg1 deficient mice by chronic oxidative stress. Carcinogenesis 23, 2005–2010. Bailly, V., and Verly, W. G. (1987). Escherichia coli endonuclease III is not an endonuclease but a beta-elimination catalyst. Biochem. J. 242, 565–572.
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Bandaru, V., Sunkara, S., Wallace, S. S., and Bond, J. P. (2002). A novel human DNA glycosylase that removes oxidative DNA damage and is homologous to Escherichia coli endonuclease VIII. DNA Repair 1, 517–529. Barrett, T. E., Savva, R., Panayotou, G., Barlow, T., Brown, T., Jiricny, J., and Pearl, L. H. (1998). Crystal structure of a G:T/U mismatch-specific DNA glycosylase: Mismatch recognition by complementary-strand interactions. Cell 92, 117–129. Barrett, T. E., Scharer, O. D., Savva, R., Brown, T., Jiricny, J., Verdine, G. L., and Pearl, L. H. (1999). Crystal structure of a thwarted mismatch glycosylase DNA repair complex. EMBO J. 18, 6599–6609. Beard, B. C., Wilson, S. H., and Smerdon, M. J. (2003). Suppressed catalytic activity of base excision repair enzymes on rotationally positioned uracil in nucleosomes. Proc. Natl. Acad. Sci. USA 100, 7465–7470. Benjamin, R. C., and Gill, D. M. (1980). Poly(ADP-ribose) synthesis in vitro programmed by damaged DNA. A comparison of DNA molecules containing different types of strand breaks. J. Biol. Chem. 255, 10502–10508. Berdal, K. G., Johansen, R. F., and Seeberg, E. (1998). Release of normal bases from intact DNA by a native DNA repair enzyme. EMBO J. 17, 363–367. Bergoglio, V., Pillaire, M. J., Lacroix-Triki, M., Raynaud-Messina, B., Canitrot, Y., Bieth, A., Gares, M., Wright, M., Delsol, G., Loeb, L. A., Cazaux, C., and Hoffmann, J. S. (2002). Deregulated DNA polymerase beta induces chromosome instability and tumorigenesis. Cancer Res. 62, 3511–3514. Bernards, A. S., Miller, J. K., Bao, K. K., and Wong, I. (2002). Flipping duplex DNA inside out: A double base-flipping reaction mechanism by Escherichia coli MutY adenine glycosylase. J. Biol. Chem. 277, 20960–20964. Bhagwat, M., and Gerlt, J. A. (1996). 30 - and 50 -strand cleavage reactions catalyzed by the Fpg protein from Escherichia coli occur via successive beta- and delta-elimination mechanisms, respectively. Biochemistry 35, 659–665. Bhakat, K. K., Izumi, T., Yang, S. H., Hazra, T. K., and Mitra, S. (2003). Role of acetylated human AP-endonuclease (APE1/Ref-1) in regulation of the parathyroid hormone gene. EMBO J. 22, 6299–6309. Boldogh, I., Milligan, D., Lee, M. S., Bassett, H., Lloyd, R. S., and McCullough, A. K. (2001). hMYH cell cycle-dependent expression, subcellular localization and association with replication foci: Evidence suggesting replication-coupled repair of adenine:8-oxoguanine mispairs. Nucleic Acids Res. 29, 2802–2809. Boon, E. M., Livingston, A. L., Chmiel, N. H., David, S. S., and Barton, J. K. (2003). DNA-mediated charge transport for DNA repair. Proc. Natl. Acad. Sci. USA 100, 12543–12547. Bregeon, D., Doddridge, Z. A., You, H. J., Weiss, B., and Doetsch, P. W. (2003). Transcriptional mutagenesis induced by uracil and 8-oxoguanine in Escherichia coli. Mol. Cell. 12, 959–970. Bruner, S. D., Nash, H. M., Lane, W. S., and Verdine, G. L. (1998). Repair of oxidatively damaged guanine in Saccharomyces cerevisiae by an alternative pathway. Curr. Biol. 8, 393–403. Bruner, S. D., Norman, D. P., and Verdine, G. L. (2000). Structural basis for recognition and repair of the endogenous mutagen 8-oxoguanine in DNA. Nature 403, 859–866. Burrows, C. J., and Muller, J. G. (1998). Oxidative nucleobase modifications leading to strand scission. Chem. Rev. 98, 1109–1152. Caldecott, K. W., Aoufouchi, S., Johnson, P., and Shall, S. (1996). XRCC1 polypeptide interacts with DNA polymerase beta and possibly poly (ADP-ribose) polymerase,
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and DNA ligase III is a novel molecular ‘nick-sensor’ in vitro. Nucleic Acids Res. 24, 4387–4394. Caldecott, K. W., McKeown, C. K., Tucker, J. D., Ljungquist, S., and Thompson, L. H. (1994). An interaction between the mammalian DNA repair protein XRCC1 and DNA ligase III. Mol. Cell. Biol. 14, 68–76. Canitrot, Y., Cazaux, C., Frechet, M., Bouayadi, K., Lesca, C., Salles, B., and Hoffmann, J. S. (1998). Overexpression of DNA polymerase beta in cell results in a mutator phenotype and a decreased sensitivity to anticancer drugs. Proc. Natl. Acad. Sci. USA 95, 12586–12590. Cao, C., Kwon, K., Jiang, Y. L., Drohat, A. C., and Stivers, J. T. (2003). Solution structure and base perturbation studies reveal a novel mode of alkylated base recognition by 3-methyladenine DNA glycosylase I. J. Biol. Chem 278, 48012–48020. Cappelli, E., Taylor, R., Cevasco, M., Abbondandolo, A., Caldecott, K., and Frosina, G. (1997). Involvement of XRCC1 and DNA ligase III gene products in DNA base excision repair. J. Biol. Chem. 272, 23970–23975. Chan, E., and Weiss, B. (1987). Endonuclease IV of Escherichia coli is induced by paraquat. Proc. Natl. Acad. Sci. USA 84, 3189–3193. Chang, D. Y., and Lu, A. L. (2002). Functional interaction of MutY homolog with proliferating cell nuclear antigen in fission yeast, Schizosaccharomyces pombe. J. Biol. Chem. 277, 11853–11858. Chapados, B. R., Hosfield, D. J., Han, S., Qiu, J., Yelent, B., Shen, B., and Tainer, J. A. (2004). Structural basis for FEN-1 substrate specificity and PCNA-mediated activation in DNA replication and repair. Cell 116, 39–50. Cheadle, J. P., and Sampson, J. R. (2003). Exposing the MYtH about base excision repair and human inherited disease. Hum. Mol. Genet. 12, R159–R165. Chen, D., Lucey, M. J., Phoenix, F., Lopez-Garcia, J., Hart, S. M., Losson, R., Buluwela, L., Coombes, R. C., Chambon, P., Schar, P., and Ali, S. (2003). T:G mismatch-specific thymine-DNA glycosylase potentiates transcription of estrogen-regulated genes through direct interaction with estrogen receptor alpha. J. Biol. Chem. 278, 38586–38592. Chmiel, N. H., Golinelli, M. P., Francis, A. W., and David, S. S. (2001). Efficient recognition of substrates and substrate analogs by the adenine glycosylase MutY requires the C-terminal domain. Nucleic Acids Res. 29, 553–564. Chmiel, N. H., Livingston, A. L., and David, S. S. (2003). Insight into the functional consequences of inherited variants of the hMYH adenine glycosylase associated with colorectal cancer: Complementation assays with hMYH variants and presteady-state kinetics of the corresponding mutated E. coli enzymes. J. Mol. Biol. 327, 431–443. Choi, Y., Gehring, M., Johnson, L., Hannon, M., Harada, J. J., Goldberg, R. B., Jacobsen, S. E., and Fischer, R. L. (2002). DEMETER, a DNA glycosylase domain protein, is required for endosperm gene imprinting and seed viability in arabidopsis. Cell. 110, 33–42. Chou, K. M., and Cheng, Y. C. (2002). An exonucleolytic activity of human apurinicapyrimidinic endonuclease on 30 mispaired DNA. Nature 415, 655–659. Coquerelle, T., Dosch, J., and Kaina, B. (1995). Overexpression of N-methylpurine-DNA glycosylase in Chinese hamster ovary cells renders them more sensitive to the production of chromosomal aberrations by methylating agents—a case of imbalanced DNA repair. Mutat. Res. 336, 9–17. Cullis, P. M., Malone, M. E., and Merson-Davies, L. A. (1996). Guanine radical cations are precursors of 7,8-dihydro-8-oxo-20 -deoxyguanosine but are not precursors of immediate strand breaks in DNA. J. Am. Chem. Soc. 118, 2775–2781.
BASE EXCISION REPAIR
31
Dantzer, F., de La Rubia, G., Menissier-De Murcia, J., Hostomsky, Z., de Murcia, G., and Schreiber, V. (2000). Base excision repair is impaired in mammalian cells lacking Poly(ADP-ribose) polymerase-1. Biochemistry 39, 7559–7569. Dantzer, F., Schreiber, V., Niedergang, C., Trucco, C., Flatter, E., De La Rubia, G., Oliver, J., Rolli, V., Menissier-de Murcia, J., and de Murcia, G. (1999). Involvement of poly(ADP-ribose) polymerase in base excision repair. Biochimie. 81, 69–75. de Murcia, G., Huletsky, A., Lamarre, D., Gaudreau, A., Pouyet, J., Daune, M., and Poirier, G. G. (1986). Modulation of chromatin superstructure induced by poly(ADP-ribose) synthesis and degradation. J. Biol. Chem. 261, 7011–7017. Demple, B., Herman, T., and Chen, D. S. (1991). Cloning and expression of APE, the cDNA encoding the major human apurinic endonuclease: Definition of a family of DNA repair enzymes. Proc. Natl. Acad. Sci. USA 88, 11450–11454. Dianova, I. I., Bohr, V. A., and Dianov, G. L. (2001). Interaction of human AP endonuclease 1 with flap endonuclease 1 and proliferating cell nuclear antigen involved in long-patch base excision repair. Biochemistry 40, 12639–12644. Dinner, A. R., Blackburn, G. M., and Karplus, M. (2001). Uracil-DNA glycosylase acts by substrate autocatalysis. Nature 413, 752–755. Dodson, M. L., Michaels, M. L., and Lloyd, R. S. (1994). Unified catalytic mechanism for DNA glycosylases. J. Biol. Chem. 269, 32709–32712. Dodson, M. L., Schrock, R. D. 3rd, and Lloyd, R. S. (1993). Evidence for an imino intermediate in the T4 endonuclease V reaction. Biochemistry 32, 8284–8290. Drohat, A. C., and Stivers, J. T. (2000). NMR evidence for an unusually low N1 pKa for uracil bound to uracil DNA glycosylase: Implications for catalysis. J. Am. Chem. Soc. 122, 1840–1841. Duncan, B. K., and Miller, J. H. (1980). Mutagenic deamination of cytosine residues in DNA. Nature 287, 560–561. Eichman, B. F., O’Rourke, E. J., Radicella, J. P., and Ellenberger, T. (2003). Crystal structures of 3-methyladenine DNA glycosylase MagIII and the recognition of alkylated bases. EMBO J. 22, 4898–4909. Elder, R. H., and Dianov, G. L. (2002). Repair of dihydrouracil supported by base excision repair in mNTH1 knock-out cell extracts. J. Biol. Chem. 277, 50487–50490. Engelward, B. P., Boosalis, M. S., Chen, B. J., Deng, Z., Siciliano, M. J., and Samson, L. D. (1993). Cloning and characterization of a mouse 3-methyladenine/7-methylguanine/3-methylguanine DNA glycosylase cDNA whose gene maps to chromosome 11. Carcinogenesis 14, 175–181. Engelward, B. P., Weeda, G., Wyatt, M. D., Broekhof, J. L., de Wit, J., Donker, I., Allan, J. M., Gold, B., Hoeijmakers, J. H., and Samson, L. D. (1997). Base excision repair deficient mice lacking the Aag alkyladenine DNA glycosylase. Proc. Natl. Acad. Sci. USA 94, 13087–13092. ESCODD(2002). Comparative analysis of baseline 8-oxo-7,8-dihydroguanine in mammalian cell DNA, by different methods in different laboratories: An approach to consensus. Carcinogenesis 23, 2129–2133. Fan, Z., Beresford, P. J., Zhang, D., Xu, Z., Novina, C. D., Yoshida, A., Pommier, Y., and Lieberman, J. (2003). Cleaving the oxidative repair protein Ape1 enhances cell death mediated by granzyme A. Nature Immunol. 4, 145–153. Flaherty, D. M., Monick, M. M., and Hunninghake, G. W. (2001). AP endonucleases and the many functions of Ref-1. Am. J. Respir. Cell Mol. Biol. 25, 664–667. Foote, R. S., Mitra, S., and Pal, B. C. (1980). Demethylation of O6-methylguanine in a synthetic DNA polymer by an inducible activity in Escherichia coli. Biochem. Biophys. Res. Commun. 97, 654–659.
32
FROMME AND VERDINE
Fortini, P., Pascucci, B., Parlanti, E., Sobol, R. W., Wilson, S. H., and Dogliotti, E. (1998). Different DNA polymerases are involved in the short- and long-patch base excision repair in mammalian cells. Biochemistry 37, 3575–3580. Francis, A. W., and David, S. S. (2003). Escherichia coli MutY and Fpg utilize a processive mechanism for target location. Biochemistry 42, 801–810. Fritz, G., Grosch, S., Tomicic, M., and Kaina, B. (2003). APE/Ref-1 and the mammalian response to genotoxic stress. Toxicology 193, 67–78. Fritz, G., and Kaina, B. (1999). Phosphorylation of the DNA repair protein APEREF-1 by CKII affects redox regulation of AP-1. Oncogene 18, 1033–1040. Fromme, J. C., Banerjee, A., Huang, S. J., and Verdine, G. L. (2004a). Structural basis for removal of adenine mispaired with 8-oxoguanine by MutY adenine DNA glycosylase. Nature 427, 652–656. Fromme, J. C., Banerjee, A., and Verdine, G. L. (2004b). DNA glycosylase recognition and catalysis. Curr. Opin. Struct. Biol. 14, 43–49. Fromme, J. C., Bruner, S. D., Yang, W., Karplus, M., and Verdine, G. L. (2003). Product-assisted catalysis in base-excision DNA repair. Nature Struc. Biol. 10, 204–211. Fromme, J. C., and Verdine, G. L. (2003a). DNA lesion recognition by the bacterial repair enzyme MutM. J. Biol. Chem. 278, 51543–51548. Fromme, J. C., and Verdine, G. L. (2003b). Structure of a trapped endonuclease IIIDNA covalent intermediate. EMBO J. 22, 3461–3471. Frosina, G., Fortini, P., Rossi, O., Carrozzino, F., Raspaglio, G., Cox, L. S., Lane, D. P., Abbondandolo, A., and Dogliotti, E. (1996). Two pathways for base excision repair in mammalian cells. J. Biol. Chem. 271, 9573–9578. Gaiddon, C., Moorthy, N. C., and Prives, C. (1999). Ref-1 regulates the transactivation and pro-apoptotic functions of p53 in vivo. EMBO J. 18, 5609–5621. Ghavidel, A., and Schultz, M. C. (2001). TATA binding protein-associated CK2 transduces DNA damage signals to the RNA polymerase III transcriptional machinery. Cell 106, 575–584. Girard, P. M., Guibourt, N., and Boiteux, S. (1997). The Ogg1 protein of Saccharomyces cerevisiae: A 7,8-dihydro-8-oxoguanine DNA glycosylase/AP lyase whose lysine 241 is a critical residue for catalytic activity. Nucleic Acids Res. 25, 3204–3211. Glassner, B. J., Rasmussen, L. J., Najarian, M. T., Posnick, L. M., and Samson, L. D. (1998). Generation of a strong mutator phenotype in yeast by imbalanced base excision repair. Proc. Natl. Acad. Sci. USA 95, 9997–10002. Gong, Z., Morales-Ruiz, T., Ariza, R. R., Roldan-Arjona, T., David, L., and Zhu, J.-K. (2002). ROS1, a repressor of transcriptional gene silencing in Arabidopsis, encodes a DNA glycosylase/lyase. Cell 111, 803–814. Grollman, A. P., and Moriya, M. (1993). Mutagenesis by 8-oxoguanine: An enemy within. Trends Genet. 9, 246–249. Guillet, M., and Boiteux, S. (2002). Endogenous DNA abasic sites cause cell death in the absence of Apn1, Apn2 and Rad1/Rad10 in Saccharomyces cerevisiae. EMBO J. 21, 2833–2841. Guillet, M., and Boiteux, S. (2003). Origin of endogenous DNA abasic sites in Saccharomyces cerevisiae. Mol. Cell. Biol. 23, 8386–8394. Hang, B., Singer, B., Margison, G. P., and Elder, R. H. (1997). Targeted deletion of alkylpurine-DNA-N-glycosylase in mice eliminates repair of 1,N6-ethenoadenine and hypoxanthine but not of 3,N4-ethenocytosine or 8-oxoguanine. Proc. Natl. Acad. Sci. USA 94, 12869–12874.
BASE EXCISION REPAIR
33
Hardeland, U., Steinacher, R., Jiricny, J., and Schar, P. (2002). Modification of the human thymine-DNA glycosylase by ubiquitin-like proteins facilitates enzymatic turnover. EMBO J. 21, 1456–1464. Hazra, T. K., Izumi, T., Kow, Y. W., and Mitra, S. (2003). The discovery of a new family of mammalian enzymes for repair of oxidatively damaged DNA, and its physiological implications. Carcinogenesis 24, 155–157. Hendrich, B., Hardeland, U., Ng, H. H., Jiricny, J., and Bird, A. (1999). The thymine glycosylase MBD4 can bind to the product of deamination at methylated CpG sites. Nature 401, 301–304. Higley, M., and Lloyd, R. S. (1993). Processivity of uracil DNA glycosylase. Mutat. Res. 294, 109–116. Higuchi, K., Katayama, T., Iwai, S., Hidaka, M., Horiuchi, T., and Maki, H. (2003). Fate of DNA replication fork encountering a single DNA lesion during oriC plasmid DNA replication in vitro. Genes Cells 8, 437–449. Hollis, T., Ichikawa, Y., and Ellenberger, T. (2000). DNA bending and a flip-out mechanism for base excision by the helix-hairpin-helix DNA glycosylase, Escherichia coli AlkA. EMBO J. 19, 758–766. Holmes, E. W., Bingham, C. M., and Cunningham, M. L. (2002). Hepatic expression of polymerase beta, Ref-1, PCNA, and Bax in WY 14,643-exposed rats and hamsters. Exp. Mol. Pathol. 73, 209–219. Hosfield, D. J., Guan, Y., Haas, B. J., Cunningham, R. P., and Tainer, J. A. (1999). Structure of the DNA repair enzyme endonuclease IV and its DNA complex: Double-nucleotide flipping at abasic sites and three-metal-ion catalysis. Cell 98, 397–408. Hsieh, M. M., Hegde, V., Kelley, M. R., and Deutsch, W. A. (2001). Activation of APERef-1 redox activity is mediated by reactive oxygen species and PKC phosphorylation. Nucleic Acids Res. 29, 3116–3122. Huang, L. E., Arany, Z., Livingston, D. M., and Bunn, H. F. (1996). Activation of hypoxia-inducible transcription factor depends primarily upon redox-sensitive stabilization of its alpha subunit. J. Biol. Chem. 271, 32253–32259. Ikeda, S., Biswas, T., Roy, R., Izumi, T., Boldogh, I., Kurosky, A., Sarker, A. H., Seki, S., and Mitra, S. (1998). Purification and characterization of human NTH1, a homolog of Escherichia coli endonuclease III. Direct identification of Lys-212 as the active nucleophilic residue. J. Biol. Chem. 273, 21585–21593. Izumi, T., Wiederhold, L. R., Roy, G., Roy, R., Jaiswal, A., Bhakat, K. K., Mitra, S., and Hazra, T. K. (2003). Mammalian DNA base excision repair proteins: Their interactions and role in repair of oxidative DNA damage. Toxicology 193, 43–65. Jayaraman, L., Murthy, K. G., Zhu, C., Curran, T., Xanthoudakis, S., and Prives, C. (1997). Identification of redox/repair protein Ref-1 as a potent activator of p53. Genes Dev. 11, 558–570. Jiang, Y. L., Ichikawa, Y., Song, F., and Stivers, J. T. (2003). Powering DNA repair through substrate electrostatic interactions. Biochemistry 42, 1922–1929. Jiang, Y. L., and Stivers, J. T. (2002). Mutational analysis of the base-flipping mechanism of uracil DNA glycosylase. Biochemistry 41, 11236–11247. Jiang, Y. L., Stivers, J. T., and Song, F. (2002). Base-flipping mutations of uracil DNA glycosylase: Substrate rescue using a pyrene nucleotide wedge. Biochemistry 41, 11248–11254. Jones, S., Emmerson, P., Maynard, J., Best, J. M., Jordan, S., Williams, G. T., Sampson, J. R., and Cheadle, J. P. (2002). Biallelic germline mutations in MYH predispose to
34
FROMME AND VERDINE
multiple colorectal adenoma and somatic G:C–>T:A mutations. Hum. Mol. Genet. 11, 2961–2967. Jost, J. P., Oakeley, E. J., Zhu, B., Benjamin, D., Thiry, S., Siegmann, M., and Jost, Y. C. (2001). 5-Methylcytosine DNA glycosylase participates in the genome-wide loss of DNA methylation occurring during mouse myoblast differentiation. Nucleic Acids Res. 29, 4452–4461. Kakolyris, S., Kaklamanis, L., Engels, K., Turley, H., Hickson, I. D., Gatter, K. C., and Harris, A. L. (1997). Human apurinic endonuclease 1 expression in a colorectal adenoma-carcinoma sequence. Cancer Res. 57, 1794–1797. Kinoshita, T., Miura, A., Choi, Y., Kinoshita, Y., Cao, X., Jacobsen, S. E., Fischer, R. L., and Kakutani, T. (2004). One-way control of FWA imprinting in Arabidopsis endosperm by DNA methylation. Science 303, 521–523. Klungland, A., Hoss, M., Gunz, D., Constantinou, A., Clarkson, S. G., Doetsch, P. W., Bolton, P. H., Wood, R. D., and Lindahl, T. (1999a). Base excision repair of oxidative DNA damage activated by XPG protein. Mol. Cell 3, 33–42. Klungland, A., and Lindahl, T. (1997). Second pathway for completion of human DNA base excision-repair: Reconstitution with purified proteins and requirement for DNase IV (FEN1). EMBO J. 16, 3341–3348. Klungland, A., Rosewell, I., Hollenbach, S., Larsen, E., Daly, G., Epe, B., Seeberg, E., Lindahl, T., and Barnes, D. E. (1999b). Accumulation of premutagenic DNA lesions in mice defective in removal of oxidative base damage [In Process Citation]. Proc. Natl. Acad. Sci. USA 96, 13300–13305. Kouchakdjian, M., Bodepudi, V., Shibutani, S., Eisenberg, M., Johnson, F., Grollman, A. P., and Patel, D. J. (1991). NMR structural studies of the ionizing radiation adduct 7-hydro-8-oxodeoxyguanosine (8-oxo-7H-dG) opposite deoxyadenosine in a DNA duplex. 8-Oxo-7H-dG(syn).dA(anti) alignment at lesion site. Biochemistry 30, 1403–1412. Kow, Y. W., and Wallace, S. S. (1987). Mechanism of action of Escherichia coli endonuclease III. Biochemistry 26, 8200–8206. Kubota, Y., Nash, R. A., Klungland, A., Schar, P., Barnes, D. E., and Lindahl, T. (1996). Reconstitution of DNA base excision-repair with purified human proteins: Interaction between DNA polymerase beta and the XRCC1 protein. EMBO J. 15, 6662–6670. Kuo, C. F., McRee, D. E., Fisher, C. L., O’Handley, S. F., Cunningham, R. P., and Tainer, J. A. (1992). Atomic structure of the DNA repair [4Fe-4S] enzyme endonuclease III. Science 258, 434–440. Lau, A. Y., Wyatt, M. D., Glassner, B. J., Samson, L. D., and Ellenberger, T. (2000). Molecular basis for discriminating between normal and damaged bases by the human alkyladenine glycosylase, AAG. Proc. Natl. Acad. Sci. USA 97, 13573–13578. Lavrik, O. I., Prasad, R., Sobol, R. W., Horton, J. K., Ackerman, E. J., and Wilson, S. H. (2001). Photoaffinity labeling of mouse fibroblast enzymes by a base excision repair intermediate. Evidence for the role of poly(ADP-ribose) polymerase-1 in DNA repair. J. Biol. Chem. 276, 25541–25548. Lazebnik, Y. A., Kaufmann, S. H., Desnoyers, S., Poirier, G. G., and Earnshaw, W. C. (1994). Cleavage of poly(ADP-ribose) polymerase by a proteinase with properties like ICE. Nature 371, 346–347. Le Page, F., Klungland, A., Barnes, D. E., Sarasin, A., and Boiteux, S. (2000). Transcription coupled repair of 8-oxoguanine in murine cells: The ogg1 protein is required for repair in nontranscribed sequences but not in transcribed sequences. Proc. Natl. Acad. Sci. USA 97, 8397–8402.
BASE EXCISION REPAIR
35
Li, X., Wright, P. M., and Lu, A. L. (2000). The C-terminal domain of MutY glycosylase determines the 7,8-dihydro-8-oxo-guanine specificity and is crucial for mutation avoidance. J. Biol. Chem. 275, 8448–8455. Lindahl, T. (1993). Instability and decay of the primary structure of DNA. Nature 362, 709–715. Ljungquist, S., and Lindahl, T. (1977). Relation between Escherichia coli endonucleases specific for apurinic sites in DNA and exonuclease III. Nucleic Acids Res. 4, 2871–2879. Loeb, L. A., and Preston, B. D. (1986). Mutagenesis by apurinic/apyrimidinic sites. Annu. Rev. Genet. 20, 201–230. Lu, A. L., Li, X., Gu, Y., Wright, P. M., and Chang, D. Y. (2001). Repair of oxidative DNA damage: Mechanisms and functions. Cell Biochem. Biophys. 35, 141–170. Marenstein, D. R., Chan, M. K., Altamirano, A., Basu, A. K., Boorstein, R. J., Cunningham, R. P., and Teebor, G. W. (2003). Substrate specificity of human endonuclease III (hNTH1). Effect of human APE1 on hNTH1 activity. J. Biol. Chem. 278, 9005–9012. Marsin, S., Vidal, A. E., Sossou, M., Menissier-de Murcia, J., Le Page, F., Boiteux, S., de Murcia, G., and Radicella, J. P. (2003). Role of XRCC1 in the coordination and stimulation of oxidative DNA damage repair initiated by the DNA glycosylase hOGG1. J. Biol. Chem. 278, 44068–44074. Masson, M., Niedergang, C., Schreiber, V., Muller, S., Menissier-de Murcia, J., and de Murcia, G. (1998). XRCC1 is specifically associated with poly(ADP-ribose) polymerase and negatively regulates its activity following DNA damage. Mol. Cell. Biol. 18, 3563–3571. Matsumoto, Y., Kim, K., and Bogenhagen, D. F. (1994). Proliferating cell nuclear antigen-dependent abasic site repair in Xenopus laevis oocytes: An alternative pathway of base excision DNA repair. Mol. Cell. Biol. 14, 6187–6197. Matsumoto, Y., Kim, K., Hurwitz, J., Gary, R., Levin, D. S., Tomkinson, A. E., and Park, M. S. (1999). Reconstitution of proliferating cell nuclear antigen-dependent repair of apurinic/apyrimidinic sites with purified human proteins. J. Biol. Chem. 274, 33703–33708. Mazumder, A., Gerlt, J. A., Absalon, M. J., Stubbe, J., Cunningham, R. P., Withka, J., and Bolton, P. H. (1991). Stereochemical studies of the beta-elimination reactions at aldehydic abasic sites in DNA: Endonuclease III from Escherichia coli, sodium hydroxide, and Lys-Trp-Lys. Biochemistry 30, 1119–1126. McAuley-Hecht, K. E., Leonard, G. A., Gibson, N. J., Thomson, J. B., Watson, W. P., Hunter, W. N., and Brown, T. (1994). Crystal structure of a DNA duplex containing 8-hydroxydeoxyguanine-adenine base pairs. Biochemistry 33, 10266–10270. McCullough, A. K., Sanchez, A., Dodson, M. L., Marapaka, P., Taylor, J. S., and Lloyd, R. S. (2001). The reaction mechanism of DNA glycosylase/AP lyases at abasic sites. Biochemistry 40, 561–568. Memisoglu, A., and Samson, L. (2000). Base excision repair in yeast and mammals. Mutat. Res. 451, 39–51. Millar, C. B., Guy, J., Sansom, O. J., Selfridge, J., MacDougall, E., Hendrich, B., Keightley, P. D., Bishop, S. M., Clarke, A. R., and Bird, A. (2002). Enhanced CpG mutability and tumorigenesis in MBD4-deficient mice. Science 297, 403–405. Minowa, O., Arai, T., Hirano, M., Monden, Y., Nakai, S., Fukuda, M., Itoh, M., Takano, H., Hippou, Y., Aburatani, H., Masumura, K., Nohmi, T., Nishimura, S., and Noda, T. (2000). Mmh/Ogg1 gene inactivation results in accumulation of 8-hydroxyguanine in mice. Proc. Natl. Acad. Sci. USA 97, 4156–4161.
36
FROMME AND VERDINE
Mitomo, K., Nakayama, K., Fujimoto, K., Sun, X., Seki, S., and Yamamoto, K. (1994). Two different cellular redox systems regulate the DNA-binding activity of the p50 subunit of NF-kappa B in vitro. Gene. 145, 197–203. Mol, C. D., Arvai, A. S., Begley, T. J., Cunningham, R. P., and Tainer, J. A. (2002). Structure and activity of a thermostable thymine-DNA glycosylase: Evidence for base twisting to remove mismatched normal DNA bases. J. Mol. Biol. 315, 373–384. Mol, C. D., Hosfield, D. J., and Tainer, J. A. (2000a). Abasic site recognition by two apurinic/apyrimidinic endonuclease families in DNA base excision repair: The 30 ends justify the means. Mutat. Res. 460, 211–229. Mol, C. D., Izumi, T., Mitra, S., and Tainer, J. A. (2000b). DNA-bound structures and mutants reveal abasic DNA binding by APE1 and DNA repair coordination. Nature 403, 451–456. Moore, D. H., Michael, H., Tritt, R., Parsons, S. H., and Kelley, M. R. (2000). Alterations in the expression of the DNA repair/redox enzyme APE/ref-1 in epithelial ovarian cancers. Clin. Cancer Res. 6, 602–609. Morland, I., Rolseth, V., Luna, L., Rognes, T., Bjoras, M., and Seeberg, E. (2002). Human DNA glycosylases of the bacterial Fpg/MutM superfamily: An alternative pathway for the repair of 8-oxoguanine and other oxidation products in DNA. Nucleic Acids Res. 30, 4926–4236. Myrnes, B., Giercksky, K. E., and Krokan, H. (1982). Repair of O6-methyl-guanine residues in DNA takes place by a similar mechanism in extracts from HeLa cells, human liver, and rat liver. J. Cell. Biochem. 20, 381–392. Nash, H. M., Bruner, S. D., Scharer, O. D., Kawate, T., Addona, T. A., Spooner, E., Lane, W. S., and Verdine, G. L. (1996). Cloning of a yeast 8-oxoguanine DNA glycosylase reveals the existence of a base-excision DNA-repair protein superfamily. Curr. Biol. 6, 968–980. Nash, H. M., Lu, R., Lane, W. S., and Verdine, G. L. (1997). The critical active-site amine of the human 8-oxoguanine DNA glycosylase, hOgg1: Direct identification, ablation and chemical reconstitution. Chem. Biol. 4, 693–702. Nicholl, I. D., Nealon, K., and Kenny, M. K. (1997). Reconstitution of human base excision repair with purified proteins. Biochemistry 36, 7557–7566. Nicholson, D. W., Ali, A., Thornberry, N. A., Vaillancourt, J. P., Ding, C. K., Gallant, M., Gareau, Y., Griffin, P. R., Labelle, M., Lazebnik, Y. A., Munday, N. A., Raju, S. M., Smulson, M. E., Yamin, T. -T., Yu, V. L., and Miller, D. K. (1995). Identification and inhibition of the ICE/CED-3 protease necessary for mammalian apoptosis. Nature 376, 37–43. Nilsen, H., Haushalter, K. A., Robins, P., Barnes, D. E., Verdine, G. L., and Lindahl, T. (2001). Excision of deaminated cytosine from the vertebrate genome: Role of the SMUG1 uracil-DNA glycosylase. EMBO J. 20, 4278–4286. Nilsen, H., Lindahl, T., and Verreault, A. (2002). DNA base excision repair of uracil residues in reconstituted nucleosome core particles. EMBO J. 21, 5943–5952. Nilsen, H., Rosewell, I., Robins, P., Skjelbred, C. F., Andersen, S., Slupphaug, G., Daly, G., Krokan, H. E., Lindahl, T., and Barnes, D. E. (2000). Uracil-DNA glycosylase (UNG)deficient mice reveal a primary role of the enzyme during DNA replication. Mol. Cell 5, 1059–1065. Nilsen, H., Stamp, G., Andersen, S., Hrivnak, G., Krokan, H. E., Lindahl, T., and Barnes, D. E. (2003). Gene-targeted mice lacking the Ung uracil-DNA glycosylase develop B-cell lymphomas. Oncogene 22, 5381–5386. Noll, D. M., Gogos, A., Granek, J. A., and Clarke, N. D. (1999). The C-terminal domain of the adenine-DNA glycosylase MutY confers specificity for 8-oxoguanine.adenine
BASE EXCISION REPAIR
37
mispairs and may have evolved from MutT, an 8-oxo-dGTPase. Biochemistry 38, 6374–6379. O’Brien, P. J., and Ellenberger, T. (2003). Dissecting the broad substrate specificity of human 3-methyladenine DNA glycosylase. J. Biol. Chem. 279, 9750–9757. Ocampo, M. T., Chaung, W., Marenstein, D. R., Chan, M. K., Altamirano, A., Basu, A. K., Boorstein, R. J., Cunningham, R. P., and Teebor, G. W. (2002). Targeted deletion of mNth1 reveals a novel DNA repair enzyme activity. Mol. Cell. Biol. 22, 6111–6121. Offer, H., Milyavsky, M., Erez, N., Matas, D., Zurer, I., Harris, C. C., and Rotter, V. (2001a). Structural and functional involvement of p53 in BER in vitro and in vivo. Oncogene. 20, 581–589. Offer, H., Zurer, I., Banfalvi, G., Reha’k, M., Falcovitz, A., Milyavsky, M., Goldfinger, N., and Rotter, V. (2001b). p53 modulates base excision repair activity in a cell cyclespecific manner after genotoxic stress. Cancer Res. 61, 88–96. Okazaki, T., Chung, U., Nishishita, T., Ebisu, S., Usuda, S., Mishiro, S., Xanthoudakis, S., Igarashi, T., and Ogata, E. (1994). A redox factor protein, ref1, is involved in negative gene regulation by extracellular calcium. J. Biol. Chem. 269, 27855–27862. Olsson, M., and Lindahl, T. (1980). Repair of alkylated DNA in Escherichia coli. Methyl group transfer from O6-methylguanine to a protein cysteine residue. J. Biol. Chem. 255, 10569–10571. Parikh, S. S., Mol, C. D., Slupphaug, G., Bharati, S., Krokan, H. E., and Tainer, J. A. (1998). Base excision repair initiation revealed by crystal structures and binding kinetics of human uracil-DNA glycosylase with DNA. EMBO J. 17, 5214–5226. Parikh, S. S., Putnam, C. D., and Tainer, J. A. (2000). Lessons learned from structural results on uracil-DNA glycosylase. Mutat. Res. 460, 183–199. Parker, A., Gu, Y., Mahoney, W., Lee, S. H., Singh, K. K., and Lu, A. L. (2001). Human homolog of the MutY repair protein (hMYH) physically interacts with proteins involved in long patch DNA base excision repair. J. Biol. Chem. 276, 5547–5555. Parsons, J. L., and Elder, R. H. (2003). DNA N-glycosylase deficient mice: A tale of redundancy. Mutat. Res. 531, 165–175. Pascucci, B., Stucki, M., Jonsson, Z. O., Dogliotti, E., and Hubscher, U. (1999). Long patch base excision repair with purified human proteins. DNA ligase I as patch size mediator for DNA polymerases delta and epsilon. J. Biol. Chem. 274, 33696–33702. Pierson, C. E., Prasad, R., Wilson, S. H., and Lloyd, R. S. (1996). Evidence for an imino intermediate in the DNA polymerase beta deoxyribose phosphate excision reaction. J. Biol. Chem. 271, 17811–17815. Piersen, C. E., Prince, M. A., Augustine, M. L., Dodson, M. L., and Lloyd, R. S. (1995). Purification and cloning of Micrococcus luteus ultraviolet endonuclease, an N-glycosylase/abasic lyase that proceeds via an imino enzyme-DNA intermediate. J. Biol. Chem. 270, 23475–23484. Popoff, S. C., Spira, A. I., Johnson, A. W., and Demple, B. (1990). Yeast structural gene (APN1) for the major apurinic endonuclease: Homology to Escherichia coli endonuclease IV. Proc. Natl. Acad. Sci. USA 87, 4193–4197. Pourquier, P., Ueng, L. M., Kohlhagen, G., Mazumder, A., Gupta, M., Kohn, K. W., and Pommier, Y. (1997). Effects of uracil incorporation, DNA mismatches, and abasic sites on cleavage and religation activities of mammalian topoisomerase I. J. Biol. Chem. 272, 7792–7796. Prasad, R., Beard, W. A., Chyan, J. Y., Maciejewski, M. W., Mullen, G. P., and Wilson, S. H. (1998). Functional analysis of the amino-terminal 8-kDa domain of DNA polymerase beta as revealed by site-directed mutagenesis. DNA binding and 50 -deoxyribose phosphate lyase activities. J. Biol. Chem. 273, 11121–11126.
38
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Prasad, R., Lavrik, O. I., Kim, S. J., Kedar, P., Yang, X. P., Vande Berg, B. J., and Wilson, S. H. (2001). DNA polymerase beta-mediated long patch base excision repair. Poly(ADP-ribose)polymerase-1 stimulates strand displacement DNA synthesis. J. Biol. Chem. 276, 32411–32414. Prasad, R., Singhal, R. K., Srivastava, D. K., Molina, J. T., Tomkinson, A. E., and Wilson, S. H. (1996). Specific interaction of DNA polymerase beta and DNA ligase I in a multiprotein base excision repair complex from bovine testis. J. Biol. Chem. 271, 16000–16007. Robertson, K. A., Bullock, H. A., Xu, Y., Tritt, R., Zimmerman, E., Ulbright, T. M., Foster, R. S., Einhorn, L. H., and Kelley, M. R. (2001). Altered expression of Ape1/ ref-1 in germ cell tumors and overexpression in NT2 cells confers resistance to bleomycin and radiation. Cancer Res. 61, 2220–2225. Rogers, S. G., and Weiss, B. (1980). Exonuclease III of Escherichia coli K-12, an AP endonuclease. Methods Enzymol. 65, 201–211. Sakumi, K., Tominaga, Y., Furuichi, M., Xu, P., Tsuzuki, T., Sekiguchi, M., and Nakabeppu, Y. (2003). Ogg1 knockout-associated lung tumorigenesis and its suppression by Mth1 gene disruption. Cancer Res. 63, 902–905. Sampson, J. R., Dolwani, S., Jones, S., Eccles, D., Ellis, A., Evans, D. G., Frayling, I., Jordan, S., Maher, E. R., Mak, T., Maynard, J., Pigatto, F., Shaw, J., and Cheadle, J. P. (2003). Autosomal recessive colorectal adenomatous polyposis due to inherited mutations of MYH. Lancet 362, 39–41. Sansom, O. J., Zabkiewicz, J., Bishop, S. M., Guy, J., Bird, A., and Clarke, A. R. (2003). MBD4 deficiency reduces the apoptotic response to DNA-damaging agents in the murine small intestine. Oncogene 22, 7130–7136. Seawell, P. C., Smith, C. A., and Ganesan, A. K. (1980). den V gene of bacteriophage T4 determines a DNA glycosylase specific for pyrimidine dimers in DNA. J. Virol. 35, 790–796. Sedgwick, B. (2004). Repairing DNA-methylation damage. Nature Rev. Mol. Cell. Biol. 5, 148–157. Shibutani, S., Takeshita, M., and Grollman, A. P. (1991). Insertion of specific bases during DNA synthesis past the oxidation-damaged base 8-oxodG. Nature 349, 431–434. Sidorkina, O. M., and Laval, J. (2000). Role of the N-terminal proline residue in the catalytic activities of the Escherichia coli Fpg protein. J. Biol. Chem. 275, 9924–9929. Sieber, O. M., Lipton, L., Crabtree, M., Heinimann, K., Fidalgo, P., Phillips, R. K., Bisgaard, M. L., Orntoft, T. F., Aaltonen, L. A., Hodgson, S. V., Thomas, H. J., and Tomlinson, I. P. (2003). Multiple colorectal adenomas, classic adenomatous polyposis, and germ-line mutations in MYH. N. Engl. J. Med. 348, 791–799. Slupphaug, G., Mol, C. D., Kavli, B., Arvai, A. S., Krokan, H. E., and Tainer, J. A. (1996). A nucleotide-flipping mechanism from the structure of human uracil-DNA glycosylase bound to DNA. Nature 384, 87–92. Stivers, J. T., and Drohat, A. C. (2001). Uracil DNA glycosylase: Insights from a master catalyst. Arch. Biochem. Biophys. 396, 1–9. Stivers, J. T., and Jiang, Y. L. (2003). A mechanistic perspective on the chemistry of DNA repair glycosylases. Chem. Rev. 103, 2729–2759. Sun, B., Latham, K. A., Dodson, M. L., and Lloyd, R. S. (1995). Studies on the catalytic mechanism of five DNA glycosylases. Probing for enzyme-DNA imino intermediates. J. Biol. Chem. 270, 19501–19508.
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39
Sved, J., and Bird, A. (1990). The expected equilibrium of the CpG dinucleotide in vertebrate genomes under a mutation model. Proc. Natl. Acad. Sci. USA 87, 4692–4696. Takao, M., Kanno, S., Kobayashi, K., Zhang, Q. M., Yonei, S., van der Horst, G. T., and Yasui, A. (2002a). A back-up glycosylase in Nth1 knock-out mice is a functional Nei (endonuclease VIII) homologue. J. Biol. Chem. 277, 42205–42213. Takao, M., Kanno, S., Shiromoto, T., Hasegawa, R., Ide, H., Ikeda, S., Sarker, A. H., Seki, S., Xing, J. Z., Le, X. C., Weinfeld, M., Kobayashi, K., Miyazaki, J., Muijtjens, M., Hoeijmakers, J. H., van der Horst, G., and Yasui, A. (2002b). Novel nuclear and mitochondrial glycosylases revealed by disruption of the mouse Nth1 gene encoding an endonuclease III homolog for repair of thymine glycols. EMBO J. 21, 3486–3493. Tchou, J., and Grollman, A. P. (1995). The catalytic mechanism of Fpg protein. Evidence for a Schiff base intermediate and amino terminus localization of the catalytic site. J. Biol. Chem. 270, 11671–11677. Thayer, M. M., Ahern, H., Xing, D., Cunningham, R. P., and Tainer, J. A. (1995). Novel DNA binding motifs in the DNA repair enzyme endonuclease III crystal structure. EMBO J. 14, 4108–4120. Thompson, L. H., and West, M. G. (2000). XRCC1 keeps DNA from getting stranded. Mutat. Res. 459, 1–18. Tini, M., Benecke, A., Um, S. J., Torchia, J., Evans, R. M., and Chambon, P. (2002). Association of CBPp300 acetylase and thymine DNA glycosylase links DNA repair and transcription. Mol. Cell 9, 265–277. Vassylyev, D. G., Kashiwagi, T., Mikami, Y., Ariyoshi, M., Iwai, S., Ohtsuka, E., and Morikawa, K. (1995). Atomic model of a pyrimidine dimer excision repair enzyme complexed with a DNA substrate: Structural basis for damaged DNA recognition. Cell 83, 773–782. Verdine, G. L., and Bruner, S. D. (1997). How do DNA repair proteins locate damaged bases in the genome? Chem. Biol. 4, 329–334. Vidal, A. E., Boiteux, S., Hickson, I. D., and Radicella, J. P. (2001a). XRCC1 coordinates the initial and late stages of DNA abasic site repair through protein-protein interactions. EMBO J. 20, 6530–6539. Vidal, A. E., Hickson, I. D., Boiteux, S., and Radicella, J. P. (2001b). Mechanism of stimulation of the DNA glycosylase activity of hOGG1 by the major human AP endonuclease: Bypass of the AP lyase activity step. Nucleic Acids Res. 29, 1285–1292. Walker, L. J., Robson, C. N., Black, E., Gillespie, D., and Hickson, I. D. (1993). Identification of residues in the human DNA repair enzyme HAP1 (Ref-1) that are essential for redox regulation of Jun DNA binding. Mol. Cell. Biol. 13, 5370–5376. Waters, T. R., Gallinari, P., Jiricny, J., and Swann, P. F. (1999). Human thymine DNA glycosylase binds to apurinic sites in DNA but is displaced by human apurinic endonuclease 1. J. Biol. Chem. 274, 67–74. Wei, S. J., Botero, A., Hirota, K., Bradbury, C. M., Markovina, S., Laszlo, A., Spitz, D. R., Goswami, P. C., Yodoi, J., and Gius, D. (2000). Thioredoxin nuclear translocation and interaction with redox factor-1 activates the activator protein-1 transcription factor in response to ionizing radiation. Cancer Res. 60, 6688–6695. Weiss, B. (1976). Endonuclease II of Escherichia coli is exonuclease III. J. Biol. Chem. 251, 1896–1901.
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Weiss, B., and Grossman, L. (1987). Phosphodiesterases involved in DNA repair. Adv. Enzymol. 60, 1–34. Werner, R. M., and Stivers, J. T. (2000). Kinetic isotope effect studies of the reaction catalyzed by uracil DNA glycosylase: Evidence for an oxocarbenium ion-uracil anion intermediate. Biochemistry 39, 14054–14064. Wibley, J. E., Waters, T. R., Haushalter, K., Verdine, G. L., and Pearl, L. H. (2003). Structure and specificity of the vertebrate anti-mutator uracil-DNA glycosylase SMUG1. Mol. Cell. 11, 1647–1659. Wilson, D. M., 3rd, and Barsky, D. (2001). The major human abasic endonuclease: Formation, consequences and repair of abasic lesions in DNA. Mutat. Res. 485, 283–307. Wilson, S. H. (1998). Mammalian base excision repair and DNA polymerase beta. Mutat Res. 407, 203–215. Wilson, S. H., and Kunkel, T. A. (2000). Passing the baton in base excision repair. Nature Struc. Biol. 7, 176–178. Wu, P., Qiu, C., Sohail, A., Zhang, X., Bhagwat, A. S., and Cheng, X. (2003). Mismatch repair in methylated DNA. Structure and activity of the mismatch-specific thymine glycosylase domain of methyl-CpG-binding protein MBD4. J. Biol. Chem. 278, 5285–5291. Wu, X., Li, J., Li, X., Hsieh, C. L., Burgers, P. M., and Lieber, M. R. (1996). Processing of branched DNA intermediates by a complex of human FEN-1 and PCNA. Nucleic Acids Res. 24, 2036–2043. Wyatt, M. D., Allan, J. M., Lau, A. Y., Ellenberger, T. E., and Samson, L. D. (1999). 3-methyladenine DNA glycosylases: structure, function, and biological importance. Bioessays 21, 668–676. Xanthoudakis, S., Miao, G., Wang, F., Pan, Y. C., and Curran, T. (1992). Redox activation of Fos-Jun DNA binding activity is mediated by a DNA repair enzyme. EMBO J. 11, 3323–3335. Xanthoudakis, S., Smeyne, R. J., Wallace, J. D., and Curran, T. (1996). The redox/DNA repair protein, Ref-1, is essential for early embryonic development in mice. Proc. Natl. Acad. Sci. USA 93, 8919–8923. Xiao, W., Gehring, M., Choi, Y., Margossian, L., Pu, H., Harada, J. J., Goldberg, R. B., Pennell, R. I., and Fischer, R. L. (2003). Imprinting of the MEA Polycomb gene is controlled by antagonism between MET1 methyltransferase and DME glycosylase. Dev. Cell. 5, 891–901. Yakovlev, A. G., Wang, G., Stoica, B. A., Boulares, H. A., Spoonde, A. Y., Yoshihara, K., and Smulson, M. E. (2000). A role of the Ca2þ/Mg2þ-dependent endonuclease in apoptosis and its inhibition by Poly(ADP-ribose) polymerase. J. Biol. Chem. 275, 21302–21308. Yoshida, K., Wang, H. G., Miki, Y., and Kufe, D. (2003). Protein kinase Cdelta is responsible for constitutive and DNA damage-induced phosphorylation of Rad9. EMBO J. 22, 1431–1441. Yu, S. L., Lee, S. K., Johnson, R. E., Prakash, L., and Prakash, S. (2003). The stalling of transcription at abasic sites is highly mutagenic. Mol. Cell. Biol. 23, 382–388. Zharkov, D. O., Golan, G., Gilboa, R., Fernandes, A. S., Gerchman, S. E., Kycia, J. H., Rieger, R. A., Grollman, A. P., and Shoham, G. (2002). Structural analysis of an Escherichia coli endonuclease VIII covalent reaction intermediate. EMBO J. 21, 789–800. Zharkov, D. O., Rieger, R. A., Iden, C. R., and Grollman, A. P. (1997). NH2-terminal proline acts as a nucleophile in the glycosylase/AP-lyase reaction catalyzed by
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Escherichia coli formamidopyrimidine-DNA glycosylase (Fpg) protein. J. Biol. Chem. 272, 5335–5341. Zhou, J., Ahn, J., Wilson, S. H., and Prives, C. (2001). A role for p53 in base excision repair. EMBO J. 20, 914–923. Zhu, B., Zheng, Y., Hess, D., Angliker, H., Schwarz, S., Siegmann, M., Thiry, S., and Jost, J. P. (2000). 5-methylcytosine-DNA glycosylase activity is present in a cloned GT mismatch DNA glycosylase associated with the chicken embryo DNA demethylation complex. Proc. Natl. Acad. Sci. USA 97, 5135–5139. Zurer, I., Hofseth, L. J., Cohen, Y., Xu-Welliver, M., Hussain, S. P., Harris, C. C., and Rotter, V. (2004). The role of p53 in base excision repair following genotoxic stress. Carcinogenesis 25, 11–19.
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NUCLEOTIDE EXCISION REPAIR IN E. COLI AND MAN By AZIZ SANCAR AND JOYCE T. REARDON Department of Biochemistry and Biophysics, University of North Carolina School of Medicine, Chapel Hill, North Carolina, 27599
I. Introduction . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . II. Damage Recognition . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . III. Mechanism of Excision Repair.. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Excision Repair in E. coli . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. Excision Repair in Humans . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . IV. Transcription-Coupled Repair. .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Transcription-Coupled Repair in E. coli . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. Transcription-Coupled Repair in Human Cells .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . V. Repair of Chromatin. . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . VI. Conclusion . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . References . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . .
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I. INTRODUCTION Nucleotide excision repair (excision repair) is a universal repair system that eliminates DNA damage by dual incisions bracketing the lesion. In nucleotide excision repair, the damage is removed in the form of a 12–13-nucleotide (nt)-long oligomer in prokaryotes and in a 24–32-nt-long oligomer in eukaryotes (Huang et al., 1992; Sancar and Rupp, 1983) (Fig. 1) Excision repair comprises three basic steps: damage recognition, dual incisions and release of the excised oligomer, and resynthesis to fill in the gap and ligation (Sancar, 1996; Sancar et al., 2004; Wood, 1997). Nucleotide excision repair is the primary repair system for bulky DNA adducts such as the cyclobutane pyrimidine dimer (Pyr<>Pyr), (6–4) photoproduct, benzo[a]pyrene-guanine adduct, acetylaminofluorene-guanine (AAF-G), and cisplatin-d(GpG) diadduct. In addition, it plays a back-up role for base excision repair by removing nonbulky DNA lesions such as thymine glycols and 8-oxoguanine at a slow but physiologically relevant rate. Finally, excision repair is required for interstrand cross-link repair in Escherichia coli (Van Houten et al., 1986) and participates in one pathway of cross-link repair in yeast and humans (Bessho et al., 1997; Zheng et al., 2003). A defect in nucleotide excision repair causes extreme ultraviolet (UV) sensitivity in E. coli and Saccharomyces cerevisiae. In humans, defects in 43 ADVANCES IN PROTEIN CHEMISTRY, Vol. 69
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FIG. 1. Schematic illustration of nucleotide excision repair in prokaryotes and eukaryotes. The basic steps are conserved: damage recognition and dual incisions to excise DNA damage, helicase activity to displace excised oligomer and repair factors, and resynthesis/ligation to restore the integrity of the DNA molecule. (See Color Insert.)
excision repair cause the inherited disease xeroderma pigmentosum (XP) (Cleaver, 1968). There are eight XP complementation groups, XP-A through XP-G and XP-V (variant), and a mutation in any of the genes can cause the disease. The signs and symptoms of XP include extreme sensitivity to sunlight, about 10,000-fold increase in skin cancer, and mental and developmental abnormalities in some cases (Cleaver and Kraemer, 1989). Transcription stimulates excision repair both in E. coli (Mellon and Hanawalt, 1989) and in humans (Bohr et al., 1985; Mellon et al., 1987) in a process dependent on proteins called transcription-repair coupling factors (TRCFs). In E. coli the mfd gene encodes the TRCF (Selby and Sancar, 1991, 1993; Selby et al., 1991), and cells mutated in this gene exhibit modest UV sensitivity but a disproportionately increased rate of UV-induced mutations and lack of mutation frequency decline (mfd) on holding in minimal medium after irradiation and before plating (Witkin, 1994). In humans, mutations in the CSA and CSB genes abolish transcriptioncoupled repair (Venema et al., 1990a). Cells from Cockayne Syndrome (CS) patients exhibit phenotypic properties similar to E. coli mfd mutants, which
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include moderate UV sensitivity and an increased rate of UV mutations that are mostly caused by lesions in the template strand of transcribed genes (see Hanawalt, 2002). CS is associated with developmental retardation and neurological abnormalities but not with an increased incidence of skin cancer, although the patients are hypersensitive to sunlight. In addition to mutations in the CSA and CSB genes, some mutations in XPB, XPD, and XPG also give rise to Cockayne syndrome. In these latter cases, the patients exhibit a mixed XP/CS phenotype. Because XPB, XPD, and XPG are involved in transcription in addition to repair and CS is never observed in the strictly repair-defective XP-A cases, CS is considered a transcription defect syndrome rather than a repair deficiency disease. Finally, a rare genetic disease called trichothiodystrophy (TTD) is also associated with mutations in the XPB and XPD genes. TTD patients have scaly, fishlike skin and brittle hair and nails as a result of a defect in synthesis of sulfur-rich proteins. TTD patients exhibit some of the developmental and neurological abnormalities associated with Cockayne syndrome. In this chapter, we will first address the general problem of damage recognition specificity by an enzyme system with an essentially infinite substrate range. Then the mechanisms of DNA repair by the E. coli and human excision nucleases, as representatives of the prokaryotic and eukaryotic excision nucleases, will be summarized. The mechanism of excision repair in S. cerevisiae, which is quite similar to human excision repair, has been reviewed elsewhere (Prakash and Prakash, 2000) and will not be discussed here.
II. DAMAGE RECOGNITION The biological dilemma in DNA repair is the virtually infinite number of DNA lesions that can and often do form within the lifespan of the cell, the necessity of repairing these lesions within a lifetime, and the theoretical limitation on what fraction of the genomic encoding capacity can be dedicated to repair. It appears that in nature, this problem has been solved by two main approaches. In one, as in the case of photolyase, a single enzyme of near-absolute specificity repairs a single, but relatively abundant, DNA lesion (Sancar, 2003). In the second general scheme, either a single polypeptide (O’Brien and Ellenberger, 2004) or a multiprotein enzyme (Sancar et al., 2004) acts on many, often structurally dissimilar, substrates. The excision nuclease falls into the category of a multiprotein system with an infinite substrate range, and it must deal with the physiological necessity of removing all types of lesions, the biological imperative to avoid normal bases, and the evolutionary requirement of keeping the mutation load at an optimum.
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Three strategies are used by excision nucleases to achieve physiologically acceptable specificity at a biologically relevant rate: cooperativity, molecular matchmaking, and kinetic proofreading (Fig. 2). In cooperative DNA–protein interactions, the binding of one protein to DNA facilitates the binding of either the second subunit of the same protein (homotropic) or of an unrelated protein (heterotropic) by protein–protein interaction. The binding sites of the monomers could be adjacent or overlapping, and cooperative binding may involve more than two proteins. In qualitative terms, the binding of one protein to DNA facilitates the binding of a second protein that has a binding site adjacent to the first one by increasing the local protein concentration of the second one by specific protein– protein interactions. Quantitatively, the binding of the second protein is enhanced by the first protein by a factor approximately equal to the square root of the equilibrium association constant of the two proteins (Giedroc et al., 1987; Kelly et al., 1976). Cooperative DNA binding is important for all forms of DNA–protein interactions in both prokaryotes and eukaryotes,
FIG. 2. Three mechanisms used to achieve specificity in DNA damage recognition and repair. (A) Cooperative binding enhances the binding of a second protein to DNA. (B) Kinetic proofreading achieves specificity by using energy to introduce irreversible steps (a time delay) between ES and E þ P steps. (C) A molecular matchmaker promotes the binding of a second protein that is not able to specifically bind DNA on its own. (See Color Insert.)
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and it is the predominant transcriptional regulatory mechanism in eukaryotes (Levine and Tjian, 2003). In molecular matchmaking, a protein uses the energy released from ATP hydrolysis to promote the DNA binding of another protein that is incapable of binding specifically on its own. A molecular matchmaker is a protein that brings two compatible yet solitary macromolecules together in an ATP-dependent reaction, promotes their association, and then leaves the complex so that the new complex can engage in productive transactions (Orren et al., 1992; Sancar and Hearst, 1993). Because several types of proteins, including adaptors and mediators, are known to facilitate DNA–protein interactions, a molecular matchmaker has been defined narrowly to differentiate it from other facilitators. A protein must fulfill five criteria to be identified as a matchmaker (Sancar and Hearst, 1993). First, in the absence of the matchmaker, the affinity of the matched protein to its target DNA site must be low enough to be physiologically insignificant. Second, the matchmaker must promote stable complex formation between DNA and the cognate protein. Third, the matchmaker or the cognate protein must hydrolyze ATP to generate the energy needed for complex formation. Fourth, the matchmaker must make a ternary complex with the matched DNA and protein, causing a conformational change but no covalent modification. Fifth, after stable complex formation, the matchmaker must dissociate to allow the matched protein to carry out its effector function. In nucleotide excision repair, UvrA in E. coli and XPC in humans fulfill all five criteria of molecular matchmakers. Molecular matchmaking provides a stepwise recognition mechanism and ATP hydrolysis-dependent exposure of the interacting groups on DNA and protein to achieve specificity. However, neither cooperativity nor molecular matchmaking are sufficient to confer the specificity and rate requisite for a biological system. Thus, a third mechanism, kinetic proofreading, complements these thermodynamic molecular mechanisms to achieve the necessary specificity. Kinetic proofreading is the biochemical version of information theory, whereby energy is converted into information. Kinetic proofreading is a specificity mechanism that achieves high fidelity beyond the level that can be achieved by the free-energy difference between correct and incorrect interactions, through the presence of (unidirectional) energy-using intermediate steps at each of which the reaction can be aborted (Hopfield, 1974; Ninio, 1975). This specificity mechanism is equivalent to the introduction of a time delay between ES complex formation and the formation of product in an otherwise normal Michaelis–Menton scheme. When applied to specific biological systems, there are variations on the original scheme (Burgess and Guthrie, 1993; MacGlashan, 2001).
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However, for a particular cellular response to occur, kinetic proofreading in general requires that the enzyme–substrate (or DNA–protein) complex complete a series of irreversible reactions (such as ATP hydrolysis or protein phosphorylation) while a moderate-specifcity ligand is bound; this temporally separates the binding step from the effector step (in nucleotide excision repair, the dual incision event). If the ligand dissociates before the full set of either covalent modifications or conformational changes is completed, the reaction is aborted and must restart at the beginning. Thus, the kinetic proofreading mechanism differs from other multiplestep kinetic reaction schemes of the Michaelis–Menton type in which every step but the ultimate one (EP ! E þ P) is reversible. As an illustration of the power of kinetic proofreading in conferring specificity, a 10-fold difference between the off rates (e.g., 0.1 s1 and 1.0 s1) between specific and nonspecific complexes can provide 104-fold difference in the effector reaction by interposing five steps between binding and catalysis steps. In nucleotide excision repair, UvrA and UvrB in E. coli and XPB and XPD in humans are ATPases that hydrolyze ATP to create irreversible intermediates between the initial low-specificity binding and the final dualincision steps and, thus, perform kinetic proofreading. It must be obvious, however, from this brief description of kinetic proofreading that the specificity conferred by this mechanism is not absolute. Actually, some degree of nonspecific effector reaction is intrinsic to the kinetic proofreading scheme, and the production of side products is a hallmark of a reaction pathway that employs kinetic proofreading. In the case of nucleotide excision repair, the kinetic proofreading scheme would predict excision of undamaged bases. Indeed, both in E. coli and in humans, excision repair acts on undamaged DNA excising damage-free oligomers 12–13 nt and 24–32 nt in length, respectively (Branum et al., 2001).
III. MECHANISM OF EXCISION REPAIR In E. coli, dual incisions are accomplished by three proteins (UvrA, UvrB, and UvrC), and in humans 15 polypeptides in six repair factors carry out the same task. The properties of the E. coli and human excision repair factors are summarized in Tables I and II, respectively. Of significance, in contrast to all other repair systems, the prokaryotic and eukaryotic excision repair factors are evolutionarily not related and show no sequence homology to one another. However, the basic strategies for the prokaryotic and eukaryotic excision nucleases are similar. First, damage is recognized by an ATP-independent mechanism to form an unstable DNA–protein complex. Then this complex is converted to a stable preincision form by ATPase subunits that hydrolyze ATP and unwind the
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TABLE I Subunits of the Escherichia coli Excision Nuclease Protein
Mr
Motif/Domain
Function
UvrA
(104)2
UvrB
78
ABC superfamily 2 zinc fingers Helicase motif
UvrC
69
Damage recognition Molecular matchmaker Damage recognition DNA unwinding Contributes to 30 incision GIY-YIG: 30 incision EndoV: 50 incision
GIY-YIG nuclease EndoV nuclease
TABLE II Subunits of the Human Excision Nuclease Factor XPA RPA
XPC TFIIH
XPG XPFERCC1
Proteins XPA/p31 p70 p32 p14 p106 HR23B/p58 XPB/ERCC3/p89 XPD/ERCC2/p80 p62 p52 p44 p34 XPG/ERCC5/p135 XPF/ERCC4/p112 ERCC1/p33
Activity
Role in repair
DNA binding DNA binding Replication factor
Damage recognition Damage recognition
DNA binding
Damage recognition Molecular matchmaker DNA unwinding Kinetic proofreading
DNA-dependent ATPase Helicase General transcription factor Nuclease Nuclease
30 incision 50 incision
duplex to promote the formation of more intimate protein–DNA contacts. Finally, the dual incisions are made in a concerted, but asynchronous, manner such that the 30 incision precedes the 50 incision.
A. Excision Repair in E. coli In E. coli, the dual-incision activity is carried out by the sequential and partially overlapping functions of UvrA, UvrB, and UvrC (Fig. 3), and the activity is referred to as (A)BC excision nuclease or (A)BC excinuclease (Orren and Sancar, 1989; Sancar and Rupp, 1983). The name is descriptive of the reaction performed by the enzyme system (excising an
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FIG. 3. Model for excision repair in E. coli. UvrA dimerizes (cooperativity) and interacts with UvrB to form an A2B1 heterotrimer, the damage recognition factor. UvrA delivers UvrB to the damage site and then dissociates (molecular matchmaker). UvrC recognizes and binds to the UvrB–DNA complex, in which the DNA is bent and locally unwound. ATP hydrolysis introduces irreversible intermediates at steps along the pathway leading to dual incision, and the reaction may be aborted at any step (kinetic proofreading). Dual incisions release the damage in a 12–13-nt-long oligomer. UvrD (Helicase II) displaces UvrC and the excised oligomer, and then DNA polymerase I displaces UvrB during resynthesis to fill in the gap; newly synthesized DNA is ligated to complete the repair reaction. (See Color Insert.)
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oligomer from DNA) and is preferable over other names such as UvrABC endonuclease or UvrABC nuclease, which give the connotation of nicking or degrading the DNA. In an ATP-dependent reaction, UvrA and UvrB make an A2B1 heterotrimer that is the damage recognition complex (Orren and Sancar, 1989). The damaged DNA preference of this complex is conferred by UvrA, which even on its own is capable of preferentially binding to bulky lesions such as a psoralen-thymine monoadduct (Van Houten et al., 1987) and acetylaminofluorene-guanine adduct (Bertrand-Burggraf et al., 1991; Delagoutte et al., 1997, 2002). Thus, the A2B1 complex, guided by the preference of UvrA for damaged DNA, binds at the damage site and the ‘‘helicase’’ activity of UvrB unwinds DNA at this site by about 5 bp and kinks it by 130 degrees. This kinking and unwinding is accompanied by significant conformational changes in UvrB as well (Delagoutte et al., 2002; Goosen et al., 1998; Hsu et al., 1995; Shi et al., 1992; Zou and Van Houten, 1999). These changes lead to formation of a stable UvrB–DNA complex and dissociation of (UvrA)2 from the DNA (Orren and Sancar, 1989; 1990). Once UvrA dissociates, UvrC binds with high affinity and specificity to the B1-DNA complex, and the dual incisions are made (Orren and Sancar, 1989; Orren et al., 1992). The 30 incision is made by the GIY-YIG homing endonucleaserelated N-terminal domain of UvrC (Aravind et al., 1999; Kowalski et al., 1999; Verhoeven et al., 2000) with participation of some residues from UvrB (Lin and Sancar, 1991; Lin et al., 1992). This is rapidly followed by the 50 incision made by the catalytic ‘‘triad’’ comprising Asp399, Asp438, Asp466, and His538 in the C-terminal EndoV domain of UvrC (Lin and Sancar, 1992). The 30 incision is made at the fourth through the sixth phosphodiester bond, and the 50 incision is made at the eighth phosphodiester bond. Thus, regardless of the type of damage, the lesion is excised in the form of a 12–13-nt-long oligomer. Interestingly, E. coli and some other bacteria contain a protein that is homologous to the N-terminal half of UvrC (Moolenaar et al., 2002). This protein is called Cho (UvrC homologue) and, acting in conjunction with UvrA and UvrB, makes the 30 incision at the ninth phosphodiester bond 30 to the lesion but cannot make the 50 incision. The deletion of cho has no measurable effect on UV survival of wild-type E. coli and only a minor effect on E. coli lacking the UvrC protein. It appears that Cho, working coordinately with UvrC, makes some contribution to DNA repair by the (A)BC excinuclease. Following the dual incisions, UvrB, UvrC, and the ‘‘excised’’ oligomer remain in the postincision complex, although the excised oligomer is no longer hydrogen-bonded to the complementary strand (Orren et al., 1992). UvrC is not very stably bound in this complex, and its dissociation is
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accelerated by UvrD (helicase II), which also releases the excised oligomer. UvrB remains bound in the excision gap and is displaced by DNA polymerase I concomitant with gap filling to produce a repair patch exactly matching the size of the excised oligomer (Sibghat-Ullah et al., 1990). Finally, the newly synthesized DNA is ligated to complete the repair reaction. As noted above, the (A)BC excinuclease has a virtually infinite substrate range; however, it is most effective on lesions such as (6–4) photoproducts and AAF–guanine adducts that destabilize the duplex, as well as, interestingly, on the thymine–psoralen monoadduct that stabilizes the helix (see Petit and Sancar, 1999). The enzyme system uses cooperativity, molecular matchmaking, and kinetic proofreading to achieve specificity. The cooperative interactions include facilitation of UvrA binding to DNA by photolyase (Sancar et al., 1984) and the dimerization of UvrA on DNA. UvrA is the molecular matchmaker that delivers UvrB to DNA, and UvrA must dissociate before UvrC can bind to UvrB–DNA and make the dual incisions. Thus, UvrA contributes to specificity both in selective loading of UvrB and in dissociation from the UvrB–DNA complex. Kinetic proofreading encompasses the unwinding of DNA by the A2B1 complex, hydrolysis of ATP by UvrB in the UvrB–DNA complex (Delagoutte et al., 2002), and differential affinities of UvrC to various UvrB–DNA complexes. No quantitative estimates are currently available for the contributions of the various mechanisms to specificity. However, it is likely that kinetic proofreading is the predominant determinant of specificity because the thermodynamic discrimination by UvrA (or A2B1) between undamaged and damaged DNA is almost nonexistent for such important lesions as cyclobutane pyrimidine dimers (Bertrand-Burggraf et al., 1991) that, in the absence of light or photolyase, can be repaired only by (A)BC excinuclease. As indicated above, kinetic proofreading is a powerful mechanism for achieving specificity, but by its very own design it has a built-in error rate such that nonsubstrates are also processed at significant rates. This is true for (A)BC excinuclease as well. The enzyme attacks undamaged DNA at about 10-4 the rate of cyclobutane pyrimidine dimer, excising damagefree 12–13-nt-long oligomers. This excision results in ‘‘gratuitous repair’’ (Branum et al., 2001), which may be stimulated by specific DNA structures or DNA dynamics such as transcription (Hanawalt, 2002), and that might be a source of spontaneous mutagenesis.
B. Excision Repair in Humans Excision repair in humans is carried out by six repair factors: RPA, XPA, XPC, TFIIH, XPG, and XPFERCC1 (Table II). RPA, XPA, and XPC recognize the damage; TFIIH unwinds the duplex around the damage;
NUCLEOTIDE EXCISION REPAIR
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and XPG and XPFERCC1 make the 30 and 50 incisions, respectively (Evans et al., 1997; Mu et al., 1995, 1996). Of the three proteins known to recognize DNA damage (RPA, XPA, and XPC), there has been serious debate as to which binds damage first and therefore should be called the ‘‘damage sensor.’’ Using either AAF–guanine adducts or (6–4) photoproducts as substrates for repair assays, it has been variously concluded that XPC or RPA and XPA are the sensors (Missura et al., 2001; Sugasawa et al., 1998; Wakasugi and Sancar, 1998, 1999). Although the causes of the discrepancies among the various groups remain to be investigated, it is important to note that all groups working on human excision repair agree that XPC does not recognize cyclobutane pyrimidine dimers (Batty et al., 2000; Reardon and Sancar, 2003; Sugasawa et al., 2001). In fact, there is a report indicating that XPC prefers undamaged DNA over cyclobutane pyrimidine dimer-containing DNA, and thus it appears XPC avoids this lesion (Hey et al., 2002). There seems to be a consensus that the most important substrate for the human excision nuclease is not recognized by XPC, and therefore, in the case of Pyr<>Pyr repair, XPC cannot be the initiator. However, XPA and RPA are equally inefficient in recognizing Pyr<>Pyr, and by this criterion, they cannot be the initiators either (Reardon and Sancar, 2003). The damaged DNA-binding protein (DDB) is a p127p48 heterodimer, and the small subunit of DDB is encoded by the XPE gene (Nichols et al., 1996). It has been reported that DDB binds to Pyr<>Pyr with modest affinity (Wakasugi et al., 2001; 2002), and hence it might be the ‘‘initiator’’ for repair of Pyr<>Pyr. However, this report is at odds with findings that CHO cell extracts, which do not contain DDB because of promoter silencing of the p48(DDB2) gene, are quite efficient at excising Pyr<>Pyr (Reardon et al., 1997a) and that supplementing the extracts with DDB has no effect on repair at low concentrations and inhibits repair at high concentrations (Reardon and Sancar, 2003). With this background, the following scheme has been proposed for the initial assembly at Pyr<>Pyr and at all other lesions: RPA, XPA, and XPC [usually in the form of XPCTFIIH (Drapkin et al., 1994)] have some preference for all DNA lesions, and any of the three may bind first. These proteins also have affinities to one another: XPA binds to both RPA and the XPCTFIIH complex (Li et al., 1995; Park et al., 1995). Thus, the three damage recognition components act cooperatively and assemble in random order at damage sites. Cooperative interaction increases the specificity somewhat, but the discrimination between damaged and undamaged DNA by each of the components and the affinities of the three factors to one another are of insufficient magnitude to confer a physiologically relevant specificity. A model consistent with all existing data, incorporating cooperativity, molecular matchmaking, and kinetic proofreading, has
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FIG. 4. Model for excision repair in man. The damage recognition factors, RPA, XPA, and XPCTFIIH, assemble at the damage site in a random order but in a cooperative manner to form an unstable ‘‘closed’’ complex. ATP hydrolysis by
NUCLEOTIDE EXCISION REPAIR
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been proposed for human excision nuclease (Reardon and Sancar, 2003, 2004; Wakasugi and Sancar, 1998) and is as follows (Fig. 4). The damage recognition factors, RPA, XPA, and XPCTFIIH, assemble at the damage site in a random order but in a cooperative manner and form an unstable ‘‘closed complex.’’ The moderate specificity achieved by cooperative binding is amplified by the kinetic proofreading activity of TFIIH, a six-subunit transcription/repair factor with both 30 ! 50 and 50 ! 30 helicase activity (Egly, 2001). Two subunits of TFIIH, XPB and XPD, hydrolyze ATP and unwind the duplex at the damage site to form a repair bubble of about 20 nt (Evans et al., 1997; Mu et al., 1997). This unwinding is accompanied by significant conformational changes in all components of the complex, leading to a new set of interactions that produces a rather stable complex called preincision complex 1 (PIC1). XPC is a molecular matchmaker that uses the ATP hydrolysis activity of TFIIH to promote entry of XPG into the complex as XPC leaves. The resulting complex is called PIC2. Finally, XPFERCC1 binds PIC2 to form PIC3 in which XPG makes the 30 incision first, followed by the 50 incision made by XPFERCC1. The first incision is made at the sixth 3 phosphodiester bond 30 to the damage and the second incision is made at the twentieth 5 phosphodiester bond 50 to the lesion to generate a damage-containing oligomer of 24–32 nt (Huang et al., 1992). The excised oligomer and most of the repair factors dissociate from the duplex (Mu et al., 1996, 1997), leaving RPA in the gap. Then repair synthesis proteins RFC/PCNA and Pol /" fill in the gap, and the repair patch is sealed by DNA ligase 1. As in the case with E. coli excision repair, the repair patch exactly matches the size of the excision gap (Reardon et al., 1997a). The human excision nuclease has an essentially infinite substrate range (Branum et al., 2001; Huang et al., 1994; Reardon et al., 1997b), and as in the case with E. coli, the excision nuclease employs cooperativity, molecular matchmaking, and kinetic proofreading to remove damage while minimizing the attack on undamaged DNA. Because of the larger genome size, human cells make more extensive use of both cooperativity and kinetic proofreading in DNA repair. Damage recognition involves TFIIH unwinds the duplex around the lesion, causing formation of a stable complex called preincision complex 1 (PIC1). XPC is a molecular matchmaker that helps to recruit/deliver XPG to PIC1, and XPC leaves before formation of PIC2, which comprises XPA, RPA, TFIIH, and XPG. Finally, this complex is recognized by XPFERCC1, leading to formation of PIC3 and the dual incision event, which releases damage in a 24–32-nt-long oligomer. ATP hydrolysis is required for PIC1–PIC3 formation, and the reaction may be aborted at any step along the pathway (kinetic proofreading) leading to dual incision. The repair gap is filled in by polymerase /", with the aid of RFC/PCNA, and sealed by DNA ligase. (See Color Insert.)
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cooperative interactions among RPA, XPA, and XPCTFIIH, and there are at least three proofreading steps at the formation of PIC1, PIC2, and PIC3, where ATP hydrolysis creates intermediates that cannot revert to the previous step but that may revert to the preassembly step at each of the stages if assembly occurred at a site with no DNA damage. At present, we do not have quantitative data on the relative contributions of cooperativity and kinetic proofreading to the specificity of excision repair in human cells. As in the case with the E. coli excinuclease, the human excision nuclease attacks and excises undamaged DNA at a significant rate (Branum et al., 2001; Reardon and Sancar, 2003). Clearly, the biological necessity of repairing all DNA lesions within the confines of the cellular limitation on the number of enzymes comes at a cost, which is a low frequency of mutations that inevitably occur when the gaps formed by excising undamaged DNA are filled in by DNA polymerases.
IV. TRANSCRIPTION-COUPLED REPAIR Excision repair is affected by other DNA transactions, including binding of regulatory proteins, compaction into chromatin, replication, recombination, and transcription. It has been found that transcription stimulates excision repair both in E. coli and in humans (Bohr et al., 1985; Mellon and Hanawalt, 1989; Mellon et al., 1987). Moreover, in the majority of cases, transcription stimulates the repair of only the transcribed strand (Mellon et al., 1987), and it may actually inhibit repair of the transcribed strand in the absence of an active mechanism coupling the two processes (Selby and Sancar, 1990). In the case of E. coli, the mechanism of transcriptioncoupled repair is reasonably well understood. In contrast, there is no in vitro system for eukaryotic transcription-coupled repair, and hence the mechanistic aspects of this process remain to be elucidated.
A. Transcription-Coupled Repair in E. coli Transcription-coupled repair is responsible for reduction of the mutation frequency (mutation frequency decline) in E. coli cells that are held in minimal medium after UV irradiation before plating on a rich selection medium (Li et al., 1999; Witkin, 1994). Transcription is coupled to excision repair through the intermediacy of the ‘‘transcription-repair coupling factor’’ (TRCF) encoded by the mfd gene (Selby et al., 1991). TRCF is a 130-kDa monomer, possesses helicase motifs, and functions as a translocase on RNA polymerase, causing its progression at temporary pause sites but releasing it when elongation is blocked by DNA damage or by a tightly
NUCLEOTIDE EXCISION REPAIR
57
bound protein (Park et al., 2002; Selby and Sancar, 1995; Washburn et al., 2003). Transcription-coupled repair in E. coli proceeds as follows (Selby and Sancar, 1993) (Fig. 5): E. coli RNA polymerase is unaffected by a DNA lesion such as Pyr<>Pyr in the nontranscribed strand, but when the damage is in the transcribed strand, elongation is blocked. The ternary complex that forms at a damage site is very stable with a half-life of greater than 20 hours and inhibits excision repair by interfering with the binding of the A2B1 complex. The TRCF recognizes both the stalled RNA polymerase and UvrA in the A2B1 complex. It releases RNA polymerase and the truncated transcript while simultaneously recruiting the A2B1 complex to the damage site. After the delivery of A2B1 to the lesion, TRCF dissociates, enabling UvrA to load UvrB onto the lesion, followed by binding of UvrC and excision of the damage. Because damage recognition is presumed to be the rate-limiting step in excision repair, and because a stalled RNA polymerase is a high-affinity target for TRCF, the overall effect of the process is an increase in the rate of repair of the transcribed strand relative to the coding (nontranscribed) strand and nontranscribed DNA. Thus, in transcription-coupled repair, RNA polymerase plays the role of a damage recognition subunit of the excision nuclease. This phenomenon has been called ‘‘recognition by proxy’’ (Sancar et al., 2004). In mfd mutants, repair of the transcribed strand is inhibited by the stalled RNA polymerase and, as a consequence, the coding strand and nontranscribed DNA are repaired more efficiently than the template strand of transcribed genes.
B. Transcription-Coupled Repair in Human Cells Transcription-coupled repair occurring at genes transcribed by RNA polymerase II (RNAPII) requires both excision repair factors and the CSA and CSB proteins (see Hanawalt, 2002; Venema et al., 1990a). Interestingly, XPC is not needed for this process (Venema et al., 1990b). The CSA protein belongs to the WD40 family of proteins (Henning et al., 1995), but how CSA may function in coupling transcription to repair is not clear. In contrast, the CSB protein, like the E. coli TRCF, possesses helicase motifs (Troelstra et al., 1992) and has some properties analogous to E. coli TRCF: it is an ATPase but not a helicase, and it has a translocase activity that enables RNAPII to progress through natural transcription pause sites (Selby and Sancar, 1997a,b). However, in contrast to the E. coli TRCF, the CSB protein does not disrupt the ternary complex of stalled RNA polymerase (Selby and Sancar, 1997b). At present, there is no in vitro system in which transcription by RNAPII stimulates excision repair.
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FIG. 5. Model for transcription-coupled repair in E. coli. A lesion in the nontranscribed (NT) strand has no effect on RNA polymerase (RNAP) (left side), but a lesion in the transcribed strand blocks progression of RNAP (right). Transcription-repair coupling factor (TRCF) recognizes and binds to the stalled
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However, three observations are relevant to how transcription may stimulate repair. First, a bubble structure 30 to a Pyr<>Pyr, such as might occur during transcription, is a substrate for human excision nuclease without XPC (Mu and Sancar, 1997). Second, surprisingly, RNAPII stalled at a Pyr<>Pyr does not inhibit excision of the lesion by the reconstituted human excision nuclease (Selby et al., 1997). Third, the human transcription termination factor 2 (TTF2) releases RNAPI and RNAPII stalled at a lesion and, in this regard, functions like the E. coli TRCF (Hara et al., 1999). Taking these facts into account, the following are two plausible models for transcription-coupled repair in humans (Fig. 6). The DNA in human cells is packed into chromatin, a structural feature that inhibits excision repair (see following). Transcribed genes are characterized by an open chromatin conformation, and stalling of RNAPII at a lesion may preserve this open structure, thus targeting the transcription-blocking lesion for rapid repair and simultaneously avoiding the inhibitory effect of chromatin compaction. Although CSB does not disrupt the ternary complex, it does interact with essential repair factors (Iyer et al., 1996; Selby and Sancar, 1997a), and thus an active role in transcription-coupled repair cannot be ruled out. It is possible that TTF2, which releases RNAPI and RNAPII stalled at a lesion (Hara et al., 1999), is the human TRCF but there is no evidence that TTF2 interacts with and recruits repair factors to sites of DNA damage. Clearly, more research is required to distinguish among these possibilities and determine the molecular mechanism of transcription-coupled repair in human cells, including the biochemical functions of CSA, CSB and TTF2 in the process.
V. REPAIR OF CHROMATIN Eukaryotic chromosomes are packaged into chromatin, a compact structure made up, at the first level of compaction, of DNA tightly wrapped around a histone octamer (nucleosome) that is joined to neighboring nucleosomes through linker DNA associated with a linker histone (Kornberg and Lorch, 1999; Wolffe, 1997). This structural organization has a significant influence on the distribution of UV-induced damage within chromatin (nucleosome vs. linker) and within the nucleosome core
polymerase and also binds to UvrA in the A2B1 complex. In an ATP-dependent reaction, TRCF displaces RNAP and the truncated transcript and, as RNAP leaves, UvrA2B1 replaces it at the damage site. UvrA delivers UvrB, UvrC binds to the UvrB–DNA complex, and excision proceeds as illustrated in Fig. 3. (See Color Insert.)
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FIG. 6. Model for transcription-coupled repair in human cells. Transcribing RNA polymerase (RNAP) is blocked by DNA lesions. Two scenarios are plausible and consistent with the available data. To the left is a pathway in which repair factors (XPC is not required) assemble at the lesion site and excise the damage, unaffected by the presence of the stalled polymerase and its transcript. After repair, RNA polymerase continues to translocate and transcribe. The pathway illustrated on the right involves transcription termination factor 2 (TTF2), which in an ATP-dependent reaction, displaces the stalled polymerase; TTF2 leaves the DNA, having performed its function. Repair is initiated and proceeds as illustrated in Fig. 4. Mechanistic details for the involvement of CSA and CSB are not known, and thus, they are shown ambiguously between the two pathways. (See Color Insert.)
(Gale and Smerdon, 1990; Gale et al., 1987; Mitchell et al., 1990; see Smerdon, 1991). In addition to this effect on adduct distribution, packaging of DNA into nucleosomes represses various DNA transactions by interfering with the accessibility of DNA-processing enzymes, including repair factors (Meijer and Smerdon, 1999; Moggs and Almouzni, 1999; Thoma, 1999). Indeed, packing DNA into minichromosomes results in less efficient repair than that observed in naked DNA, presumably because of reduced accessibility of the repair factors (Sugasawa et al., 1993; Wang et al., 1991).
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These studies used randomly damaged DNA and could not distinguish between inhibition of repair in the nucleosome core or linker DNA. More recent studies have used the in vitro assembly of nucleosomes containing site-specific DNA lesions within the core particle and either purified repair factors or mammalian cell extracts to examine in more detail the effect of chromatin structure on excision repair. It was determined that UVinduced photolesions as well as AAF- and cisplatin-modified bases located in nucleosome cores are repaired at a 5- to 10-fold reduced rate relative to the same lesions within the same sequence context in naked DNA (Hara and Sancar, 2002, 2003; Hara et al., 2000; Wang et al., 2003). These results are consistent with an effect of chromatin on protein accessibility, a problem that has been extensively studied with respect to transcription (Workman and Kingston, 1998). There are two major classes of chromatin-modifying factors that increase the accessibility of transcription factors to DNA in chromatin and thus, by analogy, may enhance the accessibility of repair factors to DNA damage within nucleosome cores (Aalfs and Kingston, 2000). The first class alters DNA-histone interactions through covalent modification of histones (Strahl and Allis, 2000). The second class encompasses several multisubunit ATP-dependent chromatin-remodeling complexes, including SWI/SNF2, ISWI, and Mi-like complexes. In one study, it was determined that ACF (ISWI-like) stimulated excision of (6–4) photoproducts in the linker region but had no effect on repair in the nucleosome core (Ura et al., 2001). In contrast, SWI/SNF (Kassabov et al., 2003; Yudkovsky et al., 1999) stimulated the repair of AAF-G adducts and (6–4) photoproducts, but not cyclobutane pyrimidine dimers, located in the nucleosome core (Hara and Sancar, 2002, 2003). It was found that the three damage recognition factors, RPA, XPA, and XPC, stimulate the remodeling activity of SWI/SNF, which in turn enhances excision of DNA lesions in the nucleosome core. The data indicate a plausible model for the role of SWI/SNF in excision repair (Hara and Sancar, 2002) (Fig. 7): repair factors locate the damage and facilitate recruitment of SWI/SNF, which remodels the nucleosome and facilitates the entry of XPG and XPFERCC1, leading to dual incisions and release of the damage-containing oligomer. Alternatively, remodeling by SWI/SNF may facilitate the assembly of repair factors at the damage site. More work is needed to distinguish between the two possibilities. Repair of chromatin is a relatively new and unexplored aspect of DNA repair in human cells, and future research will provide insight into the participation of various chromatin-modifying factors on repair in nucleosomes and in higher orders of DNA compaction.
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FIG. 7. Model for the role of chromatin remodeling by SWI/SNF in excision repair. Two pathways are presented: repair factors first or SWI/SNF first. To the left is a pathway in which damage recognition factors assemble at the damage site and then recruit SWI/ SNF to remodel the nucleosome. To the right is an alternative pathway in which remodeling by SWI/SNF accelerates the assembly of repair factors at the damage site. In both cases, dual incisions require the full complement of repair factors, as illustrated in Fig. 4, and after repair synthesis, the nucleosomes are reassembled. (See Color Insert.)
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VI. CONCLUSION Nucleotide excision repair is the major cellular pathway for removal of bulky lesions such as those introduced by UV irradiation or chemical carcinogens. Failure to remove DNA damage can result in increased mutagenesis, cancer, and cell death. Compared to the prokaryotic excision nuclease, the eukaryotic system is more complex, requiring 15 polypeptides for the basal reaction (recognition and removal of damage in naked DNA), a process that is accomplished by three proteins in bacteria. Although the human excision nuclease components show no homology to the prokaryotic proteins, the overall strategy is the same: damage recognition, localized helix unwinding, dual incisions to remove the lesion, and resynthesis/ligation to restore the DNA molecule. The mechanistic details of the dual-incision event are well characterized, especially in bacteria, so it was quite surprising when Goosen and colleagues reported the discovery of Cho, a previously unknown UvrC homolog that functions in E. coli excision repair (see Van Houten et al., 2002). Are there other repair protein homologs, particularly in the more complex human cell, and what role might they have in excision repair? Both prokaryotic and eukaryotic excision nucleases recognize and repair a wide spectrum of lesions, albeit with different efficiencies. Precisely how a DNA-binding protein distinguishes between normal and abnormal bases (base pairs) is not known. Thermodynamic destabilization by damage is a commonly proposed mechanism (see Geacintov et al., 2002), but such a mechanism disregards helix-stabilizing lesions such as those introduced by psoralen that are repaired efficiently (see Isaacs and Spielmann, 2004). Continued investigations are necessary to characterize the structural features that permit damage recognition factors to discriminate damaged from nondamaged DNA. Aside from this very basic question of what structural features make an adducted base abnormal, the more general question of damage recognition in human cells remains unresolved. Although there is a single damage recognition factor in E. coli (the A2B1 heterotrimer), human cells have three essential repair factors that show some affinity for various types of DNA damage: RPA, XPA, and XPC. In contrast to previous models that assigned a specific protein the role of ‘‘initiator,’’ we recently suggested that the three damage recognition factors assemble randomly at sites of DNA damage (Reardon and Sancar, 2003). Furthermore, we proposed a model in which this random assembly is accompanied by cooperative DNA binding, molecular matchmaking, and kinetic proofreading to achieve the requisite specificity in damage recognition and repair. This is a model to be tested, and further research is needed
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to ascertain the relative contributions of cooperativity and kinetic proofreading to the specificity of excision repair in human, as well as bacterial, cells. Excision repair is modulated by transcription in both eukaryotic and prokaryotic cells. The phenomenon of transcription-coupled repair (TCR) was first identified in mammalian cells, but in contrast to the well-established mechanistic details of TCR in E. coli, we have only a rudimentary understanding of the analogous system in human cells. Elucidation of the mechanistic aspects of this process first requires the development of an in vitro system in which both transcription and excision repair are accomplished at the efficient levels necessary for detailed biochemical studies. The problem of repair in chromatin is unique to eukaryotic cells and is likely relevant to the issue of transcription-coupled repair in human cells. Although recent work has provided insight into how lesions located in inaccessible regions of chromatin are repaired, continued research will reveal new details of this intriguing aspect of DNA repair. Eukaryotic cells have a complex system of checkpoints that delay or arrest cell cycle progression in response to DNA damage (see Sancar et al., 2004). How DNA repair is integrated into this response is an area for future study. With the exception of the XP-E complementation group, the gene products of all XP genes have well-defined roles in either excision repair or translesion synthesis (XP-V). The XPE gene encodes the small subunit of DDB, an abundant damaged DNA-binding protein. Much is known about DDB (see Tang and Chu, 2002), but its role in the cellular response to DNA damage has not been resolved and remains an active area of research. Among human proteins, DDB has the best discriminatory power between damaged and undamaged DNA, but it does not have a major role in excision repair, as evidenced by near-normal levels of excision and repair synthesis in vivo and by the in vitro reconstitution of excision repair without DDB (see Reardon and Sancar, 2003). How, then, is DDB involved in the cellular response to DNA damage? It has been suggested that DDB functions in the repair of lesions in chromatin as part of a multiprotein complex that performs chromatin remodeling in vivo (see Wittschieben and Wood, 2003). Although DDB does not stimulate repair of nucleosomal DNA in vitro (Hara et al., 2000), it possibly functions in this capacity in vivo, where there are higher orders of DNA compaction. DDB seems to be involved in other cellular processes, including transcription and the replication checkpoint, and it may function as a tumor suppressor by regulation of the p53 protein that controls cell-cycle progression and apoptosis following DNA damage (Itoh et al., 2003; see Hanawalt, 2002). Clearly, much work is needed to determine the in vivo functions of DDB. Such
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studies will also provide additional insight into the mechanisms of DNA repair in human cells and how this very important enzymatic pathway contributes to the avoidance of cancer.
REFERENCES Aalfs, J. D., and Kingston, R. E. (2000). What does ‘‘chromatin remodeling’’ mean? Trends Biochem. Sci. 25, 548–555. Aravind, L., Walker, D. R., and Koonin, E. V. (1999). Conserved domains in DNA repair proteins and evolution of repair systems. Nucleic Acids Res. 27, 1223–1242. Batty, D., Rapic-Otrin, V., Levine, A. S., and Wood, R. D. (2000). Stable binding of human XPC complex to irradiated DNA confers strong discrimination for damaged sites. J. Mol. Biol. 300, 275–290. Bertrand-Burggraf, E., Selby, C. P., Hearst, J. E., and Sancar, A. (1991). Identification of the different intermediates in the interaction of (A)BC excinuclease with its substrates by Dnase I footprinting on two uniquely modified oligonucleotides. J. Mol. Biol. 219, 27–36. Bessho, T., Mu, D., and Sancar, A. (1997). Initiation of DNA interstrand cross-link repair in humans: The nucleotide excision repair system makes dual incisions 50 to the cross-linked base and removes a 22- to 28-nucleotide long damage free strand. Mol. Cell. Biol. 17, 6822–6830. Bohr, V. A., Smith, C. A., Okumoto, D. S., and Hanawalt, P. C. (1985). DNA repair in an active gene: Removal of pyrimidine dimers from the DHFR gene of CHO cells is much more efficient than in the genome overall. Cell 40, 359–369. Branum, M. E., Reardon, J. T., and Sancar, A. (2001). DNA repair excision nuclease attacks undamaged DNA. A potential source of spontaneous mutations. J. Biol. Chem. 276, 25421–25426. Burgess, S. M., and Guthrie, C. (1993). Beat the clock: paradigms for NTPases in the maintenance of biological fidelity. Trends Biochem. Sci. 18, 381–384. Cleaver, J. E. (1968). Defective repair replication of DNA in xeroderma pigmentosum. Nature 218, 652–656. Cleaver, J. E., and Kraemer, K. H. (1989). Xeroderma pigmentosum. In ‘‘The Metabolic Basis of Inherited Disease’’ (C. R. Scriver, A. L. Beaudet, W. S. Sly, and D. Valle, Eds.). Vol. 2, pp. 2949–2971. McGraw-Hill, New York. Delagoutte, E., Bertrand-Burggraf, E., Dunand, J., and Fuchs, R. P. P. (1997). Sequence dependent modulation of nucleotide excision repair: the efficiency of the incision reaction is inversely correlated with the stability of the preincision UvrB DNA complex. J. Mol. Biol. 266, 703–710. Delagoutte, E., Fuchs, R. P. P., and Bertrand-Burggraf, E. (2002). The isomerization of the UvrB DNA preincision complex couples the UvrB and UvrC activities. J. Mol. Biol. 320, 73–84. Drapkin, R., Reardon, J. T., Ansari, A., Huang, J.-C., Zawel, L., Ahn, K., Sancar, A., and Reinberg, D. (1994). Dual role of TFIIH in DNA excision repair and in transcription by RNA polymerase II. Nature 368, 769–772. Egly, J.-M. (2001). TFIIH: From transcription to clinic. FEBS Lett. 24884, 124–128. Evans, E., Moggs, J. G., Hwang, J. R., Egly, J.-M., and Wood, R. D. (1997). Mechanism of open complex and dual incision formation by human nucleotide excision repair factors. EMBO J. 16, 6559–6573.
66
SANCAR AND REARDON
Gale, J. M., and Smerdon, M. J. (1990). UV-induced (6–4) photoproducts are distributed differently than cyclobutane dimers in nucleosomes. Photochem. Photobiol. 51, 411–417. Gale, J. M., Nissen, K. A., and Smerdon, M. J. (1987). UV-induced formation of pyrimidine dimers in nucleosome core DNA is strongly modulated with a period of 10.3 bases. Proc. Natl. Acad. Sci. USA 84, 6644–6648. Geacintov, N. E., Broyde, S., Buterin, T., Naegeli, H., Wu, M., Yan, S., and Patel, D. J. (2002). Thermodynamic and structural factors in the removal of bulky DNA adducts by the nucleotide excision repair machinery. Biopolymers 65, 202–210. Giedroc, D. P., Keating, K. M., Williams, K. R., and Coleman, J. E. (1987). The function of zinc in gene 32 protein from T4. Biochemistry 26, 5251–5259. Goosen, N., Moolenaar, G. F., Visse, R., and van de Putte, P. (1998). Functional domains of the E. coli UvrABC proteins in nucleotide excision repair. In ‘‘Nucleic Acids and Molecular Biology: DNA Repair’’ (F. Eckstein and D. M. J. Lilley, Eds.), pp. 103–123. Springer, Berlin. Hanawalt, P. C. (2002). Subpathways of nucleotide excision repair and their regulation. Oncogene 21, 8949–8956. Hara, R., and Sancar, A. (2002). The SWI/SNF chromatin-remodeling factor stimulates repair by human excision nuclease in the mononucleosome core particle. Mol. Cell. Biol. 22, 6779–6787. Hara, R., and Sancar, A. (2003). Effect of damage type on stimulation of human excision nuclease by SWI/SNF chromatin remodeling factor. Mol. Cell. Biol. 23, 4121–4125. Hara, R., Selby, C. P., Liu, M., Price, D. H., and Sancar, A. (1999). Human transcription release factor 2 dissociates RNA polymerases I and II stalled at a cyclobutane thymine dimer. J. Biol. Chem. 274, 24779–24786. Hara, R., Mo, J., and Sancar, A. (2000). DNA damage in the nucleosome core is refractory to repair by human excision nuclease. Mol. Cell. Biol. 20, 9173–9181. Henning, K. A., Li, L., Iyer, N., McDaniel, L. D., Reagan, M. S., Legerski, R., Schultz, R. A., Stefanini, M., Lehmann, A. R., Mayne, L. V., and Friedberg, E. C. (1995). The Cockayne syndrome group A gene encodes a WD repeat protein that interacts with CSB protein and a subunit of RNA polymerase II TFIIH. Cell 82, 555–564. Hey, T., Lipps, G., Sugasawa, K., Iwai, S., Hanaoka, F., and Krauss, G. (2002). The XPCHR23B complex displays high affinity and specificity for damaged DNA in a true equilibrium fluorescence assay. Biochemistry 41, 6583–6587. Hopfield, J. J. (1974). Kinetic proofreading. A new mechanism for reducing errors in biosynthetic processes requiring high specificity. Proc. Natl. Acad. Sci. USA 71, 4135–4139. Hsu, D. S., Kim, S.-T., Sun, Q., and Sancar, A. (1995). Structure and function of the UvrB protein. J. Biol. Chem. 270, 8319–8327. Huang, J.-C., Svoboda, D. L., Reardon, J. T., and Sancar, A. (1992). Human nucleotide excision nuclease removes thymine dimers from DNA by incising the 22nd phosphodiester bond 50 and the 6th phosphodiester bond 30 to the photodimer. Proc. Natl. Acad. Sci. USA 89, 3664–3668. Huang, J.-C., Hsu, D. S., Kazantsev, A., and Sancar, A. (1994). Substrate spectrum of human excinuclease: Repair of abasic sites, methylated bases, mismatches, and bulky adducts. Proc. Natl. Acad. Sci. USA 91, 12213–12217. Isaacs, R. J., and Spielmann, H. P. (2004). A model for initial DNA lesion recognition by NER and MMR based on local conformational flexibility. DNA Repair. 3, 455–464.
NUCLEOTIDE EXCISION REPAIR
67
Itoh, T., O’Shea, C., and Linn, S. (2003). Impaired regulation of tumor suppressor p53 caused by mutations in the xeroderma pigmentosum DDB2 gene: mutual regulatory interactions between p48DDB2 and p53. Mol. Cell. Biol. 23, 7540–7553. Iyer, N., Reagan, M. S., Wu, K.- J., Canagarajah, B., and Friedberg, E. C. (1996). Interactions involving the human RNA polymerase II transcription/nucleotide excision repair complex TFIIH, the nucleotide excision repair protein XPG, and Cockayne syndrome group B (CSB) protein. Biochemistry 35, 2157–2167. Kassabov, S. R., Zhang, B., Persinger, J., and Bartholomew, B. (2003). SWI/SNF unwraps, slides, and rewraps the nucleosome. Molec. Cell 11, 391–403. Kelly, R. C., Jensen, D. E., and von Hippel, P. H. (1976). DNA ‘‘melting’’ proteins. IV. Fluorescence measurements of binding parameters for bacteriophage T4 gene 32protein to mono, oligo, and polynucleotides J. Biol. Chem. 251, 7240–7250. Kornberg, R. D., and Lorch, Y. (1999). Twenty-five years of the nucleosome, fundamental particle of the eukaryotic chromosome. Cell 98, 285–294. Kowalski, J. C., Belfort, M., Stapleton, M. A., Holpert, M., Dansereau, J. T., Pietrokovski, S., Baxter, S. M., and Derbyshire, V. (1999). Configuration of the catalytic GIY-YIG domain of intron endonuclease I-TevI: coincidence of computational and molecular findings. Nucleic Acids Res. 27, 2115–2125. Levine, M., and Tjian, R. (2003). Transcription regulation and animal diversity. Nature 424, 147–151. Li, L., Lu, X., Peterson, C. A., and Legerski, R. J. (1995). An interaction between the DNA repair factor XPA and replication protein A appears essential for nucleotide excision repair. Mol. Cell. Biol. 15, 5396–5402. Li, B.-H., Ebbert, A., and Bockrath, R. (1999). Transcription-modulated repair in Escherichia coli evident with UV-induced mutation spectra in supF. J. Mol. Biol. 294, 35–48. Lin, J.-J., and Sancar, A. (1991). The C-terminal half of UvrC protein is sufficient to reconstitute (A)BC excinuclease. Proc. Natl. Acad. Sci. USA 88, 6824–6828. Lin, J.-J., and Sancar, A. (1992). Active site of (A)BC excinuclease. I. Evidence for 50 incision by UvrC through a catalytic site involving Asp399, Asp438, Asp466, and His538 residues. J. Biol. Chem. 267, 17688–17692. Lin, J.-J., Phillips, A. M., Hearst, J. E., and Sancar, A. (1992). Active site of (A)BC excinuclease. II. Binding, bending, and catalysis mutants of UvrB reveal a direct role in 30 and an indirect role in 50 incision. J. Biol. Chem. 267, 17693–17700. MacGlashan, D. (2001). Signaling cascades: escape from kinetic proofreading. Proc. Natl. Acad. Sci. USA 98, 6989–6990. Meijer, M., and Smerdon, M. J. (1999). Accessing DNA damage in chromatin: Insights from transcription. BioEssays 21, 596–603. Mellon, I., and Hanawalt, P. C. (1989). Induction of the Escherichia coli lactose operon selectively increases repair of its transcribed DNA strand. Nature 342, 95–98. Mellon, I., Spivak, G., and Hanawalt, P. C. (1987). Selective removal of transcriptionblocking DNA damage from the transcribed strand of the mammalian DHFR gene. Cell 51, 241–249. Missura, M., Buterin, T., Hindges, R., Hu¨bscher, U., Kaspa´rkova´, J., Brabec, V., and Naegeli, H. (2001). Double check probing of DNA bending and unwinding by XPA-RPA: An architectural function in DNA repair. EMBO J. 20, 3554–3564. Mitchell, D. L., Nguyen, T. D., and Cleaver, J. E. (1990). Nonrandom induction of pyrimidine-pyrimidone (6–4) photoproducts in ultraviolet-irradiated human chromatin. J. Biol. Chem. 265, 5353–5356.
68
SANCAR AND REARDON
Moggs, J. G., and Almouzni, G. (1999). Chromatin rearrangements during nucleotide excision repair. Biochimie 81, 45–52. Moolenaar, G. F., van Rossum-Fikkert, S., van Kesteren, M., and Goosen, N. (2002). Cho, a second endonuclease involved in Escherichia coli nucleotide excision repair. Proc. Natl. Acad. Sci. USA 99, 1467–1472. Mu, D., and Sancar, A. (1997). Model for XPC-independent transcription coupled repair of pyrimidine dimers in humans. J. Biol. Chem. 272, 7570–7573. Mu, D., Park, C.-H., Matsunaga, T., Hsu, D. S., Reardon, J. T., and Sancar, A. (1995). Reconstitution of human DNA repair excision nuclease in a highly defined system. J. Biol. Chem. 270, 2415–2418. Mu, D., Hsu, D. S., and Sancar, A. (1996). Reaction mechanism of human DNA repair excision nuclease. J. Biol. Chem. 271, 8285–8294. Mu, D., Wakasugi, M., Hsu, D. S., and Sancar, A. (1997). Characterization of reaction intermediates of human excision repair nuclease. J. Biol. Chem. 272, 28971–28979. Nichols, A. F., Ong, P., and Linn, S. (1996). Mutations specific to the xeroderma pigmentosum group E Ddb phenotype. J. Biol. Chem. 271, 24317–24320. Ninio, J. (1975). Kinetic amplification of enzyme discrimination. Biochimie 57, 587–595. O’Brien, P. J., and Ellenberger, T. (2004). Dissecting the broad substrate specificity of human 3-methyladenine-DNA glycosylase. J. Biol. Chem. 279, 9750–9757. Orren, D. K., and Sancar, A. (1989). The (A)BC excinuclease of Escherichia coli has only the UvrB and UvrC subunits in the incision complex. Proc. Natl. Acad. Sci. USA 86, 5237–5241. Orren, D. K., and Sancar, A. (1990). Formation and enzymatic properties of the UvrBDNA complex. J. Biol. Chem. 265, 15796–15803. Orren, D. K., Selby, C. P., Hearst, J. E., and Sancar, A. (1992). Post-incision steps of nucleotide excision repair in Escherichia coli. Disassembly of the UvrBC-DNA complex by helicase II and DNA polymerase I. J. Biol. Chem. 267, 780–788. Park, C.-H., Mu, D., Reardon, J. T., and Sancar, A. (1995). The general transcriptionrepair factor TFIIH is recruited to the excision repair complex by the XPA protein independent of the TFIIE transcription factor. J. Biol. Chem. 270, 4896–4902. Park, J.-S., Marr, M. T., and Roberts, J. W. (2002). E. coli transcription repair coupling factor (Mfd protein) rescues arrested complexes by promoting forward translocation. Cell 109, 757–767. Petit, C., and Sancar, A. (1999). Nucleotide excision repair: From E. coli to man. Biochimie 81, 15–25. Prakash, S., and Prakash, L. (2000). Nucleotide excision repair in yeast. Mutation Res. 451, 13–24. Reardon, J. T., and Sancar, A. (2003). Recognition and repair of the cyclobutane thymine dimer, a major cause of skin cancers, by the human excision nuclease. Genes Dev. 17, 2539–2551. Reardon, J. T., and Sancar, A. (2004). Thermodynamic cooperativity and kinetic proofreading in DNA damage recognition and repair. Cell Cycle. 3, 141–144. Reardon, J. T., Thompson, L. H., and Sancar, A. (1997a). Rodent UV sensitive mutant cell lines in complementation groups 6–10 have normal general excision repair activity. Nucleic Acids Res. 25, 1015–1021. Reardon, J. T., Bessho, T., Kung, H. C., Bolton, P. H., and Sancar, A. (1997b). In vitro repair of oxidative DNA damage by human nucleotide excision repair system: Possible explanation for neurodegeneration in xeroderma pigmentosum patients. Proc. Natl. Acad. Sci. USA 94, 9463–9468. Sancar, A. (1996). DNA excision repair. Annu. Rev. Biochem. 65, 43–81.
NUCLEOTIDE EXCISION REPAIR
69
Sancar, A. (2003). Structure and function of DNA photolyase and cryptochrome bluelight photoreceptors. Chem. Rev. 103, 2203–2238. Sancar, A., and Hearst, J. E. (1993). Molecular matchmakers. Science 259, 1415–1420. Sancar, A., and Rupp, D. (1983). A novel repair enzyme: UVRABC excision nuclease of Escherichia coli cuts a DNA strand on both sides of the damaged region. Cell 33, 249–260. Sancar, A., Franklin, K. A., and Sancar, G. B. (1984). Escherichia coli DNA photolyase stimulates uvrABC excision nuclease in vitro. Proc. Natl. Acad. Sci. USA 81, 7397–7401. ¨ nsal-Kac¸maz, K., and Linn, S. (2004). Molecular Sancar, A., Lindsey-Boltz, L. A., U mechanisms of mammalian DNA repair and the DNA damage checkpoints. Annu. Rev. Biochem. 73, 39–85. Selby, C. P., and Sancar, A. (1990). Transcription preferentially inhibits nucleotide excision repair of the template DNA strand in vitro. J. Biol. Chem. 265, 21330–21336. Selby, C. P., and Sancar, A. (1991). Gene- and strand-specific repair in vitro: Partial purification of a transcription-repair coupling factor. Proc. Natl. Acad. Sci. USA 88, 8232–8236. Selby, C. P., and Sancar, A. (1993). Molecular mechanism of transcription-repair coupling. Science 260, 53–58. Selby, C. P., and Sancar, A. (1995). Structure and function of transcription-repair coupling factor.II. catalytic properties. J. Biol. Chem. 270, 4890–4895. Selby, C. P., and Sancar, A. (1997a). Human transcription-repair coupling factor CSB/ ERCC6 is a DNA-stimulated ATPase but is not a helicase and does not disrupt the ternary transcription complex of stalled RNA polymerase II. J. Biol. Chem. 272, 1885–1890. Selby, C. P., and Sancar, A. (1997b). Cockayne syndrome group B protein enhances elongation by RNA polymerase II. Proc. Natl. Acad. Sci. USA 94, 11205–11209. Selby, C. P., Witkin, E. M., and Sancar, A. (1991). Escherichia coli mfd mutant deficient in ‘‘mutation frequency decline’’ lacks strand-specific repair: In vitro complementation with purified coupling factor. Proc. Natl. Acad. Sci. USA 88, 11574–11578. Selby, C. P., Drapkin, R., Reinberg, D., and Sancar, A. (1997). RNA polymerase II stalled at a thymine dimer: Footprint and effect on excision repair. Nucleic Acids Res. 25, 787–793. Shi, Q., Thresher, R., Sancar, A., and Griffith, J. (1992). Electron microscopic study of (A)BC excinuclease. DNA is sharply bent in the UvrB-DNA complex. J. Mol. Biol. 226, 425–432. Sibghat-Ullah, Sancar, A., and Hearst, J. E. (1990). The repair patch of E. coli (A)BC excinuclease. Nucleic Acids Res. 18, 5051–5053. Smerdon, M. J. (1991). DNA repair and the role of chromatin structure. Curr. Opin. Cell Biol. 3, 422–428. Strahl, B. D., and Allis, D. (2000). The language of covalent histone modifications. Nature 403, 41–45. Sugasawa, K., Masutani, C., and Hanaoka, F. (1993). Cell-free repair of UV-damaged simian virus 40 chromosomes in human cell extracts. I. Development of a cell-free system detecting excision repair of UV-irradiated SV40 chromosomes. J. Biol. Chem. 268, 9098–9104. Sugasawa, K., Ng, J. M. Y., Masutani, C., Iwai, S., van der Spek, P. J., Eker, A. P. M., Hanaoka, F., Bootsma, D., and Hoeijmakers, J. H. J. (1998). Xeroderma
70
SANCAR AND REARDON
pigmentosum group C protein complex is the initiator of global genome nucleotide excision repair. Mol. Cell 2, 223–232. Sugasawa, K., Okamoto, T., Shimizu, Y., Masutani, C., Iwai, S., and Hanaoka, F. (2001). A multistep damage recognition mechanism for global genomic nucleotide excision repair. Genes Dev. 15, 507–521. Tang, J., and Chu, G. (2002). Xeroderma pigmentosum complementation group E and UV-damaged DNA-binding protein. DNA Repair. 1, 601–616. Thoma, F. (1999). Light and dark in chromatin repair: Repair of UV-induced DNA lesions by photolyase and nucleotide excision repair. EMBO J. 18, 6585–6598. Troelstra, C., van Gool, A., de Wit, J., Vermeulen, W., Bootsma, D., and Hoeijmakers, J. H. J. (1992). ERCC6, a member of a subfamily of putative helicases, is involved in Cockayne’s syndrome and preferential repair of active genes. Cell 71, 939–953. Ura, K., Araki, M., Saeki, H., Masutani, C., Ito, T., Iwai, S., Mizukoshi, T., Kaneda, Y., and Hanaoka, F. (2001). ATP-dependent chromatin remodeling facilitates nucleotide excision repair of UV-induced DNA lesions in synthetic dinucleosomes. EMBO J. 20, 2004–2014. Van Houten, B., Gamper, H., Holbrook, S. R., Hearst, J. E., and Sancar, A. (1986). Action mechanism of ABC excision nuclease on a DNA substrate containing a psoralen crosslink at a defined position. Proc. Natl. Acad. Sci. USA 83, 8077–8081. Van Houten, B., Gamper, H., Sancar, A., and Hearst, J. E. (1987). DNase I footprint of ABC excinuclease. J. Biol. Chem. 262, 13180–13187. Van Houten, B., Eisen, J. A., and Hanawalt, P. C. (2002). A cut above: Discovery of an alternative excision repair pathway in bacteria. Proc. Natl. Acad. Sci. USA 99, 2581–2583. Venema, J., Mullenders, L. H. F., Natarajan, A. T., van Zeeland, A. A., and Mayne, L. V. (1990a). The genetic defect in Cockayne syndrome is associated with a defect in repair of UV-induced DNA damage in transcriptionally active DNA. Proc. Natl. Acad. Sci. USA 87, 4707–4711. Venema, J., van Hoffen, A., Natarajan, A. T., van Zeeland, A. A., and Mullenders, L. H. (1990b). The residual repair capacity of xeroderma pigmentosum complementation group C fibroblasts is highly specific for transcriptionally active DNA. Nucleic Acids Res. 18, 443–448. Verhoeven, E. E. A., van Kesteren, M., Moolenaar, G. F., Visse, R., and Goosen, N. (2000). Catalytic sites for 30 and 50 incision of Escherichia coli nucleotide excision repair are both located in UvrC. J. Biol. Chem. 275, 5120–5123. Wakasugi, M., and Sancar, A. (1998). Assembly, subunit composition, and footprint of human DNA repair excision nuclease. Proc. Natl Acad. Sci. USA 95, 6669–6674. Wakasugi, M., and Sancar, A. (1999). Order of assembly of human DNA repair excision nuclease. J. Biol. Chem. 274, 18759–18768. Wakasugi, M., Shimizu, M., Morioka, H., Linn, S., Nikaido, O., and Matsunaga, T. (2001). Damaged DNA binding protein DDB stimulates the excision of cyclobutane pyrimidine dimers in vitro in concert with XPA and replication protein A. J. Biol. Chem. 276, 15434–15440. Wakasugi, M., Kawashima, A., Morioka, H., Linn, S., Sancar, A., Mori, T., Nikaido, O., and Matsunaga, T. (2002). DDB accumulates at DNA damage sites immediately after UV irradiation and directly stimulates nucleotide excision repair. J. Biol. Chem. 277, 1637–1640. Wang, Z., Wu, X., and Friedberg, E. C. (1991). Nucleotide excision repair of DNA by human cell extracts is suppressed in reconstituted nucleosomes. J. Biol. Chem. 266, 22472–22478.
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Wang, D., Hara, R., Singh, G., Sancar, A., and Lippard, S. J. (2003). Nucleotide excision repair from site-specifically platinum-modified nucleosomes. Biochemistry 42, 6747–6753. Washburn, R. S., Wang, Y., and Gottesman, M. E. (2003). Role of E. coli transcriptionrepair coupling factor Mfd in Nun-mediated transcription termination. J. Mol. Biol. 329, 655–662. Witkin, E. M. (1994). Mutation frequency decline revisited. BioEssays 16, 437–444. Wittschieben, B. Ø., and Wood, R. D. (2003). DDB complexities. DNA Repair. 2, 1065–1069. Wolffe, A. P. (1997). ‘‘Chromatin: Structure and Function.’’ Academic Press, New York. Wood, R. D. (1997). Nucleotide excision repair in mammalian cells. J. Biol. Chem. 272, 23465–23468. Workman, J. L., and Kingston, R. E. (1998). Alteration of nucleosome structure as a mechanism of transcriptional regulation. Annu. Rev. Biochem. 67, 545–579. Yudkovsky, N., Logie, C., Hahn, S., and Peterson, C. (1999). Recruitment of the SWI/ SNF chromatin remodeling complex by transcriptional activators. Genes Dev. 13, 2369–2374. Zheng, H., Wang, X., Warren, A. J., Legerski, R. J., Nairn, R. S., Hamilton, J. W., and Li, L. (2003). Nucleotide excision repair and polymerase -mediated error prone removal of mitomycin C interstrand cross links. Mol. Cell. Biol. 23, 754–761. Zou, Y., and van Houten, B. (1999). Strand opening by the UvrA2B complex allows dynamic recognition of DNA damage. EMBO J. 18, 4889–4901.
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PHOTOLYASE AND CRYPTOCHROME BLUE-LIGHT PHOTORECEPTORS By AZIZ SANCAR Department of Biochemistry and Biophysics, University of North Carolina, Chapel Hill, North Carolina, 27599
I. II. III. IV.
Introduction . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . Phylogenetics . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . Structure of Photolyase. . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . Reaction Mechanism of Photolyase . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Binding of Photolyase to Substrate. . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. Catalysis by Photolyase. . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . V. (6–4) Photolyase. . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Binding of (6–4) Photolyase. . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. Catalysis by (6–4) Photolyase . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . VI. Cryptochrome . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Structure. . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. Function of Cryptochrome . . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . VII. Conclusion . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . References . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . .
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I. Introduction Photolyase repairs ultraviolet (UV)-induced DNA damage using nearUV/blue-light as energy source or cosubstrate (Fig. 1). The enzyme is a monomeric protein that contains two chromophore/cofactors (Sancar, 1994, 2003). One of the chromophores, which in the majority of photolyases is methenyltetrahydrofolate (MTHF) and in a limited number of species that can synthesize 5-deazaflavin is 8-hydroxy-7,8-didmethyl-5deazariboflavin (8-HDF) (Fig. 2), is located on the surface of the protein and functions as a photoantenna. The catalytic cofactor, located in the core of the globular enzyme, is the two-electron reduced and deprotonated flavin, FADH. Cryptochrome is defined as a photolyaselike molecule with no DNA repair activity (Cashmore, 2003; Sancar, 2000, 2003). Cryptochromes regulate growth and development in response to blue-light in plants and control the circadian clock in animals by lightdependent and light-independent mechanisms. The function of cryptochromes in bacteria is not known at present.
73 ADVANCES IN PROTEIN CHEMISTRY, Vol. 69
Copyright 2004, Elsevier Inc. All rights reserved. 0065-3233/04 $35.00
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Fig. 1. (A) Photoreactivation. Escherichia coli cells exposed to the indicated doses of 254-nm ultraviolet (UV) were either plated directly (closed circles) or exposed to a white-light flash of about 1 ms before plating (open circle). Following incubation at 37 C for 24 hours, colonies were counted and relative survival was calculated. (B) Molecular basis of UV killing and photoreactivation. Far UV (200–280 nm) induces two major photoproducts in DNA: the cyclobutane pyrimidine dimer (Pyr<>Pyr) and the pyrimidine–pyrimidone (6–4) photoproduct. Thymine is the most common pyrimidine in both types of photoproducts. These photoproducts are reversed to normal bases by photolyases (PL) with the aid of blue-light.
II. Phylogenetics At this time, the photolyase/cryptochrome family has three members: photolyase (cyclobutane pyrimidine dimer photolyase), (6–4) photolyase, and cryptochrome. A number of phylogenetic trees based on sequence comparisons of more than 100 members of the family across the three
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Fig. 2. Structures of photolyase/cryptochrome cofactors. All photolyases and cryptochromes contain FAD, and all either contain or are thought to contain 5,10-methenyltetrahydrofolate (MTHF) or 8-hydroxy-7,8-didemethyl-5-deazariboflavin (8-HDF) as the second chromophore.
biological kingdoms have been generated (Brudler et al., 2003; Cashmore et al., 1999), and a simple phylogenetic tree is shown in Figure 3. Some interesting points that emerge from these analyses will be briefly summarized. Photolyase has been found in many species from the prokaryotic, eukaryotic, and archaeal kingdoms, and even in some viruses (Sancar, 2003; Srinivasan et al., 2001; Willer et al., 1999) (Table I). However, it has also been found that many species from all three kingdoms lack the enzyme. Prokaryotic organisms that possess the enzyme include Escherichia coli and Bacillus firmus. Other prokaryotes, such as Haemophilus influenzae and Bacillus subtilis, do not have photolyase. Among eukaryotic organisms that are used as model systems, Saccharomyces cerevisiae contains photolyase, but Schizosaccharomyces pombe does not. Of multicellular organisms, Caenorhabditis elegans and garter snake lack photolyase, but rattlesnake, zebrafish, and goldfish contain the enzyme. Many vertebrates appear to have photolyase. Curiously, however, when mammals were separated into marsupials and placentals, the former retained photolyase, and the latter did not. Thus, opossum has photolyase but raccoons and humans do not.
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Fig. 3. Phylogentic relationship among select members of the photolyase/ cryptochrome family. Sequences were aligned with Clustal W, and the tree was produced by the Neighbor-Joining method using MEGA 2.1. Bootstrap confidence values are shown (values for interior branches 95% are statistically significant). At, Arabidopsis thaliana; Dm, Drosophila melangaster; Ec, Escherichia coli; Hs, Homo sapiens; Vc, Vibrio cholerae.
Table I Distribution of Photolyase and Cryptochromes in the Biological World Enzyme/Photoreceptor
Eubacteria B. subtilis E. coli V. cholerae Archaea M. janaschii M. thermoautotrophicum Eukarya S. pombe C. elegans S. cerevisiae D. melanogaster H. sapiens A. thaliana Viruses Shoppe (rabbit) fibroma virus Fowlpox virus
Photolyase
(6–4) Photolyase
Cryptochrome
No Yes Yes
No No No
No No Yes(2)
No Yes
No No
No No
No No Yes Yes No Yes
No No No Yes No Yes
No No No Yes Yes (2) Yes (3)
Yes Yes
No No
No No
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All plants tested so far, including Arabidopsis thaliana, tobacco, and soybean, do have photolyase. The (6–4) photolyase was first discovered in D. melanogaster (Todo et al., 1993, 1996). Subsequently it was found in Xenopus laevis, rattlesnake (Kim et al., 1994), zebrafish, and A. thaliana, among many other species (Todo, 1999). Of special interest, the enzyme has not been found in birds and mammals (Hsu et al., 1996). Thus, humans lack both photolyase and (6–4) photolyase and cannot carry out photorepair of UV-induced DNA damage. They rely solely on nucleotide excision repair for eliminating these potentially mutagenic and carcinogenic lesions from their DNA. Cryptochrome was first discovered in plants (Ahmad and Cashmore, 1993; Malhotra et al., 1995), and subsequently in humans (Hsu et al., 1996). It has since been found in organisms ranging from Vibrio cholerae (Worthington et al., 2003) to Drosophila (Sancar, 2000; Todo, 1999) to zebrafish, which contains six cryptochrome genes (Brudler et al., 2003; Todo, 1999). It has been found in all insect and bird species tested. Of organisms that are used as model systems, Drosophila has one cryptochrome, humans and mice have two, and Arabidopsis has three cryptochromes. Most bacteria including E. coli lack cryptochromes, Synechocystis possess one (Brudler et al., 2003; Ng and Pakrasi, 2001), and V. cholerae has two cryptochromes. Caenorhabditis elegans, which lacks photolyase and (6–4) photolyase, also does not have a cryptochrome.
III. Structure of Photolyase Photolyases are monomeric proteins of 450–550 amino acids and two noncovalently bound cofactors (Johnson et al., 1988; Jorns et al., 1984). One of the cofactors is always FAD. The other cofactor, which is also called the second chromophore, is methenyltetrahydrofolate (MTHF) in the majority of photolyases and 8-hydroxy-7,8-didemethyl-5-deazariboflavin (8-HDF) in a limited number of species (some archaea, Anacystis nidulans, and some ferns that synthesize this cofactor) (Eker et al., 1988, 1990; Johnson et al., 1988; Kiener et al., 1989; Malhotra et al., 1992). Crystal structures of photolyases from E. coli (Park et al., 1995), A. nidulans (Tamada et al., 1997), and Thermus thermophilus (Komori et al., 2001) have been solved. Although they possess different second chromophores (E. coli photolyase has MTHF and the latter two contain 8-HDF), and the level of overall sequence homology among the three enzymes is only about 25% sequence identity, the structures of all three are remarkably similar. Here, the structure of E. coli photolyase will be presented and the minor differences of the other two photolyases from this structure will be briefly mentioned.
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E. coli photolyase is made up of two well-defined domains (Fig. 4): an Nterminal / domain (residues 1–132) and a C-terminal -helical domain (residues 204–471). The two domains are connected to one another with a long loop (residues 132–203) that wraps around the / domain (Fig. 4). The MTHF photoantenna is located in a shallow cleft between the two domains and is partially exposed to solvent. In contrast, the FAD cofactor is deeply buried within the -helical domain and has the unusual Ushaped (or cis) conformation, in which the flavin and the adenine rings are stacked on top of one another. The flavin is accessible to the flat surface of the -helical domain through a hole in the middle of this domain. The hole has the right dimensions and polarity to allow the entry of a thymine dimer to within van der Waals contact distance to the isoalloxazine ring of FAD. Surface potential representation of the enzyme reveals a positively charged groove running the length of the molecule and passing through the entrance of the hole. These structural features led to the suggestion that photolyase binds to the backbone of the damaged strand and ‘‘flips’’ the cyclobutane dimer into the active site within the hole so that high-efficiency electron transfer from the flavin to the pyrimidine dimer can be effected by light (Park et al., 1995). The crystal structure shows that the center-to-center distance between MTHF and FAD is 16.8 A˚. Surprisingly, the planes of the two chromophores, and hence presumably
Fig. 4. Structure of Escherichia coli photolyase. (A) Ribbon diagram representation. The MTHF antenna is exposed on the surface, whereas the FADH catalytic cofactor is buried within the core of the -helical domain. (B) Surface potential representation. Blue, basic residues; red, acidic residues; white, hydrophobic residues. Note the positively charged groove running diagonally the length of the protein and the hole (marked by a square) with asymmetric charge distribution along the side walls and leading to the flavin located in the bottom. (See Color Insert.)
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the angle between their transition dipole moments, are nearly perpendicular to one another, which is not conducive for high-efficiency energy transfer. The structure of A. nidulans photolyase is very similar to that of the E. coli enzyme, with one important exception: the 8-HDF photoantenna is deeply buried into the interdomain cleft and the center-to-center distance between the two chromophores is 17.5 A˚. However, the planes of the two chromophores are nearly parallel, allowing for more efficient energy transfer from the second chromophore to FAD, even though they are farther apart than the two chromophores in E. coli photolyase (Kim et al., 1992; Tamada et al., 1997). The T. thermophilus photolyase has a shorter interdomain loop and is overall more compact, with more extensive interdomain contacts between the two domains, consistent with its thermostability (Komori et al., 2001).
IV. Reaction Mechanism of Photolyase The reaction mechanism of photolyase has been investigated in considerable detail (Sancar, 1994, 2003). In classical enzymological terminology, photolyase performs catalysis by a ‘‘sequential ordered mechanism’’ (Fig. 5): the enzyme must bind to one substrate (Pyr<>Pyr) first before it can bind (absorb) the second substrate (a photon) and carry out catalysis. In contrast to all other enzymes, however, the second substrate is not another molecule, but a photon that can excite the cofactors of the enzyme (binding equivalent) in a femtosecond. This unique property of photolyase has been used advantageously to analyze the various kinetic steps both in vivo and in vitro by carrying out the binding under yellow light that does not excite the enzyme, and then delivering the photoreactivating photon in light pulses of duration ranging from 20 fs to 1 ms (Kim et al., 1991; Langenbacher et al., 1997; MacFarlane and Stanley, 2003). Remarkably, when the kinetic constants for the reaction k1
h EþP EþSÐ ES ! kp k 2
obtained by flash photolysis in vivo and in vitro are compared, the agreement between the two sets of values is excellent (Sancar et al., 1987). Such a comparison can be made for only a very limited enzyme system at present, and the results obtained by photolyase validate the relevance of in vitro thermodynamic and kinetic parameters to the in vivo reactions. In the following text, we will analyze the binding of and catalysis by
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Fig. 5. Reaction mechanism of photolyase. (A) Binding (dark reaction). The enzyme binds DNA containing a T<>T by random collision and flips out the dimer into the active site pocket. Enzyme–substrate complex formation is a thermal reaction (kT), independent of light. Following the light (hv) reaction, the repaired dinucleotide is ejected from the active site cavity and DNA dissociates from the enzyme. (B) Catalysis (light reaction). The photoantenna chromophore MTHF absorbs a photon and transfers the excitation energy to FADH, which then transfers an electron to T<>T to generate a biradical. The cyclobutane ring is split, and the electron returns to FADH to regenerate catalytically active FADH. The repaired thymine dinucleotide is extruded from the active site, and the enzyme dissociates from DNA. (See Color Insert.)
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photolyase separately because these two events are temporally separated under natural reaction conditions.
A. Binding of Photolyase to Substrate Photolyase binds to Pyr<>Pyr in DNA with a second-order rate constant (kon 108107 M1 sec1) consistent with target location by three-dimensional diffusion; there is no evidence for a diffusion-controlled reaction in reduced dimensionality (Husain and Sancar, 1987), as has been found for some sequence-specific DNA binding proteins. The specific binding constant to a T<>T in DNA is KS ¼ 109 M, and the nonspecific binding constant for a dinucleotide in undamaged DNA is KNS 104 M (Husain and Sancar, 1987). Thus, the selectivity factor of the enzyme for Pyr<>Pyr is KS/KNS 105. This high selectivity is achieved by the somewhat unique backbone structure of DNA containing a Pyr<>Pyr, and by the dinucleotide flipping of Pyr<>Pyr in the active site in preference to a Pyr–Pyr dinucleotide. Finally, absolute specificity is conferred by the chemical proofreading step, whereby the excited-state flavin can donate an electron to a Pyr<>Pyr but not to a nondamaged base that may happen to be in the active site. The presence of a T<>T in a duplex causes 9 degree unwinding and about 30 degree kinking into the major groove, as revealed by both solution (Husain et al., 1988) and x-ray crystallographic analyses (Park et al., 2002). The crystal structure of a decamer duplex with a T<>T was solved recently and is shown in Figure 6. This unique structure of the duplex, and in particular that of the damaged strand with which photolyase makes nearly all of its contacts (Husain et al., 1987), provides a considerable degree of specificity. However, similar but not identical backbone distortions are caused by other DNA lesions, and as a consequence, such lesions that are not repairable by photolyase can constitute high-affinity binding sites. For example, E. coli photolyase binds to cisplatin-d(GpG) diadduct with affinity close to its affinity for a Pyr<>Pyr, and it stimulates the repair of both lesions by nucleotide excision repair (Sancar ¨ zer et al., 1995). Thus, backbone distortion is an important et al., 1984; O but insufficient structural determinant of specific binding of photolyase to a Pyr<>Pyr in DNA. The second level of specificity is achieved by what might be called an induced-fit mechanism. It has been proposed that photolyase, in a manner similar to DNA methyltransferases (Roberts and Cheng, 1998), ‘‘flips out’’ its substrate from within the duplex to the active site in the enzyme (Park et al., 1995). The size and distribution of the polar, charged, and aromatic residues within the hole leading to flavin are such that only a cyclobutane pyrimidine dimer can be accommodated
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Fig. 6. Schematic diagram illustrating the kink induced in DNA by T<>T (Park et al., 2002). Regular B-DNA decamer and a T<>T containing decamer are depicted in green and blue, respectively. The thymines making up the cyclobutane dimer are drawn in red. The view (A) shows part of the major groove, and the view (B) shows the minor groove of the duplex. The phosphodeoxyribose backbone shows a sharply kinked or pinched structure (courtesy of Dr. ChulHee Kang). (See Color Insert.)
within the active site. As a consequence, the enzyme can flip out only a pyrimidine dimer into the active site hole, and even though a cocrystal structure is not available at present, most likely some conformational change in the enzyme itself and in the dimer occurs during dinucleotide flipping to optimize the flavin–dimer contacts. It must be noted that because of the loss of aromaticity, the pyrimidine moieties of the dimer are no longer planar and have lost stacking interactions. As a consequence, a Pyr<>Pyr is structurally quite different from a Pyr–Pyr dinucleotide, and the latter most likely cannot be flipped out into the active site cavity. Moreover, the Pyr–Pyr dinucleotide that forms following repair no longer fits into the cavity and is ejected from the active site. The binding of photolyase to its substrate has been extensively investigated with DNase and chemical footprinting methods (Baer and Sancar, 1989; Husain et al., 1987; Kiener et al., 1989) and with substrates ranging in
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size from 2 to 50 nt (Husain et al., 1987; Jorns et al., 1984; Kim and Sancar, 1991). These investigations have revealed that photolyase binds to a T<>T within ssDNA and dsDNA with equal affinity and that it contacts the phosphate 50 to the T<>T and the three phosphates 30 to the dimer, but not the intradimer phosphate. The enzyme has essentially the same affinity for a substrate in the form of NpT<>TpNpNpN as for a substrate of about 50 bp with a T<>T. Thus, the hexameric substrate has all the structural determinants necessary for high-affinity and high-specificity binding. However, photolyase binds to a T<>T as well as other Pyr<>Pyr dinucleotides with a KD 105M, indicating that the cyclobutane pyrimidine dimer itself contributes about half of the binding free energy. Thus, it appears that half of the binding free energy is contributed by the interaction of the positively charged groove on the enzyme with the distorted backbone of the damaged strand, and the other half is provided by dinucleotide flipping into the active site cavity where ionic, stacking, and van der Waals interactions contribute to the stability and specificity of the complex. To recapitulate, photolyase locates Pyr<>Pyr by three-dimensional diffusion, and the positively charged groove on the enzyme surface makes a low-specificity complex by ionic interactions with the 30-degree kinked damaged strand causing further distortion, resulting in the flipping out of the dimer into the active site cavity, lined at the bottom with flavin and at the sides with two tryptophans. This drastic conformational change leads to the development of a new set of interactions between the enzyme and substrate and the formation of a stable and specific complex. However, the ultimate specificity is achieved at the chemical step: Even if an undamaged dipyrimidine or another DNA lesion were to be placed in the active site, as far as is known, the enzyme can transfer an electron only to cyclobutane pyrimidine dimers, and hence the chemical step provides near-absolute specificity of photolyase for Pyr<>Pyr.
B. Catalysis by Photolyase Photolyase catalyzes light-initiated ( s2 þ s2) cycloreversion of the cyclobutane ring joining the two pyrimidine moiety in a pyrimidine dimer. Catalysis occurs by a photo-induced cyclic electron transfer reaction that does not cause a net change in the redox state of the enzyme and substrate/product at the end of the catalytic cycle (Li et al., 1991; Payne and Sancar, 1990; Sancar, 2003). The basic features of catalysis are as follows (Fig. 5B): A 300–500-nm photon is absorbed by MTHF (or 8-HDF). The excited MTHF singlet, 1MTHF*, transfers energy by fluorescence resonance energy transfer to FADH to generate 1(FADH)*, which
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within 50 ps transfers an electron to Pyr<>Pyr to generate the FADH Pyr<>Pyr biradical. The dimer radical splits to two canonical pyrimidines concomitant with back electron transfer within 0.5–2 ns to FADH to restore it to the catalytically competent FADH form. The splitting of the cyclobutane ring is thought to be by a concerted but asynchronous cleavage of the C5–C5 and C6–C6 bonds of the cyclobutane ring. Splitting of the dimer causes a considerable change in the structure of the dinucleotide that makes up the dimer and in the structure of the DNA backbone in the immediate vicinity of the dimer. As a consequence, the two pyrimidines are ejected from the active site cavity, the interaction of the positively charged groove on the photolyase surface with the DNA backbone weakens, and the enzyme dissociates from DNA to enter new rounds of catalysis. The photochemical reactions from absorbing a photon by MTHF to splitting of Pyr<>Pyr and restoration of FADH to FADH by back-electron transfer are very fast and are expected to be completed within 0.5–2.0 ns to close the photocycle. The substrate binding and product dissociation reactions are relatively slower than the photochemical reaction and therefore are the rate-determining steps in the overall catalytic cycle.
1. Quantum Yield Quantum yield, in photochemical reactions, is the ratio of the number of chemical reactions caused by light to the number of photons absorbed by the chemical species. With the exception of some rare photochemical processes in bio-inorganic chemistry, in which chain reactions initiated by absorption of a single photon result in multiple catalytic events and hence quantum yield greater than unity, in the vast majority of photochemical reactions and in all known photobiological reactions such as photosynthesis, vision, and phototropism the quantum yield is less than 1.0. The quantum yield of DNA repair by photolyase (the number of cyclobutane pyrimidine dimers split by the enzyme for each photon absorbed by the enzyme in the enzyme-substrate complex) ranges from 0.7 to 1.0. It should be noted, however, that in photolyase FADH is the catalytic cofactor and MTHF (or 8-HDF) is the photoantenna. As a consequence, the quantum yield of photolyase is the product of three reactions (Payne and Sancar, 1990): energy transfer from 1MTHF* (or 8-HDF) to FADH, electron transfer from 1(FADH)* to the Pyr<>Pyr, and finally splitting of Pyr<>Pyr . The latter two reactions are very efficient and occur with nearly 100% efficiency, at least in the case of T<>T. Therefore, the critical determinant of overall quantum yield of repair is the quantum yield of energy transfer from the photoantenna to the catalytic cofactor (Kim et al., 1991, 1992). The efficiency of energy transfer by Fo¨rster radiationless
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transfer mechanism is inversely proportional to the distance between the donor and the acceptor, and to the angle between the transition dipole moments of the donor and acceptor. Optimum efficiency is achieved when the interchromophore distance is short and the transition dipole moments are parallel. The interchromophore distance in E. coli photolyase is 16.8 A˚, and in the A. nidulans photolyase it is 17.5 A˚. However, despite the greater distance between the chromophores in A. nidulans, photolyase energy transfer from 8-HDF to FADH occurs with nearly 100% efficiency because the planes of the two chromophores (and hence presumably the transition dipole moments) are nearly parallel. In contrast, in E. coli photolyase, the transition dipole moments of MTHF and FADH are nearly perpendicular to one another, and as a consequence, energy transfer from MTHF to FADH occurs with 70%–75% efficiency. Because the quantum yields for subsequent reactions for both enzymes are identical and near unity (Kim et al., 1991, 1992), the efficiency of interchromophore energy transfer determines the overall quantum yield of repair. Thus, for E. coli photolyase and other folate class photolyases, the overall quantum yield of repair is 0.7–0.75 (Malhotra et al., 1994; Payne and Sancar, 1990), and that for deazaflavin class enzymes is very close to 1.0; that is, for every photon absorbed by the enzyme, one pyrimidine dimer is repaired.
2. Action Spectrum An action spectrum is a plot of the rate of a photochemical reaction as a function of the wavelength of light effecting the reaction. In general, the action spectrum has the shape of the absorption spectrum of the photoactive pigment catalyzing the reaction. In photolyase, an enzyme with FADH and no MTHF (or 8-HDF) is capable of repairing DNA, albeit less efficiently than the holoenzyme (Kim et al., 1992; Payne and Sancar, 1990), but enzymes containing MTHF (or 8-HDF) but no FADH are catalytically inert (Kim et al., 1991, 1992). Despite this central role of FADH in catalysis, under physiological conditions, more than 90% of the photons used for catalysis are absorbed by MTHF (or 8-HDF), and the absorption spectrum of the second chromophore determines the shape of the action spectrum of photolyase for two reasons. First, the FADH has an absorption maximum around 360 nm and an extinction coefficient of 5,000 M1 cm1 at this wavelength. In contrast, MTHF and 8-HDF have much higher extinction coefficients and absorb at longer wavelengths: The extinction coefficient of MTHF is 25,000 M1 cm1, and its absorption maximum ranges from 377 to 415 nm, depending on the particular enzyme; the extinction coefficient of 8-HDF is 44,000 M1 cm1, and its absorption maximum is at 440 nm. Second, the fraction of photons in
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Fig. 7. Absorption and action spectra of DNA photolyases. Left, Escherichia coli photolyase. Solid and broken lines represent the absorption spectra of the E-MTHFFADH and the E-FADH forms of the enzyme, and the triangles and squares represent the photolytic cross sections ( x) of the two forms. Right, Anacystis nidulans photolyase. The solid and broken lines are the absorption spectra of the E-8-HDF-FADH and the E-FADH forms of the enzyme, and the circles and triangles represent photolytic cross sections of the corresponding forms at selected wavelengths.
sunlight in the 300–350-nm range reaching the earth surface is very low compared to those >350 nm. As a consequence, most repair in nature is mediated by photons absorbed by the second chromophore, even though a photon absorbed directly by FADH is certainly more efficient in photorepair. As a general rule, MTHF class photolyases have an essentially symmetrical action spectra, with max 375–415 nm, and those in the 8-HDF class have an action spectrum nearly identical to the absorption spectrum of enzyme-bound 8-HDF, with a peak at 444 nm (Fig. 7).
V. (6–4) Photolyase The (6–4) photoproduct is the second most abundant lesion induced in DNA by UV light, constituting 10%–20% of total UV photoproducts (Taylor, 1994). In contrast to cyclobutane pyrimidine dimers that are formed from the excited triplet state of pyrimidines, the (6–4) photoproducts are formed from the pyrimidine excited singlet state. In the (6–4) photoproduct, the C6 of the 50 pyrimidine makes a sigma bond with the C4 of the 30 pyrimidine, and the OH (or NH2) group at the C4
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of the 30 pyrimidine is transferred to the C5 of the 50 pyrimidine. As a consequence, breaking the C6–C4 sigma bond either thermally or photochemically does not repair the DNA lesion but actually converts a dinucleotide adduct to two adjacent damaged bases (see Fig. 1B). The (6–4) photoproduct distorts the DNA more severely than the cyclobutane dimer and is recognized and repaired by both the bacterial (Svoboda et al., 1993) and the human (Reardon and Sancar, 2003) excision nuclease systems five- to 10-fold more efficiently than the cyclobutane dimer pyrimidine dimer. The classical photolyase neither recognizes nor repairs the (6–4) photoproduct (Brash et al., 1985). However, there is a (6–4) photoproduct-specific photolyase that reverses this lesion in a lightdependent reaction (Todo et al., 1993, 1996). The (6–4) photolyase was first discovered in Drosophila (Todo et al., 1993) and was subsequently found in many other species (Chen et al., 1994; Kim et al., 1994; Todo, 1999). The (6–4) photolyases exhibit a high level of sequence identity to photolyase, and those that have been characterized biochemically appear to contain both chromophores (Zhao et al., 1997). However, as the enzyme has been isolated only as recombinant protein expressed in heterologous sources, often the cofactors are present at substoichiometric levels. Thus, the X. laevis (6–4) photolyase expressed in E. coli contains nearly stoichiometric FAD but no detectable folate (Hitomi et al., 1997). Similarly, the D. melanogaster (6–4) photolyase expressed in E. coli contains both FAD and folate, but the former occurs at a stoichiometry of 0.01–0.05, and the latter at even lower levels relative to the apoenzyme (Zhao et al., 1997). To date, (6–4) photolyase has not been found in organisms that synthesize 5-deazaflavin, and hence there is no evidence for the presence of (6–4) photolyases that use 8-HDF or any chromophore other than folate as a photoantenna.
A. Binding of (6–4) Photolyase The Drosophila and Xenopus (6–4) photolyases appear to bind DNA containing a (6–4) photoproduct by three-dimensional diffusion and to make contacts around the lesion quite similar to the contacts made by photolyase with DNA containing a cyclobutane pyrimidine dimer (Hitomi et al., 1997; Zhao et al., 1997). The (6–4) photolyase, like the cyclobutane photolyase, binds to its cognate lesion in ssDNA and dsDNA with essentially equal affinities (Zhao et al., 1997). When bound to a dsDNA substrate, the enzyme confers single-strandedness to a 4-bp region around the lesion, and the presence of a mismatch across the (6–4) photoproduct increases the affinity of the enzyme for the substrate (Zhao et al., 1997). These three features of binding, that is, binding to substrate in ssDNA with
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high affinity, conferring single-strandedness to the bases immediately around the target, and binding with higher affinity when the target base is in the context of a mismatch, are standard criteria for a base-flipping mechanism (Roberts and Cheng, 1998), and hence it was proposed that (6–4) photolyase, like classical photolyase, employs a dinucleotide flipping mechanism to achieve specificity following the low-specificity interactions with the distorted DNA backbone at the site of the lesion (Zhao et al., 1997). Indeed, a molecular modeling study of Xenopus (6–4) photolyase using the E. coli photolyase C backbone as a template revealed a positively charged groove on the surface of the enzyme and a pocket in the center of the groove leading to the FADH in the core of the -helical domain, and the cavity appears to have the appropriate size and charge distribution to accommodate a (6–4) photoproduct (Todo, 1999). The equilibriumbinding constants of (6–4) photolyases of Drosophila and Xenopus are in the range of KD ¼ 0.5–1.0 109 M, and the dissociation rate constant is koff 103 s1 to 105 s1 (t1/2 10 to 100 min). Thus, it has been concluded that the formation of the enzyme–substrate complex is diffusion controlled and the main determinant of high specificity is the slow off rate of dissociation of the enzymes from complexes formed at the damage site. The dissociation rate following repair has not been determined, but it is expected to be much faster than that of unrepaired substrate. Increasing the off rate several orders of magnitude would still be much slower than the photochemical reaction, which is most likely complete within a nanosecond or less. Hence, under substrate saturating (damage and photon) conditions for (6–4) photolyase, as in the case of photolyase, the rate-determining step in the overall reaction is the dissociation of the repaired product.
B. Catalysis by (6–4) Photolyase Catalysis by (6–4) photolyase must accomplish two chemical tasks: cleavage of the C6–C4 sigma bond, and transfer of the OH (or NH2) group from the C5 of the 50 base to the C4 of the 30 base. Because formation of the (6–4) photoproduct is presumed to proceed through a four-membered oxetane or azetidine intermediate, it has been proposed that (6–4) photolyase first converts the ‘‘open’’ form of the (6–4) photoproduct to the four-membered ring by a thermal reaction, and then the four-membered ring is cleaved by retro [2+2] reaction photochemically (Kim et al., 1994; Zhao et al., 1997). A site-directed mutagenesis study has identified two histidine residues in the active site that may participate in conversion of the (6–4) photoproduct to the oxetane intermediate by general acid– base catalysis (Hitomi et al., 2001). A current model for catalysis by (6–4) photolyase is as follows (Fig. 8): The enzyme binds DNA and flips out the
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Fig. 8. Reaction mechanism of (6–4) photolyase. The enzyme binds to DNA containing a (6–4) photoproduct and flips out the dinucleotide adduct into the active site cavity, where the ‘‘open’’ form of the photoproduct is converted to the oxetane intermediate by a light-independent general acid-base mechanism. Catalysis is initiated by light; MTHF absorbs a photon and transfers energy to FADH, which then transfers an electron to the oxetane intermediate; bond rearrangement in the oxetane radical regenerates two canonical pyrimidines, and back-electron transfer restores the flavin radical to catalytically competent FADH form. The repaired dipyrimidine flips back into the DNA duplex, and the enzyme is dissociated from the substrate.
(6–4) photoproduct into the active site cavity, where the photoproduct is converted into the oxetane form thermally. A 350–450-nm photon is absorbed by the folate photoantenna, which transfers energy to FADH. The 1(FADH)* transfers an electron to the oxetane ring initiating the cycloreversion reaction, which is followed by back-electron transfer to restore the flavin radical (Zhao et al., 1997). This is a plausible model; however, at present, direct evidence for energy transfer from the photoantenna to flavin is lacking. Evidence for electron transfer from flavin to substrate was obtained by demonstration of a requirement for reduction of flavin either chemically or photochemically for catalysis (Hitomi et al., 1997; Zhao et al., 1997). Strong support for the proposed mechanism was provided by a study with a model system (Cichon et al., 2002): an oxetane ring was covalently linked to flavin, and its cleavage by
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light was investigated under a variety of conditions. It was found that only two-electron reduced and deprotonated flavin induced photosplitting of the oxetane ring at a significant rate. Clearly, all indications are that (6–4) photolyase binds DNA and repairs its substrate by a mechanism quite similar to that of classical photolyase. However, there appears to be a fundamental difference in the photochemical reaction catalyzed by the two enzymes. The quantum yield of repair by excited singlet-state flavin by classical photolyase is near unity, whereas the quantum yield of repair by excited flavin in (6–4) photolyase is 0.05–0.10. Whether this low quantum yield of repair by (6–4) photolyase is a result of the low efficiency of formation of the oxetane intermediate thermally, low efficiency of electron transfer from the flavin to the photoproduct, or low efficiency splitting of the oxetane anion coupled with high rate of back electron transfer is not known at present. Furthermore, it was found that (6–4) photolyase can photorepair the Dewar valence isomer of the (6–4) photoproduct (Taylor, 1994) that cannot form an oxetane intermediate, casting some doubt about the basic premise of the retro [2þ2] reaction. However, the Dewar isomer is repaired with 300–400 lower quantum yield than the (6–4) photoproduct, and it has been proposed (Zhao et al., 1997) that the Dewar isomer may be repaired by the enzyme through a two-photon reaction in which the first photon converts the Dewar isomer to the Kekule form and a second electron transfer reaction initiated by the second photon promotes the retro [2þ2] reaction.
VI. Cryptochrome Cryptochrome was originally used as a generic term for blue-light photoreceptors that were known to exist and to regulate a variety of light responses in plants but whose identities remained cryptic for over a century. At present, at least three blue-light-specific receptors have been identified in plants including phototropin, FKF1, and a receptor related to photolyase (Briggs and Huala, 1999; Cashmore, 2003; Sancar, 2000). The photolyase-like receptor was the first blue-light receptor identified in plants and hence was called cryptochrome (Lin et al., 1996). When two photolyase-like proteins with no photolyase activity were discovered in humans, it was suggested that these may function as blue-light photoreceptors that regulate the circadian clock, and they were named cryptochrome 1 and cryptochrome 2 as well (Hsu et al., 1996). At present, the term ‘‘cryptochrome’’ has acquired a precise meaning: a photolyase-like protein with no DNA repair activity but with known or presumed blue-light receptor functions (Sancar, 2000).
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Cryptochrome has been found in bacteria, plants, and animals (Cashmore, 2003). Its role in bacteria is not known (Brudler et al., 2003; Ng and Pakrasi, 2001; Worthington et al., 2003), though it regulates growth and development in plants (Cashmore, 2003; Lin and Shalitin, 2003) and the circadian clock in animals (Sancar, 2003; Thompson and Sancar, 2002). Both the circadian clock and DNA repair play a role in ameliorating the harmful effects of sunlight. Thus, it is conceivable that in the distant past, when more UV light reached the surface of the earth, an ancestor of modern lifeforms contained a photoactive flavo-protein that acted as a photosensory pigment that regulated the movement of the organisms from the water surface to a depth inaccessible to UV, and at the same time repaired the UV-induced DNA lesions that may have been induced before escape. Apparently, during evolution, these two functions diverged so that in present-day organisms, photoreactivation and circadian photoentrainment are mediated by evolutionarily and structurally related blue-light photoreceptors called photolyase and cryptochrome, respectively (Sancar et al., 2000).
A. Structure Cryptochromes exhibit 20%–40% sequence identities to photolyases (Cashmore et al., 1999; Todo, 1999). With the exception of V. cholerae cryptochrome 1 (Worthington et al., 2003), all cryptochromes characterized to date have only been isolated by expressing the cryptochrome genes in heterologous systems, mainly in E. coli (Hsu et al., 1996; Malhotra et al., 1995) and in insect cells (Lin et al., 1996). When heterologously expressed cryptochromes are purified from such sources, they contain very little or no folate and usually substoichiometric flavin in an oxidized state. The only exception is V. cholerae cryptochrome 1, which, when purified as a recombinant protein expressed in either E. coli or in V. cholerae, contains both the folate and the flavin chromophores in essentially one-to-one stoichiometry with the apoenzyme and the flavin in the two-electron reduced state (Worthington et al., 2003). The biochemical properties of VcCry1 are the strongest evidence to date that cryptochromes may function in a manner analogous to photolyases. The crystal structure of the Synechocystis cryptochrome obtained by molecular replacement, using the E. coli photolyase as a template (Brudler et al., 2003), and the three-dimensional structure of human cryptochrome 2 obtained by molecular modeling onto the C backbone of E. coli ¨ zgu¨r and Sancar, 2003) reveals a basically photolyase-like photolyase (O structure including, somewhat surprisingly, the positively charged groove involved in binding to DNA and the hole in the middle of this groove leading to the flavin in the core of the molecule (Fig. 9). It appears,
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Fig. 9. Model for human cryptochrome 2. The model was computer generated using the Escherichia coli DNA photolyase as a template; the C-terminal 80 amino acids of hCRY2 were excluded. Left, ribbon representation. Right, surface potential representation. Note the presence of the positively charged groove on the surface and passing through the hole leading to the FAD cofactor in the core of the -helical domain. (See Color Insert.)
however, that the FAD in cryptochromes is more accessible to solvents than the FAD of photolyases. Most cryptochromes contain 20–200–amino acid C-terminal extensions beyond the photolyase homology region. Interestingly, it has been found that expression of this C-terminal domain of Arabidopsis cryptochromes confers a constitutive ‘‘light-on’’ phenotype (Yang et al., 2001). Thus, it appears that the C terminus acts as the effector function that is somewhat repressed by the photolyase-like region in the dark and relieved in the light.
B. Function of Cryptochrome In contrast to photolyases, the photochemical reaction carried out by cryptochrome is not known. As a consequence, despite overwhelming genetic evidence that cryptochromes function as photoreceptors, a legitimate argument can be made that cryptochromes are simply molecules involved in phototransduction but not photoreception (Sancar, 2000; Van Gelder, 2002). The following reactions have been detected by in vivo and in vitro biochemical experiments: first, A. thaliana CRY 2 but not CRY 1 is degraded on exposure to light (Lin and Shalitin, 2003), and presumably the
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same is also true for D. melanogaster cryptochrome (Stanewsky et al., 1998). Second, human and Drosophila cryptochromes bind to their cognate clock proteins called Tim and Per (see Reppert and Weaver, 2002). It was reported that dCry-dTim interaction in the yeast two-hybrid system was light dependent (Griffin et al., 1999) but that the interactions of the human cryptochromes with Per1 and Per2 proteins in the same system were light independent. It was therefore suggested that the Drosophila, but not the human cryptochrome, has a photoreceptor function (Ceriani et al., 1999). However, when considering the fact that human cryptochromes expressed in heterologous systems contain either grossly substoichiometric or no cofactors, these findings are open to alternative interpretations. Indeed, a more recent study found that dCry lacking the C-terminal 20 amino acids interacts with both dTim and dPer in a light-independent manner in the yeast two-hybrid assay (Rosato et al., 2001). Thus, it appears that although the yeast two-hybrid system is useful in detecting protein-protein interactions, it is prone to artifacts when used for proteins with intrinsic chromophores. Third, it has been reported that AtCRY 1, At CRY 2, and human CRY 1 are serine/threonine-specific protein kinases, and moreover, the kinase activity of Arabidopsis cryptochromes was strongly stimulated by light (Bouly et al., 2003; Shalitin et al., 2003). These are intriguing findings; however, how autophosphorylation may initiate or regulate signal transduction is unclear. Indeed, many years after the discovery of autophosphorylating kinase activities of two classes of light receptors in plants, phytochrome and phototropin, the significance of the kinase activities of these proteins to their photoreception/phototransduction functions remains controversial (Briggs and Huala, 1999). Fourth, it has been found that human CRY 2 interacts with serine/threonine phosphatase PP5 and modulates its activity (Zhao and Sancar, 1997). Similarly, Arabidopsis CRY 1 interacts with the PP7 serine/threonine phosphatase PP7, and this interaction is necessary for blue-light response (Moller et al., 2003). Fifth, cryptochromes bind to nucleic acids. Human CRY 2 binds to DNA with modest affinity and with higher affinity to DNA containing a (6–4) photoproduct; however, binding to the ¨ zgu¨r and photolesion, in contrast to photolyases, is not affected by light (O Sancar, 2003). Similarly, V. cholerae cryptochrome 1 binds to RNA (Worthington et al., 2003), although the significance and specificity of this binding is unknown at present. In contrast to the paucity of biochemical data on the photosensory functions of cryptochromes, there are extensive genetic and cell biology data on the roles of cryptochromes in blue light, photoreception in plants and animals, and circadian clock regulation in animals (Cashmore, 2003; Lin and Shalitin, 2003; Sancar, 2003). In Arabidopsis, blue light inhibits elongation of hypocotyls in a cryptochrome-dependent manner. In animals,
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cryptochromes were first discovered in humans and were hypothesized to be the photosensory pigment for regulating the circadian clock (Hsu et al., 1996). Indeed, both mammalian cryptochromes are expressed at relatively high levels in the inner retina, a region capable of synchronizing the circadian clock with the daily light–dark cycle, independent of the outer retina that is required for vision (Miyamoto and Sancar, 1998; Thompson et al., 2003) (Fig. 10A). Both cryptochromes are also expressed in all mammalian cells at moderate levels, and mCRY 1 is expressed at a particularly high level in the master circadian pacemaker in an area in the hypothalamus above the optic nerve, called the suprachiasmatic nuclei (SCN). Moreover, the expression of both CRY 1 and CRY 2 exhibits circadian (daily) rhythmicity, peaking at about 2:00 p.m. in the SCN and reaching a nadir at about 2:00 a.m.; expression in peripheral organs is 4–6 hours out of phase with that of the SCN expression (Miyamoto and Sancar, 1999). Mice lacking cryptochromes are seriously compromised in photoreception/phototransduction to the SCN (Selby et al., 2000; Thompson et al., 2004; Thresher et al., 1998; Vitaterna et al., 1999) but are not circadian blind because of functional redundancy between the cryptochromes and opsins that are expressed in the outer and inner retina. Mutation in the sole Drosophila cryptochrome has a similar effect on circadian photoreception in this organism (Stanewsky et al., 1998), consistent with functional redundancy of opsins and cryptochromes in circadian photoreception. In addition to their light-dependent effect of unknown mechanism, mammalian cryptochromes have a light-independent function that is necessary for normal functioning of the circadian clock. Thus, wild-type mice kept in constant darkness maintain a circadian rhythm of activity and rest phases with 23.7 hour periodicity. In mice lacking Cry2, the period is longer, at 24.7 hours (Thresher et al., 1998); in mice lacking Cry1 the period is shorter, 22.5 hours (Van der Horst et al., 1999; Vitaterna et al., 1999); and mice lacking both cryptochromes are arrhythmic (Van der Horst et al., 1999; Vitaterna et al., 1999). Co-immunoprecipitation and reporter gene assays have led to a considerable insight into the ‘‘clock function’’ of the cryptochrome. In mice, the molecular clock, which engenders the behavioral clock, is made up of transcription factors CLOCK, BMAL1, PER1, PER2, and CRY 1 and CRY 2 (Reppert and Weaver, 2002) (Fig. 10B). CLOCK and BMAL1 make a heterodimer that binds to the E-box in the promoters of PER and CRY genes and stimulate their transcription; the CRY and PER proteins make combinatorial heterodimers in the cytoplasm, which translocate into the nucleus and bind to the CLOCK protein, interfering with its activator function. Because of necessity for posttranslational modification of the various components of the molecular clock and the delay between protein synthesis and nuclear translocation, the transcription-stimulating activity
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Fig. 10. Circadian photoreception and the molecular clock in mammals. (A) Circadian and visual photoreception/phototransduction in mammals. Light signal received by the rods and cones in the outer retina is transmitted to the visual cortex by the optic nerve (blue). Light signal received by cryptochromes and inner retinal opsins is transmitted to the circadian center in the midbrain, an area called the suprachiasmatic nuclei (SCN) by a subset of the optic nerve fibers (red). (B) The molecular clock. The transcription factors clock, and BMal1 make a heterodimer that acts on the promoter of Cry and Per genes activating their transcription. The CRY and Per proteins heterodimerize in the cytoplasm, undergo posttranslational modification, and translocate into the nucleus, where they interfere with the CLOCK-BMAL1 activity and repress their own transcription as well as those of other genes regulated by clockBMal1. The transcriptional activation/inhibition cycle has a period of about 24 hours resulting in daily oscillation of clock-controlled functions. (See Color Insert.)
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of the BMAL1-CLOCK heterodimer exhibits periodicity of about 24 hours. In the absence of cryptochromes, there is no inhibition of BMAL1-CLOCK heterodimer, which causes constitutively high levels of clock gene transcripts (Vitaterna et al., 1999), resulting in molecular and behavioral arrhythmicity. As is apparent from this summary, the lightindependent function of cryptochrome is well understood in animals, but the photoreceptive function remains ill defined.
VII. Conclusion The photolyase/cryptochrome blue-light photoreceptor family encompasses a large group of proteins from all three biological kingdoms. These proteins absorb near-UV/blue light and use the light energy to repair far UV-induced DNA damage or to reset the circadian clock. Both photolyase and cryptochrome also perform light-independent functions in DNA repair and in generating the molecular circadian clock, respectively. In addition, cryptochromes regulate blue-light-dependent growth and development in plants. Finally, cryptochromes have now been identified in the nonphotosynthetic bacterium, V. cholerae, which has no known photoresponses. Future work is likely to uncover novel light-dependent and light-independent functions mediated by cryptochromes.
Acknowledgments I thank my student Carrie L. Partch for her critical comments on the manuscript and for preparing the figures. I am grateful to Professor ChulHee Kang and Dr. H. Park for providing Figure 6. This work was supported by NIH grant GM31082.
References Ahmad, M., and Cashmore, A. R. (1993). HY4 gene of A. thalina encodes a protein with characteristic of a blue-light photoreceptor. Nature 366, 162–166. Baer, M., and Sancar, G. B. (1989). Photolyases from Saccaromyces cerevisiae and Escherichia coli recognize common binding determinants in DNA containing pyrimidine dimers. Mol. Cell. Biol. 9, 4777–4788. Bouly, J. P., Giovani, B., Djamei, A., Mueller, M., Zeugner, A., Dudkin, E. A., Batschauer, A., and Ahmad, M. (2003). Novel ATP-binding and autophosphorylation activity associated with Arabidopsis and human cryptochrome-1. Eur. J. Biochem. 270, 2921–2928. Brash, D. E., Franklin, W. A., Sancar, G. B., Sancar, A., and Haseltine, W. A. (1985). E. coli photolyase reverses cyclobutane pyrimidine dimers but not pyrimidine-pyrimidone (6–4) photoproducts. J. Biol. Chem. 260, 11438–11441. Briggs, W. R., and Huala, E. (1999). Blue-light photoreceptors in higher plants. Annu. Rev. Cell Dev. Biol. 15, 33–62.
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Brudler, R., Hitomi, K., Daiyasu, H., Toh, H., Kucho, K., Ishiura, M., Kanehisa, M., Roberts, V. A., Todo, T., Tainer, J. A., and Getzoff, E. D. (2003). Identification of a new cryptochrome class: Structure, function, and evolution. Mol. Cell 11, 59–67. Cashmore, A. R. (2003). Cryptochromes: enabling plants and animals to determine circadian time. Cell 114, 537–543. Cashmore, A. R., Jarillo, J. A., Wu, Y. J., and Liu, D. (1999). Cryptochromes: Blue light receptors for plants and animals. Science 284, 760–765. Ceriani, M. F., Darlington, T. K., Staknis, D., Mas, P., Petti, A. A., Weitz, C. J., and Kay, S. A. (1999). Light-dependent sequestration of TIMELESS by CRYPTOCHROME. Science 285, 553–556. Chen, J. J., Mitchell, D. L., and Britt, A. B. (1994). A light-dependent pathway for elimination of UV-induced pyrimidine (6–4) pyrimidone photoproducts of Arabidopsis. Plant Cell 6, 1311–1317. Eker, A. P. M., Hessels, J. K. C., and Van de Velde, J. (1988). Photoreactivating enzyme from the green alga Scendesmus acutus. Evidence for the presence of two different flavin chromophores. Biochemistry 27, 1758–1765. Eker, A. P. M., Kooiman, P., Hessels, J. K. C., and Yasui, A. (1990). DNA photoreactivating enzyme from the cyanobacterium Anacystis nidulans. J. Biol. Chem. 265, 8009–8015. Griffin, E. A., Jr., Staknis, D., and Weitz, C. J. (1999). Light-independent role of CRY1 and CRY2 in the mammalian circadian clock. Science 286, 768–781. Hitomi, K., Kim, S. T., Iwai, S., Jarima, N., Otoshi, E., Ikenaga, M., and Todo, T. (1997). Binding and catalytic properties of Xenopus (6–4) photolyase. J. Biol. Chem. 272, 32591–32598. Hitomi, K., Nakamura, H., Kim, S. T., Mizukoshi, T., Ishikawa, T., Iwai, S., and Todo, T. (2001). Role of two histidines in the (6–4) photolyase reaction. J. Biol. Chem. 276, 10103–10109. Husain, I., Griffith, J., and Sancar, A. (1988). Thymine dimers bend DNA. Proc. Natl. Acad. Sci. USA 85, 2258–2262. Husain, I., and Sancar, A. (1987). Binding of E. coli DNA photolyase to defined substrate containing a single T<>T dimer. Nucleic Acids Res. 15, 1109–1120. Husain, I., Sancar, G. B., Holbrook, S. R., and Sancar, A. (1987). Mechanism of damage recognition by E. coli DNA photolyase. J. Biol. Chem. 262, 13188–13197. Hsu, D. S., Zhao, X., Zhao, S., Kazantsev, A., Wang, R. P., Todo, T., Wei, Y. F., and Sancar, A. (1996). Putative human blue-light photoreceptors hCRY1 and hCRY 2 are flavoproteins. Biochemistry 35, 13871–13877. Johnson, J. L., Hamm-Alvarez, S., Payne, G., Sancar, G. B., Rajagopalan, K. V., and Sancar, A. (1988). Identification of the second chromophore of Escherichia coli and yeast DNA photolyases as 5, 10-methenyltetrahydrofolate. Proc. Natl. Acad. Sci. USA 85, 2046–2050. Jorns, M. S., Sancar, G. B., and Sancar, A. (1984). Identification of a neutral flavin radical and characterization of a second chromophore in E. coli DNA photolyase. Biochemistry 23, 2673–2679. Kiener, A., Husain, I., Sancar, A., and Walsh, C. (1989). Purification and properties of Methanobacterium thermoautotrophicum DNA photolyase. J. Biol. Chem. 264, 13880–13887. Kim, S. T., Heelis, P. F., Okamura, T., Hirata, Y., Mataga, N., and Sancar, A. (1991). Determination of rates and yields of interchromophore energy transfer and intermolecular electron transfer in E. coli photolyase by time-resolved fluorescence and absorption spectroscopy. Biochemistry 30, 11262–11270.
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Kim, S. T., Heelis, P. F., and Sancar, A. (1992). Energy transfer (deazaflavin ! FADH2) and electron transfer (FADH2 ! T<>T) kinetics in Anacystics nidulans photolyase. Biochemistry 31, 11244–11248. Kim, S. T., Malhotra, K., Smith, C. A., Taylor, J. S., and Sancar, A. (1994). Characterization of (6-4) photoproduct DNA photolyase. J. Biol. Chem. 269, 8535–8540. Kim, S. T., and Sancar, A. (1991). Effect of base, pentose, and phosphodiester backbone structures on binding and repair of pyrimidine dimers by E. coli DNA photolyase. Biochemistry 30, 8623–8630. Kleine, T., Lockhart, P., and Batschauer, A. (2003). An Arabidopsis protein closely related to Synechocystis cryptochrome is targeted to organelles. Plant J. 35, 93–103. Komori, H., Masui, R., Kuramitsu, S., Yokoyama, S., Shibata, T., Inoue, Y., and Miki, K. (2001). Crystal structure of thermostable DNA photolyase: Pyrimidine dimer recognition mechanism. Proc. Natl. Acad. Sci. USA 98, 13560–13565. Kume, K., Zylka, M. J., Sriram, S., Shearman, L. P., Weaver, D. R., Jin, X., Maywood, W. S., Hastings, M. H., and Reppert, S. M. (1999). mCry1 and mCry2 are essential components of the negative limb of the circadian feedback loop. Cell 98, 193–205. Langenbacher, T., Zhao, X., Bieser, G., Heelis, P. F., Sancar, A., and Michel-Beyerle, M. E. (1997). Substrate and temperature dependence of DNA photolyase repair activity examined with ultrafast spectroscopy. J. Am. Chem. Soc. 119, 10532–10536. Li, Y. F., Heelis, P. F., and Sancar, A. (1991). Active site of DNA photolyase: Trp306 is the intrinsic H-atom donor essential for flavin radical photoreduction and DNA repair in vitro. Biochemistry 30, 6322–6329. Li, Y. F., Kim, S. T., and Sancar, A. (1993). Evidence for lack of DNA photoreactivating enzyme in humans. Proc. Natl. Acad. Sci. USA 90, 4389–4393. Lin, C., Ahmad, M., and Cashmore, A. R. (1996). Arabidopsis cryptochrome1 is a soluble protein mediating blue light-dependent regulation of plant growth and development. Plant J. 10, 893–902. Lin, C., and Shalitin, D. (2003). Cryptochrome structure and signal transduction. Annu. Rev. Plant Biol. 54, 469–496. MacFarlane, A. W., IV, and Stanley, R. J. (2003). Cis-syn thymidine dimer repair by DNA photolyase in real time. Biochemistry 42, 8558–8568. Malhotra, K., Kim, S. T., Walsh, C. T., and Sancar, A. (1992). Roles of FAD and 8hydroxy-5-deazaflavin chromophores in photoreactivation by Anacystis nidulans DNA photolyase. J. Biol. Chem. 267, 15406–15411. Malhorta, K., Kim, S. T., and Sancar, A. (1994). Characterization of a medium wavelength type DNA photolyase from Bacillus firmus. Biochemistry 33, 8712–8718. Malhotra, K., Kim, S. T., Batschauer, A., Dawut, L., and Sancar, A. (1995). Putative bluelight photoreceptors from Arabidopsis thaliana and Sinapsis alba with a high degree of sequence homology to DNA photolyase cofactors but lack DNA repair activity. Biochemistry 34, 6892–6899. Miyamoto, Y., and Sancar, A. (1998). Vitamin B2-based blue-light photoreceptors in the retinohypothalamic tract as the photoactive pigments for setting the circadian clock in mammals. Proc. Natl. Acad. Sci. USA 95, 6097–6102. Miyamoto, Y., and Sancar, A. (1999). Circadian regulation of the cryptochrome genes in the mouse. Mol. Brain Res. 71, 248–253. Moller, S. G., Kim, Y. S., Kunkel, T., and Chua, N. H. (2003). PP7 is a positive regulator of blue-light signaling in Arabidopsis. Plant Cell 15, 1111–1119. Ng, W. O., and Pakrasi, H. B. (2001). DNA photolyase homologs are the major UV resistance factors in the Cyanobacterium Synechocystis sp. PCC6803. Mol. Gen. Genet. 264, 924–930.
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¨ zgu¨r, S., and Sancar, A. (2003). Purification and properties of human blue-light O photoreceptor cryptochrome 2. Biochemistry 42, 2926–2932. ¨ zer, Z., Reardon, J. T., Hsu, D., Malhorta, K., and Sancar, A. (1995). The other O function of DNA photolyase: stimulation of excision repair of chemical damage to DNA. Biochemistry 34, 15886–15889. Park, H. J., Zhang, K., Ren, Y., Nadji, S., Sinha, N., Taylor, J. S., and Kang, C. (2002). Crystal structure of a DNA decamer containing a thymine dimer. Proc. Natl. Acad. Sci. USA 99, 15965–15970. Park, H. W., Kim, S. T., Sancar, A., and Deisenhofer, J. (1995). Crystal structure of DNA photolyase from Escherichia coli. Science 268, 1866–1872. Payne, G., and Sancar, A. (1990). Absolute action spectrum of E-FADH2 and E-FADH2MTHF forms of Escherichia coli DNA photolyase. Biochemistry 29, 7715–7727. Reppert, S. M., and Weaver, D. R. (2002). Coordination of circadian timing in mammals. Nature 418, 935–941. Roberts, R. J., and Cheng, X. (1998). Base flipping. Annu. Rev. Biochem. 67, 181–198. Sancar, A. (1994). Structure and function of DNA photolyase. Biochemistry 33, 2–9. Sancar, A. (2000). Cryptochrome: The second photoactive pigment in the eye and its role in circadian photoreception. Annu. Rev. Biochem. 69, 31–67. Sancar, A. (2003). Structure and function of DNA photolyase and cryptochrome bluelight photoreceptors. Chem. Rev. 103, 2203–2237. Sancar, A., Franklin, K. A., and Sancar, G. B. (1984). Escherichia coli DNA photolyase stimulates UvrABC excision nuclease in vitro. Proc. Natl. Acad. Sci. USA 81, 7397–7401. Sancar, A., and Sancar, G. B. (1984). Escherichia coli DNA photolyase is a flavoprotein. J. Molec. Biol. 172, 223–227. ¨ zgu¨r, S., Vagas, E., Sancar, A., Thompson, C. L., Thresher, R. J., Araujo, F., Mo, J., O Dawut, L., and Selby, C. P. (2000). Photolyase/cryptochrome family blue-light photoreceptors use light energy to repair DNA or set the circadian clock. Cold Spring Harbor Symp. Quant. Biol. 65, 157–171. Sancar, G. B., Jorns, M. S., Payne, G., Fluke, D. J., Rupert, C. S., and Sancar, A. (1987). Action mechanism of DNA photolyase.III. Photolysis of the ES complex and the absolute action spectrum. J. Biol. Chem. 262, 492–498. Selby, C. P., Thompson, C., Therese, S. M., Van Gelder, R. N., and Sancar, A. (2000). Functional redundancy of cryptochromes and classical photoreceptors for nonvisual ocular photoreception in mice. Proc. Natl. Acad. Sci. USA 97, 14697–14702. Shalitin, D., Yu, X., Maymon, M., Mockler, T., and Lin, C. (2003). Blue-light dependent in vivo and in vitro phosphorylation of Arabidopsis cryptochrome 1. Plant Cell 15, 2421–2429. Srinivasan, V., Schnitzlein, W. M., and Tripathy, D. K. (2001). Fowlpox encodes a novel repair enzyme, CPD-photolyase, that restores infectivity of UV light-damaged virus. J. Virol. 75, 1681–1688. Stanewsky, R., Kaneko, M., Emery, P., Beretta, B., Wager-Smith, K., Kay, S. A., Rosbash, M., and Hall, J. C. (1998). The cryb mutation identifies cryptochrome as a circadian photoreceptor in Drosophila. Cell 95, 681–692. Svoboda, D. L., Smith, C. A., Taylor, J. -S., and Sancar, A. (1993). Effect of sequence, adduct type, and opposing lesions on the binding and repair of UV photodamage by DNA photolyase and (A)BC exinuclease. J. Biol. Chem. 268, 10694–10700. Tamada, T., Kitadokoro, K., Higuchi, Y., Inaka, K., Yasui, A., de Ruiter, P. E., Eker, A. P. M., and Miki, K. (1997). Crystal structure of DNA photolyase from Anacystis nidulans. Nat. Struct. Biol. 4, 887–891. Taylor, J. S. (1994). Unraveling the molecular pathway from sunlight to skin cancer. Acc. Chem. Res. 27, 76–82.
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Thompson, C. L., Blaner, W. S., Van Gelder, R. N., Lai, K., Quadro, L., Colantuoni, V., Gottesman, M. E., and Sancar, A. (2001). Preservation of light-signaling to the suprachiasmatic nucleus in vitamin A-deficient mice. Proc. Natl. Acad. Sci. USA 98, 11708–11713. Thompson, C. L., Bowes Rickman, C., Shaw, S. J., Ebright, J. N., Kelly, U., Sancar, A., and Rickman, D. W. (2003). Expression of the cryptochrome blue-light photoreceptors in the human retina. Invest. Ophthalmol. Vis. Sci. 44, 4515–4521. Thompson, C. L., and Sancar, A. (2002). Photolyase/cryptochrome blue-light photoreceptors use photon energy to repair DNA and reset the circadian clock. Oncogene 21, 9043–9056. Thompson, C. L., Selby, C. P., Partch, C. L., Plante, D. T., Thresher, R. J., Araujo, F., and Sancar, A. (2004). Further evidence for the role of cryptochromes in retinohypothalamic photoreception/phototransduction. Molec. Brain Res. 122, 158–166. Thresher, R. J., Vitaterna, M. H., Miyamoto, Y., Kazantsev, A., Hsu, D. S., Petit, C., Selby, C. P., Dawut, L., Smithies, O., Takahashi, J. S., and Sancar, A. (1998). Role of mouse cryptochrome blue-light photoreceptor in circadian photoresponses. Science 282, 1490–1494. Todo, T. (1999). Functional diversity of the DNA photolyase/blue light receptor family. Mutat. Res. 236, 89–97. Todo, T., Ryo, H., Yamamoto, K., Toh, H., Inui, T., Ayaki, H., Nomura, T., and Ikenage, M. (1996). Similarity among the Drosophila (6–4) photolyase, a human photolyase homolog and the DNA photolyase blue light photoreceptor family. Science 272, 109–112. Todo, T., Takemori, H., Ryo, H., Ihara, M., Matsunaga, T., Nikaido, O., Sato, K., and Nomura, T. (1993). A new photoreactivating enzyme that specifically repairs ultraviolet induced (6–4) photoproducts. Nature 361, 371–374. Van der Horst, G. T. J., Muijtens, M., Kobayashi, K., Takano, R., Kanno, S., Takao, M., de Wit, J., Verkerk, A., Eker, A. P. M., van Leenen, D., Buijs, R., Bootsma, D., Hoeijmakers, J. H. J., and Yasui, A. (1999). Mammalian Cry1 and Cry2 are essential for maintenance of circadian rhythms. Nature 398, 627–630. Vitaterna, M. H., Selby, C. P., Todo, T., Niwa, H., Thompson, C., Fruechte, E. M., Hitomi, K., Thresher, R. J., Ishikawa, T., Miyazaki, J., Takahashi, J. S., and Sancar, A. (1999). Differential regulation of mammalian Period genes and circadian rhythmicity by cryptochrome 1 and 2. Proc. Natl. Acad. Sci. USA 96, 12114–12119. Van Gelder, R. N. (2002). Tales from the crypt (ochromes). J. Biol. Rhythms 17, 110–120. Wang, H., Ma, L. G., Li, J. M., Zhao, H. Y., and Deng, X. W. (2001). Direct interaction of Arabidopsis cryptochromes with COP1 in light control development. Science 294, 154–158. Willer, D. O., McFadden, G., and Evans, D. H. (1999). The complete genome sequence of Shoppe (rabbit) fibroma virus. Virology 264, 319–343. Worthington, E. N., Kavakli, I. H., Berrocal-Tito, G., Bondo, B. E., and Sancar, A. (2003). Purification and characterization of three members of the photolyase/ cryptochrome family blue-light photoreceptors from Vibrio cholerae. J. Biol. Chem. 278, 39143–39154. Yang, H. Q., Wu, Y. J., Tang, R. H., Liu, Y., and Cashmore, A. R. (2000). The C termini of Arabidopsis cryptochromes mediate a constitutive light response. Cell 103, 815–827. Zhao, S., and Sancar, A. (1997). Human blue-light photoreceptor hCRY2 specifically interacts with protein serine/threonine phosphatase 5 (PP5) and modulates its activity. Photochem. Photobiol. 66, 727–731. Zhao, X., Liu, J., Hsu, D. S., Zhao, S., Taylor, J. S., and Sancar, A. (1997). Reaction mechanism of (6–4) photolyase. J. Biol. Chem. 272, 32580–32590.
COORDINATION OF REPAIR, CHECKPOINT, AND CELL DEATH RESPONSES TO DNA DAMAGE By JEAN Y. J. WANG AND SARAH K. CHO Division of Biological Sciences and the Moores Cancer Center, University of California, San Diego, La Jolla, California 92093
I. Introduction . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . II. Overview of Biological Responses to DNA Damage. .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. General Framework of DNA Damage Responses. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. Temporal Coordination of DNA Damage Responses. . . . . . . . . . . . . . . . . . . .. . . . . . C. Other Comments on the General Framework . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . III. Molecular Components for the Initiation of DNA Damage Responses. . . .. . . . . . A. DNA Damage Sensor . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. Master Switch: PIKK Family of Protein Kinases. .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . C. Adaptors and Mediators. . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . D. Effector Kinases: Chk1 and Chk2. . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . E. From Components to Mechanisms . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . IV. Apoptotic Effectors in DNA Damage Response . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Role of p53 in DNA Damage Response . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. p53-Related Proteins: p63 and p73 . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . C. Abl Tyrosine Kinase. . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . D. Stress-Activated Protein Kinases: JNK and p38 . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . V. DNA Repair Proteins in Damage Signaling . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Mismatch Repair Proteins. .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. UV-DDB Complex. . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . C. MRE11-RAD50 Complex . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . VI. Alternative Models for the Temporal Coordination of DNA Damage Responses . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Integrative Surveillance. . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. Autonomous Pathways . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . VII. Future Prospects. . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . .
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I. Introduction The continual DNA damage that occurs from physiological and environmental agents is a threat to the integrity of the genome, viability of the cell, and survival of the organism. Therefore, cells contain a variety of DNA repair mechanisms (discussed in this book) for self-preservation and self-protection. A large body of evidence has amply demonstrated the importance of DNA repair to life. For example, certain base excision repair and double-stranded-break repair functions are required for viability in mice, whereas deficiencies in nucleotide excision repair or 101 ADVANCES IN PROTEIN CHEMISTRY, Vol. 69
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mismatch repair cause developmental defects and predispose mice and men to cancer (Friedberg and Meira, 2003; see also relevant chapters in this book). At steady states, DNA repair mechanisms are able to counter continual damage, protecting an organism from the consequence of an aberrant cell and protecting cells from the consequence of an aberrant genome. During the lifetime of a cell, circumstances may arise that cause excessive DNA damage, calling for an increase in the capacity of repair. These are the circumstances under which the cellular responses to DNA damage have been studied. Experimentally, the investigation of DNA damage responses (i.e., activation of DNA repair, inhibition of cell cycle progression, and induction of apoptosis), has been conducted in eukaryotic cells under conditions of genotoxic stress. The perturbations employed in these experimental protocols reveal how cells respond to an acute increase in the level of DNA lesions. Therefore, DNA damage responses described in the literature have mostly been observed following exposure to nonphysiologic levels of ultraviolet (UV) radiation, ionizing radiation, oxidative stress, DNA modifying chemicals, and drugs that inhibit DNA metabolic enzymes. Interestingly, a number of these agents, including ionizing radiation, DNA modifying chemicals (alkylating or crosslinking agents), and drugs that inhibit topoisomerases or DNA polymerases have been applied empirically in cancer therapy with some—unpredictable—efficacy. Therefore, studies on how cells respond to genotoxic stress are of relevance to cancer therapy. Moreover, these stress responses are likely to reflect in part the normal workings of the cell to preserve and protect the genome under physiological levels of damage. Our current knowledge of cellular responses to genotoxic stress is largely derived from genetic studies in budding and fission yeast (Melo and Toczyski, 2002; Nyberg et al., 2002). These studies have identified a number of genes that are conserved in all eukaryotes to regulate cellular responses to DNA damage (see below). A separate body of work on human and mouse cell lines has identified additional genes not found in yeast, most notably the tumor suppressor, p53, that play critical roles in DNA damage responses. It is unclear whether all the players in the DNA damage response network have been identified. Of the genes that have already been identified, we know the functions of their products (see following). However, we have not completely ascertained the rules that govern the interplay among these gene products. It is typically assumed that a ‘‘generic’’ program is launched on genotoxic stress regardless of the type of lesion generated in the DNA. This assumption is based on the observations in yeast cells that mutations
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Table I Conserved Core Components in DNA Damage Signal Transduction Name
Biochemical activity
Rad17-RFC (s.c. RAD24-RFC) 9-1-1 (s.p. Rad9-Hus1-Rad1) (also known as RHR) (s.c. DDC1, MEC1, RAD17) ATR, ATM, DNA-PK (s.c. MEC1, TEL1; s.p. Rad3)
Loading of the 9-1-1 clamp onto damaged DNA Heterotrimeric DNA clamp May provide platform for signaling complex Protein kinase of the PI3-kinase superfamily, Phosphorylates target substrates at SQ or TQ motifs Protein–protein interactions for complex formation, and subnuclear localization Specific interaction domains include the BRCT domain and the FHA domain Protein kinase Protein kinase with FHA domain
BRCA1, p53-BP1, NBS1, MDC1, (s.c. RAD9; s.p. Crb2)
Chk1 (also in s.c. and s.p.) Chk2 (s.c. RAD53 and DUN1; s.p. Cds1)
Classification of putative role Damage sensor Damage sensor
Master switch
Adaptor/mediator
Effector Effector
s.c. for Saccharomyces cerivisiae; s.p. for Schizosaccharomyces pombe.
in genes such as RAD9 or MEC1 (see Table I) can affect responses to a variety of genotoxic agents. In fact, many of these ‘‘general response’’ genes are conserved from yeast to man (see Table I), further supporting the notion of a ‘‘generic’’ program. Although these findings point to a common core in the DNA damage response network, this common core is but one component of the overall program. It should be noted that different types of DNA lesions are recognized and repaired by different proteins and enzymes. In addition, genotoxic agents can cause harm to cellular components other than genomic DNA. Thus, the overall cellular responses to UV, ionizing radiation, or cisplatin are not necessarily identical. Nevertheless, the cumulative data do support a common theme in how cells react to genotoxic stress. The recurring theme appears to be a temporally coordinated activation of DNA repair and cell cycle checkpoints, followed by recovery, survival, or death. This chapter is focused on a general discussion of how these temporal events may be coordinated.
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II. Overview of Biological Responses to DNA Damage A. General Framework of DNA Damage Responses Because our current knowledge on DNA damage response has mostly been gained from studies of individual gene products, this body of knowledge has not permitted us to predict how a particular cell type would respond to a specific type of DNA-damaging agent. The current knowledge does indicate that the DNA damage-response network involves some versatile players that can act on different stages with specified contexts and cooperative strategies. Because it is not possible to deduce the DNA damage response network with fine details at this time, we take the approach of laying down a general framework that includes the various responses (Fig. 1). Construction of this framework is motivated by the consideration of two overall biological objectives. The first objective of DNA damage response is self-preservation, which is achieved by protecting the genome and allowing for recovery of damaged cells. The second objective is self-protection, which is achieved by eliminating damaged cells. In this framework, we distinguish the various biological responses to DNA damage by their reversibility. The transition of a biochemical reaction from a reversible state to an irreversible state allows for the temporal regulation of the DNA damage response (Fig. 1A).
B. Temporal Coordination of DNA Damage Responses Temporally, we divide the biological responses to DNA damage into three stages: the immediate early responses, which most likely occur without the requirement for new gene expression; the early responses; and the late responses, which are mostly irreversible (Fig. 1B).
1. Immediate-Early Responses To protect the genome and the damaged cell, DNA repair is activated and cell-cycle checkpoints initiated on DNA damage (Fig. 1B). A DNA-damage signaling mechanism, composed of protein components that are conserved from yeasts to mammals (see below), activates the immediate early responses (Fig. 1B). The G2/M checkpoint, which prevents the onset of mitosis, is installed through the phosphorylation/ inhibition of Cdc25C, a dual-specificity phosphatase that is required for the activation of MPF (mitosis-promoting factor); that is, the kinase complex of Cdc2/Cyclin B in mammalian cells (Fig. 2) (Zhou and Elledge,
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Fig. 1. A general framework for the temporal coordination of DNA damage responses. (A) Biological responses to DNA damage occur, temporally, from reversible to irreversible states. (B) Multiple outcomes are organized to depict immediate-early, early, and late responses to DNA damage, as indicated by shaded boxes (see text for discussion).
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Fig. 2. Initiation and prolongation of cell-cycle checkpoints in response to DNA damage. The G1/S and G2/M checkpoints are initiated by the phosphorylation of dual-specificity phosphatases Cdc25A and Cdc25C to inactivate their functions. Phosphorylation and inactivation of Cdc25A and Cdc25C are mediated by the Chk1 and/or Chk2 kinases, which are activated by DNA damage through the combined actions of sensors, PIKKs, adaptors/mediators. The G1/S and G2/M arrest responses can be prolonged by the accumulation of p21Cip1 or 14–3–3 sigma proteins, which result from the transcriptional upregulation of these genes through p53, which is stabilized and activated by the combined actions of the same proteins that initiate the checkpoints.
2000). A parallel checkpoint, which prevents the onset of DNA replication, is installed through the phosphorylation/degradation of Cdc25A, which dephosphorylates/activates the mammalian Cdk2/Cyclin E (Fig. 2) (Bartek and Lukas, 2003; Falck et al., 2002). Degradation of Cdc25A can prevent S-phase entry and the continued progression through S-phase in response to DNA damage (Bartek and Lukas, 2003).
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2. Early Responses Clonogenic survival occurs when damage is repaired and the cell cycle checkpoints are lifted. If the damage is severe, either because of repair defects or extreme destruction of the genome, death may occur within a short period of time as a result of a general failure of metabolism. This type of passive cell death may not be ‘‘regulated’’ by the damage response network, and thus, it is not included in this framework. If repair is slow, the ongoing repair causes the checkpoints to be prolonged. In mammalian cells, the G1/S and G2/M arrest responses offer two examples for how we distinguish between checkpoint initiation and prolongation. The G1/S arrest can be initiated immediately after DNA damage by biochemical mechanisms that block the firing of replication origins (Bartek and Lukas, 2003) (Fig. 2). The G1/S arrest can be prolonged by the up-regulation of p21Cip1, a heat-stable inhibitor of Cdk2/Cyclin-E (Fig. 2). The accumulation of p21Cip1 protein is driven by an increase in its mRNA, which is dependent on activation of p53 by DNA damage (Fig. 2). Transcription and translation of p21Cip1 require more time; therefore, we consider it to be an early response that prolongs rather than initiates the G1/S arrest (Fig. 2). The G2/M arrest can be initiated immediately after DNA damage by biochemical mechanisms that block the onset of mitosis (Zhou and Elledge, 2000) (Fig. 2). The G2/M arrest can be prolonged by the upregulation of 14–3–3-sigma, which is an adaptor protein that sequesters Cdc2/Cyclin B in the cytoplasm (Chan et al., 1999) (Fig. 2). The accumulation of 14–3–3-sigma protein is driven by an increase in its mRNA, which is dependent on p53 (Fig. 2). Therefore, we consider the transcriptional activation of 14–3–3-sigma to be an early response that prolongs rather than initiates the G2/M arrest (Fig. 2).
3. Late Responses to DNA Damage The late responses to DNA damage are irreversible. These irreversible outcomes include clonogenic survival, adaptation, and death (mitotic death, apoptotic death, mitotic catastrophe).
a. Clonogenic Survival vs. Mitotic Death/Premature Senescence The prolongation of cell cycle checkpoints does not have to be permanent. When damage is properly repaired, even with slow kinetics, the ‘‘checkpoint prolongation’’ mechanism reserves the option of allowing resumption of cell proliferation, leading to clonogenic survival. However, when the prolongation of cell cycle checkpoints may become permanent, the
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damaged cell irreversibly withdraws from proliferation. This permanent cell cycle arrest has been observed with fibroblasts exposed to high-dose ionizing radiation and is customarily referred to in the radiation biology literature as ‘‘mitotic death’’ (Hendry and West, 1997). Permanent withdrawal of damaged cells from the cell division cycle has also been referred to as ‘‘premature senescence’’ (Di Leonardo et al., 1994). Mechanistically, the shutdown of cell cycle machinery in mitotic death or premature senescence appears to involve the transcriptional repression of cell cycle genes (e.g., cyclin-B and Cdc2) and requires the RB-family of transcription corepressors (Badie et al., 2000; Taylor et al., 2001; Wang et al., 2001). It thus appears that the permanent cell cycle arrest involves a program that is triggered by DNA damage but that has a built-in temporal mechanism to delay its execution.
b. Apoptosis vs. Necrosis Genotoxic stress can lead to the activation of apoptosis, mediated by the release of proapoptotic factors from the mitochondria (Lassus et al., 2002; Wang, 2001). Transduction of DNA damage signal to mitochondria requires the tumor suppressor p53 (Fridman and Lowe, 2003) (see following). Because p53-dependent apoptosis involves the induction of transcription, the apoptotic response is delayed relative to the onset of checkpoints. Activation of apoptosis eliminates damaged cells and achieves the biological objective of protecting the organism. Apoptotic cell death also protects the organism from the inflammation that can be triggered by necrotic cell death. Of course, necrotic cell death resulting from metabolic failure in damaged cells also protects the organism. Therefore, apoptosis is but one of many modes of death that may result from genotoxic stress (Fig. 1B). The cellular decision to commit suicide is highly dependent on the cell context. The temporal regulation of apoptotic response to DNA damage is discussed further in later sections of this chapter. c. Adaptation Mitotic Catastrophe, and Genomic Instability In contrast to ‘‘clonogenic survival,’’ which describes recovery of a damaged cell with fully restored genome, ‘‘adaptation’’ describes the resumption of cell cycle despite the persistence of DNA damage. The phenomenon of adaptation to DNA damage has been observed in two yeast systems (Sandell and Zakian, 1993; Toczyski et al., 1997). In one system, a galactose-inducible endonuclease resulted in telomere loss; however, it was found that yeast cells continued to divide despite the persistence of damaged telomeres (Sandell and Zakian, 1993). In another system, an irreparable double-stranded break was created in the budding yeast using an inducible HO-endonuclease in a mutant RAD52 background (Toczyski
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et al., 1997). The double-stranded break triggered the G2/M checkpoint, but the break persisted because of the lack of RAD52 function. With time, an adaptation to the persisting double-stranded break was observed as yeast cells resumed mitosis despite this lesion (Toczyski et al., 1997). Adaptation response to DNA damage can lead to catastrophic death caused by mitotic failure (Fig. 1B). Adaptation response may also increase the probability of necrotic cell death (Fig. 1B). If the adapted cells escape mitotic catastrophe and resume cell proliferation, this would put the genomes of daughter cells at risk. Thus, adaptation is a DNA damage response that increases genomic instability and may precondition a cell for tumorigenesis (Galgoczy and Toczyski, 2001). Further investigation of adaptation to DNA damage in mammalian cells may yield important insights on the mutability associated with malignant cancer cells.
C. Other Comments on the General Framework We emphasize that the processes depicted in Figure 1 and discussed above are more dynamic than could be presented in a two-dimensional diagram. We would also emphasize that this framework is not a ‘‘molecular pathway’’—none of the arrows represent a specific biochemical mechanism. Rather, this framework temporally organizes the many cellular responses that have been observed after cells incur DNA damage. In particular, the framework highlights the importance of transitions from reversible to irreversible events and how the status of DNA repair may control these transitions. At the end of the chapter, we will discuss two alternative models for how DNA repair may coordinate the checkpoint and death responses to damage. Before those discussions, we will summarize briefly the current knowledge on several key players in the DNA damage response network.
III. Molecular Components for the Initiation of DNA Damage Responses As discussed above, the studies in yeasts have led to the identification of genes that play critical roles in initiating DNA damage responses (Nyberg et al., 2002). In the framework proposed in Figure 1, we would consider these as the ‘‘immediate-early response’’ genes, which are best understood at the molecular level. At present, the proteins and enzymes involved in activating the immediate-early responses are thought to interact in a ‘‘signal transduction pathway’’ that relates the occurrence of DNA lesions to repair proteins and to the cell-cycle engine. This signal transduction
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pathway has been reviewed extensively in recent years (Melo and Toczyski, 2002; Rouse and Jackson, 2002; Zhou and Elledge, 2000). Table I summarizes some of these molecules required to initiate DNA damage response, each of which is identified in yeast genetic studies. The molecular functions of these yeast genes are conserved in mammalian cells (Table I).
A. DNA Damage Sensor A hetero-trimeric DNA clamp known as the 9–1–1 complex is an important ‘‘sensor’’ of damaged DNA. The 9–1–1 complex resembles the homotrimeric PCNA in structure and possibly also in function (Venclovas and Thelen, 2000). The 9–1–1 complex derived its name from the fission yeast genes Rad9, Rad1, and Hus1, each encoding a subunit of this heterotrimer, and this complex is conserved in mammalian cells (Caspari et al., 2000; St Onge et al., 1999; Volkmer and Karnitz, 1999). The heterotrimeric 9–1–1 complex is loaded onto DNA at or near the damaged site by an alternative RFC complex containing the Rad17 protein as its largest subunit (Bermudez et al., 2003; Majka and Burgers, 2003). The biochemical mechanism of clamp loading by the Rad17-RFC is similar to the loading of PCNA by the regular RFC at the origin of replication. The Rad17-RFC is likely to recognize a structural cue resulting from the processing of DNA lesions for loading the 9–1–1 clamp (Kai et al., 2001; Zou et al., 2003). However, the precise biochemical mechanism for how Rad17-RFC ‘‘senses’’ damaged DNA to load the 9–1–1 complex requires further investigation. During DNA replication, the PCNA clamp functions as a processivity factor for DNA polymerases. PCNA also provides a platform for the recruitment of enzymes that complete the lagging strand synthesis from Okazaki fragments (Hosfield et al., 1998). When loaded at or near the damaged DNA site, the 9–1–1 clamp is thought to provide a platform for the assembly of a damage-signaling complex (Osborn et al., 2002; Zou et al., 2002). Whether the 9–1–1 clamp can also function as a processivity factor of repair polymerases is still an open question.
B. Master Switch: PIKK Family of Protein Kinases A family of protein kinases characterized by their large size (>200 kd) and a highly conserved C-terminal kinase domain relating the phosphotidylinositol-30 -kinase is found in all eukaryotic cells to regulate DNA damage response (Hoekstra, 1997; Shiloh, 2003). These protein kinases are commonly referred to as members of the PIKK family, which includes the budding yeast MEC1 and TEL1, the fission yeast Rad3, the mammalian ATR, ATM, and DNA-dependent protein kinase (DNA-PK). The PIKK
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family members have been reviewed extensively in recent years because they play essential roles in orchestrating cellular responses to DNA damage (Shiloh, 2003). Moreover, the mammalian PIKK-family members are involved in diseases; for example, ATM mutation in ataxia-telangiectasia, and DNA-PK defect in severe combined immune deficiency (SCID) (Hoekstra, 1997; Shiloh, 2003). In mammalian cells, the ATM kinase is activated by ionizing radiation and chromatin stress in vivo (Bakkenist and Kastan, 2003; Shiloh, 2003). The mammalian ATR kinase can interact with and be activated by UV-damaged DNA in vitro (Unsal-Kacmaz et al., 2002). A cofactor of ATR, ATRIP, can interact with a single-stranded DNA/RPA complex (Zou and Elledge, 2003). These observations support the conclusion of ATM and ATR kinases as transducers of the DNA damage signal. The current research efforts on the PIKK family members have been focused on the identification of their protein substrates. The PIKK members phosphorylate serine or threonine residues followed by a glutamine in the SQ or TQ motifs (Kastan and Lim, 2000). These simple motifs may not be sufficient to determine substrate specificity; other protein–protein interactions may also be required for these PIKK to select their substrates (Shiloh, 2003; Zou et al., 2002). To date, a large number of proteins of diverse functions have been found to be phosphorylated at SQ or TQ motifs following DNA damage in a PIKKdependent manner (Kastan and Lim, 2000; Shiloh, 2003). These include DNA replication and repair proteins such as the MRE11-complex (see below), transcription factors such as p53 in mammalian cells (see below), and chromatin components such as histone H2AX (Burma et al., 2001) and SMC1 (Kim et al., 2002). Importantly, PIKK members such as the yeast MEC1 and the mammalian ATM can phosphorylate proteins that are required for the initiation of DNA damage response, including the adaptors (e.g., RAD9, p53BP1), mediators (e.g., MDC1), and effector kinases (Chk1 and Chk2), thus underscoring their roles as master switches.
C. Adaptors and Mediators In mitogenic signal transduction, adaptor proteins play important roles in the assembly of signaling complexes. Generally speaking, adaptor proteins are composed of protein–protein interaction domains and function as scaffolds to promote signal-dependent formation of specific protein complexes. DNA damage appears to also induce the formation of protein complexes, which involve scaffolding proteins that have been called ‘‘adaptors’’ or ‘‘mediators.’’ Two specific protein–protein interaction domains, that is, the BRCT (breast cancer C-terminal) domain and the FHA (forkhead associated) domain, have been found in several adaptors/mediators
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of DNA damage signal transduction (Bork et al., 1997; Durocher and Jackson, 2002). The FHA domain preferentially binds peptides with a phospho-threonine epitope (Durocher et al., 2000). The BRCT domain can interact with another BRCT domain (Soulier and Lowndes, 1999) or with phosphorylated serine or threonine residues (Manke et al., 2003; Rodriguez et al., 2003; Yu et al., 2003). Of course, not all proteins with the BRCT or the FHA domains are directly involved in the initiation of immediate early responses to DNA damage (Bork et al., 1997). The FHA domain, in particular, is widely distributed in nature and is found in prokaryotic proteins that do not play any roles in DNA damage response (Durocher and Jackson, 2002). The budding yeast RAD9 gene product, which plays an essential role in checkpoint activation, is a prototypical adaptor in DNA damage signal transduction. The RAD9 protein contains a pair of BRCT domains at its C terminus. In the budding yeast, DNA damage activates the phosphorylation of RAD9 through MEC1 and TEL1, and this phosphorylation event is required for the activation of a downstream effector kinase RAD53 (Schwartz et al., 2002). Interestingly, there is not a mammalian protein with contiguous sequence homology to the budding yeast RAD9. (The mammalian Rad9 in the 9–1–1 complex is homologous to the fission yeast Rad9 and the budding yeast DDC1.) In the fission yeast, Crb2 has a similar function as RAD9. The fission yeast Crb2 protein contains a pair of BRCT domains at its C terminus, and Crb2 is required for DNA damage to activate a downstream effector kinase Chk1 (Mochida et al., 2004). A number of BRCT-containing proteins are found in mammalian cells (Bork et al., 1997). Among them, p53-BP1 (DiTullio et al., 2002; Wang et al., 2002), MDC1 (also known as Kiaa0170) (Goldberg et al., 2003; Stewart et al., 2003), BRCA1 (breast cancer associated 1) (Scully et al., 1997), and NBS1 (Nijme gen breakage syndrome 1, or nibrin) (Lim et al., 2000) are likely to play a RAD9-like role in transducing DNA damage signals. Each of these mammalian BRCT proteins is phosphorylated in response to DNA damage, and they contribute to the activation of DNA repair and cell-cycle checkpoints. These mammalian BRCT-proteins are likely to have distinct as well as redundant functions in DNA damage–signal transduction. In yeasts, the FHA domain is found in the downstream effector kinases RAD53 (budding yeast), DUN1 (budding yeast), and Cds1 (fission yeast), and in Xrs2 of the MRE11/RAD50/Xrs2 complex. In mammalian cells, the FHA domain is found in Chk2, in NBS1 of the MRE11/RAD50/NBS1 (MRN) complex, and in MDC1. The FHA domain of RAD53 interacts with phosphorylated RAD9, and such interaction is required for RAD53 activation by DNA damage (Sun et al., 1998). The FHA domain of Chk2 mediates phosphorylation-dependent oligomerization and activation of its
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kinase activity (Ahn et al., 2002; Xu et al., 2002). The FHA domain of NBS1 is required for relocalization of the MRN complex into nuclear foci in damaged cells (Cerosaletti and Concannon, 2003). However, the FHA domain of NBS1 appears to be dispensable for S-phase checkpoint activation, an event that requires the MRN complex (see following). The mammalian MDC1 and NBS1 proteins contain both the BRCT and the FHA domains. MDC1 was named a ‘‘mediator’’ to distinguish it from the BRCT-containing, RAD9-related ‘‘adaptors.’’ The MDC1 FHA domain and BRCT domain appear to interact with distinct protein partners (Xu and Stern, 2003); thus, MDC1 can function as a bridging protein to assemble multiprotein megacomplexes. With NBS1, specific partners for its FHA versus BRCT domains have not been described. Taken together, the current evidence supports the idea that BRCT and FHA domains engage in the formation of specific protein complexes in response to DNA damage.
D. Effector Kinases: Chk1 and Chk2 Members of the PIKK family are not the only protein kinases involved in DNA damage response. Two other protein kinases, Chk1 and Chk2, are also conserved through evolution and play important roles in regulating DNA damage responses (Bartek and Lukas, 2003).
1. Chk1 The Chk1 kinase was discovered in the fission yeast as a protein kinase that is activated by DNA damage to initiate the G2/M checkpoint (Walworth et al., 1993). The classical G2/M checkpoint pathway activated by DNA damage involves a Chk1-dependent phosphorylation of Cdc25 phosphatase, which is the universal activator of M-phase promoting factor (Cdc2/Cyclin B) (Zhou and Elledge, 2000). Phosphorylation of Cdc25 prevents it from dephosphorylating Cdc2, thereby preventing the activation of MPF (Zhou and Elledge, 2000). As a result, cells remain in the G2 phase of the cell cycle, allowing time for DNA repair before the onset of cell division. This Chk1-mediated G2/M checkpoint pathway is conserved in metazoa (Bartek and Lukas, 2003; Melo and Toczyski, 2002; Zhou and Elledge, 2000). Interestingly, the budding yeast Chk1 kinase does not target Cdc25 but, rather, phosphorylates Pds1 to activate a M-phase checkpoint in response to DNA damage (Liu et al., 2000b). Whether the budding yeast Chk1-mediated M-phase checkpoint pathway is also conserved in metazoan is currently unknown. The knockout of Chk1 causes early embryonic lethality (Liu et al., 2000a), indicating that the mammalian Chk1 kinase is required for viability. This essential function of Chk1 may reflect a continual dependence of
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mammalian cells on the G2/M checkpoint pathway; however, it cannot be ruled out that the mammalian Chk1 kinase may directly regulate cell survival and that its essential function might be unrelated to the regulation of Cdc25 phosphatase.
2. Chk2 The Chk2 kinase was identified in mammalian cells by virtue of its homology to the fission yeast Cds1; hence, Chk2 is also known as the hCds1 kinase (Bartek et al., 2001). The budding yeast RAD53 and DUN1 kinases are related to Cds1 and Chk1 in structure (Bartek et al., 2001). It should be noted that although Chk1 and Chk2 belong to the protein kinase superfamily, with the characteristic kinase domain consisting of an ATP-binding lobe, a substrate binding lobe, and an activation loop, they differ significantly outside of the kinase domain (Chen et al., 2000; Li et al., 2002). The Chk2 subfamily of protein kinases contains the FHA domain, with RAD53 containing two FHA domains (Bartek et al., 2001). As discussed above, one of the FHA domains of RAD53 interacts with phosphorylated RAD9 in response to DNA damage, and this interaction is required for the phosphorylation and activation of RAD53 by MEC1 (Schwartz et al., 2002; Sun et al., 1998). In the budding yeast, RAD53 is essential to the activation of DNA damage response (Melo and Toczyski, 2002; Zhou and Elledge, 2000). The fission yeast Cds1, by contrast, has redundant function to that of Chk1, in that both kinases can phosphorylate Cdc25 phosphatase (Melo and Toczyski, 2002). As discussed above, the FHA domain of mammalian Chk2 also mediates phosphorylationdependent protein–protein interaction and plays a critical role in the regulation of Chk2 kinase activity (Ahn et al., 2002; Xu et al., 2002). Unlike Chk1, the mammalian Chk2 knockout does not cause cell lethality, but compromises checkpoint and apoptotic responses to DNA damage and predisposes to tumor formation (Hirao et al., 2002; Jack et al., 2002; Takai et al., 2002). Interestingly, an inherited mutation of the CHK2 gene in humans is also associated with cancer predisposition (Bell et al., 1999). Taken together, the current results indicate that Chk1 and Chk2, although conserved through evolution, may be employed to serve redundant or distinctive roles in DNA damage signal transduction, depending on the organisms and cell types.
E. From Components to Mechanisms Descriptors such as ‘‘sensors,’’ ‘‘master switches,’’ ‘‘adaptors/mediators,’’ and ‘‘effector kinases’’ suggest an understanding of the mechanism by which DNA damage signal is transduced to regulate cellular processes.
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However, it should be emphasized that such a mechanism is only implied, rather than elucidated. The current understanding is based on genetic studies, which provide insights on the hierarchical relationship among these gene products but do not illuminate the biochemical nature of their interactions. Therefore, the various descriptors should not be taken literally as components of a linear pathway. It is prudent to conclude at this time that the mechanism of DNA damage–signaling transduction is not known. A recent study has examined the subnuclear location and phosphorylation of some of the signaling components, including the NBS1 and Chk2, during the first hour of genotoxic stress (Lukas et al., 2003). The results confirm that these signal transducers are activated within minutes of cellular exposure to genotoxic stress and describe the interesting spatial segregation of damage sensors versus effector kinases in the nucleus (Lukas et al., 2003). Exactly how these signaling molecules regulate the late responses (i.e., adaptation or cell death) to DNA damage will await further investigation.
IV. Apoptotic Effectors in DNA Damage Response The apoptotic response to DNA damage has been extensively investigated in mammalian cells because it involves the protein product encoded by the p53 gene, one of the most frequently mutated genes in sporadic human cancer. These studies have established that p53-dependent alteration in gene expression is required for DNA damage to activate the apoptosis machinery (Fridman and Lowe, 2003; Oren, 2003). Although p53 function is necessary for the induction of apoptosis by DNA damage, it is not sufficient. In addition, recent results indicate that p53 may activate apoptosis through posttranscriptional mechanisms (Chipuk et al., 2004; Mihara et al., 2003). The roles of p53 and several other signal transducers in DNA damage-induced apoptosis are briefly discussed.
A. Role of p53 in DNA Damage Response The p53 homo-tetramer is a transcription factor, with a defined DNA binding domain, oligomerization domain, and transactivation domain (Brooks and Gu, 2003; Oren, 2003). In response to DNA damage, p53 is rapidly phosphorylated by the PIKK, Chk1, and Chk2 kinases (Brooks and Gu, 2003; Giaccia and Kastan, 1998). Phosphorylation leads to the stabilization of the p53 protein and activation of its transcriptional function. A large number of p53-regulated genes have been identified, and they can be loosely classified into four groups: DNA repair (e.g., Rnr), cell cycle
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(e.g., p21Cip1, cyclin G, 14–3–3-sigma), redox regulation (the PIG genes), and apoptosis (Bax, Noxa, Puma) (Oren, 2003). Therefore, p53 is not a ‘‘dedicated’’ regulator of cell death; instead, p53 is a general regulator of the transcriptional response to DNA damage. The p53-dependent upregulation of p21Cip1, a heat-stable inhibitor of cyclin-dependent kinase (Cyclin E/Cdk2, CyclinA/Cdk2), is responsible for DNA damage–induced G1 arrest in mammalian cells. Up-regulation of p21Cip1 may also contribute to DNA damage–induced premature senescence (mitotic death). As discussed above, permanent withdrawal from the cell cycle induced by high-dose ionizing radiation (IR) or other genotoxic agents involves the transcription repression of Cdk and cyclins through tumor suppressor RB and related p107, p130 ‘‘pocket proteins’’ (Badie et al., 2000; Taylor et al., 2001; Wang et al., 2001). (The RB family of proteins each contains several peptide-binding sites, that is, pockets, to assemble transcription repression complexes at specific promoters; Chau and Wang, 2003). The p21Cip1 protein can inhibit the phosphorylation of RB, p107, and p130 by Cdk/cyclin; as a result, these pocket proteins are able to assemble transcription repression complexes and thus suppress the expression of cell cycle genes. Therefore, up-regulation of p21Cip1 not only prolongs the G1 checkpoint but may also help to establish premature senescence in collaboration with the RB family of transcription repressors. The p53-dependent up-regulation of Bax, Noxa, and Puma contributes to the induction of apoptosis because these proapoptotic Bc12 family members can cause mitochondrial permeability transition to release several apoptotic inducers including cytochrome c, AIF, Samc, and Omi (Wang, 2001). Cells deficient in p53 are highly resistant to DNA damage– induced apoptosis. However, p53-dependent up-regulation of Bax, Noxa, and Puma could be observed in damaged cells that do not undergo apoptosis (Takaoka et al., 2003). Thus, transcriptional activation of the Bax, Noxa, or Puma gene expression is necessary but probably not sufficient for DNA damage–induced apoptosis. Recent studies have suggested that a fraction of the p53 protein exits the nucleus and binds to the antiapoptotic Bc12 family members in the cytoplasm (Chipuk et al., 2004; Mihara et al., 2003). This interaction is proposed to also be important for altering the balance between the proapoptotic (Bax, Bak) and the antiapoptotic (BclxL, Bcl-2) proteins, thus allowing the disruption of mitochondrial integrity (Chipuk et al., 2004; Mihara et al., 2003). The conclusion that p53 can either cause cell-cycle arrest or apoptosis in response to DNA damage is without dispute. However, we have not completely elucidated the rules that govern the selection of these two cell fates (Oren, 2003). There is emerging evidence to indicate p53 can selectively bind to different promoters, depending on the status of its
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covalent modifications (phosphorylation and acetylation) (Brooks and Gu, 2003; Giaccia and Kastan, 1998; Oren, 2003). In another view, the choice between cell cycle arrest and apoptosis may not be made by p53 alone. Clearly, DNA damage signals activate other transcription factors or posttranscriptional regulatory mechanisms, which may also contribute to the decision between life without parole or death. This view is supported by the identification of proteins that are activated by DNA damage to cause p53-independent apoptosis. A few of these proteins are discussed below; however, we must emphasize that this discussion is by no means comprehensive.
B. p53-Related Proteins: p63 and p73 In the mammalian genome, p53 is one of three related genes that encode transcription factors with shared structures and functions. The other two genes are p63 and p73, each encoding several protein products through alternative promoter usage at the 50 end and alternative splicing at the 30 end (Levrero et al., 2000; Melino et al., 2002). Genotoxic stress induces the stabilization of p63 and p73 proteins and activates their transcriptional functions (Flores et al., 2002; Gong et al., 1999). Current evidence indicates that all three p53 family members contribute to DNA damage–induced apoptosis. In fact, p63/p73 double-knockout mouse cells are as defective as p53-knockout mouse cells in their apoptotic response to doxorubicin or ionizing radiation (Flores et al., 2002). With p63 or p73 single-knockout mouse cells, apoptosis to DNA damage is reduced but not abolished, indicating that these two proteins may have redundant functions in mediating apoptotic response to genotoxic stress (Flores et al., 2002). Recent studies have identified genes that are commonly regulated by the three members of the p53 family as well as genes that are uniquely regulated by p63 or p73 and not by p53 (Melino et al., 2002). The transcriptional program that specifies the commitment to apoptosis in damaged cells is therefore likely to be more complex than previously thought. Stabilization and activation of p73 by DNA damage require the Ab1 tyrosine kinase, which is activated by DNA damage (see following). The Chk1 kinase also phosphorylates p73 to activate its function (Gonzalez et al., 2003). At present, it is not known how DNA damage signals regulate the p63 protein. The p63 and p73 genes, unlike p53, are seldom mutated in sporadic human cancer (Melino et al., 2002). Because p73 can mediate a p53-independent apoptotic response to DNA damage, the expression of p73 in tumor cells is correlated with increased sensitivity to chemotherapeutic agents that cause genotoxic stress (Irwin et al., 2003). Interestingly,
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p73 single-knockout cells also show reduced apoptotic response to tumor necrosis factor-alpha (TNF-) (Chau et al., 2004). TNF-induced apoptosis does not require new gene expression; on the contrary, actinomycin D or cycloheximide, which inhibit transcription and translation, actually enhance the apoptotic response to TNF. Therefore, p73 may also activate the apoptosis machinery through mechanisms other than the activation of gene expression. The finding of p63/p73 and their role in apoptosis regulation has informed us that studies of all three members of the p53 family are required to solve the mechanisms of DNA damage–induced apoptosis.
C. Abl Tyrosine Kinase The Abl protein is ubiquitously expressed in a variety of mammalian cell types. This nonreceptor tyrosine kinase contains a number of functional domains that endow it with cytoplasmic and nuclear functions (Wang, 2004). Abl contains nuclear localization and nuclear export signals and distributes itself in both compartments in a dynamic equilibrium of import and export. In the cytoplasm, Abl responds to signals from growth factors and extracellular matrix to regulate F-actin (Woodring et al., 2003). In the nucleus, Abl responds to signals from DNA damage and activates apoptosis (Wang, 2000). Activation of nuclear Abl tyrosine kinase by ionizing radiation requires a functional ATM kinase (Baskaran et al., 1997). Activation of nuclear Abl tyrosine kinase by cisplatin requires the mismatch repair protein, MLH1 (Gong et al., 1999; Nehme et al., 1999). Thus, Abl can be considered a downstream effector kinase in DNA damage–signal transduction. Abl is not present in yeast cells. The Abl tyrosine kinase of lower eukaryotes, such as Caenorhabditis elegans and Drosophila, does not appear to have a nuclear function. Hence, participation of nuclear Abl in DNA damage response may be limited to mammalian cells. The activated nuclear Abl tyrosine kinase has been shown to phosphorylate RNA polymerase II at its C-terminal repeated domain (Baskaran et al., 1999), possibly to regulate gene expression following DNA damage. Nuclear Abl tyrosine kinase also phosphorylates p73 to stabilize the protein and activate its apoptotic function (Costanzo et al., 2002). In myoblasts, genotoxic stress activates nuclear Abl tyrosine kinase to phosphorylate and inhibit MyoD function, resulting in a temporary halt of differentiation on DNA damage (Puri et al., 2002). In addition, nuclear Abl tyrosine kinase has been implicated in the tyrosine phosphorylation of DNA-dependent protein kinase (DNA-PK), a PIKK family member involved in the nonhomologous end-joining repair of double-stranded breaks (Kharbanda et al., 1997). Abl has also been shown to phosphorylate BRCA1 (Foray et al.,
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2002), Rad51 (Yuan et al., 1998), Rad52 (Kitao and Yuan, 2002), and UV-DDB2 (Cong et al., 2002). The effects of these tyrosine phosphorylation events on DNA repair, cell-cycle checkpoints, or cell death are not understood. Interestingly, activation of nuclear Abl tyrosine kinase by DNA damage is dependent on cell adhesion (Truong et al., 2003). When fibroblasts are detached from extracellular matrix (ECM) and then exposed to genotoxic agents, nuclear Abl kinase is no longer activated (Truong et al., 2003). As a result, detached fibroblasts are more resistant to DNA damage–induced apoptosis than fibroblasts that are attached to ECM. This finding indicates that Abl can integrate adhesion and DNA damage signals to regulate apoptosis, and it implies that the delayed apoptotic response to DNA damage may involve signal inputs other than DNA lesions.
D. Stress-Activated Protein Kinases: JNK and p38 The stress-activated proteins kinases (SAPKs) are classical MAP kinases that are regulated by a kinase cascade, i.e., MAPKKK, MAPKK, and MAPK (Morrison and Davis, 2003; Weston and Davis, 2002). A large body of literature has accumulated on the kinase cascades that regulate SAPK members such as JNK and p38; discussion of that literature is beyond the scope of this chapter. It is sufficient to say that JNK and p38 are activated by a variety of physiological signals as well as metabolic stresses, including those that induce DNA damage (Weston and Davis, 2002). At steady state, SAPK members are mostly localized to the cytoplasm. On activation, SAPK members can translocate into the nucleus wherein they phosphorylate transcription factors to regulate gene expression. The UVmediated activation of JNK involves cytoplasmic events that can occur in enucleated cells, indicating that UV-induced DNA lesions are dispensable in JNK activation (Rosette and Karin, 1996). Clearly, genotoxic agents can damage other cellular components or generate reactive oxygen species to activate SAPK members without input from DNA lesions. However, this does not rule out the possibility that signals from DNA lesions might directly activate a minor fraction of the SAPK members in the nucleus. Activated SAPKs phosphorylate a number of transcription factors to regulate gene expression (Weston and Davis, 2002). Some of the SAPK members, notably p38, have also been shown to directly regulate cell cycle progression by phosphorylating the Cdc25 phosphatase (Bulavin et al., 2001). Other SAPK members, notably JNK, can directly phosphorylate Bc12 family members to cause mitochondria-dependent apoptosis (Lei and Davis, 2003). With mouse embryo fibroblasts (MEFs), JNK1 and JNK2 are required for UV to induce apoptosis. MEFs derived from
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Jnk1/2-double-knockout embryos do not undergo apoptosis following UV irradiation, despite the up-regulation of p53 (Tournier et al., 2000). The requirement for JNK1/2 in the apoptotic response to UV and other genotoxic agents indicates that the damage of other cellular components or oxidative stress may be important cofactors in determining the death response to DNA damage.
V. DNA Repair Proteins in Damage Signaling Given the notion that DNA damage signaling begins with the detection of DNA lesions, it is not difficult to imagine the involvement of repair proteins in this process because repair also begins with lesion detection. At least three different lesion detection proteins, identified through the study of DNA repair, have been linked to the regulation of DNA damage responses. They are briefly discussed here.
A. Mismatch Repair Proteins Mutations of mismatch repair proteins (i.e., MSH2, MSH6, MLH1, or PMS2) contribute to microsatellite instability and increased mutation rate in cancer cells. The MSH2/MSH6 heterodimer binds to mismatched bases and several other types of DNA lesions (e.g., bases with bulky adducts or platination; Bellacosa, 2001); thus, they can detect lesions. The MLH1/ PMS2 heterodimer binds to the MSH2/6 heterodimer. Analogous to the bacterial mismatch repair mechanisms, MLH1/PMS2 are likely to participate in the recruitment of repair enzymes. Generally speaking, cells with repair defects are more likely to die upon DNA damage. Therefore, it is counterintuitive to find that MMR-deficient cells are more resistant to DNA damage–induced apoptosis. Cancer cell lines or knockout mouse cells deficient in MSH2, MSH6, MLH1, or PMS2 have each been shown to exhibit defects in cell-cycle checkpoint or apoptotic responses, particularly to chemotherapeutic agents such as temozolomide and cisplatin (Bellacosa, 2001). There are two general models for how MMR proteins regulate cell-cycle checkpoints and apoptosis. The first is based on the concept of ‘‘futile repair’’; that is, mismatch repair attempts to correct lesions that cannot be repaired and that during this process generate double-stranded breaks, which then trigger checkpoint and apoptosis (Li, 1999). The second is based on the notion that mismatch repair proteins directly participate in signaling. When encountering mismatched bases during DNA replication, the MSH2/6-MLH1/PMS2 complex recruits repair enzymes to correct
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such replication mistakes. It is conceivable that on encountering bulky adducts or cross-linked bases, the MSH2/6-MLH1/PMS2 complex adopts a different conformation that attracts signaling proteins to activate checkpoints or apoptosis (Bellacosa, 2001). Two lines of evidence have supported the second model (i.e., mismatch repair proteins directly participate in DNA damage signaling). The first line of evidence comes from the observed interactions among mismatch repair proteins and components of the DNA damage–signaling pathway. For example, MSH2/6 and MLH1 can associate with ATR and ATM (Brown et al., 2003), two members of the mammalian PIKK family, and PMS2 can associate with p73 (Shimodaira et al., 2003), an inducer of apoptosis. The second line of evidence comes from the characterization of MSH2 mutant protein; in particular, G674A (Lin et al., 2004). Cells expressing this mutant MSH2 are defective in mismatch repair but proficient in apoptotic response to cisplatin (Lin et al., 2004). This result indicates that the repair function of MSH2 can be separated from its signaling function. Although these two lines of evidence indicate mismatch repair proteins to participate in DNA damage signaling, they certainly have not ruled out a role for futile repair in the induction of apoptosis.
B. UV-DDB Complex UV irradiation induces cyclobutane pyrimidine dimers and (6–4) pyrimidine–pyrimidone photoadducts in DNA, lesions that interfere with transcription and DNA replication. The nucleotide excision repair machinery (NER), composed of several lesion-recognition proteins, helicases, and nucleases, corrects UV-induced lesions. Among the components of the NER machinery is a UV-damage DNA binding protein (DDB) complex with two subunits: DDB1 and DDB2 (Keeney et al., 1993). Because DDB directly binds to UV-induced lesions, it can function as a specific sensor of UV damage. This sensor can recruit the NER machinery to facilitate repair (Wakasugi et al., 2002). Alternatively, this sensor may recruit a signaling complex to regulate the DNA damage response. Interestingly, cells derived from Ddb2-knockout mice are resistant to UV-induced apoptosis, exhibiting a delayed activation of caspase on UV-irradiation (Itoh et al., 2004). This phenotype is similar to the resistance of mismatch repair – deficient cells to cisplatin-induced apoptosis, as discussed above. At present, there is no evidence to indicate that DDB contributes to futile repair of UV lesions. Hence, DDB may not induce apoptosis through the creation of double-stranded breaks at UV lesions. The alternative mechanism, that DDB functions as a sensor of UV-induced lesions and directly participates
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in DNA damage signaling is implied, by the finding that Abl tyrosine kinase associates with DDB and can phosphorylate DDB2 (Cong et al., 2002). UV-induced apoptosis requires JNK activity as discussed above (Tournier et al., 2000). The nuclear Abl tyrosine kinase has been suggested to modulate JNK activity under conditions of genotoxic stress (Cross et al., 2000). Interaction between DDB and Abl might therefore play a role in controlling the apoptotic response to UV damage. A recent study has found that DDB associates with Cu14A-Roc1, a cofactor in polyubiquitination of selected substrates (Groisman et al., 2003). Whether the DDB– Cu14A complex participates in DNA damage signaling or the repair of UV lesions by the NER is not known at this time.
C. MRE11-RAD50 Complex The stable complex of MRE11 and RAD50 is found in prokaryotic and eukaryotic cells. The crystal structure of the RAD50 subunit has revealed a Zn-coordinated dimerization mechanism, which can join two molecules of RAD50 with their globular N-terminal domains potentially separated by 1200 A˚ (Hopfner et al., 2002). This structural design indicates that a dimeric MRE11-RAD50 complex may bring together two DNA molecules during nonhomologous end joining or homologous recombination reactions, thus explaining its essential role in the repair of DNA ends. Because the MRE11-RAD50 complex can recognize DNA ends, it can function as a specific sensor of broken DNA in the genome (Petrini and Stracker, 2003). Indeed, the MRE11 complex has been shown to be required for S-phase checkpoint activation in mammalian cells (Falck et al., 2002; Petrini, 2000). As discussed above, the mammalian MRE11-RAD50 complex (MRN) contains a third subunit NBS1, which is encoded by the gene that is mutated (although not completely lost) in the human genetic disorder Nijmegen breakage syndrome (Williams et al., 2002). In mice, the knockout of Mre11, Rad50, or Nbs1 causes early embryonic lethality, and each single knockout leads to the disappearance of the MRN complex (Luo et al., 1999; Williams et al., 2002). In human, hypomorphic mutation of MRE11 is associated with ATLD (Ataxia telangiectasia–like disease), and that of NBS1 is associated with Nimejin Breakage Syndrome, with pathological defects similar to Ataxia telangiectasia (Lee et al., 2003). As discussed above, Nbs1 contains BRCT and FHA domains that mediate the formation of protein complexes in response to double-stranded breaks. Recent studies have suggested that an intact MRN complex is required for ionizing radiation (IR) to activate the ATM kinase, one of the master switches in orchestrating cellular responses to IR (Uziel et al.,
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2003). Thus, the MRE11-RAD50-NBS1 complex appears to play multiple roles, including lesion detection, DNA repair, and signal transduction. The notion that lesion-binding proteins can function as ‘‘sensors’’ of DNA damage to activate downstream signal transduction pathways is gaining support but is far from proven. Further investigation is required to directly link Mismatch Repair, UV-DDB, MRN, or other repair proteins to the activation of checkpoints or apoptosis.
VI. Alternative Models for the Temporal Coordination of DNA Damage Responses We propose two alternative mechanistic models for the temporal regulation of cellular responses to DNA damage. We describe them as the ‘‘integrative surveillance’’ (IS) and the ‘‘autonomous pathway’’ (AP) models (Fig. 3). These two hypothetical models represent two opposing philosophical views, but they may not be mutually exclusive.
A. Integrative Surveillance The IS model proposes the existence of a ‘‘regulatory hub’’ that actively surveys the genome for lesions and for the status of repair to control the cell cycle and the apoptosis machineries (Fig. 3a). The ‘‘regulatory hub’’ continuously monitors the extent of damage and the progress of repair, integrates the information, and then inhibits cell cycle or activates apoptosis. In the IS model, cell fate decisions are made logically based on the rate of lesion accumulation/reduction. When lesions are increasing (i.e., the rate of lesion accumulation exceeds the rate of repair), the regulatory hub activates repair and cell-cycle checkpoints. When lesions are decreasing, (i.e., the rate of repair exceeds the rate of lesion accumulation), the regulatory hub maintains checkpoints and inhibits apoptosis. When lesions persist (i.e., the levels do not reduce for an extended time), the regulatory hub activates apoptosis. The proteins and enzymes discussed above in this chapter can conceivably function in a ‘‘regulatory hub.’’ Protein complexes such as the 9-1-1heterotrimeric clamp, the MSH2/MSH6 heterodimer, the DDB heterodimer, and the MRE11-complex can each function as a ‘‘sensor’’ of DNA lesions; thus, they are able to monitor the levels of damage in the genome. The PIKK master switch kinases can also directly or indirectly sense damaged DNA. The Chk1 and Chk2 kinases, which are activated by DNA lesions through the actions of damage sensors, PIKK members, and adaptors/mediators, directly regulate the cell cycle machinery. The PIKK members, Chk1 and Chk2, directly phosphorylate p53 to activate its
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Fig. 3. Alternative models for the temporal coordination of DNA damage responses. (A) The ‘‘integrative surveillance’’ (IS) model proposes that signals indicating levels of DNA damage and the progress of repair are integrated by a regulatory hub to either activate or inhibit cell-cycle checkpoints versus apoptosis. The temporal coordination of different biological outcomes is the result of deliberated decisions made by the regulatory hub. (B) The ‘‘autonomous pathway’’ (AP) model proposes that DNA
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apoptotic function. Therefore, it could be argued that these proteins function in an integrative surveillance mechanism. The current data do not explain how the damage sensors can monitor the ‘‘rate’’ of accumulation or reduction in the levels of lesions. The integrative surveillance model does not provide an explanation for adaptation, which is illogical because adaptation allows a damaged cell to resume proliferation. Although these questions remain, they have by no means ruled out the idea of a ‘‘regulatory hub’’ in coordinating DNA damage responses.
B. Autonomous Pathways The AP model proposes that the temporal regulation of cell-cycle checkpoints, adaptation, apoptosis, and mitotic death is not achieved by a logical ‘‘brainlike’’ mechanism but is the result of independent pathways with built-in feedback mechanisms to cause delayed outcomes autonomously (Fig. 3b). In the AP model, DNA damage signal activates repair, checkpoints, or cell death through distinct pathways, without the continuous input from a ‘‘regulatory hub.’’ The DNA repair status affects the checkpoint or the cell-death pathways solely by eliminating lesions and therefore extinguishing the signals that trigger these pathways. The current knowledge on DNA damage signaling is also compatible with the AP model. The sensors, master switch, and effector kinases respond to damaged DNA and transmit that information to the autonomous pathways. Together, these proteins send signals as long as lesions are present in the genome (as illustrated by the gear wheel in Fig. 3b). The signals are eliminated when lesions are removed by DNA repair. In the AP model, cell-cycle checkpoints can be reversed through the built-in feedback mechanism while the gear wheel is still running, thus providing an explanation for ‘‘adaptation’’ to DNA damage.
1. Autonomous Checkpoint Pathway In the AP model, the checkpoint pathway has a time-dependent negative feedback loop (Fig. 3b). Damaged DNA activates this pathway to install the checkpoints. This pathway then proceeds through a series of damage activates independent pathways with self-regulated feedback mechanisms. There is not a central hub to coordinate the biological outcomes. A time-delayed negative feedback loop in the checkpoint pathway can allow resumption of proliferation despite persistent damage. By contrast, the cell death pathway is immediately inactivated by a negative feedback loop. Only repeated activation of the death pathway can eventually lead to cell death, either by apoptosis or mitotic death. The current data can be accommodated by either model (see text for discussion).
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time-dependent steps, leading eventually to inactivation of the checkpoints. Progression of this negative feedback loop can follow two possible scenarios. In one, the feedback cycle can be set into motion by a DNA damage signal and then run autonomously to the end. In this scenario, cells recover from checkpoints at a predetermined time interval after genotoxic stress, regardless of the extent of damage. If the DNA lesions are eliminated within this predetermined time interval, cells recover and undergo ‘‘clonogenic survival.’’ If the lesions are not completely eliminated within this time period, cells undergo ‘‘adaptation’’ with damage and are at risk for mitotic catastrophe, necrosis, or genome instability. In an alternative scenario, progression through the negative feedback loop is designed to require continual input of the DNA damage signal. If the damage signal is eliminated, the feedback cycle is aborted, and cells resume proliferation to undergo ‘‘clonogenic survival.’’ If DNA damage signal persists long enough to allow completion of the entire feedback cycle, cells undergo ‘‘adaptation.’’
2. Autonomous Death Pathway In the AP model, cell death (either by apoptosis or through mitotic death) is regulated by a series of feedback loops, which must be activated multiple times to achieve death (Fig. 3b). The cell-death pathway is activated immediately on DNA damage, but to no avail, because of a built-in feedback inhibition (illustrated as the small negative feedback loops in the diagram, Fig. 3b). At the completion of the first feedback loop, the cell death pathway can be activated again if damage signal persists. The repeated cycles of activation and inactivation can proceed as long as the damage signal is present. The negative feedback mechanism may be weakened through the successive rounds of activation. When the feedback mechanism eventually fails, cell death occurs. The choice between apoptosis or mitotic death is determined by other factors relating to the overall cellular context, but not through a ‘‘logical’’ evaluation of the rate of lesion accumulation or the rate of DNA repair.
3. Negative Feedback Regulation of p53 DNA damage–induced death, either through mitotic death or apoptosis, requires p53 (Oren, 2003). The AP model would indicate p53 to be regulated by an intrinsic negative feedback loop (Fig. 3b). Indeed, such a feedback loop is well established, and it involves Mdm2, the inhibitor of p53 (Michael and Oren, 2003). In nonstressed cells, p53 levels are kept low through an Mdm2-mediated polyubiquitination and proteosome-dependent degradation (Michael and Oren, 2003). Moreover, Mdm2 directly binds to p53 and inhibits its transactivation function (Michael
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and Oren, 2003). DNA damage leads to the phosphorylation of p53 and Mdm2, disrupting their interaction, allowing p53 to accumulate and to activate the transcription of Mdm2 (Shiloh, 2003). This feedback loop has been described at the level of single cells (Lahav et al., 2004). DNA damage induces a wave of p53: its up-regulation occurs rapidly after ionizing radiation, followed approximately 5–6 h later by its down-regulation correlating with the up-regulation of Mdm2 (Lahav et al., 2004). The single cell–based study has revealed a previously unknown property of the p53 self-regulatory loop; that is, the peak height and the duration of this p53 wave is not affected by the dose of ionizing radiation (Lahav et al., 2004). Instead, IR dose affects the number of p53 waves a cell can generate. A single p53 wave is observed following 2 Gy of IR, a dose that rarely activates apoptosis or mitotic death in cultured mammalian cells. Following 10 Gy of IR, a dose that activates apoptosis or mitotic death, two or more waves of p53 are observed (Lahav et al., 2004). These observations indicate that more than one wave of p53 activation, through the persistence of DNA lesions, may be required to issue the cell death command. The repeated activation of p53 may eventually wear down the autoinhibitory loop to allow the accumulation of sufficient p53 to execute cell death. Alternatively, the repeated activation of p53 may execute a multilayered genetic program to trigger cell death without disrupting the autoinhibitory loop. Future studies at the level of single cells will provide insights on the temporal regulation of p53 and p53-dependent cell death response to DNA damage.
VII. Future Prospects Three objectives should propel the field of DNA damage response forward. The first is to continue the identification of genes involved in this process. The second is to increase our understanding of the biochemical functions of the known gene products. The third is to elucidate the mechanism of DNA damage signal transduction. These objectives can be achieved with the conventional approaches already in practice. However, new technologies and high-throughput methods will be required to provide a more coherent description of the DNA damage response program. A systematic comparison of responses to different genotoxic agents in one cell context will be helpful. These studies should include not only biological recordings of cell viability, cell cycle profiles, and apoptotic phenotypes but also molecular recordings of DNA lesions, the activity of repair enzymes, protein kinases, and the extent of phosphorylation of critical substrates. Examination of gene expression alteration as a function of genotoxic dose and throughout a biologically relevant time course
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needs to be performed so that we can have a better understanding of the entire protective program, not simply as a linear signal transduction pathway but as a multilayered network. Many of these measurements may have to be performed at the level of single cells, because recording the averaged response of an entire population may miss important feedback loops that do not operate synchronously. Clearly, the field faces many new challenges, which when met, will pay dividends that are worthy of the investment.
Acknowledgments We are supported by grants from the National Institutes of Health to J.Y.J.W. and a postdoctoral fellowship from the Lady Tata Foundation to S.K.C. J.Y.J.W. is the Herbert Stern Professor of Biology at the University of California, San Diego. We wish to thank members of the Wang laboratory for critical comments, and La Jolla Scientific Management (
[email protected]) for the graphic work.
References Ahn, J. Y., Li, X., Davis, H. L., and Canman, C. E. (2002). Phosphorylation of threonine 68 promotes oligomerization and autophosphorylation of the Chk2 protein kinase via the forkhead-associated domain. J. Biol. Chem. 277, 19389–19395. Badie, C., Itzhaki, J. E., Sullivan, M. J., Carpenter, A. J., and Porter, A. C. (2000). Repression of CDK1 and other genes with CDE and CHR promoter elements during DNA damage-induced G(2)/M arrest in human cells. Mol. Cell. Biol. 20, 2358–2366. Bakkenist, C. J., and Kastan, M. B. (2003). DNA damage activates ATM through intermolecular autophosphorylation and dimer dissociation. Nature 421, 499–506. Bartek, J., Falck, J., and Lukas, J. (2001). CHK2 kinase–a busy messenger. Nat. Rev. Mol. Cell Biol. 2, 877–886. Bartek, J., and Lukas, J. (2003). Chk1 and Chk2 kinases in checkpoint control and cancer. Cancer Cell 3, 421–429. Baskaran, R., Escobar, S. R., and Wang, J. Y. (1999). Nuclear c-Abl is a COOH-terminal repeated domain (CTD)-tyrosine (CTD)-tyrosine kinase-specific for the mammalian RNA polymerase II: Possible role in transcription elongation. Cell Growth Differ. 10, 387–396. Baskaran, R., Wood, L. D., Whitaker, L. L., Canman, C. E., Morgan, S. E., Xu, Y., Barlow, C., Baltimore, D., Wynshaw-Boris, A., Kastan, M. B., and Wang, J. Y. (1997). Ataxia telangiectasia mutant protein activates c-Abl tyrosine kinase in response to ionizing radiation. Nature 387, 516–519. Bell, D. W., Varley, J. M., Szydlo, T. E., Kang, D. H., Wahrer, D. C., Shannon, K. E., Lubratovich, M., Verselis, S. J., Isselbacher, K. J., Fraumeni, J. F., Birch, J. M., Li, F. P., Garber, J. E., and Habur, D. A. (1999). Heterozygous germ line hCHK2 mutations in Li-Fraumeni syndrome. Science 286, 2528–2531. Bellacosa, A. (2001). Functional interactions and signaling properties of mammalian DNA mismatch repair proteins. Cell Death Differ. 8, 1076–1092.
RESPONSES TO DNA DAMAGE
129
Bermudez, V. P., Lindsey-Boltz, L. A., Cesare, A. J., Maniwa, Y., Griffith, J. D., Hurwitz, J., and Sancar, A. (2003). Loading of the human 9-1-1 checkpoint complex onto DNA by the checkpoint clamp loader hRad17-replication factor C complex in vitro. Proc. Natl. Acad. Sci. USA 100, 1633–1638. Bork, P., Hofmann, K., Bucher, P., Neuwald, A. F., Altschul, S. F., and Koonin, E. V. (1997). A superfamily of conserved domains in DNA damage-responsive cell cycle checkpoint proteins. FASEB J. 11, 68–76. Brooks, C. L., and Gu, W. (2003). Ubiquitination, phosphorylation and acetylation: the molecular basis for p53 regulation. Curr. Opin. Cell Biol. 15, 164–171. Brown, K. D., Rathi, A., Kamath, R., Beardsley, D. I., Zhan, Q., Mannino, J. L., and Baskaran, R. (2003). The mismatch repair system is required for S-phase checkpoint activation. Nat. Genet. 33, 80–84. Bulavin, D. V., Higashimoto, Y., Popoff, I. J., Gaarde, W. A., Basrur, V., Potapova, O., Appella, E., and Fornace, A. J., Jr. (2001). Initiation of a G2/M checkpoint after ultraviolet radiation requires p38 kinase. Nature 411, 102–107. Burma, S., Chen, B. P., Murphy, M., Kurimasa, A., and Chen, D. J. (2001). ATM phosphorylates histone H2AX in response to DNA double-strand breaks. J. Biol. Chem. 276, 42462–42467. Caspari, T., Dahlen, M., Kanter-Smoler, G., Lindsay, H. D., Hofmann, K., Papadimitriou, K., Sunnerhagen, P., and Carr, A. M. (2000). Characterization of Schizosaccharomyces pombe Hus1: a PCNA-related protein that associates with Rad1 and Rad9. Mol. Cell. Biol. 20, 1254–1262. Cerosaletti, K. M., and Concannon, P. (2003). Nibrin forkhead-associated domain and breast cancer C-terminal domain are both required for nuclear focus formation and phosphorylation. J. Biol. Chem. 278, 21944–21951. Chan, T. A., Hermeking, H., Lengauer, C., Kinzler, K. W., and Vogelstein, B. (1999). 14-3-3Sigma is required to prevent mitotic catastrophe after DNA damage. Nature 401, 616–620. Chau, B. N., and Wang, J. Y. (2003). Coordinated regulation of life and death by RB. Nat. Rev. Cancer 3, 130–138. Chau, B. N., Chen, T. T., Wan, Y. Y., DeGregor, J., and Wang, J. Y. (2004). Mol. Cell. Biol. 24, 4438–4477. Chen, P., Luo, C., Deng, Y., Ryan, K., Register, J., Margosiak, S., Tempczyk-Russell, A., Nguyen, B., Myers, P., Lundgren, K., Kan, C. C., and O’Connor, P. M. (2000). The 1.7 A crystal structure of human cell cycle checkpoint kinase Chk1: Implications for Chk1 regulation. Cell 100, 681–692. Chipuk, J. E., Kuwana, T., Bouchier-Hayes, L., Droin, N. M., Newmeyer, D. D., Schuler, M., and Green, D. R. (2004). Direct activation of Bax by p53 mediates mitochondrial membrane permeabilization and apoptosis. Science 303, 1010–1014. Cong, F., Tang, J., Hwang, B. J., Vuong, B. Q., Chu, G., and Goff, S. P. (2002). Interaction between UV-damaged DNA binding activity proteins and the c-Abl tyrosine kinase. J. Biol. Chem. 277, 34870–34878. Costanzo, A., Merlo, P., Pediconi, N., Fulco, M., Sartorelli, V., Cole, P. A., Fontemaggi, G., Fanciulli, M., Schiltz, L., Blandino, G., Balsano, C., and Levrero, M. (2002). DNA damage-dependent acetylation of p73 dictates the selective activation of apoptotic target genes. Mol. Cell 9, 175–186. Cross, T. G., Scheel-Toellner, D., Henriquez, N. V., Deacon, E., Salmon, M., and Lord, J. M. (2000). Serine/threonine protein kinases and apoptosis. Exp. Cell Res. 256, 34–41.
130
WANG AND CHO
Di Leonardo, A., Linke, S. P., Clarkin, K., and Wahl, G. M. (1994). DNA damage triggers a prolonged p53-dependent G1 arrest and long-term induction of Cip1 in normal human fibroblasts. Genes Dev. 8, 2540–2551. DiTullio, R. A., Jr., Mochan, T. A., Venere, M., Bartkova, J., Sehested, M., Bartek, J., and Halazonetis, T. D. (2002). 53BP1 functions in an ATM-dependent checkpoint pathway that is constitutively activated in human cancer. Nat. Cell Biol. 4, 998–1002. Durocher, D., and Jackson, S. P. (2002). The FHA domain. FEBS Lett. 513, 58–66. Durocher, D., Taylor, I. A., Sarbassova, D., Haire, L. F., Westcott, S. L., Jackson, S. P., Smerdon, S. J., and Yaffe, M. B. (2000). The molecular basis of FHA domain:phosphopeptide binding specificity and implications for phospho-dependent signaling mechanisms. Mol. Cell 6, 1169–1182. Falck, J., Petrini, J. H., Williams, B. R., Lukas, J., and Bartek, J. (2002). The DNA damage-dependent intra-S phase checkpoint is regulated by parallel pathways. Nat. Genet. 30, 290–294. Flores, E. R., Tsai, K. Y., Crowley, D., Sengupta, S., Yang, A., McKeon, F., and Jacks, T. (2002). p63 and p73 are required for p53-dependent apoptosis in response to DNA damage. Nature 416, 560–564. Foray, N., Marot, D., Randrianarison, V., Venezia, N. D., Picard, D., Perricaudet, M., Favaudon, V., and Jeggo, P. (2002). Constitutive association of BRCA1 and c-Abl and its ATM-dependent disruption after irradiation. Mol. Cell. Biol. 22, 4020–4032. Fridman, J. S., and Lowe, S. W. (2003). Control of apoptosis by p53. Oncogene 22, 9030–9040. Friedberg, E. C., and Meira, L. B. (2003). Database of mouse strains carrying targeted mutations in genes affecting biological responses to DNA damage. Version 5. DNA Repair (Amst) 2, 501–530. Galgoczy, D. J., and Toczyski, D. P. (2001). Checkpoint adaptation precedes spontaneous and damage-induced genomic instability in yeast. Mol. Cell. Biol. 21, 1710–1718. Giaccia, A. J., and Kastan, M. B. (1998). The complexity of p53 modulation: Emerging patterns from divergent signals. Genes Dev. 12, 2973–2983. Goldberg, M., Stucki, M., Falck, J., D’Amours, D., Rahman, D., Pappin, D., Bartek, J., and Jackson, S. P. (2003). MDC1 is required for the intra-S-phase DNA damage checkpoint. Nature 421, 952–956. Gong, J. G., Costanzo, A., Yang, H. Q., Melino, G., Kaelin, W. G., Jr., Levrero, M., and Wang, J. Y. (1999). The tyrosine kinase c-Abl regulates p73 in apoptotic response to cisplatin-induced DNA damage. Nature 399, 806–809. Gonzalez, S., Prives, C., and Cordon-Cardo, C. (2003). p73alpha regulation by Chk1 in response to DNA damage. Mol. Cell. Biol. 23, 8161–8171. Groisman, R., Polanowska, J., Kuraoka, I., Sawada, J., Saijo, M., Drapkin, R., Kisselev, A. F., Tanaka, K., and Nakatani, Y. (2003). The ubiquitin ligase activity in the DDB2 and CSA complexes is differentially regulated by the COP9 signalosome in response to DNA damage. Cell 113, 357–367. Hendry, J. H., and West, C. M. (1997). Apoptosis and mitotic cell death: Their relative contributions to normal-tissue and tumour radiation response. Int. J. Radiat. Biol. 71, 709–719. Hirao, A., Cheung, A., Duncan, G., Girard, P. M., Elia, A. J., Wakeham, A., Okada, H., Sarkissian, T., Wong, J. A., Sakai, T., De Stanchina, E., Bristow, R. G., Suda, T., Lowe, S. W., Jeggo, P. A., Elledge, S. J., and Mak, T. W. (2002). Chk2 is a tumor suppressor that regulates apoptosis in both an ataxia telangiectasia mutated (ATM)-dependent and an ATM-independent manner. Mol. Cell. Biol. 22, 6521–6532.
RESPONSES TO DNA DAMAGE
131
Hoekstra, M. F. (1997). Responses to DNA damage and regulation of cell cycle checkpoints by the ATM protein kinase family. Curr. Opin. Genet. Dev. 7, 170–175. Hopfner, K. P., Craig, L., Moncalian, G., Zinkel, R. A., Usui, T., Owen, B. A., Karcher, A., Henderson, B., Bodmer, J. L., McMurray, C. T. et al. (2002). The Rad50 zinchook is a structure joining Mre11 complexes in DNA recombination and repair. Nature 418, 562–566. Hosfield, D. J., Mol, C. D., Shen, B., and Tainer, J. A. (1998). Structure of the DNA repair and replication endonuclease and exonuclease FEN-1: Coupling DNA and PCNA binding to FEN-1 activity. Cell 95, 135–146. Irwin, M. S., Kondo, K., Marin, M. C., Cheng, L. S., Hahn, W. C., and Kaelin, W. G., Jr. (2003). Chemosensitivity linked to p73 function. Cancer Cell 3, 403–410. Itoh, T., Cado, D., Kamide, R., and Linn, S. (2004). DDB2 gene disruption leads to skin tumors and resistance to apoptosis after exposure to ultraviolet light but not a chemical carcinogen. Proc. Natl. Acad. Sci. USA 101, 2052–2057. Jack, M. T., Woo, R. A., Hirao, A., Cheung, A., Mak, T. W., and Lee, P. W. (2002). Chk2 is dispensable for p53-mediated G1 arrest but is required for a latent p53-mediated apoptotic response. Proc. Natl. Acad. Sci. USA 99, 9825–9829. Kai, M., Tanaka, H., and Wang, T. S. (2001). Fission yeast Rad17 associates with chromatin in response to aberrant genomic structures. Mol. Cell. Biol. 21, 3289–3301. Kastan, M. B., and Lim, D. S. (2000). The many substrates and functions of ATM. Nat. Rev. Mol. Cell Biol. 1, 179–186. Keeney, S., Chang, G. J., and Linn, S. (1993). Characterization of a human DNA damage binding protein implicated in xeroderma pigmentosum E. J. Biol. Chem. 268, 21293–21300. Kharbanda, S., Pandey, P., Jin, S., Inoue, S., Bharti, A., Yuan, Z. M., Weichselbaum, R., Weaver, D., and Kufe, D. (1997). Functional interaction between DNA-PK and c-Abl in response to DNA damage. Nature 386, 732–735. Kim, S. T., Xu, B., and Kastan, M. B. (2002). Involvement of the cohesin protein, Smc1, in Atm-dependent and independent responses to DNA damage. Genes Dev. 16, 560–570. Kitao, H., and Yuan, Z. M. (2002). Regulation of ionizing radiation-induced Rad52 nuclear foci formation by c-Abl-mediated phosphorylation. J. Biol. Chem. 277, 48944–48948. Lahav, G., Rosenfeld, N., Sigal, A., Geva-Zatorsky, N., Levine, A. J., Elowitz, M. B., and Alon, U. (2004). Dynamics of the p53-Mdm2 feedback loop in individual cells. Nat. Genet. 36, 147–150. Lassus, P., Opitz-Araya, X., and Lazebnik, Y. (2002). Requirement for caspase-2 in stress-induced apoptosis before mitochondrial permeabilization. Science 297, 1352–1354. Lee, J. H., Ghirlando, R., Bhaskara, V., Hoffmeyer, M. R., Gu, J., and Paull, T. T. (2003). Regulation of Mre11/Rad50 by Nbs1: Effects on nucleotide-dependent DNA binding and association with ataxia-telangiectasia-like disorder mutant complexes. J. Biol. Chem. 278, 45171–45181. Lei, K., and Davis, R. J. (2003). JNK phosphorylation of Bim-related members of the Bcl2 family induces Bax-dependent apoptosis. Proc. Natl. Acad. Sci. USA 100, 2432–2437. Levrero, M., De Laurenzi, V., Costanzo, A., Gong, J., Wang, J. Y., and Melino, G. (2000). The p53/p63/p73 family of transcription factors: Overlapping and distinct functions. J. Cell Sci. 113, 1661–1670.
132
WANG AND CHO
Li, G. M. (1999). The role of mismatch repair in DNA damage-induced apoptosis. Oncol. Res. 11, 393–400. Li, J., Williams, B. L., Haire, L. F., Goldberg, M., Wilker, E., Durocher, D., Yaffe, M. B., Jackson, S. P., and Smerdon, S. J. (2002). Structural and functional versatility of the FHA domain in DNA-damage signaling by the tumor suppressor kinase Chk2. Mol. Cell 9, 1045–1054. Lim, D. S., Kim, S. T., Xu, B., Maser, R. S., Lin, J., Petrini, J. H., and Kastan, M. B. (2000). ATM phosphorylates p95/nbs1 in an S-phase checkpoint pathway. Nature 404, 613–617. Lin, D. P., Wang, Y., Scherer, S. J., Clark, A. B., Yang, K., Avdievich, E., Jin, B., Werling, U., Parris, T., Kurihara, N., Umar, A., Kucherlapati, R., Lipkin, M., Kunkel, T. A., and Edelmann, W. (2004). An Msh2 point mutation uncouples DNA mismatch repair and apoptosis. Cancer Res. 64, 517–522. Liu, Q., Guntuku, S., Cui, X. S., Matsuoka, S., Cortez, D., Tamai, K., Luo, G., Carattini-Rivera, S., DeMayo, F., Bradley, A., Donehower, L. A., and Elledge, S. J. (2000a). Chk1 is an essential kinase that is regulated by Atr and required for the G(2)/M DNA damage checkpoint. Genes Dev. 14, 1448–1459. Liu, Y., Vidanes, G., Lin, Y. C., Mori, S., and Siede, W. (2000b). Characterization of a Saccharomyces cerevisiae homologue of Schizosaccharomyces pombe Chk1 involved in DNA-damage-induced M-phase arrest. Mol. Gen. Genet. 262, 1132–1146. Lukas, C., Falck, J., Bartkova, J., Bartek, J., and Lukas, J. (2003). Distinct spatiotemporal dynamics of mammalian checkpoint regulators induced by DNA damage. Nat. Cell Biol. 5, 255–260. Luo, G., Yao, M. S., Bender, C. F., Mills, M., Bladl, A. R., Bradley, A., and Petrini, J. H. (1999). Disruption of mRad50 causes embryonic stem cell lethality, abnormal embryonic development, and sensitivity to ionizing radiation. Proc. Natl. Acad. Sci. USA 96, 7376–7381. Majka, J., and Burgers, P. M. (2003). Yeast Rad17/Mec3/Ddc1: A sliding clamp for the DNA damage checkpoint. Proc. Natl. Acad. Sci. USA 100, 2249–2254. Manke, I. A., Lowery, D. M., Nguyen, A., and Yaffe, M. B. (2003). BRCT repeats as phosphopeptide-binding modules involved in protein targeting. Science 302, 636–639. Melino, G., De Laurenzi, V., and Vousden, K. H. (2002). p73: Friend or foe in tumorigenesis. Nat. Rev. Cancer 2, 605–615. Melo, J., and Toczyski, D. (2002). A unified view of the DNA-damage checkpoint. Curr. Opin. Cell Biol. 14, 237–245. Michael, D., and Oren, M. (2003). The p53-Mdm2 module and the ubiquitin system. Semin. Cancer Biol. 13, 49–58. Mihara, M., Erster, S., Zaika, A., Petrenko, O., Chittenden, T., Pancoska, P., and Moll, U. M. (2003). p53 has a direct apoptogenic role at the mitochondria. Mol. Cell 11, 577–590. Mochida, S., Esashi, F., Aono, N., Tamai, K., O’Connell, M. J., and Yanagida, M. (2004). Regulation of checkpoint kinases through dynamic interaction with Crb2. EMBO J. 23, 418–428. Morrison, D. K., and Davis, R. J. (2003). Regulation of MAP kinase signaling modules by scaffold proteins in mammals. Annu. Rev. Cell Dev. Biol. 19, 91–118. Nehme, A., Baskaran, R., Nebel, S., Fink, D., Howell, S. B., Wang, J. Y., and Christen, R. D. (1999). Induction of JNK and c-Abl signalling by cisplatin and oxaliplatin in mismatch repair-proficient and -deficient cells. Br. J. Cancer 79, 1104–1110.
RESPONSES TO DNA DAMAGE
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Nyberg, K. A., Michelson, R. J., Putnam, C. W., and Weinert, T. A. (2002). Toward maintaining the genome: DNA damage and replication checkpoints. Annu. Rev. Genet. 36, 617–656. Oren, M. (2003). Decision making by p53: Life, death and cancer. Cell Death Differ. 10, 431–442. Osborn, A. J., Elledge, S. J., and Zou, L. (2002). Checking on the fork: The DNAreplication stress-response pathway. Trends Cell. Biol. 12, 509–516. Petrini, J. H. (2000). The Mre11 complex and ATM: Collaborating to navigate S phase. Curr. Opin. Cell Biol. 12, 293–296. Petrini, J. H., and Stracker, T. H. (2003). The cellular response to DNA double-strand breaks: Defining the sensors and mediators. Trends Cell Biol. 13, 458–462. Puri, P. L., Bhakta, K., Wood, L. D., Costanzo, A., Zhu, J., and Wang, J. Y. (2002). A myogenic differentiation checkpoint activated by genotoxic stress. Nat. Genet. 32, 585–593. Rodriguez, M., Yu, X., Chen, J., and Songyang, Z. (2003). Phosphopeptide binding specificities of BRCA1 COOH-terminal (BRCT) domains. J. Biol. Chem. 278, 52914–52918. Rosette, C., and Karin, M. (1996). Ultraviolet light and osmotic stress: Activation of the JNK cascade through multiple growth factor and cytokine receptors. Science 274, 1194–1197. Rouse, J., and Jackson, S. P. (2002). Interfaces between the detection, signaling, and repair of DNA damage. Science 297, 547–551. Sandell, L. L., and Zakian, V. A. (1993). Loss of a yeast telomere: Arrest, recovery, and chromosome loss. Cell 75, 729–739. Schwartz, M. F., Duong, J. K., Sun, Z., Morrow, J. S., Pradhan, D., and Stern, D. F. (2002). Rad9 phosphorylation sites couple Rad53 to the Saccharomyces cerevisiae DNA damage checkpoint. Mol. Cell 9, 1055–1065. Scully, R., Chen, J., Ochs, R. L., Keegan, K., Hoekstra, M., Feunteun, J., and Livingston, D. M. (1997). Dynamic changes of BRCA1 subnuclear location and phosphorylation state are initiated by DNA damage. Cell 90, 425–435. Shiloh, Y. (2003). ATM and related protein kinases: Safeguarding genome integrity. Nat. Rev. Cancer 3, 155–168. Shimodaira, H., Yoshioka-Yamashita, A., Kolodner, R. D., and Wang, J. Y. (2003). Interaction of mismatch repair protein PMS2 and the p53-related transcription factor p73 in apoptosis response to cisplatin. Proc. Natl. Acad. Sci. USA 100, 2420–2425. Soulier, J., and Lowndes, N. F. (1999). The BRCT domain of the S. cerevisiae checkpoint protein Rad9 mediates a Rad9-Rad9 interaction after DNA damage. Curr. Biol. 9, 551–554. St Onge, R. P., Udell, C. M., Casselman, R., and Davey, S. (1999). The human G2 checkpoint control protein hRAD9 is a nuclear phosphoprotein that forms complexes with hRAD1 and hHUS1. Mol. Biol. Cell 10, 1985–1995. Stewart, G. S., Wang, B., Bignell, C. R., Taylor, A. M., and Elledge, S. J. (2003). MDC1 is a mediator of the mammalian DNA damage checkpoint. Nature 421, 961–966. Sun, Z., Hsiao, J., Fay, D. S., and Stern, D. F. (1998). Rad53 FHA domain associated with phosphorylated Rad9 in the DNA damage checkpoint. Science 281, 272–274. Takai, H., Naka, K., Okada, Y., Watanabe, M., Harada, N., Saito, S., Anderson, C. W., Appella, E., Nakanishi, M., Suzuki, H., Nagashima, K., Sawa, H., Ikeda, K., and Motoyama, N. (2002). Chk2-deficient mice exhibit radioresistance and defective p53-mediated transcription. EMBO J. 21, 5195–5205.
134
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Takaoka, A., Hayakawa, S., Yanai, H., Stoiber, D., Negishi, H., Kikuchi, H., Sasaki, S., Imai, K., Shibue, T., Honda, K., and Taniguchi, T. (2003). Integration of interferon-alpha/beta signalling to p53 responses in tumour suppression and antiviral defence. Nature 424, 516–523. Taylor, W. R., Schonthal, A. H., Galante, J., and Stark, G. R. (2001). p130/E2F4 binds to and represses the cdc2 promoter in response to p53. J. Biol. Chem. 276, 1998–2006. Toczyski, D. P., Galgoczy, D. J., and Hartwell, L. H. (1997). CDC5 and CKII control adaptation to the yeast DNA damage checkpoint. Cell 90, 1097–1106. Tournier, C., Hess, P., Yang, D. D., Xu, J., Turner, T. K., Nimnual, A., Bar-Sagi, D., Jones, S. N., Flavell, R. A., and Davis, R. J. (2000). Requirement of JNK for stressinduced activation of the cytochrome c-mediated death pathway. Science 288, 870–874. Truong, T., Sun, G., Doorly, M., Wang, J. Y., and Schwartz, M. A. (2003). Modulation of DNA damage-induced apoptosis by cell adhesion is independently mediated by p53 and c-Abl. Proc. Natl. Acad. Sci. USA 100, 10281–10286. Unsal-Kacmaz, K., Makhov, A. M., Griffith, J. D., and Sancar, A. (2002). Preferential binding of ATR protein to UV-damaged DNA. Proc. Natl. Acad. Sci. USA 99, 6673–6678. Uziel, T., Lerenthal, Y., Moyal, L., Andegeko, Y., Mittelman, L., and Shiloh, Y. (2003). Requirement of the MRN complex for ATM activation by DNA damage. EMBO J. 22, 5612–5621. Venclovas, C., and Thelen, M. P. (2000). Structure-based predictions of Rad1, Rad9, Hus1 and Rad17 participation in sliding clamp and clamp-loading complexes. Nucleic Acids Res. 28, 2481–2493. Volkmer, E., and Karnitz, L. M. (1999). Human homologs of Schizosaccharomyces pombe rad1, hus1, and rad9 form a DNA damage-responsive protein complex. J. Biol. Chem. 274, 567–570. Wakasugi, M., Kawashima, A., Morioka, H., Linn, S., Sancar, A., Mori, T., Nikaido, O., and Matsunaga, T. (2002). DDB accumulates at DNA damage sites immediately after UV irradiation and directly stimulates nucleotide excision repair. J. Biol. Chem. 277, 1637–1640. Walworth, N., Davey, S., and Beach, D. (1993). Fission yeast chk1 protein kinase links the rad checkpoint pathway to cdc2. Nature 363, 368–371. Wang, B., Matsuoka, S., Carpenter, P. B., and Elledge, S. J. (2002). 53BP1, a mediator of the DNA damage checkpoint. Science 298, 1435–1438. Wang, J. Y. (2000). Regulation of cell death by the Abl tyrosine kinase. Oncogene 19, 5643–5650. Wang, J. Y. (2004). Controlling Abl: Auto-inhibition and co-inhibition? Nat. Cell Biol. 6, 3–7. Wang, J. Y., Naderi, S., and Chen, T. T. (2001). Role of retinoblastoma tumor suppressor protein in DNA damage response. Acta Oncol. 40, 689–695. Wang, X. (2001). The expanding role of mitochondria in apoptosis. Genes Dev. 15, 2922–2933. Weston, C. R., and Davis, R. J. (2002). The JNK signal transduction pathway. Curr. Opin. Genet. Dev. 12, 14–21. Williams, B. R., Mirzoeva, O. K., Morgan, W. F., Lin, J., Dunnick, W., and Petrini, J. H. (2002). A murine model of Nijmegen breakage syndrome. Curr. Biol. 12, 648–653.
RESPONSES TO DNA DAMAGE
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Woodring, P. J., Hunter, T., and Wang, J. Y. (2003). Regulation of F-actin-dependent processes by the Abl family of tyrosine kinases. J. Cell Sci. 116, 2613–2626. Xu, X., and Stern, D. F. (2003). NFBD1/MDC1 regulates ionizing radiation-induced focus formation by DNA checkpoint signaling and repair factors. FASEB J. 17, 1842–1848. Xu, X., Tsvetkov, L. M., and Stern, D. F. (2002). Chk2 activation and phosphorylationdependent oligomerization. Mol. Cell. Biol. 22, 4419–4432. Yu, X., Chini, C. C., He, M., Mer, G., and Chen, J. (2003). The BRCT domain is a phospho-protein binding domain. Science 302, 639–642. Yuan, Z. M., Huang, Y., Ishiko, T., Nakada, S., Utsugisawa, T., Kharbanda, S., Wang, R., Sung, P., Shinohara, A., Weichselbaum, R., and Kufe, D. (1998). Regulation of Rad51 function by c-Abl in response to DNA damage. J. Biol. Chem. 273, 3799–3802. Zhou, B. B., and Elledge, S. J. (2000). The DNA damage response: Putting checkpoints in perspective. Nature 408, 433–439. Zou, L., Cortez, D., and Elledge, S. J. (2002). Regulation of ATR substrate selection by Rad17-dependent loading of Rad9 complexes onto chromatin. Genes Dev. 16, 198–208. Zou, L., and Elledge, S. J. (2003). Sensing DNA damage through ATRIP recognition of RPA-ssDNA complexes. Science 300, 1542–1548. Zou, L., Liu, D., and Elledge, S. J. (2003). Replication protein A-mediated recruitment and activation of Rad17 complexes. Proc. Natl. Acad. Sci. USA 100, 13827–13832.
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FUNCTIONS OF DNA POLYMERASES By KATARZYNA BEBENEK AND THOMAS A. KUNKEL Laboratory of Molecular Genetics and Laboratory of Structural Biology, National Institute of Environmental Health Sciences, Research Triangle Park, North Carolina, 27709
I. II. III. IV.
V. VI. VII. VIII. IX. X. XI. XII. XIII. XIV.
DNA Polymerase Families. . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . Structures and Compositions of DNA Polymerases . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . Functions of DNA Polymerases . . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . Polymerases for DNA Repair . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Polymerases for Nucleotide Excision Repair . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. Polymerases for Base Excision Repair . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . C. Polymerases for Interstand Cross-Link Repair . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . D. Polymerases for Nonhomologous End-Joining of Double-Strand Breaks . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . Polymerases for Replicating Undamaged DNA. . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . Polymerases for Sister Chromatid Cohesion . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . Mitochondrial DNA Replication . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . Polymerases for Replicating Damaged DNA . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . Polymerases and Cell-Cycle Checkpoints . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . Polymerases for Replication Restart and Homologous Recombination . . .. . . . . . Polymerases for DNA Mismatch Repair. . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . Polymerases in the Development of the Immune System . . . . . . . . . . . . . . . . . . .. . . . . . Biological Consequences of Polymerase Dysfunction. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . Closing Comments . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . .
137 139 141 141 143 143 148 149 150 151 152 152 155 155 156 156 157 158 159
I. DNA Polymerase Families DNA polymerases are central players in DNA repair and replication, the processes that duplicate genomes and maintain their integrity to ensure faithful transmission of genetic information from one generation to the next. Our appreciation for the enormous complexity of repair and replication processes has grown significantly in the past few years with the discovery of a large number of DNA polymerases. Five polymerases are now recognized in Escherichia coli, nine in Saccharomyces cerevisiae, and 16 in humans (Table I) (Goodman, 2002; Hubscher et al., 2002; Shcherbakova et al., 2003). Based on differences in the primary structure of their catalytic subunits, DNA polymerases are classified into several distinct families (Figs. 1 and 2). Family A is named after the E. coli polA gene that encodes Pol I. Family A members also include the well-known bacteriophage T7 replicative polymerase and eukaryotic mitochondrial polymerase , as well 137 ADVANCES IN PROTEIN CHEMISTRY, Vol. 69
138
Table I DNA Polymerases in Escherichia coli, Saccharomyces cerivisiae, and Humans Family
Bacterial gene
Ec Pol I
(gamma) (theta) (nu) Ec Pol II (alpha) (delta) " (epsilon) (zeta) Ec Pol III
A
pol A
C
(beta)
X
(lambda) (mu) TdT (sigma) Ec Pol IV Ec PolV (eta) (iota) (kappa) Rev1 a
B
Y
Human gene
Yeast gene
Mol. Wt. (kDa)a
30 Exo
Other activities
103 140 290 100 89 165 125 225 353 130
þ þ þ þ þ (separate subunit)
50 Exonuclease dRPlyase ATPase, helicase
POLG POLQ POLN
MIP1
POLA POLD1 POLE POLZ (REV3)
POL1 (CDC17) POL3 (CDC3) POL2 REV3
POLB
39
POLL POLM TdT POLS (TRF4-1)
POL4 (POLX)
POLH (RAD30A, XPV) POLI (RAD30B) POLK (DINB) REV1
RAD30
66 55 56 60 40 46 78 80 76 138
polB
dnaE
TRF4
dinB umuC
Deduced from protein primary structure.
REV1
Primase
dRP lyase AP lyase dRP lyase, TdT TdT
dRP lyase
BEBENEK AND KUNKEL
Name
FUNCTIONS OF DNA POLYMERASES
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as two newly identified polymerases in human cells, Pol (Seki et al., 2003; Sharief et al., 1999) and Pol (Marini et al., 2003). Family B includes E. coli Pol II, the product of the polB gene, and homologous such as the replicative polymerases of bacteriophages T4 and RB69 and the eukaryotic polymerases , , ", and . Family C includes the E. coli replicative polymerase Pol III, whose catalytic subunit is encoded by the polC (dnaE) gene, and homologous polymerases present in most Gram-positive bacteria (Bruck et al., 2003; Dervyn et al., 2001). Family D (not shown) contains heterodimeric euryarchaea DNA polymerases (Pol II or PolD) (Cann and Ishino, 1999). Family X includes the well-characterized mammalian Pol ; more recently discovered eukaryotic polymerases , , and ; and a template-independent polymerase, terminal transferase (TdT). Members of the most recently named family Y include E. coli Pol IV and Pol V; eukaryotic polymerases , and ; and a template-dependent deoxycytidyl transferase, Rev1.
II. Structures and Compositions of DNA Polymerases X-ray crystallographic studies (Beard and Wilson, 2003, and many references therein) indicate that the catalytic subunits of polymerases in different families share three common subdomains, often called the fingers, palm, and thumb (Fig. 3; Ollis et al., 1985). These subdomains form a cleft whose bottom is formed by the palm, which harbors three catalytic residues (asterisks within red regions in Fig. 1) that coordinate with two divalent metal ions. All DNA polymerases are believed to use a common two-metal ion mechanism to catalyze the phosphoryl transfer reaction for nucleotide addition (Steitz et al., 1994). Beyond these basic features, polymerases are highly diverse, both between and even within families. Their catalytic subunits range from relatively small proteins like the 39-kDa human Pol to those as large as the 353-kDa human Pol (Table I, Fig. 1). They interact with a variety of different accessory proteins needed for repair or replication via noncatalytic domains and motifs, such as the BRCT domain (the C-terminal domain of BRCA1, the product encoded by the breast cancer susceptibility gene) or PCNA binding motifs located in C-terminal regions of Pol and . Polymerases can also have remarkably different polymerization properties. For example, the efficiency with which different polymerases insert correct nucleotides varies over an incredible 107-fold range (Beard et al., 2002), the number of nucleotides they incorporate per binding event varies from one to more than 10,000, and their fidelity varies by as much as 100,000-fold (Table II; Kunkel, 2004). Polymerase substrate preferences also vary, from preferential use of single-nucleotide gaps (Pol ) to preferential copying
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FUNCTIONS OF DNA POLYMERASES
141
of damaged DNA (Pol ; McCulloch et al., 2004), to coordinated synthesis of leading and lagging DNA strands by multiple polymerases acting as integral components of a complex multiprotein replication machine (McHenry, 2003). Some polymerases have additional enzymatic activities such as 30 ! 50 exonuclease (yellow regions in Fig. 1), 50 ! 30 exonuclease, 50 -deoxyribose phosphate (dRP) lyase, ATPase (Pol ), or primase (Pol ). These activities are located either in separate domains of the polymerase polypeptide (e.g., yellow and gray domains in Fig. 3A–C) or reside in separate but tightly associated subunits. Examples of the latter include the 30 ! 50 exonuclease activity in the subunit of E. coli Pol III and the primase activity of Pol (Figs. 1 and 2).
III. Functions of DNA Polymerases A complex network of DNA transactions must occur in cells to maintain the appropriate balance between accurately maintaining genetic information over many generations and permitting some diversity for the evolution of species, for the increased survival of microbes when subjected to changing environments, and for the development of a normal immune system. The physical and biochemical differences among polymerases imply that each protein has evolved to fulfill specific roles in maintaining this balance. In the following sections, we discuss the possible biological roles of various polymerases, as suggested by their diverse properties.
IV. Polymerases for DNA Repair We begin with repair transactions needed to provide clean substrates for the replication fork. Normal cellular metabolism and exposure of cells to exogenous genotoxicants produces DNA damage such as the loss or modification of bases, single-strand and double-strand breaks in DNA, and intra- and interstrand cross links. Multiple repair pathways exist to repair DNA damage, and the specific repair pathway employed partly depends on the type of damage experienced (Friedberg et al., 1995). A number of
Fig. 1. Modular organization of polymerases in different families. The modular organization of the five Escherichia coli and 16 human polymerases is shown with the number of amino acid residues in each polypeptide, as indicated. The polymerase domains are colored red, and other domains or functional motifs are color coded as indicated in the legend. A color gradient is used (see Pol III) to indicate that the domain boundaries are not defined. (See Color Insert.)
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these repair processes involve excision of damaged DNA followed by a resynthesis step that requires a polymerase. However, the substrates for DNA synthesis differ among the repair pathways, leading to different polymerase requirements (Fig. 4).
A. Polymerases for Nucleotide Excision Repair A variety of helix-distorting lesions, including ultraviolet (UV) radiation–induced damage and bulky chemical adducts, are removed by nucleotide excision repair (NER), which can occur by distinct subpathways. NER in prokaryotes and eukaryotes involves the same basic steps of recognizing the damage, unwinding the DNA duplex containing the damage, incision on both sides of the lesion to remove a damaged oligomer, resynthesis of DNA to fill the gap, and ligation. In E. coli, excision of the damage-containing oligonucleotide generates a 12–13nucleotide single-strand gap that is filled by Pol I. In eukaryotes, the gap size is 30 nucleotides, and there is substantial evidence that the gap is filled by one or both of the B-family polymerases, Pol or Pol ".
B. Polymerases for Base Excision Repair When bases are lost by depurination or depyrimidination; when bases are modified by alkylation, oxidation, or deamination; and when abnormal bases (e.g., dUTP, 8-oxo-dGTP) are incorporated into DNA, the resulting lesions are repaired by base excision repair (BER), which, like NER, can occur via distinct subpathways (see Chapter 1; also see Beard and
Fig. 2. Subunit composition of polymerases in different families. The subunit composition of E. coli and human polymerases is shown. The polymerase catalytic subunits are depicted in red, and the accessory subunits are in other colors. The abbreviations used are Kf, the large (Klenow) fragment of E. coli Pol I; T7, bacteriophage T7 DNA polymerase, Klentaq, the large fragment of Thermus aquaticus DNA polymerase I equivalent to Kf; Bs, Bacillus stearothermophilus DNA polymerase I large fragment; RB69, replicative polymerase from bacteriophage RB69; Tgo, Thermococcus gorgonarius DNA polymerase; 9 N-7, DNA polymerase from hyperthermo philic archaeon Thermococcus sp. 9 N-7; ASFV, African Swine Fever Virus family X DNA polymerase; Dpo4, Sulfolobus solfataricus P2 DNA polymerase IV; Dbh, catalytically active fragment of Sulfolobus solfataricus P1 DNA polymerase IV. 1Goldsby et al., 2001; 2Bemark et al., 2000; Esposito et al., 2000; Wittschieben et al., 2000; 3Gu et al., 1994; 4Bertocci et al., 2002; 5Bertocci et al., 2002; 2003; 6Gilfillan et al., 1993; 7McDonald et al., 2003; 8 Schenten et al., 2002. (See Color Insert.)
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Fig. 3. Structures of DNA polymerases in different families. The images of T7 Pol, Rb69 Pol, Pol , and Dpo4 were created based on their crystal structures in complex with duplex DNA using coordinates from the Protein Data Bank under the accession numbers 1T7P, 1IG9, 1BPY, and 1JX4 for T7 Pol, RB69 Pol, Pol , and Dpo4, respectively. In the image of T7 Pol, RB69 Pol, and Dpo4, the subdomains palm, fingers, and thumb of the polymerase domain are colored red, orange, and pink, respectively. In Pol , the fingers are pink and the thumb is orange. The 30 exonuclease domains (Exo 30 ) are colored yellow, and other domains are depicted in gray. The DNA template strand (T) is navy blue, and the primer strand (P) is blue. The position of the 30 hydroxyl of the primer terminus is indicated by 30 . (See Color Insert.)
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Table II Error Rates of Polymerases in Different Families Error rate 105
DNA polymerase
Exonuclease 30 ! 50
Family
Escherichia coli Pol III Escherichia coli Pol II Pol " Pol Kf(Pol I) Pol Pol Pol Pol Pol Dpo4 Pol Pol
Yes
C
0.6–1.2
0.025–1
Yes
B
0.2
0.1
Yes Yes Yes Yes No No No No No No No
B B A A B X X Y Y Y Y
1 1 0.8 1 16 67 90 580 650 3500 72,000 (TdGTP) 22 (misinsertion at A)
0.5 2 0.05 0.6 3 13 450 180 230 240 —
Substitution
1 deletions
Wilson, 2000; Bohr and Dianov, 1999; Lindahl et al., 1997). BER is initiated by a DNA glycosylase, which recognizes the damaged base and removes it by cleaving the N-glycosylic bond, leaving an apurinic/apyrimidinic (AP) site. An AP site generated by a monofunctional DNA glycosylase (one lacking an intrinsic AP lyase activity) is subsequently cleaved by an AP endonuclease to create a nick with a 30 -OH and 50 -dRP terminus. Alternatively, BER may be initiated by a bifunctional DNA glycosylase that also has intrinsic AP lyase activity. After it cleaves the phosphodiester bond 30 of the AP site, subsequent action of an AP endonuclease generates a singlenucleotide gap with 30 -OH and 50 -phosphate termini. DNA-damaging agents can also generate single-strand breaks that possess modified termini that need to be processed into substrates for polymerases or ligases (reviewed in Caldecott, 2003a). In E. coli, gap filling during BER is performed by Pol I. In mammals, BER can involve different DNA polymerases. In the major mammalian BER pathway, Pol inserts a single nucleotide onto the 30 -OH and then removes the 50 -dRP group, using its dRP lyase activity. The resulting nick is sealed by DNA ligase, completing the ‘‘short-patch’’ repair process. If the dRP is modified or not cleaved by the dRP lyase activity of Pol , strand displacement synthesis may generate a 2–13-nucleotide single-stranded
146 BEBENEK AND KUNKEL
Fig. 4. DNA polymerases involved in DNA repair and replication. See text for description. (See Color Insert.)
FUNCTIONS OF DNA POLYMERASES
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DNA flap that is removed by FEN1 flap endonuclease. Polymerases implicated in synthesis during ‘‘long-patch’’ BER include Pol , Pol , and Pol ". Which polymerase is actually used for any particular circumstance is the subject of active investigations. S. cerevisiae lacks Pol , such that the majority of BER in yeast may occur via the long-patch pathway, using Pol ", Pol , and Pol (Wang et al., 1993). For further details on BER, see Beard and Wilson (2000), Caldecott (2003b), Lindahl and Wood (1999), and Chapter 1 of this book. Like Pol , human Pol , Pol , and Pol also have a dRP lyase activity (Table I), indicating that they too might participate in repair processes that require removal of a dRP group. DNA Pol (Aoufouchi et al., 2000; Garcia-Diaz et al., 2000) is a member of the X family. Pol and Pol have similar domain organization and three-dimensional structures (GarciaDiaz et al., 2004), as well as several enzymatic properties in common. Like Pol and other family X members, Pol lacks an intrinsic 30 ! 50 exonuclease activity, and it has low processivity when extending a primer on a single-strand template. As for Pol , the processivity of Pol increases when filling short gaps with a phosphate on the 50 end, and Pol can substitute for Pol in reconstituted BER of uracil-containing DNA in vitro (Garcia-Diaz et al., 2001, 2002). Thus, Pol is a likely candidate for BER synthesis, perhaps being partially redundant with Pol or participating in specialized BER reactions involving a subset of damaged bases, specific cell or tissue types, or specific phases of the cell cycle. For example, the fact that Pol has a high affinity for dNTPs is consistent with the hypothesis that it might participate in BER or other repair processes under conditions in which the nucleotide triphosphate pools are low (e.g., in quiescent calls). In agreement with this, the expression of Pol is higher in cells undergoing S to M phase transition and in quiescent cells. As a member of the Y family, Pol has been implicated in the bypass of lesions that block DNA replication (see following). In addition, some properties of Pol are consistent with its possible role in specialized BER processes. Pol has low processivity and can fill 1–5-nucleotide gaps, and it can substitute for Pol to repair G–U and A–U pairs in DNA. In considering which lesions Pol might possibly repair, it is notable that Pol has a very unusual nucleotide incorporation specificity. Pol incorporates dTMP opposite template A much more efficiently than it forms the three other correct Watson–Crick base pairs, and its insertion fidelity opposite template A is relatively high (2 104), similar to that of Pol . These observations led us to suggest that Pol could participate in BER of uracil resulting from incorporation of dUMP during DNA replication (Bebenek et al., 2001). Consistent with this idea is the fact that Pol interacts with PCNA and colocalizes to replication foci (see Chapter 7).
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Pol also has the unprecedented ability to misinsert dGMP opposite a template T at a rate that exceeds that of correct dAMP incorporation. Furthermore, on templates that contain two or more consecutive Ts, preferential dGMP incorporation opposite the first T is followed by preferential incorporation of A opposite the second template T. This remarkable specificity led us to speculate that Pol may function in a specialized BER reaction, replacing dGs that are inadvertently removed by a DNA glycosylase from G–T or G–U mismatches that arise by deamination of 5-methyl-cytosine or cytosine (Bebenek et al., 2001). The fourth human polymerase with dRP lyase activity is DNA polymerase (Longley et al., 1998). As the only polymerase known to be present in mitochondria, Pol is thought to be responsible for all DNA synthesis in this organelle, including DNA replication and repair. Together with other proteins in mitochondria implicated in BER, Pol may participate in mitochondrial BER to remove damage resulting from reactive oxygen species generated during oxidative phosphorylation.
C. Polymerases for Interstrand Cross-Link Repair Interstrand cross links (ICLs) are highly cytotoxic lesions generated by agents such as nitrogen mustard, psoralen, diepoxybutane, and cisplatin. Repair of ICLs presents a special challenge, as both DNA strands are damaged and neither strand retains the correct genetic information. In E. coli, one repair pathway that has been well characterized (Van Houten, 1990) involves nucleotide excision repair, homologous recombination, and the action of UvrD DNA helicase and Pol I. Another E. coli pathway has been also described that depends on the function of NER and Pol II (Berardini et al., 1999). The two pathways do not seem to be functionally redundant, as cells deficient in either were hypersensitive to nitrogen mustard; however, the exact function of each pathway is not yet determined. Repair of ICLs in mammalian cells is not yet well defined. On the basis of studies showing that mutant alleles of some genes confer sensitivity to cross-link-inducing agents in model organisms, polymerases implicated in ICL repair include yeast Pol (see Chapter 6) and the Drosophila mus308 gene product. The latter has helicase motifs, and its C-terminal region contains polymerase motifs with homology to E. coli Pol I. Mutations in the mus308 gene cause hypersensitivity of cells to cross-linking agents and give rise to chromosomal aberrations in treated mutant cells consistent with a role of mus308 in repair of interstrand cross-links. Recently, two human polymerases, Pol (Seki et al., 2003; Sharief et al., 1999) and Pol (Marini et al., 2003), both members of family A, were identified by homology to the
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Drosophila mus308 protein, which indicates their possible involvement in ICL repair. Analysis of different mouse and human tissues indicated that the expression of both genes is highest in testis. The recombinant proteins showed polymerase activity on nicked double-strand DNA and on primed single-strand DNA. Pol also has an intrinsic ATP-ase activity and helicase motifs at the N terminus, although helicase activity has not been demonstrated. On the basis of primer extension assays in the presence of a single nucleotide, it was suggested that Pol incorporates G opposite template T more readily than other family A polymerases.
D. Polymerases for Nonhomologous End-Joining of Double-Strand Breaks During repair of double-strand breaks by nonhomologous end-joining (NHEJ, reviewed in Critchlow and Jackson, 1998; Lieber, 1999), broken DNA ends can be aligned using microhomology to create duplexes with short gaps that need to be filled by a DNA polymerase. Filling of short gaps during XRCC4-LigaseIV-dependent rejoining of double-strand breaks in HeLa cell extracts requires Pol , including its N-terminal BRCT domain (Lee et al., 2004). These and other observations (references in Lee et al., 2004) suggest that, in addition to a likely role in BER, Pol also participates in NHEJ in human cells. Genetic (Leem et al., 1994; Wilson and Lieber, 1999) and biochemical (Tseng and Tomkinson, 2002) studies indicate that yeast Pol IV, the homolog of human Pol , is also involved in NHEJ. Small gaps formed by the alignment of linear duplex DNA molecules are preferential substrates for yeast Pol IV; the protein interacts physically and functionally with Dn14/Lif1 complex, a core NHEJ factor; and this interaction is mediated by the BRCT domain of Pol IV. Another mammalian family X member, Pol , can extend primers containing up to four mismatches (Zhang et al., 2001) and even perform template-independent synthesis (Dominguez et al., 2000). Similar to Pol , Pol has a BRCT domain. It interacts with the Ku heterodimer, a major NHEJ factor, and it stably associates with DNA in the presence of Ku and XRCC4-ligaseIV. This complex can perform an end-joining reaction involving annealing of partially overlapping DNA ends and the filling of a single-nucleotide gap (Mahajan et al., 2002). Exposure of human cells to ionizing radiation results in increased Pol levels, and Pol localizes in nuclear foci containing double-strand breaks, indicating that it may also be involved in a NHEJ pathway. Pol also has the remarkable ability to efficiently incorporate ribonucleotides into DNA in vitro (Nick McElhinny and Ramsden, 2003; Ruiz et al., 2003). The significance of this activity is not yet well understood, but it has been suggested to permit activity under conditions of low dNTP pools.
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V. Polymerases for Replicating Undamaged DNA Efficient removal of DNA damage by the repair pathways mentioned above, combined with destruction of damaged dNTPs by enzymes (e.g., MutT, dUTPase) that sanitize dNTP pools (Ishibashi et al., 2003; Sekiguchi, 1996), provides undamaged substrates for the replication machinery. During chromosomal replication, the two antiparallel DNA strands are coordinately replicated (Kornberg and Baker, 1992; McHenry, 2003). Because polymerases only synthesize DNA in the 50 to 30 direction, one strand is replicated first as the leading strand, and the other is replicated slightly later as the lagging strand. Leading-strand replication is largely continuous, whereas lagging-strand synthesis is discontinuous and requires multiple cycles of RNA priming and DNA synthesis to generate Okazaki fragments, which are sealed after removal of the RNA. To perform efficient and accurate replication, replicative polymerases function with multiple accessory proteins (Fig. 2). In E. coli, the multisubunit Pol III holoenzyme is responsible for the bulk of synthesis on both DNA strands during chromosomal replication, whereas Pol I is involved in the processing of the Okazaki fragments. Pol III comprises a core complex containing the catalytic subunit, the subunit that has 30 ! 50 exonuclease activity for proofreading errors, and the subunit that stabilizes the subunit (Fig. 2). Association of the core polymerase with the clamp loader complex and the clamp forms the highly processive and accurate holoenzyme that functions as an asymmetric dimer for coordinated leading- and lagging-strand replication (Glover and McHenry, 2001; Kelman and O’Donnell, 1995). In most Gram-positive bacteria, two replicative polymerases are present, PolC and DnaE (Bruck and O’Donnell, 2000). PolC and DnaE have high sequence similarity to the subunit of E. coli Pol III. The subunit of PolC has an intrinsic 30 ! 50 proofreading exonuclease, whereas DnaE lacks an intrinsic exonuclease and appears to have relatively low fidelity (Bruck et al., 2003). PolC and DnaE function in a complex with the clamp and clamp loader, and it has been suggested that PolC replicates the leading strand while DnaE replicates the lagging strand (Bruck et al., 2003). In eukaryotic cells, replication of undamaged chromosomal DNA in the nucleus is performed by at least three DNA polymerases: Pol Pol , and Pol ". The catalytic subunit of Pol is not highly processive and lacks 30 ! 50 exonuclease activity for proofreading errors, but it does have a tightly associated primase activity for de novo synthesis of short RNA primers at replication origins and for initiation of Okazaki fragments on the lagging strand. Pol elongates these RNA primers to provide a short DNA primer, and then a switch occurs to allow the bulk of chain
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elongation by Pol or Pol ". Both Pol and Pol interact with the eukaryotic sliding clamp PCNA, and both enzymes can synthesize DNA processively. Both are accurate enzymes (Table II), because of the high nucleotide selectivity of the polymerase active site and proofreading by intrinsic 30 ! 50 exonuclease activities. Thus, both enzymes are well suited for replicating large eukaryotic genomes. Yeast strains with a deletion of the catalytic domain of Pol are inviable and are clearly defective in replication. A strain with a deletion of the N-terminal region encoding the polymerase activity of Pol is viable, albeit with severe growth and replication defects, so long as the C-terminal region is expressed (Kesti et al., 1999). Thus, another polymerase can partially substitute for Pol ". However, a strain with a deletion of the C-terminal, noncatalytic domain of Pol is inviable, indicating that this region is essential for some function other than polymerization per se (Dua et al., 1999). The exact contributions of Pol and Pol to leading and lagging strand replication remain an area of active investigation. The fact that they differ in primary structure (Fig. 1) and protein partnerships (Fig. 2) implies that they have distinct roles. Their functions may be differentiated for synthesis on opposite DNA strands (e.g., Pol for the lagging strand and Pol for the leading strand, or vice versa). Consistent with this hypothesis are data indicating that 30 ! 50 exonucleases associated with Pol and Pol can proofread replication errors on opposite DNA strands during replication (Shcherbakova and Pavlov, 1996). Alternatively, or in addition, Pol and Pol functions may be distinct for copying templates that differ by sequence, timing in S phase, or chromosomal region (e.g., euchromatin versus heterochromatin; see Fuss and Linn, 2002).
VI. Polymerases for Sister Chromatid Cohesion To ensure accurate segregation of chromosomes to daughter cells during mitosis, sister chromatids produced during replication are held together by cohesion complexes until their separation in anaphase. The product of the S. cerevisiae TRF4 gene, one of the proteins required for establishing sister chromatid cohesion in S phase, has been reported to be a family X DNA polymerase (Wang et al., 2000b). Originally designated as Pol , it has since been designated Pol and is suggested to perform DNA replication through the cohesion sites that could present an obstacle for the replicative polymerases and ". Interestingly, it was recently reported that the C-terminal domain of Pol interacts with Pol . This indicates that Pol might be involved in coupling DNA replication and sister chromatid cohesion. In addition to TRF4, eukaryotic genomes encode homologs of
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TRF4 (e.g., TRF5 in S. cerevisiae and humans and Cid genes in fission yeast). Cid1 was suggested to be a nucleotidyl transferase (Wang et al., 2000a), and Cid13 has been demonstrated to have poly(A) polymerase activity (Saitoh et al., 2002). In the latter study (Saitoh et al., 2002), the TRF4 gene product was also shown to have poly(A) polymerase activity, leading to the suggestion that an important function of these nucleotidyl transferases is to polyadenylate mRNA.
VII. Mitochondrial DNA Replication DNA polymerase is a member of family A and is the only cellular polymerase known to be present in mitochondria (Kaguni, 2004). Thus, it is believed to be responsible for replication of mitochondrial DNA, as well as for any repair that occurs in mitochondria (e.g., BER; see above). Human Pol is an accurate enzyme (Table II) and has an intrinsic 30 ! 50 proofreading exonuclease. It forms a tight complex with the p55 accessory subunit (Carrodeguas et al., 2001; Lim et al., 1999), which increases DNA binding affinity, stimulates the polymerase and exonuclease activities, and increases processivity.
VIII. Polymerases for Replicating Damaged DNA DNA repair systems are not perfect and leave some lesions in DNA. Moreover, some damage occurring during S phase may simply not be repaired quickly enough to avoid an encounter with the replication machinery. Many types of DNA damage, such as AP sites, UV photoproducts, and adducts generated by polycyclic aromatic hydrocarbons distort DNA helix geometry or alter base-coding potential. These lesions can stall normal replication conducted by the major replicative polymerases, which require correct base-pairing geometry for accurate and efficient replication (reviewed in Kunkel, 2004). To overcome the replication barrier posed by such lesions, cells harbor multiple polymerases capable of Translesion Synthesis (TLS). Most TLS polymerases belong to family Y, members of which are found in organisms from bacteria to humans (Ohmori et al., 2001). The properties of different family Y members are described in Chapters 6–9. Here we present a brief overview. Family Y polymerases typically (but not invariably) have relatively low processivity, low catalytic efficiency, and low fidelity when copying undamaged DNA templates (Table II). The low fidelity reflects their lack of 30 exonucleolytic proofreading activity and also the intrinsically low nucleotide selectivity of the polymerase active site. X-ray crystal structures (Friedberg et al., 2001; Ling et al., 2001; Silvian et al., 2001; Trincao et al.,
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2001; Zhou et al., 2001) indicate that the family Y polymerases may be able to accommodate lesions because they have unusually small fingers and thumb subdomains and because their active sites comprise smaller, uncharged side chains. They may be more flexible and their active sites more open and solvent accessible than polymerases in other families. In fact, the active site of Sso Dpo4, a family Y polymerase, can simultaneously accommodate two undamaged template nucleotides (Ling et al., 2001), a covalently linked cis-syn thymine–thymine dimer (Ling et al., 2003), or a bulky benzo[a]pyrene diol epoxide adduct (Ling et al., 2004). Family Y TLS polymerases can have very different properties, chief among them being which lesions are or are not bypassed. Some of these differences may depend on differences in an additional DNA binding domain distinct to family Y enzymes, the little finger domain (Fig. 3), also called the wrist or polymerase-associate domain. Data accumulated thus far indicate that, depending on the DNA polymerase, the type of lesion, and the local DNA sequence, translesion synthesis may either avoid or contribute to mutagenesis. Three E. coli polymerases are implicated in TLS, the family Y members Pol IV (DinB) and Pol V (UmuD2C), and the family B member, Pol II (Fig. 4). All three are induced as part of the SOS response to environmental stress, and all three modulate the ability of E. coli to survive during long periods in stationary phase (Goodman, 2002). Recent studies in E. coli with plasmids bearing different types of site-specific lesions show that all three polymerases are involved in TLS and can modulate lesion-dependent mutagenesis (Page´s and Fuchs, 2002). Human cells contain five TLS polymerases: Pols , Pol , Pol , Pol , and REV1 (Fig. 4). Among these, Pol is a member of family B and the others belong to the Y family. In addition, DNA Pol has been shown to perform TLS synthesis in vitro (Havener et al., 2003; Zhang et al., 2002), although there is as yet no evidence that this ability is related to its in vivo function. Of these six polymerases, only three are found in S. cerivisiae : Pol , Pol , and REV1. Bypass of some lesions may be conducted by one polymerase that can insert bases opposite the lesion and also extend the resulting primer terminus (Fig. 5A). Other lesions may require two TLS polymerases for bypass (Fig. 5B): one for insertion and another for extension, (e.g., Pol ; see Chapter 6). Thus, translesion synthesis likely requires multiple switches among polymerases and perhaps between polymerases and 30 exonucleases (e.g., intrinsic to Pol or Pol ") to allow proofreading of errors introduced by the TLS pols (Matsuda et al., 2000), thus ensuring efficient and accurate TLS. The mechanisms responsible for these enzymatic switches are under active investigation (e.g., see other chapters and also McCulloch et al., 2004; Pham et al., 2001a). Coordination of
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Fig. 5. Models for DNA polymerase switching during translesion synthesis. (A) Model for lesion bypass by a single TLS polymerase. (B) Model for lesion bypass by two TLS polymerases, wherein the first polymerase inserts a nucleotide opposite the damaged site and the second extends the aberrant primer terminus. (See Color Insert.)
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multiple polymerases during bypass likely involves the participation of several polymerase accessory proteins. These include REV1, which interacts with multiple TLS polymerases (Guo et al., 2003), and the E. coli clamp and eukaryotic PCNA, which have been shown to interact with most of the TLS polymerases (Fig. 3). In E. coli and S. cerevisiae, studies have show that bypass synthesis depends on this polymerase-clamp interaction (see Haracska et al., 2001; Page´s and Fuchs, 2002). Moreover, trafficking among multiple polymerases may be modulated by posttranslational modifications of clamp proteins. For example, recent work indicates that the protein–protein interactions of PCNA in replication and DNA repair can be differentially modulated by distinct DNA damage–induced ubiquitination and sumoylation of PCNA (Hoege et al., 2002; Ulrich, 2004) and see Chapter 10.
IX. Polymerases and Cell-Cycle Checkpoints Replication stalled by an elongation barrier such as a blocking lesion can initiate checkpoint responses in yeast and mammalian cells. Certain checkpoint responses depend on Pol (Navas et al., 1995, 1996), specifically the noncatalytic C-terminal residues of Pol (Fig. 1) that encode a putative zinc finger (Dua et al., 1998, 1999) and that interact with the MDM2 protein (Vlatkovic et al., 2000), thereby enhancing polymerase activity (Asahara et al., 2003). Defects in DNA damage checkpoints are observed with mutants in the polymerase catalytic subunit (D’Urso et al., 1995) and in the primase subunit of Pol (Marini et al., 1997), with a mutant that inactivates the 30 exonuclease activity of Pol (Datta et al., 2000), and with mutants in the fission yeast gene Cid1, a putative nucleotidyltransferase in the TRF4/Pol family (Wang et al., 2000a). It is also notable that one subunit of Pol shares significant homology and interacts with MAD2, a key protein involved in the spindle assembly checkpoint pathway (Murakumo et al., 2000). This checkpoint ensures that cells do not enter mitosis and that chromosome segregation does not occur until all chromosomes are properly attached to the mitotic spindle (reviewed recently in Musacchio and Hardwick, 2002).
X. Polymerases for Replication Restart and Homologous Recombination When replication forks are stalled, they may be restarted by a ‘‘fork regression’’ process, a rearrangement of DNA strands that allows the complementary undamaged daughter strand to temporarily act as a
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template for limited synthesis, followed by reestablishment of a normal fork that ultimately results in accurate lesion bypass (see Fig. 5 in review by Goodman, 2002). The DNA synthesis associated with this type of replication restart in E. coli is thought to be performed by Pol II (Pham et al., 2001b), and in eukaryotes it may be a major replicative polymerase (e.g., Pol "; see discussion in Asahara et al., 2003). Alternatively, stalled replication may lead to double-strand breaks that can be repaired by homologous recombination. The major replicative polymerases are likely to conduct the DNA synthesis associated with homologous recombination (Holmes and Haber, 1999; Jessberger et al., 1993). It has also been proposed that Pol functions in replication-associated repair in early S phase and also in replicating DNA in heterochromatin in late S phase (Fuss and Linn, 2002).
XI. Polymerases for DNA Mismatch Repair Polymerization errors that escape proofreading can be corrected by postreplicative DNA mismatch repair. This repair pathway recognizes base–base and addition/deletion mismatches. Repair involves excision of a region of the newly synthesized DNA strand containing the mismatch, followed by accurate resynthesis of DNA. In E. coli, the enzyme responsible for this DNA synthesis is Pol III. In human cells, resynthesis is performed by a replicative polymerase (e.g., Pol ; Longley et al., 1997). In addition, it has been suggested that exonucleases associated with Pol and Pol may be involved in the excision step of the mismatch repair process. Mismatches can also result from DNA damage, such as G–U and G–T mismatches generated by deamination of cytosine and 5-methyl-cytosine, respectively. In such cases, repair of the mismatch is initiated by a DNA glycosylase, and the gap is filled by a BER polymerase such as Pol .
XII. Polymerases in the Development of the Immune System The wide variety of immunoglobulins required for a full immune response in humans and mice results from the combinatorial joining of immunoglobulin gene (Ig) V, D, and J gene segments; from class switch recombination; and from somatic hypermutation of variable (V) regions (see Chapter 11). Several polymerases have been implicated in the DNA synthesis required for development of a normal immune system. Mammalian cells contain a template-independent family X polymerase called terminal deoxynucleotidyl transferase (TdT). TdT contains a
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BRCT domain characteristic of polymerases involved in DNA repair. It is expressed in lymphoid tissue, and there it is involved in a specialized polymerization reaction. In a template-independent manner, TdT inserts nucleotides (so-called N-regions) at the junctions between the V, D, and J elements during recombination to assemble expressed Ig heavy-chain genes. This results in junctional diversity in the coding sequence, thereby increasing the repertoire of immunoglobulins (Gellert, 2002; Neuberger et al., 2003; Thompson, 1995). Pol is closely related to TdT, and it too is highly expressed in lymphoid tissue. As mentioned above, it is thought to be involved in NHEJ of double-strand DNA breaks. Studies with Pol / mice indicate that Pol is involved in the processing of DNA ends during Ig light-chain gene rearrangement at a stage when TdT is no longer expressed (Bertocci et al., 2003). The somatic hypermutation (SHM) process is characterized by the frequent occurrence of base substitution mutations within a DNA segment of approximately one to two thousand bases of V regions in Ig genes. These mutations are generated at a frequency that is perhaps a million-fold higher than expected, given the very low rate of spontaneous mutation throughout the eukaryotic genome. The enzymatic mechanisms responsible for SHM are the subject of very active investigation (see Chapter 11). The mutations are generated in two distinct phases that have different base substitution specificity. SHM is likely to be initiated by enzymatic cytosine deamination by the activation-induced cytosine deaminase (AID), followed by replicative-type or repair-type DNA synthesis. Current biochemical and genetic evidence (reviewed in Chapter 11) indicates that the polymerases responsible for this synthesis may include members of family B, such as Pol , Pol , and Pol ", as well as members of family Y, such as Pol or Pol .
XIII. Biological Consequences of Polymerase Dysfunction There are now several examples in which mutations in polymerase genes that inactivate or modify enzymatic functions have consequences for human health (Kunkel, 2003). For example, several mutations in the polymerase and exonuclease domains of human Pol have recently been associated with progressive external ophthalmoplegia (PEO) (Copeland and Longley, 2003; Ponamarev et al., 2002). PEO is a rare disease characterized by muscle dysfunction resulting from the accumulation of point mutations and large deletions in mitochondrial DNA that eventually lead to loss of mitochondrial function. Humans carrying mutations in the XPV (POLH) gene that inactivate the function of Pol suffer from
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Xeroderma pigmentosum, a rare disease characterized by increased susceptibility to sunlight-induced skin cancer. Mice carrying a point mutation that inactivates the 30 to 50 exonuclease of Pol and eliminates its proofreading function have a recessive mutator phenotype (Goldsby et al., 2002, 2001). The mice also have a recessive cancer phenotype characterized by reduced lifespan (median survival, 10 months) and several tumor types, predominantly of epithelial-cell origin. This implies that DNA polymerase errors that are not proofread contribute to carcinogenesis. Thus, the consequences of loss of proofreading during replication generally conform to the mutator hypothesis for the origins of cancer, which posits that an early event in tumorigenesis is the expression of a mutator phenotype resulting from mutations in genes that normally function to maintain genome stability (Loeb, 2001). These connections between polymerase dysfunction and disease indicate that it will be worthwhile in the future to determine whether polymorphisms in DNA polymerase genes are associated with adverse human health.
XIV. Closing Comments A half century ago, in their seminal article describing the structure of the DNA double helix, Watson and Crick (Watson and Crick, 1953) wrote, ‘‘It has not escaped our notice that the specific pairing we have postulated immediately suggests a possible copying mechanism for the genetic material.’’ Very soon thereafter, E. coli Pol I and human Pol were discovered (reviewed in Kornberg and Baker, 1992). Fifty years later, we still do not completely understand the functions of these two polymerases, so it is no surprise that great uncertainty remains as to the precise functions of the many other polymerases discovered since then, many of which were only found in the last 5 years. What was certainly not appreciated 50 years ago was the large number and amazing diversity of transactions involving DNA synthesis required to faithfully replicate genomes that are diverse in sequence, in functional composition, and in organization, and to stably maintain them while they are being used for transcription and constantly being insulted by normal cellular metabolism and by the external environment. Although our knowledge of the existence and properties of DNA polymerases has greatly expanded, there are many exciting questions remaining to answer regarding the biological functions of each polymerase and the mechanisms by which they are regulated so as to function in the right place and at the right time. Many of the key issues, and our current understanding of them, are addressed in the following chapters and in the review articles liberally cited throughout this chapter.
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Acknowledgments We thank William Copeland, Matthew Longley, and Miguel Garcia-Diaz for critically reading of this chapter and for offering thoughtful suggestions. We also thank Miguel GarciaDiaz for help in preparing the figues. TAK dedicates this chapter to the memory of Dale W. Mosbaugh, an outstanding nucleic acid biochemist, a kind and generous human being, and a very dear friend.
References Aoufouchi, S., Flatter, E., Dahan, A., Faili, A., Bertocci, B., Storck, S., Delbos, F., Cocea, L., Gupta, N., Weill, J. C., and Reynaud, C. A. (2000). Two novel human and mouse DNA polymerases of the polX family. Nucleic Acids Res. 28, 3684–3693. Asahara, H., Li, Y., Fuss, J., Haines, D. S., Vlatkovic, N., Boyd, M. T., and Linn, S. (2003). Stimulation of human DNA polymerase epsilon by MDM2. Nucleic Acids Res. 31, 2451–2459. Beard, W. A., Shock, D. D., Vande Berg, B. J., and Wilson, S. H. (2002). Efficiency of correct nucleotide insertion governs DNA polymerase fidelity. J. Biol. Chem. 277, 47393–47398. Beard, W. A., and Wilson, S. H. (2000). Structural design of a eukaryotic DNA repair polymerase: DNA polymerase beta. Mutat. Res. 460, 231–244. Beard, W. A., and Wilson, S. H. (2003). Structural insights into the origins of DNA polymerase fidelity. Structure 11, 489–496. Bebenek, K., Tissier, A., Frank, E. G., McDonald, J. P., Prasad, R., Wilson, S. H., Woodgate, R., and Kunkel, T. A. (2001). 50 -Deoxyribose phosphate lyase activity of human DNA polymerase iota in vitro. Science 291, 2156–2159. Bemark, M., Khamlichi, A. A., Davies, S. L., and Neuberger, M. S. (2000). Disruption of mouse polymerase zeta (Rev3) leads to embryonic lethality and impairs blastocyst development in vitro. Curr. Biol. 10, 1213–1216. Berardini, M., Foster, P. L., and Loechler, E. L. (1999). DNA polymerase II (polB) is involved in a new DNA repair pathway for DNA interstrand cross-links in Escherichia coli. J. Bacteriol. 181, 2878–2882. Bertocci, B., De Smet, A., Berek, C., Weill, J. C., and Reynaud, C. A. (2003). Immunoglobulin kappa light chain gene rearrangement is impaired in mice deficient for DNA polymerase mu. Immunity 19, 203–211. Bertocci, B., De Smet, A., Flatter, E., Dahan, A., Bories, J. C., Landreau, C., Weill, J. C., and Reynaud, C. A. (2002). Cutting edge: DNA polymerases mu and lambda are dispensable for Ig gene hypermutation. J. Immunol. 168, 3702–3706. Bohr, V. A., and Dianov, G. L. (1999). Oxidative DNA damage processing in nuclear and mitochondrial DNA. Biochimie 81, 155–160. Bruck, I., Goodman, M. F., and O’Donnell, M. (2003). The essential C family DnaE polymerase is error-prone and efficient at lesion bypass. J. Biol. Chem. 278, 44361–44368. Bruck, I., and O’Donnell, M. (2000). The DNA replication machine of a gram-positive organism. J. Biol. Chem. 275, 28971–28983. Caldecott, K. W. (2003a). Protein-protein interactions during mammalian DNA singlestrand break repair. Biochem. Soc. Trans. 31, 247–251. Caldecott, K. W. (2003b). XRCC1 and DNA strand break repair. DNA Repair (Amst.) 2, 955–969.
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Cann, I. K., and Ishino, Y. (1999). Archaeal DNA replication: Identifying the pieces to solve a puzzle. Genetics 152, 1249–1267. Carrodeguas, J. A., Theis, K., Bogenhagen, D. F., and Kisker, C. (2001). Crystal structure and deletion analysis show that the accessory subunit of mammalian DNA polymerase gamma, Pol gamma B, functions as a homodimer. Mol. Cell 7, 43–54. Copeland, W. C., and Longley, M. J. (2003). DNA polymerase gamma in mitochondrial DNA replication and repair. Scientific WorldJournal 3, 34–44. Critchlow, S. E., and Jackson, S. P. (1998). DNA end-joining: From yeast to man. Trends Biochem. Sci. 23, 394–398. D’Urso, G., Grallert, B., and Nurse, P. (1995). DNA polymerase alpha, a component of the replication initiation complex, is essential for the checkpoint coupling S phase to mitosis in fission yeast. J. Cell Sci. 108(Pt 9), 3109–3118. Datta, A., Schmeits, J. L., Amin, N. S., Lau, P. J., Myung, K., and Kolodner, R. D. (2000). Checkpoint-dependent activation of mutagenic repair in Saccharomyces cerevisiae pol3-01 mutants. Mol. Cell 6, 593–603. Dervyn, E., Suski, C., Daniel, R., Bruand, C., Chapuis, J., Errington, J., Janniere, L., and Ehrlich, S. D. (2001). Two essential DNA polymerases at the bacterial replication fork. Science 294, 1716–1719. Dominguez, O., Ruiz, J. F., Lain de Lera, T., Garcia-Diaz, M., Gonzalez, M. A., Kirchhoff, T., Martinez, A. C., Bernad, A., and Blanco, L. (2000). DNA polymerase mu (Pol mu), homologous to TdT, could act as a DNA mutator in eukaryotic cells. EMBO J. 19, 1731–1742. Dua, R., Levy, D. L., and Campbell, J. L. (1998). Role of the putative zinc finger domain of Saccharomyces cerevisiae DNA polymerase epsilon in DNA replication and the S/M checkpoint pathway. J. Biol. Chem. 273, 30046–30055. Dua, R., Levy, D. L., and Campbell, J. L. (1999). Analysis of the essential functions of the C-terminal protein/protein interaction domain of Saccharomyces cerevisiae pol epsilon and its unexpected ability to support growth in the absence of the DNA polymerase domain. J. Biol. Chem. 274, 22283–22288. Esposito, G., Godindagger, I., Klein, U., Yaspo, M. L., Cumano, A., and Rajewsky, K. (2000). Disruption of the Rev31-encoded catalytic subunit of polymerase zeta in mice results in early embryonic lethality. Curr. Biol. 10, 1221–1224. Friedberg, E. C., Fischhaber, P. L., and Kisker, C. (2001). Error-prone DNA polymerases: Novel structures and the benefits of infidelity. Cell 107, 9–12. Friedberg, E. C., Walker, G. C., and Siede, W. (1995). DNA repair and mutagenesis. Washington, DC: ASM Press. Fuss, J., and Linn, S. (2002). Human DNA polymerase epsilon colocalizes with proliferating cell nuclear antigen and DNA replication late, but not early, in S phase. J. Biol. Chem. 277, 8658–8666. Garcia-Diaz, M., Bebenek, K., Krahn, J. M., Blanco, L., Kunkel, T. A., and Pedersen, L. C. (2004). Structural solution for the DNA polymerase -dependent repair of DNA gaps with minimal homology. Mol. Cell 13, 561–572. Garcia-Diaz, M., Bebenek, K., Kunkel, T. A., and Blanco, L. (2001). Identification of an intrinsic 50 -deoxyribose-5-phosphate lyase activity in human DNA polymerase lambda. A possible role in base excision repair. J. Biol. Chem. 276, 34659–34663. Garcia-Diaz, M., Bebenek, K., Sabariegos, R., Dominguez, O., Rodriguez, J., Kirchhoff, T., Garcia-Palomero, E., Picher, A. J., Juarez, R., Ruiz, J. F., Kunkel, T. A., and Blanco, L. (2002). DNA polymerase lambda, a novel DNA repair enzyme in human cells. J. Biol. Chem. 277, 13184–13191.
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Garcia-Diaz, M., Dominguez, O., Lopez-Fernandez, L. A., de Lera, L. T., Saniger, M. L., Ruiz, J. F., Parraga, M., Garcia-Ortiz, M. J., Kirchhoff, T., del Mazo, J., Bernad, A., and Blanco, L. (2000). DNA polymerase lambda (Pol lambda), a novel eukaryotic DNA polymerase with a potential role in meiosis. J. Mol. Biol. 301, 851–867. Gellert, M. (2002). V(D)J recombination: RAG proteins, repair factors, and regulation. Annu. Rev. Biochem. 71, 101–132. Gilfillan, S., Dierich, A., Lemeur, M., Benoist, C., and Mathis, D. (1993). Mice lacking TdT: Mature animals with an immature lymphocyte repertoire. Science 261, 1175–1178. Glover, B. P., and McHenry, C. S. (2001). The DNA polymerase III holoenzyme: An asymmetric dimeric replicative complex with leading and lagging strand polymerases. Cell 105, 925–934. Goldsby, R. E., Hays, L. E., Chen, X., Olmsted, E. A., Slayton, W. B., Spangrude, G. J., and Preston, B. D. (2002). High incidence of epithelial cancers in mice deficient for DNA polymerase delta proofreading. Proc. Natl. Acad. Sci. USA 99, 15560–15565. Goldsby, R. E., Lawrence, N. A., Hays, L. E., Olmsted, E. A., Chen, X., Singh, M., and Preston, B. D. (2001). Defective DNA polymerase-delta proofreading causes cancer susceptibility in mice. Nat. Med. 7, 638–639. Goodman, M. F. (2002). Error-prone repair DNA polymerases in prokaryotes and eukaryotes. Annu. Rev. Biochem. 71, 17–50. Gu, H., Marth, J. D., Orban, P. C., Mossmann, H., and Rajewsky, K. (1994). Deletion of a DNA polymerase beta gene segment in T cells using cell type-specific gene targeting. Science 265, 103–106. Guo, C., Fischhaber, P. L., Luk-Paszyc, M. J., Masuda, Y., Zhou, J., Kamiya, K., Kisker, C., and Friedberg, E. C. (2003). Mouse Rev1 protein interacts with multiple DNA polymerases involved in translesion DNA synthesis. EMBO J. 22, 6621–6630. Haracska, L., Kondratick, C. M., Unk, I., Prakash, S., and Prakash, L. (2001). Interaction with PCNA is essential for yeast DNA polymerase eta function. Mol. Cell 8, 407–415. Havener, J. M., McElhinny, S. A., Bassett, E., Gauger, M., Ramsden, D. A., and Chaney, S. G. (2003). Translesion synthesis past platinum DNA adducts by human DNA polymerase mu. Biochemistry 42, 1777–1788. Hoege, C., Pfander, B., Moldovan, G. L., Pyrowolakis, G., and Jentsch, S. (2002). RAD6dependent DNA repair is linked to modification of PCNA by ubiquitin and SUMO. Nature 419, 135–141. Holmes, A. M., and Haber, J. E. (1999). Double-strand break repair in yeast requires both leading and lagging strand DNA polymerases. Cell 96, 415–424. Hubscher, U., Maga, G., and Spadari, S. (2002). Eukaryotic DNA polymerases. Annu. Rev. Biochem. 71, 133–163. Ishibashi, T., Hayakawa, H., and Sekiguchi, M. (2003). A novel mechanism for preventing mutations caused by oxidation of guanine nucleotides. EMBO Rep. 4, 479–483. Jessberger, R., Podust, V., Hubscher, U., and Berg, P. (1993). A mammalian protein complex that repairs double-strand breaks and deletions by recombination. J. Biol. Chem. 268, 15070–15079. Kaguni, L. S. (2004). DNA polymerase gamma, the mitochondrial replicase. Annu. Rev. Biochem. 73, 293–320. Kelman, Z., and O’Donnell, M. (1995). DNA polymerase III holoenzyme: Structure and function of a chromosomal replicating machine. Annu. Rev. Biochem. 64, 171–200.
162
BEBENEK AND KUNKEL
Kesti, T., Flick, K., Keranen, S., Syvaoja, J. E., and Wittenberg, C. (1999). DNA polymerase epsilon catalytic domains are dispensable for DNA replication, DNA repair, and cell viability. Mol. Cell 3, 679–685. Kornberg, A., and Baker, T. (1992). DNA replication.2nd ed. New York: Freeman. Kunkel, T. A. (2003). Considering the cancer consequences of altered DNA polymerase function. Cancer Cell 3, 105–110. Kunkel, T. A. (2004). DNA replication Fidelity. J. Biol. Chem 279, 16895–16898. Lee, J. W., Blanco, L., Zhou, T., Garcia-Diaz, M., Bebenek, K., Kunkel, T. A., Wang, Z., and Povirk, L. F. (2004). Implication of DNA polymerase lambda in alignmentbased gap filling for nonhomologous DNA end joining in human nuclear extracts. J. Biol. Chem. 279, 805–811. Leem, S. H., Ropp, P. A., and Sugino, A. (1994). The yeast Saccharomyces cerevisiae DNA polymerase IV: Possible involvement in double strand break DNA repair. Nucleic Acids Res. 22, 3011–3017. Lieber, M. R. (1999). The biochemistry and biological significance of nonhomologous DNA end joining: An essential repair process in multicellular eukaryotes. Genes Cells 4, 77–85. Lim, S. E., Longley, M. J., and Copeland, W. C. (1999). The mitochondrial p55 accessory subunit of human DNA polymerase gamma enhances DNA binding, promotes processive DNA synthesis, and confers N-ethylmaleimide resistance. J. Biol. Chem. 274, 38197–38203. Lindahl, T., Karran, P., and Wood, R. D. (1997). DNA excision repair pathways. Curr. Opin. Genet. Dev. 7, 158–169. Lindahl, T., and Wood, R. D. (1999). Quality control by DNA repair. Science 286, 1897–1905. Ling, H., Boudsocq, F., Plosky, B. S., Woodgate, R., and Yang, W. (2003). Replication of a cis-syn thymine dimer at atomic resolution. Nature 424, 1083–1087. Ling, H., Boudsocq, F., Woodgate, R., and Yang, W. (2001). Crystal structure of a Y-family DNA polymerase in action: A mechanism for error-prone and lesionbypass replication. Cell 107, 91–102. Ling, H., Sayer, J. M., Plosky, B. S., Yagi, H., Boudsocq, F., Woodgate, R., Jerina, D. M., and Yang, W. (2004). Crystal structure of a benzo[a]pyrene diol epoxide adduct in a ternary complex with a DNA polymerase. Proc. Natl. Acad. Sci. USA 101, 2265–2269. Loeb, L. A. (2001). A mutator phenotype in cancer. Cancer Res. 61, 3230–3239. Longley, M. J., Pierce, A. J., and Modrich, P. (1997). DNA polymerase delta is required for human mismatch repair in vitro. J. Biol. Chem. 272, 10917–10921. Longley, M. J., Prasad, R., Srivastava, D. K., Wilson, S. H., and Copeland, W. C. (1998). Identification of 50 -deoxyribose phosphate lyase activity in human DNA polymerase
and its role in mitochondrial base excision repair in vitro. Proc. Natl. Acad. Sci. USA 95, 12244–12248. Mahajan, K. N., Nick McElhinny, S. A., Mitchell, B. S., and Ramsden, D. A. (2002). Association of DNA polymerase mu (pol mu) with Ku and ligase IV: Role for pol mu in end-joining double-strand break repair. Mol. Cell. Biol. 22, 5194–5202. Marini, F., Kim, N., Schuffert, A., and Wood, R. D. (2003). POLN, a nuclear PolA family DNA polymerase homologous to the DNA cross-link sensitivity protein Mus308. J. Biol. Chem. 278, 32014–32019. Marini, F., Pellicioli, A., Paciotti, V., Lucchini, G., Plevani, P., Stern, D. F., and Foiani, M. (1997). A role for DNA primase in coupling DNA replication to DNA damage response. EMBO J. 16, 639–650.
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163
Matsuda, T., Bebenek, K., Masutani, C., Hanaoka, F., and Kunkel, T. A. (2000). Low fidelity DNA synthesis by human DNA polymerase-eta. Nature 404, 1011–1013. McCulloch, S. D., Kokoska, R. J., Masutani, C., Iwai, S., Hanaoka, F., and Kunkel, T. A. (2004). Preferential cis-syn thymine dimer bypass by DNA polymerase h occurs with biased fidelity. Nature 427, 97–100. McDonald, J. P., Frank, E. G., Plosky, B. S., Rogozin, I. B., Masutani, C., Hanaoka, F., Woodgate, R., and Gearhart, P. J. (2003). 129-derived strains of mice are deficient in DNA polymerase iota and have normal immunoglobulin hypermutation. J. Exp. Med. 198, 635–643. McHenry, C. S. (2003). Chromosomal replicases as asymmetric dimers: Studies of subunit arrangement and functional consequences. Mol. Microbiol. 49, 1157–1165. Murakumo, Y., Roth, T., Ishii, H., Rasio, D., Numata, S., Croce, C. M., and Fishel, R. (2000). A human REV7 homolog that interacts with the polymerase zeta catalytic subunit hREV3 and the spindle assembly checkpoint protein hMAD2. J. Biol. Chem. 275, 4391–4397. Musacchio, A., and Hardwick, K. G. (2002). The spindle checkpoint: Structural insights into dynamic signalling. Nat. Rev. Mol. Cell Biol. 3, 731–741. Navas, T. A., Sanchez, Y., and Elledge, S. J. (1996). RAD9 and DNA polymerase epsilon form parallel sensory branches for transducing the DNA damage checkpoint signal in Saccharomyces cerevisiae. Genes Dev. 10, 2632–2643. Navas, T. A., Zhou, Z., and Elledge, S. J. (1995). DNA polymerase epsilon links the DNA replication machinery to the S phase checkpoint. Cell 80, 29–39. Neuberger, M. S., Harris, R. S., Di Noia, J., and Petersen-Mahrt, S. K. (2003). Immunity through DNA deamination. Trends Biochem. Sci. 28, 305–312. Nick McElhinny, S. A., and Ramsden, D. A. (2003). Polymerase mu is a DNA-directed DNA/RNA polymerase. Mol. Cell. Biol. 23, 2309–2315. Ohmori, H., Friedberg, E. C., Fuchs, R. P., Goodman, M. F., Hanaoka, F., Hinkle, D., Kunkel, T. A., Lawrence, C. W., Livneh, Z., Nohmi, T., Prakash, L., Prakash, S., Todo, T., Walker, G. C., Wang, Z., and Woodgate, R. (2001). The Y-family of DNA polymerases. Mol. Cell 8, 7–8. Ollis, D. L., Brick, P., Hamlin, R., Xuong, N. G., and Steitz, T. A. (1985). Structure of large fragment of Escherichia coli DNA polymerase I complexed with dTMP. Nature 313, 762–766. Page´s, V., and Fuchs, R. P. (2002). How DNA lesions are turned into mutations within cells. Oncogene 21, 8957–8966. Pham, P., Bertram, J. G., O’Donnell, M., Woodgate, R., and Goodman, M. F. (2001a). A model for SOS-lesion-targeted mutations in Escherichia coli. Nature 409, 366–370. Pham, P., Rangarajan, S., Woodgate, R., and Goodman, M. F. (2001b). Roles of DNA polymerases V and II in SOS-induced error-prone and error-free repair in Escherichia coli. Proc. Natl. Acad. Sci. USA 98, 8350–8354. Ponamarev, M. V., Longley, M. J., Nguyen, D., Kunkel, T. A., and Copeland, W. C. (2002). Active site mutation in DNA polymerase gamma associated with progressive external ophthalmoplegia causes error-prone DNA synthesis. J. Biol. Chem. 277, 15225–15228. Ruiz, J. F., Juarez, R., Garcia-Diaz, M., Terrados, G., Picher, A. J., Gonzalez-Barrera, S., Fernandez de Henestrosa, A. R., and Blanco, L. (2003). Lack of sugar discrimination by human Pol mu requires a single glycine residue. Nucleic Acids Res. 31, 4441–4449.
164
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Saitoh, S., Chabes, A., McDonald, W. H., Thelander, L., Yates, J. R., and Russell, P. (2002). Cid13 is a cytoplasmic poly(A) polymerase that regulates ribonucleotide reductase mRNA. Cell 109, 563–573. Schenten, D., Gerlach, V. L., Guo, C., Velasco-Miguel, S., Hladik, C. L., White, C. L., Friedberg, E. C., Rajewsky, K., and Esposito, G. (2002). DNA polymerase kappa deficiency does not affect somatic hypermutation in mice. Eur. J. Immunol. 32, 3152–3160. Seki, M., Marini, F., and Wood, R. D. (2003). POLQ (Pol theta), a DNA polymerase and DNA-dependent ATPase in human cells. Nucleic Acids Res. 31, 6117–6126. Sekiguchi, M. (1996). MutT-related error avoidance mechanism for DNA synthesis. Genes Cells 1, 139–145. Sharief, F. S., Vojta, P. J., Ropp, P. A., and Copeland, W. C. (1999). Cloning and chromosomal mapping of the human DNA polymerase theta (POLQ), the eighth human DNA polymerase. Genomics 59, 90–96. Shcherbakova, P. V., Bebenek, K., and Kunkel, T. A. (2003). Functions of eukaryotic DNA polymerases. Sci. Aging Knowledge Environ 2003, RE3. Shcherbakova, P. V., and Pavlov, Y. I. (1996). 30 !50 exonucleases of DNA polymerases epsilon and delta correct base analog induced DNA replication errors on opposite DNA strands in Saccharomyces cerevisiae. Genetics 142, 717–726. Silvian, L. F., Toth, E. A., Pham, P., Goodman, M. F., and Ellenberger, T. (2001). Crystal structure of a DinB family error-prone DNA polymerase from Sulfolobus solfataricus. Nat. Struct. Biol. 8, 984–989. Steitz, T. A., Smerdon, S. J., Jager, J., and Joyce, C. M. (1994). A unified polymerase mechanism for nonhomologous DNA and RNA polymerases. Science 266, 2022–2025. Thompson, C. B. (1995). New insights into V(D)J recombination and its role in the evolution of the immune system. Immunity 3, 531–539. Trincao, J., Johnson, R. E., Escalante, C. R., Prakash, S., Prakash, L., and Aggarwal, A. K. (2001). Structure of the catalytic core of S. cerevisiae DNA polymerase eta: Implications for translesion DNA synthesis. Mol. Cell 8, 417–426. Tseng, H. M., and Tomkinson, A. E. (2002). A physical and functional interaction between yeast Pol4 and Dn14-Lif1 links DNA synthesis and ligation in nonhomologous end joining. J. Biol. Chem. 277, 45630–45637. Ulrich, H. D. (2004). How to activate a damage-tolerant polymerase: Consequences of PCNA modifications by ubiquitin and SUMO. Cell Cycle 3, 15–18. Van Houten, B. (1990). Nucleotide excision repair in Escherichia coli. Microbiol. Rev. 54, 18–51. Vlatkovic, N., Guerrera, S., Li, Y., Linn, S., Haines, D. S., and Boyd, M. T. (2000). MDM2 interacts with the C-terminus of the catalytic subunit of DNA polymerase epsilon. Nucleic Acids Res. 28, 3581–3586. Wang, S. W., Toda, T., MacCallum, R., Harris, A. L., and Norbury, C. (2000a). Cid1, a fission yeast protein required for S-M checkpoint control when DNA polymerase delta or epsilon is inactivated. Mol. Cell. Biol. 20, 3234–3244. Wang, Z., Castano, I. B., De Las Penas, A., Adams, C., and Christman, M. F. (2000b). Pol kappa: A DNA polymerase required for sister chromatid cohesion. Science 289, 774–779. Wang, Z., Wu, X., and Friedberg, E. C. (1993). DNA repair synthesis during base excision repair in vitro is catalyzed by DNA polymerase epsilon and is influenced
FUNCTIONS OF DNA POLYMERASES
165
by DNA polymerases alpha and delta in Saccharomyces cerevisiae. Mol. Cell. Biol. 13, 1051–1058. Watson, J. D., and Crick, F. H. C. (1953). Molecular structure of nucleic acids; a structure for deoxyribose nucleic acid. Nature 171, 737–738. Wilson, T. E., and Lieber, M. R. (1999). Efficient processing of DNA ends during yeast nonhomologous end joining. Evidence for a DNA polymerase beta (Pol4)dependent pathway. J. Biol. Chem. 274, 23599–23609. Wittschieben, J., Shivji, M. K., Lalani, E., Jacobs, M. A., Marini, F., Gearhart, P. J., Rosewell, I., Stamp, G., and Wood, R. D. (2000). Disruption of the developmentally regulated Rev31 gene causes embryonic lethality. Curr. Biol. 10, 1217–1220. Zhang, Y., Wu, X., Guo, D., Rechkoblit, O., Taylor, J. S., Geacintov, N. E., and Wang, Z. (2002). Lesion bypass activities of human DNA polymerase mu. J. Biol. Chem. 277, 44582–44587. Zhang, Y., Wu, X., Yuan, F., Xie, Z., and Wang, Z. (2001). Highly frequent frameshift DNA synthesis by human DNA polymerase mu. Mol. Cell. Biol. 21, 7995–8006. Zhou, B. L., Pata, J. D., and Steitz, T. A. (2001). Crystal structure of a DinB lesion bypass DNA polymerase catalytic fragment reveals a classic polymerase catalytic domain. Mol. Cell 8, 427–437.
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CELLULAR FUNCTIONS OF DNA POLYMERASE z AND REV1 PROTEIN By CHRISTOPHER W. LAWRENCE Department of Biochemistry and Biophysics, University of Rochester School of Medicine and Dentistry, Rochester, New York, 14642
I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . II. Enzymological Studies with Pol and Rev1p. . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Properties of Pol and Rev1p .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. In Vitro Studies of Pol and Rev1p on Lesion-Containing Templates . . . .. . . . . . III. Genetic Analysis . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . IV. Processes Other than General Translesion Replication that Employ Pol and Rev1p . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Somatic Hypermutation. . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. Double-Strand Break Repair. . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . V. Regulation of Pol and Rev1p and Interactions with Other Proteins . . . . . . . .. . . . . . VI. Conclusions and Speculations . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . References . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . .
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I. Introduction DNA polymerase (Pol) and Rev1 protein (Rev1p) perform a variety of important and, in some cases, essential, functions in eukaryotes, including roles in DNA damage tolerance, in the development of diversity among immunoglobulin genes, and in the repair of double-strand breaks by homologous recombination. Originally discovered in budding yeast, Saccharomyces cerevisiae, the genes encoding these enzymes have been found in all fully sequenced eukaryotic genomes, including those of other microbial organisms, nematodes, mammals, and plants, suggesting that they are present in all eukaryotes (Lawrence, 2002; Lawrence et al., 2000). Of the various processes in which they participate, their roles in translesion replication are probably the most widespread, though they may not be universal. Like other DNA damage tolerance mechanisms, translesion replication is concerned with overcoming the major consequence of unrepaired damage in the genome; namely, its ability to block the progress of replicases and thus prevent complete replication of the genome. Translesion replication achieves this with the aid of Pol and Rev1p, together with other specialized DNA polymerases, such as Pol. Although the activities of Pol and Rev1p contribute only modestly to resistance to DNA-damaging agents in yeast, they participate in processes that generate 167 ADVANCES IN PROTEIN CHEMISTRY, Vol. 69
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the majority of spontaneous mutations and almost all of those induced by overt mutagenic treatment. Interestingly, these enzymes appear to contribute more substantially to damage resistance in at least some vertebrates, raising the question of the molecular basis for this increased importance. Even so, vertebrate Pol and Rev1p appear to be involved in the production of most spontaneous and DNA damage–induced mutations, like their yeast counterparts. Understanding the molecular processes underlying translesion replication and mutagenesis is therefore likely to provide important insights into cancerogenesis and other genetic diseases in which mutations figure prominently. In this context, it is also significant that rev1 and rev3 yeast mutants are sensitive to many, though not all, of a large set of anticancer agents (Simon et al., 2000). The yeast REV3 gene, later shown to encode the catalytic subunit of Pol, and the REV1 gene were first identified by Lemontt (1971) using a screen for mutants with reduced frequencies of UV (ultraviolet)-induced mutagenesis. REV7, encoding a second subunit of Pol, was subsequently identified using a similar procedure (Lawrence et al., 1985a,b). The PSO1 locus, independently identified in a screen for mutants sensitive to 8-methoxy-psoralen plus 365 nm UV, was later found to be the REV3 gene (Cassier-Chauvat and Moustacchi, 1988). Sequence analysis of cloned REV3 (Morrison et al., 1989) revealed a predicted protein, with the motifs of an enzyme belonging to the B family of DNA polymerases in the classification of Ito and Braithwaite (Braithwaite and Ito, 1993; Ito and Braithwaite, 1991), distantly related to DNA polymerase . Analysis of the cloned REV1 gene (Larimer et al., 1989) identified a predicted protein containing a 152-residue region with 25% identity and 42% similarity to a segment of the E. coli UmuC protein, both of which were subsequently designated as members of the recently designated Y family of DNA polymerases (Ohmori et al., 2001). Clones of the human homologs of REV3 (Gibbs et al., 1998; Lin et al., 1998; Morelli et al., 1998; Xiao et al., 1998), REV1 (Gibbs et al., 2000; Lin et al., 1999) and REV7 (Murakumo et al., 2000) have been isolated subsequently, as well as clones of mouse REV3 (Van Sloun et al., 1999) and mouse REV1 (Gibbs and Lawrence, unpublished; Masuda et al., 2002). REV3 homologs have also been characterized in Aspergillus nidulans (Han et al., 1998), Drosophila melanogaster (Eeken et al., 2001), Neurospora crassa (Sakai et al., 2002), Arabidopsis thaliana (Sakamoto et al., 2003), and chickens (Sonoda et al., 2003). The properties of Neurospora REV1 and REV7 homologs (Sakai et al., 2003), and the REV1 homolog of chicken (Simpson and Sale, 2003), have also been investigated. The domain structure of Rev1p, Rev3p, and Rev7p in budding yeast, humans, and Arabidopsis thaliana is diagrammed in Fig. 1.
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Fig. 1. Domain structure and percentage identity of Rev1, Rev3, and Rev7 proteins from yeast, humans, and Arabidopsis. A.t., Arabidopsis thaliana, S.c., Saccharomyces cerevisiae, H.s., Homo sapiens. Numbers between lines connecting homologous domains indicate percentage identity. Roman numerals I to VI above the Rev3 polymerase domain indicate conserved sequence motifs.
Following the initial characterization of the yeast rev mutant phenotypes, further investigations showed that they are deficient not only for mutagenesis induced by exposure to 254 nm UV (Lawrence and Christensen, 1976, 1978, 1979; Lawrence et al., 1984, 1985a; Lemontt, 1971, 1972) but also for mutagenesis induced by treatments with a wide array of contrasting agents that also damage DNA. Rev3 and rev7 mutants are deficient for mutagenesis induced by ionizing radiation (McKee and Lawrence, 1979a,b), 4-nitroquinoline-1-oxide (Prakash, 1976), methyl methane sulfonate and N-methyl-N 0 -nitro-N-nitrosoguanidine (Lawrence et al., 1985b), ethyl methane sulfonate (Lemontt, 1972; Prakash, 1976), 8-methoxy psoralen plus 365 nm UV light (Henriques and Moustacchi, 1980), and different alkylating agents (Ruhland and Brendel, 1979).
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Although fewer studies have been made with rev1 mutants, their phenotype appears to be similar (Lawrence and Christensen, 1976, 1978; Lawrence et al., 1984; Lemontt, 1971, 1972; McKee and Lawrence, 1979a,b). Combined results from a variety of data suggest that Pol function is required for the generation of 98%, and Rev1p function for 95%, of UV-induced base-pair substitutions. However, although Pol function is required for the generation of 90% of UV-induced frameshifts, Rev1p function is more variably involved in the production of these events; for some, it is required to a similar extent as Pol, whereas for others its involvement is much less (Lawrence and Christensen, 1978; Lawrence et al., 1984). As well as possessing a deficiency with respect to mutagenesis induced by many DNA-damaging agents, rev1 and rev3 mutants of yeast are also antimutators, with spontaneous mutation rates that are half to a quarter of those seen in REV þ strains. Although this aspect of the rev3 mutant phenotype was first uncovered in a screen specifically designed to recover antimutator mutants (Quah et al., 1980), and during the characterization of pso1 mutants (Cassier et al., 1980), Pol activity was later found to be responsible for the enhanced spontaneous mutagenesis observed in a range of different genetic circumstances, which include the mutator phenotypes associated with rad1, rad6, rad18, and rad52 mutants (Roche et al., 1994, 1995), with transcription (Datta and Jinks-Robertson, 1995) and with double-strand break repair (Holbeck and Strathern, 1997). Rev1p, as well as Pol, was found to be responsible for the increased spontaneous mutation rate associated with overproduction of the 3-methyladenine DNA glycosylase encoded by MAG1 (Glassner et al., 1998). Although the spontaneous mutations observed in the above investigations were principally base-pair substitutions, Pol and Rev1p activities were also observed to be largely responsible for the occurrence of spontaneous frameshift mutations found in rad1, rad2, rad14, and rad52 strains and to be entirely responsible for the complex events arising in these strains, in which from one to five base-pair substitutions occurred in the region of the þ1 insertion that led to the reversion of the lys2A746 allele employed (Harfe and Jinks-Robertson, 2000). Pol was also responsible for the enhanced frequencies of lys2BglII revertants found in stationary phase cells of rad14 and rad16 strains (Heidenreich et al., 2004). Investigations of REV gene function in organisms other than budding yeast, although fewer in number, also support a role for these genes in DNA damage–induced mutagenesis, and consequently in translesion replication. The organisms studied include filamentous fungi, plants, and mammals, encouraging the conclusion that the properties observed are likely to be found in almost all eukaryotes. Mutations in the UVSI gene of
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Aspergillus nidulans, which encodes a REV3 homolog (Han et al., 1998), are defective for UV mutagenesis (Chae and Kafer, 1993), and a similar mutant phenotype has been observed for the upr-1 mutant of Neurospora crassa, which encodes a REV3 homolog in this organism (Sakai et al., 2002). REV1 and REV7 homologs have also been identified in Neurospora and were shown to possess the same mutant phenotype (Sakai et al., 2003). Disruption of the REV3 homolog in A. thaliana led to increased sensitivity of the plants to UV-B irradiation (>280 nm, peak 312 nm), -rays, methyl methane sulfonate, and mitomycin C, though an influence on mutagenesis was not tested (Sakamoto et al., 2003). Reduction of cellular levels of human Rev3p (Gibbs et al., 1998; Li et al., 2002) or Rev1p (Gibbs et al., 2000) by high expression of antisense RNA decreased the frequencies of 6-thioguanine resistant mutants induced in human fibroblasts by 254 nm UV by about sevenfold and more than 20-fold, respectively. High expression of REV3 antisense RNA also reduced the frequency of 6-thioguanineresistant mutants induced by benzo[a]pyrene diol epoxide by four- to sixfold in these cells (Li et al., 2002). In a related approach, reduction of Rev1p expression with a ribozyme construct decreased the frequency of 254-nm UV-induced 6-thioguanine-resistant mutants in human cells by two- to threefold (Clark et al., 2003). Last, the frequency of spontaneous 6-thioguanine-resistant mutants was decreased several fold by high levels of REV3 antisense RNA in an msh6 human fibroblast cell line (X. Li and V.M. Maher, unpublished data, cited in Lawrence et al., 2000). Results from each of these investigations therefore resemble, at least qualitatively, those observed with yeast, and indicate that the functions of Pol and Rev1p are conserved from fungi to plants and humans. However, the involvement of Pol in translesion replication may not be universal. Although Drosophila melanogaster possesses a REV3 homolog, no evidence could be found in this organism for its involvement in forward mutagenesis induced by x-rays, 4-nitroquinoline-1-oxide, or methyl methane sulfonate; instead, it appears to be concerned with repair (Eeken et al., 2001). This chapter reviews information about the properties and functions of Pol and Rev1p in the diverse processes within which they play a part. Although most of these data come from yeast, important results have also been obtained from a variety of other species, including the mouse, humans, and chickens. A combination of investigations over the last several years, examining the enzymology, genetics, and cell biology of Pol and Rev1p, has done much to advance our understanding of their cellular functions, but conflicting interpretations remain with respect to several important issues. A second aim of this chapter is, therefore, to review the different models proposed for Pol and Rev1p function, and examine the data that are used to support them.
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II. Enzymological Studies With Pol And Rev1p In vitro studies of the purified yeast enzymes and, for Rev1p, its mammalian counterpart have established some of the basic properties of these proteins. They have also been used to investigate the role of these enzymes, often in combination with other DNA polymerases, in replication past various lesions, leading to proposals concerning the in vivo function of the enzymes. It is likely, however, that such reactions lack important factors present within cells, emphasizing the need for validation of the models with in vivo studies, as discussed in Section III.
A. Properties of Pol z and Rev1p Although yeast REV3 encoded a predicted protein that contained motifs characteristic of a DNA polymerase, biochemical demonstration of this activity was obtained only after identification of a second subunit, encoded by REV7 (Nelson et al., 1996a; Torpey et al., 1994). All in vitro studies of Pol have employed this two-subunit enzyme from yeast; efforts to produce it from mammalian genes, or those from other organisms, have so far been unsuccessful. Because native Pol has not been isolated from any species, probably the consequence of very low cellular levels, it is uncertain whether other subunits exist, or whether the two-subunit enzyme possesses a fully normal activity. In particular, a variety of evidence suggests that Rev1p is essential for Pol activity, but a functional association with this protein has not yet been reconstructed, perhaps because of a requirement for other proteins. As expected for a B-family polymerase, the activity of the two-subunit enzyme is DNA-template dependent and employs all four dNTPs. It is, however, poorly processive, adding three or fewer nucleotides per binding event, and it lacks a 30 –50 proofreading exonuclease activity (Nelson et al., 1996a). Compared to other B-family DNA polymerases, Pol has an unusual facility for extending primers with terminal mismatches ( Johnson et al., 2000; Lawrence and Hinkle, 1996; Lawrence et al., 2000), suggesting a general capability for elongation from structurally abnormal termini of the kind presented by DNA lesions. Apparent extension efficiencies from terminal mismatches, as determined by the method of Mendelman et al. (1990), were strongly sequence dependent, but were always greater for Pol than for Pol, which, like Pol, also lacks proofreading activity. In one sequence context, Pol was twofold to 1265-fold more efficient than Pol within the complete set of mismatches examined, and in another, Pol was fivefold to 1,226-fold more efficient. For some mismatches, extension by Pol was only a few fold less efficient than extension from the correctly
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paired terminus. In the most extreme case, in which a primer terminal G was mispaired with template T, apparent extension efficiencies using Pol were 54% and 35% of the values for the AT match in the two sequence contexts examined, whereas for Pol, the corresponding efficiencies were 1.7% and 0.33% (Lawrence et al., 2000). In addition to an unusual facility for extension from mismatched ends of primers, Pol appears to incorporate nucleotides with only a moderate fidelity. On lesion-free templates, Pol was found to have an incorporation error rate of between 4 103 and 2 105, depending on the particular template base and incoming nucleotide examined, with most values lying between 104 and 105 ( Johnson et al., 2000). Although the authors suggest that these values approach those for Pol, citing data from Thomas et al. (1991), results in the latter paper indicate that the error rates for Pol are at least 10-fold lower, with values of between 2.9 105 and 3.6 106, depending on the site studied. However, because the data from Thomas and coworkers were calculated from LacZ forward mutation rates during gap-filling synthesis on an M13mp2 template, a very different procedure to that used by Johnson and coworkers, it would be useful to directly compare the two enzymes using the same primed templates and methodology. On the basis of these data, it has been suggested that Pol is incapable of inserting nucleotides opposite lesions, but extends termini resulting from insertions by other enzymes ( Johnson et al., 2000; Prakash and Prakash, 2002), a model that is discussed in Section VI. Yeast Rev1p has two distinct functions, neither of which is fully understood. The most general, with respect to different DNA lesions, is its apparent requirement for Pol activity (see Section III), and a possible mechanistic model for this function is given in Section V. The second, more specialized, function is a DNA template-dependent deoxycytidyl transferase activity. In the original qualitative study of this activity, Rev1p was found to preferentially insert dCMP opposite a template abasic site and, less efficiently, opposite template guanine and adenine nucleotides (Nelson et al., 1996b). The reaction was highly specific for dCMP and could not use ribonucleoside triphosphates. This finding explained a prior observation (Gibbs and Lawrence, 1995), that dCMP was inserted in 60%–85% of the bypass events at a site-specific abasic residue in vivo, a preference that was later shown to be entirely dependent on Rev1p (Nelson et al., 2000). Similar to yeast Rev1p, both the human (Lin et al., 1999; Zhang et al., 2002) and the mouse enzyme (Masuda et al., 2002) possess a deoxycytidyl transferase activity, though low levels of dGMP and dTMP incorporation were also observed with these enzymes. Steady-state kinetic studies with each of these three Rev1p (Haracska et al., 2002; Masuda et al., 2002; Zhang et al., 2002) indicated that dCMP insertion
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was most efficient opposite sites of template guanine, though insertion at an abasic site was more efficient than at template A, T, or C, and the degree of preference for template guanine varied between studies. Because the efficiency of dCMP insertion by mouse Rev1p was influenced strongly by sequence context, the quantitatively disparate results may arise from this variable, though there may also be intrinsic differences between the proteins from each organism. The extent of the preference for dCMP incorporation at template guanine rather than an abasic site was particularly great in results with the yeast enzyme, leading the authors to propose that Rev1p is a G-templatespecific DNA polymerase (Haracska et al., 2002). Because there is clear evidence for Rev1p-dependent insertion of dCMP opposite an abasic site in vivo in yeast (Gibbs and Lawrence, 1995; Nelson et al., 2000; Otsuka et al., 2002a; 2004), such a designation appears inappropriate. Moreover, it seems unlikely that eukaryotes require an inefficient, highly unprocessive, Y-family enzyme to incorporate dCMP opposite normal guanine residues in the presence of highly efficient and accurate enzymes such as Pol. The apparent conflict between the in vivo and in vitro results may perhaps be explained by Rev1p existing within a complex in the cell, in which dCMP insertion is inhibited at template sites other than those with abasic residues and, perhaps, some other lesions. Finally, the deoxycytidyl transferase activity, even if not essential for the bypass of abasic sites, nevertheless doubles the efficiency of replication past this lesion in yeast (Otsuka et al., 2002b; see also Section III), perhaps explaining why the activity has been maintained throughout evolution.
B. In Vitro Studies of Polz and Rev1p on Lesion-Containing Templates A variety of investigations has been carried out to examine the roles of Pol and Rev1p on lesion-containing templates, usually in combination with other DNA polymerases or proteins concerned with translesion replication. Oligonucleotide templates containing a diverse variety of lesions have been used in such studies, including those carrying an abasic site, thymine–thymine pyrimidine (6–4) pyrimidinone adduct [T-T (6–4) photoadduct], thymine–thymine cis-syn cyclobutane dimer (T-T dimer), acetyl aminofluorene-guanine adduct (AAF-G), (þ) and () anti-benzo[a]pyrene diol epoxide (BPDE), 7,8-dihydro-8-oxoguanine (8-oxoG), O6-methyl guanine (6MeG), thymine glycol, and acrolein derivatives of guanine. Pol appears to play a part in the bypass of all of these lesions, apart from the T-T dimer, though the particular role may vary. Abasic residues are one of the most abundant types of DNA damage, with 10,000 of these lesions arising daily in mammalian genomes
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(Lindahl, 1993). At the same time, they constitute a fairly severe block to continued replication. The identity of the enzymes used by yeast to replicate past an abasic site has been investigated by several groups. Nelson et al. (1996b) showed that although Pol alone replicated past an abasic residue very inefficiently, it readily extended the primer resulting from incorporation of dCMP opposite the lesion by Rev1p. Yeast Pol, however, was unable to extend from such an insertion. This result is consistent with in vivo data showing dependence of such bypass on Pol and Rev1p, and preferential incorporation of dCMP opposite the abasic site (Gibbs and Lawrence, 1995; Nelson et al., 2000). Yuan et al. (2000). On the other hand, examined insertion opposite the lesion by Pol, which was unable to further extend the primer, followed by elongation with Pol. Pol principally inserted dAMP and dGMP opposite the abasic residue, though less frequently it incorporated the other two nucleotides. In contrast, a steadystate kinetic analysis (Haracska et al., 2001a) indicated that insertion opposite this lesion by either yeast or human Pol was very inefficient. This was partly alleviated by the presence of PCNA, RPA, and RFC (Haracska et al., 2001a), but Pol could not extend from these insertions. Yuan and coworkers (2000) also presented data indicating that yeast Pol was capable of bypassing an abasic site, principally incorporating dAMP, but such bypass was achieved only with high enzyme-to-template ratios that are probably uncharacteristic of in vivo conditions. The identity of the enzymes responsible for replication past abasic residues was also investigated by measuring bypass product yields resulting from in vitro reactions containing combinations of Pol, with either Pol or Rev1p (Haracska et al., 2001c). None of these enzymes alone was found to be capable of replicating past the abasic residue, though, under the conditions used, Pol could insert dAMP opposite the lesion, and Rev1p, as expected, could insert dCMP, with Pol capable of extending from both of these nucleotides. Of these two combinations, it was concluded that bypass of this lesion in vivo entailed insertion opposite the abasic site by Pol, rather than by Rev1p, followed by extension of the primer by Pol, because the reaction efficiency was greater. In a running-start assay, where the primer was set back 15 nucleotides from the abasic site, the amount of bypass product yielded by the Pol/Rev1p combination was 33% of that produced by Pol/Pol, whereas in a standing-start assay, where the primer abutted the lesion, the relative yield was 55%. Of these two estimates, the latter is probably the better one because Pol is a much less processive enzyme than Pol, suggesting that the efficiencies of the enzyme combinations differ by less than twofold. A difference this small does not seem to provide decisive support for the author’s hypothesis; its significance is difficult to evaluate, both because the specific activity of the proteins is unknown and because it
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is unclear whether the two-subunit Pol used fully reconstitutes the in vivo activity of this enzyme. In addition to these data, Haracska and coworkers also supported their model with a variety of genetic results, which are discussed in Section VI. Because the function of pol is known to be required for UV-induced mutagenesis (reviewed in Lawrence, 2002), the role of this enzyme in replication past UV lesions has also been investigated (Guo et al., 2001; Johnson et al., 2000, 2001). Before the discovery of Pol, Pol was originally described as having a modest capability for replicating past a T-T cyclobutane dimer (Nelson et al., 1996a), but later work indicated a much lower bypass frequency, and Pol is now known to be solely responsible for replication past this lesion (Gibbs et al., 2004; Johnson et al., 1999). Such is not the case with the T-T (6–4) UV photoproduct, which, unlike the dimer, severely distorts local DNA structure and possesses no capability for base-pairing. Johnson and coworkers (Johnson et al., 2001) found that Pol was incapable of insertion opposite the 30 T of the photoproduct. It could, however, efficiently extend from insertions carried out by either yeast or human Pol, of which both preferentially incorporated dGMP at this site. In the work of Guo and coworkers (Guo et al., 2001), in contrast, Pol was found to inefficiently bypass the T-T (6–4), inserting dAMP and dTMP, and more rarely dGMP opposite the 30 T, and inserting predominantly dAMP opposite the 50 T. Insertion at this site was most efficient following dGMP incorporation. Pol was not examined in this study. The apparent lack of agreement between these two investigations with respect to Pol can probably be ascribed to differences in experimental procedure; in the first, reactions contained a several-fold molar excess of primer/template over enzymes and were terminated after 5 minutes of incubation, whereas in the second investigation, reactions contained a fourfold excess of enzyme over primer/template and were incubated for 30 minutes, the latter conditions favoring insertion. As discussed in section III, in vivo experiments with yeast support the involvement of Pol in the bypass of the T-T (6–4), but the extent of this involvement varies from substantial to very small in different studies, the latter case raising the question of whether Pol or some other enzyme performs insertion opposite the 30 T. Such work also shows that replication past a T-T (6–4) photoadduct in vivo is at best very inefficient, as might be expected with such a distorting lesion, with a bypass efficiency of only 4%. As a consequence, even enzymes that bypass this lesion only inefficiently in vitro cannot be eliminated as candidates for this function in vivo. In addition to UV photoproducts, a variety of DNA lesions that result from treatment with chemical mutagens has also been examined as being potential substrates for Pol. AAF-guanine was found by Guo et al. (2001)
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to substantially inhibit this enzyme, with only a low efficiency of insertion opposite the lesion, and very little extension beyond it, despite a fourfold molar excess of enzyme over primer/template. However, Pol was able to extend primers that overlapped the lesion, most efficiently with that creating a terminal AGAAF mispair, though the correctly paired CGAAF was extended almost as well. Pol was found to bypass both (þ) and () enantiomers of benzo[a]pyrene diol epoxide-deoxyguanosine adducts on 18% of templates containing these lesions. Bypass was accurate, with insertion of only dCMP in >300 samples analyzed per lesion (Simhadri et al., 2002). However, the bypass frequencies observed are of questionable relevance to in vivo conditions, because the reactions lasted 90 minutes and were carried out with high levels of the enzyme. In another study of these lesions, and also the corresponding adducts of adenine, none of them were detectably bypassed by Pol in reactions containing a modest molar excess of Pol and incubation periods of up to 30 minutes (Rechkoblit et al., 2002). The possible role of Pol and Rev1p in the bypass of propanodeoxyguanosine, a lesion derived from acrolein that is replicated accurately in vivo despite its lack of base-pairing capacity, has also been investigated (Yang et al., 2003). Both Rev1p and Pol were found to efficiently incorporate dCMP opposite this adduct, as needed for accurate bypass, but were both unable to extend from this terminus, indicating that this function might be carried out by Pol. However, Pol extended from the inserted dCMP less efficiently than from dAMP, dGMP, or dTMP insertions. Because Pol was unable to bypass the lesion, the basis for the accurate replication was left unexplained. Steady-state kinetic data suggest that Pol is also capable of extension from misinsertions of dAMP or dTMP opposite 8-oxoG and O6-methylguanine by Pol, thought to be involved in the bypass of these lesions because of reduced mutation frequencies induced by MNNG in strains deleted for the Pol subunit, Pol32p (Haracska et al., 2000, 2003). However, as discussed in Section VI, it is doubtful that the involvement of the polymerase function of Pol can be inferred from the pol32 mutant phenotype, and the biological relevance of the data is questionable. Unlike its capability with other DNA lesions, Pol can not only extend from nucleotides inserted opposite a thymine glycol lesion but can also carry out the insertion step ( Johnson et al., 2003). dAMP is inserted preferentially, with 13% of the efficiency for dAMP insertion opposite undamaged thymine, and extension is also preferentially from this incorporated residue, with 50% of the efficiency of this process for dAMP paired with normal thymine, resulting overall in a marked capability of the enzyme for accurate replication past this lesion. Johnson and coworkers nevertheless suggest that Pol, rather than Pol, performs the insertion step because it reaches the lesion first. Pol, in
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contrast, is very inefficient at both insertion and extension with thymine glycol. Steady-state kinetic analysis with the human and yeast Rev1p was also used to examine insertion efficiencies opposite a variety of lesions, including 8-oxoG, (þ)-trans-anti benzo[a]pyrene diol epoxide –N2-dG adduct, ()-trans-anti benzo[a]pyrene diol epoxide –N2-dG, acetylaminofluorenedG, and 1, N6-ethenoadenine with the human enzyme (Zhang et al., 2002), and O6-methyl guanine and 8-oxoguanine with the yeast protein (Haracska et al., 2002). In the first of these investigations, in which human Rev1p was used, dCMP was inserted opposite a template 8-oxoG with 38% of the efficiency of insertion opposite an undamaged template guanine, and with 15% and 18% efficiency opposite the (þ) and () benzo[a]pyrene diol epoxide adducts, respectively. dCMP was preferentially inserted in each of these cases. For the acetylaminofluorene-dG adduct, the efficiency relative to template guanine was <0.1%. Interestingly, however, dCMP was inserted opposite a template ethenoadenine adduct with an efficiency of 125%, relative to a template adenine, suggesting that Rev1p activity may be an important source of mutations with this lesion. In the second study, in which yeast Rev1p was employed, insertion efficiencies opposite O6-methylguanine and 8-oxoguanine lesions were found to be 2.5% and 0.25% respectively, relative to template guanine. The marked difference between the results for the 8-oxoG lesion in the two studies may reflect differences between the human and yeast enzymes, but it could also result from the influence of other factors such as sequence context. In aggregate, these observations indicate that Pol possesses a marked facility for extension from abnormal primer ends of many kinds. They also show that Pol can, in some cases, insert nucleotides opposite damaged bases, suggesting that the role of this enzyme may not be restricted exclusively to extension. These results also confirm the strong preference of Rev1p for dCMP insertion, even though other nucleotides can also be used at much reduced efficiencies.
III. Genetic Analysis Although the enzymatic properties of Pol and Rev1p in vitro and their requirement for DNA damage induced mutagenesis in vivo strongly suggest that the proteins they encode are employed in translesion replication, this conclusion is most directly demonstrated by transformation of rev mutant strains, with plasmid constructs containing a single defined lesion at a specific location. Using budding yeast, such experiments yield estimates of the frequency of lesion bypass and, with the aid of strains deleted for genes encoding Pol, Pol, Rev1p, or other relevant proteins, the relative contributions of these enzymes to replication past various lesions.
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Sequence analysis of replicated plasmids identifies the nucleotide insertions accompanying lesion bypass. This approach has been used to investigate the roles of REV1, REV3, and REV7 in the bypass of an AAF-G adduct, a T-T dimer, a T-T (6–4) photoadduct, and an abasic site (Baynton et al., 1998, 1999; Bresson and Fuchs, 2002; Gibbs et al. 2004; Nelson et al., 2000). With the exception of the T-T dimer, the bypass of each of these lesions required the function of REV3. The function of REV7 was also found to be necessary for the bypass of AAF-G, the only lesion examined with this gene. However, because it is essential for Pol activity (Nelson et al., 1996a), a requirement for REV7 is likely to follow that of REV3 in the bypass of other types of DNA damage. These results indicate an essential role for Pol in replication past these lesions. In the sequences examined (50 -CACGAAFTTTC-30 , 50 -ATAGAAFCCC-30 ), bypass of the AAF-G adduct can occur via a misaligned (slipped) primer terminus, leading to a 1 frameshift in the first sequence or þ1 frameshift in the second, or via a correctly aligned (nonslipped) primer, resulting in accurate bypass in both cases. Interestingly, though both of these events were abolished in rev3 or rev7 strains, only bypass via the nonslipped intermediate was substantially reduced in the rev1 mutant; with the slipped intermediate, bypass frequencies were 92% or 56% of the wild-type frequency in the two sequence contexts examined (Baynton et al., 1999). The molecular basis for the latter result is unclear, particularly as in other circumstances Rev1p appears to be essential for Pol activity in vivo (Gibbs et al., 2004; Nelson et al., 2000). In addition to requiring Pol, however, replication past the AAF-G adduct was also found to employ Pol; in the first sequence, bypass frequencies for both slipped and nonslipped intermediates were reduced to a quarter of the wild-type frequency in a rad30 (Pol deficient) mutant, and in the second sequence, they were reduced to 10% of this frequency (Bresson and Fuchs, 2002). Pol, together with Pol and Rev1p, was also involved in replication past a T-T (6–4) photoadduct, though the extent of Pol involvement varied greatly in the two experiments in which it has been investigated (Bresson and Fuchs, 2002; Gibbs et al., 2004). In the Bresson and Fuchs study, mutagenic events bypassing the T-T (6–4) photoadduct, which constituted 60% of the total and resulted exclusively from the insertion of dGMP opposite the 30 T of this lesion, were reduced to 9% of the wild-type frequency in a Pol-deficient mutant, whereas the frequency of the remaining error-free bypass events was unaffected by the absence of this enzyme. In the study of Gibbs and coworkers, mutagenic bypass events with alterations at the site of the 30 T of the T-T (6–4) lesion constituted only 15% of the total and resulted from incorporation of dCMP and dTMP as well as dGMP. In the Pol -deficient mutant, the bypass frequency was 93% of that found in the wild type, and the
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frequency of dGMP insertion opposite the 30 T of the T-T (6–4) lesion was reduced from the 10% observed in the wild-type to 4% of total insertions. In addition, mutations resulting from insertions opposite the 50 T were also observed. Such results indicate that the intervention of Pol in the bypass of this lesion is highly dependent on some factor such as sequence context, which was 50 -ACAAT[6–4]TGAAC-30 in the Bresson and Fuchs experiment, and 50 -GCAAGT[6–4]TGGAG-30 in the work of Gibbs et al. In addition to these investigations, in which yeast strains were transformed with plasmids that carry a specifically located lesion, similar experiments have been carried out using transformation with lesioncontaining oligonucleotides (Otsuka et al., 2000, 2002a,b,c, 2004). This interesting and innovative approach is based on the observation (Moerschell et al., 1988) that yeast strains carrying a cyc1-31 mutation can be reverted to wild-type by transformation with oligonucleotides that have the CYC1þ sequence spanning the mutant site. The cycl-31 mutation carries a base substitution and adjacent single-nucleotide deletion generating a stop codon, which, in the absence of transformation, reverts at only a very low frequency. Although it is not known how the sequence present in the single-stranded oligonucleotide is integrated into the yeast chromosome, it presumably requires invasion of the genomic DNA and some kind of recombination or gene conversion event. This experimental method therefore either achieves or closely approaches the ideal of placing a single defined lesion at a specified genomic location. Transformation frequencies using oligonucleotides that carried a site-specific T-T (6–4) photoadduct were only 3% of the transformation frequencies with their damage-free counterpart (Otsuka et al., 2002a), a value close to those observed in the plasmid experiments; all experiments therefore indicate that this lesion strongly blocks replication. Seventy-seven percent of the transformants carried mutations at the lesion site, 81% of which were 30 T ! C substitutions. The frequency of these mutations was much reduced in a rad30 strain, but the bypass frequency, though lower, was not reduced proportionately because of a partially compensating increase in bypass events resulting from the insertion of other nucleotides (Otsuka et al., 2004). No transformants were found in a rev1 strain, indicating that Rev1p is essential for the bypass of this lesion. This reflects a requirement for the second function of Rev1p, rather than its deoxycytidyl transferase activity, because bypass was just as efficient in a rev1 mutant strain producing protein that lacks transferase activity by virtue of D467A, E468A substitutions (Otsuka et al., 2004). Oligonucleotide transformation was also used to investigate the genetic requirements and mutagenic outcome of replication past either a natural (deoxyribose) abasic site or its synthetic tetrahydrofuran analog (Otsuka et al., 2002b). A series of isogenic strains,
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comprising a wild-type and derivatives carrying deletions of APN1 or APN2 (both encoding apurinic endonucleases), RAD30, or REV1, or both APN1 and REV1, were transformed with oligonucleotides containing one or the other of these lesions or a deoxyuracil residue. Results from oligonucleotides carrying the latter base or the natural abasic site were essentially identical, indicating that endogenous uracil-DNA-glycosylase rapidly and efficiently removes the uracil before replication. Replication past the natural abasic site was unchanged in the rad30 strain but was much reduced in the rev1 mutant, indicating that Rev1p, though not Pol, is necessary for the bypass of this lesion. In REV1þ strains, dCMP was preferentially incorporated opposite the abasic site, with an average of 63% insertions being of this kind. This class of event was much reduced in rev1 strains, however, showing that it results from the deoxycytidyl transferase activity of Rev1p. Although this activity stimulates replication past the abasic site, a reduced frequency of bypass can still occur in strains that contain Rev1 mutant protein lacking transferase activity because of D467A and E468A substitutions (K. Negishi, personal communication; Otsuka et al., 2002c). Bypass of the abasic site in this strain occurs at about half the frequency found in the wild type, a reduction that is accounted for by the marked decrease in bypass events resulting from dCMP insertion. In the absence of the transferase activity, the principal residue incorporated was dTMP, though low frequencies of insertion of the other nucleotides, including dCMP, were observed. These insertions were presumably carried out by either Pol or Pol, both of which are no doubt less efficient at this task than wild-type Rev1p. However, the second function of Rev1p is also needed, as shown by the absence of bypass in the rev1 strain. This function is probably a requirement of Rev1p for Pol activity, perhaps as an intermediary to anchor it to PCNA (see Section V). In the presence of wild-type Rev1, Pol, also essential for the bypass of abasic sites, can then extend the primer to complete lesion bypass. Bypass frequencies for the tetrahydrofuran residue were substantially lower than for the natural abasic site in REV1þ strains, except in the apn1 strain, indicating that the synthetic analog is a better substrate for the APN1 apurinic endonuclease than the natural lesion. Deletion of APN1, though not APN2, also increased the transformation efficiencies of oligonucleotides carrying the natural abasic site but did not alter bypass frequencies in the mutants relative to those in the wild-type control. As with the natural abasic site, bypass of the tetrahydrofuran residue in the apn1 strain also requires Rev1p, as bypass is essentially abolished in the apn1 rev1 double mutant. However, the Rev1p transferase activity appears to be much less important with this lesion, indicating that the tetrahydrofuran abasic site is a relatively poor substrate for dCMP insertion; 48% of the bypass events
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result from the incorporation of dAMP, 32% from dGMP incorporation, and only 20% from dCMP incorporation. No incorporation of dTMP was detected. Although insertion of dCMP is much less frequent opposite the tetrahydrofuran lesion than the natural abasic site, it is still dependent on the Rev1p deoxycytidyl transferase activity, as such insertion is absent in an apn1 mutant that possesses the D467A, E468A Rev1 mutant protein. Despite this, the bypass frequency is almost unaffected by this mutant protein (Otsuka et al., 2004). The role of Rev1p in the bypass of the tetrahydrofuran lesion therefore appears to depend principally on its second, Pol-supporting, function.
IV. Processes Other than General Translesion Replication that Employ Pol and Rev1p Although most attention has been given to the involvement of Pol and Rev1p in translesion replication in the genome at large, these enzymes also appear to function in other cellular processes, most notably in the repair of double-strand breaks and, among vertebrates, in somatic hypermutation.
A. Somatic Hypermutation Somatic hypermutation is the phenomenon in which a high frequency of point mutations are generated within a 1–2-kb segment in the variable region of expressed immunoglobulin genes in response to the presence of an antigen. High-affinity immunoglobulins are generated by selection among variants generated over about 20 rounds of replication, in each of which the region specific mutation frequency is about one mutant per kilobase, resulting in up to 2% nucleotide substitutions within the target region overall (Diaz and Storb, 2003). These mutations are produced by inaccurate DNA polymerases, including pol and Rev1p, that bypass damage within the target region initiated by the activation induced cytidine deaminase (Muramatsu et al., 2000), which, following the conversion of cytosine to uracil and the action of uracil-DNA-glycosylase (Di Noia and Neuberger, 2002; Rada et al., 2002), results in the production of abasic sites. Although at least some of the GC ! AT mutations are likely to arise during replication past uracil by accurate enzymes (Di Noia and Neuberger, 2002), the remaining substitutions presumably arise from bypass by inaccurate bypass polymerases. Pol appears to play a major role in the latter process. Somatic hypermutation frequencies within VHDJH and bcl-6 genes in a human B-cell line were reduced to about 27% of control frequencies
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following treatment with phosphorothioate-modified 15-mer oligonucleotides that substantially reduced REV3 mRNA levels (Zan et al., 2001). A similar result was found for 6-thioguanine resistant mutants induced in a UV-irradiated pSP189 shuttle vector transfected into the oligonucleotidetreated cells (Zan et al., 2001). Interestingly, Pol (REV3) transcript levels were found to be upregulated, and Pol mRNA levels downregulated, in cells undergoing somatic hypermutation, whereas the transcript levels for Pol, Pol, Pol, Pol, Pol", Pol, and Pol all remained unchanged. The involvement of Pol in somatic hypermutation is also evident in experiments with transgenic mice expressing high levels of antisense RNA to mouse REV3, which concomitantly reduced levels of endogenous REV3 mRNA transcript (Diaz et al., 2001). Mutation frequencies in memory B-cell clones in these mice were between 56% and 72% of the frequencies in cells from control animals, with a marked deficiency of clones that contained more than three mutations. Such figures are likely to appreciably underestimate the role of Pol in this process. Few transgene mice were recovered, and only 1 of 10 expressed high levels of antisense RNA, indicating selection for animals expressing at least a minimal amount of Pol, a conclusion also predicted from the embryonic lethality of mice homozygous for null disruptions (Bemark et al., 2000; Esposito et al., 2000; Wittschieben et al., 2000). As expected from its requirement for Pol activity, Rev1p also appears to be necessary for somatic hypermutation in DT-40 chicken cells (Simpson and Sale, 2003). Disruption of the REV1 gene in this cell line reduces nontemplated mutations, resulting from de novo mutation, to 0.8 104 mutations per base pair, compared to a frequency of 5.5 104 mutations per base pair in the REV1þ control. Almost all untemplated mutations are therefore generated by a Rev1pdependent process, though it cannot be excluded that a small fraction arose in a Rev1p-independent manner, as the calculated background error frequency from the PCR procedure used was 0.4 104 mutations per base pair. The frequency of templated changes, resulting from gene conversion, was not decreased. Disruption of REV1 reduced the frequency of all types of base substitutions, consistent with loss of the Pol-supporting function of Rev1p. However, DNA sequence results from the REV1þ cell line (Sale et al., 2001) are also consistent with the frequent employment of the Rev1p deoxycytidyl transferase activity during somatic hypermutation. In this clone, 51% of the mutations were C ! G or G ! C mutations, as expected from the sequential action of the AID cytidine deaminase and uracil-DNA-glycosylase, followed by dCMP incorporation opposite the resulting abasic site. The remaining substitutions presumably arise from insertions by other DNA polymerases, such as Pol, Pol, Pol, or Pol; however, chickens do not appear to possess Pol ( J. McDonald and
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R. Woodgate, personal communication). All of these polymerases may be employed in somatic hypermutation to some degree in mammals. Data from in vitro experiments suggest that Pol has a marked facility for insertion opposite abasic sites (Vaisman et al., 2002), and a variety of evidence indicates that Pol is employed in somatic hypermutation at AT base pairs and certain hotspots (Rogozin et al., 2001; Zeng et al., 2001). An extended discussion of somatic hypermutation is given in Chapter 11 of this book.
B. Double-Strand Break Repair Evidence for the use of Pol during the repair of double-strand breaks has been found in budding yeast, DT-40 chicken cells, and the mouse, though it appears to be essential for such repair only in vertebrates and is merely associated with it in yeast. In a REV3þ diploid strain of the latter organism, repair of a specifically located double-strand break in one of the chromosome III homologs was accompanied by a 56- to 76-fold increase in the spontaneous reversion rate of the trp1-488 nonsense allele located 314 base pairs from the break site. In a rev3 mutant, however, trp1-488 reversion frequency in cells repairing the double-strand break was only 1%–2% of the wild-type value, even though the induced recombination frequency was about the same as in the REV3 strain (Holbeck and Strathern, 1997). The reversion rates of two trp1 frameshift alleles, even though much increased in cells repairing the double-strand break, were not dependent on REV3, however. The types of REV3-dependent mutations induced by double-strand break repair was further examined in a subsequent investigation (Rattray et al., 2002), using forward mutations to canavanine resistance to monitor mutagenesis rather than reversion of trp1 alleles. As before, double-strand breaks were induced by inserting an HO-endonuclease recognition sequence within one of the chromosome III homologs, together with regulated expression of the endonuclease from an HO gene controlled by the GAL1 galactose-inducible promoter. The frequency of base-pair substitutions induced in break repair by homologous recombination was about ninefold lower in the rev3 mutant than REV3þ strain, whereas for frame-shift mutants the decrease was only about twofold, indicating that half of the latter events are produced by a DNA polymerase other than Pol. Interestingly, the frequency of can1R mutations associated with break repair by what appeared to be nonhomologous end joining rather than homologous recombination were mostly produced by Pol, with a more than 50-fold decrease in their frequency in the rev3 mutant compared to the REV3þ strain. This result suggests that Pol may be employed in a greater variety of double-strand break
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repair events than just homologous recombination. It would be interesting to know whether Rev1p is also required for recombination-associated mutagenesis in yeast. A variety of evidence suggests that Pol, as well as participating in translesion replication, may be directly involved in double-strand break repair in DT-40 chicken cells, rather than just associated with it (Sonoda et al., 2003). In these cells, a REV3/ homozygote carrying disrupted alleles is viable but suffers a threefold higher frequency of spontaneously occurring chromosome aberrations than its isogenic REV3þ/þ counterpart. The REV3/ clone also exhibits higher frequencies of chromosome aberrations induced by ionizing and UV radiations. Although, as expected from their deficiency in translesion replication, REV3/ mutant cells are much more sensitive than the wild-type to ionizing radiation in G1-early S phase, they are also more sensitive in the G2 phase and, indeed, are at least as sensitive in this phase as a DT-40 RAD54/ clone deficient in homologous recombination (Bezzubova et al., 1997). Moreover, DT-40 REV3/ mutant cells are substantially more sensitive to ionizing radiation than RAD18/ mutants, and a little more sensitive to UV, unlike in yeast where rev3 mutants are much less sensitive to UV, and a little less sensitive to ionizing radiation, than rad18 mutants (Lawrence and Christensen, 1976; McKee and Lawrence, 1979a,b). Because Rad18p, together with Rad6p, regulates translesion replication by Pol in yeast (Bailly et al., 1994), the greater sensitivity of REV3/ relative to RAD18/ strains in the DT-40 cells implies that Pol is employed additionally in processes other than translesion replication in chickens. This additional process is likely to be repair of double-strand breaks by homologous recombination because gene targeting, which depends on this measure, is much reduced in the REV3/ clone, whereas nonhomologous end joining appears to occur normally (Sonoda et al., 2003). A DT-40 REV3/ clone has a very similar profile of sensitivities to different DNA damaging agents as the REV3/ mutant (Simpson and Sale, 2003), suggesting that the Rev3 and Rev1 proteins act together in this process, as they do in translesion replication in yeast. Similarities between the consequences of REV3/ disruptions in the mouse and DT-40 cells suggest that mouse Pol may also be employed in double-strand break repair. Such disruptions cause embryonic lethality (Bemark et al., 2000; Esposito et al., 2000; Wittschieben et al., 2000), with embryonic cells showing high frequencies of apoptosis that cannot be relieved by a disruption of p53, which also failed to rescue embryos from the lethality (van Sloun et al., 2002). Cytological examination of cells from 11.5-day embryos showed that disruption of REV3 led to a marked increase in double-strand breaks and other chromosome aberrations, with 14% of
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the cells containing these compared to 0.7% in cells from the REV3þ/þ or REV3þ/ control clone, a result reminiscent of those from chicken DT-40 cells.
V. Regulation of Pol and Rev1p and Interactions with Other Proteins The regulation of Pol and Rev1p, together with the other bypass polymerases, is poorly understood, but it presumably addresses several problems such enzymes pose for the cell. Of these problems, a major issue is the need to recruit the appropriate enzyme, or sequence of enzymes, to a replication fork stalled at a particular lesion, a problem that is particularly acute in mammals because of the number of polymerases they potentially can employ. Another important problem is the necessity of curtailing the activity of the enzymes in circumstances where they are not required, to minimize the production of mutations. A variety of evidence indicates that PCNA and Rev1p are important elements in the recruiting problem and that specific recruiting together with tight regulation of the amount of protein expressed and its activity is part of the means used to limit unwanted activity in lesion-free DNA. There is, however, little evidence in yeast for regulation of REV3 at the level of transcription, either during the mitotic cell cycle or in response to DNA damage, where UV irradiation led at best to only a small increase in transcript level (Singhal et al., 1992). REV1 also appears to possess a promoter that is unresponsive to such damage (Larimer et al., 1989). There was, however, a marked increase in REV3 transcript in meiosis, with a maximal level that was 18fold above the premeiotic level, perhaps in response to a need for Pol in the repair of the double-strand breaks responsible for homologous recombination in meiotic cells. The high levels of human (Lin et al., 1999) and mouse (Van Sloun et al., 1999) REV3 transcript in testis and ovary may reflect the same phenomenon. Although REV3 transcript levels are only marginally increased in UV-irradiated cells of budding yeast, more substantial responses to such treatment were observed in Neurospora (Sakai et al., 2002, 2003). No transcripts for upr-1 (encoding Neurospora REV3), ncrev1, or ncrev7 were detected in unirradiated cells, though they were easily observed after 15–30 minutes in wild-type cells irradiated with 100 J/ m2 UV. An increase of mouse REV3 transcript in response to stress was also suggested by the recovery of a partial REV3 clone from a differential screen of a cDNA library isolated from mouse fetal cortical cells treated with the drug pentylenetetrazol, which induces epileptic-like seizures in mice (Kajiwara et al., 1996).
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Rev3 protein levels appear to be very low in both yeast and mammals, indicating that this may be common to all organisms. Low levels of Rev3p in yeast appear to result, at least in part, from the presence of an outof-frame ATG codon, in a good context for translation (Kozak, 1986), 10 nucleotides before the open-reading frame ATG, a feature that is expected to reduce the translation efficiency of the REV3 reading frame by a factor of at least a hundred. Out-of-frame ATG codons in good contexts are also present in the 50 untranslated region of the human REV3 and REV1 genes (Gibbs et al., 1998, 2000). Moreover, 40%–50% of the human and mouse cDNAs analyzed contained an insertion of 128 bp between nucleotides þ139 and þ140 of the REV3 reading frame, the result of alternate splicing, which introduces an immediate in-frame stop codon (Gibbs et al., 1998; Murakumo et al., 2000; Van Sloun et al., 1999), reducing the proportion of translatable transcript. These features, and perhaps others, appear to curtail production of Rev3p and Rev1p to very low levels, presumably to limit their inappropriate synthesis on undamaged templates. This end is also likely to be served by mechanisms that target Pol and other polymerases to the sites of blocked forks. As discussed at greater length in Chapter 10, this is believed to be initiated in yeast, humans, and perhaps all eukaryotes by modification of lysine 164 in PCNA (Hoege et al., 2002; Stelter and Ulrich, 2003). Modification at this site acts as a molecular mechanism to switch cells between three different modes of replication; translesion replication, replication dependent on the error-free damage tolerance process, and normal replication. Addition of ubiquitin to lysine 164 by the Rad6p E2 ubiquitin conjugase is thought to promote translesion replication by Pol, Rev1p, Pol, and, presumably, other Y-family enzymes. The error-free DNA damage tolerance process, on the other hand, is promoted by polyubiquitination at lysine 164, in which ubiquitin is conjugated to the lysine 63 residue of ubiquitin itself, using the ubiquitin conjugase activity of the Ubc13p/Mms2p/Rad5p complex. The addition of SUMO to lysine 164 in PCNA appears to promote normal replication. The way these modifications of PCNA implement the recruitment of the various DNA polymerases or other proteins is not known, but it presumably depends, either directly or indirectly, on modification-specific association with the polymerases or some intermediate protein. An investigation of the yeast rev6-1 mutant indicates that Rev1p may perform such an intermediary function. The rev6-1 mutant was isolated in a screen for strains deficient for UV-induced reversion of the his4-38 frameshift allele (Lawrence et al., 1985c). Initial characterization showed that rev6-1 strains were substantially deficient in the UV-induced reversion of the ochre allele arg4-17 and the missense allele ilv1-92, as well as of his4-38, and they were also more sensitive to UV than other rev
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mutants. Rates of spontaneous reversion of ilv1-92 were also found to be 10-fold higher in rev6-1 mutants than in the wild-type. Recent studies (H. Zhang, P. E. M. Gibbs, and C. W. Lawrence, unpublished data) show that rev6-1 mutants are incapable of replicating past a site-specific T-T cyclobutane dimer or abasic site, and that they are also deficient for the RAD6—dependent error-free tolerance process, but grow at normal rates. These studies also found that the rev6-1 mutation is an allele of POL30, encoding the sliding clamp protein PCNA, which results in a G178S substitution in this protein, which is located in a -sheet at the interface between monomers in this homotrimeric structure. When mapped onto this structure, the serine side chain at this site appears to cause an unfavorable steric interaction with tyrosine 114 in the adjacent monomer, indicating that clamp structure may be slightly abnormal, though clearly not sufficiently to interfere with normal replication. Because the rev6-1 phenotype appears to indicate loss of all damagetolerance processes, the G178S substitution may prohibit conjugation of ubiquitin at lysine 164, preventing recruitment of polymerases or other proteins that promote their association with PCNA. Results of a two-hybrid analysis (H. Zhang, P. E. M. Gibbs, and C. W. Lawrence, unpublished data) indicate that Rev1p may be a protein of this kind. These data indicate that Rev1p associates with wild-type PCNA, but not with the G178S mutant protein. In addition, no association was observed between wild-type PCNA and Rev1p containing a G193R substitution in the N-terminal BRCT domain of this protein, a region believed to be concerned with protein– protein interactions. The rev1-1 mutant expressing the G193R protein retains a substantial fraction of its deoxycytidyl transferase activity but is substantially impaired with respect to replication past a site-specific T-T (6-4) photoadduct or abasic site (Nelson et al., 2000) and with respect to induced mutagenesis in general (Lawrence and Christensen, 1978; Lemontt, 1971, 1972; McKee and Lawrence, 1979a). The marked similarity between the REV1 and REV3 mutant phenotypes, and in particular the observation that deletion of either gene can almost interchangeably abolish lesion bypass, indicate that Rev1p is required for Pol activity in vivo, which is probably the function lost in the rev1-1 mutant. It is therefore possible that in binding to PCNA, Rev1p recruits Pol to a replication fork stalled at the site of a lesion. It is also possible that Rev3p alone, rather than Rev3p/Rev7p, is bound to Rev1p in this circumstance, an association that might enhance Pol activity. Because Rev1p is not employed in the error-free process of DNA damage tolerance, the G178S substitution in PCNA may also abrogate an association with proteins other than Rev1p. In addition to regulation by means of PCNA modification, budding yeast cells possess what appears to be a regulatory cascade initiated by a
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DNA damage-sensing mechanism dependent on the checkpoint genes RAD9, RAD17, RAD24, and MEC3. In an excision-deficient background, deletion of any one of these genes greatly reduces UV-induced forward mutation to canavanine resistance, down to a frequency characteristic of rev mutants (Paulovitch et al., 1998), an observation that suggests the existence of a signaling pathway initiated by unrepaired DNA damage that leads to the activation or mobilization of Pol and Rev1p. Induced mutation frequencies were reduced, but to a lesser extent, in an excision proficient background. Associations among Revp, or between them and other proteins, also suggest the possibility of some forms of regulatory control, although the mechanisms by which such control might be accomplished are not yet known. Because yeast Pol activity in vitro is strongly dependent on the binding of Rev7p to Rev3p (Nelson et al., 1996a), and human Rev7p has been shown to associate with hRev3p (Murakumo et al., 2000), the Rev7p subunit of Pol may perform some regulatory function. Moreover, hRev7p associates with at least two other proteins, hRev1p and hMad2p. hRev7 binds to a site at the C terminus of hRev1p ( Murakumo, 2002; Murakumo et al., 2001), producing a stable complex but producing no change in the deoxycytidyl transferase activity (Masuda et al., 2003). hRev7p also binds to the spindle assembly checkpoint protein hMad2, with which it shares significant homology, and such an association appears to suggest regulation tied to the cell cycle (Murakumo et al., 2001). Mouse Pol, Pol, and Pol, each bind to mouse Revp1 at the same site as Rev7p, though, unlike the association of Rev7p and Rev3p, their enzymatic activity appears to be unchanged (Guo et al., 2003). Similar results have been observed with the human counterparts of these genes (Ohashi et al., 2004). Competitive binding to Rev1p therefore appears to play some important regulatory role in mammals, though how this is effected is not known. As judged by sequence comparisons, yeast Rev1p appears not to have such a binding site, though a direct test for association between Rev1p and Pol has not yet been reported. Last, two-hybrid screens of a human library, using different segments of hRev3p as bait, have uncovered an association of the tumorsuppressor protein p33ING, encoded by the ING1 gene, with the amino terminus of hRev3p (R. Murante, O. Uluckan, and C.W. Lawrence, unpublished data). This protein has been shown to suppress growth and apoptosis, promote repair of UV damage to DNA, associate with p53, and participate in the p53 signaling pathway. Counterparts of this protein in yeast interact with histone acetyltransferase complexes and may therefore modulate transcription (Cheung and Li, 2001; Cheung et al., 2001; Garkavtsev et al., 1998; Loewith et al., 2000). Although this association appears to indicate integration of Pol in a network regulating checkpoints, growth control, and other
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aspects of responses to DNA damage, much remains to be learned about the way this might be implemented.
VI. Conclusions and Speculations As discussed above, there is a large amount of experimental evidence showing that Pol and Rev1p perform important functions in replication past a great variety of DNA lesions. There are, however, disagreements about the particular nature of the functions of both of these enzymes. According to one model ( Johnson et al., 2000, 2001; Prakash and Prakash, 2002), the function of Pol is exclusively to extend from insertions opposite lesions, whether of the correct or incorrect nucleotide, whereas according to another model (Lawrence, 2002; Lawrence et al., 2000), Pol can perform both insertion and extension activities with some lesions. The ability of this enzyme to extend from terminal mismatches ( Johnson et al., 2000; Lawrence and Hinkle, 1996; Lawrence et al., 2000), and from nucleotides incorporated opposite lesions by other polymerases (Guo et al., 2001; Haracska et al., 2003; Johnson et al., 2003; Nelson et al., 1996b), has been clearly demonstrated, but its capabilities for insertion at lesion sites are not well established. The conclusion that Pol cannot perform the latter function ( Johnson et al., 2000, 2001; Prakash and Prakash, 2002) is based on in vitro assays using the two-subunit enzyme and primed oligonucleotide templates that carry defined lesions, most of which indicate that Pol inserts only inefficiently opposite the lesions. The extent to which these results with the two-subunit enzyme reflect the in vivo activity of the polymerase is still uncertain, however. It is not known, for example, whether functional association with Rev1p, thought to occur in cells, might alter Pol activity. In addition, incorporation opposite lesions by Pol has been thought to be unlikely because it is believed to be a highly accurate enzyme, having greater fidelity than Pol and nearly the same fidelity as Pol ( Johnson et al., 2000). This conclusion was reached by comparing error rates in the LacZ gene in M13mp2 replicated by purified calf thymus Pol or Hela cell Pol (Thomas et al., 1991) with those from a steady-state kinetic analysis of the fidelity of nucleotide incorporation on lesion-free templates by Pol ( Johnson et al., 2000). Because the experimental procedures were very different, the validity of comparing these two sets of data is open to question. But, more importantly, the data in Thomas et al. (1991) do not appear to support the conclusion reached by Johnson and coworkers: Pol is a little less accurate than Pol, and much less accurate than Pol. The fidelity of Pol is greater than the fidelity of Pol by twofold or more in 7 of the 12 template base/incoming mispaired nucleotide combinations studied, and it is approximately the
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same in the remaining 5. The fidelity of Pol is at least 10-fold greater than the fidelity of Pol overall and in the same set of comparisons, by twofold to >700-fold; Pol does not, by these measures, appear to be an unusually accurate polymerase, and it may in fact be the least accurate of the B-family enzymes. Further support for the conclusion that Pol is able to incorporate nucleotides opposite lesions is provided by the occurrence of Polindependent replication past various site-specific lesions in yeast. For example, although the proportion of Pol-independent events that bypass a T-T (6-4) photoadduct varies in different experiments (Bresson and Fuchs, 2002; Gibbs et al., 2004; Otsuka et al., 2002a, 2004), an appreciable fraction of this kind is nevertheless found in each case, and in the work of Gibbs et al. (2004) this fraction was over 90%. In this circumstance, which polymerase might be responsible for nucleotide insertion opposite the 30 T of the lesion? Although Pol cannot be formally excluded, the most likely candidate is Pol. As an enzyme concerned with lesion bypass, Pol is much better adapted than a replicase to cope with distorted templates, a property that is demonstrated in the extension reactions it carries out. Pol is also a better candidate because, unlike Pol, it lacks a 30 to 50 exonuclease proofreading function, which could inhibit nucleotide insertion. Such an effect is seen with E. coli DNA polymerase III. In cells lacking all other DNA polymerases, this enzyme can bypass a T-T cyclobutane dimer, but only if proofreading is disabled by the mutD5 mutation (Vanderwiele et al., 1998). Pol is also a much more suitable enzyme for incorporation because, as discussed above, it is a much less accurate than Pol, again consistent with a relaxed requirement for normal template structure. For the same reasons, Pol is a good candidate for insertion opposite the 30 T of a T-T cyclobutane dimer in Pol-deficient cells, which results in an error rate of 9%, an error rate atypical of a replicase (Gibbs et al., 2004). Pol is also likely to be responsible for the bypass of abasic sites that is Pol and Rev1p independent (Gibbs et al., 2004). A second major difference of interpretation concerns the question of whether the Rev1p deoxycytidyl transferase activity plays any part in the bypass of an abasic site in yeast. Although investigations using plasmids and oligonucleotides carrying a specifically located natural abasic site all show that bypass of this lesion in yeast is chiefly accomplished by Rev1pdependent insertion of dCMP (Gibbs et al., 2004; Nelson et al., 2000; Otsuka et al., 2002b) Haracska and colleagues (2001c) have concluded that such bypass depends principally on the incorporation of dAMP by Pol, followed by extension from this terminus by Pol. In addition to the relative efficiencies of the Pol/Pol and Pol/Rev1p combinations for bypass of an abasic site in vitro (see section IIB), evidence cited by these
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authors to support this model includes results from MMS-induced mutagenesis in an apn1 apn2 mutant, a critique of plasmid-based methods, and inference from the phenotype of the pol32 mutant. The interpretation of each of these pieces of evidence is open to question, however, and the case made does not seem compelling. Evidence interpreted as showing preferential insertion of dAMP, and not dCMP, opposite abasic sites in the genome was obtained by treating an apn1 apn2 yeast strain with MMS, followed by sequence analysis of the forward mutations to canavanine resistance (canR), which this treatment induced. Based on the spectrum of base-pair substitutions induced, and assuming that these resulted from abasic residues at sites of template guanine or adenine, it was calculated that 64% of the mutations resulted from dAMP insertion, with lesser and approximately equal frequencies of insertions of the remaining nucleotides. A basic problem with this procedure is that it is incapable of providing unbiased estimates of nucleotide insertion frequencies because it only detects mutagenic events and cannot, unlike experiments with site-specific abasic sites, estimate the frequencies of insertions that restore the correct sequence. In particular, it cannot detect the insertion of dCMP opposite the site of a lesion derived from a template guanine, the predominant event expected with Rev1p. This is likely to have resulted in a serious underestimation of dCMP insertion in the experiment of Haracska and coworkers (2001c) because 79% (22/28) of the base substitutions sequenced resulted from alterations at GC sites, implying that the majority of abasic sites arose from the loss of a guanine base. At the same time, it is unclear how many of the mutations were in fact induced by abasic residues, rather than by methylation adducts or other kinds of DNA damage. Abasic sites are likely to be highly labile in the absence of the protection provided by bound AP endonucleases (Mol et al., 2000), and they probably undergo a spontaneous -elimination reaction, resulting in a nicked strand that is a substrate for repair rather than replication, providing a greater opportunity for mutagenesis by alkylated bases that have escaped repair. As a further argument against preferential incorporation of dCMP opposite abasic sites in yeast, Haracska and coworkers cite results from another plasmid-based experiment (Kunz et al., 1994) that appeared to indicate preferential insertion of dGMP. A reevaluation of these data in the light of later work indicates, however, that the results may not in fact be inconsistent with dCMP insertion, again because procedures that monitor insertion events by selecting mutations, as used in this experiment, fail to detect events that restore the wild-type sequence. At the same time, the identity of mutagenic lesions is not well defined in these experiments, unlike those using the site-specific techniques. In the experiment of Kunz
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and coworkers, the basis for the mutator phenotype in an apn1 deletion mutant of yeast was investigated by sequence analysis of spontaneous mutations that inactivate the SUP4-o tRNA suppressor, carried on a plasmid. Because spontaneous mutation was much reduced in an apn1 mag1 strain, such mutations were thought to have arisen at abasic sites produced by the action of N3-methyladenine (Mag1) glycosylase on endogenously alkylated adenine. Because the largest increase of spontaneous mutations in the apn1 mutant were AT ! CG substitutions, most of the mutations were consequently interpreted as having arisen from the insertion of dGMP, rather than dCMP opposite abasic sites generated in this way. However, the work of Guillet and Boiteux (2003) indicates that a high proportion of the abasic sites may have arisen not by removal of adenine but by removal of deoxyuracil that was inserted opposite template adenine in the apn1 strain, indicating that dCMP, rather than dGMP, incorporation was indeed the major event, exactly as found with the site-specific abasic residue. Guillet and Boiteux observed that the lethality of an apn1 apn2 rad1 yeast strain can be relieved by an ung1 mutation, deficient in uracil-DNA-glycosylase, or by overexpression of DUT1, encoding the dUTP pyrophosphatase, but not by a deletion of MAG1, OGG1, NTG1, or NTG2. This result indicates that many of the highly toxic abasic sites in an apn1 strain arise from incorporation of dUTP opposite a template adenine in the genome, followed by its removal by uracil-DNA-glycosylase. Moreover, abasic sites may have also arisen at abasic sites generated by removal of template guanine. In these cases, insertion of dCMP cannot be detected by the SUP4-o system, because it restores the normal base-pair rather than generating a mutation. Removal of guanine is predicted because N3 methyladenine glycosylases, of the kind encoded by yeast MAG1, discriminate relatively poorly between alkylated and normal bases, principally releasing guanine from nonalkylated DNA, though also releasing adenine and pyrimidines at fivefold and 20-fold lower frequencies, respectively (Berdal et al., 1998). As a consequence, overproduction of Mag1 leads to a strong mutator phenotype (Glassner et al., 1998), and some guanine release is likely even when Mag1 expression is normal. It is far from clear, therefore, that the results of Kunz et al. (1994) are in fact different from those using plasmids carrying a site-specific abasic residue. Lastly, Haracska and coworkers implicate Pol in the bypass of abasic sites because deletion of POL32, which encodes a nonessential subunit of this enzyme, essentially abolishes the induction of canR mutations by MMS. This observation is unlikely to indicate the involvement of the catalytic function of Pol in the bypass of abasic sites, however, because such a function is by necessity present in the pol32 strain. More probably, the loss of the Pol32 subunit has an indirect influence on translesion
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replication, dependent on its association with the Srs2 helicase, as detected by two-hybrid analysis (Huang et al., 2000). Srs2 disrupts the Rad51 nucleoprotein filament (Krejci et al., 2003; Veaute et al., 2003), thereby preventing homologous recombination repair and presumably maintaining the blocked fork structures as substrates for RAD6-dependent DNA damage tolerance activities. In the absence of Srs2, blocked forks are diverted into the Rad52-dependent recombination repair pathway (Schiestl et al., 1990), preventing processing by the Rad6 pathway, and hence the generation of mutations. It is probably for this reason that the frequency of induced mutations is reduced in both srs2 and pol32 mutants (Aboussekhra et al., 1989, 1992). Although incorporation of dCMP by Rev1p appears to be the major event during bypass of an abasic site in yeast, such a preference is less common in vertebrates, perhaps because a greater variety of DNA polymerases may participate in this event. Of these, Pol in particular has a marked facility for insertion opposite this lesion (Vaisman et al., 2002), and the efficiency of this enzyme may out-compete that of Rev1p. In some cases (Avkin et al., 2002; Takeshita and Eisenberg, 1994), the absence of a dCMP preference may well result from the use of synthetic analogs of natural abasic sites; as shown by Otsuka and coworkers (2002b), such analogs are poor substrates for the deoxycytidyl transferase activity of Rev1p. Nevertheless, preferential incorporation of dCMP is not necessarily observed in mammalian cells even when a natural abasic site is used (Cabral-Neto et al., 1994). The maintenance throughout evolution of the Rev1p transferase activity in mammalian cells, even though it does not appear to be employed, is puzzling: perhaps altered levels of expression of the various Y-family enzymes, uncharacteristic of the levels in the animal, occurred during establishment of the cell lines, an issue for further investigation. However, preferential incorporation of dCMP opposite abasic sites is found in at least one vertebrate system; namely, the immunoglobulin genes of DT-40 chicken cells (Sale et al., 2001; see Section IV.A), in which natural abasic sites are generated by the sequential action of the activation-induced cytidine deaminase and uracil-DNA-glycosylase. Although it is perplexing that mammalian cells can maintain a deoxcytidyl transferase activity, but not apparently use it, it is also puzzling why eukaryotes should possess such an activity in the first place. Why do enzymes in the Rev1 branch of the Y family only use dCTP, and not all four dNTPs, as do members of the other branches of this family, from which the Rev1p branch was presumably evolutionarily derived? One possible explanation is that restricting incorporation to dCMP, although inevitably causing mutations at abasic sites previously occupied by nucleotides other than guanine, nevertheless still results in a lower mutation
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frequency than would occur from insertion by DNA polymerases that use all four dNTPs. It will be interesting to identify the particular residues and structural features of Rev1p that exclude incorporation of dNTPs other than dCTP, structures that are presumably absent in the members of the other branches of the Y family. Finally, perhaps the greatest problem with enzymatic studies of Pol and Rev1p concerns the properties and structure of the native enzymes. As in the case of Pol, for which a considerable enhancement of insertion efficiency was observed following association with PCNA and the presence of RPA (Haracska et al., 2001b,d), the properties of Pol and Rev1p may also be enhanced by such factors. In particular, the apparent inefficiency of Pol for insertion opposite lesions in vitro may reflect their absence. A variety of evidence suggests that Pol and Rev1p are associated with one another in a multiprotein complex of some kind, possibly because Rev1p acts as a structural link between Pol and PCNA (see Section V). Identifying and investigating the enzymatic properties of such a complex is likely to be needed for a fuller characterization of these proteins.
Acknowledgments This work was supported by Public Health Service grant GM60652 from the National Institutes of Health.
References Aboussekhra, A., Chanet, R., Zgaga, Z., Cassier-Chauvat, C., Heude, M., and Fabre, F. (1989). RADH, a gene of Saccharomyces cerevisiae encoding a putative DNA helicase involved in DNA repair. Characteristics of radH mutants and sequence of the gene. Nucleic Acids Res. 18, 7211–7219. Aboussekhra, A., Chanet, R., Adjiri, A., and Fabre, F. (1992). Semidominant suppressors of Srs2 helicase mutations of Saccharomyces cerevisiae map in the RAD51 gene, whose sequence predicts a protein with similarities to procaryotic RecA proteins. Mol. Cell. Biol. 12, 3224–3234. Avkin, S., Adar, S., and Livneh, Z. (2002). Quantitative measurement of translesion replication in human cells: Evidence for bypass of an abasic site by a replicative DNA polymerase. Proc. Natl. Acad. Sci. USA 99, 3764–3769. Bailly, V., Lamb, J., Sung, P., Prakash, S., and Prakash, L. (1994). Specific complex formation between yeast RAD6 and RAD18 proteins: A potential mechanism for targeting RAD6 ubiquitin-conjugating activity to DNA damage sites. Genes Dev. 8, 811–820. Baynton, K., Bresson-Roy, A., and Fuchs, R. P. P. (1998). Analysis of damage tolerance pathways in Saccharomyces cerevisiae: A requirement for Rev3 DNA Polymerase in translesion synthesis. Mol. Cell. Biol. 18, 960–966. Baynton, K., Bresson-Roy, A., and Fuchs, R. P. P. (1999). Distinct roles for Rev1p and Rev7p during translesion synthesis in Saccharomyces cerevisiae. Mol. Micro. 34, 124–133.
196
LAWRENCE
Bemark, M., Khamlichi, A. A., Davies, S. L., and Neuberger, M. S. (2000). Disruption of mouse polymerase (Rev3) leads to embryonic lethality and impairs blastocyst development in vitro. Curr. Biol. 10, 1213–1216. Berdal, K. G., Johansen, R. F., and Seeberg, E. (1998). Release of normal bases from intact DNA by a native DNA repair enzyme. EMBO J. 17, 363–367. Bezzubova, O., Silbergleit, A., Yamaguchi-Iwai, Y., Takeda, S., and Buerstedde, J.-M. (1997). Reduced X-ray resistance and homologous recombination frequencies in a RAD54/ mutant of the chicken DT40 cell line. Cell 89, 185–193. Bresson, A., and Fuchs, R. P. P. (2002). Lesion bypass in yeast cells: Pol participates in a multi-DNA polymerase process. EMBO J. 21, 3881–3887. Braithwaite, D. K., and Ito, J. (1993). Compilation, alignment, and phylogenetic relationships of DNA polymerases. Nucleic Acids Res. 21, 787–802. Cabral-Neto, J. B., Caseira-Cabral, R. E., Margot, A., Le Page, F., Sarasin, A., and Gentil, A. (1994). Coding properties of a unique apurinic/apyrimidinic site replicated in mammalian cells. J. Mol. Biol. 240, 416–420. Cassier, C., Chanet, R., Henriques, J. A. P., and Moustacchi, E. (1980). The effect of three PSO genes on induced mutagenesis: A novel class of mutationally defective yeast. Genetics 96, 841–857. Cassier-Chauvat, C., and Moustacchi, E. (1988). Allelism between psol-1 and rev3-1 mutants and between pso2-1 and snm1 mutants in Saccharomyces cerevisiae. Curr. Genet. 13, 37–40. Chae, S.-K., and Kafer, E. (1993). uvs-1 mutants defective in UV mutagenesis define a fourth epistatic group of uvs genes in Aspergillus. Curr. Genet. 24, 67–74. Cheung, K.-J., and Li, G. (2001). The tumor suppressor INGI: Structure and function. Exp. Cell Res. 268, 1–6. Cheung, K.-J., Mitchell, D., Lin, P., and Li, G. (2001). The tumor suppressor candidate p33INGI mediates repair of UV-damaged DNA. Cancer Res. 61, 4974–4977. Clark, D. R., Zacharias, W., Panaitescu, L., and McGregor, W. G. (2003). Ribozymemediated REV1 inhibition reduces the frequency of UV-induced mutations in the human HPRT gene. Nucleic Acids Res. 31, 4981–4988. Datta, A., and Jinks-Robertson, S. (1995). Association of increased spontaneous mutation rates with high levels of transcription in yeast. Science 268, 1616–1619. Diaz, M., and Storb, U. (2003). A novel cytidine deaminase AIDs in the delivery of error-prone polymerases to immunoglobulin genes. DNA Repair 2, 623–627. Diaz, M., Verkoczy, L. K., Flajnik, M. F., and Klinman, N. R. (2001). Decreased frequency of somatic hypermutation and impaired affinity maturation but intact germinal center formation in mice expressing antisense RNA to DNA polymerase . J. Immunology 167, 327–335. Di Noia, J., and Neuberger, M. (2002). Altering the pathway of immunoglobulin hypermutation by inhibiting uracil-DNA glycosylase. Nature 419, 1–3. Eeken, J. C. J., Romeijn, R. J., de Jong, A. W. M., Pastink, A., and Lohman, P. H. M. (2001). Isolation and genetic characterization of the Drosophila homologue of (SCE)REV3, encoding the catalytic subunit of DNA polymerase . Mutat. Res. 485, 237–253. Esposito, G., Godin, I., Klein, U., Yaspo, M.-L., Cumano, A., and Rajewsky, K. (2000). Disruption of the Rev3l-encoded catalytic subunit of polymerase in mice results in early embryonic lethality. Curr. Biol. 10, 1221–1224. Garkavtsev, I., Grigorian, I. A., Ossovskaya, V. S., Chernov, M. V., Chumakov, P. M., and Gudkov, A. V. (1998). The candidate tumor suppressor p33ING1 cooperates with p53 in cell growth control. Nature 391, 295–298.
DNA POLYMERASE AND REV1 PROTEIN
197
Gibbs, P. E. M., and Lawrence, C. W. (1995). Novel mutagenic properties of abasic sites in Saccharomyces cerevisiae. J. Mol. Biol. 251, 229–236. Gibbs, P. E. M., McGregor, W. G., Maher, V. M., Nisson, P., and Lawrence, C. W. (1998). A human homolog of the Saccharomyces cerevisiae REV3 gene, which encodes the catalytic subunit of DNA polymerase . Proc. Natl. Acad. Sci. USA 95, 6876–6880. Gibbs, P. E. M., Wang, X.-D., Li, Z., McManus, T. P., McGregor, W. G., Lawrence, C. W., and Maher, V. M. (2000). The function of the human homolog of Saccharomyces cerevisiae REV1 is required for mutagenesis induced by UV light. Proc. Natl. Acad. Sci. USA 97, 4186–4191. Gibbs, P. E. M., MacDonald, J., Woodgate, R., and Lawrence, C. W. (2004). The relative roles in vivo of Saccharomyces cerevisiae Pol , Pol , Rev1 protein, and Pol 32 in the bypass and mutation induction of an abasic site, T-T (6-4) photoadduct, and T-T cis-syn cyclobutane dimer. Genetics. (in press). Glassner, B. J., Rasmussen, L. J., Najarian, M. T., Posnick, L. M., and Samson, L. D. (1998). Generation of a strong mutator phenotype in yeast by imbalanced base excision repair. Proc. Nat. Acad. Sci. USA 95, 9997–10002. Guillet, M., and Boiteux, S. (2003). Origin of endogenous DNA abasic sites in Saccharomyces cerevisiae. Mol. Cell. Biol. 23, 8366–8394. Guo, D., Wu, X., Rajpal, D. K., Taylor, J.-S., and Wang, Z. (2001). Translesion synthesis by yeast DNA polymerase from templates containing lesions of ultraviolet radiation and acetylaminofluorene. Nucleic Acids Res. 29, 2875–2883. Guo, C., Fischhaber, P. L., Luk-Paszyc, M. J., Masuda, Y., Zhou, J., Kamiya, K., Kisker, C., and Friedberg, E. C. (2003). Mouse Rev1 protein interacts with multiple DNA polymerases involved in translesion DNA synthesis. EMBO J. 22, 6621–6630. Han, K.-Y., Chae, S.-K., and Han, D.-M. (1998). The uvsI gene of Aspergillus nidulans required for UV-mutagenesis encodes a homolog to REV3, a subunit of the DNA polymerase of yeast involved in translesion DNA synthesis. FEMS Microbiol. Lett. 164, 13–19. Haracska, L., Prakash, S., and Prakash, L. (2000). Replication past O6-methylguanine by yeast and human DNA polymerase . Mol. Cell. Biol. 20, 8001–8007. Haracska, L., Washington, M. T., Prakash, S., and Prakash, L. (2001a). Inefficient bypass of an abasic site by DNA polymerase . J. Biol. Chem. 276, 6861–6866. Haracska, L., Kondratick, C. M., Unk, I., Prakash, S., and Prakash, L. (2001b). Interaction with PCNA is essential for yeast DNA polymerase function. Mol. Cell 8, 407–415. Haracska, L., Unk, I., Johnson, R. E., Johansson, E., Burgers, P. M. J., Prakash, S., and Prakash, L. (2001c). Roles of yeast DNA polymerase and and of Rev1 in the bypass of abasic sites. Genes Dev. 15, 945–954. Haracska, L., Johnson, R. E., Unk, I., Phillips, B., Hurwitz, J., Prakash, L., and Prakash, S. (2001d). Physical and functional interactions of human DNA polymerase with PCNA. Mol. Cell. Biol. 21, 7199–7206. Haracska, L., Prakash, S., and Prakash, L. (2002). Yeast Rev1 protein is a G template specific DNA polymerase. J. Biol. Chem. 277, 15546–15551. Haracska, L., Prakash, S., and Prakash, L. (2003). Yeast DNA polymerase is an efficient extender of primer ends opposite from 7,8-dihydro-8-oxoguanine and O6-methyl guanine. Mol. Cell. Biol. 23, 1453–1459. Harfe, B. D., and Jinks-Robertson, S. (2000). DNA polymerase introduces multiple mutations when bypassing spontaneous DNA damage in Saccharomyces cerevisiae. Mol. Cell 6, 1491–1499.
198
LAWRENCE
Heidenreich, E., Holzmann, V., and Eisler, H. (2004). Polymerase dependency of increased adaptive mutation frequencies in nucleotide excision repair-deficient yeast strains. DNA Repair 3, 395–402. Henriques, J. A. P., and Moustacchi, E. (1980). Isolation and characterization of pso mutants sensitive to photo-addition of psoralen derivatives in Saccharomyces cerevisiae. Genetics 95, 273–288. Hoege, C., Pfander, B., Moldovan, G.-L., Pyrowolakis, G., and Jentsch, S. (2002). RAD6dependent DNA repair is linked to modification of PCNA by ubiquitin and SUMO. Nature 419, 135–141. Holbeck, S. L., and Strathern, J. N. (1997). A role for REV3 in mutagenesis during double-strand break repair in Saccharomyces cerevisiae. Genetics 147, 1017–1024. Huang, M.-E., de Caligon, A., Nicolas, A., and Galibert, F. (2000). Pol32, a subunit of the Saccharomyces cerevisiae DNA polymerase , defines a link between DNA replication and the mutagenic bypass repair pathway. Curr. Genet. 38, 178–187. Ito, J., and Braithwaite, D. K. (1991). Compilation and alignment of DNA polymerase sequences. Nucleic Acids Res. 19, 4045–4057. Johnson, R. E., Prakash, S., and Prakash, L. (1999). Efficient bypass of a thyminethymine dimer by yeast DNA polymerase, Pol. Science 283, 1001–1004. Johnson, R. E., Washington, M. T., Haracska, L., Prakash, S., and Prakash, L. (2000)). Eukaryotic Polymerases and act sequentially to bypass DNA lesions. Nature 406, 1015–1019. Johnson, R. E., Haracska, L., Prakash, S., and Prakash, L. (2001). Role of DNA polymerase in the bypass of a (6–4) TT photoproduct. Mol. Cell. Biol. 21, 3558–3563. Johnson, R. E., Yu, S.-L., Prakash, S., and Prakash, L. (2003). Yeast DNA polymerase zeta () is essential for error-free replication past thymine glycol. Genes Dev. 17, 77–87. Kajiwara, K., Nagawawa, H., Shimizu-Nishikawa, K., Ookuri, T., Kimura, M., and Sugaya, E. (1996). Molecular characterization of seizure-related genes isolated by differential screening. Biochem. Biophys. Res. Comm. 219, 795–799. Krejci, L., Van Komen, S., Li, Y., Villemain, J., Reddy, M. S., Klein, H., Ellenberger, T., and Sung, P. (2003). DNA helicase Srs2 disrupts the Rad51 presynaptic filament. Nature 423, 305–309. Kozak, M. (1986). Point mutations define a sequence flanking the AUG initiator codon that modulates translation by eukaryotic ribosomes. Cell 44, 283–292. Kunz, B. A., Hensen, E. S., Roches, H., Ramotor, D., Nunoshiba, T., and Demple, B. (1994). Specificity of the mutator caused by the deletion of the yeast structural gene [APN1] for the major apurinic endonuclease. Proc. Nat. Acad. Sci. USA 91, 8165–8169. Larimer, F. W., Perry, J. R., and Hardigree, A. A. (1989). The REV1 gene of Saccharomyces cerevisiae: Isolation, sequence, and functional analysis. J. Bacteriol 171, 230–237. Lawrence, C. W. (2002). Cellular roles of DNA polymerase and Rev1 protein. DNA Repair 1, 425–435. Lawrence, C. W., and Christensen, R. B. (1976). UV mutagenesis in radiation-sensitive strains of yeast. Genetics 82, 207–232. Lawrence, C. W., and Christensen, R. B. (1978). Ultraviolet-induced reversion of cyc1 alleles in radiation-sensitive strains of yeast. I. rev1 mutant strains. J. Mol. Biol. 122, 1–21. Lawrence, C. W., and Christensen, R. B. (1979). Ultraviolet-induced reversion of cyc1 alleles in radiation-sensitive strains of yeast. III. rev3 mutant strains. Genetics 92, 397–408.
DNA POLYMERASE AND REV1 PROTEIN
199
Lawrence, C. W., and Hinkle, D. C. (1996) DNA Polymerase and the control of DNA damage induced mutagenesis in eukaryotes. In ‘‘Cancer Surveys’’ (T. Lindahl, Ed.), Vol. 28, pp. 21–31. Genetic Stability in Cancer. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. Lawrence, C. W., O’Brien, T., and Bond, J. (1984). UV-induced reversion of his4 frameshift mutations in rad6, rev1, and rev3 mutants of yeast. Mol. Gen. Genet. 195, 487–490. Lawrence, C. W., Das, G., and Christensen, R. B. (1985a). REV7, a new gene concerned with UV mutagenesis in yeast. Mol. Gen. Genet. 200, 80–85. Lawrence, C. W., Nisson, P. E., and Christensen, R. B. (1985b). UV and chemical mutagenesis in rev7 mutants of yeast. Mol. Gen. Genet. 200, 86–91. Lawrence, C. W., Kraus, B. R., and Christensen, R. B. (1985c). New mutations affecting induced mutagenesis in yeast. Mutat. Res. 150, 211–216. Lawrence, C. W., Gibbs, P. E. M., Murante, R. S., Wang, X.-D., Li, Z., McManus, T. P., McGregor, W. G., Nelson, J. R., Hinkle, D. C., and Maher, V. M. (2000) Roles of DNA polymerase and Rev1 protein in eukaryotic mutagenesis and translesion replication. In ‘‘Cold Spring Harbor Symposia on Quantitative Biology,’’ Vol. 65, pp. 61–69. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. Lemontt, J. F. (1971). Mutants of yeast defective in mutation induced by ultraviolet light. Genetics 68, 21–33. Lemontt, J. F. (1972). Induction of forward mutations in mutationally defective yeast. Mol. Gen. Genet. 119, 27–42. Li, Z., Zhang, H., Mcmanus, T. P., McCormick, J. J., Lawrence, C. W., and Maher, V. M. (2002). hREV3 is essential for error-prone translesion synthesis past UV or benzo[a]pyrene diol epoxide-induced DNA lesions in human fibroblasts. Mutat. Res. 510, 71–80. Lin, W., Wu, X., and Wang, Z. (1998). A full-length cDNA of hREV3 is predicted to encode DNA polymerase for damage-induced mutagenesis in humans. Mutat. Res. 433, 89–98. Lin, W., Xin, H., Zhang, Y., Wu, X., Yuan, F., and Wang, Z. (1999). The human REV1 gene codes for a DNA template-dependent dCMP transferase. Nucleic Acids Res. 27, 4468–4475. Lindahl, T. (1993). Instability and decay of the primary structure of DNA. Nature 362, 709–715. Loewith, R., Meijer, M., Lees-Miller, S. P., Riabowol, K., and Young, D. (2000). Three yeast proteins related to the human candidate tumor suppressor p33ING1 are associated with histone acetyltransferase activities. Mol. Cell. Biol. 20, 3807–3816. Masuda, Y., Takahashi, M., Fukuda, S., Sumii, M., and Kamiya, K. (2002). Mechanisms of dCMP transferase reactions catalyzed by mouse Rev1 protein. J. Biol. Chem. 277, 3040–3046. Masuda, Y., Ohmae, M., Masuda, K., and Kamiya, K. (2003). Structure and enzymatic properties of a stable complex of the human REV1 and REV7 proteins. J. Biol. Chem. 278, 12356–12360. Mendelman, L. V., Petruska, J., and Goodman, M. F. (1990). Base mispair extension kinetics. J. Biol. Chem. 265, 2338–2346. McKee, R. H., and Lawrence, C. W. (1979a). Genetic analysis of gamma-ray mutagenesis in yeast I. Reversion in radiation-sensitive strains. Genetics 93, 361–373. McKee, R. H., and Lawrence, C. W. (1979b). Genetic analysis of gamma-ray mutagenesis in yeast II. Allele-specific control of mutagenesis. Genetics 93, 375–381.
200
LAWRENCE
Moerschell, R. P., Tsunasawa, S., and Sherman, F. (1988). Transformation of yeast with synthetic oligonucleotides. Proc. Natl. Acad. Sci. USA 85, 524–528. Mol, C. D., Izumi, T., Mitra, S., and Tainer, J. A. (2000). DNA-bound structures and mutants reveal abasic DNA binding by APE1 DNA repair and coordination. Nature 403, 451–456. Morelli, C., Mungall, A. J., Negrini, M., Barbanti-Brodano, G., and Croce, C. M. (1998). Alternate splicing, genomic structure, and fine chromosome localization of Rev3L, Cytogenet. Cell Genet. 83, 18–20. Morrison, A., Christensen, R. B., Alley, J., Beck, A. K., Bernstine, E. G., Lemontt, J. F., and Lawrence, C. W. (1989). REV3, a Saccharomyces cerevisiae gene whose function is required for induced mutagenesis, is predicted to encode a nonessential DNA polymerase. J. Bacteriol. 171, 5659–5667. Murakumo, Y. (2002). The property of DNA polymerase : REV7 is a putative protein involved in translesion DNA synthesis and cell cycle control. Mutat. Res. 510, 37–44. Murakumo, Y., Roth, T., Ishii, H., Rasio, D., Numata, S.-I., Croce, C. M., and Fishel, R. (2000). A human REV7 homolog that interacts with the polymerase catalytic subunit hREV3 and the spindle assembly checkpoint protein hMAD2. J. Biol. Chem. 275, 4391–4397. Murakumo, Y., Ogura, Y., Ishii, H., Numata, S.-I., Ichihara, M., Croce, C. M., Fishel, R., and Takahashi, M. (2001). Interactions in the error-prone post replication repair proteins hRev1, hRev3, and hRev7. J. Biol. Chem. 276, 35644–35651. Muramatsu, M., Kinoshita, K., Fagarasan, S., Yamada, S., Shinkai, Y., and Honjo, T. (2000). Class switch recombination and somatic hypermutation require activationinduced cytidine deaminase (AID), a potential RNA editing enzyme. Cell 102, 553–563. Nelson, J. R., Lawrence, C. W., and Hinkle, D. C. (1996a). Thymine-thymine dimer bypass by yeast DNA polymerase . Science 272, 1646–1649. Nelson, J. R., Lawrence, C. W., and Hinkle, D. C. (1996b). Deoxycytidyl transferase activity of yeast REV1 protein. Nature 382, 729–731. Nelson, J. R., Gibbs, P. E. M., Nowicka, A. M., Hinkle, D. C., and Lawrence, C. W. (2000). Evidence for a second function for Saccharomyces cerevisiae Rev1p. Mol. Micro. 37, 549–554. Ohashi, E., Murakumo, Y., Kanjo, N., Akagi, J.-I., Masutani, C., Hanaoka, F., and Ohmori, H. (2004). Interaction of hREV1 with three human Y-family DNA polymerases. Genes Cells 9, 523–531. Ohmori, H., Friedberg, E. C., Fuchs, R. P. P., Goodman, M. F., Hanaoka, F., Hinkle, D., Kunkel, T. A., Lawrence, C. W., Livneh, Z., Nohmi, T., Prakash, L., Prakash, S., Todo, T., Walker, G. C., Wang, Z., and Woodgate, R. (2001). The Y-family of DNA polymerases. Mol. Cell 8, 7–8. Otsuka, C., Kobayashi, K., Kawaguchi, N., Kunitomi, N., Moriyama, K., Hata, Y., Iwai, S., Loakes, D., Noskov, V. N., Pavlov, Y., and Negishi, K. (2002a). Use of yeast transformation by oligonucleotides to study DNA lesion bypass in vivo. Mutat. Res. 502, 53–60. Otsuka, C., Sanadai, S., Hata, Y., Okuto, H., Noskov, V. N., Loakes, D., and Negishi, K. (2002b). Difference between deoxyribose- and tetrahydrofuran-type abasic sites in the in vivo mutagenic response in yeast. Nucleic Acids Res. 30, 5129–5135. Otsuka, C., Loakes, D., and Negishi, K. (2002c). The role of deoxycytidyl transferase activity of yeast Rev1 protein in the bypass of abasic sites. Nucleic Acids Res. 2, 87–88.
DNA POLYMERASE AND REV1 PROTEIN
201
Otsuka, C., Kunitomi, N., Iwai, S., Loakes, D., and Negishi, K. (2004). Role of REV1 polymerase domain and DNA polymerase h in translesion DNA synthesis in yeast in vivo. Submitted. Paulovich, A. G., Armout, C. D., and Hartwell, L. H. (1998). The Saccharomyces cerevisiae RAD9, RAD17, RAD24, and MEC3 genes are required for tolerating irreparable, ultraviolet-induced damage. Genetics 150, 75–93. Prakash, L. (1976). Effect of genes controlling radiation sensitivity on chemically induced mutations in Saccharomyces cerevisiae. Genetics 83, 285–301. Prakash, S., and Prakash, L. (2002). Translesion DNA synthesis in eukaryotes: A one- or two-polymerase affair. Genes Dev. 16, 1872–1883. Quah, S.-K., von Borstel, R. C., and Hastings, P. J. (1980). The origin of spontaneous mutation in Saccharomyces cerevisae. Genetics 96, 819–839. Rada, C., Williams, G., Nilsen, H., Barnes, D., Lindahl, T., and Neuberger, M. (2002). Immunoglobulin isotype switching is inhibited and somatic hypermutation perturbed in UNG-deficient mice. Curr. Biol. 12, 1748–1755. Rattray, A. J., Shafer, B. K., McGill, C. B., and Strathern, J. N. (2002). The roles of REV3 and RAD57 in double-strand-break-repair-induced-mutagenesis of Saccharomyces cerevisiae. Genetics 162, 1063–1077. Rechkoblit, O., Zhang, Y., Guo, D., Wang, Z., amin, S., Krzeminsky, J., Louneva, N., and Geacintov, N. E. (2002). Translesion synthesis past bulky benzo[a]pyrene diol epoxide N2-dG and N6-dA lesions catalyzed by DNA bypass polymerases. J. Biol. Chem. 277, 30488–30494. Roche, H. R., Gietz, R. D., and Kunz, B. A. (1994). Specificity of the rev3 antimutator and REV3 dependency of the mutator resulting from a defect (rad1) in nucleotide excision repair. Genetics 137, 637–646. Roche, H. R., Gietz, R. D., and Kunz, B. A. (1995). Specificities of the Saccharomyces cerevisiae rad6, rad18, and rad52 mutators exhibit different degrees of dependence on the REV3 gene product, a putative nonessential DNA polymerase. Genetics 140, 443–456. Rogozin, I. B., Pavlov, Y. I., Bebenek, K., Matsuda, T., and Kunkel, T. A. (2001). Somatic mutation hotspots correlate with DNA polymerase error spectrum. Nat. Immunol. 2, 530–536. Ruhland, A., and Brendel, M. (1979). Mutagenesis by cytostatic alkylating agents in yeast strains of differing repair capacities. Genetics 92, 83–97. Sakai, W., Ishii, C., and Inoue, H. (2002). The upr-1 gene encodes a catalytic subunit of the DNA polymerase which is involved in damage-induced mutagenesis in Neurospora crassa. Mol. Genet. Genomics 267, 401–408. Sakai, W., Wada, Y., Naoi, Y., Ishii, C., and Inoue, H. (2003). Isolation and genetic characterization of the Neurospora crassa REV1 and REV7 homologs: Evidence for involvement in damage-induced mutagenesis. DNA Repair. 2, 337–346. Sakamoto, A., Lan, V. T. T., Hase, Y., Shikazono, N., Matsunaga, T., and Tanaka, A. (2003). Disruption of the AtREV3 gene causes hypersensitivity to ultraviolet B light and -rays in Arabidopsis: Implications of the presence of a translesion synthesis mechanism in plants. Plant Cell 15, 2042–2057. Sale, J. E., Calandrini, D. M., Takata, M., Takeda, S., and Neuberger, M. (2001). Ablation of XRCC2/3 transforms immunoglobulin V gene conversion into somatic hypermutation. Nature 412, 921–926. Schiestl, R. H., Prakash, S., and Prakash, L. (1990). The SRS2 suppressor of Rad6 mutations of Saccharomyces cerevisiae acts by channeling DNA lesions into the RAD52 DNA repair pathway. Genetics 124, 817–831.
202
LAWRENCE
Simhadri, S., Kramata, P., Zajc, B., Sayer, J. M., Jerina, D. M., Hinkle, D. C., and Wei, C. S.-J. (2002). Benzo[a]pyrene diol epoxide-deoxyguanosine adducts are accurately bypassed by yeast DNA polymerase in vitro. Mutat. Res. 508, 137–145. Simon, J. A., Szankasi, P., Nguyen, D. K., Ludlow, C., Dunstan, H. M., Roberts, C. J., Jensen, E. L., Hartwell, L. H., and Friend, S. H. (2000). Differential toxicities of anticancer agents among DNA repair and checkpoint mutants of Saccharomyces cerevisiae. Cancer Res. 60, 328–333. Simpson, L. J., and Sale, J. E. (2003). Rev1 is essential foe DNA damage tolerance and non-teplated immunogobulin gene mutation in a vertebrate cell line. EMBO J. 22, 1654–1664. Singhal, R. K., Hinkle, D. C., and Lawrence, C. W. (1992). The REV3 gene of Saccharomyces cerevisiae is regulated more like a repair gene than one encoding a DNA polymerase. Mol. Gen. Genet. 236, 17–24. Sonoda, E., Okada, T., Zhao, G. Y., Tateishi, S., Araki, K., Yamaizumi, M., Yagi, T., Verkaik, N. S., van Gent, D. C., Takata, M., and Takeda, S. (2003). Multiple roles of Rev3, the catalytic subunit of pol in maintaining genome stability in vertebrates. EMBO J. 22, 3189–3197. Stelter, P., and Ulrich, H. D. (2003). Control of spontaneous and damage-induced mutagenesis by SUMO and ubiquitin conjugation. Nature 425, 188–191. Takeshita, M., and Eisenberg, W. (1994). Mechanism of mutation on DNA templates containing synthetic abasic sites: study with a double strand vector. Nucleic Acids Res. 22, 1897–1902. Thomas, D. C., Roberts, J. D., Sabatino, R. D., Myers, T. W., Tan, C.-K., Downey, K. M., So, A. G., Bambara, R. A., and Kunkel, T. A. (1991). Fidelity of mammalian DNA replication and replicative DNA polymerases. Biochem. 30, 11751–11759. Torpey, L. E., Gibbs, P. E. M., Nelson, J. R., and Lawrence, C. W. (1994). Cloning and sequence of REV7, a gene whose function is required for DNA damage-induced mutagenesis in Saccharomyces cerevisiae. Yeast 10, 1503–1509. Vanderwiele, D., Borden, A., O’Grady, P. I., Woodgate, R., and Lawrence, C. W. (1998). Efficient translesion replication in the absence of Escherichia coli Umu proteins and 30 –50 exonuclease proofreading function. Proc. Natl. Acad. Sci. USA 95, 15519–15534. Van Sloun, P. P., Romeijn, R. J., and Eeken, J. C. (1999). Molecular cloning, expression and chromosomal location of the mouse Rev31 gene, encoding the catalytic subunit of polymerase . Mutat. Res. 109, 109–116. Van Sloun, P. P., Varlet, I., Sonneveld, E., Boei, J. J., Romeijn, R. J., Eeken, J. C., and De Wind, N. (2002). Involvement of Mouse Rev3 in tolerance of endogenous and exogenous DNA damage. Mol. Cell. Biol. 22, 2159–2169. Vaisman, A., Frank, E. G., McDonald, J. P., Tissier, A., and Woodgate, R. (2002). Poldependent lesion bypass in vitro. Mutat. Res. 510, 9–22. Veaute, X., Jeusset, J., Soustelle, C., Kowalczkowski, S. C., Le Cam, E., and Fabre, F. (2003). The Srs2 helicase prevents recombination by disrupting Rad51 nucleoprotein filaments. Nature 423, 309–312. Wittschieben, J., Shivji, M. K., Lalani, E., Jacobs, M. A., Marini, F., Gearhart, P. J., Rosewell, I., Stamp, G., and Wood, R. D. (2000). Disruption of the developmentally regulated Rev3l gene causes embryonic lethality. Curr. Biol. 10, 1217–1220. Xiao, W., Lechler, T., Chow, B. L., Fontanie, T., Agustus, M., Carter, K. C., and Wei, Y.-F. (1998). Identification, chromosomal mapping and tissue-specific expression of hREV3 encoding a putative human DNA polymerase . Carcinogenesis 19, 945–949.
DNA POLYMERASE AND REV1 PROTEIN
203
Yang, I.-Y., Miller, H., Wang, Z., Frank, E. G., Ohmori, H., Hanaoka, F., and Moriya, M. (2003). Mammalian translesion DNA synthesis across an acrolein-derived deoxyguanosine adduct. J. Biol. Chem. 278, 13989–13994. Yuan, F., Zhang, Y., Rajpal, D. K., Wu, X., Guo, D., Wang, M., Taylor, J.-S., and Wang, Z. (2000). Specificity of DNA lesion bypass by the yeast DNA polymerase . J. Biol. Chem. 275, 8233–8239. Zan, H., Komori, A., Li, Z., Cerutti, A., Schaffer, A., Fajnok, M. F., Diaz, M., and Casali, P. (2001). The translesion DNA polymerase plays a major role in Ig and bcl-6 somatic hypermutation. Immunity 14, 643–653. Zeng, X., Winter, D. B., Kasmer, C., Kraemer, K. H., Lehmann, A. R., and Gearhart, P. J. (2001). DNA polymerase is an AT mutator in somatic hypermutation of immunoglobulin variable genes. Nature Immunology 2, 537–541. Zhang, Y., Wu, X., Rechkoblit, O., Geacintov, N. E., Taylor, J.-S., and Wang, Z. (2002). Response of human Rev1 to different DNA damage: Preferential dCMP insertion opposite the lesion. Nucleic Acids Res. 30, 1630–1638.
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DNA POLYMERASES h AND i By ALEXANDRA VAISMAN,* ALAN R. LEHMANN,À AND ROGER WOODGATE* *Laboratory of Genomic Integrity, National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, Maryland, 20892-2725 À Genome Damage and Stability Centre, University of Sussex, Falmer, Brighton BN1 9RQ, United Kingdom
I. II. III. IV. V. VI. VII. VIII. IX. X.
Historical Perspective . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . Identification of RAD30 and its Orthologs. . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . Biochemical Properties of Pol and Pol . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . Translesion Synthesis by Pol and Pol . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . Structure of the Catalytic Core of S. Cerevisiae Pol .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . Regulation and Localization of Pol and Pol. . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . Mutations in Pol in XP Variants. . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . Pols and and the Polymerase Switch: Interactions with PCNA and Rev1 . . . Protection from Cellular Effects of DNA Damage . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . Roles of Pol and Pol in Somatic Hypermutation . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . .
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I. HISTORICAL PERSPECTIVE Based on genetic studies in Escherichia coli, it was believed for many years that damage-induced mutations in DNA were generated in a two-step process commonly referred to as Translesion DNA Synthesis (TLS) (for a general review, see Friedberg et al., 1995). The first step of this process is (mis)incorporation and involves the physical insertion of a nucleotide opposite the DNA lesion. The second step involves extension of the (mis)incorporated base, such that the replication-blocking lesion is completely bypassed. It was originally hypothesized that both steps were performed by the cell’s high-fidelity replicase, Pol III, with the assistance of accessory proteins, such as RecA and UmuDC, which together facilitated highly error-prone DNA replication through various DNA lesions (Bridges and Woodgate, 1984, 1985a,b). The first clue that cells may actually contain specific polymerases specialized in lesion bypass came in 1996 with the purification of Pol, a heterodimer consisting of Rev3/Rev7, which has the ability to replicate past ultraviolet (UV) induced thymine–thymine cis–syn cyclobutane dimers (CPDs) (Nelson et al., 1996b) (reviewed in detail in Chapter 6). At the same time, it was found that the product of the Saccharomyces cerevisiae REV1 gene long implicated in the mutagenic process has deoxycytidyl transferase activity (Nelson et al., 1996a). Although the REV3 gene encoding the catalytic subdomain of Pol belongs to the B-family of DNA polymerases, 205 ADVANCES IN PROTEIN CHEMISTRY, Vol. 69
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the REV1 gene exhibits no sequence or motif homology to any of the previously identified DNA polymerase families. Instead, it shares homology with the umuC gene (Larimer et al., 1989) that is required for damageinduced mutagenesis in E. coli (Kato and Shinoura, 1977; Steinborn, 1978). Because at that time Rev1 was believed to use only dCTP as a substrate, the possibility that UmuC or phylogenetically related proteins might actually possess DNA polymerase activity was largely overlooked. However, that view began to change in the late 1990s, when it was shown that a highly purified preparation of heterodimeric E. coli UmuD0 2C was able to bypass a synthetic abasic site unassisted (Tang et al., 1998). Although at that time there was a possibility that the UmuD0 2C preparation contained a trace amount of endogenous PolIII, the biochemical studies on UmuD0 2C provided the first hint that many of the proteins implicated in the mutagenic process might actually be bona fide DNA polymerases. The intrinsic polymerase activity of the UmuD0 2C complex was subsequently confirmed (Tang et al., 1999) and was shown to reside in the UmuC subunit of the complex (Reuven et al., 1999). Amazingly, within a short, 18-month period, DNA polymerase activity was also described for several proteins homologous to UmuC and Rev1. These include E. coli PolIV, encoded by the dinB gene (Wagner et al., 1999) and described in detail in Chapter 8; its human homolog Pol ( Johnson et al., 2000a; Ohashi et al., 2000; Zhang et al., 2000c), described in Chapter 9; and two eukaryotic paralogs of S. cerevisiae RAD30, Pol ( Johnson et al., 1999b; Masutani et al., 1999) and Pol ( Johnson et al., 2000b; Tissier et al., 2000b; Zhang et al., 2000b), which are described in detail later. Collectively, these proteins are now referred to as the Y-family of DNA polymerases (Ohmori et al., 2001). Over 200 proteins that share homology to the Y-family polymerases have since been identified in prokaryotes, archaea, and eukayrotes. Interestingly, many organisms often possess more than one family member, suggesting that Y-family polymerases play important roles in cellular survival or evolutionary ‘‘fitness’’ (Friedberg et al., 2002; Yeiser et al., 2002). Indeed, defects in human Pol result in the sunlight-sensitive and cancer-prone Xeroderma pigmentosum variant (XP-V) syndrome ( Johnson et al., 1999a; Masutani et al., 1999) (see following), whereas mutations in E. coli dinB reduces the cell’s ability to undergo adaptive mutagenesis in stationary phase (McKenzie et al., 2001; Tompkins et al., 2003).
II. IDENTIFICATION OF RAD30
AND ITS
ORTHOLOGS
The RAD30 gene was identified in a search of the S. cerevisiae genome for homologs of prokaryotic DinB- and UmuC-like proteins and for eukaryotic Revl-like proteins (McDonald et al., 1997; Roush et al., 1998). Similar to its
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E. coli counterparts, RAD30 is damage inducible, and Rad30-disrupted yeast strains are mildly sensitive to UV light (McDonald et al., 1997; Roush et al., 1998). Epistasis analysis suggested an involvement of the Rad30 protein in the postreplication repair pathway (McDonald et al., 1997), and an important step in understanding the cellular function of Rad30 came with its overproduction and purification ( Johnson et al., 1999b). Biochemical characterization revealed that the Rad30 protein is a bona fide DNA polymerase, which uses all four nucleoside triphosphates in a template-dependent reaction. Strikingly, however, Rad30 was shown to bypass a thymine–thymine CPD exceptionally efficiently and relatively accurately ( Johnson et al., 1999b). As the sixth eukaryotic polymerase reported in the literature at that time, the S. cerevisiae Rad30 protein was designated as Pol. Shortly afterward, a human homolog of RAD30 was identified and shown to be the enzyme mutated or missing in cells from humans exhibiting the XP-V phenotype ( Johnson et al., 1999a; Masutani et al., 1999). XP-V cells are proficient in nucleotide excision repair but defective in postreplicative repair (they have a reduced ability to make intact daughter DNA strands on damaged templates) (Lehmann et al., 1975). This defect results in an increase in UV-induced mutagenesis and leads to a high incidence of skin cancer (Cleaver and Carter, 1973). The name approved by the Human Genome Organization for the human RAD30 ortholog is POLH, although it is also commonly referred to as the XPV or RAD30A gene. Soon after the discovery of human POLH, a second human homolog of RAD30 was identified and initially designated RAD30B (McDonald et al., 1999). Subsequent studies demonstrated that, similar to the other phylogenetically related proteins, the Rad30B protein has DNA polymerase activity. The protein was therefore renamed DNA polymerase and is encoded by the human POLI gene (Tissier et al., 2000b). POLH orthologs are evolutionally conserved in a wide variety of eukaryotes from yeast to human, and it is thought that POLI most likely arose as a gene duplication of POLH during evolution of the species. POLI ’s distribution is much more limited than that of POLH, and it is found mostly in higher eukaryotes.
III. BIOCHEMICAL PROPERTIES OF POL AND POL Similar to other members of the Y-family, both Pol and Pol are highly error prone and lack intrinsic exonuclease activity, meaning they cannot proofread any of the errors they make when copying DNA. Both enzymes are distributive and incorporate only a few deoxynucleotides before dissociating from the extended product. There are, however, significant
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differences in the biochemical properties of these phylogenetically related enzymes. For example, a property that distinguishes human Pol from Pol and other members of Y-family DNA polymerases is its intrinsic 50 -deoxyribose phosphate (dRP) lyase activity (Bebenek et al., 2001b; Prasad et al., 2003); that is, the ability to catalyze excision of 50 -dRP from DNA during base-excision repair. Furthermore, Pol has stronger dRP lyase activity relative to its polymerase activity than does the main BER polymerase, Pol (Prasad et al., 2003). The fidelity of both yeast and human Pol is very low, with in vitro misincorporation frequencies ranging from 102 to 103 (Fig. 1A; Johnson et al., 2000c; Matsuda et al., 2000; Washington et al., 1999), which is at least 100 times less accurate than that of replicative DNA polymerases (Kunkel, 2004). As is usually seen with other DNA polymerases, error rates vary depending on mismatch composition and template sequence context (Goodman and Fygenson, 1998; Goodman et al., 1993; Mendelman et al., 1989; Zhang and Mathews, 1995). For example, higher misincorporation rates are observed when the base-pair 50 of the error is T:A or A:T compared to C:G or G:C (Matsuda et al., 2000). Pol’s misincorporation frequency and specificity is unprecedented, even when compared to Pol. In general, Pol forms most mispairs more frequently than Pol (Fig. 1A). Furthermore, Pol is characterized by a significantly broader range of misincorporation frequencies for different mispairs compared to Pol (Fig. 1, compare distribution of points along x - and y -axes). The fidelity of Pol is uniquely template dependent. On primed singlestranded DNA, the most accurate nucleotide incorporation is observed opposite template A (with a misinsertion frequency of about 1 104) (Tissier et al., 2000b). The preference for the correct nucleotide in this case is ensured by tighter binding and faster incorporation compared to the binding and incorporation of the incorrect nucleotide (Washington et al., 2004). In contrast, Pol makes an extremely high level of errors at template T. In fact, depending on the sequence context, the wobble base, G, is incorporated 3 to 11 times more often opposite T than the Watson– Crick base, A ( Johnson et al., 2000b; Tissier et al., 2000b; Vaisman et al., 2001). On DNA templates with a 1-nucleotide 50 overhang, the pattern of nucleotide misincorporation by Pol is entirely different from that on primed single-stranded DNA templates. In this case, Pol is most inaccurate on template C, where C and A are misinserted three- to eight-fold more often than the correct base, G (Frank et al., 2001). Pol and Pol are characterized not only by high levels of nucleotide misinsertion but also by their relatively high efficiency of mismatch extension compared to classical DNA polymerases. The range of single mismatch extension frequencies by Pol and both yeast and human Pol are similar
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FIG. 1. Comparison of insertion and extension fidelities of human Pols and . (A) Efficiencies of misincorporation ( finc) by Pol (Tissier et al., 2000b) were plotted versus efficiencies of misincorporation by Pol ( Johnson et al., 2000c; Washington et al., 1999), which is at least 100 times less accurate than that of replicative DNA polymerases (Kunkel, 2004 #1346; Washington et al., 1999). The dashed line corresponds to finc Pol ¼ finc Pol . Points above the dashed line correspond to a higher frequency of nucleotide misinsertion catalyzed by Pol compared with that of Pol, whereas points below the dashed line represent the reverse situation. In the base mispairs shown, the first base is the incoming nucleotide and the second base is in the template. (B) 0 ) by Pol (Vaisman et al., 2001) were plotted versus Efficiencies of mispair extension ( fext the efficiencies of mispair extension by Pol (Washington et al., 2001). The dashed line corresponds to f 0ext Pol ¼ f 0ext Pol . Points above the dashed line correspond to a higher relative efficiency of mispair extension catalyzed by Pol compared with that of Pol, whereas points below the dashed line represent the reverse situation. In the base mispairs shown, the first base at the 30 primer terminus and the second base is in the template.
(Fig. 1B; Vaisman et al., 2001; Washington et al., 2001). Mismatch extension by human Pol is somewhat less efficient than the incorporation of the wrong nucleotide for most mispairs (Fig. 1; Washington et al., 2001). For Pol, the correlation between relative efficiencies of mismatch formation and extension is more dependent on the identity of a mispair and the next template nucleotide encountered (Fig. 1; Vaisman et al., 2001). More mispairs are extended by Pol, with a higher efficiency than by Pol (Fig. 1B; Matsuda et al., 2000; Vaisman et al., 2001; Washington et al., 2001). Although Pol is able to extend a variety of mismatches quite efficiently, a ‘‘buried’’ mispair located 2–3 bases from the nascent primer chain significantly inhibits further primer elongation and therefore restricts Pol synthesis to short regions of DNA (Vaisman et al., 2001). Pol is also essentially
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blocked by tandem mispairs, whereas human Pol extends tandem doublemismatched termini at rates similar to those at which other polymerases extend single mismatches (Matsuda et al., 2000). Human Pol also generates a remarkable variety of nucleotide deletion and addition errors with high frequencies (Matsuda et al., 2000). These frameshift errors are explained by strand slippage, often preceded by nucleotide misincorporation.
IV. TRANSLESION SYNTHESIS BY POL AND POL The ability of Pol and Pol to bypass a variety of structurally diverse and potentially mutagenic lesions in vitro has been analyzed extensively. These biochemical studies revealed that the efficiency and pattern of nucleotide incorporation opposite DNA damaged sites is distinct for the two Rad30 paralogs. In some cases, Pol readily bypasses a lesion, whereas Pol does it inefficiently, or not at all. An example of Pol’s and Pol’s disparate TLS activities is observed at thymine CPDs and cisplatin-induced dG intrastrand crosslinks. Pol efficiently facilitates robust bypass of both CPDs ( Johnson et al., 2000c; Masutani et al., 2000; McCulloch et al., 2004) and platinum-GG cross links (Vaisman et al., 2000) in vitro, whereas Pol bypasses the CPD in a limited manner (Tissier et al., 2000a) that is sequence context dependent (Vaisman et al., 2003) and is incapable of nucleotide incorporation opposite a cisplatin adduct (McDonald et al., 2001). On the contrary, Pol appears to be limited to insertion opposite the 30 T of UV-induced 6-4 pyrimidine-pyrimidone dimers (6-4PP) and is significantly inhibited after nucleotide incorporation at abasic sites and N-2-acetylaminofluorene (AAF)-modified guanine adducts (Haracska et al., 2001c; Kusumoto et al., 2002; Zhang et al., 2000a), whereas Pol is significantly more efficient at incorporating across from both Ts of the 6-4PP (Tissier et al., 2000a; Vaisman et al., 2003) and opposite an abasic site ( Johnson et al., 2000b; McDonald et al., 2001). When encountering certain other lesions, the enzymes behave in a similar manner. For example, both Pols can readily bypass an 8-oxoG lesion (Vaisman and Woodgate, 2001; Zhang et al., 2000a), and both are limited to inefficient incorporation opposite acrolein-derived deoxyguanosine (Minko et al., 2003; Yang et al., 2003) and benzo[a]pyrene (BP) adducts (Chiapperino et al., 2002; Frank et al., 2002; Rechkoblit et al., 2002). Thus, TLS not only depends on the particular lesion encountered but also on the polymerase attempting TLS. The accuracy with which any given lesion is bypassed is also uniquely dependent on each polymerase. The fact that humans lacking Pol have an increased incidence of skin cancers clearly indicates that Pol plays an
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important role in the accurate postreplication repair of UV-induced CPDs in vivo. Depending on the point of view taken, the fidelity of bypass of CPDs in vitro by Pol can be regarded as error prone or relatively error free. Using a template containing a CPD, ‘‘incorrect bases’’ opposite the Ts are inserted at a frequency of approximately 102 ( Johnson et al., 2000c; Masutani et al., 2000; McCulloch et al., 2004), which could be regarded as error prone. However, this means that in 99% of instances, Pol inserts the ‘‘correct’’ bases opposite these lesions. No other polymerase displays any comparable accuracy. With other lesions, Pol frequently misinserts erroneous bases. For example, dG is predominantly misinserted at the 30 T of the 6-4PP before DNA synthesis is aborted ( Johnson et al., 2001; Masutani et al., 2000). Pol also predominantly misincorporates dA, and to a lesser degree dG and dT, at acrolein-derived dG (Yang et al., 2003), dA at cyclodeoxyadenosine (Kuraoka et al., 2001), dT at O6-methylguanine (Haracska et al., 2002), and dG and dA opposite BP-G adducts (Chiapperino et al., 2002). On the basis of highly error-prone nature of the enzyme at these specific lesions in vitro, one has to speculate that either these errors are removed in vivo by exonucleolytic proofreading (Bebenek et al., 2001a) or TLS of the lesions is facilitated by a more accurate enzyme. The specificity of nucleotide misincorporation opposite many of the lesions tested is very different for Pol compared to that of Pol. Pol is characterized by unique nucleotide selectivity for most of the damaged sites and often incorporates the wrong nucleotide more efficiently than the correct one. Despite the fact that coding properties of damaged bases essentially depend on chemical structure, the miscoding potential of DNA lesions for Pol appears to be related to its fidelity on undamaged DNA. The selection of nucleotides by Pol is most unpredictable at template T. Depending on the surrounding sequence context and the type of damage, A, T, or G is predominantly inserted. An unusually accurate incorporation is observed at the 30 T of 6-4PP in an ATTC sequence context (Vaisman et al., 2003). This exception is particularly interesting because structural features of 6-4PP-containing duplex DNA suggest that 30 T of 6-4PP forms a more stable base pair with G than with A (Kim and Choi, 1995; Lee et al., 1999), which is consistent with the misincorporation specificity of other DNA polymerases, including Pol ( Johnson et al., 2001). In contrast to the extremely promiscuous behavior of Pol on template T, the enzyme is very accurate at template A, which is either undamaged or contains various stereoisomers of BP adducts placed in different sequence contexts (Frank et al., 2002). Although the misincorporation frequency increases upon formation of BP adducts at adenosine, the fidelity of Pol remains high (Frank et al., 2002). Thus, it is possible that while Pol has
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evolved to protect us from damage occurring at thymine, Pol may similarly help protect us from mutations at damaged adenosines. In addition to yeast and human Pol and human Pol, Drosophila and mouse Pol and Pol enzymes have been overproduced and characterized (Ishikawa et al., 2001; Matsuda et al., 2001; McDonald et al., 2003). Although the enzymatic properties of the various Pols were similar from all sources, the properties of Drosophila Pol were very different from either the human or mouse Pol enzymes. In fact, Drosophila Pol is more akin to Pol in its ability to traverse a CPD accurately (Ishikawa et al., 2001). Such observations lend support to the notion that Pol arose as a duplication of Pol and that evolutionary selective pressures acted on Pol over the millennia, so as to distinguish it from Pol.
V. STRUCTURE OF THE CATALYTIC CORE OF S.
CEREVISIAE
POL
Although human Pols and are only 23% identical to each other at the primary sequence level, they nevertheless have conserved blocks of amino acids (called motif I–V in Kulaeva et al., 1996), which are readily identifiable when their primary amino acid sequences are aligned to other Y-family polymerases (Fig. 2A). These five motifs are located in the N-terminal half of the two proteins, which is now known to contain the catalytic active site of all Y-family polymerases (Ling et al., 2001; Silvian et al., 2001; Trincao et al., 2001; Zhou et al., 2001). Remarkably, even though there is little to no similarity in the primary amino acid sequence of Yfamily polymerases with those previously identified from the A-, B-, C-, D-, or X-polymerase families, the three-dimensional structure of Pol is similar to those previously reported for other DNA polymerases, in that it retains the overall topology of a right hand that is capable of holding DNA in its grasp (Fig 2B). Similar to the classical polymerases, the catalytic core of Pol is composed of a thumb subdomain, which is thought to have a primary role in DNA duplex binding and polymerase processivity; a fingers subdomain, which is likely to be important for nucleoside triphosphate (dNTP) selection; and a palm subdomain, which contains the catalytically important residues for the nucleotidyl transfer reaction. The core of the palm domain of Pol includes a four-stranded antiparallel sheet flanked by two small helices and an -helical structure situated at the base of the palm, which is superimposable with the core of the palm domain from high-fidelity polymerases. Akin to the active sites of the polymerases from the A-, B-, C-, and X-families, the palm domain of Pol contains three carboxylates (Asp30 and Asp155-Glu156), which coordinate two catalytically essential metal ions assisting the nucleotidyl transferase reaction. Although the palm subdomain is nearly superimposable
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FIG. 2. Cartoon representation of the conserved regions in Pols and and the crystal structure of the catalytic core of Saccharomyces cerevisiae Pol. (A): Schematic alignment of hPOLH and hPOLI. Conserved amino acid sequences found in all Y-family polymerases are represented by the five colored boxes (motifs I–V) and by white boxes indicating unique sequences. Gaps have been introduced in the sequences for optimal alignment. The length of Pols and are indicated by the number of amino acids on the right site of the diagram. A conserved zinc-binding motif in the C terminus of Pol is indicated as C2H2 in the gray box. Structural domains of Pol shown in blue (finger, F), red (palm, P), green (thumb, T), and purple (little finger, LF) are indicated above the alignment. The positions of the nuclear localization signal (NLS), the domain involved in polymerase localizing into replication foci (FS), and regions responsible for the interaction with other proteins are also indicated. (B) Structural domains of Pol are shown in red (palm), green (thumb), blue (finger), and purple (little finger) as a ribbon diagram. The acidic residues (Asp30, Asp155, and Glu156) that make up the active site are shown as gold rods. S. cerevisiae Pol has an insert of 70 amino acids in the palm domain that is absent in the archaeal Y-family polymerases (Ling et al., 2001; Silvian et al., 2001; Zhou et al., 2001), and this region is shown in pink. This figure was made with ribbons (Carson, 1987). (See Color Insert.)
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on the palm of high-fidelity polymerases, the fingers and the thumb domains of Pol are small and stubby when compared to the fingers and thumb of polymerases from other families. These structural features provide a basis for the tolerance by Pol for mismatch- or lesion-induced geometric distortions in the DNA. The fingers subdomain in Pol consists of a -sheet and three small helices (Fig. 2). The ‘‘O helices,’’ which are believed to play a key role in fidelity enhancement by tightening the interface between the polymerase and the replicating base pair, are absent in the fingers of Pol. Instead, only a short loop is positioned to potentially interact with the replicating base pair. The thumb subdomain of Pol is composed of six helices that are structurally distinct from helices in other DNA polymerases (Fig. 2B). The surface area in the Pol active site enclosed by the palm, fingers, and thumb domains is reduced compared to classical polymerases and appears to be too small to allow for efficient binding and replicating DNA. To increase its binding area, Pol employs an additional C-terminal domain, which mimics an extra set of fingers and has been termed the polymerase-associated domain (PAD) (Trincao et al., 2001). This domain is not found in DNA polymerases from other families but is structurally conserved within the Y-family polymerases. In archaeal DinB homologs Dpo4 and Dbh, it has been termed the ‘‘little finger’’ (Ling et al., 2001) and the ‘‘wrist,’’ respectively (Silvian et al., 2001), and we will refer to this domain as the little finger (LF) throughout the rest of this chapter. The LF domain has been proposed to play a role similar to that of the accessory factors employed by other polymerases, so as to increase the processivity of the polymerase (Boudsocq et al., 2004). This domain in Pol consists of an antiparallel sheet and two long helices packed next to the fingers subdomain and is connected to the thumb by an extended peptide (Fig. 2B). The crystal structure of S. cerevisiae Pol lacked 120 C-terminal residues, which, although unnecessary for polymerase activity in vitro, are likely to be essential for protein–protein interactions with accessory factors, such as PCNA (see below). The corresponding C-terminal domain in human Pol is much longer (280 vs. 120 amino acids [aa]). The crystal structure of Pol was determined in the absence of DNA, but the location of DNA within the polymerase was determined by fitting the crystal structure of Pol to the previously reported template-primerddNTP ternary complex of T7 DNA polymerase (Trincao et al., 2001). Using such an approach, it was shown that the binding pocket of the polymerase around the templating base is likely to be open or extended, thereby providing an explanation for the low fidelity of the polymerase and suggesting a mechanism by which altered bases in the template can be
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accommodated within the active site. In particular, modeling of a covalently linked thymine dimer into the active site of Pol suggests it has the capacity to accommodate two templating residues. Support for this model comes from recent structural studies of Dpo4, a Y-family polymerase from Sulfolobus solfataricus, whose structure has been solved in a ternary complex with CDP-containing DNA and an incoming nucleotide (Ling et al., 2003). The Dpo4-CPD structure clearly showed that the covalently linked CPD was readily accommodated within the active site of Dpo4. Modeling of Pol onto the Dpo4 structure further suggested that the finger, thumb, and LF polymerase domains undergo a conformational change from an ‘‘open,’’ DNA-free complex to a ‘‘closed,’’ CPDcontaining complex, which facilitates efficient bypass of the CPD (Ling et al., 2003). The possibility that Pol retains both thymine bases of the CPD within its active site and directs incorporation of the correct nucleotides by using the intrinsic base-pairing ability of the lesion is also supported by kinetic studies (Washington et al., 2003b). Furthermore, structural and biochemical data suggest that intact Watson–Crick base pairing rather than the geometric fit of the incoming nucleotide with the templating base within the polymerase active site plays a crucial role in the efficiency and fidelity of DNA synthesis by Pol. Thus, Pol accommodates a CPD lesion because of the openness and flexibility of the active site, but also because Watson– Crick base pairing, although weakened, remains intact at the CPD. Similarly, yeast Pol readily traverses through 8-oxoG with the preferential formation of a C:8-oxoG base pair, which retains the three hydrogen bonds found in a normal C:G base pair (Haracska et al., 2000). In contrast to the CPD and 8-oxoG, an abasic site or 6-4PP lesion represents a strong obstacle to replication by Pol, as any hydrogen bonding potential is completely absent at an abasic site and is significantly disrupted at the 30 side of the 6-4PP. The importance of the ability to form normal hydrogen bonds during Pol-catalyzed DNA synthesis is further supported by the drastic decrease in insertion fidelity of yeast Pol in replication reactions with difluorotoluene, a nonpolar isosteric analog of thymine that is unable to form Watson–Crick hydrogen bonds with adenine but is virtually identical to dT in shape, size, and conformation (Washington et al., 2003a).
VI. REGULATION AND LOCALIZATION OF POL
AND
POL
The human POLH gene is expressed in all tissues, with small increases in expression in proliferating tissues such as testis, thymus, and skin, and also in the liver (Yamada et al., 2000). In mouse cells, Pol expression increased when serum-starved cells were stimulated to proliferate (Yamada et al.,
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2000). However, Pol is not required for normal DNA replication, as XP-V cells proliferate normally in the absence of DNA damage (Lehmann et al., 1975). Although the level of POLH mRNA decreases temporarily after UV irradiation (Yamada et al., 2000), protein levels are unaffected (P. Kannouche and ARL, unpublished observations). Both human and murine POLI are similarly expressed in all tissues, with the highest level of expression observed in postmeiotic spermatids from testis (Frank et al., 2001; McDonald et al., 1999). To gain insight into how Pol and Pol function inside the cell, localization studies have been carried out using eGFP-tagged polymerase constructs (Kannouche et al., 2001, 2002). This work showed that both polymerases were localized exclusively in the nucleus and were uniformly distributed through the nucleus in the majority of cells in an asynchronous culture of SV40-transformed human fibroblasts. However, in a small proportion of the cells (typically 15%), the polymerases accumulated in many bright foci, where they colocalized both with each other and with PCNA. These foci are thought to be replication factories, in which DNA replication is occurring. After treatment with UV irradiation, the number of cells containing bright foci of colocalizing Pol, Pol, and PCNA increased up to about 70% of the population (Fig. 3). It is believed that the number of foci increases because UV-induced DNA damage blocks progression of the replication forks and slows down cell passage through S phase. The resulting gradual accumulation of cells in S phase accounts for the increase in the number of cells with foci containing Pol, Pol, and PCNA, rather than DNA damage-induced relocalization of these proteins, as it was initially postulated. Consistent with this idea, accumulation of nuclei with foci containing Pol and Pol has also been observed after
FIG. 3. Colocalization of Pols and in response to UV irradiation. Foci formation after UV damage is shown for Pol (red) (left) and Pol (green) (middle). Both polymerases form foci in the same time and space within a cell following UV irradiation, indicating that the accumulation of Pol and Pol at replication forks stalled at sites of UV damage is coordinated in vivo. The colocalization of Pol and Pol is indicated by a yellow pattern in the merged figure (right). This figure is reproduced with permission from Kannouche et al., 2002. (See Color Insert.)
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other treatments that block progression of the replication fork, such as methyl methanesulfonate or N-acetoxy-acetylaminofluorene, which generate DNA damage, and hydroxyurea, which arrests cells in S phase, without causing DNA damage. The localization of Pol in replication foci is partly dependent on the presence of Pol because in XP-V cells, Pol remains nuclear, but its accumulation in replication foci is reduced approximately three- to six-fold (Kannouche et al., 2002). Deletion analysis has identified the domains required for correct localization of the polymerases (Fig. 2A). For Pol, the last 70 aa, containing a bipartite nuclear localization signal, are sufficient for nuclear localization, but not for accumulation into foci. However, the last 120 aa are sufficient for localization both in the nucleus and in nuclear foci (Kannouche et al., 2001). This 120-aa region contains a C2H2 zinc finger motif and, at the extreme C terminus, a PCNA-binding motif (Fig. 2A). Both of these are required for localization into foci (P. Kannouche and ARL, unpublished results). Pol does not contain any obvious motifs in the unconserved Cterminal part of the protein, and similar analyses were much less clear-cut. Localization into the nucleus required sequences contained within aa 219–451 at the end of the polymerase domain and the beginning of the nonconserved domain, whereas foci formation required the C-terminal 220 aa (Fig. 2A; Kannouche et al., 2002). Pol and Pol were shown to interact directly, and this same C-terminal 220 aa of Pol were required for the interaction with Pol. Taken together with the partial dependence on Pol for foci formation, these data suggest that Pol helps transport Pol into nuclear foci. The cellular role of aa 430–600 of Pol, which is poorly conserved among the Pol orthologs, is currently unknown.
VII. MUTATIONS IN POL IN XP VARIANTS The two initial studies in which POLH was discovered to be the gene defective in XP-V cells also identified several nonsense mutations in human patients with XP-V ( Johnson et al., 1999a; Masutani et al., 1999). Subsequent work identified many further mutations in XP-V patients (Broughton et al., 2002; Itoh et al., 2000), and these are summarized in Figure 4. The majority of the mutations cause truncations in Pol that are located within the conserved polymerase domain found in all Y-family polymerases and that are likely to produce a completely inactive protein. This suggests that Pol is not required for human life. A few truncating mutations are located close to the C terminus, outside of the polymerase domain. Extracts from these cells have TLS activity characteristic for Pol (Broughton et al., 2002), but we anticipate that the truncated Pol
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FIG. 4. Locations of mutations in POLH that lead to the Xeroderma pigmentosum variant (XP-V) phenotype. The locations of the amino acid changes identified in patients with XP-V are shown relative to the different domains of the polymerase shown in Fig. 2B. The structural domains of Pol are shown in blue (finger, F), red (palm, P), green (thumb, T), and purple (little finger, LF) in the central part of the diagram. The nuclear localization signal (NLS) and the domain involved in localizing Pol into replication foci (URS) are also indicated in black and brown, respectively. Cell strains are designated in boxes. Subscripts 1 and 2 specify different alleles. Deletions are shown using horizontal lines. Single amino acid changes are shown in the upper part of the diagram, and truncations are shown in the lower part. (See Color Insert.)
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molecules would probably not reside in the replisomes, as they are missing the C-terminal domain required for localization in the nucleus and in nuclear foci. Several missense mutations, mostly located within the conserved catalytic domain, have also been identified (Fig. 4, top). These were modeled onto the three-dimensional structure of Dpo4 from Sulfolobus solfataricus. Two of the mutated amino acids, G263 and R361, are involved in interactions with the DNA, whereas the other mutations are likely to disrupt the conformations of the different domains (Broughton et al., 2002). Interestingly, a Japanese patient is a compound heterozygote for two mutations, K535E and K589T, both located in the poorly conserved central domain between the polymerase and localization domains (Itoh et al., 2000). K535 is, however, in a run of nine aa conserved in mammalian species of Pol, and K589 is conserved in mouse and rat, demonstrating important functions for this part of the protein. As mentioned above, the severely truncating mutations suggest that Pol is not an essential gene in humans, and in support of this idea, a transgenic mouse lacking Pol has recently been generated (Fumio Hanaoka, personal communication).
VIII. POLS AND AND THE POLYMERASE SWITCH: INTERACTIONS WITH PCNA AND REV1 A topic of major interest that is also discussed in more detail in other chapters of this book is the mechanism by which the replicative DNA polymerase is displaced at a replication fork blocked by DNA damage and is replaced with a TLS polymerase. All three of the E. coli TLS polymerases have motifs involved in the interaction with the sliding clamp that helps to recruit polymerases to their site of action (Becherel et al., 2002). In eukaryotic cells, proliferating cell nuclear antigen (PCNA) plays the role of a sliding clamp, and analyses of the primary aa sequences of Pols and reveal putative PCNA-binding motifs. Yeast two-hybrid assays confirmed that there is a physical interaction between Pol and PCNA in vivo (Haracska et al., 2001a); and in vitro replication assays revealed that the catalytic activities of both Pol and Pol on undamaged and damaged DNA templates were increased by the presence of PCNA along with clamp loader, replication factor C (RFC), and RPA (replication factor A) (Haracska et al., 2001a,b). As discussed in detail in Chapter 10, PCNA becomes both mono- and polyubiquitinated in yeast cells following treatment with MMS (Hoege et al., 2002). Based on these findings, it was postulated that PCNA modification was involved in the polymerase switch (Hoege et al., 2002), and this
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idea was supported by genetic evidence showing that ubiquitination of PCNA was epistatic with POL30 (Stelter and Ulrich, 2003). In mammalian cells, in response to UV irradiation, PCNA is only monoubiquitinated, and this ubiquitination is not dependent on NER or on Pol. The monoubiquitinated PCNA is found exclusively in chromatin, where it interacts physically and specifically with Pol (Kannouche et al., 2004). This interaction, detected by coimmunoprecipitation, was found only with the modified form of PCNA. No interaction was detected with unmodified PCNA in cell extracts. The increased affinity for Pol of monoubiquitinated PCNA generated at blocked replication forks therefore provides an attractive mechanism for mediating the polymerase switch at the sites of DNA damage. As discussed at the beginning of this chapter, Rev1 is phylogenetically related to the Y-family polymerases, but it primarily inserts dCMP, and not the three remaining dNTPs. Recently, several groups have independently demonstrated that Pol aa residues 370–492 interact with the C-terminal 100 aa of Rev1 (Fig. 2A) (Guo et al., 2003; Ohashi et al., 2004; A. Tissier et al., 2004). The same C-terminal fragment of Rev1 also interacts with Pol, Pol (see Chapter 9), and Rev7 (see Chapter 6) (Guo et al., 2003; Ohashi et al., 2004; Tissier et al., 2004). Revl, like Pols and , is localized in replication factories (Tissier et al., 2004). These findings suggest that Revl might play a role as a molecular scaffold during the bypass of DNA lesions, facilitating aspects of the polymerase switching mechanisms.
IX. PROTECTION FROM CELLULAR EFFECTS OF DNA DAMAGE Early work showed that, in contrast to NER-defective XP cells, XP-V fibroblasts were only marginally sensitive to killing by UV light (although they are specifically sensitized by caffeine). They were, however, like NERdeficient XP cells, highly sensitive to the induction of mutations by UV light (Maher et al., 1976). We may now interpret these data as suggesting that TLS by Pol plays only a minor role in protecting human cells from killing by UV light but has a major role in safeguarding us from mutations. This is absolutely consistent with the finding that Pol in most instances inserts the ‘‘correct’’ nucleotides opposite CPDs (discussed above, in section III), and that in its absence, the substituting mechanism is much more error prone. In recent work supporting these findings, cell lines have been derived in which XP-V cells stably express a transfected POLH gene. UV mutagenesis was measured using two different systems, and the mutant frequency was restored to normal levels in the Pol-transfected cells (Stary et al., 2003; Taylor et al., 2003).
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The UV mutation spectra in XP-V cells differ significantly from those in normal or NER-defective XP cells. In particular, whereas most UV-induced mutations in the latter two cell types are C to T transitions, in XP-V cells there is a greater proportion of mutations at thymines (Wang et al., 1993). Thus, in the absence of Pol, the substituting enzymes make more errors at A or T residues. On the basis of biochemical data, it seems reasonable to hypothesize that Pol might be the enzyme responsible for such phenotype. Although the ability of Pol to replicate CPD-containing templates is much lower than that of Pol, it is higher than that of any other known eukaryotic polymerase and is very similar to that observed for the well-characterized E. coli translesion polymerase, PolV (Tang et al., 2000; Vaisman et al., 2002). In addition, the highly error-prone behavior of Pol at template T (Tissier et al., 2000a), and the increased accuracy of nucleotide incorporation opposite deaminated cytosines compared to other polymerases (Vaisman and Woodgate, 2001), is consistent with the mutation spectra observed in XP-V cells.
X. ROLES OF POL AND POL IN SOMATIC HYPERMUTATION Somatic hypermutation (SHM) is one of the mechanisms by which antibody diversity is generated and is the subject of Chapter 11. Models for the mechanism of SHM implicate an error-prone polymerase, and several groups have investigated the involvement of different Y-family polymerases in this process. In lymphocytes from three XP-V patients, the frequency of SHM was normal but the types of base changes were different: There was a decrease in mutations at A and T and a concomitant rise in mutations at G and C. These results suggest that more than one polymerase contributes to SHM, and that if one is absent, others compensate. The data suggest that Pol is involved in generating errors that occur predominantly at A and T and that another polymerase may preferentially generate errors opposite G and C (Zeng et al., 2001). In support of this idea, the SHM hotspots at A and T residues correlate with the Pol error spectrum (Rogozin et al., 2001). With regard to Pol, the data presently available are inconclusive. Targeted inactivation of the POLI gene in the Burkitt’s lymphoma BL2 cell line abolished SHM, suggesting that Pol is absolutely required for SHM in these cells (Faili et al., 2002). In contrast, the frequency and spectrum of SHM were indistinguishable in a 129-derived mouse strain that contains a stop codon early in the Poli gene and in Polþ mice (McDonald et al., 2003). These latter data suggest that Pol is dispensable for SHM. However, both systems have their peculiarities, and so extrapolation to the human population must be treated with caution. It therefore remains plausible that Pol, like Pol, may play a role in SHM, but that the various polymerases have redundant functions.
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REFERENCES Bebenek, K., Matsuda, T., Masutani, C., Hanaoka, F., and Kunkel, T. A. (2001a). Proofreading of DNA polymerase -dependent replication errors. J. Biol. Chem. 276, 2317–2320. Bebenek, K., Tissier, A., Frank, E. G., McDonald, J. P., Prasad, R., Wilson, S. H., Woodgate, R., and Kunkel, T. A. (2001b). 50 -Deoxyribose phosphate lyase activity of human DNA polymerase in vitro. Science 291, 2156–2159. Becherel, O. J., Fuchs, R. P., and Wagner, J. (2002). Pivotal role of the -clamp in translesion DNA synthesis and mutagenesis in E. coli cells. DNA Repair 1, 703–708. Boudsocq, F., Kokoska, R. J., Plosky, B. S., Vaisman, A., Ling, H., Kunkel, T. A., Yang, W., and Woodgate, R. (2004). Investigating the role of the little finger domain of Y-family DNA polymerases in low-fidelity synthesis and translesion replication. J. Biol. Chem. 279, 32932–32940. Bridges, B. A., and Woodgate, R. (1984). Mutagenic repair in Escherichia coli, X. The umuC gene product may be required for replication past pyrimidine dimers but not for the coding error in UV mutagenesis. Mol. Gen. Genet. 196, 364–366. Bridges, B. A., and Woodgate, R. (1985a). Mutagenic repair in Escherichia coli: Products of the recA gene and of the umuD and umuC genes act at different steps in UVinduced mutagenesis. Proc. Natl. Acad. Sci. USA 82, 4193–4197. Bridges, B. A., and Woodgate, R. (1985b). The two-step model of bacterial UV mutagenesis. Mutat. Res. 150, 133–139. Broughton, B. C., Cordonnier, A., Kleijer, W. J., Jaspers, N. G., Fawcett, H., Raams, A., Garritsen, V. H., Stary, A., Avril, M. F., Boudsocq, F., Masutani, C., Hanaoka, F., Fuchs, R. P., Sarasin, A., and Lehmann, A. R. (2002). Molecular analysis of mutations in DNA polymerase in xeroderma pigmentosum-variant patients. Proc. Natl. Acad. Sci. USA 99, 815–820. Carson, M. (1987). Ribbon model of macromolecules. J. Mol. Graphics 5, 103–106. Chiapperino, D., Kroth, H., Kramarczuk, I. H., Sayer, J. M., Masutani, C., Hanaoka, F., Jerina, D. M., and Cheh, A. M. (2002). Preferential misincorporation of purine nucleotides by human DNA polymerase opposite benzo[a]pyrene 7,8-diol 9,10epoxide deoxyguanosine adducts. J. Biol. Chem. 277, 11765–11771. Cleaver, J. E., and Carter, D. M. (1973). Xeroderma pigmentosum variants: Influence of temperature on DNA repair. J. Invest. Dermatol. 60, 29–32. Faili, A., Aoufouchi, S., Flatter, E., Gueranger, Q., Reynaud, C. A., and Weill, J. C. (2002). Induction of somatic hypermutation in immunoglobulin genes is dependent on DNA polymerase . Nature 419, 944–947. Frank, E. G., Sayer, J. M., Kroth, H., Ohashi, E., Ohmori, H., Jerina, D. M., and Woodgate, R. (2002). Translesion replication of benzo[a]pyrene and benzo[c]phenanthrene diolexpoxide adducts of deoxyadenosine and deoxyguanosine by human DNA polymerase . Nucleic Acids Res. 30, 5284–5292. Frank, E. G., Tissier, A., McDonald, J. P., Rapic-Otrin, V., Zeng, X., Gearhart, P. J., and Woodgate, R. (2001). Altered nucleotide misinsertion fidelity associated with poldependent replication at the end of a DNA template. EMBO J. 20, 2914–2922. Friedberg, E. C., Wagner, R., and Radman, M. (2002). Specialized DNA polymerases, cellular survival, and the genesis of mutations. Science 296, 1627–1630. Friedberg, E. C., Walker, G. C., and Siede, W. (1995). DNA repair and mutagenesis. American Society for Microbiology, Washington, DC. Goodman, M. F., Creighton, S., Bloom, L. B., and Petruska, J. (1993). Biochemical basis of DNA replication fidelity. Crit. Rev. Biochem. Mol. Biol. 28, 83–126.
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Goodman, M. F., and Fygenson, K. D. (1998). DNA polymerase fidelity: from genetics toward a biochemical understanding. Genetics 148, 1475–1482. Guo, C., Fischhaber, P. L., Luk-Paszyc, M. J., Masuda, Y., Zhou, J., Kamiya, K., Kisker, C., and Friedberg, E. C. (2003). Mouse Rev1 protein interacts with multiple DNA polymerases involved in translesion DNA synthesis. EMBO J. 22, 6621–6630. Haracska, L., Johnson, R. E., Unk, I., Phillips, B., Hurwitz, J., Prakash, L., and Prakash, S. (2001a). Physical and functional interactions of human DNA polymerase with PCNA. Mol. Cell Biol. 21, 7199–7206. Haracska, L., Johnson, R. E., Unk, I., Phillips, B. B., Hurwitz, J., Prakash, L., and Prakash, S. (2001b). Targeting of human DNA polymerase to the replication machinery via interaction with PCNA. Proc. Natl. Acad. Sci. USA 98, 14256–14261. Haracska, L., Prakash, S., and Prakash, L. (2002). Replication past O6-methylguanine by yeast and human DNA polymerase . Mol. Cell. Biol. 20, 8001–8007. Haracska, L., Washington, M. T., Prakash, S., and Prakash, L. (2001c). Inefficient bypass of an abasic site by DNA polymerase . J. Biol. Chem. 276, 6861–6866. Haracska, L., Yu, S. L., Johnson, R. E., Prakash, L., and Prakash, S. (2000). Efficient and accurate replication in the presence of 7,8-dihydro-8-oxoguanine by DNA polymerase . Nat. Genet. 25, 458–461. Hoege, C., Pfander, B., Moldovan, G. L., Pyrowolakis, G., and Jentsch, S. (2002). RAD6dependent DNA repair is linked to modification of PCNA by ubiquitin and SUMO. Nature 419, 135–141. Ishikawa, T., Uematsu, N., Mizukoshi, T., Iwai, S., Masutani, C., Hanaoka, F., Ueda, R., Ohmori, H., and Todo, T. (2001). Mutagenic and non-mutagenic bypass of DNA lesions by Drosophila DNA polymerases dpol and dpol. J. Biol. Chem. 276, 15155–15163. Itoh, T., Linn, S., Kamide, R., Tokushige, H., Katori, N., Hosaka, Y., and Yamaizumi, M. (2000). Xeroderma pigmentosum variant heterozygotes show reduced levels of recovery of replicative DNA synthesis in the presence of caffeine after ultraviolet irradiation. J. Invest. Dermatol. 115, 981–985. Johnson, R. E., Haracska, L., Prakash, S., and Prakash, L. (2001). Role of DNA polymerase in the bypass of a (6-4) TT photoproduct. Mol. Cell. Biol. 21, 3558–3563. Johnson, R. E., Kondratick, C. M., Prakash, S., and Prakash, L. (1999a). hRAD30 mutations in the variant form of Xeroderma pigmentosum. Science 285, 263–265. Johnson, R. E., Prakash, S., and Prakash, L. (1999b). Efficient bypass of a thyminethymine dimer by yeast DNA polymerase, po. Science 283, 1001–1004. Johnson, R. E., Prakash, S., and Prakash, L. (2000a). The human DINB1 gene encodes the DNA polymerase Pol. Proc. Natl. Acad. Sci. USA 97, 3838–3843. Johnson, R. E., Washington, M. T., Haracska, L., Prakash, S., and Prakash, L. (2000b). Eukaryotic polymerases and act sequentially to bypass DNA lesions. Nature 406, 1015–1019. Johnson, R. E., Washington, M. T., Prakash, S., and Prakash, L. (2000c). Fidelity of human DNA polymerase . J. Biol. Chem. 275, 7447–7450. Kannouche, P., Broughton, B. C., Volker, M., Hanaoka, F., Mullenders, L. H., and Lehmann, A. R. (2001). Domain structure, localization, and function of DNA polymerase , defective in xeroderma pigmentosum variant cells. Genes Dev. 15, 158–172. Kannouche, P., Ferna´ndez de Henestrosa, A. R., Coull, B., Vidal, A., Gray, C., Zicha, D., Woodgate, R., and Lehmann, A. R. (2002). Localisation of DNA polymerases and
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to the replication machinery is tightly co-ordinated in human cells. EMBO J. 21, 6246–6256. Kannouche, P. L., Wing, J., and Lehmann, A. R. (2004). Interaction of human DNA polymerase with monoubiquitinated PCNA; a possible mechanism for the polymerase switch in response to DNA damage. Mol. Cell 14, 491–500. Kato, T., and Shinoura, Y. (1977). Isolation and characterization of mutants of Escherichia coli deficient in induction of mutations by ultraviolet light. Mol. Gen. Genet. 156, 121–131. Kim, J. K., and Choi, B. S. (1995). The solution structure of DNA duplex-decamer containing the (6-4) photoproduct of thymidyly(30 ->50 )thymidine by NMR and relaxation matrix refinement. Eur. J. Biochem. 228, 849–854. Kulaeva, O. I., Koonin, E. V., McDonald, J. P., Randall, S. K., Rabinovich, N., Connaughton, J. F., Levine, A. S., and Woodgate, R. (1996). Identification of a DinB/UmuC homolog in the archeon Sulfolobus. solfataricus. Mutat. Res. 357, 245–253. Kunkel, T. A. (2004). DNA replication fidelity. J. Biol. Chem. 279, 16895–16898. Kuraoka, I., Robins, P., Masutani, C., Hanaoka, F., Gasparutto, D., Cadet, J., Wood, R. D., and Lindahl, T. (2001). Oxygen free radical damage to DNA. Translesion synthesis by human DNA polymerase and resistance to exonuclease action at cyclopurine deoxynucleoside residues. J. Biol. Chem. 276, 49283–49288. Kusumoto, R., Masutani, C., Iwai, S., and Hanaoka, F. (2002). Translesion synthesis by human DNA polymerase across thymine glycol lesions. Biochemistry 41, 6090–6099. Larimer, F. W., Perry, J. R., and Hardigree, A. A. (1989). The REV1 gene of Saccharomyces cerevisiae : Isolation, sequence and functional analysis. J. Bacteriol. 171, 230–237. Lee, J. H., Hwang, G. S., and Choi, B. S. (1999). Solution structure of a DNA decamer duplex containing the stable 30 T:G base pair of the pyrimidine(6-4)pyrimidone photoproduct [(6-4) adduct]: implications for the highly specific 30 T-> C transition of the (6-4) adduct. Proc. Natl. Acad. Sci. USA 96, 6632–6636. Lehmann, A. R., Kirk-Bell, S., Arlett, C. F., Paterson, M. C., Lohman, P. H., de WeerdKastelein, E. A., and Bootsma, D. (1975). Xeroderma pigmentosum cells with normal levels of excision repair have a defect in DNA synthesis after UV-irradiation. Proc. Natl. Acad. Sci. USA 72, 219–223. Ling, H., Boudsocq, F., Plosky, B. S., Woodgate, R., and Yang, W. (2003). Replication of a cis-syn thymine dimer at atomic resolution. Nature 424, 1083–1087. Ling, H., Boudsocq, F., Woodgate, R., and Yang, W. (2001). Crystal structure of a Yfamily DNA polymerase in action: a mechanism for error-prone and lesion-bypass replication. Cell 107, 91–102. Maher, V. M., Ouellette, L. M., Curren, R. D., and McCormick, J. J. (1976). Frequency of ultraviolet light-induced mutations is higher in Xeroderma pigmentosum variant cells than in normal human cells. Nature 261, 593–595. Masutani, C., Kusumoto, R., Iwai, S., and Hanaoka, F. (2000). Mechanisms of accurate translesion synthesis by human DNA polymerase . EMBO J. 19, 3100–3109. Masutani, C., Kusumoto, R., Yamada, A., Dohmae, N., Yokoi, M., Yuasa, M., Araki, M., Iwai, S., Takio, K., and Hanaoka, F. (1999). The XPV (Xeroderma pigmentosum variant) gene encodes human DNA polymerase . Nature 399, 700–704. Matsuda, T., Bebenek, K., Masutani, C., Hanaoka, F., and Kunkel, T. A. (2000). Low fidelity DNA synthesis by human DNA polymerase-. Nature 404, 1011–1013.
DNA POLYMERASES AND
225
Matsuda, T., Bebenek, K., Masutani, C., Rogozin, I. B., Hanaoka, F., and Kunkel, T. A. (2001). Error rate and specificity of human and murine DNA polymerase eta. J. Mol. Biol. 312, 335–346. McCulloch, S. D., Kokoska, R. J., Masutani, C., Iwai, S., Hanaoka, F., and Kunkel, T. A. (2004). Preferential cis-syn thymine dimer bypass by DNA polymerase occurs with biased fidelity. Nature 428, 97–100. McDonald, J. P., Frank, E. G., Plosky, B. S., Rogozin, I. B., Masutani, C., Hanaoka, F., Woodgate, R., and Gearhart, P. J. (2003). Identification of a nonsense mutation in DNA polymerase from 129-derived strains of mice and its effect on somatic hypermutation. J. Exp. Med. 198, 635–643. McDonald, J. P., Levine, A. S., and Woodgate, R. (1997). The Saccharomyces cerevisiae RAD30 gene, a homologue of Escherichia coli dinB and umuC, is DNA damage inducible and functions in a novel error-free postreplication repair mechanism. Genetics 147, 1557–1568. McDonald, J. P., Rapic-Otrin, V., Epstein, J. A., Broughton, B. C., Wang, X., Lehmann, A. R., Wolgemuth, D. J., and Woodgate, R. (1999). Novel human and mouse homologs of Saccharomyces cerevisiae DNA polymerase . Genomics 60, 20–30. McDonald, J. P., Tissier, A., Frank, E. G., Iwai, S., Hanaoka, F., and Woodgate, R. (2001). DNA polymerase iota and related Rad30-like enzymes. Phil. Trans. R. Soc. Lond. B Biol. Sci. 356, 53–60. McKenzie, G. J., Lee, P. L., Lombardo, M. J., Hastings, P. J., and Rosenberg, S. M. (2001). SOS mutator DNA polymerase IV functions in adaptive mutation and not adaptive amplification. Mol. Cell 7, 571–579. Mendelman, L. V., Boosalis, M. S., Petruska, J., and Goodman, M. F. (1989). Nearest neighbor influences on DNA polymerase insertion fidelity. J. Biol. Chem. 264, 14415–14423. Minko, I. G., Washington, M. T., Kanuri, M., Prakash, L., Prakash, S., and Lloyd, R. S. (2003). Translesion synthesis past acrolein-derived DNA adduct, gammahydroxypropanodeoxyguanosine, by yeast and human DNA polymerase . J. Biol. Chem. 278, 784–790. Nelson, J. R., Lawrence, C. W., and Hinkle, D. C. (1996a). Deoxycytidyl transferase activity of yeast REV1 protein. Nature 382, 729–731. Nelson, J. R., Lawrence, C. W., and Hinkle, D. C. (1996b). Thymine-thymine dimer bypass by yeast DNA polymerase . Science 272, 1646–1649. Ohashi, E., Murakumo, Y., Kanjo, N., Akagi, J., Masutani, C., Hanaoka, F., and Ohmori, H. (2004). Interaction of hREV1 with three human Y-family DNA polymerases. Genes Cells 9, 523–531. Ohashi, E., Ogi, T., Kusumoto, R., Iwai, S., Masutani, C., Hanaoka, F., and Ohmori, H. (2000). Error-prone bypass of certain DNA lesions by the human DNA polymerase . Genes Dev. 14, 1589–1594. Ohmori, H., Friedberg, E. C., Fuchs, R. P. P., Goodman, M. F., Hanaoka, F., Hinkle, D., Kunkel, T. A., Lawrence, C. W., Livneh, Z., Nohmi, T., Prakash, L., Prakash, S., Todo, T., Walker, G. C., Wang, Z., and Woodgate, R. (2001). The Y-family of DNA polymerases. Mol. Cell 8, 7–8. Prasad, R., Bebenek, K., Hou, E., Shock, D. D., Beard, W. A., Woodgate, R., Kunkel, T. A., and Wilson, S. H. (2003). Localization of the deoxyribose phosphate lyase active site in human DNA polymerase by controlled proteolysis. J. Biol. Chem. 278, 29649–29654. Rechkoblit, O., Zhang, Y., Guo, D., Wang, Z., Amin, S., Krzeminsky, J., Louneva, N., and Geacintov, N. E. (2002). trans-Lesion synthesis past bulky benzo[a]pyrene diol
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epoxide N2-dG and N6-dA lesions catalyzed by DNA bypass polymerases. J. Biol. Chem. 277, 30488–30494. Reuven, N. B., Arad, G., Maor-Shoshani, A., and Livneh, Z. (1999). The mutagenesis protein UmuC is a DNA polymerase activated by UmuD0 , RecA, and SSB and Is specialized for translesion replication. J. Biol. Chem. 274, 31763–31766. Rogozin, I. B., Pavlov, Y. I., Bebenek, K., Matsuda, T., and Kunkel, T. A. (2001). Somatic mutation hotspots correlate with DNA polymerase error spectrum. Nat. Immunol. 2, 530–536. Roush, A. A., Suarez, M., Friedberg, E. C., Radman, M., and Siede, W. (1998). Deletion of the Saccharomyces cerevisiae gene RAD30 encoding an Escherichia coli DinB homolog confers UV radiation sensitivity and altered mutability. Mol. Gen. Genet. 257, 686–692. Silvian, L. F., Toth, E. A., Pham, P., Goodman, M. F., and Ellenberger, T. (2001). Crystal structure of a DinB family error-prone DNA polymerase from Sulfolobus. solfataricus. Nat. Struct. Biol. 8, 984–989. Stary, A., Kannouche, P., Lehmann, A. R., and Sarasin, A. (2003). Role of DNA polymerase in the UV mutation spectrum in human cells. J. Biol. Chem. 278, 18767–18775. Steinborn, G. (1978). Uvm mutants of Escherichia coli K12 deficient in UV mutagenesis. I. Isolation of uvm mutants and their phenotypical characterization in DNA repair and mutagenesis. Mol. Gen. Genet. 165, 87–93. Stelter, P., and Ulrich, H. D. (2003). Control of spontaneous and damage-induced mutagenesis by SUMO and ubiquitin conjugation. Nature 425, 188–191. Tang, M., Bruck, I., Eritja, R., Turner, J., Frank, E. G., Woodgate, R., O’Donnell, M., and Goodman, M. F. (1998). Biochemical basis of SOS-induced mutagenesis in Escherichia coli: reconstitution of in vitro lesion bypass dependent on the UmuD0 2C mutagenic complex and RecA. Proc. Natl. Acad. Sci. USA 95, 9755–9760. Tang, M., Pham, P., Shen, X., Taylor, J.-S., O’Donnell, M., Woodgate, R., and Goodman, M. (2000). Roles of E. coli DNA polymerases IV and V in lesion-targeted and untargeted SOS mutagenesis. Nature 404, 1014–1018. Tang, M., Shen, X., Frank, E. G., O’Donnell, M., Woodgate, R., and Goodman, M. F. (1999). UmuD0 2C is an error-prone DNA polymerase, Escherichia coli, DNA pol V. Proc. Natl. Acad. Sci. USA 96, 8919–8924. Taylor, E. R., Dornan, E. S., Boner, W., Connolly, J. A., McNair, S., Kannouche, P., Lehmann, A. R., and Morgan, I. M. (2003). The fidelity of HPV16 E1/E2-mediated DNA replication. J. Biol. Chem. 278, 52223–52230. Tissier, A., Frank, E. G., McDonald, J. P., Iwai, S., Hanaoka, F., and Woodgate, R. (2000a). Misinsertion and bypass of thymine-thymine dimers by human DNA polymerase . EMBO J. 19, 5259–5266. Tissier, A., Kannouche, P., Reck, M. P., Lehmann, A. R., Fuchs, R. P., and Cordonnier, A. (2004). Co-localization in replication foci and interaction of human Y-family members, DNA polymerase pol, and REV1 protein. DNA Repair 3, 1503–1514. Tissier, A., McDonald, J. P., Frank, E. G., and Woodgate, R. (2000b). pol, a remarkably error-prone human DNA polymerase. Genes Dev. 14, 1642–1650. Tompkins, J. D., Nelson, J. L., Hazel, J. C., Leugers, S. L., Stumpf, J. D., and Foster, P. L. (2003). Error-prone polymerase, DNA polymerase IV, is responsible for transient hypermutation during adaptive mutation in Escherichia coli. J. Bacteriol. 185, 3469–3472. Trincao, J., Johnson, R. E., Escalante, C. R., Prakash, S., Prakash, L., and Aggarwal, A. K. (2001). Structure of the catalytic core of S. cerevisiae DNA polymerase . Implications for translesion DNA synthesis. Mol. Cell 8, 417–426.
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Vaisman, A., Frank, E. G., Iwai, S., Ohashi, E., Ohmori, H., Hanaoka, F., and Woodgate, R. (2003). Sequence context-dependent replication of DNA templates containing UV-induced lesions by human DNA polymerase . DNA Repair 2, 991–1006. Vaisman, A., Frank, E. G., McDonald, J. P., Tissier, A., and Woodgate, R. (2002). Poldependent lesion bypass in vitro. Mutat. Res. 510, 9–22. Vaisman, A., Masutani, C., Hanaoka, F., and Chaney, S. G. (2000). Efficient translesion replication past Oxaliplatin and Cisplatin GpG adducts by human DNA polymerase . Biochemistry 39, 4575–4580. Vaisman, A., Tissier, A., Frank, E. G., Goodman, M. F., and Woodgate, R. (2001). Human DNA polymerase promiscuous mismatch extension. J. Biol. Chem. 276, 30615–30622. Vaisman, A., and Woodgate, R. (2001). Unique misinsertion specificity of pol may decrease the mutagenic potential of deaminated cytosines. EMBO J. 20, 6520–6529. Wagner, J., Gruz, P., Kim, S. R., Yamada, M., Matsui, K., Fuchs, R. P. P., and Nohmi, T. (1999). The dinB gene encodes a novel Escherichia coli DNA polymerase (DNA pol IV) involved in mutagenesis. Mol. Cell 4, 281–286. Wang, Y. C., Maher, V. M., Mitchell, D. L., and McCormick, J. J. (1993). Evidence from mutation spectra that the UV hypermutability of xeroderma pigmentosum variant cells reflects abnormal, error-prone replication on a template containing photoproducts. Mol. Cell. Biol. 13, 4276–4283. Washington, M. T., Helquist, S. A., Kool, E. T., Prakash, L., and Prakash, S. (2003a). Requirement of Watson-Crick hydrogen bonding for DNA synthesis by yeast DNA polymerase . Mol. Cell. Biol. 23, 5107–5112. Washington, M. T., Johnson, R. E., Prakash, L., and Prakash, S. (2004). Human DNA Polymerase utilizes different nucleotide incorporation mechanisms dependent upon the template base. Mol. Cell. Biol. 24, 936–943. Washington, M. T., Johnson, R. E., Prakash, S., and Prakash, L. (1999). Fidelity and processivity of Saccharomyces cerevisiae DNA polymerase . J. Biol. Chem. 274, 36835–36838. Washington, M. T., Johnson, R. E., Prakash, S., and Prakash, L. (2001). Mismatch extension ability of yeast and human DNA polymerase . J. Biol. Chem. 276, 2263–2266. Washington, M. T., Prakash, L., and Prakash, S. (2003b). Mechanism of nucleotide incorporation opposite a thymine-thymine dimer by yeast DNA polymerase . Proc. Natl. Acad. Sci. USA 100, 12093–12098. Yamada, A., Masutani, C., Iwai, S., and Hanaoka, F. (2000). Complementation of defective translesion synthesis and UV light sensitivity in xeroderma pigmentosum variant cells by human and mouse DNA polymerase . Nucleic Acids Res. 28, 2473–2480. Yang, I. Y., Miller, H., Wang, Z., Frank, E. G., Ohmori, H., Hanaoka, F., and Moriya, M. (2003). Mammalian translesion DNA synthesis across an acrolein-derived deoxyguanosine adduct. Participation of DNA polymerase in error-prone synthesis in human cells. J. Biol. Chem. 278, 13989–13994. Yeiser, B., Pepper, E. D., Goodman, M. F., and Finkel, S. E. (2002). SOS-induced DNA polymerases enhance long-term survival and evolutionary fitness. Proc. Natl. Acad. Sci. USA 99, 8737–8741. Zeng, X., Winter, D. B., Kasmer, C., Kraemer, K. H., Lehmann, A. R., and Gearhart, P. J. (2001). DNA polymerase is an A-T mutator in somatic hypermutation of immunoglobulin variable genes. Nat. Immunol. 2, 537–541.
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Zhang, X., and Mathews, C. K. (1995). Natural DNA precursor pool asymmetry and base sequence context as determinants of replication fidelity. J. Biol. Chem. 270, 8401–8404. Zhang, Y., Yuan, F., Wu, X., Rechkoblit, O., Taylor, J. S., Geacintov, N. E., and Wang, Z. (2000a). Error-prone lesion bypass by human DNA polymerase . Nucleic Acids Res. 28, 4717–4724. Zhang, Y., Yuan, F., Wu, X., and Wang, Z. (2000b). Preferential incorporation of G opposite template T by the low-fidelity human DNA polymerase . Mol. Cell. Biol. 20, 7099–7108. Zhang, Y., Yuan, F., Xin, H., Wu, X., Rajpal, D. K., Yang, D., and Wang, Z. (2000c). Human DNA polymerase synthesizes DNA with extraordinarily low fidelity. Nucleic Acids Res. 28, 4147–4156. Zhou, B., Pata, J. D., and Steitz, T. A. (2001). Crystal structure of a DinB lesion bypass DNA polymerase catalytic fragment reveals a classic polymerase catalytic domain. Mol. Cell 8, 427–437.
PROPERTIES AND FUNCTIONS OF ESCHERICHIA COLI: POL IV AND POL V ˆ ME WAGNER By ROBERT P. FUCHS, SHINGO FUJII, AND JE´RO Cance´rogene`se et Mutagene`se Mole´culaire et Structurale, CNRS ESBS, 67400 Strasbourg, France
I. DNA Pol IV, the dinB Gene Product . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Discovery of the dinB Gene Product Activity . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. Biochemical Properties of DinB/Pol IV. . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . C. In Vivo Functions of Pol IV . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . D. Regulating the Access of Pol IV to Replication Intermediates . . . . . . . . . . .. . . . . . II. DNA Polymerase V, the umuDC Gene Product . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Genetic Requirements of Induced Mutagenesis . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. Biochemical Properties of Pol V . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . C. Switches Between Replicative and Specialized DNA Polymerases During Lesion Bypass . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . References . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . .
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Abstract Escherichia coli possesses two members of the newly discovered class of Y DNA polymerases (Ohmori et al., 2001): Pol IV (dinB) and Pol V (umuD 0 C). Polymerases that belong to this family are often referred to as specialized or error-prone DNA polymerases to distinguish them from the previously discovered DNA polymerases (Pol I, II, and III) that are essentially involved in DNA replication or error-free DNA repair. Y-family DNA polymerases are characterized by their capacity to replicate DNA, through chemically damaged template bases, or to elongate mismatched primer termini. These properties stem from their capacity to accommodate and use distorted primer templates within their active site and from the lack of an associated exonuclease activity. Even though both belong to the Y-family, Pol IV and Pol V appear to perform distinct physiological functions. Although Pol V is clearly the major lesion bypass polymerase involved in damage-induced mutagenesis, the role of Pol IV remains enigmatic. Indeed, compared to a wild-type strain, a dinB mutant exhibits no clear phenotype with respect to survival or mutagenesis following treatment with DNA-damaging agents. Subtler dinB phenotypes will be discussed below. Moreover, despite the fact that both dinB and umuDC loci are controlled by the SOS response, their constitutive and induced levels of expression are dramatically different. In noninduced cells, Pol V is undetectable by Western analysis. In contrast, it is estimated that there are about 250 copies 229 ADVANCES IN PROTEIN CHEMISTRY, Vol. 69
Copyright 2004, Elsevier Inc. All rights reserved. 0065-3233/04 $35.00
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of Pol IV per cell. On SOS induction, it is believed that only about 15 molecules of Pol V are assembled per cell (S. Sommer, personal communication), whereas Pol IV levels reach 2500 molecules. In fact, despite extensive knowledge of the individual enzymatic properties of all five E. coli DNA polymerases, much more work is needed to understand how their activities are orchestrated within a living cell.
I. DNA POL IV,
THE DINB
GENE PRODUCT
A. Discovery of the dinB Gene Product Activity The dinB (din stands for DNA damage inducible) locus was originally described by Kenyon and Walker in the early 1980s (Kenyon and Walker, 1980) when searching for E. coli genes whose expression is upregulated by DNA damage. Some 6 years later, Brotcorne-Lannoye and Maenhaut-Michel (1986) found the first dinB phenotype; namely, the inability of an E. coli strain carrying the original Mu d1(Ap Lac)–dinB fusion (isolated by Kenyon and Walker, 1980), which disrupted the dinB locus, to promote untargeted mutagenesis of bacteriophage . Untargeted mutagenesis is defined as mutations occurring in undamaged lambda phage as a result of being replicated within an ultraviolet (UV)-irradiated host cell. This pathway was found to be independent of UmuD0 C and RecA, the key players involved in induced chromosomal mutagenesis. Almost 10 years later, as part of the E. coli genome sequencing project in Japan, Ohmori et al. (1995) identified an ORF that they called dinP, with a putative LexA (the repressor of the SOS response) binding site within its promoter region. The 1056 bp long dinP ORF was subsequently shown to be identical to the dinB locus and encodes for a 351–amino acid, rather basic protein with a calculated pI and molecular weight of 9.4 and 39,516 Da, respectively (Kim et al., 1997). These authors further showed that disruption of the dinP gene abolishes the untargeted mutagenesis of phage and that overexpression of DinB dramatically increases the spontaneous mutation frequency in E. coli cells (hereafter designated the DinB mutator phenotype). Sequence analyses reveal that the dinB gene product shares strong local sequence homologies with UmuC-like proteins, which include Saccharomyces cerevisiae REV1 protein (Kulaeva et al., 1996; Larimer et al., 1989; Ohmori et al., 1995). Multiple sequence alignment of the UmuC-like proteins revealed five conserved motifs. Figure 1 shows the alignment of motifs I–V (Kulaeva et al., 1996) from six representative members of the UmuC-like proteins.
POL IV AND POL V IN E. COLI
FIG. 1. The five conserved domains in the DinB/UmuC/Rev1-like proteins. Multiple alignment (Clustal W; Thompson et al., 1994) of the Escherichia coli (eubacterial) DinB (Pol IV) and UmuC (Pol V), Sulfolobus solfataricus and Sulfolobus acidocaldarius (archeal) Dpo4 and Dbh, Saccharomyces cerevisiae (lower eukaryote), Rev1 and Homo sapiens (higher eukaryote) RAD30 A (XPV or Pol ) shows five conserved motifs in all these proteins. Arrows mark specific amino acids discussed within the text. Numbers indicate the distances (in amino acids) separating the conserved domains and the distances from the proteins termini. The bottom line (conservation) indicates the strictly conserved residues, two dots (:) indicate full conservation among groups of strongly related amino-acids (STA, NEQK, NHQK, NDEQ, QHRK, MILV, MILF, HY, or FYW); one dot (.) indicates conservation among groups of weakly related amino-acids (CSA, ATV, SAG, STNK, STPA, SGND, SNDEQK, NDEQHK, NEQHRK, FVLIM, or HFY).
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Kulaeva et al. (1996) also noted that ‘‘the most conserved portion of motif III, with adjacent invariant aspartic acid and glutamic acid residues preceded by a stretch of hydrophobic residues, resembled the Mg2þ-binding site that is conserved in a variety of ATPases’’ and that ‘‘the high level of sequence conservation between UmuC-like proteins from bacteria, archaea and eukaryotes suggests that these proteins may have an enzymatic activity, the nature of which remains to be determined’’ (pp. 250–251). This hypothesis was shown to be true later the same year when C. Lawrence and coworkers showed that the REV1 protein was endowed with a highly specific deoxycitidyl transferase activity in vitro (Nelson et al., 1996a). These two studies therefore provided the first hint that all members of this family might similarly possess a nucleotidyl transferase activity. Moreover, although no direct homology between UmuC/DinB/REV1 family members and known nucleotidyl transferases could be found at the primary sequence level, the few strictly conserved acidic residues in motifs I and III, as well as the positively charged residue (R or K) in motif II, are actually the same as those found in conserved motifs A, B, and C of known DNA polymerases (Delarue et al., 1990). Altogether, such observations provided the first clues as to the true biochemical functions of the DinB/UmuC-like proteins; namely, that they are DNA polymerases. As discussed in the following sections of this chapter, such activity was demonstrated not only for both dinB and umuC gene products of E. coli but also for other homologues from all three domains of life (see Chapter 6 by Lawrence; Chapter 7 by Vaisman, Lehmann, and Woodgate; and Chapter 9 by Ohashi, Ohmori, and Ogi). These polymerases form the so-called Y family of DNA polymerases and are sometimes referred to as error-prone, specialized, or TLS (for translesion synthesis) polymerases. Although these specialized DNA polymerases are clearly involved in the replication of damaged templates and are, as such, responsible for a high proportion of induced mutations, it should be kept in mind that some lesions can be replicated by the so-called replicative DNA polymerases, thus causing mutations as well (see later discussion).
B. Biochemical Properties of DinB/Pol IV 1. dinB Gene Encodes a Bona Fide Template-Directed DNA Polymerase, DNA Pol IV When the catalytic activity of a highly purified HisTag-DinB fusion protein was directly tested in the laboratory of T. Nohmi (NIHS, Tokyo) in 1998, it was found to possess a bona fide template-directed DNA polymerase activity (Fig. 2; Wagner et al., 1999), rather than a highly specific nucleotidyl transferase, as observed previously for REV1 (Nelson et al.,
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FIG. 2. The dinB gene encodes a bona fide template-directed DNA polymerase. The specificity of nucleotide incorporation by highly purified histidine-tagged DinB was investigated by reacting 30 nM of the specified substrates with 10 nM enzyme, with or without dNTP (125 M), as indicated for 10 minutes at 37 C. (Adapted from Wagner et al., 1999, with permission.)
1996) and Mgþþ ions were found to be essential for DNA polymerase activity. As noted above, primary sequence analysis of DinB and its homolog identified conserved residues potentially critical for the catalytic activity of the protein. Specifically, Asp8, Asp103, and Glu104 may correspond to the three acidic residues of ‘‘classic’’ DNA polymerases necessary to coordinate the divalent cations engaged in catalysis. Mutating any of these residues abolished the catalytic activity of this protein in vitro (Wagner et al., 1999), as well as the previously documented DinB mutator phenotype in vivo (Kim et al., 1997), thus directly linking the polymerase activity of DinB with its mutagenic properties. Likewise, mutating the highly conserved Arg49 in motif II severely compromised the activity of DinB (Wagner et al., 1999). Since then, structural studies performed on eukaryotic and prokaryotic homologs of DinB confirmed the critical roles of the three acidic residues in motifs I and III that are coordinating the divalent cations and of the positively charged residue in motif II that interacts with the incoming dNTP (Boudsocq et al., 2002; Ling et al., 2001; Silvian et al., 2001; Trincao et al., 2001; Zhou et al., 2001). Similarly, a highly conserved aromatic residue in Y polymerases (F13 and F12 in Pol IV of E. coli and in Dbh of Sulfolobus acidocaldarius [Boudsocq et al., 2004], respectively) allows discrimination against the incorporation of ribonucleotides (DeLucia et al., 2003; Shimizu et al., 2003; Zhou et al., 2001). Pre-steady-state studies of Y-family polymerases indicate that for nucleotidyl transfer reaction to occur, the polymerases have to undergo
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conformational changes before the chemical step (Fiala and Suo, 2004b; Washington et al., 2001). It was thus concluded that Y-family polymerases perform a fidelity check, involving an ‘‘induced-fit’’ mechanism. The classic induced-fit mechanism is defined as the finger subdomain undergoing a large conformational change from an ‘‘open’’ state to a ‘‘closed’’ state on binding of a correct incoming nucleotide to the polymerase (Johnson, 1993). If the incoming nucleotide does not match the template base, the proper ‘‘closed’’ state cannot be attained, which in turn prevents the chemical reaction. However, crystal structures of the Y-family polymerases, including two archaeal members (Dbh and Dpo4) and yeast Pol , determined to date indicate different conformational changes (see review by Yang, 2003). The catalytic core of the two closely related Y polymerases, Dbh and Dpo4, is virtually superimposable, despite the fact that Dbh is free of substrates and Dpo4 is complexed with DNA and a correct incoming nucleotide. Comparison of the Dbh apoprotein structure with the replicative DNA polymerases indicates that Dbh is already in the ‘‘closed’’ state, even without any substrate (Zhou et al., 2001). Structural changes of Dpo4 on association with DNA do occur, as the crystal structures reveal (Ling et al., 2004), but the domain that moves is not the finger subdomain but, rather, the ‘‘little finger,’’ far away from the replicating base pair. Even though the finger subdomain of Pol may undergo some conformational changes (Ling et al., 2003), the largest movement still occurs with the little finger. Therefore, the conformational step observed in Dpo4 and Pol by pre-steady-state kinetic analyses may differ from the classic ‘‘induced-fit’’ mechanism.
2. Processivity of Pol IV and Modulation of Its Activity Through Interaction with the Clamp When examined in a classical primer-extension assay using a simple primer-template DNA substrate, Pol IV produces a ladder-like pattern (see Fig. 2) illustrating its poor processivity. In fact, Pol IV appears to be strictly distributive, catalyzing the incorporation of only 1 nucleotide per binding event (Wagner et al., 1999). This feature may be related to an extremely low affinity of Pol IV for the 30 -OH extremity of a naked primer template substrate (Gruz et al., 2001; Wagner et al., 2000). The fact that no stable complex between DNA and Pol IV has been observed indicates that accessory factors allow the formation of a more stable polymerase/DNA substrate complex (Wagner et al., 2000). The clamp, which is also known as the processivity subunit of the replicative DNA polymerase (Pol III), fulfills this role. Once loaded onto DNA by the multisubunit clamp loader,
complex, the clamp confers the requested high processivity to the replicative DNA polymerase. It turns out that this sliding clamp dramatically
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alters the activity of Pol IV, increasing its affinity for the substrate, synthesis efficiency, and apparent processivity by two to four order of magnitude (Table I; Tang et al., 2000; Wagner et al., 2000). The dissociation rate of the Pol IV--DNA complex has been determined to be 0.005 sec1, which corresponds to a half-life of the complex of about 140 seconds. Thus, with a measured kpol of 2 nucleotides per second, thecalculated processivity of Pol IV- is 300–400 nucleotides. As discussed later in this chapter, the interaction between Pol IV and is essential for its role in mutagenesis. However, whether Pol IV synthesizes hundreds of nucleotide-long tracks in vivo remains to be determined.
3. Fidelity of Pol IV As opposed to classical E. coli DNA polymerases Pol I, Pol II, and Pol III core, Pol IV is devoid of any intrinsic 30 to 50 exonuclease (proofreading) activity (Wagner et al., 1999). This implies that the Pol IV fidelity is achieved solely by its capacity to discriminate between correct and incorrect base-pair formation. As shown in Fig. 2, Pol IV preferentially catalyses the incorporation of the correct nucleotides opposite all four template bases. Actually, steady-state kinetic analysis of the misinsertion capacities of Pol IV opposite all four template bases showed that Pol IV is, on average, only four- to fivefold less accurate than the catalytic -subunit of the replicative Pol III (Kobayashi et al., 2002; Tang et al., 2000). This observation suggests that, despite dinB being associated with mutagenesis, Pol IV is, at least at the incorporation step, a relatively faithful enzyme. Recent pre-steady-state kinetic studies of S. solfataricus DNA Pol IV (Dpo4) show that
TABLE I Modifications of Kinetic Parameters of Pol IV on Interaction with
koff (S1) kpol (S1) Processivity (nucleotides) KmDNA (nM) KmdNTP (M) Vmax (min1) Vmax/Km (M1 min1) a
Pol IV
Pol IV þ
Fold stimulation
n.d. n.d. 1a 7180b >2 000c n.d. 1.1 103c
0.005a 2a 400a 0.2a 0.12a 0.3a 2.5a
n.a. n.a. 400 35 900 >16 000 n.a. 2 270
Data from Wagner et al. (2000). Data from Gruz et al. (2001). c Data from Tang et al. (2000). (n.d.) not determined, (n.a.) not available. b
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the low fidelity of this enzyme mainly results from a weak discrimination between the correct and incorrect incoming nucleotide at the initial nucleotide binding step (Fiala and Suo, 2004b). This property is related to the observed solvent accessible and open structure of the Dpo4 active site (Ling et al., 2001). Another aspect of DNA polymerase low fidelity resides in the capacity to elongate mismatched primer/template termini. Although Pol IV is able to extend mismatches, it is less efficient than its human counterpart Pol (Kobayashi et al., 2002). However, as discussed next, efficient extension of mismatches by Pol IV and its homologs may be sequence context specific. Thus, it is worth noting that although Y-family DNA polymerases are generally considered as ‘‘error-prone’’ enzymes, huge differences in their specific properties are observed. The efficiencies of Pol IV in incorporation and extension of each mismatched base pair are summarized in Fig. 3 (Kobayashi et al., 2002), where the mismatches appearing in the upper-right corner of the panel are most efficiently generated and extended. These data are in relatively
FIG. 3. In vitro fidelity of Pol IV. Nucleotide misinsertion efficiencies (fins) are plotted versus mismatch extension efficiencies (fext). Data points above the line indicate lower values for the misinsertion efficiency compared to mismatch extension; inversely, data falling below the line indicate higher values for misinsertion compared to mismatch extension. (Reproduced from Kobayashi et al., 2002, with permission.)
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good agreement with the mutation spectra determined in vitro or in vivo, excepting T to A transversions that are not observed in vivo (Kobayashi et al., 2002; Wagner and Nohmi, 2000). The dinB-dependent untargeted mutagenesis pathway, as well as the DinB mutator activity, are associated with a specific increase in 1 frameshift mutagenesis within homopolymeric runs of 6 and more G:C base pairs (Kim et al., 1997; Wood and Hutchinson, 1984). The capacity of Pol IV to elongate frameshift intermediates was also confirmed in vitro (Wagner et al., 1999). However, analysis of forward mutational spectra obtained either in vivo or in vitro highlighted other specificities of Pol IV–induced mutagenesis. First, 1 frameshifts are strongly induced within both G:C and A:T single-nucleotide repeats, and more important, short repeats and nonrepetitive sequences are also susceptible to deletion events catalyzed by Pol IV (Kobayashi et al., 2002; Wagner and Nohmi, 2000). Second, base substitutions are also strongly induced by PolIV, representing 18%–30% of all mutagenic events observed in vitro and in vivo, respectively (Kobayashi et al., 2002; Wagner and Nohmi, 2000). Remarkably, changes toward G:C pairs represent 70% of all Pol IV–induced base substitutions. The mutational spectrum obtained in vivo by overexpression of Pol IV in E. coli indicates that a vast majority of both base substitutions toward G:C pairs as well as 1 frameshifts occur within a 50 -GX sequence context, in which X represents the deleted or mutated base (Wagner and Nohmi, 2000). Actually, kinetic and structural studies strongly support the notion that Pol IV and homologs such as the S. solfataricus Dpo4 and S. acidocaldarius Dbh polymerases may be particularly prone to ‘‘dNTP-stabilized misalignment’’ (Fig. 4) (Fiala and Suo, 2004a; Kobayashi et al., 2002; Kokoska et al., 2002; Potapova et al., 2002). Ling et al. (2001) described a crystal structure of a Dpo4 ternary complex that accommodates two undamaged template bases in its active site, with the incoming nucleotide paired with the second (50 ) template base. Recently, the same group reported a crystal structure of Dpo4 complexed with an abasic lesion, in which Dpo4 loops out the abasic site and uses the base 50 to the lesion to direct nucleotide incorporation (Ling et al., 2004). These structures illustrate two alternative mechanisms to generate a 1 frameshift invoked in the ‘‘dNTP stabilized misalignment’’ model presented in Fig. 4. Why this specific sequence context (50 -GX) represents a ‘‘hot spot’’ for mutagenesis induced by Pol IV remains to be determined.
4. Involvement of Pol IV in Lesion-Induced Mutagenesis and Translesion Synthesis In Vitro Although the umuDC genes products (Pol V) were known to be required for base substitution mutagenesis triggered by UV light or abasic sites (review by Smith and Walker, 1998), the discovery of the Y family of DNA
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FIG. 4. A ‘‘dNTP-stabilized misalignment’’ model for the Pol IV polymerases induced mutagenesis at 50 GX sequences. (a, b): An incoming dCTP residue is noncomplementary to the template cytosine residue in bold but can form a Watson– Crick base pair with the 50 adjacent guanosine, leaving the template cytosine unpaired. Such an intermediate, which is referred to as a ‘‘dNTP-stabilized misalignment’’ intermediate, has been observed in a ternary complex in which an incoming dGTP was unable to pair with the template G but could form a base pair with the 50 adjacent C residue (Ling et al., 2001). If a phophodiester bond between the misaligned incoming dNTP and the 30 -OH extremity of the primer can be formed (c), it can lead either to a 1 frameshift mutation (direct extension, pathway F) or to a base substitution after realignment and mismatch extension (pathway B). (Reproduced from Kokoska et al., 2002, with permission.)
polymerases led to the evaluation of the implication of other polymerases in lesion-induced mutagenesis and TLS. As far as E. coli Pol IV and archaeal Dbh and Dpo4 homologs are concerned, several studies addressed this question through in vitro experiments involving purified polymerases and, eventually, accessory proteins
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such as the processivity clamp (Boudsocq et al., 2001; Gruz et al., 2001; Kobayashi et al., 2002; Maor-Shoshani et al., 2003; Shen et al., 2002; Suzuki et al., 2001; Tang et al., 2000). From these studies, it turns out that Pol IV has the capacity to perform in vitro DNA synthesis across a wide panel of base modifications [8-oxoguanine; O6-methylguanine, uracil, abasic sites, N-2-acetylaminofluorene and N-2-aminofluorene modified guanines, cissyn cyclobutane dimers and 6-4 pyrimidine-pyrimidone TT photoproducts, 1,2-cisplatinated guanine adduct, and the major benzo[a]pyrene diol epoxide-N2-guanine (B[a]P guanine) adduct] with various efficiencies. In vitro bypass of abasic sites and B[a]P guanine adducts by Pol IV are particularly efficient (Maor-Shoshani et al., 2003; Shen et al., 2002). However, although Pol IV is clearly involved in the bypass of a site-specific B[a]P guanine adduct (see Section I.C.3 and Napolitano et al., 2000), it does not seem to be involved in the in vivo bypass of a site-specific AP site (Maor-Shoshani et al., 2003). The discrepancies between in vitro and in vivo data, with respect to AP site bypass, strongly indicate that the activity of these enzymes is regulated in vivo. In addition to TLS, direct incorporation of modified nucleotides into DNA constitutes a real threat to genome stability (Ames and Gold, 1991). Interestingly, oxidized DNA precursors 8-OH-dGTP and 2-OH-dATP are efficiently and erroneously incorporated into DNA by E. coli Pol IV and its archaeal homologs, Dbh and Dpo4 (Shimizu et al., 2003; T. Nohmi, personal communication). Actually, these polymerases preferentially incorporate 8-OH-dGTP opposite a template adenine, and 2-OH-dATP opposite a template guanine. Moreover, these events do not prevent further elongation of the nascent chain by these polymerases. As concluded by Shimizu et al. (2003), it turns out that Pol IV and homologs may promote mutagenesis through three distinct pathways that are spontaneous replication errors, TLS, and misincorporation of modified dNTPs during DNA synthesis.
C. In Vivo Functions of Pol IV 1. In Vivo Expression Level of Pol IV As noted above, the expression of the chromosomal dinB gene coding for Pol IV is regulated as part of the SOS response. However, the LexA-box located within the promoter region of dinB differs by 7 nucleotides from the 20-nucleotide consensus sequence (Ohmori et al., 1995), possibly explaining the high level of constitutive Pol IV expression (about 250 molecules per cell). Induction of the SOS response further increases its expression about 10-fold, yielding 2500 molecules per bacteria (Kim
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et al., 2001). Moreover, Layton and Foster (2003) recently showed that the general stress response sigma factor 38, encoded by the rpoS gene, also participates in the regulation of the Pol IV intracellular level. The maximal amount of Pol IV expressed from its single chromosomal locus actually occurs in stationary phase. Remarkably, it is 30-fold higher than the constitutive level, representing as much as 7500 Pol IV molecules per cell (Layton and Foster, 2003). Thus, if one combines the high affinity of Pol IV for -loaded DNA (see Section B.2. of this chapter) with its high cellular concentration, it is tempting to speculate that Pol IV may be constitutively part of the replisome, a notion that fits well with a previously suggested role for Pol IV in assisting the replicative polymerase during synthesis of particularly difficult sequence contexts (Wagner et al., 1999). It is difficult to evaluate whether such a role for Pol IV is compatible with the lack of a strong phenotype in dinB mutants.
2. Involvement of Pol IV in Long-Term Survival and Evolutionary Fitness of E. coli The high expression level of Pol IV indicates that there is an important and basic function for Pol IV in general metabolism that remains, however, to be discovered. In addition to the strict requirement for a functional dinB gene in the untargeted mutagenesis of phage, a function that provides no increase in bacterial fitness (Brotcorne-Lannoye and Maenhaut-Michel, 1986), two other functions of Pol IV have been illustrated in vivo. One is the involvement of this polymerase in TLS of various DNA lesions (see next paragraph), and the other is that the stationary phase-dependent induction of Pol IV may correlate with the capacity of the enzyme to enhance the long-term survival and evolutionary fitness of E. coli (Yeiser et al., 2002). This property is also shared with the other two SOSinduced DNA polymerases (Pol II and Pol V) and is characterized by the inability of individual SOS polymerase mutant strains to maintain themselves when cocultured with wild-type cells during long-term stationary phase incubation. Also, such mutant strains show partial defects in expressing the ‘‘growth advantage in stationary phase’’ (GASP) phenotype. The GASP phenotype, defined as the capacity of ‘‘aged’’ cells (that have experienced a long stationary phase period) to take over a ‘‘young’’ cell population depends on the appearance of new mutations that confer such a competitive advantage. Because the culture conditions used in the GASP experiments mimic long periods of nutrient stress that bacteria experience in natural environments (Morita, 1993), such phenotypes may, in fact, be physiologically relevant. The fact that high expression levels of Pol IV substantially increase spontaneous mutagenesis in both growing cells (untargeted mutagenesis) and nonproliferating cells under nonlethal
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selection (stationary phase or ‘‘adaptive’’ mutagenesis; (Kim et al., 1997; McKenzie et al., 2001; Slechta et al., 2003; Strauss et al., 2000; Wagner and Nohmi, 2000) fits well with the suggestion that Pol IV and homologs may contribute to adaptive strategies of bacteria, including pathogens (McKenzie and Rosenberg, 2001).
3. Pol IV–Dependent Translesion Synthesis and Mutagenesis In Vivo In E. coli, the expression of Pol II, a proofreading-proficient B-family DNA polymerase, and Pols IV and V, the two Y-family polymerases, is stimulated through the induction of the SOS response identified in this bacterium. All three polymerases are thought to participate in TLS in vivo (Napolitano et al., 2000). In particular, Pol IV participates in TLS and mutagenesis induced by B[a]P-guanine adduct, 4-nitroquinoline N-oxyde (4-NQO)–induced DNA lesions and oxidative DNA damages (Kim et al., 2001; LenneSamuel et al., 2000; Napolitano et al., 2000; Shen et al., 2002; Wagner et al., 2002). It should also be noted that lesion bypass is by no means restricted to Y-family DNA polymerases: Both replicative and specialized DNA polymerases allow cells to deal with the large diversity of ‘‘lesion/sequence contexts’’ situations that are encountered by the replication fork (Table II). This complexity is particularly well illustrated by the bypass of a sitespecific B[a]P-guanine adducts in E. coli. Within the GGA sequence context (the adducted base shown in bold), both ‘‘error-free’’ and 1 frameshift bypass pathways depend on both Pol IV and Pol V, whereas the G to T
TABLE II All Three SOS Polymerases are Involved in TLS Inducible polymerases involved in Lesion T(6–4)a BPDEa AAFa AAFa AAFb Oxydative lesionb Oxydative lesionb a
Sequence context
Error-free TLS
Mutagenic TLS (mutation type)
50 -AATT50 -GGG50 -GGG50 -gGCGCc50 -gGCGCGCc50 -gGCGCc-
Pol V Pol V þ Pol IV Pol V Pol V n.d. n.d.
Pol V (T to C) Pol V þ Pol IV (1) Pol V (1) Pol II (2) Pol II or Pol V (2) Pol V þ Pol IV (2)
50 -gGCGCGCc-
n.d.
Pol V þ [Pol IV or II] (2)
Data obtained using site specifically mono-modified plasmid DNA. Data obtained using randomly modified plasmid DNA. (n.d.) not determined. (Adapted from Wagner et al., 2002, with permission.) b
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transversion pathway requires neither of them and is most probably performed by Pol III (Lenne-Samuel et al., 2000; Napolitano et al., 2000). Interestingly, G to T transversions induced by the same BaP adduct within a TGT context (the adducted base shown in bold) were found to exclusively require Pol V (Yin et al., 2004). The requirement of multiple polymerases during lesion bypass, described as the DNA polymerase switch model (Cordonnier and Fuchs, 1999; Woodgate, 1999), raises the question as to the access of these polymerases to the different replication intermediates.
D. Regulating the Access of Pol IV to Replication Intermediates 1. Interaction between SOS Polymerases and Clamp is Essential for Translesion Synthesis and Mutagenesis As discussed in Section B.2, the Pol III holoenzyme processivity factor (the clamp) interacts directly with Pol IV, largely increasing its affinity for the substrates, synthesis efficiency, and apparent processivity in vitro. A small peptide (LVLGL), at the extreme C terminus of Pol IV, was shown to be essential for this interaction (Lenne-Samuel et al., 2002). Because the deletion of this motif abolishes the interaction between Pol IV and without affecting Pol IV’s intrinsic polymerase activity, it was possible to directly test the potential role of this interaction in vivo. The results obtained clearly demonstrate that the C-terminal peptide is essential for both B[a]P-guanine adduct bypass and the spontaneous mutator phenotype of Pol IV in vivo (Lenne-Samuel et al., 2002). Concomitant to these studies, Dalrymple et al. (2001) defined a consensus sequence, QL(S/D)LF, for most of the known eubacterial -interacting proteins, including E. coli Pol II and V. This allowed the construction of polB and umuC alleles that are mutated in the clamp interacting motifs. These mutants led to the demonstration that, similar to Pol IV, Pols II and V lesion bypass activities require a functional interaction with the clamp (Becherel et al., 2002). Similar interactions between the eukaryotic PCNA processivity clamp and eukaryotic Y polymerases , , and have also been demonstrated (see chapters by Vaisman, Lehmann and Woodgate and Ohashi and Ohmori). From these studies it can be inferred that, in vivo, the processivity clamps play a central role in lesion bypass.
2. Structural and Biochemical Studies Suggest Competition between Polymerases for Binding to the clamp All five DNA polymerases of E. coli interact with the clamp (Bonner et al., 1992; Lopez de Saro and O’Donnell, 2001; Stukenberg et al., 1991; Tang et al., 2000; Wagner et al., 2000). With the exception of Pol I, all these
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interactions involve an identified motif that resembles the consensus sequence defined by Dalrymple et al. (2001), thereby indicating competition of these polymerases for binding to the clamp. In an effort to gain details on the interaction between Pol IV and , Burnouf et al. (2004) solved the crystal structure of a complex between the clamp and the 16 residues C-terminal peptide (P16) of Pol IV, which contains the five last amino acids essential for the interaction (Lenne-Samuel et al., 2002). The authors showed that the seven C-terminal residues bind to a hydrophobic pocket located at the surface of the molecule between subdomains II and III (Fig. 5). Interestingly, this region strictly corresponds to the previously identified binding site of another ligand, the subunit of the complex ( Jeruzalmi et al., 2001). The subunit binds to via the previously mentioned consensus sequence (QAMSLF), and both peptides adopt a very similar conformation (Fig. 5). Biochemical studies further showed that all five E. coli DNA polymerases (as well as the subunit) bind to the same site on (Burnouf et al., 2004; Lopez de Saro et al., 2003). Particularly, the Pol IV P16 peptide is able to completely inhibit stimulation by the clamp of Pol III subunit DNA synthesis in vitro (Fig. 6; Burnouf et al., 2004). Similarly, a peptide derived from the binding site of Pol III subunit to the clamp inhibits the loading of the clamp by the complex as well as the -dependent Pol I DNA synthesis (Lopez de Saro et al., 2003). The fact that the same binding site on the clamp mediated the interaction with many enzymes involved in DNA metabolism, including essential ones such as the Pol III and subunits, leads the authors of these studies to conclude that this hydrophobic pocket may represent an attractive target for the development of a novel class of therapeutic agents (Burnouf et al., 2004; Lenne-Samuel et al., 2002; Lopez de Saro et al., 2003). Similarly, a ‘‘common’’ interacting pocket is conserved in PCNA, the eukaryotic counterpart of the clamp, and may be used as a target for the design of DNA replication inhibitors (Burnouf et al., 2004). These studies also point to the fact that regulating access of the different specialized polymerases to a replication fork may be simply mediated by the relative affinities of the different partners for the processivity clamp. Additional partners, such as RecA protein in E. coli, may also selectively modulate the access of specialized polymerases to bypass intermediates (see Section II). Moreover, posttranslational modifications of eukaryotic PCNA (Haracska et al., 2004; Hoege et al., 2002; Kannouche et al., 2004; Plosky and Woodgate, 2004; Stelter and Ulrich, 2003) or the existence of alternative clamps such as the Rad9—Rad1-Hus1 complex in eukaryotes (Kai and Wang, 2003, and references herein) and * in prokaryotes
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FIG. 5. Structure of Dpo4 polymerase and of the Pol IV and subunit -binding peptides bound to the clamp. (A) Crystal structure of Dpo4. The DNA and nucleotide in the ternary complex with Dpo4 are removed for clarity (Ling et al., 2001). The four structural domains common among Y polymerase are shown in red (palm), blue
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(Paz-Elizur et al., 1996; Skaliter et al., 1996a,b) potentially offer additional ways to differentially recruit specialized DNA polymerases. The multimeric nature of processivity clamps (review by Bruck and O’Donnell, 2001) such as the subunit in E. coli (homo-dimer) and PCNA in eukaryotes (homo-trimer) directly led to the formal possibility that two or three potentially distinct, interacting proteins may bind simultaneously to the clamp or PCNA, respectively thus constituting a ‘‘tool-belt’’ (Becherel et al., 2002). Although this possibility remains to be demonstrated, some studies support this notion. First, as mentioned above, homo-trimeric * clamps, which arise from the translation of the Cterminal 2/3 of the dnaN gene after the induction of the SOS response (Paz-Elizur et al., 1996; Skaliter et al., 1996a,b), may offer an additional binding site to -interacting proteins such as MutS, Lig1, and the SOS polymerases. Second, Bunting et al. (2003) recently solved the crystal structure of a complex between the E. coli clamp and the C-terminal domain of Pol IV, comprising the so called ‘‘little finger domain’’ (LF domain; Ling et al., 2001). This study shows that, in addition to the essential interaction between the C-terminal Pol IV peptide and the hydrophobic binding pocket on mentioned above, Pol IV makes an additional contact with the clamp through an external face of the LF domain. Interestingly, superimposition of the LF domain from a previously determined structure of Dpo4 polymerase onto the ‘‘Pol IV-LF/’’ complex reveals that Pol IV does not interact with the DNA primer template (‘‘OFF’’ position, Fig. 7; Bunting et al., 2003). To gain access to the DNA, the additional protein– protein interface between the LF domain and must be disrupted while the interaction via the C-terminal peptide is maintained (‘‘ON’’ position, Fig. 7). (finger), green (thumb), and purple (little finger). The five conserved sequence motifs shared by the Y-family polymerases are located in the palm, finger, and thumb subdomains, which form the catalytic core domain. The peptide at the C terminus that binds to the clamp is disordered in the crystal structure of Dpo4-DNA complex and is therefore represented by the yellow dashed line. This figure is generated using RIBBONS (Carson, 1987). (B) ribbon representation of the -ring with one P16 peptide (the 16 C-terminal residues of Pol IV) bound at the interface of subdomains II and III of monomer B. Only the seven C-terminal residues of the peptide were structured and modeled in the density map (colored in yellow). (C) detailed stereo view of peptide P16/ monomer interaction and comparison with the subunit. The accessible surface of the atoms in the -binding pocket is shown with hydrophobic residues in white, oxygen atoms in red, nitrogen atoms in blue, and sulphur atoms in orange. P16 peptide is shown in yellow, and the -interacting peptide of subunit is in blue. Part B was made with Pymol (DeLano, 2002). Residues 10–16 from P16 correspond to residues 345–351 of fulllength Pol IV. (Adapted from Burnouf et al., 2004, with permission.) (See Color Insert.)
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FIG. 6. Inhibition of the -dependent activity of Pol III subunit by the Pol IV binding peptide (P16). A SSB-coated synthetic 32P-labeled primer/template duplex (1 nM) allowing stable loading of the clamp is preincubated with (5 nM as a dimer; lanes 5–8 and 13–16) or without (lanes 1–4 and 9–12) the processivity factor, the clamp loader (1 nM), and increasing amounts (0, 1, 10, and 25 M final concentrations) of either control peptide (that does not contain the -binding motif; lanes 1–8) or P16 (the 16 C-terminal residues of Pol IV, which include the -binding motif; lanes 9–16). The DNA synthesis activity of the Pol III subunit on these substrates is then assayed in the presence of all four dNTPS (200 M) for 1 minute at room temperature. As shown in the left panel, the -independent activity of Pol III subunit, characterized by the appearance of elongation products of no more than 12 nucleotides, is not affected by the presence of either control of P16 peptides (lanes 1–4 and 9–12). In contrast, although the -dependent activity of the subunit (characterized by elongation products longer than 12 nucleotides) is not altered by the presence of the control peptide (lanes 5–8), increasing amounts of P16 peptide lead to almost complete inhibition of this specific activity (lanes 13–16). A quantitative analysis of the experiment is shown in the right part of the figure (black and open squares represent the calculated ratio of -dependent to -independent Pol III subunit activity in the presence of indicated concentrations of control and P16 peptides, respectively). (Adapted from Burnouf et al., 2004, with permission.)
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FIG. 7. ‘‘ON-OFF’’ model for Pol IV bound to the clamp. Model of a Pol IV type-Y DNA polymerase (red) bound to the clamp (gold) in the inactive, ‘‘locked-down’’ position (‘‘OFF’’ position, left part of the figure). The position of the polymerase was modeled by superimposing the little finger (LF) domain of the archaeal Dpo4 enzyme from the DNA complex (PDB code: 1JXL) onto the Pol IV-LF (blue) in the complex with the clamp described in Bunting et al. (2003). Modeled in this position, the polymerase makes no steric clashes with the clamp but cannot access the primer– template junction (primer strand in purple, template strand in green). As far as the clamp surface is not obstructed, such a complex may accommodate a second polymerase bound to the second monomer of . Transition to a productive complex (‘‘ON’’ position, right part of the figure) necessitates the disruption of the substantial protein–protein interface between the clamp and the LF domain of the Pol IV polymerase. In this ‘‘ON’’ configuration, contact with the clamp is maintained by the C-terminal clamp-binding peptide (blue), which tethers the enzyme to the replication complex. (Bunting, K. A., Roe, S. M., and Pearl, L. H. (EMBO) 20, 5883–5892 (2003); advance online publication, [doi:10.1038/nature xxxxx]) (See Color Insert.)
The ‘‘OFF’’ position may also be seen as a way to bind more than one ligand on a single multimeric clamp, with the polymerase being maintained away from the primer terminus while being present at a high local concentration. Finally, the occurrence of such a ‘‘tool belt’’ structure is strongly supported by the recent findings of Dionne et al. (2003), who showed that the heterotrimeric equivalent of PCNA in the archeon S. solfataricus is able to accommodate the simultaneous binding of three distinct partners, DNA polymerase B1, ligase I, and Fen I. Such a multifunctional complex ‘‘would facilitate a tight coupling of DNA synthesis and Okazaki fragment processing’’ in this archeon (Dionne et al., 2003).
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II. DNA POLYMERASE V,
THE UMUDC
GENE PRODUCT
A. Genetic Requirements of Induced Mutagenesis Chemical and UV irradiation physically alter DNA bases that can be converted into mutations on replication. Although many ‘‘bulky’’ lesions block DNA replication, some base modifications may simply change the coding properties of bases without blocking replication, thus inducing mutations during normal replication without any additional proteins (direct-acting mutagens such as alkylating agents producing O6Me-guanine or 04-Me-thymine adducts). In addition to these ‘‘small’’ alkylation lesions, replicative DNA polymerases may perform bypass of ‘‘bulky’’ adducts in vivo on their own. For instance, the adduct formed by the chemical carcinogen N-2-aminofluorene bound to the C8 position of guanine (G-AF), despite its bulkiness, is easily bypassed by the replicative polymerases (Bichara and Fuchs, 1985; Koffel-Schwartz et al., 1996). In contrast, the closely related G-AAF adducts, which contains an additional N-acetyl group, is a strong replication block that cannot be bypassed by replicative polymerases (Belguise-Valladier et al., 1994; Lindsley and Fuchs, 1994). For all lesions that block replication (indirect-acting mutagens such as UV-light induced lesions, polycyclic hydrocarbon adducts, etc.), genetic studies have revealed that mutagenesis requires additional gene products; namely, those encoded by the umuDC and recA loci in E. coli (Blanco et al., 1982; Dutreix et al., 1989; Kato and Shinoura, 1977; Steinborn, 1978; Sweasy et al., 1990). Although most studies were initially based on UV light as the mutagenic agent, many chemicals were subsequently shown to exhibit similar genetic requirements. As far as replication-blocking agents are concerned, RecA protein is the central component of the cellular response to this class of DNA-damaging agents (for a recent review, see Courcelle and Hanawalt, 2003). The key molecular intermediate is RecA*, the so-called activated form of RecA, which is produced when RecA monomers polymerize along single-stranded DNA, forming a nucleoprotein filament. Mutagenesis was found to require both UmuC and UmuD0 , a proteolytic fragment of UmuD (Burckhardt et al., 1988; Nohmi et al., 1988; Shinagawa et al., 1988). The processing of UmuD into UmuD0 requires the interaction of UmuD2 with the RecA nucleoprotein filament. This interaction stimulates a latent ability of UmuD2 to autodigest, resulting in the removal of its N-terminal 24 amino acids. In addition to derepression of the SOS response by mediating cleavage of the LexA repressor and activating UmuD0 by mediating cleavage of UmuD, it is believed that RecA protein has an additional and direct role in promoting mutagenesis (Dutreix et al., 1989; Sweasy et al., 1990). More recently, the clamp, the replication processivity
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factor, was shown to be an essential cofactor for umuDC-mediated mutagenesis (Becherel et al., 2002). On the basis of a wealth of genetic data, it was generally assumed that the role in mutagenesis of the umuDC gene products was to transiently alter the properties of the replication complex as to allow the replication machinery to proceed through damaged template bases. For many years, the major barrier in studying umuDC-dependent TLS in vitro was the difficulty in obtaining soluble UmuC protein. UmuC was first purified from insoluble inclusion bodies in the laboratory of the late Harison Echols by using a denaturation–renaturation protocol (Woodgate et al., 1989). Using renatured UmuC, it was demonstrated that monomeric UmuC interacts with a homodimer of UmuD0 to form a heterotrimeric UmuD0 2C complex (Woodgate et al., 1989). Replication experiments with UmuD0 2C demonstrated that the Umu complex, along with RecA and PolIII, could bypass a synthetic abasic site in vitro (Rajagopalan et al., 1992). A native UmuD0 2C complex was first purified by expressing recombinant UmuD0 and UmuC (Bruck et al., 1996) in a strain carrying chromosomal deletion of the umu locus (Woodgate, 1992). Analysis of the biochemical properties of the native UmuD0 2C complex allowed Tang et al. (1998) to recapitulate the earlier in vitro studies with denatured/ renatured UmuC from the Echols lab, but they also raised the intruiging possibility that the Umu complex may possess intrinsic polymerase activity (Tang et al., 1998). However, at that time, it was impossible to rule out the possibility that the polymerase activity was not from a minor contaminant of Pol II or Pol III. Indeed, using a different approach in which soluble UmuC was expressed as a Maltose binding protein (MBP) fusion, Reuven et al., (1998) did not detect any intrinsic polymerase activity associated with the MBP-UmuC fusion (Reuven et al. 1998). However, the following year Tang et al., purified native UmuD0 2C from strains lacking Pol II and carrying a thermosensitive Pol III and clearly demonstrated that UmuD0 2 possessed intrinsic polymerase activity and, as a consequence, was called Pol V (Tang et al., 1999). Similarly, the MBP-UmuC fusion protein was also found to possess intrinsic polymerase activity (Reuven et al., 1999). With the discovery of DNA polymerases specialized in TLS, it became clear that lesion bypass and mutagenesis would entail several DNA polymerase switches, with the replicative DNA polymerase being transiently replaced in the vicinity of the lesion by one (or several) specialized polymerases before resuming high-fidelity replication. This new paradigm for the mechanism of TLS and mutagenesis was dubbed the ‘‘DNA polymerase switch model’’ (Cordonnier and Fuchs, 1999). Next, we summarize the properties of Pol V and discuss some of the conflicting results that have been reported over the last several years with
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respect to Pol V biochemistry. We will describe a ‘‘minimal’’ lesion bypass assay that involves a long, circular, single-stranded template onto which the essential cofactors for Pol V-mediated TLS, that is, the clamp and RecA, are stably assembled. In a second part, we will summarize a recent attempt to reconstitute the whole process of TLS in vitro, recapitulating the switches between the replicative Pol III holoenzyme and the polymerases specialized in lesion bypass. Other aspects of umuDC biology, namely, its regulation via proteolysis and its activity in a checkpoint-type response, will not be discussed in this chapter (see the following recent papers for these aspects: Gonzalez and Woodgate, 2002; Opperman et al., 1999; Sutton et al., 1999).
B. Biochemical Properties of Pol V In vitro, Pol V was shown to be able to bypass a TT cis-syn cyclobutane dimer and TT (6-4) photoproduct with insertion specificities similar to those observed in vivo, thus establishing the physiological significance of Pol V (Tang et al., 2000). As already discussed in the literature (Sutton et al., 2000; Walker, 1998), because of major differences in their experimental design, the groups of Goodman and Livneh reported different biochemical requirements for Pol V–mediated in vitro bypass. We have recently reinvestigated the biochemical properties of native Pol V (Fujii et al., 2004). Our results shed some light on the previously published discrepancies, as we will outline below. In addition to obvious differences in their respective Pol V preparations as mentioned above, the groups of Livneh and Goodman used different DNA substrates (Fig. 8). Livneh and colleagues used a gapped circular plasmid with a single lesion located in a 350-nucleotide-long, single-stranded DNA gap (Tomer and Livneh, 1999). In contrast, Goodman and colleagues used primed linear templates, either a 240-mer synthetic oligonucleotide or a 7.2kb linear single-stranded phage DNA. In both cases, the single lesion was located only about 50 nucleotides from the 50 -end of the single-stranded template (Tang et al., 1999). Our own experimental design involves a primed, 2.7-kb-long, single-stranded circular plasmid DNA (Napolitano and Fuchs, 1997). Several common replication-blocking lesions were used by the three groups; namely, synthetic AP sites (Reuven et al., 1999; Tang et al., 1999), TT cis-syn cyclobutane dimer (Fujii et al., 2004; Tang et al., 2000), TT (6-4) photoproduct (Fujii et al., 2004; Tang et al., 2000), and the covalent adduct formed by the chemical carcinogen N-2-acetylaminofluorene with guanine (G-AAF) (Fujii et al., 2004). Although native Pol V (Fujii et al., 2004; Tang et al., 2000) and UmuC protein alone (Reuven et al., 1999) are able to copy
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FIG. 8. Basic requirements for Pol V–mediated translesion synthesis in vitro: sorting out the published literature. The groups of Dr. Goodman (Goodman, 2002), Dr. Livneh (Livneh, 2001), and ourselves (Fujii et al., 2004) have published work on the reconstitution of lesion bypass using Pol V and accessory factors. As outlined in this figure, the three studies differ in (i) the nature of the Pol V preparation and (ii) the structure of the primer template. The length of the single-stranded region downstream from the lesion site that appears to be a critical parameter (see text) is indicated for each substrate. Except for RecA protein that was found to be essential in all three studies, differences with respect to the requirements of SSB protein, ATP or ATP- S, and the clamp have been reported as discussed in the text and summarized in this figure.
undamaged DNA without any additional cofactor, TLS requires additional factors. All three groups found that RecA protein is essential for lesion bypass; however, some discrepancies with respect to the nature of the nucleotide triphosphate cofactor that is required to activate the RecA filament were reported. With the natural ATP cofactor, Goodman and Livneh groups reported low or high Pol V TLS activity, respectively (Pham et al., 2001; Reuven et al., 2001). With the slowly hydrolysable ATP- S analog, opposite results were reported, Livneh’s group reported severe inhibition, whereas Goodman’s group found high Pol V activity (Pham et al., 2001; Reuven et al., 2001). In our hands, both ATP and ATP- S were found to support high Pol V–mediated TLS activity provided that SSB protein was added when the RecA filament is activated with ATP-gammaS (Fujii et al., 2004). We suggest that SSB is required to disrupt the
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‘‘stablilized RecA filament’’ (Goodman, 2002; Pham et al., 2001) that is formed in the presence of ATP- S. In contrast, in the presence of the physiologically relevant RecA/ATP filament, SSB is not required for Pol V bypass activity (Fig. 9). This latter finding is in striking contrast with the findings of both Livneh and Goodman groups, who reported SSB to be an essential component of the Pol V–mediated lesion bypass reaction (Pham et al., 2001; Reuven et al., 2001). Goodman and coworkers suggested that SSB is necessary, acting together with the advancing Pol V molecule as a ‘‘locomotive cowcatcher’’ that dissociates RecA monomers from the 30 end of the filament (Pham et al., 2001). In contrast, our data indicate that Pol V can copy a single-stranded template covered with RecA in the absence of SSB (Fujii et al., 2004). To maintain a permanent contact between Pol V and the tip of the RecA filament, we suggest that the RecA filament ‘‘slides back’’ in a 30 ! 50 direction, with the RecA monomer dissociating from the 50 -end of the filament in an ATP-catalyzed reaction (Fig. 10b). In vivo, another cofactor essential for Pol V–mediated lesion bypass is the clamp, the replication processivity factor (Becherel et al., 2002). The clamp is an annular structure that encircles and freely slides along DNA (Kong et al., 1992). Therefore, its stable loading is best achieved on a circular DNA substrate that prevents it from sliding off the template. Using such a circular template, we found that Pol V–mediated TLS is highly stimulated ( 100-fold) by the presence of the clamp (Fujii et al., 2004). Initially, Livneh reported that Pol V–mediated TLS only required a basic four-component reaction set, including UmuC, UmuD0 , RecA, and SSB (Reuven et al., 1999). In a subsequent study, this group observed a threefold stimulation of TLS in the presence of the clamp (Maor-Shoshani and Livneh, 2002). Goodman’s group initially observed that and highly stimulate Pol V’s bypass activity (Tang et al., 1999), but that the stimulation was partially abrogated by the use of a RecA filament stabilized with ATP- S (Tang et al., 2000). In conclusion, it appears that Pol V bypasses efficiently all lesions that have been tested so far (AP sites, TT cyclobutane dimers, TT (6-4) photoproducts, G-AAF adducts), provided specific cofactors are present at the lesion site. These cofactors are RecA and the clamp. Although it can be speculated that the role of the clamp is to maintain Pol V in the vicinity of the substrate to allow it synthesize a TLS patch that is sufficiently long to allow subsequent elongation of the nascent primer by Pol III holoenzyme (see next paragraph), the exact role of RecA still remains to be discovered. One possibility may be that on interaction with the clamp, Pol V is locked in a position that prevents it from contacting the primer-template, an ‘‘OFF’’ position, as recently described for Pol IV (Fig. 7; Bunting et al., 2003). The specific role of RecA may be to interact with Pol V, thus
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FIG. 9. SSB protein is not required for robust Pol V-mediated TLS activity in the presence of a RecA/ATP filament. In these experiments, the amount of RecA protein (2 M) is stoichiometric with respect to the amount of single-stranded DNA present in the reaction mixture ( 5.4 M expressed in nucleotides), given that one RecA protein covers 3 nucleotides. While TLS efficiently occurs in the absence of SSB protein (34%), a low amount of SSB protein (10 nM corresponding, to less than 10% of the amount required for template saturation) stimulates the TLS reaction (66%). In contrast, increasing amounts of SSB (300 nM) strongly inhibit lesion bypass. The reaction conditions are as follows. A circular single-stranded template ( 2.7 kb) containing a single G-AAF adduct is primed with a 50 -32P-labeled 25-mer oligonucleotide, the 30 extremity being located 8 nucleotides upstream from the lesion site (L8 primer). A typical reaction mixture contains the DNA substrate (2 nM) in 20 mM Tris-HCl (pH 7.5), 4% glycerol, 8 mM DTT, 80 g/ml BSA, 8 mM MgCl2, dATP, dTTP, dGTP, dCTP (each 0.1 mM) and ATP (2.5 mM). The DNA substrate is preincubated with -clamp (50 nM as a dimer), -complex (10 nM), RecA protein, and SSB at the indicated concentrations at 30 C for 10 minutes. Reactions were initiated by adding Pol V (100 nM), and incubated for 20 minutes at 30 C. The reaction products were digested by EcoR I and analyzed by electrophoresis on a 10% denaturing polyacrylamide gel. The data are quantified as follows: primer utilization efficiency (initiation percentage) is calculated as the ratio of all products above the primer divided by the amount of total primer input. TLS efficiency (TLS%) is the ratio of all products above L0 (excluded) divided by the extent of primer utilization (sum of all bands above the primer band).
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FIG. 10. Switches between replicative and specialized DNA polymerases during translesion synthesis. (a) Pol III holoenzyme is able to elongate a primer in the vicinity of a lesion provided its 30 -extremity is located four or more nucleotides downstream from the lesion site. If the primer is shorter, the proofreading exonuclease that is associated with Pol III degrades the primer (Fujii and Fuchs, 2004). (b) Minimal conditions for robust Pol V-mediated TLS. Two cofactors are essential for efficient Pol V-mediated lesion bypass: (i) a DNA substrate onto which the clamp is stably loaded,
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correctly positioning the polymerase to engage the nascent primer terminus (‘‘ON’’ position). Although a long RecA filament activated with ATP is most likely to be the physiologically relevant cofactor, it has recently been suggested that mechanistically, a single RecA monomer may be sufficient for Pol V–mediated bypass (Pham et al., 2002).
C. Switches Between Replicative and Specialized DNA Polymerases During Lesion Bypass In vivo, the process of lesion bypass involves the recruitment of one or several specialized DNA polymerases that will temporarily replace the replicative polymerase in the vicinity of the lesion site. Indeed, for most replication-blocking lesions, the replicative polymerase will stop at the position preceding the lesion in the template (position L-1). Although for some lesions replicative polymerases may, in fact, be able to incorporate a nucleotide opposite the damaged template base, the kinetic barrier for further elongation will trigger its excision by the associated proofreading function, thus leading to futile insertion/excision cycles. DNA polymerases specialized in lesion bypass have the intrinsic capacity to copy lesion-containing templates. In addition, as these polymerases lack an associated proofreading exonuclease, they will not remove the newly incorporated nucleotides. A key factor for successful translesion synthesis will be to endowing these highly distributive enzymes with enough ‘‘processivity’’ to add several nucleotides onto the primer at once. Let us define and (ii) an extended single-stranded RecA/ATP filament assembled downstream from the lesion site. For efficient bypass, Pol V needs to interact simultaneously with the clamp and the 30 tip of the RecA filament. Formation of an extended RecA/ATP filament and stable loading of the clamp are best achieved on long single-stranded circular DNA templates. Under these conditions, SSB protein is not required for the bypass reaction. We suggest that to maintain a permanent contact between the advancing Pol V polymerase molecule and the 30 tip of the RecA filament, RecA molecules dissociate from the 50 end of the filament. (c) Overall lesion bypass scenario (Fujii and Fuchs, 2004): when Pol III associated with its clamp encounters, a noncoding lesion in the template a RecA filament formed on the single-stranded DNA region downstream the lesion site. This RecA filament, together with the clamp, forms a structure-specific template to which the bypass polymerase Pol V binds and mediates, in a single binding event—a TLS patch 20 nucleotides long on average (Fujii and Fuchs, 2004). The size distribution of the patches generated by Pol V is such that
75% of the patches are longer than 5 nucleotides, thus allowing subsequent elongation by the replicative polymerase. In contrast, the patches that are less than 5 nucleotides long ( 25%) are degraded by the Pol III–associated proofreading function, leading to an aborted bypass process.
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the ‘‘TLS patch’’ as being the fragment of DNA that is made by the specialized DNA polymerase opposite a template lesion. The TLS patch needs to be sufficiently long to prevent it from being degraded by the proofreading function on rebinding of the replicative polymerase. As will be discussed below, the interaction of the specialized DNA polymerases with the general replication processivity factor, that is, the clamp, is essential for this purpose. First, we determined the patch size made by Pol V, in a singlebinding event, under optimal conditions; that is, in the presence of an activated RecA/ATP filament and the clamp (as in Fig. 9). For a single G-AAF adduct, under standing-start conditions, we find that Pol V produces a large distribution of TLS patches, ranging from 1 to 60 nucleotides long, with the average length being about 20 nucleotides (Fujii and Fuchs, 2004). Similar results were obtained with the TT cyclobutane and the TT (6-4) photoproduct. In the absence of clamp, Pol V is completely distributive and is therefore unable to participate in a successful TLS event (Fujii and Fuchs, 2004). Second, we tested the capacity of Pol III holoenzyme (Pol III HE), the replicative machinery in E. coli, to extend a primer when a lesion is present in the template strand. It turns out that for a set of different lesions (AP site, TT cyclobutane dimer, TT(6-4) photoproduct, and G-AAF adduct), efficient primer elongation by Pol III HE requires the primer to reach 4–5 nucleotides beyond the lesion site (Fujii and Fuchs, 2004). On the basis of structural studies, it was suggested that replicative-type DNA polymerases possess a minor groove recognition domain that acts as a ‘‘sensor’’ capable of discriminating between correctly and incorrectly paired nucleotides within the four to five last base pairs (Kiefer et al., 1998). Disruption of these minor-groove interactions caused by the presence of the lesion within the four last base pairs of the nascent double-stranded DNA ( Johnson et al., 2003) is likely to cause stalling of DNA synthesis and, as a consequence, to activate degradation via proofreading. Under our experimental conditions, it turns out that about 75% of the TLS patches made by Pol V when it bypasses a single G-AAF adduct are longer than 5 nucleotides and will therefore be converted into TLS events on rebinding of Pol III replicase (Fujii and Fuchs, 2004). Conversely, about 25% of TLS patches are shorter than 5 nucleotides and will be degraded by the proofreading function of Pol III. These aborted TLS events will enter a new bypass attempt. The average patch size of 20 nucleotides may appear as a reasonable trade-off between ensuring efficient TLS (75%, in a single attempt) and preventing a heavy load of untargeted mutations. The level of untargeted mutation made by Pol V can be estimated as follows: Given its fidelity on undamaged DNA (103 to 104, Maor-Shoshani et al., 2000; Tang et al., 2000), Pol V will produce only one untargeted mutation per 50–500 TLS events, a low level of point mutations that will potentially be corrected by mismatch repair. We suggest
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that the process of polymerase switching during TLS as described here in E. coli may represent a simplified paradigm for lesion bypass in eukaryotes. As already discussed here in section I.D.3, the control of polymerase switching in yeast and human cells appears to be much more complicated, as it involves PCNA and its posttranslationally modified forms as well as alternative sliding clamps.
Acknowledgments We gratefully thank Drs. Roger Woodgate and Wei Yang for critical reading and suggestions.
REFERENCES Ames, B. N., and Gold, L. S. (1991). Endogenous mutagens and the causes of aging and cancer. Mutat. Res. 250, 3–16. Becherel, O. J., Fuchs, R. P. P., and Wagner, J. (2002). Pivotal role of the -clamp in translesion DNA synthesis and mutagenesis in E. coli cells. DNA Repair 1, 703–708. Belguise-Valladier, P., Maki, H., Sekiguchi, M., and Fuchs, R. P. P. (1994). Effect of single DNA lesions on in vitro replication with DNA polymerase III holoenzyme: Comparison with other polymerases. J. Mol. Biol. 236, 151–164. Bichara, M., and Fuchs, R. P. P. (1985). Binding and mutation spectra of the carcinogen N-2-aminofluorene. A correlation between the conformation of the premutagenic lesion and the mutation specificity. J. Mol. Biol. 183, 341–351. Blanco, M., Herrera, G., Collado, P., Rebollo, J. E., and Botella, L. M. (1982). Influence of RecA protein on induced mutagenesis. Biochimie 64, 633–636. Bonner, C. A., Stukenberg, P. T., Rajagopalan, M., Eritja, R., O’Donnell, M., McEntee, K., Echols, H., and Goodman, M. F. (1992). Processive DNA synthesis by DNA polymerase II mediated by DNA polymerase III accessory proteins. J. Biol. Chem. 267, 11431–11438. Boudsocq, F., Iwai, S., Hanaoka, F., and Woodgate, R. (2001). Sulfolobus solfataricus P2 DNA polymerase IV (Dpo4): An archaeal DinB-like DNA polymerase with lesionbypass properties akin to eukaryotic pol . Nucleic Acids Res. 29, 4607–4616. Boudsocq, F., Kokoska, R. J., Plosky, B. S., Vaisman, A., Ling, H., Kunkel, T. A., Yang, W., and Woodgate, R. (2004). Investigating the role of the little finger domain of Yfamily DNA polymerases in low-fidelity synthesis and translesion replication. J. Biol. Chem 279, 32932–32940. Boudsocq, F., Ling, H., Yang, W., and Woodgate, R. (2002). Structure-based interpretation of missense mutations in Y-family DNA polymerases and their implications for polymerase function and lesion bypass. DNA Repair (Amst) 1, 343–358. Brotcorne-Lannoye, A., and Maenhaut-Michel, G. (1986). Role of RecA protein in untargeted UV mutagenesis of bacteriophage lambda: Evidence for the requirement for the dinB gene. Proc. Natl. Acad. Sci. USA 83, 3904–3908. Bruck, I., and O’Donnell, M. (2001). The ring-type polymerase sliding clamp family. Genome Biol. 2, reviews3001.1–reviews3001.3.
258
FUCHS ET AL.
Bruck, I., Woodgate, R., McEntee, K., and Goodman, M. F. (1996). Purification of a soluble UmuD’C complex from Escherichia coli. Cooperative binding of UmuD’C to single-stranded DNA. J. Biol. Chem. 271, 10767–10774. Bunting, K. A., Roe, S. M., and Pearl, L. H. (2003). Structural basis for recruitment of translesion DNA polymerase Pol IV/DinB to the beta-clamp. EMBO J. 22, 5883–5892. Burckhardt, S. E., Woodgate, R., Scheuermann, R. H., and Echols, H. (1988). UmuD mutagenesis protein of Escherichia coli: Overproduction, purification, and cleavage by RecA. Proc. Natl. Acad. Sci. USA 85, 1811–1815. Burnouf, D. Y., Olieric, V., Wagner, J., Fujii, S., Reinbolt, J., Fuchs, R. P., and Dumas, P. (2004). Structural and biochemical analysis of sliding clamp/ligand interactions suggest a competition between replicative and translesion DNA polymerases. J. Mol. Biol. 335, 1187–1197. Carson, M. (1987). Ribbon models of macromolecules. J. Mol. Graphics 5, 103–106. Cordonnier, A., Lehmann, A. R., and Fuchs, R. P. P. (1999). Impaired translesion synthesis in Xeroderma pigmentosum variant extracts. Mol. Cell. Biol. 19, 2206–2211. Cordonnier, A. M., and Fuchs, R. P. (1999). Replication of damaged DNA: Molecular defect in Xeroderma pigmentosum variant cells. Mutat Res. 435, 111–119. Courcelle, J., and Hanawalt, P. C. (2003). RecA-dependent recovery of arrested DNA replication forks. Annu. Rev. Genet. 37, 611–646. Dalrymple, B. P., Kongsuwan, K., Wijffels, G., Dixon, N. E., and Jennings, P. A. (2001). A universal protein-protein interaction motif in the eubacterial DNA replication and repair systems. Proc. Natl. Acad. Sci. USA 98, 11627–11632. DeLano, W. L. (2002). The PyMOL molecular graphics system. DeLano Scientific, San Carlos, CA,USA. http://pymol.sourceforge.net Delarue, M., Poch, O., Tordo, N., Moras, D., and Argos, P. (1990). An attempt to unify the structure of polymerases. Protein Eng. 3, 461–467. DeLucia, A. M., Grindley, N. D., and Joyce, C. M. (2003). An error-prone family Y DNA polymerase (DinB homolog from Sulfolobus solfataricus) uses a ‘‘steric gate’’ residue for discrimination against ribonucleotides. Nucleic Acids Res. 31, 4129–4137. Dionne, I., Nookala, R. K., Jackson, S. P., Doherty, A. J., and Bell, S. D. (2003). A heterotrimeric PCNA in the hyperthermophilic archaeon Sulfolobus solfataricus. Mol. Cell 11, 275–282. Dutreix, M., Moreau, P. L., Bailone, A., Galibert, F., Battista, J. R., Walker, G. C., and Devoret, R. (1989). New recA mutations that dissociate the various RecA protein activities in Escherichia coli provide evidence for an additional role for RecA protein in UV mutagenesis. J. Bact. 171, 2415–2423. Fiala, K. A., and Suo, Z. (2004a). Pre-steady-state kinetic studies of the fidelity of Sulfolobus solfataricus P2 DNA polymerase IV. Biochemistry 43, 2106–2115. Fiala, K. A., and Suo, Z. (2004b). Mechanism of DNA polymerization catalyzed by Sulfolobus solfataricus P2 DNA polymerase IV. Biochemistry 43, 2116–2125. Fujii, S., and Fuchs, R. P. (2004). Defining the position of the switches between replicative and bypass DNA polymerases. EMBO. J. in press. Fujii, S., Gasser, V., and Fuchs, R. P. (2004). The biochemical requirements of DNA polymerase V-mediated translesion synthesis revisited. J. Mol. Biol. 341, 405–417. Gonzalez, M., and Woodgate, R. (2002). The ‘‘tale’’ of UmuD and its role in SOS mutagenesis. Bioessays 24, 141–148.
POL IV AND POL V IN E. COLI
259
Goodman, M. F. (2002). Error-prone repair DNA polymerases in prokaryotes and eukaryotes. Annu. Rev. Biochem. 71, 17–50. Gruz, P., Pisani, F. M., Shimizu, M., Yamada, M., Hayashi, I., Morikawa, K., and Nohmi, T. (2001). Synthetic activity of Sso DNA polymerase Y1, an archaeal DinB-like DNA polymerase, is stimulated by processivity factors proliferating cell nuclear antigen and replication factor C. J. Biol. Chem. 276, 47394–47401. Haracska, L., Torres-Ramos, C. A., Johnson, R. E., Prakash, S., and Prakash, L. (2004). Opposing effects of ubiquitin conjugation and SUMO modification of PCNA on replicational bypass of DNA lesions in Saccharomyces cerevisiae. Mol. Cell. Biol. 24, 4267–4274. Hoege, C., Pfander, B., Moldovan, G. L., Pyrowolakis, G., and Jentsch, S. (2002). RAD6dependent DNA repair is linked to modification of PCNA by ubiquitin and SUMO. Nature 419, 135–141. Jeruzalmi, D., Yurieva, O., Zhao, Y., Young, M., Stewart, J., Hingorani, M., O’Donnell, M., and Kuriyan, J. (2001). Mechanism of processivity clamp opening by the delta subunit wrench of the clamp loader complex of E. coli DNA polymerase III. Cell 106, 417–428. Johnson, K. A. (1993). Conformational coupling in DNA polymerase fidelity. Annu. Rev. Biochem. 62, 685–713. Johnson, S. J., Taylor, J. S., and Beese, L. S. (2003). Processive DNA synthesis observed in a polymerase crystal suggests a mechanism for the prevention of frameshift mutations. Proc. Natl. Acad. Sci. USA 100, 3895–3900. Kai, M., and Wang, T. S. (2003). Checkpoint activation regulates mutagenic translesion synthesis. Genet Dev. 17, 64–76. Kannouche, P. L., Wing, J., and Lehmann, A. R. (2004). Interaction of human DNA polymerase with monoubiquitinated PCNA; a possible mechanism for the polymerase switch in response to DNA damage. Mol. Cell 14, 491–500. Kato, T., and Shinoura, Y. (1977). Isolation and characterization of mutants of E. coli deficient in induction of mutagenesis by UV light. Mol. Gen. Genet. 156, 121–131. Kenyon, C. J., and Walker, G. C. (1980). DNA-damaging agents stimulate gene expression at specific loci in Escherichia coli. Proc. Natl. Acad. Sci. USA 77, 2819–2823. Kiefer, J. R., Mao, C., Braman, J. C., and Beese, L. S. (1998). Visualizing DNA replication in a catalytically active Bacillus DNA polymerase crystal. Nature 391, 304–307. Kim, S. R., Maenhaut, M. G., Yamada, M., Yamamoto, Y., Matsui, K., Sofuni, T., Nohmi, T., and Ohmori, H. (1997). Multiple pathways for SOS-induced mutagenesis in Escherichia coli: An overexpression of dinB/dinP results in strongly enhancing mutagenesis in the absence of any exogenous treatment to damage DNA. Proc. Natl. Acad. Sci. USA 94, 13792–13797. Kim, S. R., Matsui, K., Yamada, M., Gruz, P., and Nohmi, T. (2001). Roles of chromosomal and episomal dinB genes encoding DNA pol IV in targeted and untargeted mutagenesis in Escherichia coli. Mol. Genet. Genomics 266, 207–215. Kobayashi, S., Valentine, M. R., Pham, P., O’Donnell, M., and Goodman, M. F. (2002). Fidelity of Escherichia coli DNA polymerase IV. Preferential generation of small deletion mutations by dNTP-stabilized misalignment. J. Biol. Chem. 277, 34198–34207. Koffel-Schwartz, N., Coin, F., Veaute, X., and Fuchs, R. P. P. (1996). Cellular strategies for accomodating replication-hindering adducts in DNA: Control by the SOS response in E. coli. Proc. Natl. Acad. Sci. USA 93, 7805–7810.
260
FUCHS ET AL.
Kokoska, R. J., Bebenek, K., Boudsocq, F., Woodgate, R., and Kunkel, T. A. (2002). Low fidelity DNA synthesis by a Y family DNA polymerase due to misalignment in the active site. J. Biol. Chem. 277, 19633–19638. Kong, X. P., Onrust, R., O’Donnell, M., and Kuriyan, J. (1992). Three-dimensional structure of the subunit of E. coli DNA polymerase III holoenzyme: A sliding DNA clamp. Cell 69, 425–437. Kulaeva, O. I., Koonin, E. V., McDonald, J. P., Randall, S. K., Rabinovich, N., Connaughton, J. F., Levine, A. S., and Woodgate, R. (1996). Identification of a DinB/UmuC homolog in the archeon Sulfolobus solfataricus. Mutat. Res. 357, 245–253. Larimer, F. W., Perry, J. R., and Hardigree, A. A. (1989). The REV1 gene of Saccharomyces cerevisiae : Isolation, sequence, and functional analysis. J. Bacteriol. 171, 230–237. Layton, J. C., and Foster, P. L. (2003). Error-prone DNA polymerase IV is controlled by the stress-response sigma factor, RpoS, in Escherichia coli. Mol. Microbiol. 50, 549–561. Lenne-Samuel, N., Janel-Bintz, R., Kolbanovskiy, A., Geacintov, N. E., and Fuchs, R. P. (2000). The processing of a Benzo(a)pyrene adduct into a frameshift or a base substitution mutation requires a different set of genes in Escherichia coli. Mol. Microbiol. 38, 299–307. Lenne-Samuel, N., Wagner, J., Etienne, H., and Fuchs, R. P. (2002). The processivity factor controls DNA polymerase IV traffic during spontaneous mutagenesis and translesion synthesis in vivo. EMBO Rep. 3, 45–49. Lindsley, J. E., and Fuchs, R. P. P. (1994). Use of single-turnover kinetics to study adduct bypass by T7 DNA polymerase. Biochemistry 33, 764–772. Ling, H., Boudsocq, F., Plosky, B. S., Woodgate, R., and Yang, W. (2003). Replication of a cis-syn thymine dimer at atomic resolution. Nature 424, 1083–1087. Ling, H., Boudsocq, F., Woodgate, R., and Yang, W. (2001). Crystal structure of a Y-family DNA polymerase in action: A mechanism for error-prone and lesion-bypass replication. Cell 107, 91–102. Ling, H., Boudsocq, F., Woodgate, R., and Yang, W. (2004). Snapshots of replication through an abasic lesion; structural basis for base substitutions and frameshifts. Mol. Cell 13, 751–762. Livneh, Z. (2001). DNA damage control by novel DNA polymerases: Translesion replication and mutagenesis. J. Biol. Chem. 276, 25639–25642. Lopez de Saro, F. J., Georgescu, R. E., Goodman, M. F., and O’Donnell, M. (2003). Competitive processivity-clamp usage by DNA polymerases during DNA replication and repair. EMBO J. 22, 6408–6418. Lopez de Saro, F. J., and O’Donnell, M. (2001). Interaction of the beta sliding clamp with MutS, ligase, and DNA polymerase I. Proc. Natl. Acad. Sci. USA 98, 8376–8380. Maor-Shoshani, A., Bacher Reuven, N., Tomer, G., and Livneh, Z. (2000). Highly mutagenic replication by DNA polymerase V (UmuC) provides a mechanistic basis for SOS untargeted mutagenesis. Proc. Natl. Acad. Sci. USA 97, 565–570. Maor-Shoshani, A., Hayashi, K., Ohmori, H., and Livneh, Z. (2003). Analysis of translesion replication across an abasic site by DNA polymerase IV of Escherichia coli. DNA Repair (Amst.) 2, 1227–1238. Maor-Shoshani, A., and Livneh, Z. (2002). Analysis of the stimulation of DNA polymerase V of Escherichia coli by processivity proteins. Biochemistry 41, 14438–14446.
POL IV AND POL V IN E. COLI
261
McKenzie, G. J., Lee, P. L., Lombardo, M. J., Hastings, P. J., and Rosenberg, S. M. (2001). SOS mutator DNA polymerase IV functions in adaptive mutation and not adaptive amplification. Mol. Cell 7, 571–579. McKenzie, G. J., and Rosenberg, S. M. (2001). Adaptive mutations, mutator DNA polymerases and genetic change strategies of pathogens. Curr. Opin. Microbiol. 4, 586–594. Morita, R. Y. (1993). Bioavailability of energy and the starvation state. In ‘‘Starvation in Bacteria,’’ (S. Kjelleberg, Ed.), pp. 1–23. Plenum Press, New York. Napolitano, R., Janel-Bintz, R., Wagner, J., and Fuchs, R. P. (2000). All three SOSinducible DNA polymerases (Pol II, Pol IV, and Pol V) are involved in induced mutagenesis. EMBO J. 19, 6259–6265. Napolitano, R. L., and Fuchs, R. P. P. (1997). New strategy for the construction of single stranded plasmids with single mutagenic lesions. Chem. Res. Toxicol. 10, 667–671. Nelson, J. R., Lawrence, C. W., and Hinkle, D. C. (1996a). Deoxycytidyl transferase activity of yeast REV1 protein. Nature 382, 729–731. Nohmi, T., Battista, J. R., Dodson, L. A., and Walker, G. C. (1988). RecA-mediated cleavage activates UmuD for mutagenesis: mechanistic relationship between transcriptional derepression and posttranslational activation. Proc. Natl. Acad. Sci. USA 85, 1816–1820. Ohmori, H., Friedberg, E. C., Fuchs, R. P., Goodman, M. F., Hanaoka, F., Hinkle, D., Kunkel, T. A., Lawrence, C. W., Livneh, Z., Nohmi, T., Prakash, L., Prakash, S., Todo, T., Walker, G. C., Wang, Z., and Woodgate, R. (2001). The Y-family of DNA polymerases. Mol. Cell 8, 7–8. Ohmori, H., Hatada, E., Qiao, Y., Tsuji, M., and Fukuda, R. (1995). dinP, a new gene in Escherichia coli, whose product shows similarities to UmuC and its homologues. Mutat. Res. 347, 1–7. Opperman, T., Murli, S., Smith, B. T., and Walker, G. C. (1999). A model for a umuDCdependent prokaryotic DNA damage checkpoint. Proc. Natl. Acad. Sci. USA 96, 9218–9223. Paz-Elizur, T., Skaliter, R., Blumenstein, S., and Livneh, Z. (1996). Beta*, a UV-inducible smaller form of the beta subunit sliding clamp of DNA polymerase III of Escherichia coli. I. Gene expression and regulation. J. Biol. Chem. 271, 2482–2490. Pham, P., Bertram, J. G., O’Donnell, M., Woodgate, R., and Goodman, M. F. (2001). A model for SOS-lesion-targeted mutations in Escherichia coli. Nature 409, 366–370. Pham, P., Seitz, E. M., Saveliev, S., Shen, X., Woodgate, R., Cox, M. M., and Goodman, M. F. (2002). Two distinct modes of RecA action are required for DNA polymerase V-catalyzed translesion synthesis. Proc. Natl. Acad. Sci. USA 99, 11061–11066. Plosky, B. S., and Woodgate, R. (2004). Switching from high-fidelity replicases to lowfidelity lesion-bypass polymerases. Curr. Opin. Genet. Dev. 14, 113–119. Potapova, O., Grindley, N. D., and Joyce, C. M. (2002). The mutational specificity of the Dbh lesion bypass polymerase and its implications. J. Biol. Chem. 277, 28157–28166. Rajagopalan, M., Lu, C., Woodgate, R., O’Donnell, M., Goodman, M. F., and Echols, H. (1992). Activity of the purified mutagenesis proteins UmuC, UmuD0 , and RecA in replicative bypass of an abasic DNA lesion by DNA polymerase III. Proc. Natl. Acad. Sci. USA 89, 10777–10781. Reuven, N. B., Arad, G., Maor-Shoshani, A., and Livneh, Z. (1999). The mutagenesis protein UmuC is a DNA polymerase activated by UmuD0 , RecA, and SSB and is specialized for translesion replication. J. Biol. Chem. 274, 31763–31766.
262
FUCHS ET AL.
Reuven, N. B., Arad, G., Stasiak, A. Z., Stasiak, A., and Livneh, Z. (2001). Lesion bypass by the Escherichia coli DNA polymerase V requires assembly of a RecA nucleoprotein filament. J. Biol. Chem. 276, 5511–5517. Reuven, N. B., Tomer, G., and Livneh, Z. (1998). The mutagenesis proteins UmuD0 and UmuC prevent lethal frameshifts while increasing base substitution mutations. Mol. Cell 2, 191–199. Shen, X., Sayer, J. M., Kroth, H., Ponten, I., O’Donnell, M., Woodgate, R., Jerina, D. M., and Goodman, M. F. (2002). Efficiency and accuracy of SOS-induced DNA polymerases replicating benzo[a]pyrene-7,8-diol 9,10-epoxide A and G adducts. J. Biol. Chem. 277, 5265–5274. Shimizu, M., Gruz, P., Kamiya, H., Kim, S. R., Pisani, F. M., Masutani, C., Kanke, Y., Harashima, H., Hanaoka, F., and Nohmi, T. (2003). Erroneous incorporation of oxidized DNA precursors by Y-family DNA polymerases. EMBO Rep. 4, 269–273. Shinagawa, H., Iwasaki, H., Kato, T., and Nakata, A. (1988). RecA protein-dependent cleavage of UmuD protein and SOS mutagenesis. Proc. Natl. Acad. Sci. USA 85, 1806–1810. Silvian, L. F., Toth, E. A., Pham, P., Goodman, M. F., and Ellenberger, T. (2001). Crystal structure of a DinB family error-prone DNA polymerase from Sulfolobus solfataricus. Nat. Struct. Biol. 8, 984–989. Skaliter, R., Bergstein, M., and Livneh, Z. (1996b). Beta*, a UV-inducible shorter form of the beta subunit of DNA polymerase III of Escherichia coli. II. Overproduction, purification, and activity as a polymerase processivity clamp. J. Biol. Chem. 271, 2491–2496. Skaliter, R., Paz-Elizur, T., and Livneh, Z. (1996a). A smaller form of the sliding clamp subunit of DNA polymerase III is induced by UV irradiation in Escherichia coli. J. Biol. Chem. 271, 2478–2481. Slechta, E. S., Bunny, K. L., Kugelberg, E., Kofoid, E., Andersson, D. I., and Roth, J. R. (2003). Adaptive mutation: General mutagenesis is not a programmed response to stress but results from rare coamplification of dinB with lac. Proc. Natl. Acad. Sci. USA 100, 12847–12852. Smith, B. T., and Walker, G. C. (1998). Mutagenesis and more: umuDC and the Escherichia coli SOS response. Genetics 148, 1599–1610. Steinborn, G. (1978). Uvm mutants of E. coli K12 deficient in UV mutagenesis. 1. Isolation of uvm mutants and their phenotypical characterization in DNA repair and mutagenesis. Mol. Gen. Genet. 165, 87–93. Stelter, P., and Ulrich, H. D. (2003). Control of spontaneous and damage-induced mutagenesis by SUMO and ubiquitin conjugation. Nature 425, 188–191. Strauss, B. S., Roberts, R., Francis, L., and Pouryazdanparast, P. (2000). Role of the dinB gene product in spontaneous mutation in Escherichia coli with an impaired replicative polymerase. J. Bacteriol. 182, 6742–6750. Stukenberg, P. T., Studwell-Vaughan, P. S., and O’Donnell, M. (1991). Mechanism of the sliding beta-clamp of DNA polymerase III holoenzyme. J. Biol. Chem. 266, 11328–11334. Sutton, M. D., Opperman, T., and Walker, G. C. (1999). The Escherichia coli SOS mutagenesis proteins UmuD and UmuD0 interact physically with the replicative DNA polymerase. Proc. Natl. Acad. Sci. USA 96, 12373–12378. Sutton, M. D., Smith, B. T., Godoy, V. G., and Walker, G. C. (2000). The SOS response: Recent insights into umuDC-dependent mutagenesis and DNA damage tolerance. Annu. Rev. Genet. 34, 479–497.
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Suzuki, N., Ohashi, E., Hayashi, K., Ohmori, H., Grollman, A. P., and Shibutani, S. (2001). Translesional synthesis past acetylaminofluorene-derived DNA adducts catalyzed by human DNA polymerase kappa and Escherichia coli DNA polymerase IV. Biochemistry 40, 15176–15183. Sweasy, J. B., Witkin, E. M., Sinha, N., and Roegner-Maniscalco, V. (1990). RecA protein of Escherichia coli has a third essential role in SOS mutator activity. J. Bact. 172, 3030–3036. Tang, M., Bruck, I., Eritja, R., Turner, J., Frank, E. G., Woodgate, R., O’Donnell, M., and Goodman, M. F. (1998). Biochemical basis of SOS-induced mutagenesis in Escherichia coli: Reconstitution of in vitro lesion bypass dependent on the UmuD0 2C mutagenic complex and RecA protein. Proc. Natl. Acad. Sci. USA 95, 9755–9760. Tang, M., Pham, P., Shen, X., Taylor, J. S., O’Donnell, M., Woodgate, R., and Goodman, M. F. (2000). Roles of E. coli DNA polymerases IV and V in lesiontargeted and untargeted SOS mutagenesis. Nature 404, 1014–1018. Tang, M., Shen, X., Frank, E. G., O’Donnell, M., Woodgate, R., and Goodman, M. F. (1999). UmuD0 (2)C is an error-prone DNA polymerase, Escherichia coli pol V. Proc. Natl. Acad. Sci. USA 96, 8919–8924. Tomer, G., and Livneh, Z. (1999). Analysis of unassisted translesion replication by the DNA polymerase III holoenzyme. Biochemistry 38, 5948–5958. Trincao, J., Johnson, R. E., Escalante, C. R., Prakash, S., Prakash, L., and Aggarwal, A. K. (2001). Structure of the catalytic core of S. cerevisiae DNA polymerase : Implications for translesion DNA synthesis. Mol. Cell 8, 417–426. Wagner, J., Etienne, H., Janel-Bintz, R., and Fuchs, R. P. P. (2002). Genetics of mutagenesis in E. coli: Various combinations of translesion polymerases (Pol II, IV and V) deal with lesion/sequence context diversity. DNA Repair 1, 159–167. Wagner, J., Fujii, S., Gruz, P., Nohmi, T., and Fuchs, R. P. (2000). The clamp targets DNA polymerase IV to DNA and strongly increases its processivity. EMBO Rep. 1, 484–488. Wagner, J., Gruz, P., Kim, S.-R., Yamada, M., Matsui, K., Fuchs, R. P. P., and Nohmi, T. (1999). The dinB gene encodes a novel E. coli DNA polymerase, DNA PolIV, involved in mutagenesis. Mol. Cell 4, 281–286. Wagner, J., and Nohmi, T. (2000). Escherichia coli DNA polymerase IV mutator activity: Genetic requirements and mutational specificity. J. Bacteriol. 182, 4587–4595. Walker, G. C. (1998). Skiing the black diamond slope: progress on the biochemistry of translesion DNA synthesis. Proc. Natl. Acad. Sci. USA 95, 10348–10350. Washington, M. T., Prakash, L., and Prakash, S. (2001). Yeast DNA polymerase utilizes an induced-fit mechanism of nucleotide incorporation. Cell 107, 917–927. Wood, R. D., and Hutchinson, F. (1984). Non-targeted mutagenesis of unirradiated lambda phage in Escherichia coli host cells irradiated with ultraviolet light. J. Mol. Biol. 173, 293–305. Woodgate, R. (1992). Construction of a umuDC operon substitution mutation in E. coli. Mut. Res. 281, 221–225. Woodgate, R. (1999). A plethora of lesion-replicating DNA polymerases. Genes Dev. 13, 2191–2195. Woodgate, R., Rajagopalan, M., Lu, C., and Echols, H. (1989). UmuC mutagenesis protein of Escherichia coli: Purification and interaction with UmuD and UmuD0 . Proc. Natl. Acad. Sci. USA 86, 7301–7305. Yang, W. (2003). Damage repair DNA polymerases Y. Curr. Opin. Struct. Biol. 13, 23–30.
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Yeiser, B., Pepper, E. D., Goodman, M. F., and Finkel, S. E. (2002). SOS-induced DNA polymerases enhance long-term survival and evolutionary fitness. Proc. Natl. Acad. Sci. USA 99, 8737–8741. Yin, J., Seo, K. Y., and Loechler, E. L. (2004). A role for DNA polymerase V in G -> T mutations from the major benzo[a]pyrene N2-dG adduct when studied in a 50 -TT sequence in E. coli. DNA Repair 3, 323–334. Zhou, B. L., Pata, J. D., and Steitz, T. A. (2001). Crystal structure of a DinB lesion bypass DNA polymerase catalytic fragment reveals a classic polymerase catalytic domain. Mol. Cell 8, 427–437.
MAMMALIAN POL k: REGULATION OF ITS EXPRESSION AND LESION SUBSTRATES By HARUO OHMORI,* EIJI OHASHI,* AND TOMOO OGIÀ *Institute For Virus Research, Kyoto University, Sakyo-ku, Kyoto, 606-8507, Japan À Genome Damage and Stability Centre, University of Sussex, Farmer, Brighton BN19RR, United Kingdom
I. Structures of the Genes and Proteins. . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Regulation of Expression of the Human POLK Gene and the Mouse Polk Gene . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. Splicing Variants and Enzyme Activities . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . C. Conservation of the C2HC Zinc Cluster Sequence . . . . . . . . . . . . . . . . . . . . . .. . . . . . II. Enzymatic Properties of Pol. . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Is Pol an Inserter or an Extender?. . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. What are the Cognate DNA Lesions for Pol? . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . III. Possible Mechanisms of TLS by Pol In Vivo . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . .
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I. STRUCTURES OF THE GENES AND PROTEINS A. Regulation of Expression of the Human POLK Gene and the Mouse Polk Gene Chromosomal DNAs in living organisms are continually exposed to a vast variety of genotoxic agents from exogenous and endogenous sources. Some environmental compounds such as polycyclic aromatic hydrocarbons (PAHs) are activated as mutagens in mammalian cells (Friedberg et al., 1995). Among PAHs, benzo[a]pyrene (B[a]P) has been most extensively studied because it is believed to be a potent carcinogen, especially responsible for the p53 mutations detected in the lung tumors of smokers (Denissenko et al., 1996). PAHs are activated by intracellular processes that are mediated by the arylhydrocarbon receptor (AhR) (for a recent review, see Nebert et al., 2004). AhR is called also as a dioxin receptor, because dioxin has a much higher affinity to AhR. When a PAH compound enters mammalian cells, it binds to AhR localized in the cytoplasm. The ligandactivated AhR then moves to the nucleus, where the protein forms a complex with Arnt (AhR nuclear translocator). The AhR-Arnt complex binds to specific DNA sequences called XREs (xenobiotic responsive elements), which are present in the promoter regions of PAH-inducible genes including the mouse Cyp1a1 gene. The product of the Cyp1a1 gene is a cytochrome 265 ADVANCES IN PROTEIN CHEMISTRY, Vol. 69
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P450 protein (1A1) that metabolizes B[a]P and other PAHs to phenols and dihydrodiols to excrete such lipophilic compounds from the inside of cells. However, some of the metabolites become activated so as to attack DNA to form a covalent linkage. B[a]P diol epoxide (BPDE, see Fig. 1), the so-called ultimate carcinogen derived from B[a]P, introduces bulky adducts predominantly at the N 2 position of guanine and less frequently at the N 6 position of adenine. Analysis of the mouse Polk genomic sequence revealed that the promoter region contains two copies of XRE-like sequence (Ogi et al., 2001). In vitro experiment with partially purified AhR and Arnt proteins showed that the AhR–Arnt complex did bind to each of the two XRE-like sequences found in the Polk promoter region, although less efficiently compared with the XRE sequence in the Cyp1a1 promoter region. Furthermore, expression of the mouse Polk gene was stimulated when wild-type mice were treated with 3-methylcholanthrene (3MC), a PAH compound similar to B[a]P (see Fig. 1). In contrast, such stimulation by 3MC was not observed in AhR-knockout mice, whereas a basal level of the Polk expression was still observed. AhR-knockout mice show no detectable Cyp1a1 expression with or without 3MC treatment, and they are refractory to B[a]P-induced skin tumors (Shimizu et al., 2000). The human POLK gene also has similar XRE-like sequences in the promoter region. O-Wang et al. (2001) found
FIG. 1. Structures of benzo[a]pyrene, 3-methylcholanthrene and dioxin. Benzo[a]pyrene and 3-methylcholanthrene are PAH compounds, but dioxin is not. Benzo[a]pyrene is converted to the ultimate carcinogen BPDE by Cyp1A1.
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that Pol was frequently overexpressed in human lung cancer tissues as compared with a matched nontumor tissue counterpart. More recently, a close correlation between elevated Pol expression and p53 inactivation in human lung cancer tissues was found (Wang et al., 2004). In contrast Velasco-Miguel et al. (2003) reported that the mouse Polk gene transcription was regulated by the p53 gene both constitutively and following exposure to selected DNA-damaging agents such as ultraviolet (UV) irradiation and doxorubicin, whereas the human POLK gene expression was not induced by such agents. The detailed mechanisms by which the mouse Polk and the human POLK genes are regulated by p53 in different ways remain unclear, because no sequence similar to the consensus p53 binding sequence has been identified near their promoter regions. Further analyses of the mouse Polk and human POLK genomic sequences revealed that another gene (COL4A3BP in humans and Col4a3bp in mice) transcribed into the opposite direction is present in the immediate upstream of the genes. The initiation sites for the two divergent transcriptions in the human and mouse genomes are separated by only about 300 nucleotides, indicating that those genes share some regulatory elements to bind common transcription factors. The human COL4A3BP gene was initially identified as the GPBP (Goodpasture antigen binding protein) gene, because the product was thought to bind to the C-terminal region of the collagen IV alpha-3 chain, a Goodpasture antigen (Raya et al., 1999). However, a recent paper demonstrated that the COL4A3BP gene codes for CERT, a protein that mediates intracellular trafficking of the lipid precursor ceramide (Hanada et al., 2003). At present, it is unknown how the gene product carries out two such completely different functions. Moreover, almost nothing is known about how the GPBP/CERT gene expression is regulated, although the gene might be also under the control of AhR–Arnt. It should be noted that many eukaryotic genes, including those coding for DNA polymerase, have a gene in the immediate upstream that is transcribed into the opposite direction (see Adachi and Lieber, 2002). Another typical example for such a bidirectional gene organization related to a lesion-bypass enzyme is the human XPV and XPO5 genes, which code for Pol and Exportin 5, respectively. Both of the human POLK and mouse Polk genes are ubiquitously expressed with the highest expression in testis. Such expression patterns are observed with many other genes involved in DNA repair and recombination functions. However, one unique feature of the human and mouse genes encoding Pol is that the major transcripts found in the testes are different in size from those found in other organs; the mRNAs abundantly present in the testes are about 2.8 kb whereas those ubiquitously expressed in many organs (including testis) are 4.2 kb (Gerlach et al.,
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1999; Ogi et al., 1999). The major difference between the two distinct species of mRNAs resides in the 30 -untranslated region (30 -UTR). Although the shorter 2.8-kb transcripts have a polyA stretch in the immediate downstream of the translation stop codon, the longer 4.2-kb transcripts have a long (1.4 kb) 30 -UTR containing multiple copies of AUUUA sequence (Gerlach et al., 1999, see Fig. 2). The AUUUA sequence is an essential and minimal unit of AREs (A+U-rich elements) that are frequently found in the 30 -UTRs of highly labile mRNAs, such as those encoding cytokines, growth factors, and proto-oncogenes (Xu et al., 1997). ARE when present in 30 -UTR is considered to make mRNAs unstable. The alternative selection of the polyA site in the testes may render the 2.8-kb transcripts
FIG. 2. The genomic structure of the mouse Polk gene and splicing variants. Transcription of the mouse Polk gene starts at two different sites, between which P1a is ubiquitously used and P1b is almost exclusively in testis. Major transcripts found in testis are 2.8 kb in length. Most of them have a polyA tail immediately after the translation stop codon (1 and 2), in which the AAUGAA containing the stop codon should correspond to the polyA signal sequence. Some transcripts lacked the translation stop codon (as in variant 4), where the upstream AAUAAA sequence was used as the polyA signal sequence. About 50% of transcripts in testis lacked the exon 7 (as in variant 3). Longer transcripts of 4.2 kb in length were ubiquitously expressed, which included a long 30 -UTR with multiple copies of ARE sequence. The coding region starts within exon 2 and terminates within exon 15.
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lacking an ARE sequence more stable than the 4.2-kb transcripts and may consequently result in the abundant accumulation of the 2.8-kb transcripts in the testes. Many tissues may express only labile mRNAs to keep the amount of Pol at a low level and avoid gratuitous mutations due to the presence of an excess amount of the error-prone enzyme.
B. Splicing Variants and Enzyme Activities Testis-specific regulation of the mouse Polk transcription is also observed as splicing variants. RT-PCR experiments indicated that about 50% of the human POLK-mRNAs from testis lacked the exon 7 of 240 nucleotides in length (our unpublished results; see Fig. 2), and a minor proportion of them lacked the exon 13 (Gerlach et al., 1999). The full-length forms of the human and mouse Pol proteins consisted of 870 and 852 amino acids, respectively (Figs. 2 and 3). When the mouse testis extracts were reacted with anti-Pol antibody, which was raised against the peptide antigen corresponding to the human Pol 558–571 sequence (identical to the mouse 557–570 sequence), two different protein bands of 100 and 80 kDa were observed. The 100-kDa band probably corresponds to the intact form of the mouse Pol protein, which is observed as the sole product in other organs. The shorter, 80-kDa band was observed exclusively in testis, which seems to be too small to be the product originated from skipping of the exon 7. It is also unlikely that the 80-kDa band is derived from the splicing variant lacking the exon 13, because the antigen sequence taken to raise the antibody used in the experiment is encoded by the exon 13. The N-terminal half of Pol forms a catalytic core domain containing multiple motifs that are shared among the Y-family DNA polymerases including the Escherichia coli DinB (Pol IV) (Fig. 3). The C-terminal half is not essential for the DNA polymerase activity per se, at least in vitro, but the region would be necessary for the in vivo function because putative nuclear localization signal (NLS) and PCNA binding site (PBS) are found at the extreme C terminus. Two copies of C2HC zinc cluster sequence, which is similar to that found in RAD18 homologues, are found between the catalytic domain and the C terminus. A truncated form containing the 1–560 residues, which was overproduced by a baculovirus system, showed a DNA polymerase activity in vitro that bypasses certain DNA lesions, as does the full-length protein (Ohashi et al., 2000a,b), but a shorter form containing the 1-510 residues did not show a DNA polymerase activity (Gerlach et al., 2001). Skipping exon 13 in the human POLK-mRNA should result in the deletion of the 511–830 residues, and therefore, the protein derived from the mouse splicing variant lacking exon 13 is expected to be inactive as a DNA polymerase. Recently, we
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FIG. 3. The genomic structure of the human POLK gene and structure of the Pol protein. The human POLK gene has a genomic structure very similar to that of the mouse Polk gene. The N-terminal half of Pol contains multiple motifs (I–V) conserved among Y-family DNA polymerases, which are required for the enzyme activity. NLS and PBS at the C terminus represent putative nuclear localization signal and PCNA binding sites. Splicing variants lacking exons 7 and 13 lack a part of the motifIV and two copies of C2HC sequences, respectively. Various forms of Pol protein lacking the C-terminal half were overproduced in Escherichia coli and assayed for DNA polymerase activity.
developed a system to overproduce the human Pol protein in E. coli cells. The 1–560 protein overproduced in E. coli cells retained the DNA polymerase activity, but deletion of the 232–311 residues corresponding to the region encoded by exon 7 of the mouse Polk and human POLK genes abolished the activity. Thus, neither of the two splicing variants observed in mouse testis appears to direct a protein active as a DNA polymerase. Mammalian Pol proteins have an N-terminal extension of about 100 residues that is not conserved among the Y-family proteins. We overproduced the protein containing the 91–560 or 117–560 residues of human Pol in E. coli. When measuring DNA polymerase activity in vitro, the 91–560 protein retained the enzyme activity, although weaker than the 1–560 protein, but the 117–560 protein did not show any activity (see Fig. 3).
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C. Conservation of the C2HC Zinc Cluster Sequence Recently, Pol homologs in chicken and rat were identified (Okada et al., 2002; the XP_342179 entry in the protein database). The amino acid sequences of the C-terminal half domain in such homologs are variable, except for the two C2HC-cluster, putative NLS and PCNA binding sequences. A DinB/Pol homolog is present in Schizosaccharomyces pombe and Caenorhabditis elegans, but not in Saccharomyces cerevisiae or Drosophila melanogaster. The hypothetical Pol homologs of S. pombe and C. elegans contain the multiple motifs shared among the Y-family DNA polymerases, but their protein sequences deduced by conceptual translation (e.g., P34409 and T41397, respectively, in the Swiss-protein database) do not have a C2HC zinc cluster sequence in the C-terminal regions. However, the absence of a C2HC motif in the S. pombe and C. elegans Pol homologs seems to be a result of an error in the prediction of protein coding region by computer programs, which neglected the presence of one additional exon at the 30 -terminus in each case. The S. pombe Pol homolog actually contains a total of 547 residues with a C2HC sequence (492-CPVC-X13-HVDLC-513) (our unpublished results), instead of 493 residues, as described in the T41397 entry (see Johnson et al., 1999). Similarly, the hypothetical C. elegans Pol homolog (F22B7.6) should contain 596 residues with a C2HC motif (519-CPIC-X13-HVDEC-540), instead of 518 residues described in the P34409 entry. In fact, a hypothetical Pol homolog (CAE75018) of the closely related nematoda C. briggsae contains 612 residues with a C2HC motif (515-CPIC-X17-HVDEC-540). Thus, eukaryotic Pol homologs appear to share a common basic architecture, including a catalytic domain, C2HC (one or two copies), NLS, and PCNA binding site from the N to C terminus (Fig. 3). The function of the C2HC motif remains to be determined.
II. ENZYMATIC PROPERTIES OF POL A. Is Pol an Inserter or an Extender? Thus far, at least four different groups have reported their experimental results on in vitro TLS by Pol, all of which are consistent with stating that the enzyme cannot bypass T-T cyclobutane pyrimidine dimer (CPD) in vitro by itself (Gerlach et al., 2001; Johnson et al., 2000a; Ohashi et al., 2000a; Zhang et al., 2000a). However, Pol-defective mutants derived from mouse embryonic stem (ES) and fibroblast (MEF) cells or from chicken DT40 cells are moderately UV sensitive (Ogi et al., 2002; Okada et al., 2002; Schenten et al., 2002), indicating that Pol is somehow involved in the response to ultraviolet (UV)-induced DNA damages. Washington et al.
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FIG. 4. Structures of BPDE adducts and estrogen-derived adducts. BPDE forms covalent linkage at the N 2 of guanine (major products) and N 6 of adenine (minor products). Estrogen also forms similar adducts.
(2002) showed that Pol efficiently extended G and A (but less efficiently C and T) placed opposite the 30 T of a T-T CPD, whereas the enzyme did not insert any nucleotide opposite the 30 T of the same lesion or extend any base placed opposite the 30 T of a (6–4) T-T photoproduct. In addition, they found that Pol efficiently extended from base mispairs on undamaged DNA. Subsequently, Haracska et al. (2002a) found that Pol efficiently extended various nucleotides incorporated opposite O6-methly guanine and 8-oxoguaine by Pol . Thus, they concluded that Pol should play a role as an extender in translesion synthesis. However, the efficiency of Pol or any other TLS enzyme in inserting a correct or incorrect nucleotide opposite a DNA lesion site strongly depends on the species of the lesion. In fact, the authors also noticed that Pol inserted A in preference to other bases opposite 8-oxoguaine at 37% efficiency (in terms of relative ratio of kcat/Km), compared with nondamaged G, whereas it inserted C opposite O6-methyl guanine at 1% efficiency, still higher by more than
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two orders of magnitude than for other bases. We believe that the bypassing efficiency of each enzyme in comparison with nondamaged template should greatly vary on the extent of a distortion in the DNA structure caused by the respective DNA lesion, and that the more important datum is the relative efficiency of each lesion by a given TLS enzyme in comparison with that of other TLS and replicative DNA polymerases. As the mouse Polk gene expression was induced by 3MC, an analog of B[a]P, which is believed to be a major causative agent of human lung cancers, whether or not Pol could bypass DNA adducts generated by B[a]P was thus examined. The results obtained by two groups indeed showed that human Pol could bypass different stereoisomers of dG-N2BPDE (the major products generated by BPDE, see Fig. 4) by inserting the correct C opposite the bulky lesions in preference to other bases (Suzuki et al., 2002; Zhang et al., 2000a, 2002). Furthermore, Pol bypassed dG-N2BPDE more efficiently than human Pol and Pol (alone or in combination with the yeast Pol ) (Rechkoblit et al., 2002). Very interestingly, Pol inserted A more efficiently than C opposite dG-N2-BPDE (Chiapperino et al., 2002; Rechkoblit et al., 2002; Zhang et al., 2000b). Pol inserted C opposite dG-N2-BPDE adducts at 1% efficiency of nondamaged G, much lower when compared with the fact that Pol can insert A opposite the 30 T of a T-T CPD at the same efficiency as it does with nondamaged template (Johnson et al., 2000b). Nevertheless, we believe that Pol plays a critical role in the response to B[a]P–induced DNA damages, because mouse ES cells with a Polk-defective mutation showed a hypersensitivity of B[a]P, generating more mutations than the parental cells (Ogi et al., 2002). Furthermore, the spectrum of B[a]P-induced mutations in Polk-defective cells was different from that in the wild-type cells; G-to-T transversions predominated (70%) among the mutations observed in Polk-defective cells, whereas G-to-T and A-to-G substitutions occurred at an equal frequency of 30% of total mutations in the parental cells. Such in vivo results strongly indicated that Pol contributes to error-free bypass of dGN2-BPDE adducts and that in the absence of Pol, another enzyme, probably Pol, inserts A opposite the adducts to generate G-to-T transversions. This situation is reminiscent of that in the XPV cells lacking Pol, where other TLS polymerases are involved in error-prone bypass of UVinduced DNA damages.
B. What are the Cognate DNA Lesions for Pol? The E. coli Pol IV (DinB) protein is also able to bypass dG-N2-BPDE adduct by inserting the correct C opposite the lesion in vitro (Shen et al., 2002). The ability to correctly bypass dG-N2-BPDE adduct is apparently
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conserved among the DinB/Pol homologs from E. coli to mammals. A structure around the active sites conserved by the DinB/Pol homologs, but not by Pol or Pol, probably enables the bulky dG-N2-BPDE adduct to form a base paring with C. However, the ability of the E. coli Pol IV to bypass such lesions should have little biological role because E. coli cells do not have an enzyme to activate B[a]P into BPDE. Similarly, it seems unlikely that mammalian Pol enzymes are conserved just for bypassing B[a]P-induced DNA damages. Fischhaber et al. (2002) reported that human Pol was able to insert A opposite thymine glycol (Tg) at 0.2% 5% efficiency of nondamaged thymine in vitro. This seems to be consistent with the finding that the mouse Polk gene expression is induced by UV radiation and doxorubicin, both of which are known to generate Tg and other lesions in DNA (Velasco-Miguel et al., 2003). More recently, Suzuki et al. (2004) found that human Pol could efficiently bypass through N2-[3-methoxyestra-1,3,5(10)-trien-6-y1]-20 -deoxyguanosine (dGN2-3MeE) by inserting C opposite the lesion at 33% efficiency of nondamaged G. dG-N2-3MeE is a model compound used in place of N2-(2hydroestron-6-yl-)-20 -deoxyguanosine (2-OHE-N2-dG; see Fig. 4), a product generated by the reaction between DNA and estrogen-2,3-quinone, which is a metabolite generated by oxidation of estrogen. Because Pol is highly expressed in the adrenal cortex of embryonic mice (Velasco-Miguel et al., 2003) a site of active steroid biosynthesis, such estrogen-derived DNA adducts, especially those formed at the N2 position of guanine with some structural similarity to dG-N2-BPDE, are good candidates for endogenous DNA damages that Pol could cope with. To conclude that thymine glycol and estrogen-modified guanine are cognate lesions for Pol, in vivo experiments demonstrating that Pol-deficient cells are defective for the bypass of such adducts are required.
III. POSSIBLE MECHANISMS OF TLS
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The most important question for the mechanism of TLS in vivo is how a TLS enzyme replaces the replicative DNA polymerases stalled at a DNA lesion site, performs the lesion-bypass reaction, and is then replaced by the replicative enzyme again. One hot issue in this regard is the finding that monoubiquitination of PCNA by the RAD6-RAD18 complex is required for S. cerevisiae Pol to function in vivo at a lesion site (Hoege et al., 2002; Steler and Ulrich, 2003; see the chapter by Xiao). Thus far, the interactions between TLS enzymes and PCNA have been studied, using unmodified PCNA (Haracska et al., 2001a, b, c, and 2002b), so that they should be reexamined with ubiquitinated PCNA. In contrast, a study on the S. pombe DinB homolog suggested that its functioning needs the activity of the
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RAD9-RAD1-HUS1 (9-1-1) complex (a complex similar to PCNA) and the RAD17-RFC(2-5) complex (the loader of the 9-1-1 complex to DNA) (Kai and Wang, 2003). It is still an open question whether the human Pol prefers ubiquitinated PCNA to the unmodified one or uses the 9-1-1 complex instead of unmodified and ubiquitinated PCNA. Studies to search for Pol-interacting proteins revealed that Pol interacts with a C-terminal region of REV1 (Guo et al., 2003; Ohashi et al., 2004), which has been known to interact with REV7, the noncatalytic subunit of another TLS enzyme Pol (Murakumo et al., 2001). Interestingly, REV1 has a BRCT domain near the N terminus, which may function as an interaction domain with another protein (for more details, see the chapter by Lawrence). Because the C-terminal region of REV1 also interacts with Pol and Pol, it seems likely that REV1 plays a key role, for example, as a scaffold protein in a multiprotein complex for TLS in vivo. We are still left with many unanswered questions regarding the in vivo TLS mechanisms. For example, how is the RAD6-RAD18 complex activated on DNA damaging? Is deubiquitination of PCNA required for the replicative DNA polymerase to resume DNA synthesis when lesionbypass is completed? What is the mechanism for recruiting an appropriate enzyme to a given species of DNA lesion. Obviously, further intensive investigations are required to resolve these intriguing questions.
Acknowledgments The studies in our laboratory cited here are supported in parts by Research-in-Aid Grants (to H.O. and E.O.) from the Ministry of Education, Culture, Sports, and Science of Japan.
REFERENCES Adachi, N., and Lieber, M. R. (2002). Bidirectional gene organization: A common architectural feature of the human genome. Cell 109, 807–809. Chiapperino, D., Kroth, H., Kramarczuk, I. H., Sayer, J. M., Masutani, C., Hanaoka, F., Jerina, D. M., and Cheh, A. M. (2002). Preferential misincorporation of purine nucleotides by human DNA polymerase opposite benzo[a]pyrene 7,8-diol 9,10epoxide deoxyguanosine adducts. J. Biol. Chem. 277, 11765–11771. Denissenko, M. F., Pao, A., Tang, M., and Pfeifer, G. P. (1996). Preferential formation of benzo[a]pyrene adducts at lung cancer mutational hotspots in P53. Science 274, 430–432. Fischhaber, P. L., Gerlach, V. L., Feaver, W. J., Hatahet, Z., Wallace, S. S., and Friedberg, E. C. (2002). Human DNA polymerase bypasses and extends beyond thymine glycols during translesion synthesis in vitro, preferentially incorporating correct nucleotides. J. Biol. Chem. 277, 37604–37611. Friedberg, E. C., Walker, G. C., and Siede, W. (1995). DNA Repair and Mutagenesis. Washington, D.C: ASM Press.
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Gerlach, V. L., Aravind, L., Gotway, G., Schultz, R. A., Koonin, E. V., and Friedberg, E. C. (1999). Human and mouse homologs of Escherichia coli DinB (DNA polymerase IV), members of the UmuC/DinB superfamily. Proc. Natl. Acad. Sci. USA 96, 11922–11927. Gerlach, V. L., Feaver, W. J., Fischhaber, P. L., and Friedberg, E. C. (2001). Purification and characterization of pol, a DNA polymerase encoded by the human DINB1 gene. J. Biol. Chem. 276, 92–98. Guo, C., Fischhaber, P. L., Luk-Paszyc, M. J., Masuda, Y., Zhou, J., Kamiya, K., Kisker, C., and Friedberg, E. C. (2003). Mouse Rev1 protein interacts with multiple DNA polymerases involved in translesion DNA synthesis. EMBO J. 22, 6621–6630. Hanada, K., Kumagai, K., Yasuda, S., Miura, Y., Kawano, M., Fukasawa, M., and Nishijima, M. (2003). Molecular machinery for non-vesicular trafficking of ceramide. Nature 426, 803–809. Hoege, C., Pfander, B., Moldovan, G.-L., Pyrowolakis, G., and Jentsch, S. (2002). RAD6dependent DNA repair is linked to modification of PCNA by ubiquitin and SUMO. Nature 419, 135–141. Haracska, L., Kondratick, C. M., Unk, I., Prakash, S., and Prakash, L. (2001a). Interaction with PCNA is essential for yeast DNA polymerase function. Mol. Cell 8, 407–415. Haracska, L., Johnson, R. E., Unk, I., Phillips, B., Hurwitz, J., Prakash, L., and Prakash, S. (2001b). Physical and functional interactions of human DNA polymerase with PCNA. Mol. Cell. Biol. 21, 7199–7206. Haracska, L., Johnson, R. E., Unk, I., Phillips, B. B., Hurwitz, J., Prakash, L., and Prakash, S. (2001c). Targeting of human DNA polymerase to the replication machinery via interaction with PCNA. Proc. Natl. Acad. Sci. USA 98, 14256–14261. Haracska, L., Prakash, L., and Prakash, S. (2002a). Role of human DNA polymerase as an extender in translesion synthesis. Proc. Natl. Acad. Sci. USA 99, 16000–16005. Haracska, L., Unk, I., Johnson, R. E., Philips, B. B., Hurwitz, J., Prakash, L., and Prakash, S. (2002b). Stimulation of DNA synthesis activity of human DNA polymerase by PCNA. Mol. Cell. Biol. 22, 784–791. Johnson, R. E., Washington, M. T., Prakash, S., and Prakash, L. (1999). Bridging the gap: A family of novel DNA polymerases that replicate faulty DNA. Proc. Natl. Acad. Sci. USA 96, 12224–12226. Johnson, R. E., Prakash, S., and Prakash, L. (2000a). The human DINB1 gene encodes the DNA polymerase . Proc. Natl. Acad. Sci. USA 97, 3838–3843. Johnson, R. E., Washington, M. T., Prakash, S., and Prakash, L. (2000b). Fidelity of human DNA polymerase . J. Biol. Chem. 275, 7447–7450. Kai, M., and Wang, T. S. (2003). Checkpoint activation regulates mutagenic translesion synthesis. Genes Dev. 17, 64–76. Murakumo, Y., Ogura, Y., Ishii, H., Numata, S.-I., Ichihara, M., Croce, C. M., Fishel, R., and Takahashi, M. (2001). Interactions in the error-prone postreplication repair proteins hREV1, hREV3, and hREV7. J. Biol. Chem. 276, 35644–35651. Nebert, D. W., Dalton, T. P., Okey, A. B., and Gonzalez, F. J. (2004). Role of arylhydrocarbon receptor-mediated induction of the CYP1 enzymes in environmental toxicity and cancer. J. Biol. Chem. 279, 23847–23850. Ogi, T., Kato, T., Jr., Kato, T., and Ohmori, H. (1999). Mutation enhancement by DINB1, a mammalian homolog of the Escherichia coli mutagenesis protein DinB. Genes Cells 4, 607–618. Ogi, T., Mimura, J., Hikida, M., Fujimoto, H., Fujii-Kuriyama, Y., and Ohmori, H. (2001). Expression of human and mouse genes encoding pol: Testis-specific
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developmental regulation and AhR-dependent inducible transcription. Genes Cells 6, 943–953. Ogi, T., Shinkai, Y., Tanaka, K., and Ohmori, H. (2002). Pol protects mammalian cells against the lethal and mutagenic effects of benzo[a]pyrene. Proc. Natl. Acad. Sci. USA 99, 15548–15553. Ohashi, E., Bebenek, K., Matsuda, T., Feaver, W. F., Gerlach, V. L., Friedberg, E. C., Ohmori, H., and Kunkel, T. A. (2000a). Fidelity and processivity of DNA synthesis by DNA polymerase , the product of the human DINB1 gene. J. Biol. Chem. 275, 39678–39684. Ohashi, E., Murakumo, Y., Kanjo, N., Akagi, J., Masutani, C., Hanaoka, F., and Ohmori, H. (2004). Interaction of hREV1 with three human Y-family DNA polymerases. Genes Cells. 9, 523–531. Ohashi, E., Ogi, T., Kusumoto, R., Iwai, S., Masutani, C., Hanaoka, F., and Ohmori, H. (2000b). Error-prone bypass of certain DNA lesions by the human DNA polymerase . Genes Dev. 14, 1589–1594. Okada, T., Sonoda, E., Yamashita, Y. M., Koyoshi, S., Tateishi, S., Yamaizumi, M., Takata, M., Ogawa, O., and Takeda, S. (2002). Involvement of vertebrate Pol in Rad18-independent postreplication repair of UV damage. J. Biol. Chem. 277, 48690–48695. O-Wang, J., Kawamura, K., Tada, Y., Ohmori, H., Hideki, K., Sakiyama, S., and Tagawa, M. (2001). DNA polymerase , implicated in spontaneous and DNA damage-induced mutagenesis, is overexpressed in lung cancer. Cancer Res. 61, 5366–5369. Raya, A., Revert, F., Navarro, S., and Saus, J. (1999). Characterization of a novel type of serine/threonine kinase that specifically phosphorylates the human Goodpasture antigen. J. Biol. Chem. 274, 12642–12649. Rechkoblit, O., Zhang, Y., Guo, D., Wang, Z., Amin, S., Krzeminsky, J., Louneva, N., and Geacintov, N. E. (2002). trans-Lesion synthesis past bulky benzo[a]pyrene diol epoxide N2-dG and N6-dA lesions catalyzed by DNA bypass polymerases. J. Biol. Chem. 277, 30488–30494. Shen, X., Sayer, J. M., Kroth, H., Ponten, I., O’Donnell, M., Woodgate, R., Jerina, D. M., and Goodman, M. F. (2002). Efficiency and accuracy of SOS-induced DNA polymerases replicating benzo[a]pyrene-7,8- diol-9,10-epoxide A and G adducts. J. Biol. Chem. 277, 5265–5274. Shenten, D., Gerlach, V. L., Guo, C., Velasco-Miguel, S., Hladik, C. L., White, C. L., Friedberg, E. C., Rajewsky, K., and Esposito, G. (2002). DNA polymerase deficiency does not affect somatic hypermutation in mice. Eur. J. Immunol. 32, 3152–3160. Shimizu, Y., Nakatsuru, Y., Ichinose, M., Takahashi, Y., Kume, H., Mimura, J., Fujii-Kuriyama, Y., and Ishikawa, T. (2000). Benzo[a]pyrene carcinogenicity is lost in mice lacking the arylhydrocarbon receptor. Proc. Natl. Acad. Sci. USA 97, 779–782. Steler, P., and Ulrich, H. D. (2003). Control of spontaneous and damage-induced mutagenesis by SUMO and ubiquitin conjugation. Nature 425, 188–191. Suzuki, N., Ohashi, E., Kolbanovskiy, A., Geacintov, N. E., Grollman, A. P., Ohmori, H., and Shibutani, S. (2002). Translesion synthesis by human DNA polymerase on a DNA template containing a single stereoisomer of dG -(+)- or dG-()-anti-N2-BPDE (7,8-dihydroxy-anti-9,10-epoxy-7,8,9,10-tetrahydrobenzo[a]pyrene). Biochemistry 41, 6100–6106. Suzuki, N., Itoh, S., Poon, K., Masutani, C., Hanaoka, F., Ohmori, H., Yoshizawa, I., and Shibutani, S. (2004). Translesion synthesis past estrogen-derived DNA adducts by human DNA polymerases and . Biochemistry 43, 6304–6311.
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Velasco-Miguel, S., Richardson, J. A., Gerlach, V. L., Lai, W. C., Gao, T., Russell, L. D., Hladik, C. L., White, III C. L., and Friedberg, E. C. (2003). Constitutive and regulated expression of the mouse Dinb (Polk) gene encoding DNA polymerase . DNA Repair 2, 91–106. Washington, M. T., Johnson, R. E., Prakash, L., and Prakash, S. (2002). Human DINB1encoded DNA polymerase is a promiscuous extender of mispaired primer termini. Proc. Natl. Acad. Sci. USA 99, 1910–1914. Xu, N., Chen, C.-C. A., and Shyu, A.-B. (1997). Modulation of the fate of cytoplasmic mRNA by AU-rich elements: Key sequence features controlling mRNA deadenylation and decay. Mol. Cell. Biol. 17, 4611–4621. Wang, Y. Q., Seimiya, M., Kawamura, K., Yu, L., Ogi, T., Takenaga, K., Shishikura, T., Nakagawara, A., Sakiyama, S., Tagawa, M., and O-Wang, J. (2004). Elevated expression of DNA polymerase in human lung cancer is associated with p53 inactivation: Negative regulation of POLK promoter activity by p53. Int. J. Oncol. 25, 161–165. Zhang, Y., Wu, X., Guo, D., Rechkoblit, O., and Wang, Z. (2002). Activities of human DNA polymerase in response to the major benzo[a]pyrene DNA adduct: Errorfree lesion bypass and extension synthesis from opposite the lesion. DNA Repair 1, 559–569. Zhang, Y., Yuan, F., Wu, X., Wang, M., Rechkoblit, O., Taylor, J.-S., Geacintov, N. E., and Wang, Z. (2000a). Error-free and error-prone lesion bypass by human DNA polymerase in vitro. Nucleic Acids Res. 28, 4138–4146. Zhang, Y., Yuan, F., Wu, X., Rechkoblit, O., Taylor, J.-S., Geacintov, N. E., and Wang, Z. (2000b). Error-prone lesion bypass by human DNA polymerase . Nucleic Acids Res. 28, 4717–4724.
DNA POSTREPLICATION REPAIR MODULATED BY UBIQUITINATION AND SUMOYLATION By LANDON PASTUSHOK AND WEI XIAO Department of Microbiology and Immunology, University of Saskatchewan, Saskatoon, Canada
I. II. III. IV. V.
VI. VII. VIII. IX.
Introduction . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . DNA Postreplication Repair in Prokaryotes . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . DNA Postreplication Repair in Eukaryotes. . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . Ubiquitination . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . Protein Conjugation in PRR. . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Rad6-Rad18. . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. Mms2-Ubc13-Rad5 . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . C. Ubc9-Siz1 . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . Postreplication Repair via Covalent Modifications of PCNA . . . . . . . . . . . . . . . .. . . . . . Functional Conservation of Eukaryotic Postreplication Repair . . . . . . . . . . . . .. . . . . . Future Directions. . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . .
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I. Introduction To ensure the viability of itself and its progeny, the living cell has developed ways to reduce or avoid detrimental changes to its genetic material. Because of the numerous external and internal agents that act on and modify DNA, the incredible task of ensuring DNA fidelity and the survival of individual organisms is made possible by a variety of DNA repair and replication processes. In most cases, these processes are well understood and, as such, are the major focus of other chapters in this book. Conversely, DNA damage-tolerance pathways such as postreplication repair (PRR) in eukaryotes are not yet well defined. Part of the difficulty in this case is that PRR is not superficially considered an actual DNA repair mechanism. PRR itself does not result in the physical removal of DNA lesions but exists as a means to circumvent the severe consequences of replication blocks that otherwise lead to cell death. This has made it difficult to describe PRR in the context of a pathway or biochemical activity. Instead, PRR loosely refers to events whereby damage-induced single-stranded DNA gaps are somehow converted into double-stranded DNA after replication. To advance our understanding of the genetics and possible biochemical processes of PRR, the yeast Saccharomyces cerevisiae has been the most studied eukaryotic model organism. Recent 279 ADVANCES IN PROTEIN CHEMISTRY, Vol. 69
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experimentation in the budding yeast has finally provided a functional and mechanistic context for this often puzzling enigma. This chapter will focus on yeast PRR from a protein-based perspective, with emphasis on the covalent protein modification of proliferating cell nuclear antigen (PCNA) by ubiquitin (Ub) and a small Ub-like modifier (SUMO). Readers may wish to refer to recent reviews dealing with the genetic analysis of PRR in particular (Broomfield et al., 2001) and the DNA damage tolerance network in general (Barbour and Xiao, 2003).
II. DNA Postreplication Repair in Prokaryotes It can perhaps be argued that the mechanism of DNA damage tolerance has been best described in prokaryotes such as Escherichia coli. Indeed, research into the SOS response and the RecA protein has led to the discovery of physiological consequences comparable to PRR in S. cerevisiae. However, although parallels can be drawn between recombination and mutagenic translesion DNA synthesis (TLS) in each case, the regulatory mechanisms and molecular architecture are very different. Consequently, a brief overview of DNA damage tolerance in prokaryotes provides an interesting perspective and raises interest in the remarkable molecular evolution of PRR regulatory components in eukaryotes. The crux of DNA damage tolerance in E. coli lies in the RecA protein, which plays both a regulatory role in controlling the SOS response and a physical role in repairing and bypassing damaged DNA. When the replication machinery encounters a block, such as a lesion generated by DNA damage, synthesis is reinitiated downstream, and the result is a singlestranded DNA (ssDNA) gap (Rupp and Howard-Flanders, 1968). In its regulatory role, RecA acts as a sensor that recognizes these regions via ssDNA binding affinity and consequently transforms itself to an activated form, RecA* (Salles and Defais, 1984). RecA* acts as a coprotease that stimulates LexA to autodigest (Little, 1984). Under noninduced conditions, LexA forms a homodimer that binds to ‘‘SOS boxes’’ found in the promoter regions of SOS regulon genes and represses, in varying degrees, the expression of over 20 genes involved in many facets of DNA repair and replication, including those for postreplication repair. Thus, when RecA* stimulates the autodigestion of the LexA repressor, the SOS regulon genes including lexA and recA are expressed at increased levels. As part of a negative feedback loop, the newly synthesized LexA and RecA proteins ensure a rapid return to the repressed state once the activating signal (ssDNA gaps) is removed from the cell. Similar to its role in controlling SOS response after DNA damage, a second means of regulation by RecA in prokaryotic PRR also involves the
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coprotease activity of RecA*. The umuC and umuD genes, also under the control of SOS response, encode two subunits of a mutagenic translesion polymerase, Pol V (Reuven et al., 1999; Tang et al., 1998, 1999). On DNA-damage treatment, RecA* stimulates the cleavage of the Pol V regulatory subunit UmuD to its active UmuD0 form (Shinagawa et al., 1988). UmuD0 then forms a homodimer, which pairs with UmuC to create a fully functional Pol V (UmuD0 2UmuC). In addition to the regulatory roles above, RecA also participates directly in the DNA-damage avoidance process. In the recombination-mediated mode of DNA damage tolerance, RecA acts with RecBCD in DNA doublestrand break repair (Kuzminov, 1999) and with the RecFOR complex to stabilize stalled replication forks (Chow and Courcelle, 2004; Courcelle et al., 1997; Webb et al., 1997) and to bypass replication blocks by resuming replication using a newly synthesized homologous template (Courcelle et al., 1997; Kogoma, 1997). During Pol V–catalyzed TLS, RecA is required in two distinct ways; namely, translesion synthesis and the stimulation of nucleotide incorporation (Pham et al., 2002). In summary, as illustrated in Fig. 1, prokaryotic PRR employs RecA as a DNA damage sensor via ssDNA binding affinity, as a signal transducer at both transcriptional and posttranslational levels, and as an effector that directly participates in homologous recombination, replication restart, and TLS. Notably, the regulatory role of RecA in prokaryotic PRR involves the modification of proteins via protease cleavage. In contrast, eukaryotes have developed a very different and potentially more sophisticated strategy of PRR regulation; namely, modification by protein conjugation.
III. DNA Postreplication Repair in Eukaryotes The PRR response in eukaryotes is apparently very similar in overall strategy to that of prokaryotic cells (see Fig. 1). In each case, the endpoint is recombination-mediated damage avoidance or TLS to allow replication past DNA lesions that would otherwise result in cell death. The similarities end there, however, as the molecular architecture, biochemical activities, and signaling events in PRR are strikingly different between prokaryotes and eukaryotes. Because the PRR pathway has been best characterized in S. cerevisiae, which is the only model to date to illustrate eukaryotic PRR mechanisms, our discussions of PRR in this chapter will focus on a yeast perspective. Classical genetic analysis has led to the convention that PRR consists of branched pathways that include at least one ‘‘error-free’’ and one ‘‘errorprone’’ pathway (reviewed in Broomfield et al., 2001). The RAD6 and RAD18 genes are required for both subpathways, and mutations that
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Fig. 1. Comparison of the DNA postreplication repair response in prokaryotes and eukaryotes. Shaded blocks indicate functionally conserved steps between the Escherichia coli and Saccharomyces cerevisiae pathways.
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disrupt either gene result in the most severe DNA damage sensitivities of all PRR genes. RAD6 encodes a ubiquitin-conjugating enzyme (Ubc or E2) that forms a heterodimer with Rad18, an ssDNA binding protein with ATPase activity. The error-prone branch consists of Pol (Rev3 þ Rev7) and the UmuC homolog, Rev1, which are discussed in detail in other chapters of this book. A parallel branch consists of an E2 complex comprises Ubc13-Mms2 and Rad5, a ssDNA-dependent ATPase. Given the fact that the ubiquitination activity of both Rad6-Rad18 and Mms2-Ubc13-Rad5 complexes is essential for their PRR functions, it is necessary to briefly review the ubiquitination process. Readers are encouraged to refer to recent reviews in this field (Hershko and Ciechanover, 1998; Hochstrasser, 1996; Jentsch, 1992; Pickart, 2001).
IV. Ubiquitination Since the discovery of Ub (Schlesinger et al., 1975) more than 20 years ago, ubiquitination (Hershko et al., 1983) has become one of the cornerstones of covalent protein modification. An explosion of research in the area in the last 10 years has revealed biological roles for ubiquitination that rival the scope of phosphorylation. Because its traditional and bestcharacterized role is a fundamental biological process found in eukaryotes, namely, proteasome-dependent protein degradation, it is not surprising that ubiquitination has such a broad cellular influence. More recently, however, breakthrough discoveries in the field have revealed an even greater depth and versatility, several examples of which are encountered in the PRR pathway. In the simplest sense, ubiquitination is a three-step biochemical reaction that uses Ub, a small, globular, 76–amino acid protein to covalently modify its targets. As the name implies, Ub is found throughout eukaryotic cells, and with merely a three–amino acid difference between lower and higher eukaryotes, it is one of the most conserved proteins in nature (Ozkaynak et al., 1984). The biochemical reaction is initiated when an ATP-dependent ubiquitin-activating enzyme (Uba or E1) forms a high-energy thiolester bond between the C-terminal Gly76 residue of Ub and an internal Cys residue of the E1. Ub is next transferred from the E1 to form another thiolester bond, this time with the active-site Cys residue of an E2. The final step links the C-terminal residue of Ub to a surface "-amino group of a Lys residue on the target, forming an isopeptide bond that usually, but not always, requires a ubiquitin-ligase (Ubl or E3) that may itself interact with Ub. Although the E1 performs a rather general role and is encoded by a single or very few genes in the cell, E2s and E3s are responsible for the
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versatility and target specificity of ubiquitination. E2s comprise a family of proteins with a highly conserved core domain of approximately 150 amino acids. On the basis of various crystallographic data, the core domain forms a globular / protein with a central -sheet and surrounding -helices (Moraes et al., 2001; Pickart, 2001; see Fig. 2A). Within this region lies an obligatory Cys active site that is absolutely conserved among all E2s and that is housed in a shallow cleft in the center of the protein (VanDemark
Fig. 2. Structure and function of the Ubc13-Mms2 heterodimer. (A) Crystal structure of the human Ubc13-Mms2 heterodimer at 1.8 A˚ resolution (Moraes et al., 2001). Mms2 binds Ubc13 end-on to form a ‘‘T’’ shape. Mms2 adopts a similar / fold to Ubc13, but without two C-terminal -helices. (B) van der Waals space-fill representation of the Ubc13-Mms2 surface. Green, Mms2; blue, Ubc13; yellow, activesite cysteine of Ubc13. Residues undergoing chemical shift on Ub-Ubc13 thiolester formation, and the Ub-Mms2 noncovalent interaction are shown in tan and rose, respectively. Arrows indicate channels through which Ub passes during ubiquitination. Molecular graphics in both (A) and (B) were generated using RasTop software (Valadon, http://www.geneinfinity.org/rastop/). (C) Ubc13-Mms2-mediated ubiquitination involves Ubc13 thiolester formation with ‘‘donor’’ Ub, Mms2 orientation of the ‘‘acceptor’’ Ub, and direct or indirect delivery to the substrate. Within the yeast PRR pathway, it is believed that Rad5 acts as the E3 and PCNA is the substrate. (See Color Insert.)
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and Hill, 2002). As evidenced by multiple E2 crystal structures (e.g., see Cook et al., 1993; Moraes et al., 2001; Worthylake et al., 1998), the threedimensional organization of all E2s is similar and creates a typical Ubc fold. Although confined to these general similarities, E2 enzymes have the potential for diversification. Indeed, S. cerevisiae contains only 11 E2s that are able to covalently attach Ub to various target proteins. The diversity and specificity are believed to be achieved in several ways. Many E2s are longer than the core sequence and consist of C- or N-terminal extensions and are classified as Class II and Class III Ubcs, respectively ( Jentsch, 1992). It is believed that such E2 extensions mediate specific downstream interactions with E3 proteins or the actual target of ubiquitination (Pickart, 2001). More important, the ability of a single E2 to bind more than one E3 is probably the determining factor in specifying a given substrate, as the E3s represent the most plentiful group of proteins in ubiquitination. E3s are structurally diverse and act alone or as part of a multisubunit complex to fulfill their functions as either active or passive adapters between the E2 and substrate. To date, only two characteristic domains have been well defined in some E3 proteins. The HECT (homology to the E6-AP carboxyl terminus) domain is able to bind Ub in a thiolesterdependent manner akin to E1 and E2 enzymes (Scheffner et al., 1995). In contrast, RING finger–containing E3s do not necessarily participate in conjugation reactions but act as adapters between conjugation enzymes and their substrates via a RING finger motif (reviewed in Joazeiro and Weissman, 2000). The discovery of the RING finger was based on its similarity to the DNA-binding zinc finger motif (Lovering et al., 1993). Similar to the zinc finger, the RING finger fulfills a structural purpose to stabilize an otherwise unstructured loop region by creating a scaffold via intrapeptide binding of zinc cations. The RING finger differs in structure from the zinc finger in that it has a longer consensus sequence designated Cys-X2-Cys-X(9–39)-Cys-X(1–3)-His-X(2–3)-Cys/His-X2-Cys-X(4–48)Cys-X2-Cys, where X is any amino acid (Saurin et al., 1996), and in its propensity for binding other proteins (especially Ubcs) instead of DNA. Several crystal structures of proteins containing RING motifs have been solved, the most significant of which is the structure of the Ubc-RING finger heterodimer of UbcH7 and c-Cbl (Zheng et al., 2000). A single pass through ubiquitination results in the tagging of substrate by a single Ub moiety (mono-Ub), which in some cases is the desired end product. Alternatively, poly-Ub chains can be formed if successive reactions take place and an internal lysine of another Ub molecule is used as the subsequent site of attachment. Poly-Ub chains are most commonly conjugated via Gly76-Lys48 linkages to provide a characteristic ‘‘flag’’ for signaling protein degradation by the 26S proteasome (Hochstrasser,
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1996). However, the structure of Ub reveals six other surfaceexposed lysine residues (Vijay-Kumar et al., 1985, 1987), and thus a new and growing field recognizing alternatively linked poly-Ub chains has emerged. To date, poly-Ub chains made via Lys63 (Spence et al., 2000), Lys-11 (Baboshina and Haas, 1996), Lys29 (Arnason and Ellison, 1994), and Lys6 (Wu-Baer et al., 2003) linkages have been reported. Although the E2s and E3s provide two levels of variability in ubiquitination, the potential for different poly-Ub conjugates allows a third and fascinating level of versatility. At a fundamental level, it implies a mechanistic diversity with regard to enzyme catalysis, as the surface lysines of Ub are well dispersed. Because E2s are so conserved within their catalytic domain, the ability of one E2 to conjugate Ub through a different Lys than another E2 indicated that the first E2 either possesses corresponding intrinsic differences or functions along with other, potentially novel accessory factors. A case for the latter was recently discovered for the function of the Ubc13-Mms2 complex in PRR and will be discussed below. Another consequence of nonclassical poly-Ub conjugates is that different conjugates may lead to different physiological signals. For example, although typical Lys48 poly-Ub conjugates signal for proteasome degradation, Lys63-mediated poly-Ub conjugates signal for various cellular processes such as a stress response (Arnason and Ellison, 1994), mitochondrial inheritance (Fisk and Yaffe, 1999), plasma membrane protein endocytosis (Galan and Haguenauer-Tsapis, 1997), ribosome function (Spence et al., 2000), and PRR (Hofmann and Pickart, 1999).
V. Protein Conjugation in PRR Perhaps the most fascinating aspect of eukaryotic PRR is not simply how it has adapted to use protein conjugation in place of protein cleavage as in prokaryotes, but that it does so in ways that have set new precedents and revealed completely novel mechanisms in DNA repair. The following sections will describe the molecular framework behind the three protein conjugation complexes in PRR.
A. Rad6-Rad18 On the basis of the conventional genetic hierarchy of DNA postreplication repair, as discussed above, Rad6 is considered the hallmark and starting point for the pathway. RAD6 encodes a multifunctional Ubc (Jentsch et al., 1987) that, in addition to its roles in DNA repair, functions
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in histone modification and N-end rule protein degradation. Because the E2 activity of Rad6 is required for all its known biological roles (Sung et al., 1990, 1991), it stands to reason that Rad6 fulfills its distinct functions through interaction with different E3s or via different Ub conjugations, or both. One of these functions is revealed by the ability of S. cerevisiae Rad6 to create and attach poly-Ub chains to histones in vitro (Sung et al., 1988). Subsequent in vitro studies confirmed the E2 activity of Rad6 for histones but went on to suggest atypical poly-Ub chain catalysis (Haas et al., 1991). Although Rad6 can indeed form poly-Ub chains via Lys6 in vitro (Baboshina and Haas, 1996), the in vivo evidence remains elusive. Another curious aspect of Rad6 is its specificity for histones in vitro in the absence of an E3. Because Rad6 is a Class II E2 with a C-terminal tail that lies beyond the core Ubc domain, it has been hypothesized that the C-terminal string of 14 consecutive acidic amino acids provides affinity for positively charged histones. This view is supported by studies that show that the deletion of the Rad6 C-terminal tail abolishes its ability to ubiquitinate histones, but not its DNA repair function (Sung et al., 1988). However, the role of Rad6 in histone modification has been substantiated in vivo (Robzyk et al., 2000), and contrary to in vitro studies, Rad6 requires an E3 RING finger protein, Bre1 (Hwang et al., 2003; Wood et al., 2003). However, the interplay, if any, between the C-terminus of Rad6 and Bre1 has yet to be elucidated. Also, these studies indicate that the in vivo modification of H2B by Rad6 is by mono-Ub. Such conjugates are not sufficient for proteasome degradation and are instead implicated in stabilization or signaling. Another early study of ubiquitination by Rad6 involves a process called the ‘‘N-end rule,’’ in which the in vivo half-life of some proteins is determined by the nature of their N-terminal amino acids (Bachmair et al., 1986). The N-end rule is dependent on the E2 activity of Rad6 (Dohmen et al., 1991), and as such, Rad6 catalyzes the formation of typical Lys48 poly-Ub conjugates (Chau et al., 1989). These tagged proteins are subsequently degraded by the 26S proteasome. In contrast to histone modification, the role of Rad6 in the N-end rule has shown a clear and defined requirement for a RING finger E3 protein, Ubr1, and classical Lys48 poly-Ub chain catalysis for proteasome degradation. The multifunctionality of Rad6 is further supported by the fact that the ubr1 mutation does not affect PRR, whereas the rad18 mutant is completely defective. Indeed, Rad6 is able to form exclusive complexes with Rad18 or Ubr1 (Bailly et al., 1994), and the Rad6-Rad18 heterodimer is required for PRR (Bailly et al., 1997a). The Rad18 protein contains the aforementioned RING finger motif and has ATPase and ssDNA binding
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activities (Bailly et al., 1997b). These functions fit well with a working model in which Rad18 brings Rad6 into close proximity to ssDNA gaps so that the ubiquitination of a target protein by Rad6 can initiate the PRR process. The ssDNA binding activity of Rad18 is consistent with observations that PRR pathway mutants are sensitive to killing by a broad range of DNA-damaging agents, which may all potentially lead to a common replication-blocking end product (Broomfield et al., 2001; Prakash et al., 1993). Some insight into the Rad6-Rad18 complex (and Rad6-Ubr1) has been made possible by the yeast Rad6 crystal structure at 2.6 A˚ (Worthylake et al., 1998). As expected from the high sequence homology between E2s, Rad6 adopts the characteristic / fold of the Ubc family and does not contain structural elements that differ significantly from the other Ubcs. Notably, the highly acidic C-terminus unique to Rad6 in S. cerevisiae could not be resolved. Although the Rad6 structure is perhaps not profoundly insightful on its own, it became a valuable tool for visualizing the predicted interacting regions for Rad18 and Ubr1. Using data derived from Rad6 deletion analysis and subsequent binding studies with Ubr1 (Watkins et al., 1993) and Rad18 (Bailly et al., 1997), the residues important for interaction in each case were mapped to the Rad6 surface (Worthylake et al., 1998). Using this method, both Rad18 and Ubr1 were implicated in interacting with similar surfaces on Rad6. These regions are distal from the Rad6 active site and indicate an indirect role, if any, for these E3s in the ubiquitination reaction. This finding underlines that Rad18 and Ubr1 provide roles in defining the biological process for Rad6 and confirms the observation that Rad6 forms distinct heterodimers with each E3. These early structure and function studies demonstrate an ability of Rad6 to catalyze distinct protein modifications; namely, mono-Ub and poly-Ub chains. As a result, it remained difficult to make inferences as to the Ub modification by Rad6-Rad18 in PRR. Given the assumption that both Ubr1 and Rad18 bind the same surface of Rad6, one might assume that the ubiquitination in PRR is via Lys48 as it is in the N-end rule. However, it was very recently discovered that the Rad6-Rad18 complex targets proliferating cell nuclear antigen (PCNA) with a single Ub moiety (Hoege et al., 2002), indicating another level of variability in Rad6-mediated Ub conjugation by its binding partners.
B. Mms2-Ubc13-Rad5 The second instance of ubiquitin conjugation in PRR involves an atypical poly-Ub chain and a novel mechanism of catalysis employing a Ubc enzyme variant (Uev), Mms2. The MMS2 gene was isolated by functional
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complementation, and genetic analyses indicate that MMS2 belongs to the RAD6 pathway (Broomfield et al., 1998). However, unlike rad6 and rad18 mutants, the mms2 mutant displays moderate sensitivity to DNA-damaging agents but significantly increased spontaneous mutation rates. Although the increased mutation rate in mms2 is completely abolished by inactivation of the REV3-mediated TLS pathway, the mms2 and rev3 mutations exhibit a remarkable synergism with respect to killing by DNA damaging agents. These observations place MMS2 in an error-free branch of PRR parallel to the Pol-mediated mutagenesis pathway (Broomfield et al., 1998; Xiao et al., 1999). Uevs are similar in sequence to Ubcs but lack the obligatory active site Cys residue required for Ub conjugation. Uev homologs have been found in essentially all eukaryotic organisms examined to date, including humans (Sancho et al., 1998; Xiao et al., 1998). Phylogenetic analysis also indicates that Uevs have evolved as a distinct class of proteins from Ubcs, early in eukaryotic evolution (Villalobo et al., 2002). Because the Uev domain adopts a similar three-dimensional structure to the Ubc (Fig. 2A) but does not form Ub conjugates, it had been hypothesized that Uevs function as dominant-negative regulators of ubiquitin conjugation (Koonin and Abagyan, 1997). Given the epistatic relationship between rad6 and mms2 in DNA repair, it was suggested that Mms2 functions as an accessory protein that positively modulates the ubiquitination activity of Rad6 (Broomfield et al., 1998). However, rigorous in vivo and in vitro studies failed to demonstrate a physical link between Mms2 and Rad6 (Xiao, W., and Broomfield, S., unpublished observations) and it turned out that Mms2 forms a stable complex with a different and novel Ubc, Ubc13. The physical interaction between Ubc13-Uev/Mms2 was demonstrated by using Ubc13 and Mms2 as bait for copurification (Hofmann and Pickart, 1999) and yeast two-hybrid screens (Brown et al., 2002), respectively. More significantly, it was discovered that Ubc13 is the only known Ubc capable of Lys63 poly-Ub chain assembly, and that its associated Uev is absolutely required for this process (Hofmann and Pickart, 1999). Indeed, deletion of UBC13 in yeast cells results in phenotypes indistinguishable from those of mms2 mutants (Brusky et al., 2000), thus placing UBC13 in error-free PRR together with MMS2. Because Ubc13 consists solely of the core Ubc domain but catalyzes atypical poly-Ub chains via Lys63, the role for Mms2 as a positive regulator via its physical interaction with Ubc13 can be envisioned. Indeed, both yeast (Hofmann and Pickart, 1999) and human (McKenna et al., 2001) Ubc13-Mms2 form a 1:1 stable heterodimer with binding affinities (Kd) of approximately 400 (Ulrich, 2003) and 50 nM (McKenna et al., 2003; Ulrich, 2003), respectively. The role of Mms2 in Ubc13-mediated
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polyubiquitination is revealed using two-dimensional 1H-15N HSQC (heteronuclear single nuclear quantum coherence) such that an ‘‘acceptor’’ Ub for Lys63 attachment is bound in a noncovalent manner by Mms2. This finding leads to a model in which Mms2 orients the acceptor Ub via noncovalent contacts so that its Lys63 is made available to the ‘‘donor’’ Ub of the Ubc13 thiolester (McKenna et al., 2001 and Fig. 2C). The structural basis of the Ubc13-Mms2 complex and its unique role in Lys63 polyubiquitination is further elucidated by crystallographic analysis of yeast (VanDemark et al., 2001) and human (Moraes et al., 2001) complexes. Interestingly, despite the typical Ubc folds adopted by each protein, the heterodimer is asymmetrical and is formed with Mms2 binding Ubc13 in an ‘‘end-on’’ or ‘‘T-shaped’’ manner (Fig. 2A). The interface in each of the yeast and human Ubc13-Mms2 structures is almost completely conserved and buries nearly 1500 A˚ of solvent-accessible surface area. The hydrophobic pocket generated by Ubc13 is not conserved in other E2s but is highly conserved within Ubc13s from different organisms. Site-specific mutations of UBC13 and MMS2 that result in amino acid substitutions of some of the highly conserved residues within this hydrophobic pocket severely affect or abolish Ubc13-Mms2 complex formation (Pastushok, L., Moraes, T. F., Ellison, M. J., and Xiao, W., unpublished observations; VanDemark et al., 2001). Perhaps the greatest contribution of the Ubc13-Mms2 crystal structure analyses is the insight into the basis of Lys63-mediated poly-Ub chain formation. Notably, both studies identified three main channels or clefts converging on the Ubc13 active site, through which Ub makes contact or passes during Lys-63 Ub conjugation (Fig. 2B). Channel I refers to the typical E2 active site at which the donor Ub forms a thiolester with Ubc13. Because the Ubc13 structure does not appreciably change on binding Mms2, it has been inferred that Channel I behaves relatively independent of Mms2. In contrast, the remaining two channels for Ub interaction are dependent on the asymmetrical dimerization of Ubc13-Mms2. Channel II comprises the necessary link between donor and acceptor Ub and provides the molecular framework for Lys63 poly-Ub chain assembly. Various docking experiments indicate a common mechanism whereby Mms2 positions the acceptor Ub within a concave surface such that only the Lys63 residue is made available for conjugation by the Ubc13 active site–bound donor Ub. The last identifiable channel to converge on the Ubc13 active site is also only apparent after Ubc13-Mms2 dimerization. Channel III has been speculated to be an ‘‘outgoing’’ path for Lys63 Ub conjugates and follows in a relatively opposite direction to Channel II. Importantly, structures of the Ubc13-Mms2 heterodimer and the predicted tetramer (Ub-Ubc 13-Mms2-Ub) both indicate that the surface of Ubc13 at the end of
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Channel III remains completely available (McKenna et al., 2003). This exposed region corresponds to the surface required for an E2–E3 interaction, as evidenced by crystal structures of UbcH7 bound to the RING finger E3, c-Cb1 (Zheng et al., 2000) and the HECT domain E3, E6-AP (Huang et al., 1999). Although direct experimental evidence to support a function for Channel III in PRR is currently unavailable, the observation of a physical interaction between Ubc13 and the RING finger protein Rad5 (Ulrich and Jentsch, 2000) indicates functional relevance in vivo. RAD5 was genetically placed within the PRR pathway ( Johnson et al., 1992) and was also identified as an antimutator, rev2 (Lawrence, 1982). In addition to its RING finger motif, Rad5 contains a ssDNA-dependent ATPase domain ( Johnson et al., 1992) and Swi2/Snf2 homologous domains important for chromatin remodeling (Richmond and Peterson, 1996). Therefore, like the Rad6–Rad18 complex, the Rad5–Ubc13 interaction provides the framework for another instance of Ub conjugation in PRR at sites of damaged DNA. Indeed, two-hybrid and in vitro coimmunoprecipitation experiments revealed a physical interaction between Rad5 and Ubc13 (Ulrich and Jentsch, 2000). Functional significance for the interaction was demonstrated with a combination of in vivo localization and cross-linking experiments that showed nuclear localization of Ubc13Mms2 after DNA damage and chromatin association of Rad5-Ubc13, respectively. Substitution of critical residues within the Rad5 RING finger abolishes its binding to Ubc13 (Ulrich, 2003; Ulrich and Jentsch, 2000). To support these findings, the same study revealed specific residues of Ubc13 that mediate interaction with Rad5, and such residues are situated at the distal end of Channel III. Therefore, it may be hypothesized that the Rad5 RING finger plays an active role as an E3 in Ubc13-Mms2 Lys63 poly-Ub conjugation.
C. Ubc9-Siz1 The third and most recent finding of protein conjugation in PRR does not use Ub, but a small Ub-like modifier (SUMO). SUMO is the best-studied example of a group of Ub-like proteins (Schwartz and Hochstrasser, 2003) that can be attached to targets for posttranslational modification in a manner reminiscent of ubiquitination. SUMO-linked proteins are involved in a wide variety of cellular processes. Sumoylation is unique to and conserved throughout eukaryotes and shares similar enzymology to ubiquitination. Although the SUMO molecule shares only 18% similarity with Ub, it adopts a Ub-like fold with conserved positioning of C-terminal residues for isopeptide bond formation (Bayer et al., 1998; Sheng and Liao, 2002). Notably, a protruding flexible N terminus
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extends beyond the region of Ub homology, and the charged surface of SUMO is markedly different than Ub. Furthermore, residues of the Ub moiety responsible for poly-Ub formation are correspondingly absent from SUMO (Bayer et al., 1998). The disparity between SUMO and Ub molecules is indicative of a functionally analogous but distinct set of sumoylation enzymes. Indeed, the only known SUMO-conjugating enzyme to date was originally identified as the ubiquitin-conjugating enzyme, Ubc9 (Seufert et al., 1995), but was subsequently demonstrated to conjugate SUMO instead of Ub (Desterro et al., 1997; Johnson and Blobel, 1997; Schwarz et al., 1998). Although the overall strategy of sumoylation is comparable to ubiquitination, a notable exception is that substrates are almost always tagged with a single SUMO moiety that does not signal for degradation. Also, the crystal structure of a Ubc9-substrate interaction (Bernier-Villamor et al., 2002; Rodriguez et al., 2001) and various in vitro experiments indicate that target identification by Ubc9 can often occur independent of an E3. A further divergence from ubiquitination is evidenced by a consensus sequence found in the target proteins that is commonly used as the site of SUMO attachment (Rodriguez et al., 2001; Sampson et al., 2001). The emergence of Ubc9 in PRR was not brought about by direct studies of DNA repair but through a survey for proteins modified by SUMO in S. cerevisiae. Immunopurification of SUMO–protein conjugates revealed PCNA as a novel target of sumoylation, and by biochemical association, Ubc9 was implicated as the functional E2 (Hoege et al., 2002). Furthermore, the same study identified that SUMO modification of PCNA is mediated by Siz1, a SUMO E3 previously identified as a factor required for sumoylation of yeast septins (Takahashi et al., 2001a,b). Interestingly, as Siz1 contains a zinc-binding RING finger–like domain, the Ubc9-Siz1 relationship represents the third protein-conjugating complex in PRR mediated by a RING finger E3s.
VI. Postreplication Repair via Covalent Modifications of PCNA Because previous studies demonstrated that both Rad6 and Ubc13 ubiquitination activities are required for their PRR functions, it is expected that the covalent modification of one or more critical targets by these two ubiquitination complexes signals for their respective PRR activities (Broomfield et al., 2001). A recent study has clearly pointed to PCNA as such a critical target (Hoege et al., 2002). As discussed elsewhere in this book, PCNA (or Pol30 in S. cerevisiae) is an essential component of the eukaryotic replication machinery. The most
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notable function of PCNA occurs while bound to DNA polymerases (Pol or Pol"), whereby a PCNA trimer encircles DNA and acts as a sliding-clamp to provide processivity during replication (Paunesku et al., 2001). In addition, PCNA is involved in other forms of DNA metabolism such as nucleotide excision repair, base excision repair, mismatch repair, and postreplicative processing (Kelman and Hurwitz, 1998; Paunesku et al., 2001). The involvement of PCNA in PRR was originally implicated through the characterization of a mutant allele of POL30, pol30-46, which conferred ultraviolet sensitivity (Ayyagari et al., 1995). Genetic analysis of pol30-46 placed POL30 within the RAD6 pathway (Torres-Ramos et al., 1996), and pol30-46 appears to be defective in the error-free branch of PRR (Xiao et al., 2000). Discovery of PCNA as the target for covalent modification by PRR proteins was somewhat inadvertent. In screening for new SUMO substrates, Hoege et al. (2002) found that PCNA can be sumoylated. By systematically mutating each of the 18 Lys residues of PCNA to Arg, it was revealed that S. cerevisiae PCNA could be monosumoylated at two Lys residues; namely, Lys127 and Lys164. The authors noted that the prominent site for SUMO conjugation is Lys164, which is conserved within eukaryotes (Fig. 3). It was observed that SUMO modification of PCNA increases during S phase and that treatment of cells with excessive DNA damage resulted in a marked increase in sumoylated PCNA. In contrast, moderate DNA damage resulted in additional PCNA-specific bands that persist in ubc9-1 mutants that are defective in SUMO conjugation. Interestingly, pull-down experiments revealed the novel PCNA species to be conjugates containing one to more than four Ub molecules. Furthermore,
Fig. 3. Alignment of partial PCNA amino acid sequences from various model eukaryotic organisms. Shaded residues match the consensus. Sc, Saccharomyces cerevisiae; Sp, Schizosaccharomyces pombe; Dm, Drosophila melanogaster; Ce, Caenorhabditis elegans; At, Arabidopsis thaliana; Mm, Mus musculus; Hs, Homo sapiens. Lys127 and Lys164 residues of S. cerevisiae (indicated with arrows) are modified by SUMO only, or Ub and SUMO, respectively.
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the Ub conjugates were induced only after DNA damage, and the site of Ub attachment is identical to that of SUMO at Lys164. Because Ub-PCNA is dependent on DNA damage and PCNA was genetically linked to PRR (Torres-Ramos et al., 1996), it was expected that one of the Ubc complexes in the PRR pathway might be responsible. Indeed, covalent modification of PCNA was completely abolished in rad6 mutants, whereas sumoylation was unaffected. Surprisingly, mutants in the errorfree PRR genes UBC13, MMS2, and RAD5 led to the disappearance of polyUb conjugates without affecting the mono-Ub species. It can be thus deduced that monoubiquitination of PCNA is performed by the Rad6Rad18 complex, whereas Ubc13-Mms2-Rad5 is responsible for polyubiquitination of PCNA. Indeed, poly-Ub conjugates of PCNA were absent in yeast cells solely expressing Lys63-mutated Ub. Therefore, the poly-Ub conjugates are attributed to the rare Lys63 conjugation activity of Ubc13-Mms2. To further explore the mechanisms behind PCNA modification, Hoege et al. (2002) sought to demonstrate direct physical interactions between PRR proteins and PCNA. Using the yeast two-hybrid assay, the authors identified interactions between PCNA and the two RING finger proteins, Rad18 and Rad5. Similarly, Ubc9 was also shown to associate with Rad18 and Rad5, as well as PCNA. Considering their ssDNA-binding affinities, the new interactions involving Rad18 and Rad5 can be seen as physical links between the substrate (ssDNA), conjugation machinery (Rad6, Ubc13Mms2, and Ubc9) and target (PCNA) of PRR. Taking into account an earlier report that Rad18 and Rad5 can each form homo- and heterodimers (Ulrich and Jentsch, 2000), a large multisubunit conjugation complex and its biological effects through covalent modification of PCNA can be predicted (Fig. 4). Indeed, characterization of the pol30-K164R mutant phenotypes and its genetic interactions with other members in the RAD6 pathway supports the above proposed model. Because SUMO and Ub converge on the same conserved residue of PCNA, it is attractive to speculate that each conjugate acts as a mutually exclusive switch among different replication and PRR modes. However, although conjugates of Lys63 poly-Ub chains on PCNA can be firmly rooted in error-free PRR via Ubc13-Mms2, the interplay and function of mono-Ub and sumoylation requires further investigation. A recent study (Stelter and Ulrich, 2003) attempted to address this issue through extensive genetic analysis, and it was found that although mono-Ub of PCNA activates TLS mediated by both Pol and Pol, the effects of PCNA sumoylation appear to be more complicated. Sumoylation of PCNA contributes to the extreme sensitivity of rad6 and rad18 mutants and partially affects Pol-mediated spontaneous mutagenesis. It is argued that the
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Fig. 4. Covalent modification by Ub or SUMO modulates PCNA function and leads to different PRR pathways. Current observations are consistent with a model that unmodified and sumoylated PCNA functions in DNA replication. Mono- and Lys63-polyubiquitinated PCNA functions in error-prone and error-free damage tolerance, respectively. Solid circle with ‘‘s,’’ SUMO; open circles with ‘‘u,’’ Ub. This model is adapted from Stelter and Ulrich (2003).
sumoylation of PCNA must play a more critical role than its mutant phenotypes currently offer. One possibility is that the sumoylation of PCNA acts as regulatory antagonist in PRR by competing with ubiquitination at Lys164. It should be noted that although Lys127 conforms to the postulated SUMO conjugation consensus sequence, is sumoylated and appears to play a minor role in PPR, the residue corresponding to Lys127 is only found in S. cerevisiae. In contrast, Lys164 is invariable among PCNAs from various model organisms (Fig. 3).
VII. Functional Conservation of Eukaryotic Postreplication Repair Although the genes and enzymology in PRR are rather different in eukaryotes and prokaryotes, there is sufficient evidence indicating that the PRR process is highly conserved within eukaryotes, from yeast to human. First, almost all yeast PRR proteins have apparent homologs in higher eukaryotes (Table I). Furthermore, most, if not all, of the proteins listed have retained the functional domains or motifs that are seen in S. cerevisiae PRR proteins. The only exception is RAD5, which does not have a true homolog based on protein sequence analysis alone in higher
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Table I Sequence Conservation of Eukaryotic PRR Proteins Protein (S. cerevisiae) Rad6
Rad18
Ubc13
Mms2
PCNA
Organism
Protein name or accession number
% Identity
H. sapiens M. musculus A. thaliana D. melanogaster C. elegans S. pombe H. sapiens M. musculus A. thaliana D. melanogaster C. elegans S. pombe H. sapiens M. musculus A. thaliana D. melanogaster C. elegans S. pombe H. sapiens M. musculus A. thaliana D. melanogaster C. elegans S. pombe H. sapiens M. musculus A. thaliana D. melanogaster C. elegans S. pombe
E2a, E2b Ube2a, Ube2b Ubc2 Dhr6 Ubc-1 Rhp6 Rad18 Rad18 n/a n/a n/a NP_595423 Ubc13 Ubc13 Ubc13a, Ubc13b Bendless, NP_609715 Ubc13 Ubc13 Mms2, Uev1a Mms2 Uev1a, Uev1b, Uev1c, Uev1d NP_647959 Uev1 Mms2 PCNA PCNA PCNA PCNA PCNA PCNA
69, 69 69, 68 65 68 61 77 14 15 — — — 23 68 68 66, 65 70, 68 66 70 47, 46 47 53, 51, 53, 52 46 45 65 35 35 39 35 38 45
PRR proteins from S. cerevisiae were applied to BLAST analysis, and the highest scoring homologs from various model organisms were included in the table and shown with percentage amino acid sequence identity. n/a, not applicable (no sufficient homology found). Note: BLAST search with Rad5 failed to identify sufficient sequence homologs from any organisms examined in the database to date.
eukaryotes. Nevertheless, several potential RAD5 functional homologs are currently under investigation. It is also interesting to note that in some cases (e.g., RAD6 and MMS2), more than one homolog has been identified in the mammalian genome (Table I). Second, several genes isolated from higher eukaryotes are found to be capable of functionally complementing
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their corresponding PRR defects in budding yeast (Ashley et al., 2002; Koken et al., 1991; Xiao et al., 1998). Third, in a few limited cases, experimental inactivation of mammalian PRR genes in a cell line or in an animal model results in phenotypes reminiscent of the corresponding yeast mutants (Li et al., 2002; Roest et al., 1996; Tateishi et al., 2000). Last but not least, the Lys164 residue in PCNA appears to be conserved in all eukaryotic organisms examined so far (Fig. 3), indicating that the mechanism of covalent modification of PCNA in PRR has been conserved throughout evolution. One can infer from this analysis that postreplicative DNA repair is an important mechanism for maintaining genome stability, comparable to other well-characterized repair mechanisms such as nuclear excision repair, base excision repair, recombination repair, and mismatch repair. Certainly, more research needs to be done in multicellular model organisms and in human cells to determine the similarities and differences in PRR and to see whether defects in this pathway lead to human diseases.
VIII. Future Directions The mechanistic studies of PRR described above have been instrumental in bridging genetic data with the biochemical activities of various proteins involved in PRR and allow the development of various elegant models of the PRR pathway (Hoege et al., 2002; Stelter and Ulrich, 2003). Although genetic approaches will continue to provide invaluable insights into the PRR pathway, we anticipate an exciting era in our understanding of PRR through biochemical characterization, leading to the in vitro reconstitution of PRR activity. Such assays have become instrumental in studying other repair processes such as DNA excision repair (Kubota et al., 1996). Because reconstitution experiments are often performed with highly purified components under well-defined conditions and in a piece-by-piece manner, they can help to identify the minimal requirements of a particular process. Potentially, the precise order of molecular and regulatory events can also be elucidated. It is envisioned that three key steps lead toward an ultimate in vitro reconstitution of PRR. The first would be the in vitro covalent modification of PCNA by Ub- and SUMO-conjugating complexes and is justified from several perspectives. First, most of the proteins (or derivatives thereof ) involved in these modifications have been successfully purified. Notably, many of these proteins retain their functions after expression in prokaryotic systems, in which contaminating elements of ubiquitination and sumoylation reactions do not exist. Second, the components and cofactors required for Ub and SUMO conjugation have been well documented, and
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such in vitro reactions are commonplace. Finally, the identification of PCNA as a target of covalent modification provides a single point in the pathway that can be easily monitored. However, whether other unidentified players still exist en route to PCNA modification remains to be elucidated but can be addressed through this investigation. It is noted that Ub or SUMO conjugation to PCNA is probably a regulatory element and does not likely result in the direct processing DNA. Although the downstream effects of mono- and Lys63 polyubiquitination of PCNA are translesion synthesis and recombination or template switching, respectively, the molecular determinants that recognize the conjugates are currently unknown. This study will lead to the second and more challenging task; namely, to understand how these conjugation states of PCNA signal the physiological switch between error-free and error-prone PRR. Ultimately, the reconstitution of PRR in vitro may involve a series of reactions from the recognition of the DNA lesion substrate to the use of different strategies to bypass the lesion, with or without an undamaged homologous chromatid as a template. Two working models have been proposed (Fig. 5) to account for possible molecular events in response to Lys63 polyubiquitination of PCNA by error-free PRR proteins; however, neither has been confirmed or ruled out because of a lack of proper assays. In this regard, the reconstitution of PRR activity may pose the greatest challenge in the field of DNA replication and repair. Another difficulty concerning in vitro reconstitution of PRR is that although some of the regulatory mechanisms have now become clear, our understanding of several other aspects is still in its infancy. In our opinion, the following issues are pertinent to our understanding of how PRR is regulated in eukaryotic cells and need to be addressed. First, one of the regulatory components of PCNA conjugation is its sumoylation, which presumably competes with Rad6-Rad18-mediated monoubiquitination and is involved in promoting spontaneous mutagenesis (Stelter and Ulrich, 2003). SUMO conjugation to PCNA is increased during S phase and can also be induced by severe DNA damage (Hoege et al., 2002); however, the biological significance of this process is unknown. Interestingly, the DNA helicase Srs2, which serves as a molecular switch between PRR and recombination (Barbour and Xiao, 2003; Schiestl et al., 1990), is also cell-cycle regulated (Liberi et al., 2000). Because the entire PRR activity is dependent on functional SRS2 (Broomfield and Xiao, 2002; Ulrich, 2001), the relationship between PCNA sumoylation and Srs2 activity needs to be further investigated. Second, as discussed before, Rad5 contains predicted structural motifs indicative of involvement in chromatin remodeling. This unique feature of Rad5 among PRR proteins may explain additional phenotypes of its
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Fig. 5. Two alternative models for error-free PRR via recombinational processes. (A) A strand exchange model, and (B) a template switching model. Both models propose that progression of leading strand synthesis in the presence of replicationblocking DNA damage (represented by a triangle) requires the association of the two nascent DNA strands, followed by resolution of the intermediary structure via (A) cleavage of the Holliday junction or (B) reverse branch migration. Adapted from Broomfield et al. (2001).
mutants compared with those of ubc13/mms2. Indeed, characterization of a histone H2B mutation htb1-3 points to possible links between Rad5 and histone modification (Martini et al., 2002). Also, the ubiquitination of H2B by Rad6 is required for optimal mitotic cell growth and meiosis (Robzyk et al., 2000). Whether chromatin remodeling through histone modification plays a role in PRR remains to be determined. Third, it has been previously demonstrated that while Rad6 and Rad18 are nuclear proteins, Ubc13 and Mms2 are largely cytosolic and are translocated into the nucleus in response to DNA damage (Ulrich and Jentsch, 2000). The mechanism of this regulation is currently unknown and needs to be investigated. Fourth, although RecA in E. coli cells plays a critical role in the transcriptional regulation of SOS regulon genes in response to DNA damage, a similar phenomenon involving the Rad6–Rad18 complex has not been reported. Instead, cell-cycle checkpoint genes appear to play a similar role in eukaryotic cells (Bachant and Elledge, 1998). It would be interesting to
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know whether the central regulatory proteins in PRR such as Rad6 and Rad18 are involved in gene regulation in response to DNA damage. Fifth, because the currently known modifications of PCNA are on the same Lys residue and do not signal for degradation, it can be inferred that a reversible process may play an important role in the restoration of eukaryotic PRR. Although it is known that desumoylation of PCNA probably requires Ulp1 (Hoege et al., 2002), a Ub protease involved in deubiquitination of PCNA has not been reported. Similar to the negative feedback by the LexA repressor in the E. coli SOS response (Fig. 1), the recovery from DNA damage in eukaryotes might result in the concomitant sumoylation of PCNA to inhibit PRR.
IX. Conclusions Through the above analysis, it is conceivable that both prokaryotic and eukaryotic organisms employ a centrally controlled postreplicative survival mechanism to deal with ssDNA gaps left by replication blocks during DNA synthesis. In bacteria such as E. coli, the homologous recombinase RecA serves as a coprotease to fulfill a regulatory role at the posttranslational level. Its targets include a transcriptional repressor and a nonessential DNA polymerase. In eukaryotes, it is the E2–E3 ubiquitination complex Rad6–Rad18 that plays the central regulatory role by controlling TLS and an error-free mechanism of lesion bypass, presumably via synthesis of gapped DNA using a homologous chromatid or homologous chromosome. Although the only currently known target is PCNA, other targets may be encountered. The discovery of novel signal transduction mechanisms in PRR through sequential modification of PCNA by two E2–E3 ubiquitination complexes sets an important milestone in the research of eukaryotic DNA repair. Notably, the Rad6–Rad18 complex bridges monoubiquitination of PCNA with error-prone PRR, whereas Ubc13-Mms2-Rad5 ties novel Lys-63 polyubiquitination of PCNA with error-free PRR. In addition, another level of sophistication is revealed by the sumoylation of PCNA by Ubc9-Siz1, which underlines an elegant interplay between DNA repair and replication.
Acknowledgments We thank Michelle Hanna for proofreading the manuscript and other laboratory members for valuable comments. This work is supported by the Canadian Institutes of Health Research operating grants (OP-38104 and MOP-53240) to W.X. and a Natural Sciences and Engineering Research Council of Canada postgraduate fellowship to L.P.
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References Arnason, T., and Ellison, M. J. (1994). Stress resistance in Saccharomyces cerevisiae is strongly correlated with assembly of a novel type of multiubiquitin chain. Mol. Cell. Biol. 14, 7876–7883. Ashley, C., Pastushok, L., McKenna, S., Ellison, M. J., and Xiao, W. (2002). Roles of mouse UBC13 in DNA postreplication repair and Lys63-linked ubiquitination. Gene 285, 183–191. Ayyagari, R., Impellizzeri, K. J., Yoder, B. L., Gary, S. L., and Burgers, P. M. (1995). A mutational analysis of the yeast proliferating cell nuclear antigen indicates distinct roles in DNA replication and DNA repair. Mol. Cell. Biol. 15, 4420–4429. Baboshina, O. V., and Haas, A. L. (1996). Novel multiubiquitin chain linkages catalyzed by the conjugating enzymes E2EPF and RAD6 are recognized by 26 S proteasome subunit 5. J. Biol. Chem. 271, 2823–2831. Bachant, J. B., and Elledge, S. J. (1998). Regulatory metworks that control DNA damage-inducible genes in Saccharomyces cerevisiae. In ‘‘DNA Damage and Repair, Vol. 1: DNA Repair in Prokaryotes and Lower Eukaryotes’’ ( J. A. Nickoloff and M. F. Hoekstra, Eds.), pp. 383–410. Totowa, NJ, Humana Press. Bachmair, A., Finley, D., and Varshavsky, A. (1986). In vivo half-life of a protein is a function of its amino-terminal residue. Science 234, 179–186. Bailly, V., Lamb, J., Sung, P., Prakash, S., and Prakash, L. (1994). Specific complex formation between yeast RAD6 and RAD18 proteins: A potential mechanism for targeting RAD6 ubiquitin-conjugating activity to DNA damage sites. Genes Dev. 8, 811–820. Bailly, V., Lauder, S., Prakash, S., and Prakash, L. (1997b). Yeast DNA repair proteins Rad6 and Rad18 form a heterodimer that has ubiquitin conjugating, DNA binding, and ATP hydrolytic activities. J. Biol. Chem. 272, 23360–23365. Bailly, V., Prakash, S., and Prakash, L. (1997a). Domains required for dimerization of yeast Rad6 ubiquitin-conjugating enzyme and Rad18 DNA binding protein. Mol. Cell. Biol. 17, 4536–4543. Barbour, L., and Xiao, W. (2003). Regulation of alternative replication bypass pathways at stalled replication forks and its effects on genome stability: A yeast model. Mutat. Res. 532, 137–155. Bayer, P., Arndt, A., Metzger, S., Mahajan, R., Melchior, F., Jaenicke, R., and Becker, J. (1998). Structure determination of the small ubiquitin-related modifier SUMO-1. J. Mol. Biol. 280, 275–286. Bernier-Villamor, V., Sampson, D. A., Matunis, M. J., and Lima, C. D. (2002). Structural basis for E2-mediated SUMO conjugation revealed by a complex between ubiquitin-conjugating enzyme Ubc9 and RanGAP1. Cell 108, 345–356. Broomfield, S., Chow, B. L., and Xiao, W. (1998). MMS2, encoding a ubiquitinconjugating-enzyme-like protein, is a member of the yeast error-free postreplication repair pathway. Proc. Natl. Acad. Sci. USA 95, 5678–5683. Broomfield, S., Hryciw, T., and Xiao, W. (2001). DNA postreplication repair and mutagenesis in Saccharomyces cerevisiae. Mutat. Res. 486, 167–184. Broomfield, S., and Xiao, W. (2002). Suppression of genetic defects within the RAD6 pathway by srs2 is specific for error-free post-replication repair but not for damageinduced mutagenesis. Nucleic Acids Res. 30, 732–739. Brown, M., Zhu, Y., Hemmingsen, S. M., and Xiao, W. (2002). Structural and functional conservation of error-free DNA postreplication repair in Schizosaccharomyces pombe. DNA Repair. 1, 869–880.
302
PASTUSHOK AND XIAO
Brusky, J., Zhu, Y., and Xiao, W. (2000). UBC13, a DNA-damage-inducible gene, is a member of the error-free postreplication repair pathway in Saccharomyces cerevisiae. Curr. Genet. 37, 168–174. Chau, V., Tobias, J. W., Bachmair, A., Marriott, D., Ecker, D. J., Gonda, D. K., and Varshavsky, A. (1989). A multiubiquitin chain is confined to specific lysine in a targeted short-lived protein. Science 243, 1576–1583. Chow, K. H., and Courcelle, J. (2004). RecO acts with RecF and RecR to protect and maintain replication forks blocked by UV-induced DNA damage in Escherichia coli. J. Biol. Chem. 279, 3492–3496. Cook, W. J., Jeffrey, L. C., Xu, Y., and Chau, V. (1993). Tertiary structures of class I ubiquitin-conjugating enzymes are highly conserved: Crystal structure of yeast Ubc4. Biochemistry 32, 13809–13817. Courcelle, J., Carswell-Crumpton, C., and Hanawalt, P. C. (1997). recF and recR are required for the resumption of replication at DNA replication forks in Escherichia coli. Proc. Natl. Acad. Sci. USA 94, 3714–3719. Desterro, J. M., Thomson, J., and Hay, R. T. (1997). Ubch9 conjugates SUMO but not ubiquitin. FEBS Lett. 417, 297–300. Dohmen, R. J., Madura, K., Bartel, B., and Varshavsky, A. (1991). The N-end rule is mediated by the UBC2(RAD6) ubiquitin-conjugating enzyme. Proc. Natl. Acad. Sci. USA 88, 7351–7355. Fisk, H. A., and Yaffe, M. P. (1999). A role for ubiquitination in mitochondrial inheritance in Saccharomyces cerevisiae. J. Cell. Biol. 145, 1199–1208. Galan, J. M., and Haguenauer-Tsapis, R. (1997). Ubiquitin Lys63 is involved in ubiquitination of a yeast plasma membrane protein. EMBO J. 16, 5847–5854. Haas, A. L., Reback, P. B., and Chau, V. (1991). Ubiquitin conjugation by the yeast RAD6 and CDC34 gene products. Comparison to their putative rabbit homologs, E2(20K) and E2(32K). J. Biol. Chem. 266, 5104–5112. Hershko, A., and Ciechanover, A. (1998). The ubiquitin system. Annu. Rev. Biochem. 67, 425–479. Hershko, A., Heller, H., Elias, S., and Ciechanover, A. (1983). Components of ubiquitin-protein ligase system. Resolution, affinity purification, and role in protein breakdown. J. Biol. Chem. 258, 8206–8214. Hochstrasser, M. (1996). Ubiquitin-dependent protein degradation. Annu. Rev. Genet. 30, 405–439. Hoege, C., Pfander, B., Moldovan, G. L., Pyrowolakis, G., and Jentsch, S. (2002). RAD6dependent DNA repair is linked to modification of PCNA by ubiquitin and SUMO. Nature 419, 135–141. Hofmann, R. M., and Pickart, C. M. (1999). Noncanonical MMS2-encoded ubiquitinconjugating enzyme functions in assembly of novel polyubiquitin chains for DNA repair. Cell 96, 645–653. Huang, L., Kinnucan, E., Wang, G., Beaudenon, S., Howley, P. M., Huibregtse, J. M., and Pavletich, N. P. (1999). Structure of an E6AP-UbcH7 complex: Insights into ubiquitination by the E2-E3 enzyme cascade. Science 286, 1321–1326. Hwang, W. W., Venkatasubrahmanyam, S., Ianculescu, A. G., Tong, A., Boone, C., and Madhani, H. D. (2003). A conserved RING finger protein required for histone H2B monoubiquitination and cell size control. Mol. Cell 11, 261–266. Jentsch, S. (1992). The ubiquitin-conjugation system. Annu. Rev. Genet. 26, 179–207. Jentsch, S., McGrath, J. P., and Varshavsky, A. (1987). The yeast DNA repair gene RAD6 encodes a ubiquitin-conjugating enzyme. Nature 329, 131–134.
DNA POSTREPLICATION REPAIR
303
Joazeiro, C. A., and Weissman, A. M. (2000). RING finger proteins: Mediators of ubiquitin ligase activity. Cell 102, 549–552. Johnson, E. S., and Blobel, G. (1997). Ubc9p is the conjugating enzyme for the ubiquitin-like protein Smt3p. J. Biol. Chem. 272, 26799–26802. Johnson, R. E., Henderson, S. T., Petes, T. D., Prakash, S., Bankmann, M., and Prakash, L. (1992). Saccharomyces cerevisiae RAD5-encoded DNA repair protein contains DNA helicase and zinc-binding sequence motifs and affects the stability of simple repetitive sequences in the genome. Mol. Cell. Biol. 12, 3807–3818. Kelman, Z., and Hurwitz, J. (1998). Protein-PCNA interactions: a DNA-scanning mechanism? Trends Biochem. Sci. 23, 236–238. Kogoma, T. (1997). Is RecF a DNA replication protein? Proc. Natl. Acad. Sci. USA 94, 3483–3484. Koken, M. H., Reynolds, P., Jaspers-Dekker, I., Prakash, L., Prakash, S., Bootsma, D., and Hoeijmakers, J. H. (1991). Structural and functional conservation of two human homologs of the yeast DNA repair gene RAD6. Proc. Natl. Acad. Sci. USA 88, 8865–8869. Koonin, E. V., and Abagyan, R. A. (1997). TSG101 may be the prototype of a class of dominant negative ubiquitin regulators. Nat. Genet. 16, 330–331. Kubota, Y., Nash, R. A., Klungland, A., Schar, P., Barnes, D. E., and Lindahl, T. (1996). Reconstitution of DNA base excision-repair with purified human proteins: Interaction between DNA polymerase beta and the XRCC1 protein. EMBO J. 15, 6662–6670. Kuzminov, A. (1999). Recombinational repair of DNA damage in Escherichia coli and bacteriophage lambda. Microbiol. Mol. Biol. Rev. 63, 751–813. Lawrence, C. W. (1982). Mutagenesis in Saccharomyces cerevisiae. Adv. Genet. 21, 173–254. Li, Z., Xiao, W., McCormick, J. J., and Maher, V. M. (2002). Identification of a protein essential for a major pathway used by human cells to avoid UV-induced DNA damage. Proc. Natl. Acad. Sci. USA 99, 4459–4464. Liberi, G., Chiolo, I., Pellicioli, A., Lopes, M., Plevani, P., Muzi-Falconi, M., and Foiani, M. (2000). Srs2 DNA helicase is involved in checkpoint response and its regulation requires a functional Mec1-dependent pathway and Cdk1 activity. EMBO J. 19, 5027–5038. Little, J. W. (1984). Autodigestion of lexA and phage lambda repressors. Proc. Natl. Acad. Sci. USA 81, 1375–1379. Lovering, R., Hanson, I. M., Borden, K. L., Martin, S., O’Reilly, N. J., Evan, G. I., Rahman, D., Pappin, D. J., Trowsdale, J., and Freemont, P. S. (1993). Identification and preliminary characterization of a protein motif related to the zinc finger. Proc. Natl. Acad. Sci. USA 90, 2112–2116. Martini, E. M., Keeney, S., and Osley, M. A. (2002). A role for histone H2B during repair of UV-induced DNA damage in Saccharomyces cerevisiae. Genetics 160, 1375–1387. McKenna, S., Moraes, T., Pastushok, L., Ptak, C., Xiao, W., Spyracopoulos, L., and Ellison, M. J. (2003). An NMR-based model of the ubiquitin-bound human ubiquitin conjugation complex Mms2.Ubc13. The structural basis for lysine 63 chain catalysis. J. Biol. Chem. 278, 13151–13158. McKenna, S., Spyracopoulos, L., Moraes, T., Pastushok, L., Ptak, C., Xiao, W., and Ellison, M. J. (2001). Noncovalent interaction between ubiquitin and the human DNA repair protein Mms2 is required for Ubc13-mediated polyubiquitination. J. Biol. Chem. 276, 40120–40126.
304
PASTUSHOK AND XIAO
Moraes, T. F., Edwards, R. A., McKenna, S., Pastushok, L., Xiao, W., Glover, J. N., and Ellison, M. J. (2001). Crystal structure of the human ubiquitin conjugating enzyme complex, hMms2-hUbc13. Nat. Struct. Biol. 8, 669–673. Ozkaynak, E., Finley, D., and Varshavsky, A. (1984). The yeast ubiquitin gene: Headto-tail repeats encoding a polyubiquitin precursor protein. Nature 312, 663–666. Paunesku, T., Mittal, S., Protic, M., Oryhon, J., Korolev, S. V., Joachimiak, A., and Woloschak, G. E. (2001). Proliferating cell nuclear antigen (PCNA): Ringmaster of the genome. Int. J. Radiat. Biol. 77, 1007–1021. Pham, P., Seitz, E. M., Saveliev, S., Shen, X., Woodgate, R., Cox, M. M., and Goodman, M. F. (2002). Two distinct modes of RecA action are required for DNA polymerase V-catalyzed translesion synthesis. Proc. Natl. Acad. Sci. USA 99, 11061–11066. Pickart, C. M. (2001). Mechanisms underlying ubiquitination. Annu. Rev. Biochem. 70, 503–533. Prakash, S., Sung, P., and Prakash, L. (1993). DNA repair genes and proteins of Saccharomyces cerevisiae. Annu. Rev. Genet. 27, 33–70. Reuven, N. B., Arad, G., Maor-Shoshani, A., and Livneh, Z. (1999). The mutagenesis protein UmuC is a DNA polymerase activated by UmuD’, RecA, and SSB and is specialized for translesion replication. J. Biol. Chem. 274, 31763–31766. Richmond, E., and Peterson, C. L. (1996). Functional analysis of the DNA-stimulated ATPase domain of yeast SWI2/SNF2. Nucleic Acids Res. 24, 3685–3692. Robzyk, K., Recht, J., and Osley, M. A. (2000). Rad6-dependent ubiquitination of histone H2B in yeast. Science 287, 501–504. Rodriguez, M. S., Dargemont, C., and Hay, R. T. (2001). SUMO-1 conjugation in vivo requires both a consensus modification motif and nuclear targeting. J. Biol. Chem. 276, 12654–12659. Roest, H. P., van Klaveren, J., de Wit, J., van Gurp, C. G., Koken, M. H., Vermey, M., van Roijen, J. H., Hoogerbrugge, J. W., Vreeburg, J. T., Baarends, W. M., Bootsma, D., Grootegoed, J. A., and Hoeijmakers, J. H. (1996). Inactivation of the HR6B ubiquitin-conjugating DNA repair enzyme in mice causes male sterility associated with chromatin modification. Cell 86, 799–810. Rupp, W. D., and Howard-Flanders, P. (1968). Discontinuities in the DNA synthesized in an excision-defective strain of Escherichia coli following ultraviolet irradiation. J. Mol. Biol. 31, 291–304. Salles, B., and Defais, M. (1984). Signal of induction of recA protein in E. coli. Mutat. Res. 131, 53–59. Sampson, D. A., Wang, M., and Matunis, M. J. (2001). The small ubiquitin-like modifier-1 (SUMO-1) consensus sequence mediates Ubc9 binding and is essential for SUMO-1 modification. J. Biol. Chem. 276, 21664–21669. Sancho, E., Vila, M. R., Sanchez-Pulido, L., Lozano, J. J., Paciucci, R., Nadal, M., Fox, M., Harvey, C., Bercovich, B., Loukili, N., Ciechanover, A., Lin, S. L., Sanz, F., Estivill, X., Valencia, A., and Thomson, T. M. (1998). Role of UEV-1, an inactive variant of the E2 ubiquitin-conjugating enzymes, in in vitro differentiation and cell cycle behavior of HT-29-M6 intestinal mucosecretory cells. Mol. Cell. Biol. 18, 576–589. Saurin, A. J., Borden, K. L., Boddy, M. N., and Freemont, P. S. (1996). Does this have a familiar RING? Trends Biochem. Sci. 21, 208–214. Scheffner, M., Nuber, U., and Huibregtse, J. M. (1995). Protein ubiquitination involving an E1-E2-E3 enzyme ubiquitin thioester cascade. Nature 373, 81–83. Schiestl, R. H., Prakash, S., and Prakash, L. (1990). The SRS2 suppressor of rad6 mutations of Saccharomyces cerevisiae acts by channeling DNA lesions into the RAD52 DNA repair pathway. Genetics 124, 817–831.
DNA POSTREPLICATION REPAIR
305
Schlesinger, D. H., Goldstein, G., and Niall, H. D. (1975). The complete amino acid sequence of ubiquitin, an adenylate cyclase stimulating polypeptide probably universal in living cells. Biochemistry 14, 2214–2218. Schwartz, D. C., and Hochstrasser, M. (2003). A superfamily of protein tags: Ubiquitin, SUMO and related modifiers. Trends Biochem. Sci. 28, 321–328. Schwarz, S. E., Matuschewski, K., Liakopoulos, D., Scheffner, M., and Jentsch, S. (1998). The ubiquitin-like proteins SMT3 and SUMO-1 are conjugated by the UBC9 E2 enzyme. Proc. Natl. Acad. Sci. USA 95, 560–564. Seufert, W., Futcher, B., and Jentsch, S. (1995). Role of a ubiquitin-conjugating enzyme in degradation of S- and M-phase cyclins. Nature 373, 78–81. Sheng, W., and Liao, X. (2002). Solution structure of a yeast ubiquitin-like protein Smt3: The role of structurally less defined sequences in protein–protein recognitions. Protein Sci. 11, 1482–1491. Shinagawa, H., Iwasaki, H., Kato, T., and Nakata, A. (1988). RecA protein-dependent cleavage of UmuD protein and SOS mutagenesis. Proc. Natl. Acad. Sci. USA 85, 1806–1810. Spence, J., Gali, R. R., Dittmar, G., Sherman, F., Karin, M., and Finley, D. (2000). Cell cycle-regulated modification of the ribosome by a variant multiubiquitin chain. Cell 102, 67–76. Stelter, P., and Ulrich, H. D. (2003). Control of spontaneous and damageinduced mutagenesis by SUMO and ubiquitin conjugation. Nature 425, 188–191. Sung, P., Berleth, E., Pickart, C., Prakash, S., and Prakash, L. (1991). Yeast RAD6 encoded ubiquitin conjugating enzyme mediates protein degradation dependent on the N-end-recognizing E3 enzyme. EMBO J. 10, 2187–2193. Sung, P., Prakash, S., and Prakash, L. (1988). The RAD6 protein of Saccharomyces cerevisiae polyubiquitinates histones, and its acidic domain mediates this activity. Genes Dev. 2, 1476–1485. Sung, P., Prakash, S., and Prakash, L. (1990). Mutation of cysteine-88 in the Saccharomyces cerevisiae RAD6 protein abolishes its ubiquitin-conjugating activity and its various biological functions. Proc Natl Acad Sci USA 87, 2695–2699. Takahashi, Y., Kahyo, T., Toh, E. A., Yasuda, H., and Kikuchi, Y. (2001a). Yeast Ul11/Siz1 is a novel SUMO1/Smt3 ligase for septin components and functions as an adaptor between conjugating enzyme and substrates. J. Biol. Chem. 276, 48973–48977. Takahashi, Y., Toh-e, A., and Kikuchi, Y. (2001b). A novel factor required for the SUMO1/Smt3 conjugation of yeast septins. Gene 275, 223–231. Tang, M., Bruck, I., Eritja, R., Turner, J., Frank, E. G., Woodgate, R., O’Donnell, M., and Goodman, M. F. (1998). Biochemical basis of SOS-induced mutagenesis in Escherichia coli: reconstitution of in vitro lesion bypass dependent on the UmuD0 2C mutagenic complex and RecA protein. Proc. Natl. Acad. Sci. USA 95, 9755–9760. Tang, M., Shen, X., Frank, E. G., O’Donnell, M., Woodgate, R., and Goodman, M. F. (1999). ‘‘UmuD0 (2)C is an error-prone DNA polymerase, Escherichia coli pol V. Proc. Natl. Acad. Sci. USA 96, 8919–8924. Tateishi, S., Sakuraba, Y., Masuyama, S., Inoue, H., and Yamaizumi, M. (2000). Dysfunction of human Rad18 results in defective postreplication repair and hypersensitivity to multiple mutagens. Proc. Natl. Acad. Sci. USA 97, 7927–7932. Torres-Ramos, C. A., Yoder, B. L., Burgers, P. M., Prakash, S., and Prakash, L. (1996). Requirement of proliferating cell nuclear antigen in RAD6-dependent postreplicational DNA repair. Proc. Natl. Acad. Sci. USA 93, 9676–9681.
306
PASTUSHOK AND XIAO
Ulrich, H. D. (2001). The srs2 suppressor of UV sensitivity acts specifically on the RAD5and MMS2-dependent branch of the RAD6 pathway. Nucleic Acids Res. 29, 3487–3494. Ulrich, H. D. (2003). Protein–protein interactions within an E2-RING finger complex. Implications for ubiquitin-dependent DNA damage repair. J. Biol. Chem. 278, 7051–7058. Ulrich, H. D., and Jentsch, S. (2000). Two RING finger proteins mediate cooperation between ubiquitin-conjugating enzymes in DNA repair. EMBO J. 19, 3388–3397. VanDemark, A. P., and Hill, C. P. (2002). Structural basis of ubiquitylation. Curr. Opin. Struct. Biol. 12, 822–830. VanDemark, A. P., Hofmann, R. M., Tsui, C., Pickart, C. M., and Wolberger, C. (2001). Molecular insights into polyubiquitin chain assembly: Crystal structure of the Mms2/Ubc13 heterodimer. Cell 105, 711–720. Vijay-Kumar, S., Bugg, C. E., and Cook, W. J. (1987). Structure of ubiquitin refined at 1.8 A resolution. J. Mol. Biol. 194, 531–544. Vijay-Kumar, S., Bugg, C. E., Wilkinson, K. D., and Cook, W. J. (1985). Three-dimensional structure of ubiquitin at 2.8 A resolution. Proc. Natl. Acad. Sci. USA 82, 3582–3585. Villalobo, E., Morin, L., Moch, C., Lescasse, R., Hanna, M., Xiao, W., and BaroinTourancheau, A. (2002). A homologue of CROC-1 in a ciliated protist (Sterkiella histriomuscorum) testifies to the ancient origin of the ubiquitin-conjugating enzyme variant family. Mol. Biol. Evol. 19, 39–48. Watkins, J. F., Sung, P., Prakash, S., and Prakash, L. (1993). The extremely conserved amino terminus of RAD6 ubiquitin-conjugating enzyme is essential for amino-end rule-dependent protein degradation. Genes Dev. 7, 250–261. Webb, B. L., Cox, M. M., and Inman, R. B. (1997). Recombinational DNA repair: The RecF and RecR proteins limit the extension of RecA filaments beyond single-strand DNA gaps. Cell 91, 347–356. Wood, A., Krogan, N. J., Dover, J., Schneider, J., Heidt, J., Boateng, M. A., Dean, K., Golshani, A., Zhang, Y., Greenblatt, J. F., Johnston, M., and Shilatifard, A. (2003). Brel, an E3 ubiquitin ligase required for recruitment and substrate selection of Rad6 at a promoter. Mol. Cell 11, 267–274. Worthylake, D. K., Prakash, S., Prakash, L., and Hill, C. P. (1998). Crystal structure of the Saccharomyces cerevisiae ubiquitin-conjugating enzyme Rad6 at 2.6 A resolution. J. Biol. Chem. 273, 6271–6276. Wu-Baer, F., Lagrazon, K., Yuan, W., and Baer, R. (2003). The BRCA1/BARD1 heterodimer assembles polyubiquitin chains through an unconventional linkage involving lysine residue K6 of ubiquitin. J. Biol. Chem. 278, 34743–34746. Xiao, W., Chow, B. L., Broomfield, S., and Hanna, M. (2000). The Saccharomyces cerevisiae RAD6 group is composed of an error-prone and two error-free postreplication repair pathways. Genetics 155, 1633–1641. Xiao, W., Chow, B. L., Fontanie, T., Ma, L., Bacchetti, S., Hryciw, T., and Broomfield, S. (1999). Genetic interactions between error-prone and error-free postreplication repair pathways in Saccharomyces cerevisiae. Mutat. Res. 435, 1–11. Xiao, W., Lin, S. L., Broomfield, S., Chow, B. L., and Wei, Y. F. (1998). The products of the yeast MMS2 and two human homologs (hMMS2 and CROC-1) define a structurally and functionally conserved Ubc-like protein family. Nucleic Acids Res. 26, 3908–3914. Zheng, N., Wang, P., Jeffrey, P.D., and Pavletich, N.P. (2000). Structure of a cCb1-UbcH7 complex: RING domain function in ubiquitin-protein ligases. Cell 102, 533–539.
SOMATIC HYPERMUTATION: A MUTATIONAL PANACEA By BRIGETTE TIPPIN, PHUONG PHAM, RONDA BRANSTEITTER, AND MYRON F. GOODMAN Departments of Biological Sciences and Chemistry, University of Southern California, Los Angeles, California, 90089
I. Generation of Antibody Diversity . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . II. Somatic Hypermutation . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Discovery of a Role for AID . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. Separation of AID activities for SHM and CSR . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . C. Initiation of SHM and CSR by AID . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . D. Some Like It Hot—Biochemical Aspects of AID . . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . E. Mutational Models Depicting Two-Phase Somatic Hypermutation . . . .. . . . . . F. Involvement of MMR and BER . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . G. Regulation of AID Expression in SHM and Its Importance in Cancer Avoidance . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . III. Apobec Protein Family . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Apobec-1. . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. Antiretroviral Activity of Apobec3G, G ! A Hypermutation in HIV and Other Retroviruses. . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . IV. Biochemical Perspective. . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . .
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I. Generation of Antibody Diversity DNA damage, be it chromosomal breaks or base alterations leading to erroneous coding information, are extremely harmful when occurring randomly within the genome, but when controlled and targeted, the outcome can be extremely beneficial to the organism’s survival and overall fitness. Nature has harnessed the power of mutagenesis for productive measures most prominently during the generation of antibody diversity in the immune response of vertebrates. Humans are born with a subset of genes dedicated for the production of B-cell antibodies and T-cell receptors, the immunoglobulin genes. To achieve the tremendous diversity necessary to recognize a myriad of foreign invaders to the body, an elaborate series of mechanisms has evolved to convert the original germ-line set of immunoglobulin genes into millions of variants. Antibody variation occurs for two main purposes. The first is to allow recognition and binding of antigen that is achieved through V(D)J 307 ADVANCES IN PROTEIN CHEMISTRY, Vol. 69
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recombination and somatic hypermutation (SHM). The second is to allow antigen binding sites that are created to be expressed with the eight different constant region genes so that they can mediate many different effecter functions throughout the body by a process called class-switch recombination (CSR). V(D)J recombination is the first step in generating antibody diversity and occurs throughout early B-cell development in the bone marrow. This specialized form of recombination introduces site-specific doublestranded DNA breaks between multiple germline copies of V, D, and J segments in an immunoglobulin gene and subsequently re-assembles the fragments to produce transcriptionally competent heavy- or light-chain genes (Fig. 1). A functional antibody is composed of four distinct immunoglobulin polypeptides: two heavy chains (IgH), and two light chains of either the lambda (Ig1) or kappa (Igk) type. Diversity is achieved by the random choice of which V, D, and J segments are rejoined to one another and also through errors that are introduced at the junction sites before the breaks are sealed (reviewed in Gellert, 2002). Transcription and translation of functional heavy- and light-chain genes that have successfully undergone V(D)J recombination are essential for proper progression through B-cell developmental stages. If improper rejoining occurs in either heavy or light chains, the cell will not receive the proper signals and thus undergo apoptosis. T-cell-receptor (TCR) genes also undergo V(D)J recombination in T cells in the thymus (Krangel, 2003; Strominger, 1989). Initiation of V(D)J recombination begins by the action of site-specific recombination-activating genes (RAG1) and RAG2 (Mombaerts et al., 1992; Oettinger et al., 1990; Schatz et al., 1989; Shinkai et al., 1992) in a complex with high-mobility group (HMG) 1 proteins (Sawchuk et al., 1997; van Gent et al., 1997), which introduce double-stranded breaks at recombination signal sequences (RSS) adjacent to the multicopy V, D, and J segments (Lieber et al., 2003). Conversion of break points into hairpins by the complex protects the ends from rejoining prior to removal of unwanted gene segments. Next, Ku70/80 heterodimer (Casellas et al., 1998; Manis et al., 1998) binds to the ends, possibly displacing the RAG complex, and a different complex made up of Artemis (Moshous et al., 2001) and the DNA dependent protein kinase catalytic subunit (DNAPKcs) (Blunt et al., 1995; Kirchgessner et al., 1995; Peterson et al., 1995) is recruited by Ku70/80 to open the hairpinned V, D, or J ends (Ma et al., 2002). Artemis-DNA-PKcs acts as a nuclease, trimming the ends and increasing the diversity at the junctions (Ma et al., 2002). At this intermediate step, terminal deoxynucleotidyl transferase (TdT) (Gilfillan et al., 1993; Kohler and Milstein, 1975; Komori et al., 1993), can add extra nucleotides
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Fig. 1. High-affinity antibodies in mice and human are generated by V(D)J joining and subsequent diversification by somatic hypermutation (SHM) and class-switch recombination (CSR). The primary antibody repertoire is generated during B-cell development by site-specific recombination resulting in the fusion of germline V, D, and J gene segments. On encounter with antigens, the V(D)J rearranged genes further undergo SHM and CSR to generate high-affinity antibodies. CSR selection to downstream constant domains
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in a template-independent manner to increase diversity at the junctions, but TdT’s contribution is limited by its restricted expression in B cells at the early pro-B cell developmental stage (Li et al., 1993). During the late pre-B and immature B stages of development, DNA polymerase (mu) might be recruited by Ku70/80 (Mahajan et al., 2002) in lieu of TdT to participate in V-J end-processing at microhomologies (Bertocci et al., 2003). Finally, the combined action of XRCC4 (X-ray cross complementation 4) and DNA-ligase-IV link the two coding DNA ends to make a functional immunoglobulin gene (Gao et al., 1998; Grawunder et al., 1998; Li et al., 1995; Taccioli et al., 1993). Following the completion of V(D)J recombination in both heavy- and light-chain immunoglobulin genes, the B cell reaches maturity and expresses a complete four subunit antibody on its surface. Mature B cells migrate from the bone marrow to peripheral lymphoid tissues such as the lymph nodes, spleen, Peyer’s patches in the gut, or tonsils, where they await an encounter with foreign antigen. The initial diversity created in each individual mature B cell during V(D)J recombination provides a wide range of low-affinity binding capabilities for antigens. Once a positive interaction has occurred between an antibody on the B cell surface (e.g., IgM, IgD), signaling from a T helper cell triggers the B cell to form germinal centers (GCs) in peripheral lymph nodes. In the GC, B cells bound to antigen will undergo positive selection and rapid cell division to create more clones. During this proliferation process, the second stages of diversification begin to take place; CSR to convert antibodies into different isoforms that can better trap, neutralize, and clear antigens from the body in the immune response through unique effector functions; and SHM targeted to the V (variable) region in IgH and IgL genes that improve binding of antigen (Fig. 1). CSR occurs in the Ig heavy-chain locus in the C (constant) domain located downstream of the V(D)J variable region (Fig. 1). Eight different CH regions encode unique isotypes IgM, IgD, 4 distinct IgG, IgE, or IgA. Initially, mature B cells translate the IgH locus using the most upstream exon C to produce IgM that is expressed as a low-affinity membrane bound and secreted isotype. The expression of the C heavy chain is accomplished by mRNA splicing. Each of the other six C regions contains a cryptic promoter and repetitive switch region (S) located 50 of the exon
[ 3, 1, ( 2b and 2a not shown), , or "] allows switching from IgM or IgD to other the isotypes IgG3, IgG1, (IgG2b and IgG2a not shown), IgA, or IgE. P represents cryptic promoters adjacent to switch (S) regions, E and E30 are enhancer regulatory regions, and HS4 is an enhancer binding site within the E30 region important for CSR.
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coding sequence. After antigen engagement, one or another of the other CH exons is brought into the position adjacent to the V(D)J region (displacing C), converting the antibody to a different isotype that can be expressed on the surface or be secreted (Fig. 1). The complete mechanism of CSR is far less well understood than V(D)J recombination, but some key features and players have been defined. Unlike V(D)J recombination, CSR only occurs in B-lineage cells, and although CSR is triggered during antigen-driven clonal expansion in germinal centers (GCs), the process can also occur in other regions of the body via T-cell-dependent interactions with CD40 ligand or T-cell-dependent antigens such as lipopolysaccharide (LPS) in combination with cytokine activators such as interleukin-4 (IL-4) (Stavnezer, 2000). CSR is directed initially by transcription from the cryptic promoter 50 of the exon targeted for isotype switching by unique cellular signals (Manis et al., 2002a). The B-cell-specific transcriptional enhancer E, located between the JH and C region (Fig. 1), can stimulate transcription from the CH promoters, but this enhancer is not absolutely required (Bottaro et al., 1998) for CSR. A second cluster of enhancers located 30 of the IgH C region (in particular the HS4 enhancer element) can also interact with the activated CH promoter (Fig. 1) and appears to serve as the regulatory region for CSR (Khamlichi et al., 2000). Germline CH RNA transcripts generated from these promoters will undergo splicing and polyadenylation, but they are not translated into protein (Stavnezer, 2000; Zhang et al., 1995). Rather, the transcripts themselves appear to help target CSR machinery to a particular S region (Bottaro et al., 1994; Harriman et al., 1996; Jung, 1993; Seidl et al., 1998). S regions can vary greatly in different CH segments (Fig. 1), with some composed of tandem pentamer repeats and others having 49-base-pair repeats. The lack of a consensus break point or fusion sequence makes CSR a region-specific process (Kinoshita et al., 1999), rather than a site-specific recombination process as seen for V(D)J recombination. Various models have been suggested to explain the regional break phenomenon via higher-order structures in the S-regions. However the recent discovery of the activation-induced cytidine deaminase (AID) (Muramatsu et al., 1999) has led to the proposal that CSR is actually targeted by AID-initiated strand-breaks on single-stranded DNA generated during germline transcription (Chaudhuri et al., 2003; Yu and Lieber, 2003). Both CSR (Lee et al., 2001) and SHM require transcription (Bachl et al., 2001; Peters and Storb, 1996) as a precursor step in their mechanism, and both pathways are abrogated in the absence of AID (Muramatsu et al., 2000), but the two processes can be differentially induced in response to unique signals. For example, activation of splenic B cells by LPS leads to
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CSR and not SHM (Manis et al., 2002b). In the case of CSR, transcription through an S region would generate single-stranded stretches of DNA on which AID could deaminate C bases, any of which could be converted into a single-stranded break by the subsequent action of a uracil N-glycosylase (UNG) and AP-endonuclease (Rada et al., 2002). The conversion of single-stranded breaks into double-stranded breaks for a CSR event and subsequent end-processing by NHEJ component proteins (Casellas et al., 1998; Manis et al., 1998; Rolink et al., 1996), and perhaps DNA-PKcs (Bosma et al., 2002; Manis et al., 2002a), would ultimately leave little trace of a consensus motif by the completion of the repair process. The action of AID on any available C in a single-stranded piece of DNA could help explain the randomness of breakpoints seen. In addition, R loops that form during CSR might contain short regions of ssDNA on both strands at the transition between the R loop and the adjacent dsDNA (Chaudhuri et al., 2003; Yu and Lieber, 2003) on which AID could trigger a double-stranded break. However, what limits AID access from acting outside of the S-region during transcription is not yet understood. SHM, like CSR, is induced in immunoglobulin genes following antigen recognition by low-affinity antibody on B cells that migrate to germinal centers. SHM is specifically targeted to V regions in the immunoglobulin heavy- and light-chain genes (Lebecque and Gearhart, 1990). Mutations that result in improved affinity for antigen become positively selected, eventually leading to the development of ‘‘immunity’’ to the infectious antigen. The key elements necessary for SHM include active transcription (boosted by enhancer elements) (Maizels, 1995; Peters and Storb, 1996), high fidelity (Hi Fi), and error-prone (EP) DNA polymerases, and most prominently, AID (Fig. 2). SHM requires AID, with initiation now known to involve AID-catalyzed C ! U conversion on ssDNA (Bransteitter et al., 2003; Chaudhuri et al., 2003; Dickerson et al., 2003; Pham et al., 2003; Sohail et al., 2003), presumably within a transcription bubble on the nontranscribed strand. C ! T transitions are the most commonly observed V-gene mutations and are notably favored in WRC hotspot sequences (W ¼ A or T, R ¼ purine) that account for roughly half of all documented SHM mutations (Golding et al., 1987; Rogozin and Kolchanov, 1992), and will result if U remains in the DNA and is subsequently copied with a normal Hi Fi pol (i.e., pols or "; Fig. 3). The schematic description shown in Figure 2 provides a general picture of enzymes and pathways used during SHM, without committing to specific molecular models. However, with recent progress made in understanding the action of AID, molecular mechanisms can now begin to be investigated (Fig. 3).
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Fig. 2. Integrated model for SHM. Interactions between a transcription factor, activated by the E enhancer, and transcription complex bound at the promoter (P) lead to transcription of Ig genes. Transcription-dependent AID-mediated deamination initiates SHM by converting C to U, preferentially at 50 WRC hotspots in the transiently created transcription bubbles. Subsequent replication of U-containing DNA by high fidelity (Hi-Fi) polymerases or by error-prone repair mechanisms ‘‘fix’’ SHM mutations at 50 WRC and 50 TAA motifs. Single-strand nicks or double-strand breaks have been also observed in the V-region of Ig genes at 50 WRC hot-spot motifs, caused presumably by an as yet unidentified endonuclease (Endo). The DNA nicks or breaks may be substrates for one or more error-prone polymerases (EP pols) to bind and generate mutations. An example of EP-pols is Pol , which is responsible for mutating A and T sites, primarily within TAA motifs. MAR designates a matrix attachment region, DJ—the D and J joined with V-region by V(D)J recombination, C-region—the constant region of Ig genes.
II. Somatic Hypermutation A. Discovery of a Role for AID AID was first discovered using a subtractive hybridization screen for genes activated on induction of CSR(Muramatsu et al., 1999) and has since been shown to serve highly critical roles in a number of immune-specific processes. AID expression is restricted to activated B cells (Muramatsu et al., 1999) and is required for SHM in mice (Muramatsu et al., 2000). AID was also found to be the culprit in a human immune disorder, Hyper-IgM-2 syndrome, in which some patients exhibited AID deficiency, resulting in the abolishment of both CSR and SHM (Revy et al., 2000). Hyper-IgM syndrome (HIGM) is characterized by normal or elevated serum IgM levels associated with the absence of IgG, IgA, and IgE isotypes as a result of defective CSR (Notarangelo et al., 1992). Additional experiments in mice and B cell lines have verified this AID requirement (Martin and Scharff, 2002; Muramatsu et al., 2000; Revy et al., 2000). AID transfection studies have further shown that AID is likely to be the only B-cell-specific protein required for SHM and CSR. Indeed,
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transfection of AID into human B-cells at the improper stage of differentiation (Martin and Scharff, 2002), into non-B cells (Martin and Scharff, 2002), and surprisingly even in Escherichia coli (Petersen-Mahrt et al., 2002), is sufficient to induce hypermutation in all of these cell types, whereas ectopic expression of AID in fibroblasts induces CSR on artificial substrates (Okazaki et al., 2002). Yet another interesting role for AID is seen in chicken DT-40 cells, which are unable to carry out the process of gene conversion in the absence of AID (Arakawa et al., 2002; Flajnik, 2002; Harris et al., 2002). Gene conversion is an alternative pathway for generating antibody diversity used by chickens and rabbits in place of SHM. Inactivation of homologous recombination genes XRCC2, XRCC3, or RAD51B in chicken DT40 cells also has the same effect of impairing gene conversion, but in the presence of AID, these cells shift to using the SHM pathway that normally does not occur in this cell line (Sale et al., 2001). Because both gene-conversion and SHM pathways are inactivated when AID is lacking in DT-40, it appears that under normal conditions, when AID is present, the type of repair protein recruited to a uracil lesion may be the determinant of the mechanistic outcome, either gene conversion or SHM. Taken together, the evidence indicates that AID plays a universal role overseeing entry into all immunoglobulin gene-specific modifications that increase antibody diversification in a variety of species. AID was initially thought to function by editing mRNA that encodes an endonuclease that initiates breaks in V and S regions of Ig genes (Honjo et al., 2002), but an RNA substrate for AID in vivo has not been identified. AID, when expressed in E. coli, is able to mutate genes on the bacterial chromosome (Petersen-Mahrt et al., 2002), the first indication that the enzyme might act specifically on DNA and not RNA. Biochemical data subsequently showed that AID deaminates C residues on ssDNA (Bransteitter et al., 2003; Chaudhuri et al., 2003; Dickerson et al., 2003).
Fig. 3. SHM branched pathway involving the initiator action of AID. Shown at the left is a region of DNA undergoing transcription (RNA transcript indicated by a curved line in blue). AID acting on ssDNA exposed in the transcription bubble deaminates C in the WRC hot spot sequence to initiate the first ‘‘phase’’ of SHM. Subsequent copying of U by normal replication polymerases will result in a C ! T mutation or alternatively an error-prone (EP) polymerase can generate a C ! N mutation. A second phase of SHM (SHM diversification) can occur by using the postreplication mismatch repair (MMR) pathway to excise the U-G mismatch and generate a repair patch in which a second deamination by AID can occur, and/or EP synthesis can generate the WA mutational hotspots. SHM diversification can also occur using the base excision repair (BER) pathway triggered by the action of uracil glycosylase (UNG) and apurinic/apyrimidinic endonuclease (APE) to excise uracil and nick the DNA backbone. AID-catalyzed deamination of C is responsible for initiating both phases of SHM. (See Color Insert.)
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A transgenic approach has been taken to further investigate whether or not AID deamination of C ! U was directly involved in SHM by looking at Uracil N–glyclosylase (UNG) deficiencies. In normal cells, UNG plays a significant role in removing uracil from DNA via the base-excision repair (BER) pathway (Lindahl and Barnes, 2001). One would predict that if AID deaminates DNA, then UNG-deficient mice would fail to remove uracil from the DNA, and when replicated, this would result in an increased frequency of transition mutations, especially C!T. UNG-deficient mice and human cells do exhibit altered mutational patterns with C:G ! T:A transitions that are significantly greater than transversions (Di Noia and Neuberger, 2002; Imai et al., 2003b; Rada et al., 2002). In these same studies, CSR is also inhibited, most likely because of a lack of necessary double-stranded breaks that likely arise during the normal uracil removal process. Although other uracil DNA glycosylases such as MBD4, TDG, and SMUG are present in UNG deficient mice and humans, they do not appear to compensate for loss of UNG activity at the Ig locus. In fact, in MBD4 knockout mice, neither SHM nor CSR is perturbed (Bardwell et al., 2003). Other deficiencies of BER proteins such as DNA glycosylases AAG and OGG1, Poly (ADP-ribose) polymerase, and Pol fail to alter or impair SHM (Esposito et al., 2000; Jacobs et al., 1998; Winter et al., 2003). Aside from the XPV gene (encoding DNA polymerase ), proteins from the nucleotide excision repair (NER) pathway also fail to have any effect on SHM ( Jacobs et al., 1998). The effect of the UNG deficiencies therefore demonstrates that introduction of U into DNA by AID occurs in vivo, and recapitulates the in vitro biochemical activity found for AID.
B. Separation of AID Activities for SHM and CSR Mutations in AID suggest that it functions as a multimer of two or four identical subunits, and that specific domains in each peptide may be responsible for recruiting factors that mediate the outcome of SHM and CSR. Expression of a double mutant in the catalytic domain of AID in a Ramos B cell line has a dominant negative effect on the wild-type AID, abrogating normal activity (measured by SHM), and argues that complexes must form between mutant and wild-type proteins (Papavasiliou and Schatz, 2002). A naturally occurring dominant negative mutant was also found in a Hyper-IgM2 patient who was determined to have a heterozygous mutation within AICDA, the gene encoding AID (Kasahara et al., 2003). Typical Hyper-IgM type 2 patients contain homozygous mutations within AICDA and lack CSR as well as SHM functions (Revy et al., 2000). There are alternative forms of the syndrome that reveal possible separation of AID
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activity between CSR or SHM pathways. HIGM type 4 patients maintain SHM activity yet lack CSR (Imai et al., 2003a). Mutations within the 189–198 amino acids of the C-terminal portion of AID produce the HIGM type 4 phenotype (Barreto et al., 2003; Ta et al., 2003), implying a critical role for this region during CSR, but an expendable role in SHM. Of note, the C-terminal mutants also exhibit increased cytidine deaminase activity compared to wild-type AID when expressed in E. coli (Barreto et al., 2003). Overall, these results suggest that different domains on AID must require the interaction of additional factors to specify the pathway of AID targeting to either SHM or CSR in response to unique cellular signals. In addition to the naturally occurring mutations within AICDA, three alternative spliced forms of AID have been found. In Splice variant 1 (SV1), the intron between exon 3 and 4 is retained. In splice variant 2 (SV2), exon 4 is removed. Last, in splice variant 3 (SV3), a short neoexon in intron 3 is retained and exons 3 and 4 are spliced out (Greeve et al., 2003). SV1 and SV2 are expressed during the centrocyte B cell stage, a time when AID is typically down-regulated in the cell (McCarthy et al., 2003). In contrast, these same alternatively spliced transcripts (SV1 and SV2) have also been found in B-cell lymphomas at the centroblast stage, the normal stage for high-level AID expression (Greeve et al., 2003; McCarthy et al., 2003; Oppezzo et al., 2003). In the case of the B centroblast lymphomas, in the combined presence of the splice variants and high-level wt AID, SHM is not observed (McCarthy et al., 2003). A possible explanation for these results is that the alternative spliced forms of AID may act as dominate negative regulators, serving to downregulate AID function in healthy centrocytes, repressing SHM and CSR, but when abnormally expressed in centroblasts, may trigger B cell lymphogenesis. The splice variant studies have also revealed even further evidence of the ability to uncouple CSR and SHM through the function of AID. In follicular lymphoma B cells, the distinct pattern of SV2 and SV3 expression exhibits intact SHM, but is defective for CSR (Greeve et al., 2003). In contrast, in another study on chronic lymphoid leukemia (CLL), cells that express both wt AID and splice variants were found to have defective SHM but intact CSR (Oppezzo et al., 2003).
C. Initiation of SHM and CSR by AID Baculovirus-expressed GST-AID fusion protein revealed that not only does AID act specifically to deaminate C residues in ssDNA in vitro, but it fails to work on dsDNA, DNA/RNA hybrids or RNA alone (Bransteitter et al., 2003). The initially cryptic activity of AID was revealed after it was found that the purified GST-AID was bound to an inhibitory RNA
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molecule, but the putative biological relevance of this RNA inhibition of AID is yet to be determined (Bransteitter et al., 2003). Additional studies using partially purified AID from B cell extracts as well as AID purified from E. coli also showed deamination of ssDNA in vitro (Chaudhuri et al., 2003; Dickerson et al., 2003; Sohail et al., 2003). During activation of SHM and CSR, AID must somehow gain access to ssDNA, which is postulated to occur through opening of a transcription bubble in the V region (Fig. 2) or at one of the various CH loci. Evidence to support this model comes from several experiments. First, AID-dependent deamination during a prokaryotic RNA polymerase-driven transcription reaction exhibits a 15-fold preference for the nontranscribed strand (Pham et al., 2003; Ramiro et al., 2003; Sohail et al., 2003). Second, GST-AID activity was found to be greatest on a small 9-nt transcription-like DNA bubble (Bransteitter et al., 2003), and although any connection of this observation to SHM in the ‘‘real world’’ would obviously be naive, it may indicate a potential connection to transcriptional targeting. Third, it is well documented that both CSR and SHM require transcription to proceed (Maizels, 1995; Manis et al., 2002b; Peters and Storb, 1996), and a transcription bubble constitutes an obvious potential entry point for AID as transcription generates ssDNA regions on the nontranscribed strand. Regardless of the specific targeting mechanisms for AID to V regions for SHM and C regions for CSR, the genetic and biochemical data suggest that AID deamination on DNA in Ig genes is a critical early step in these processes.
D. Some Like It Hot—Biochemical Aspects of AID Attempts to reconstitute the entire SHM process in a test tube are in their infancy; however, there are already several biochemical observations involving AID that directly simulate some of the most salient features of SHM. A hallmark of SHM is that transition mutations are favored over transversions occurring predominantly in WRC hotspot sequences (including GYW on the opposite strand) (Rogozin and Kolchanov, 1992). Perhaps the strongest biochemical evidence for a direct role of AID on DNA during SHM was obtained through analysis of the mutational spectrum for GST-AID on ssDNA that revealed preferential deamination of dC in WRC hotspot motifs (Pham et al., 2003). AID, acting on ‘‘naked’’ ssDNA, with no additional cofactors, exhibits precisely the same sequence specificity for C ! U conversions (Pham et al., 2003). The WRC mutability index, which is defined as the number of times an oligonucleotide sequence within a segment of DNA contains a mutation divided by the number of times the sequence is expected to mutate for a mechanism with
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no sequence bias, has a value for AID of 1.85 0.26 compared to 1.88 0.59 for SHM in vivo (Pham et al., 2003). This remarkably close agreement between the biochemical specificity of AID and biological specificity of SHM indicates that the V-gene targeting at WRC hotspots may depend primarily, if not exclusively, on AID. AID also appears to act in a nondistributive manner on naked DNA and to give rise to a broad clonal distribution of mutations. The mutational spectra study shows that roughly half of lacZ reporter sequences contained between 2 and 20 mutations per clone, with the remaining half of the clones containing greater than 20 and up to as many as 80 mutations (Pham et al., 2003). An examination of individual sparsely mutated clones reveals that deamination of hot-spot C residues could occur a hundred or more nucleotides apart, with intervening hotspot C residues remaining untouched. Perhaps AID, which is probably acting as a multimer, can engage one or more of its subunits to bind distal regions along ssDNA in search of target C residues. In more heavily mutated clones, clusters of closely spaced Cs, including even adjacent C residues, are deaminated, with each clustered region generally containing at least one hotspot sequence. Perhaps these clones demonstrate the potential for each AID subunit to translocate processively over relatively short distances, catalyzing closely spaced deaminations in a clustered region allowing multiple hits by each subunit (Bransteitter et al., 2004).
E. Mutational Models Depicting Two-Phase Somatic Hypermutation The question arises as to how might mutations be generated subsequent to the action of AID, because although AID can target V-gene mutations, it cannot generate the entire spectrum observed in vivo. SHM may best be modeled as two-phase process (Petersen-Mahrt and Neuberger, 2003). The simplest picture describing the first SHM stage would have AID-catalyzed C deamination at WRC followed by faithful replication of U by a high-fidelity polymerase, resulting in a C ! T transition (Fig. 3). Alternatively, a transversion mutation could take place at WRC sites by copying the same U with an error-prone polymerase instead (Fig. 3). The model implies that an interaction between AID and RNA polymerase during transcription enables AID to load and deaminate Cs on the nontranscribed strand while confined within a moving transcription bubble. Recent biochemical data indicate the possibility of a direct interaction between AID and RNA polymerase II and RPA (Chaudhuri et al., 2004; Nambu et al., 2003). Transcription-dependent AID deamination studies performed in E. coli (Ramiro et al., 2003) and in vitro (Pham et al., 2003; Sohail et al., 2003) show that deamination is strongly favored on
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the nontranscribed strand. However, that’s not what happens in vivo, where V-gene mutations have been found to occur in roughly equal numbers on both the nontranscribed and transcribed strands (Milstein et al., 1998), nor can the action of AID alone account for SHM hotspots in WA motifs. V-gene mutations can be generated on both strands and further diversified through a second SHM phase in which AID is involved in initiating the mutational process but not necessarily in targeting the mutations to a specific site on DNA (Fig. 3). Recent evidence indicates that at least two, or perhaps more, polymerases are engaged in SHM. Although there appears to be no change in frequency of SHM in xeroderma pigmentosum patients lacking pol , P. Gearhart and colleagues (Zeng et al., 2001) found a measurable reduction in mutations at A:T base pairs and an increase in mutations at C:G base pairs in V genes obtained from the peripheral blood lymphocytes of XP patients. Furthermore, analysis of switched memory B cells in XP-V patients revealed that pol is also an A/T mutator during CSR, in both the switch region of tandem repeats as well as upstream of it, thus indicating that the same error-prone translesional polymerases might be involved, together with AID, in both SHM and CSR processes (Faili et al., 2004). These data indicate that pol is specifically an A/T mutator. Using error-prone pol to copy a gapped DNA construct containing a lacZ reporter sequence, either alone or aligned in frame with a mouse Ig light chain transgene, Kunkel and colleagues (Pavlov et al., 2002; Rogozin et al., 2001), showed that the base-substitution spectra at A-T pairs correlated with SHM WA hot spot motifs (Rogozin et al., 2001). There was, moreover, good agreement with mouse data for A ! G substitutions (but not C ! T substitutions) when error-prone pol was used to copy a mouse Ig light-chain transgene in vitro (Pavlov et al., 2002). The data also revealed that avidly mutated WA motifs were often situated close to WRC motifs (Pavlov et al., 2002). Based on the close proximity of the two types of hot spot motifs, it is possible that AID-catalyzed deamination of C residues in WRC could therefore provide a mechanism for loading an EP pol to copy nearby WA motifs. Conversion of C ! U by AID, followed by U removal by UNG þ APE enzymes, results in a nick at the 50 -end of the abasic moiety. If an errorprone repair polymerase (e.g., pol ) binds and carries out strand displacement synthesis (long-patch BER), then misincorporation of G opposite T (Fig. 3) will yield A ! G transitions in the nontranscribed strand at WA hotspots, in accordance with pol ’s mutational specificity (Pavlov et al., 2002), as shown (in red) in Fig. 3.
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In support of the EP pol model, genetic studies with knockout mice have revealed that two additional EP polymerases, pol and pol , might participate in SHM. Deficiencies of pol in a Burkitt’s lymphoma cell line dramatically impair the mutation frequency increase on activation of these cells, despite the presence of AID, and it was restored by overexpression of the polymerase (Faili et al., 2002). However, a conflicting report from mice deficient for pol indicates no change in the frequency of SHM (McDonald et al., 2003). The difference between the two studies is yet to be reconciled. Pol can substitute for pol in a BER in vitro reaction (Bebenek et al., 2001), indicating that pol could be involved in generating SHM through error-prone BER repair of G:U mismatches. During short-patch BER, pol might misinsert nucleotides opposite G templates, generating mutations at G/C sites. In addition, substitution for pol , with a relatively weak dRP lyase activity, in place of pol could result in a flap substrate (Fig. 3). Ensuing longer gap-filling synthesis (long patch BER) by error-prone pol copying nearby TW motifs on the transcribed strand should generate mutations at A-T pairs (Fig. 3). Pol may also participate in generating SHM. Knocking out the pol catalytic subunit, REV3, by specific antisense oligonucleotides impairs B-cell SHM in Ig and BCL6 genes (Zan et al., 2001). Similarly, transgenic mice, expressing antisense RNA to a REV3 gene, exhibit decreased frequency of SHM and impaired affinity maturation (Diaz, 2001). A simplified overview of the speculative two-phase mutational model (Fig. 3) is that AID is tacitly assumed to act first by converting C ! U. Then U may be copied accurately or inaccurately by a high-fidelity or low-fidelity polymerase, respectively, leading to mutations on the nontranscribed strand (Fig. 3). A possible scenario to explain the variety of mutations generated during SHM entails competition between the removal of U in DNA by BER or MMR and the possibility of direct replication of U in the DNA by a HiFi or EP pol. Removal of U through BER or MMR, coupled with the action of EP pols, could then result in mutations at different sites on the transcribed strand (Fig. 3). Yet another possibility could be that in contrast to prokaryotic transcription, the transcribed strand might be susceptible to AID-catalyzed C deamination during eukaryotic transcription.
F. Involvement of MMR and BER As illustrated by the model (Fig. 3), postreplicative mismatch repair also appears to play a role in the SHM process at A/T base pairs. Mice deficient in the MutS homologs (Msh) 2 or Msh6 have fewer mutations
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at A/T base-pairs than controls (Phung et al., 1998; Rada et al., 1998; Wiesendanger, 2000). Preferential mutations at G/C base pairs are also seen in Mlh1/ mice (Ehrenstein and Neuberger, 1999; Schrader et al., 1999) but are not as dramatic as those in Msh2/6 knockout mice, indicating that Pms2 or Mlh1 are partially redundant. SHM is also compromised in mice with a ‘‘knockin’’ G674A mutation in the Msh2 gene that inactivates the adenosine triphosphatase domain. This Msh2 mutation does not affect apoptosis signaling and allows mismatches to be recognized, but it prevents Msh2 from initiating mismatch repair (Martin et al., 2003). Thus, the effect of Msh2 on mutations at A/T base pairs is not a reduced B-cell viability resulting from a general loss of genomic stability. The genetic data argue that a UG mismatch initially generated by AID, or a GT mismatch caused by an error-prone polymerase during BER, may sometimes be targeted for repair by the MMR pathway in B cells. In this scenario, MMR would result in C ! T mutations on the transcribed strand as well as on the nontranscribed strand (Fig. 3). Unlike the BER pathway that removes U to cause a 1-base gap, the MMR pathway can form a large gap on either side of the mismatch. If relatively large MMR-generated gaps are equally likely to be formed on either DNA strand, then AID-catalyzed C ! U deaminations at WRC hotspots may occur with similar probabilities on both transcribed and nontranscribed strands, leading to the absence of mutational strand bias, as well as secondary mutations at WA sites nearby, using EP pols (Fig. 3). Not surprising is the finding that MMR deficiencies can also have an effect on CSR. MSH2-deficient mice display a two- to ten-fold reduction in CSR (Ehrenstein and Neuberger, 1999; Schrader et al., 1999), and PMS2or MHL1-deficient mice exhibit two- to four-fold reduction in CSR (Ehrenstein et al., 2001; Schrader et al., 1999). In addition, mice deficient for exonuclease 1, an enzymatic contributor within the MMR pathway, show impairments in both SHM and CSR similar to MSH2 mutant mice (Bardwell, 2004). Although UNG-deficient mice show a strong defect in CSR, neither UNG nor MMR deficiencies completely abolish CSR. The overlapping activity of the BER and MMR repair pathways may compensate for the remaining CSR function observed. The argument that is now being made is that generic DNA repair mechanisms have the potential to interject during CSR (Manis et al., 2002b) and SHM (Neuberger et al., 2003), before completion of the immune-specific pathways, and although it not fundamentally required, MMR, or perhaps BER, appears to function in broadening the final mutational spectra generated by the immune-specific processes.
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G. Regulation of AID Expression in SHM and Its Importance in Cancer Avoidance Transcription of AID is highly regulated during normal B-cell development. In humans and mice, its expression is restricted to centroblast cells within germinal centers, and mRNA production in naı¨ve B-cells, cells that have not yet encountered antigen, occurs predominantly within 48 hours after in vitro stimulation (Greeve et al., 2003; Muramatsu et al., 1999). Recent studies have identified at least three genes directly involved in regulating AID expression: E2A (Sayegh et al., 2003), Pax5, and Id2 (Gonda et al., 2003). E2A and Pax5 are highly expressed in activated B cells and are essential for their differentiation. E2A and Pax5 act by binding to DNA at an E-box or Pax-box located in the regulatory region of AID and activating its transcription (Gonda et al., 2003; Sayegh et al., 2003). The Id2 gene encodes an inhibitor protein that negatively regulates AID expression by interacting with E2A gene products, abolishing their E-box binding activity. Examination of the putative regulatory region of AID showed that the Pax5-binding site is absolutely required for AID gene expression (Gonda et al., 2003). Although antibody diversification benefits from AID deamination action, its inappropriate expression can induce a significant increase in genomic instability, which may lead to cancer. Not only are AID transcripts and their splice variants readily detectable in B-cell non-Hodgkin lymphomas (Greeve et al., 2003) and B-cell chronic lymphocytic leukemia (Albesiano et al., 2003; McCarthy et al., 2003), but constitutive expression of AID in transgenic mice also leads to the development of malignant Tcell lymphoma, micro-adenomas, and adenocarcinomas in the lungs (Okazaki et al., 2003). In addition, AID transgenic mice exhibit substantial increases in point mutations, but not translocations, in expressed T cell receptors and c-myc genes found in T cell lymphomas. Mutational distribution and specificity appear to be similar to those observed in B-cell lymphoma (Pasqualucci et al., 2001) and AID-overexpressing cells in vitro (Martin and Scharff, 2002; Martin et al., 2002; Yoshikawa et al., 2002), indicating that AID-induced mutagenesis is responsible for tumorigenesis. Similar to AID, ectopic expression of its homolog, Apobec-1, has been also shown to promote liver cancer in transgenic mice (Yamanaka et al., 1995).
III. Apobec Protein Family AID is homologous to the mRNA editing enzyme Apobec-1 (Table I), sharing 34% amino acid identity (Muramatsu et al., 1999; Navaratnam et al., 1993). Analyses of human genomic and EST data have revealed several proteins with sequence homology to zinc-dependent deaminase domain
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Table I Human Apobec Protein Family of Nucleic Acid Deaminasesa Gene
Location
Tissue expression
Function
RNA deaminase APOBEC-1
12p13.1
Small and large intestine
mRNA apoB editing
DNA deaminase AID APOBEC-3G
12p13 22q13.1
B lymphocytes Spleen, breast, heart, thymus, colon, stomach, kidney, uterus, pancreas, placenta, prostate
SHM and CSR Antiretroviruses
Unknown APOBEC-2 APOBEC-3A APOBEC-3B APOBEC-3C
6p21 22q13.1 22q13.1 22q13.1
Apobec-1 inhibitor? ? ? ?
APOBEC-3D APOBEC-3E APOBEC-3F XP_092919 XP_115170
22q13.1 22q13.1 22q13.1 22q13.1 12q23
Cardiac and skeletal muscle Keratinocytes Keratinocytes, colon Spleen, testes, heart, thymus, Prostate, ovary, uterus Head and neck cancers ? B lymphocytes ? ?
? Pseudogene Antiretroviruses? ? ?
a
Adapted from Wedekind et al., 2003.
of Apobec-1 and AID (Anant et al., 2001; Jarmuz et al., 2002; Madsen et al., 1999). A cluster of apobec-related proteins, Apobec3A to Apobec3G, and an expressed gene (XM_092919) have been found on chromosome 22. In addition, another gene (XP_115170), closely related to Apobec3G, is located at position 12q23. These proteins have distinct expression profiles and their functions remain largely unknown (Table I). Domain structures of more than half of the members of the apobec family, such as Apobec-1 and AID, are characterized by the presence of the catalytic domain (CD) and a pseudocatalytic domain (PCD) and separated by a linker region (Fig. 4). However, other members like Apobec3 variants B, F, G, and its mouse homolog CEM15, have CD-PCD-CD-PCD domain structure and have probably evolved through gene duplication and divergence. On the basis of biochemical properties, the members of this family can be classified into two subclasses: RNA deaminases (APOBEC-1) and DNA deaminases (AID and APOBEC3G) (Table I).
A. Apobec-1 Apobec-1, a founding member of this family, is responsible for C ! U editing in apolipoproteinB mRNA at nucleotides 6666 and 6802, changing
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Fig. 4. Domain structure of AID. AID is composed of a catalytic domain (CD) containing a Zn+ binding consensus sequence and a pseudocatalytic domain (PCD), separated by a linker region. The N-terminal region of AID is characterized by the enrichment of positively charged lysine and arginine residues, possibly responsible for its tight binding to nucleic acids. The C-terminal region of AID has been identified to be essential for CSR. Amino acid changes identified in patients with impaired SHM and CSR are indicated at the bottom.
a glutamine codon CAA to a stop codon UAA and a threonine codon to an isoleucine, respectively (Teng et al., 1993). In addition, Apobec-1 may also be responsible for mRNA editing of a tumor suppressor gene, neurofibromin, in approximately a quarter of neurofibromatosis Type 1 patients (Mukhopadhyay et al., 2002; Skuse et al., 1996). Site-specific C ! U conversion by Apobec-1 requires its dimerization as well as an interaction with at least one of two splice variants of ACF protein (ACF65 and ACF64). The ACF proteins bind with high affinity to an AU-rich RNA known as the mooring sequence located 30 of the edited cytidine and together with Apobec-1 form a minimal editing complex (Dance et al., 2002; Mehta and Driscoll, 2002). A number of other proteins have also been shown to interact with Apobec-1 modulating its apoB mRNA editing activity in vitro, but their involvement in vivo as auxiliary factors remains to be validated (Wedekind et al., 2003). A homolog of Apobec-1, Apobec-2, can dimerize with Apobec-1 and has been shown to inhibit its C ! U editing activity (Anant et al., 2001; Liao et al., 1999). Although Apobec-1 is capable of deaminating dC residues in vitro (Harris et al., 2003) and its overexpression in E. coli induces C ! T transitions (Harris et al., 2002), there is no evidence that Apobec-1 can deaminate DNA in human cells.
B. Antiretroviral Activity of Apobec3G, G ! A Hypermutation in HIV and Other Retroviruses One enigmatic feature of genetic variation of HIV and other retrovirus genomes was G ! A hypermutation, characterized by high level of G ! A base substitutions in the positive (mRNA) strand (Vartanian et al., 1991). G ! A hypermutation can occur throughout the HIV genome with more
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than a third of the total 2189 G residues being mutated. In some segments, G ! A mutations are observed in up to 60% of G templates (Vartanian et al., 2002). G ! A hypermutation is only seen during reverse transcription, in which the retroviral RNA genome is copied into DNA in host cells. A major clue to how G ! A hypermutation might happen came from the discovery of the potent antiviral activity of an apobec protein family member, Apobec3G, and a novel immune defense mechanism through DNA dC deamination (Harris et al., 2003; Mangeat et al., 2003; Sheehy et al., 2002; Zhang et al., 2003). Similar to AID, Apobec3G was discovered in a substractive mRNA screen for genes that specifically inhibit the replication of Vif-minus HIV virus (HIV-1 strain) in nonpermissive cell lines (Sheehy et al., 2002). Vif, virus infectivity factor, is a protein required by HIV-1 to replicate in certain nonpermissive cell lines, such as CEM15. It is now clear that Apobec3G targets HIV by altering its genome through production of massive numbers of G!A mutations (Harris et al., 2003; Mangeat et al., 2003; Zhang et al., 2003). Apobec3G is incorporated into the Vif-minus HIV particles, and upon their infection of naı¨ve T cells, the enzyme is released and works a C deaminase on the first (minus) strand of DNA during reverse transcription. As a consequene, about 1%–2% of all C residues in the minus strand are deaminated to uracils. The mutations are preferentially observed at third C residue in a 50 -YCC sequence. Besides HIV, Apobec3G also acts on other retrovirus genomes such as SIV, EIAV and MuLV (Harris et al., 2003; Mangeat et al., 2003). Recombinant Apobec3G, purified from E. coli, has been shown to deaminate dC on ssDNA in vitro (Harris et al., 2003). In addition, the enzyme, purified from baculovirus-infected insect cells, exhibits a preferential specificity for CC hotspots similar to what is seen in vivo, indicating that the enzyme itself is solely responsible for the observed in vivo activity (P. Pham, R. Bransteitter and M. F. Goodman, unpublished data). The presence of a massive number of uracil residues in the minus strand of DNA could either lead to its degradation by combined action of UNG and AP endonuclease, or to the death of HIV virus by ‘‘error catastrophe,’’ whereby hypermutation in essential genes no longer encodes functional viral proteins.
IV. Biochemical Perspective The ability of higher organisms to mount an effective biodefense to counter exposure to potentially life-threatening antigenic assault is central to individual and species survival. By generating a diverse set of antibodies, the immune response is generally capable of providing ample protection. The conversion of low-affinity to high-affinity antibodies occurring by
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somatic hypermutation and class-switch recombination allows antibodies to carry out many effector functions and be distributed throughout the body and secretions. Both processes are initiated by AID, an enzyme that catalyzes transcription-dependent deamination of C residues. The recent discoveries that the substrate for AID is single-stranded DNA and that AID acting on ssDNA simulates SHM spectra, in the absence of additional cofactors, will undoubtedly facilitate a deeper understanding into the molecular basis of CSR and SHM. This chapter has focused on the biochemical role of AID during SHM, a ‘‘surface’’ that has just barely been ‘‘scratched.’’ There is a clear path ahead to address AID sequence specificity mechanisms, using relatively straightforward measurements of AID binding affinities to ssDNA, dsDNA, and AID catalytic rates in C hotspot and coldspot sequences. AID acts nondistributively on ssDNA, and there are methods available for measuring enzyme processivity and scanning mechanisms in ‘‘simple’’ circumstances. However, all bets are off regarding the ease of analyzing processivity, because AID is believed to be composed of a multimer of identical subunits, and it is not known how many subunits can bind to and act on DNA at any one time. AID has been shown to act in a transcription-dependent manner using a model T7 RNA polymerase assay. AID has also been found to deaminate C residues more aggressively using a ‘‘transcription-like’’ bubble compared to ssDNA. The next step will be to decipher how the mutational C-targeting AID is itself targeted to active transcription complexes. Here it will be necessary to establish a eukaryotic transcription system to investigate AID interactions with RNA polymerase II, along with its numerous transcription factors. Somewhat further off is the search for the SHM ‘‘Holy Grail’’—a stem-to-stern reconstitution of SHM, incorporating MMR, BER and error-prone DNA polymerases acting on chromatin.
Acknowledgments This work was supported by grants from the National Institutes of Health, R37GM21422 and RO1ES012259. We thank Dr. Matthew D. Scharff for his patient and generous tutelage and for reading and commenting on this chapter
References Albesiano, E., Messmer, B. T., Damle, R. N., Allen, S. L., Rai, K. R., and Chiorazzi, N. (2003). Activation-induced cytidine deaminase in chronic lymphocytic leukemia B cells: Expression as multiple forms in a dynamic, variably sized fraction of the clone. Blood 102, 3333–3339. Anant, S., Mukhopadhyay, D., Sankaranand, V., Kennedy, S., Henderson, J. O., and Davidson, N. O. (2001). ARCD-1, an apobec-1-related cytidine deaminase, exerts a
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dominant negative effect on C to U RNA editing. Am. J. Phys. Cell Physiol. 281, C1904–C1916. Arakawa, H., Hauschild, J., and Buerstedde, J. M. (2002). Requirement of the activation-induced deaminase (AID) gene for immunoglobulin gene conversion. Science 295, 1301–1306. Bachl, J., Carlson, C., Gray-Schopfer, V., Dessing, M., and Olsson, C. (2001). Increased transcription levels induce higher mutation rates in a hypermutating cell line. J. Immunol. 166, 5051–5057. Bardwell, P. D., Martin, A., Wong, E., Li, Z., Edelmann, W., and Scharff, M. D. (2003). Cutting edge: The G-U mismatch glycosylase methyl-CpG binding domain 4 is dispensable for somatic hypermutation and class switch recombination. J. Immunol. 170, 1620–1624. Bardwell, P. D., Woo, C., Wei, K., Ziqiang, L., Martin, A., Stephen, S. Z., Tchaiko, P., Winfried, E., and Scharff, M. D. (2004). Altered somatic hypermutation and reduced class-switch recombination in exonuclease 1-mutant mice. Nat. Immunol. 5, 224–229. Barreto, V., Reina-San-Martin, B., Ramiro, A. R., McBride, K. M., and Nussenzweig, M. C. (2003). C-terminal deletion of AID uncouples class switch recombination from somatic hypermutation and gene conversion. Mol. Cell 12, 501–508. Bebenek, K., Tissier, A., Frank, E. G., McDonald, J. P., Prasad, R., Wilson, S. H., Woodgate, R., and Kunkel, T. A. (2001). 50 -Deoxyribose phosphate lyase activity of human DNA polymerase iota in vitro. Science 291, 2156–2159. Bertocci, B., De Smet, A., Berek, C., Weill, J., and Reynaud, C. (2003). Immunoglobulin kappa light chain gene rearrangement is impaired in mice deficient for DNA polymerase mu. Immunity 2, 203–211. Blunt, T., Finnie, N., Taccioli, G., Smith, G., Demengeot, J., Gottlieb, T., Mizuta, R., Varghese, A., Alt, F., and Jeggo, P. (1995). Defective DNA-dependent protein kinase activity is linked to V(D)J recombination and DNA repair defects associated with the murine scid mutation. Cell 80, 813–823. Bosma, G., Kim, J., Urich, T., Fath, D., Cotticelli, M., Ruetsch, N., Radic, M., and Bosma, M. (2002). DNA-dependent protein kinase activity is not required for immunoglobulin class switching. J. Exp. Med. 196, 1483–1495. Bottaro, A., Lansford, R., Xu, L., Zhang, J., Rothman, P., and Alt, F. (1994). S region transcription per se promotes basal IgE class switch recombination but additional factors regulate the efficiency of the process. EMBO J. 13, 665–674. Bottaro, A., Young, F., Chen, J., Serwe, M., Sablitzky, F., and Alt, F. (1998). Deletion of the IgH intronic enhancer and associated matrix-attachment regions decreases, but does not abolish, class switching at the mu locus. Int. Immunol. 6, 799–806. Bransteitter, R., Pham, P., Calabrese, P., and Goodman, M. F. (2004). Biochemical analysis of hyper-mutational targeting by wild type and mutant AID. J. Biol. Chem. in press. Bransteitter, R., Pham, P., Scharff, M. D., and Goodman, M. F. (2003). Activationinduced cytidine deaminase deaminates deoxycytidine on single-stranded DNA but requires the action of RNase. Proc. Natl. Acad. Sci. USA 100, 4102–4107. Casellas, R., Nussenzweig, A., Wuerffel, R. A., Pelanda, R., Reichlin, A., Suh, H., Qin, X. F., Besmer, E., Kenter, A., Rajewsky, K., and Nussenzweig, M. C. (1998). Ku80 is required for immunoglobulin isotype switching. EMBO J. 17, 2404–2411. Chaudhuri, J., Khuong, C., and Alt, F. W. (2004). Replication protein A interacts with AID to promote deamination of somatic hypermutation targets. Nature 430, 992–998. Chaudhuri, J., Tian, M., Khoung, C., Chua, K., Pinaud, E., and Alt, F. W. (2003). Transcription-targeted DNA deamination by the AID antibody diversification enzyme. Nature 421, 726–730.
SOMATIC HYPERMUTATION
329
Dance, G. S., Sowden, M. P., Cartegni, L., Cooper, E., Krainer, A. R., and Smith, H. C. (2002). Two proteins essential for apolipoprotein B mRNA editing are expressed from a single gene through alternative splicing. J. Biol. Chem. 277, 12703–12709. Di Noia, J., and Neuberger, M. S. (2002). Altering the pathway of immunoglobulin hypermutation by inhibiting uracil-DNA glycosylase. Nature 419, 43–48. Diaz, M., Verkoczy, L. K., Flajnik, M. F., and Klinman, N. R. (2001). Decreased frequency of somatic hypermutation and impaired affinity maturation but intact germinal center formation in mice expressing antisense RNA to DNA polymerase zeta. J. Immunol. 167, 327–335. Dickerson, S. K., Market, E., Besmer, E., and Papavasiliou, F. N. (2003). AID mediates hypermutation by deaminating single stranded DNA. J. Exp. Med. 197, 1291–1296. Ehrenstein, M. R., and Neuberger, M. S. (1999). Deficiency in msh2 affects the efficiency and local sequence specificity of immunoglobulin class-switch recombination: parallels with somatic hypermutation. EMBO J. 18, 3484–3490. Ehrenstein, M. R., Rada, C., Jones, A. M., Milstein, C., and Neuberger, M. S. (2001). Switch junction sequences in PMS2-deficient mice reveal a microhomologymediated mechanism of Ig class switch recombination. Proc. Natl. Acad. Sci. USA 98, 14553–14558. Esposito, G., Texido, G., Betz, U. A., Gu, H., Muller, W., Klein, U., and Rajewsky, K. (2000). Mice reconstituted with DNA polymerase beta-deficient fetal liver cells are able to mount a T cell-dependent immune response and mutate their Ig genes normally. Proc. Natl. Acad. Sci. USA 97, 1166–1171. Faili, A., Aoufouchi, S., Flatter, E., Gueranger, Q., Reynaud, C. A., and Weill, J. C. (2002). Induction of somatic hypermutation in immunoglobulin genes is dependent on DNA polymerase iota. Nature 419, 944–947. Faili, A., Aoufouchi, S., Weller, S., Vuillier, F., Stary, A., Sarasin, A., Reynaud, C., and Weill, J. (2004). DNA polymerase eta is involved in hypermutation occurring during immunoglobulin class switch recombination. J. Exp. Med. 199, 265–270. Flajnik, M. (2002). Comparative analysis of immunoglobulin genes: Suprises and portents. Nat. Rev. Immunol. 2, 688–698. Gao, Y., Sun, Y., Frank, K., Dikkes, P., Fujiwara, Y., Seidl, K., Sekiguchi, J., Rathbun, G., Swat, W., Wang, J., Bronson, R., Malynn, B., Bryans, M., Zhu, C., Chaudhuri, J., Davidson, L., Ferrini, R., Stamato, T., Orkin, S., Greenberg, M., and Alt, F. (1998). A critical role for DNA end-joining proteins in both lymphogenesis and neurogenesis. Cell 95, 891–902. Gellert, M. (2002). V(D)J recombination: RAG proteins, repair factors, and regulation. Annu. Rev. Biochem. 71, 101–132. Gilfillan, S., Dierich, A., Lemeur, M., Benoist, C., and Mathis, D. (1993). Mice lacking TdT: Mature animals with an immature lymphocyte repertoire. Science 261, 1175–1178. Golding, G. B., Gearhart, P. J., and Glickman, B. W. (1987). Patterns of somatic mutations in immunoglobulin variable genes. Genetics 115, 169–176. Gonda, H., Sugai, M., Nambu, Y., Katakai, T., Agata, Y., Mori, K. J., Yokota, Y., and Shimizu, A. (2003). The balance between Pax5 and Id2 activities is the key to AID gene expression. J. Exp. Med. 198, 1427–1437. Grawunder, U., Zimmer, D., Fugmann, S., Scharz, K., and Lieber, M. (1998). DNA ligase IV is essential for V(D)J recombination and DNA double-strand break repair in human precursor lymphocytes. Mol. Cell 2, 477–484.
330
TIPPIN ET AL.
Greeve, J., Philipsen, A., Krause, K., Klapper, W., Heidorn, K., Castle, B. E., Janda, J., Marcu, K. B., and Parwaresch, R. (2003). Expression of activation-induced cytidine deaminase in human B-cell non-Hodgkin lymphomas. Blood 101, 3574–3580. Harriman, G., Bradley, A., Das, S., Rogers-Fani, P., and Davis, A. (1996). IgA class switch in I alpha exon-deficient mice. Role of germline transcription in class switch recombination. J. Clin. Invest. 97, 477–485. Harris, R. S., Petersen-Mahrt, S. K., and Neuberger, M. S. (2002). RNA editing enzyme APOBEC1 and some of its homologs can act as DNA mutators. Mol. Cell 10, 1247–1253. Harris, R. S., Sale, J. E., Petersen-Mahrt, S. K., and Neuberger, M. S. (2002). AID is essential for immunoglobulin V gene conversion in a cultured B cell line. Curr. Biol. 12, 435–438. Harris, R. S., Sheehy, A. M., Craig, H. M., Malim, M. H., and Neuberger, M. S. (2003). DNA deamination: Not just a trigger for antibody diversification but also a mechanism for defense against retroviruses. Nat. Immunol. 4, 641–643. Honjo, T., Kinoshita, K., and Muramatsu, M. (2002). Molecular mechanism of class switch recombination: Linkage with somatic hypermutation. Annu. Rev. Immunol. 20, 165–196. Imai, K., Catalan, N., Plebani, A., Marodi, L., Sanal, O., Kumaki, S., Nagendran, V., Wood, P., Glastre, C., Sarrot-Reynauld, F., Hermine, O., Forveille, M., Revy, P., Fischer, A., and Durandy, A. (2003a). Hyper-IgM syndrome type 4 with a B lymphocyte-intrinsic selective deficiency in Ig class-switch recombination. [see comment]. J. Clin. Invest. 112, 136–142. Imai, K., Slupphaug, G., Lee, W. I., Revy, P., Nonoyama, S., Catalan, N., Yel, L., Forveille, M., Kavli, B., Krokan, H. E., Ochs, H. D., Fischer, A., and Durandy, A. (2003b). Human uracil-DNA glycosylase deficiency associated with profoundly impaired immunoglobulin class-switch recombination. Nat. Immunol. 4, 1023–1028. Jacobs, H., Fukita, Y., van der Horst, G. T. J., de Boer, J., Weeda, G., Essers, J., de Wind, N., Engelward, B. P., Samson, L., Verbeek, S., Menissier-de Murcia, J., de Murcia, G., te Riele, H., and Rajewsky, K. (1998). Hypermutation of immunoglobulin genes in memory B cells of DNA repair-deficient mice. J. Exp. Med. 187, 1735–1743. Jarmuz, A., Chester, A., Bayliss, J., Gisbourne, J., Dunham, I., Scott, J., and Navaratnam, N. (2002). An anthropoid-specific locus of orphan C to U RNA-editing enzymes on chromosome 22. Genomics 79, 285–296. Jung, S. (1993). Shutdown of class-switch recombination by deletion of a switch-region control element. Science 259, 984–987. Kasahara, Y., Kaneko, H., Fukao, T., Terada, T., Asano, T., Kasahara, K., and Kondo, N. (2003). Hyper-IgM syndrome with putative dominant negative mutation in activation-induced cytidine deaminase. J. Allergy Clin. Immunol. 112, 755–760. Khamlichi, A., Pinaud, E., Decourt, C., Chauveau, C., and Cogne, M. (2000). The 30 IgH regulatory region: A complex structure in a search for a function. Adv. Immunol. 75, 317–345. Kinoshita, K., Lee, C., Tashiro, J., Muramatsu, M., Chen, X., Yoshikawa, K., and Honjo, T. (1999). Molecular mechanism of immunoglobulin class switch recombination. Cold Spring Harb. Symp. Quant. Biol. 64, 217–226. Kirchgessner, C., Patil, C., Evans, J., Cuomo, C., Fried, L., Carter, T., Oettinger, M., and Brown, J. (1995). DNA-dependent kinase (p350) as a candidate gene for the murine SCID defect. Science 267, 1178–1183.
SOMATIC HYPERMUTATION
331
Kohler, G., and Milstein, C. (1975). Continuous cultures of fused cells secreting antibody of predefined specificity. Nature 256, 495–497. Komori, T., Okada, A., Stewart, V., and Alt, F. (1993). Lack of N regions in antigen receptor variable region genes of TdT-deficient lymphocytes. Science 261, 1171–1175. Krangel, M. (2003). Gene segment selection in V(D)J recombination: Accessibility and beyond. Nat. Immunol. 4, 624–630. Lebecque, S. G., and Gearhart, P. J. (1990). Boundaries of somatic mutation in rearranged immunoglobulin genes: 50 boundary is near the promoter, and 30 boundary is approximately 1 kb from V(D)J gene. J. Exp. Med. 172, 1717–1727. Lee, C., Kinoshita, K., Arudchandran, A., Cerritelli, S., Crouch, R., and Honjo, T. (2001). Quantitative regulation of class switch recombination by switch region transcription. J. Exp. Med. 194, 365–374. Li, Y., Hayakawa, K., and Hardy, R. (1993). The regulated expression of B lineage associated genes during B cell differentiation in bone marrow and fetal liver. J. Exp. Med. 178, 951–960. Li, Z., Otevrel, T., Gao, Y., Cheng, H., and Seed, B. (1995). The XRCC4 gene encodes a novel protein involved in DNA double-strand break repair and V(D)J recombination. Cell 83, 1079–1089. Liao, W., Hong, S. H., Chan, B. H., Rudolph, F. B., Clark, S. C., and Chan, L. (1999). APOBEC-2, a cardiac-and skeletal muscle-specific member of the cytidine deaminase supergene family. Biochem. Biophys. Res. Comm. 260, 398–404. Lieber, M., Ma, Y., Pannicke, U., and Schwarz, K. (2003). Mechanism and regulation of human non-homologous DNA end-joining. Nat. Rev. Mol. Cell. Biol. 4, 712–720. Lindahl, T., and Barnes, D. E. (2001). Repair of endogenous DNA damage. Cold Spring Harb. Symp. Quant. Biol. 65, 127–133. Ma, Y., Pannicke, U., Schwarz, K., and Lieber, M. (2002). Hairpin opening and overhang processing by an Artemis/DNA-dependent protein kinase complex in nonhomologous end joining and V(D)J recombination. Cell 108, 781–794. Madsen, P., Anant, S., Rasmussen, H. H., Gromov, P., Vorum, H., Dumanski, J. P., Tommerup, N., Collins, J. E., Wright, C. L., Dunham, I., MacGinnitie, A. J., Davidson, N. O., and Celis, J. E. (1999). Psoriasis upregulated phorbolin-1 shares structural but not functional similarity to the mRNA-editing protein apobec-1. J. Invest. Dermatol. 113, 162–169. Mahajan, K., Nick McElhinny, S., Mitchell, B., and Ramsden, D. (2002). Association of DNA polymerase mu (pol mu) with Ku and ligase IV: Role for pol mu in endjoining double-strand break repair. Mol. Cell. Biol. 14, 5194–5202. Maizels, N. (1995). Somatic hypermutation: How many mechanisms diversify V region sequences? Cell 83, 9–12. Mangeat, B., Turelli, P., Caron, G., Friedli, M., Perrin, L., and Trono, D. (2003). Broad antiretroviral defence by human APOBEC3G through lethal editing of nascent reverse transcripts. Nature 424, 99–103. Manis, J. P., Dudley, D., Kaylor, L., and Alt, F. W. (2002a). IgH class switch recombination to IgG1 in DNA-PKcs-deficient B cells. Immunity 16, 607–617. Manis, J. P., Gu, Y., Lansford, R., Sonada, E., Ferrini, R., Davidson, L., Rajewsky, K., and Alt, F. W. (1998). Ku70 is required for late B cell development and immunoglobulin heavy chain class switching. J. Exp. Med. 187, 2081–2089. Manis, J. P., Tian, M., and Alt, F. W. (2002b). Mechanism and control of class-switch recombination. Trends Immunol. 23, 31–39.
332
TIPPIN ET AL.
Martin, A., Bardwell, P. D., Woo, C. J., Fan, M., Shulman, M. J., and Scharff, M. D. (2002). Activation-induced cytidine deaminase turns on somatic hypermutation in hybridomas. Nature 415, 802–806. Martin, A., Li, Z., Lin, D., Bardwell, P., Iglesias-Ussel, M., Edelmann, W., and Scharff, M. (2003). Msh2 ATPase activity is essential for somatic hypermutation at a-T basepairs and for efficient class switch recombination. J. Exp. Med. 198, 1171–1178. Martin, A., and Scharff, M. D. (2002). Somatic hypermutation of the AID transgene in B and non-B cells. Proc. Natl. Acad. Sci. USA 99, 12304–12308. McCarthy, J., Wierda, W. G., Barron, L. L., Cromwell, C. C., Wang, J., Coombes, K. R., Rangel, R., Elenitoba-Johnson, K. S., Keating, M. J., and Abruzzo, L. V. (2003). High expression of activation-induced cytidine deaminase (AID) and splice variants is a distinctive feature of poor-prognosis chronic lymphocytic leukemia. Blood 101, 4903–4908. McDonald, J. P., Frank, E. G., Plosky, B. S., Rogozin, I. B., Masutani, C., Hanaoka, F., Woodgate, R., and Gearhart, P. J. (2003). 129-derived strains of mice are deficient in DNA polymerase iota and have normal immunoglobulin hypermutation. J. Exp. Med. 198, 635–643. Mehta, A., and Driscoll, D. M. (2002). Identification of domains in apobec-1 complementation factor required for RNA binding and apolipoprotein-B mRNA editing. RNA 8, 69–82. Milstein, C., Neuberger, M. S., and Staden, R. (1998). Both DNA strands of antibody genes are hypermutation targets. Proc. Natl. Acad. Sci. USA 95, 8791–8794. Mombaerts, P., Iacomini, J., Johnson, R., Herrup, K., Tonegawa, S., and Papaioannou, V. (1992). RAG-1-deficient mice have no mature B and T lymphocytes. Cell 68, 869–877. Moshous, D., Callebaut, I., de Chasseval, R., Corneo, B., Cavazzana-Calvo, M., Le Deist, F., Tezcan, I., Sanal, O., Bertrand, Y., Philippe, N., Fischer, A., and de Villartay, J. (2001). Artemis, a novel DNA double-strand break repair/V(D)J recombination protein, is mutated in human severe combined immune deficiency. Cell 105, 177–186. Mukhopadhyay, D., Anant, S., Lee, R. M., Kennedy, S., Viskochil, D., and Davidson, N. O. (2002). C!U editing of neurofibromatosis 1 mRNA occurs in tumors that express both the type II transcript and apobec-1, the catalytic subunit of the apolipoprotein B mRNA-editing enzyme. Am. J. Hum. Genet. 70, 38–50. Muramatsu, M., Kinoshita, K., Fagarasan, S., Yamada, S., Shinkai, Y., and Honjo, T. (2000). Class switch recombination and hypermutation require activation-induced cytidine deaminase (AID), a potential RNA editing enzyme. Cell 102, 553–563. Muramatsu, M., Sankaranand, V. S., Anant, S., Sugai, M., Kinoshita, K., Davidson, N. O., and Honjo, T. (1999). Specific expression of activation-induced cytidine deaminase (AID), a novel member of the RNA-editing deaminase family in germinal center B cells. J. Biol. Chem. 274, 18470–18476. Nambu, Y., Sugai, M., Gonda, H., Lee, C., Katakai, T., Agata, Y., Yokota, Y., and Shimizu, A. (2003). Transcription-coupled events associating with immunoglobulin switch region chromatin. Science 302, 2137–2140. Navaratnam, N., Morrison, J. R., Bhattacharya, S., Patel, D., Funahashi, T., Giannoni, F., Teng, B. B., Davidson, N. O., and Scott, J. (1993). The p27 catalytic subunit of the apolipoprotein B mRNA editing enzyme is a cytidine deaminase. J. Biol. Chem. 268, 20709–20712. Neuberger, M. S., Harris, R. S., Di Noia, J., and Petersen-Mahrt, S. K. (2003). Immunity through DNA deamination. TiBS 28, 305–312.
SOMATIC HYPERMUTATION
333
Notarangelo, L. D., Due, M., and Ugazio, A. G. (1992). Immunodeficiency with hyperIgM (HIM). Immunodefic. Rev. 3, 101–122. Oettinger, M., Schatz, D., Gorka, C., and Baltimore, D. (1990). RAG -1 and RAG -2, adjacent genes that synergistically activate V(D)J recombination. Science 248, 1517–1523. Okazaki, I. M., Hiai, H., Kakazu, N., Yamada, S., Muramatsu, M., Kinoshita, K., and Honjo, T. (2003). Constitutive expression of AID leads to tumorigenesis. J. Exp. Med. 197, 1173–1181. Okazaki, I. M., Kinoshita, K., Muramatsu, M., Yoshikawa, K., and Honjo, T. (2002). The AID enzyme induces class switch recombination in fibroblasts. Nature 416, 340–345. Oppezzo, P., Vuillier, F., Vasconcelos, Y., Dumas, G., Magnac, C., Payelle-Brogard, B., Pritsch, O., and Dighiero, G. (2003). Chronic lymphocytic leukemia B cells expressing AID display dissociation between class switch recombination and somatic hypermutation. Blood 101, 4029–4032. Papavasiliou, F. N., and Schatz, D. G. (2002). The activation-induced deaminase functions in a postcleavage step of the somatic hypermutation process. J. Exp. Med. 195, 1193–1198. Pasqualucci, L., Neumeister, P., Goossens, T., Nanjangud, G., Chaganti, R. S., Kuppers, R., and Dalla-Favera, R. (2001). Hypermutation of multiple proto-oncogenes in Bcell diffuse large-cell lymphomas. Nature 412, 341–346. Pavlov, Y. I., Rogozin, I. B., Galkin, A. P., Aksenova, A. Y., Hanaoka, F., Rada, C., and Kunkel, T. A. (2002). Correlation of somatic hypermutation specificity and A-T base pair substitution errors by DNA polymerase during copying of a mouse immunoglobulin light chain transgene. Proc. Natl. Acad. Sci. USA 99, 9954–9959. Peters, A., and Storb, U. (1996). Somatic hypermutation of immunoglobulin genes is linked to transcription initiation. Immunity 4, 57–65. Petersen-Mahrt, S. K., Harris, R. S., and Neuberger, M. S. (2002). AID mutates E. coli suggesting a DNA deamination mechanism for antibody diversification. Nature 418, 99–103. Petersen-Mahrt, S. K., and Neuberger, M. S. (2003). In vitro deamination of cytosine to uracil in single-stranded DNA by apolipoprotein B editing complex catalytic subunit 1 (APOBEC1). J. Biol. Chem. 278, 19583–19586. Peterson, S., Kurimasa, A., Oshimura, M., Dynan, W., Bradbury, E., and Chen, D. (1995). Loss of the catalytic subunit of the DNA-dependent protein kinase in DNA double-strand-break-repair mutant mammalian cells. Proc. Natl. Acad. Sci. USA 92, 3171–3174. Pham, P., Bransteitter, R., Petruska, J., and Goodman, M. F. (2003). Processive AIDcatalyzed cytosine deamination on ssDNA simulates somatic hypermutation. Nature 423, 103–107. Phung, Q., Winter, D., Cranston, A., Tarone, R., Bohr, V., Fishel, R., and Gearhart, P. (1998). Increased hypermutation ot G and C nucleotides in immunoglobulin varia ble genes from mice deficient in the MSH2 mismatch repair protein. J. Exp. Med. 187, 1745–1751. Rada, C., Ehrenstein, M. R., Neuberger, M. S., and Milstein, C. (1998). Hot spot focusing of somatic hypermutation in MSH2-deficient mice suggests two stages of mutational targeting. Immunity 9, 135–141. Rada, C., Williams, G. T., Nilsen, H., Barnes, D. E., Lindahl, T., and Neuberger, M. S. (2002). Immunoglobulin isotype switching is inhibited and somatic hypermutation perturbed in UNG-deficient mice. Curr. Biol. 12, 1748–1755.
334
TIPPIN ET AL.
Ramiro, A. R., Stavropoulos, P., Jankovic, M., and Nussenzweig, M. C. (2003). Transcription enhances AID-mediated cytidine deamination by exposing single-stranded DNA on the nontemplate strand. Nat. Immunol. 4, 452–456. Revy, P., Muto, T., Levy, Y., Geissmann, F., Plebani, A., Sanal, O., Catalan, N., Forveille, M., Dufourcq-Labelouse, R., Gennery, A., Tezcan, I., Ersoy, F., Kayserili, H., Ugazio, A. G., Brousse, N., Muramatsu, M., Notarangelo, L. D., Kinoshita, K., Honjo, T., Fischer, A., and Durandy, A. (2000). Activation-induced cytidine deaminase (AID) deficiency causes the autosomal recessive form of the Hyper-IgM syndrome (HIGM2). Cell 102, 565–575. Rogozin, I. B., and Kolchanov, N. A. (1992). Somatic hypermutagenesis in immunoglobulin genes. II. Influence of neighbouring base sequences on mutagenesis. Biochim. Biophy. Acta 1171, 11–18. Rogozin, I. B., Pavlov, Y. I., Bebenek, K., Matsuda, T., and Kunkel, T. A. (2001). Somatic mutation hotspots correlate with DNA polymerase eta error spectrum. Nat. Immunol. 2, 530–536. Rolink, A., Melchers, F., and Andersson, J. (1996). The SCID but not the RAG-2 gene product is required for S mu-S epsilon heavy chain class switching. Immunity 4, 319–330. Sale, J. E., Calandrini, D. M., Takata, M., Takeda, S., and Neuberger, M. S. (2001). Ablation of XRCC2/3 transforms immunoglobulin V gene conversion into somatic hypermutation. Nature 412, 921–924. Sawchuk, D., Weis-Garcia, F., Malik, S., Besmer, E., Bustin, M., Nussenzweig, M., and Cortes, P. (1997). V(D)J recombination: Modulation of RAG1 and RAG2 cleavage activity on 12/23 substrates by whole cell extract and DNA bending proteins. J. Exp. Med. 185, 2025–2032. Sayegh, C. E., Quong, M. W., Agata, Y., and Murre, C. (2003). E-proteins directly regulate expression of activation-induced deaminase in mature B cells. Nat. Immunol. 4, 586–593. Schatz, D. G., Oettinger, M. A., and Baltimore, D. (1989). The V(D)J recombination activating gene, Rag-1. Cell 59, 1035–1048. Schrader, C. E., Edelmann, W., Kucherlapati, R., and Stavnezer, J. (1999). Reduced isotype switching in splenic B cells from mice deficient in mismatch repair enzymes. J. Exp. Med. 190, 323–330. Seidl, K., Bottaro, A., Vo, A., Zhang, J., Davidson, L., and Alt, F. (1998). An expressed neo(r) cassette provides required functions of the 1 gamma2b exon for class switching. Int. Immunol. 11, 1683–1692. Sheehy, A. M., Gaddis, N. C., Choi, J. D., and Malim, M. H. (2002). Isolation of a human gene that inhibits HIV-1 infection and is suppressed by the viral Vif protein. Nature 418, 646–650. Shinkai, Y., Rathbun, G., Lam, K., Oltz, E., Stewart, V., Mendelsohn, M., Charron, J., Datta, M., Young, F., and Stall, A. (1992). RAG-2-deficient mice lack mature lymphocytes owing to inability to initiate V(D)J rearrangement. Cell 68, 855–867. Skuse, G. R., Cappione, A. J., Sowden, M., Metheny, L. J., and Smith, H. C. (1996). The neurofibromatosis type I messenger RNA undergoes base-modification RNA editing. Nucl. Acids Res. 24, 478–485. Sohail, A., Klapacz, J., Samaranayake, M., Ullah, A., and Bhagwhat, A. S. (2003). Human activation-induced cytidine deaminase causes transcription-dependent, strand-biased C to U deaminations. Nucl. Acids Res. 31, 2990–2994. Stavnezer, J. (2000). Molecular processes that regulate class switching. Curr. Top. Microbiol. Immunol. 245, 127–168.
SOMATIC HYPERMUTATION
335
Strominger, J. (1989). Developmental biology of T cell receptors. Science 244, 943–950. Ta, V.-T., Nagaoka, H., Catalan, N., Durandy, A., Fischer, A., Imai, K., Nonoyama, S., Tashiro, J., Ikegawa, M., Ito, S., Kinoshita, K., Muramatsu, M., and Honjo, T. (2003). AID mutant analyses indicate requirement for class-switch-specific cofactors. Nat. Immunol. 4, 843–848. Taccioli, G., Rathbun, G., Oltz, E., Stamato, T., Jeggo, P., and Alt, F. (1993). Science 260, 207–210. Teng, B., Burant, C. F., and Davidson, N. O. (1993). Molecular cloning of an apolipoprotein B messenger RNA editing protein. Science 260, 1816–1819. van Gent, D., Hiom, K., Paull, T., and Gellert, M. (1997). Stimulation of V(D)J cleavage by high mobility group proteins. EMBO J. 16, 2665–2670. Vartanian, J. P., Henry, M., and Wain-Hobson, S. (2002). Sustained G –> A hypermutation during reverse transcription of an entire human immunodeficiency virus type 1 strain Vau group O genome. J. Gen. Virol. 83, 801–805. Vartanian, J. P., Meyerhans, A., Asjo, B., and Wain-Hobson, S. (1991). Selection, recombination, and G to A hypermutation of human immunodeficiency virus type 1 genomes. J. Virol. 65, 1779–1788. Wedekind, J. E., Dance, G. S., Sowden, M. P., and Smith, H. C. (2003). Messenger RNA editing in mammals: New members of the APOBEC family seeking roles in the family business. [erratum appears in Trends Genet. 2003 Jul;19(7):369]. Trends Genet. 19, 207–216. Wiesendanger, M., Kneitz, B., Edelmann, W., and Scharff, M. D. (2000). Somatic hypermutation in MutS homologue (MSH3, MSH6 and MSH3/MSH6) deficient mice reveals a role for the MSH2-MSH6 heterodimer in modulating the base substitution pattern. J. Exp. Med. 191, 579–584. Winter, D. B., Phung, Q. H., Zeng, X., Seeberg, E., Barnes, D. E., Lindahl, T., and Gearhart, P. J. (2003). Normal somatic hypermutation of Ig genes in the absence of 8-hydroxyguanine-DNA glycosylase. J. Immunol. 170, 5558–5562. Yamanaka, S., Balestra, M. E., Ferrell, L. D., Fan, J., Arnold, K. S., Taylor, S., Taylor, J. M., and Innerarity, T. L. (1995). Apolipoprotein B mRNA-editing protein induces hepatocellular carcinoma and dysplasia in transgenic animals. Proc. Natl. Acad. Sci. USA 92, 8483–8487. Yoshikawa, K., Okazaki, I. M., Eto, T., Kinoshita, K., Muramatsu, M., Nagaoka, H., and Honjo, T. (2002). AID enzyme-induced hypermutation in an actively transcribed gene in fibroblasts. Science 296, 2033–2036. Yu, K., and Lieber, M. (2003). Nucleic acid structures and enzymes in the immunoglobulin class switch recombination mechanism. DNA Repair 2, 1163–1174. Zan, H., Komori, A., Li, Z., Cerrutti, M., Flajnik, M. F., Diaz, M., and Casali, P. (2001). The translesional polymerase zeta plays a major role in Ig and Bcl-6 somatic mutation. Immunity 14, 643–653. Zeng, X., Winter, D. B., Kasmer, C., Kraemer, K. H., Lehmann, A. R., and Gearhart, P. J. (2001). DNA polymerase eta is an A-T mutator in somatic hypermutation of immunoglobulin variable genes. Nat. Immunol. 2, 537–541. Zhang, H., Yang, B., Pomerantz, R. J., Zhang, C., Arunachalam, S. C., and Gao, L. (2003). The cytidine deaminase CEM15 induces hypermutation in newly synthesized HIV-1 DNA. [see comment]. Nature 424, 94–98. Zhang, K., Cheah, H., and Saxon, A. (1995). Secondary deletional recombination of rearranged switch region in Ig isotype-switched B cells. A mechanism for isotype stabilization. J. Immunol. 154, 2237–2247.
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AUTHOR INDEX
A Aaltonen, L. A., 26 Abagyan, R. A., 289 Abbondandolo, A., 21 Aboussekhra, A., 194 Abruzzo, L. V., 317, 323 Absalon, M. J., 7 Aburantani, H., 25 Ackerman, E. J., 21 Adachi, N., 267 Adams, C., 151 Adar, S., 194 Addona, T. A., 7, 9 Adjiri, A., 194 Agata, Y., 319, 323 Aggarwal, A. K., 152, 212, 214, 233 Agustus, M., 168 Ahern, H., 7, 9 Ahmad, M., 77, 90, 91, 93 Ahn, J. Y., 24, 113, 114 Ahn, K., 53 Akagi, J.-I., 180, 189, 220, 275 Aksenova, A. Y., 320 Albesiano, E., 323 Ali, A., 26 Ali, S., 28 Allan, J. M., 6, 25 Allen, S. L., 323 Alley, J., 168 Allinson, S. L., 19, 21 Allis, D., 61 Almouzni, G., 60 Alon, U., 125, 127 Alt, F. W., 308, 310–312, 316, 318, 322 Altamirano, A., 19, 25 Al-Tassan, N., 26 Altschul, S. F., 112 Ames, B. N., 239
Amin, N. S., 155 Amin, S., 177, 210, 273 Anant, S., 311, 313, 323–325 Andegeko, Y., 123 Andersen, S., 25 Anderson, C. W., 114 Andersson, D. I., 241 Andersson, J., 312 Angliker, H., 27 Ansari, A., 53 Aono, N., 112 Aoufouchi, S., 24, 147, 221, 320, 321 Appella, E., 114, 119 Arad, G., 206, 249–252, 281 Arai, T., 25 Arakawa, H., 315 Araki, K., 168, 185 Araki, M., 60, 61, 206, 207, 217 Arany, Z., 23 Araujo, F., 94 Aravind, L., 51, 267–269 Argos, P., 232 Ariyoshi, M., 9, 10 Ariza, R. R., 27 Arlett, C. F., 207, 216 Armout, C. D., 189 Arnason, T., 286 Arndt, A., 291, 292 Arnold, K. S., 323 Arudchandran, A., 311 Arunachalam, S. C., 326 Arvai, A. S., 10, 15 Asahara, H., 155, 156 Asano, T., 316 Ashley, C., 297 Asjo, B., 325 Ataujo, R., 91 Augustine, M. L., 7 Avdievich, E., 121 Avkin, S., 194
337
338
AUTHOR INDEX
Avril, M. F., 217, 219 Ayaki, H., 87 Ayyagari, R., 293
B Baarends, W. M., 297 Baboshina, O. V., 286, 287 Bacchetti, S., 289 Bachant, J. B., 299 Bacher Reuven, N., 256 Bachl, J., 311 Bachmair, A., 287 Badie, C., 108, 116 Baer, M., 82 Baer, R., 286 Bailly, V., 7, 185, 287, 288 Bailone, A., 248 Baker, T., 150, 158 Bakkenist, C. J., 111 Balestra, M. E., 323 Baltimore, D., 118, 308 Bambara, R. A., 173, 190 Bandaru, V., 9 Banerjee, A., 4, 10, 11, 13–15, 17 Banerjee, S. K., 180 Banfalvi, G., 23 Bankmann, M., 291 Bao, K. K., 13 Barbanti-Brodano, G., 168 Barbour, L., 280, 298 Bardwell, P. D., 316, 322, 323 Barlow, C., 118 Barlow, T., 10 Barnes, D. E., 24, 25, 182, 297, 312, 316, 322 Baroin-Tourancheau, A., 289 Barreto, V., 317 Barrett, T. E., 10 Barron, L. L., 317, 323 Bar-Sagi, D., 120, 122 Barsky, D., 7 Bartek, J., 106, 107, 112–115, 122 Bartel, B., 287 Bartholomew, B., 61 Bartkova, J., 112, 115 Barton, J. K., 16 Baskaran, R., 118, 121 Basrur, V., 119 Bassett, E., 153
Bassett, H., 16 Basu, A. K., 19, 25 Batschauer, A., 93 Battista, J. R., 248 Batty, D., 53 Baxter, S. M., 51 Bayer, P., 291, 292 Bayliss, J., 324 Baynton, K., 179 Beach, D., 113 Beard, B. C., 16 Beard, W. A., 19, 139, 143, 147, 208 Beardsley, D. I., 121 Beaudenon, S., 291 Bebenek, K., 147–149, 151, 153, 184, 208–212, 221, 237, 238, 269, 271, 320 Becherel, O. J., 219, 242, 245, 249, 252 Beck, A. K., 168 Becker, J., 291, 292 Beese, L. S., 256 Begley, T. J., 15 Belfort, M., 51 Belguise-Valladier, P., 248 Bell, D. W., 114 Bell, S. D., 247 Bellacosa, A., 120, 121 Bemark, M., 143, 183, 185 Bender, C. F., 122 Benecke, A., 28 Benjamin, D., 27 Benjamin, R. C., 21 Benoist, C., 143, 308 Berardini, M., 148 Bercovich, B., 289 Berdal, K. G., 17, 193 Berek, C., 143, 157, 310 Beresford, P. J., 26 Beretta, B., 93, 94 Berg, P., 156 Bergoglio, V., 27 Bergstein, M., 245 Berleth, E., 287 Bermudez, V. P., 110 Bernad, A., 149 Bernards, A. S., 13 Bernier-Villamor, V., 292 Bernstine, E. G., 168 Berrocal-Tito, G., 77, 91, 93 Bertocci, B., 143, 147, 157, 310 Bertram, J. G., 153, 251, 252
AUTHOR INDEX
Bertrand, Y., 308 Bertrand-Burggraf, E., 51, 52 Besmer, E., 308, 312, 316, 318 Bessho, T., 43, 55 Best, J. M., 26 Betz, U. A., 316 Bezzubova, O., 185 Bhagwat, A. S., 15, 312, 318, 320 Bhagwat, M., 7 Bhakat, K. K., 18, 23, 118 Bharati, S., 10, 19 Bharti, A., 118 Bhattacharya, S., 323 Bichara, M., 248 Bieser, G., 79 Bieth, A., 27 Bignell, C. R., 112 Bingham, C. M., 26 Bird, A., 14, 15, 25 Bisgaard, M. L., 26 Bishop, S. M., 25 Biswas, T., 7 Bjoras, M., 9 Black, E., 19 Blackburn, G. M., 17 Bladl, A. R., 122 Blanco, L., 147, 149 Blanco, M., 248 Blandino, G., 118 Blobel, G., 292 Bloom, L. B., 208, 221 Blumenstein, S., 245 Blunt, T., 308 Boateng, M. A., 287 Bockrath, R., 56 Boddy, M. N., 285 Bodepudi, V., 13 Bodmer, J. L., 122 Boei, J. J. W. A., 185 Bogenhagen, D. F., 21, 152 Bohr, V. A., 21, 44, 56, 145, 322 Boiteux, S., 7, 18, 19, 24, 25, 193 Boldogh, I., 7, 16 Bolton, P. H., 7, 22–24, 55 Bond, J. P., 9, 169, 170 Bondo, B. E., 77, 91, 93 Boner, W., 220 Bonner, C. A., 242 Boon, E. M., 16 Boone, C., 287
Boorstein, R. J., 19, 25 Boosalis, M. S., 9, 208 Bootsma, D., 53, 57, 94, 207, 216, 297 Borden, A., 191 Borden, K. L., 285 Bories, J. C., 143 Bork, P., 112 Bosma, G., 312 Bosma, M., 312 Botella, L. M., 248 Botero, A., 23 Bottaro, A., 311 Bouayadi, K., 27 Bouchier-Hayes, L., 115, 116 Boudsocq, F., 152, 153, 212–215, 217, 219, 233, 234, 236 –239, 244, 245 Boulares, H. A., 26 Bouly, J. P., 93 Bowes Rickman, C., 94 Boyd, M. T., 155, 156 Brabec, V., 53 Bradbury, C. M., 23 Bradley, A., 113, 122, 311 Braithwaite, D. K., 168 Braman, J. C., 256 Bransteitter, R., 312, 316–320 Branum, M. E., 48, 52, 55, 56 Brash, D. E., 87 Bregeon, D., 2 Brendel, M., 169 Bresson, A., 179, 180, 191 Bresson-Roy, A., 179 Brick, P., 139 Bridges, B. A., 205 Briggs, W. R., 93 Broekhof, J. L., 25 Bronson, R., 310 Brooks, C. L., 115, 117 Broomfield, S., 280, 281, 288, 289, 292, 293, 297–299 Brotcorne-Lannoye, A., 230, 240 Broughton, B. C., 207, 216, 217, 219 Brousse, N., 313, 317 Brown, J., 308 Brown, K. D., 121 Brown, M., 289 Brown, T., 10, 13 Broyde, S., 63 Bruand, C., 139 Bruck, I., 139, 150, 206, 245, 249, 281
339
340
AUTHOR INDEX
Brudler, R., 75, 77, 91 Bruner, S. D., 7, 9, 11, 12, 15, 17 Brusky, J., 289 Bryans, M., 310 Bucher, P., 112 Buerstedde, J.-M., 185, 315 Bugg, C. E., 286 Buijs, R., 94 Bulavin, D. V., 119 Bullock, H. A., 26 Buluwela, L., 28 Bunn, H. F., 23 Bunny, K. L., 241 Bunting, K. A., 245, 247, 252 Burant, C. F., 325 Burckhardt, S. E., 248 Burgers, P. M., 22, 110, 175, 191, 192, 293, 294 Burgess, S. M., 47 Burma, S., 111 Burnouf, D. Y., 243, 245, 246 Burrows, C. J., 6 Bustin, M., 308 Buterin, T., 53, 63
C Cabral-Neto, J. B., 194 Cadet, J., 211 Cado, D., 121 Calandrini, D. M., 183, 194, 315 Caldecott, K. W., 21, 24, 145, 147 Callebaut, I., 308 Campbell, J. L., 151, 155 Canagarajah, B., 59 Canitrot, Y., 27 Canman, C. E., 113, 114, 118 Cann, I. K., 139 Cao, C., 14 Cao, X., 27 Cappelli, E., 21 Cappione, A. J., 325 Carattini-Rivera, S., 113 Carlson, C., 311 Caron, G., 326 Carpenter, A. J., 108, 116 Carpenter, P. B., 112 Carr, A. M., 110 Carrodeguas, J. A., 152
Carrozzino, F., 21 Carson, M., 213, 245 Carswell-Crumpton, C., 281 Cartegni, L., 325 Carter, D. M., 207 Carter, K. C., 168 Carter, T., 308 Casali, P., 183, 321 Caseira-Cabral, R. E., 194 Casellas, R., 308, 312 Cashmore, A. R., 73, 75, 77, 90–92, 94 Caspari, T., 110 Casselman, R., 110 Cassier, C., 170 Cassier-Chauvat, C., 168, 194 Castano, I. B., 151 Castle, B. E., 317, 323 Catalan, N., 313, 316, 317 Cavazzana-Calvo, M., 308 Cazaux, C., 27 Celis, J. E., 324 Ceriani, M. F., 93 Cerosaletti, K. M., 113 Cerritelli, S., 311 Cerrutti, M., 321 Cerutti, A., 183 Cesare, A. J., 110 Cevasco, M., 21 Chabes, A., 152 Chae, S.-K., 168, 171 Chaganti, R. S., 323 Chambon, P., 28 Chan, B. H., 325 Chan, E., 18 Chan, L., 325 Chan, M. K., 19, 25 Chan, T. A., 107 Chanet, R., 170, 194 Chaney, S. G., 153, 210 Chang, D. Y., 6, 16 Chang, G. J., 121 Chapados, B. R., 21, 22 Chapuis, J., 139 Charron, J., 308 Chaskara, V., 122 Chau, B. N., 116, 118 Chau, V., 285, 287 Chaudhuri, J., 310–312, 316, 318 Chaung, W., 25 Chauveau, C., 311
AUTHOR INDEX
Cheadle, J. P., 26 Cheah, H., 311 Cheh, A. M., 210, 211, 273 Chen, B. J., 9 Chen, B. P., 111 Chen, C.-C. A., 268 Chen, D., 28 Chen, D. J., 111 Chen, D. S., 18 Chen, J., 112, 311 Chen, P., 114 Chen, T. T., 108, 116 Chen, X., 143, 158, 311 Cheng, H., 310 Cheng, L. S., 117 Cheng, X., 15, 81, 88 Cheng, Y. C., 19 Chernov, M. V., 189 Chester, A., 324 Cheung, A., 114 Cheung, K.-J., 189 Chiapperino, D., 210, 211, 273 Chini, C. C., 112 Chiolo, I., 298 Chiorazzi, N., 323 Chipuk, J. E., 115, 116 Chittenden, T., 115, 116 Chmiel, N. H., 13, 16, 26 Choi, B. S., 211 Choi, J. D., 326 Choi, Y., 27 Chou, K. M., 19 Chow, B. L., 168, 289, 293, 297 Chow, K. H., 281 Christen, R. D., 118 Christensen, R. B., 168–170, 185, 187, 188 Christman, M. F., 151 Chu, G., 64, 119, 122 Chua, K., 311, 312, 316, 318 Chua, N. H., 93 Chumakov, P. M., 189 Chung, U., 18 Chyan, J. Y., 19 Ciechanover, A., 283, 289 Citaterna, M. H., 94, 96 Clark, A. B., 121 Clark, D. R., 171 Clark, S. C., 325 Clarke, A. R., 25 Clarke, N. D., 13
Clarkin, K., 108 Clarkson, S. G., 22–24 Cleaver, J. E., 44, 60, 207 Cocea, L., 147 Cogne, M., 311 Cohen, Y., 24 Coin, F., 248 Cole, P. A., 118 Coleman, J. E., 46 Collado, P., 248 Collins, J. E., 324 Concannon, P., 113 Cong, F., 119, 122 Connaughton, J. F., 212, 230, 232 Connolly, J. A., 220 Constantinou, A., 22–24 Cook, W. J., 285, 286 Coombes, K. R., 317, 323 Coombes, R. C., 28 Cooper, E., 325 Copeland, W. C., 139, 148, 152, 157 Coquerelle, T., 27 Cordon-Cardo, C., 117 Cordonnier, A. M., 217, 219, 242, 249 Corneo, B., 308 Cortes, P., 308 Cortez, D., 110, 111, 113 Costanzo, A., 117, 118 Cotticelli, M., 312 Coull, B., 216, 217 Courcelle, J., 248, 281 Cox, L. S., 21 Cox, M. M., 255, 281 Crabtree, M., 26 Craig, H. M., 325, 326 Craig, L., 122 Cranston, A., 322 Creighton, S., 208, 221 Crick, F. H. C., 158 Critchlow, S. E., 149 Croce, C. M., 155, 168, 187, 189, 275 Cromwell, C. C., 317, 323 Cross, T. G., 122 Crouch, R., 311 Crowley, D., 117 Cui, X. S., 113 Cullis, P. M., 6 Cumano, A., 143, 183, 185 Cunningham, M. L., 26 Cunningham, R. P., 7, 9, 15, 19, 20, 25
341
342
AUTHOR INDEX
Cuomo, C., 308 Curran, T., 18, 23, 26 Curren, R. D., 220
D Dahan, A., 143, 147 Dahlen, M., 110 Daiyasu, H., 75, 77, 91 Dalla-Favera, R., 323 Dalton, T. P., 265 Daly, G., 25 Damle, R. N., 323 D’Amours, D., 112 Dance, G. S., 325 Daniel, R., 139 Dansereau, J. T., 51 Dantzer, F., 24 Dargemont, C., 292 Darlington, T. K., 93 Das, G., 168, 169 Das, S., 311 Datta, A., 155, 170 Datta, M., 308 Daune, M., 23 Davey, S., 110, 113 David, L., 27 David, S. S., 13, 16, 26 Davidson, L., 308, 310–312 Davidson, N. O., 311, 313, 323–325 Davies, D. R., 26 Davies, S. L., 143, 183, 185 Davis, A., 311 Davis, H. L., 113, 114 Davis, R. J., 119, 120, 122 Dawut, L., 91, 94 Deacon, E., 122 Dean, K., 287 de Boer, J., 316 de Chasseval, R., 308 Decourt, C., 311 Defais, M., 280 Deisenhefer, J., 77, 78, 81 de Jong, A. W. M., 168, 171 Delagoutte, E., 51, 52 de La Rubia, G., 24 Delarue, M., 232 De Las Penas, A., 151 De Laurenzi, V., 117
Delbos, F., 147 de Lera, L. T., 147 del Mazo, J., 147 Delsol, G., 27 DeLucia, A. M., 233 DeMayo, F., 113 Demengeot, J., 308 Demple, B., 18, 192, 193 de Murcia, G., 23, 24, 316 Deng, Y., 114 Deng, Z., 9 Denissenko, M. F., 265 Derbyshire, V., 51 de Ruiter, P. E., 77, 79 Dervyn, E., 139 De Smet, A., 143, 157, 310 Desnoyers, S., 26 Dessing, M., 311 Desterro, J. M., 292 Deutsch, W. A., 23 de Villartay, J., 308 Devoret, R., 248 de Weerd-Kastelein, E. A., 207, 216 de Wind, N., 185, 316 de Wit, J., 25, 57, 94, 297 Dianov, G. L., 19, 21, 25, 145 Dianova, I. I., 19, 21 Diaz, M., 182, 183, 321 Dickerson, S. K., 312, 316, 318 Dierich, A., 143, 308 Dighiero, G., 317 Dikkes, P., 310 Di Leonardo, A., 108 Ding, C. K., 26 Dinner, A. R., 17 Di Noia, J., 157, 182, 316, 322 Dionne, I., 247 Dittmar, G., 286 DiTullio, R. A., Jr., 112 Djamei, A., 93 Doddridge, Z. A., 2 Dodson, L. A., 248 Dodson, M. L., 7, 9 Doetsch, P. W., 2, 22–24 Dogliotti, E., 21 Doherty, A. J., 247 Dohmae, N., 206, 207, 217 Dohmen, R. J., 287 Dolwani, S., 26 Dominguez, O., 147, 149
343
AUTHOR INDEX
Donker, I., 25 Doorly, M., 119 Dornan, E. S, 220 Dosch, J., 27 Dover, J., 287 Downey, K. M., 173, 190 Drapkin, R., 53, 59, 122 Driscoll, D. M., 325 Drohat, A. C., 14, 17 Droin, N. M., 115, 116 Dua, R., 151, 155 Dudkin, E. A., 93 Dudley, D., 311, 318, 322 Due, M., 313 Dufourcq-Labelouse, R., 313, 317 Dumanski, J. P., 324 Dumas, G., 317 Dumas, P., 243, 245, 246 Dunand, J., 51 Duncan, B. K., 14 Duncan, G., 114 Dunham, I., 324 Dunnick, W., 122 Dunstan, H. M., 168 Duong, J. K., 112, 114 Durandy, A., 313, 316, 317 Durocher, D., 112, 114 D’Urso, G., 155 Dutreix, M., 248
E Earnshaw, W. C., 26 Ebbert, A., 56 Ebisu, S., 18 Ebright, J. N., 94 Eccles, D., 26 Echols, H., 242, 248, 249 Ecker, D. J., 287 Edelmann, W., 316, 322 Edwards, R. A., 284, 285, 290 Eeken, J. C. J., 168, 171, 185–187 Egly, J.-M., 53, 55 Ehrenstein, M. R., 316, 322 Ehrlich, S. D., 139 Eichman, B. F., 14, 17 Einhorn, L. H., 26 Eisen, J. A., 63 Eisenberg, M., 13
Eisenberg, W., 194 Eisler, H., 170 Eker, A. P. M., 53, 77, 79, 94 Elder, R. H., 25, 26 Elenitoba-Johnson, K. S., 317, 323 Elia, A. J., 114 Elias, S., 283 Elledge, S. J., 104, 110–114, 155, 299 Ellenberger, T., 6, 9–11, 14, 17, 45, 152, 194, 212–214, 233 Ellis, A., 26 Ellison, M. J., 284–286, 289, 290, 297 Elowitz, M. B., 125, 127 Emery, P., 93, 94 Emmerson, P., 26 Engels, K., 26 Engelward, B. P., 9, 25, 316 Epe, B., 25 Epstein, J. A., 207, 216 Erez, N., 24 Eritja, R., 206, 242, 249, 281 Errington, J., 139 Ersoy, F., 313, 317 Esashi, F., 112 Escalante, C. R., 152, 212, 214, 233 Escobar, S. R., 118 ESCODD, 15 Esposito, G., 143, 183, 185, 271, 316 Essers, J., 316 Estivill, X., 289 Etienne, H., 241–243 Eto, T., 323 Evan, G. I., 285 Evans, D. G., 26 Evans, D. H., 75 Evans, E., 53, 55 Evans, J., 308 Evans, R. M., 28
F Fabre, F., 194 Fagarasan, S., 182, 311, 313 Faili, A., 147, 221, 320, 321 Fajnok, M. F., 183 Falck, J., 106, 107, 112, 114, 115, 122 Falcovitz, A., 23 Fan, J., 323 Fan, M., 323
344
AUTHOR INDEX
Fan, Z., 26 Fanciulli, M., 118 Fanklin, W. A., 87 Fath, D., 312 Favaudon, V., 118 Fawcett, H., 217, 219 Fay, D. S., 112, 114 Feaver, W. J., 269, 271, 274 Fernandes, A. S., 14 Ferna´ndez de Henestrosa, A. R., 149, 216, 217 Ferrell, L. D., 323 Ferrini, R., 308, 310, 312 Feunteun, J., 112 Fiala, K. A., 234, 236, 237 Fidalgo, P., 26 Fink, D., 118 Finkel, S. E., 206, 240 Finley, D., 283, 286, 287 Finnie, N., 308 Fischer, A., 308, 313, 316, 317 Fischer, R. L., 27 Fischhaber, P. L., 152, 155, 189, 220, 269, 271, 274, 275 Fishel, R., 155, 168, 187, 189, 275, 322 Fisher, C. L., 7 Fisk, H. A., 286 Flaherty, D. M., 23 Flajnik, M. F., 183, 315, 321 Flatter, E., 24, 143, 147, 221, 321 Flavell, R. A., 120, 122 Fleming, N., 26 Flores, E. R., 117 Fluke, D. J., 79 Foiani, M., 155, 298 Fontanie, T., 168, 289 Fontemaggi, G., 118 Foote, R. S., 6 Foray, N., 118 Fornace, A. J., Jr., 119 Fortini, P., 21 Forveille, M., 313, 316, 317 Foster, P. L., 148, 206, 240 Foster, R. S., 26 Fox, M., 289 Francis, A. W., 13, 16 Francis, L., 241 Frank, E. G., 143, 147, 148, 177, 184, 194, 206–212, 216, 221, 249, 250, 252, 281, 321
Frank, K., 310 Franklin, K. A., 52, 81 Fraumeni, J. F., 114 Frayling, I., 26 Frechet, M., 27 Freemont, P. S., 285 Freidberg, E. C., 141 Fridman, J. S., 108, 115 Fried, L., 308 Friedberg, E. C., 57, 59, 61, 102, 143, 147, 152, 155, 168, 189, 205–207, 220, 229, 265, 267–269, 271, 274, 275 Friedli, M., 326 Friend, S. H., 168 Fritz, G., 19, 23, 26 Fromme, J. C., 4, 9–15, 17 Frosina, G., 21 Fruechte, E. M., 94, 96 Fuchs, R. P., 51, 52, 152, 153, 155, 168, 179, 180, 191, 206, 217, 219, 229, 232–235, 237, 239–243, 245, 246, 248–252, 256 Fugmann, S., 310 Fujii, S., 234, 235, 242, 243, 245, 246, 250–252, 256 Fujii-Kuriyama, Y., 266 Fujimoto, H., 266 Fujimoto, K., 23 Fujiwara, Y., 310 Fukao, T., 316 Fukasawa, M., 267 Fukita, Y., 316 Fukuda, M., 25 Fukuda, R., 230, 239 Fukuda, S., 168, 173 Fulco, M., 118 Funahashi, T., 323 Furuichi, M., 25 Fuss, J., 151, 155, 156 Futcher, B., 292 Fygenson, K. D., 208
G Gaarde, W. A., 119 Gaddis, N. C., 326 Gaiddon, C., 23 Galan, J. M., 286 Galante, J., 108, 116 Gale, J. M., 59, 60
AUTHOR INDEX
Galgoczy, D. J., 108, 109 Gali, R. R., 286 Galibert, F., 194, 248 Galkin, A. P., 320 Gallant, M., 26 Gallinari, P., 19 Gamper, H., 43, 51 Ganesan, A. K., 7 Gao, L., 326 Gao, T., 267, 274 Gao, Y., 310 Garcia-Diaz, M., 147, 149 Garcia-Ortiz, M. J., 147 Garcia-Palomero, E., 147 Gareau, Y., 26 Gares, M., 27 Garkavtsev, I., 189 Garritsen, V. H., 217, 219 Gary, R., 21 Gary, S. L., 293 Gasparutto, D., 211 Gasser, V., 250–252 Gatter, K. C., 26 Gaudreau, A., 23 Gauger, M., 153 Geacintov, N. E., 63, 153, 173, 177, 178, 210, 241, 242, 273 Gearhart, P. J., 143, 183–185, 208, 212, 216, 221, 312, 316, 320–322 Gehring, M., 27 Geissmann, F., 313, 317 Gellert, M., 157, 308 Gennery, A., 313, 317 Gentil, A., 194 Georgescu, R. E., 243 Gerchamn, S. E., 14 Gerlach, V. L., 143, 267–269, 271, 274 Gerlt, J. A., 7 Getzoff, E. D., 75, 77, 91 Geva-Zatorsky, N., 125, 127 Ghavidel, A., 23 Ghirlando, R., 122 Giaccia, A. J., 115, 117 Giannoni, F., 323 Gibbs, P. E. M., 167, 168, 171–176, 179, 180, 187, 188, 190, 191 Gibson, N. J., 13 Giedroc, D. P., 46 Giercksky, K. E., 6 Gietz, R. D., 170
345
Gilboa, R., 14 Gilfillan, S., 143, 308 Gill, D. M., 21 Gillespie, D., 19 Giovani, B., 93 Girard, P. M., 7, 114 Gisbourne, J., 324 Gius, D., 23 Glassner, B. J., 9–11, 27, 170, 193 Glastre, C., 316, 317 Glickman, B. W., 312 Glover, B. P., 150 Glover, J. N., 284, 285, 290 Godin, I., 183, 185 Godindagger, I., 143 Godoy, V. G., 250 Goff, S. P., 119, 122 Gogos, A., 13 Golan, G., 14 Gold, B., 25 Gold, L. S., 239 Goldberg, M., 112, 114 Goldberg, R. B., 27 Goldfinger, N., 23 Golding, G. B., 312 Goldsby, R. E., 143, 158 Goldstein, G., 283 Golinelli, M. P., 13 Golshani, A., 287 Gonda, D. K., 287 Gonda, H., 319, 323 Gong, J. G., 117, 118 Gong, Z., 27 Gonzalez, F. J., 265 Gonzalez, M., 250 Gonzalez, M. A., 149 Gonzalez, S., 117 Gonzalez-Barrera, S., 149 Goodman, M. F., 137, 139, 150, 152, 153, 156, 157, 168, 172, 206, 208, 209, 212–214, 221, 229, 233, 235–237, 239–243, 249–252, 255, 256, 273, 281, 312, 316–320 Goosen, N., 51 Goossens, T., 323 Gorka, C., 308 Goswami, P. C., 23 Gottesman, M. E., 57 Gottlieb, T., 308 Gotway, G., 267–269
346
AUTHOR INDEX
Grallert, B., 155 Granek, J. A., 13 Grawunder, U., 310 Gray, C., 216, 217 Gray-Schopfer, V., 311 Green, D. R., 115, 116 Greenberg, M., 310 Greenblatt, J. F., 287 Greeve, J., 317, 323 Griffin, E. A., Jr., 93 Griffin, P. R., 26 Griffith, J. D., 51, 81, 110, 111 Grigorian, I. A., 189 Grindley, N. D., 233, 237 Groisman, R., 122 Grollman, A. P., 6, 7, 9, 13, 14, 239, 273 Gromov, P., 324 Grootegoed, J. A., 297 Grossman, L., 7 Gruz, P., 206, 232–235, 237, 239–242 Gu, H., 143, 316 Gu, J., 122 Gu, W., 115, 117 Gu, Y., 6, 16, 19, 308, 312 Guan, Y., 19, 20 Gudkov, A. V., 189 Gueranger, Q., 221, 321 Guerrera, S., 155 Guibourt, N., 7 Guillet, M., 18, 193 Guntuku, S., 113 Gunz, D., 22, 23, 24 Guo, C., 143, 155, 189, 220, 271, 275 Guo, D., 153, 175–177, 190, 210, 273 Gupta, M., 18 Gupta, N., 147 Guthrie, C., 47 Guy, J., 25
H Haas, A. L., 286, 287 Haas, B. J., 19, 20 Haber, J. E., 156 Haguenauer-Tsapis, R., 286 Hahn, S., 61 Hahn, W. C., 117 Haines, D. S., 155, 156 Haire, L. F., 112, 114
Halazonetis, T. D., 112 Hall, J. C., 93, 94 Hamilton, J. W., 43 Hamlin, R., 139 Hamm-Alvarez, S., 77 Han, D.-M., 168, 171 Han, K.-Y., 168, 171 Han, S., 21, 22 Hanada, K., 267 Hanaoka, F., 53, 60, 61, 141, 143, 152, 153, 168, 177, 180, 189, 206–212, 215–217, 219–221, 229, 233, 239, 269, 273–275, 320, 321 Hanawalt, P. C., 44, 45, 52, 56, 57, 63, 64, 248, 281 Hang, B., 25 Hanna, M., 289, 293 Hannon, M., 27 Hanson, I. M., 285 Hara, R., 59, 61, 64 Haracska, L., 155, 172–178, 190–192, 195, 206, 208–211, 215, 219, 243, 272, 274 Harada, J. J., 27 Harada, N., 114 Harashima, H., 233, 239 Hardeland, U., 15, 28 Hardigree, A. A., 168, 186, 206, 230 Hardwich, K. G., 155 Hardy, R., 310 Harfe, B. D., 170 Harriman, G., 311 Harris, A. L., 26, 152, 155 Harris, C. C., 24 Harris, R. S., 157, 315, 322, 325, 326 Hart, S. M., 28 Hartwell, L. H., 108, 109, 168, 189 Harvey, C., 289 Hase, Y., 168, 171 Hasegawa, R., 9, 25 Haseltine, W. A., 87 Hastings, P. J., 170, 206, 241 Hata, Y., 174, 180, 191, 194 Hatada, E., 230, 239 Hatahet, Z., 274 Hauschild, J., 315 Haushalter, K. A., 10, 25 Havener, J. M., 153 Hay, R. T., 292 Hayakawa, H., 150
AUTHOR INDEX
Hayakawa, K., 310 Hayakawa, S., 116 Hayashi, I., 234, 235, 239 Hayashi, K., 239 Hays, L. E., 143, 158 Hazel, J. C., 206 Hazra, T. K., 9, 18, 23 He, M., 112 Hearst, J. E., 43, 47, 51, 52 Heelis, P. F., 79, 83–85 Hegde, V., 23 Heidenreich, E., 170 Heidorn, K., 317, 323 Heidt, J., 287 Heinimann, K., 26 Heller, H., 283 Helquist, S. A., 215 Hemmingsen, S. M., 289 Henderson, B., 122 Henderson, J. O., 324, 325 Henderson, S. T., 291 Hendrich, B., 15, 25 Hendry, J. H., 108 Henning, K. A., 57 Henriques, J. A. P., 169, 170 Henriquez, N. V., 122 Henry, M., 326 Hensen, E. S., 192, 193 Herman, T., 18 Hermeking, H., 107 Hermine, O., 316, 317 Herrera, G., 248 Herrup, K., 308 Hershko, A., 283 Hess, D., 27 Hess, P., 120, 122 Hessels, J. K. C., 77 Heude, M., 194 Hey, T., 53 Hiai, H., 323 Hickson, I. D., 19, 24, 26 Hidaka, M., 18 Hideki, K., 266 Higashimoto, Y., 119 Higley, M., 16 Higuchi, K., 18 Higuchi, Y., 77, 79 Hikida, M., 266 Hill, C. P., 285, 288 Hindges, R., 53
347
Hingorani, M., 243 Hinkle, D. C., 152, 167, 168, 171–177, 179, 186, 188–191, 205, 206, 229, 232 Hiom, K., 308 Hippou, Y., 25 Hirano, M., 25 Hirao, A., 114 Hirata, Y., 79, 84, 85 Hirota, K., 23 Hitomi, K., 75, 77, 88–89, 91, 94, 96 Hladik, C. L., 143, 267, 271, 274 Hobeck, S.L., 170, 184 Hochstrasser, M., 283, 285, 291 Hodges, A. K., 26 Hodgson, S. V., 26 Hoege, C., 155, 187, 219, 243, 274, 288, 292–294, 297, 298, 300 Hoeijmakers, J. H., 9, 25, 53, 57, 297 Hoekstra, M. F., 110–112 Hoffmann, J. S., 27 Hoffmeyer, M. R., 122 Hofmann, K., 110, 112 Hofmann, R. M., 286, 289, 290 Hofseth, L. J., 24 Hokijmakers, J. H. J., 94 Holbrook, S. R., 43, 81–83 Hollis, T., 14 Holmes, A. M., 156 Holmes, E. W., 26 Holpert, M., 51 Holzmann, V., 170 Honda, K., 116 Hong, S. H., 325 Honjo, T., 182, 311, 313, 315, 317, 323 Hoogerbrugge, J. W., 297 Hopfield, J. J., 47 Hopfner, K. P., 122 Horiuchi, T., 18 Horton, J. K., 21 Hosaka, Y., 217, 219 Hosfield, D. J., 7, 19–22, 110 Hoss, M., 22–24 Hostomsky, Z., 24 Hou, E., 208 Howard-Flanders, P., 280 Howell, S. B., 118 Howley, P. M., 291 Hrivnak, G., 25 Hryciw, T., 280, 281, 288, 289, 292, 299 Hsiao, J., 112, 114
348
AUTHOR INDEX
Hsieh, C. L., 22 Hsieh, M. M., 23 Hsu, D. S., 51, 53, 55, 77, 81, 87–91, 94 Huala, E., 93 Huang, J.-C., 43, 53, 55 Huang, L., 291 Huang, L. E., 23 Huang, M.-E., 194 Huang, S. J., 11, 13–15 Huang, Y., 119 Hubscher, U., 21, 53, 137, 156 Huibregtse, J. M., 285, 291 Huletsky, A., 23 Hunninghake, G. W., 23 Hunter, T., 118 Hunter, W. N., 13 Hurwitz, J., 21, 110, 195, 219, 274, 293 Husain, I., 77, 81–83 Hussain, S. P., 24 Hutchinson, F., 237 Hwang, B. J., 119, 122 Hwang, G. S., 211 Hwang, J. R., 53, 55 Hwang, W. W., 287
I Iacomini, J., 308 Ianculescu, A. G., 287 Ichihara, M., 189, 275 Ichikawa, Y., 14, 17 Ichinose, M., 266 Ide, H., 9, 25 Iden, C. R., 7, 9 Igarashi, T., 18 Iglesias-Ussel, M., 322 Ihara, M., 77, 87 Ikeda, S., 7 Ikegawa, M., 317 Ikenaga, M., 87, 89 Imai, K., 116, 316, 317 Impellizzeri, K. J., 293 Inaka, K., 77, 79 Inman, R. B., 281 Innerarity, T. L., 323 Inoue, H., 168, 171, 186, 297 Inoue, S., 118 Inoue, Y., 77, 79 Inui, T., 87
Irwin, M. S., 117 Isaacs, R. J., 63 Ishibashi, T., 150 Ishii, C., 168, 171, 186 Ishii, H., 155, 168, 187, 189, 275 Ishikawa, T., 88, 94, 96, 212, 266 Ishiko, T., 119 Ishino, Y., 139 Ishiura, M., 75, 77, 91 Isselbacher, K. J., 114 Ito, J., 168 Ito, S., 317 Ito, T., 61 Itoh, M., 25 Itoh, S., 274 Itoh, T., 64, 121, 217, 219 Itzhaki, J. E., 108, 116 Iwai, S., 9, 10, 18, 53, 61, 87–89, 141, 153, 174, 180, 182, 191, 206–208, 210–212, 215–217, 221, 239, 269 Iwasaki, H., 248, 281 Iyer, N., 57, 59 Izumi, T., 7, 9, 18–20, 23, 192
J Jack, M. T., 114 Jacks, T., 117 Jackson, S. P., 110, 112, 114, 149, 247 Jacobs, H., 316 Jacobs, M. A., 143, 183, 185 Jacobsen, S. E., 27 Jaenicke, R., 291, 292 Jager, J., 139 Jaiswal, A., 23 Janawalt, P. C., 44, 56 Janda, J., 317, 323 Janel-Bintz, R., 239, 241, 242 Jankovic, M., 318, 320 Janniere, L., 139 Jarillo, J. A., 91 Jarima, N., 87, 89 Jarmuz, A., 324 Jaspers, N. G., 217, 219 Jaspers-Dekker, I., 297 Jayaraman, L., 23 Jeffrey, L. C., 285 Jeffrey, P. D., 285, 291 Jeggo, P., 118, 308, 310
AUTHOR INDEX
Jensen, D. E., 46 Jensen, E. L., 168 Jentsch, S., 155, 187, 219, 243, 274, 283, 285, 286, 288, 291, 292–294, 297–300 Jerina, D. M., 153, 177, 210, 211, 239, 241, 273 Jeruzalmi, D., 243 Jessberger, R., 156 Jeusset, J., 194 Jiang, Y. L., 2, 6, 14, 16, 17 Jin, B., 121 Jin, S., 118 Jinks-Robertson, S., 170 Jiricny, J., 10, 15, 19, 28 Joachimiak, A., 293 Joazeiro, C. A., 285 Johansen, R. F., 17, 193 Johansson, E., 175, 191, 192 Johnson, A. W., 18 Johnson, E. S., 292 Johnson, F., 13 Johnson, J. L., 77 Johnson, K. A., 234 Johnson, L., 27 Johnson, P., 24 Johnson, R., 308 Johnson, R. E., 18, 152, 172, 173, 175–177, 190–192, 195, 206–212, 214, 215, 217, 219, 233, 243, 271–274 Johnson, S. J., 256 Johnston, M., 287 Jones, A. M., 322 Jones, S., 26 Jones, S. N., 120, 122 Jonson, R. E., 291 Jonsson, Z. O., 21 Jordan, S., 26 Jorns, M. S., 79, 83 Jost, J. P., 27 Jost, Y. C., 27 Joyce, C. M., 139, 233, 237 Juarez, R., 147, 149 Jung, S., 311
K Kaelin, W. G., Jr., 117, 118 Kafer, E., 171 Kaguni, L. S., 152
349
Kahyo, T., 292 Kai, M., 110, 243, 275 Kaina, B., 19, 23, 26, 27 Kajiwara, K., 186 Kakazu, N., 323 Kaklamanis, L., 26 Kakolyris, S., 26 Kakutani, T., 27 Kamath, R., 121 Kamide, R., 121, 217, 219 Kamiya, H., 233, 239 Kamiya, K., 155, 168, 173, 189, 220, 275 Kaneda, Y., 61 Kanehisa, M., 75, 77, 91 Kaneko, H., 316 Kaneko, M., 93, 94 Kang, C., 81, 82 Kang, D. H., 114 Kanjo, N., 180, 189, 220, 275 Kanke, Y., 233, 239 Kanno, S., 9, 25, 94 Kannouche, P. L., 216, 217, 220, 243 Kanter-Smoler, G., 110 Kanuri, M., 210 Karcher, A., 122 Karin, M., 119, 286 Karnitz, L. M., 110 Karplus, M., 17 Karran, P., 145 Kasahara, K., 316 Kasahara, Y., 316 Kashiwagi, T., 9, 10 Kasmer, C., 184, 221, 320 Kaspa´rkova´, J., 53 Kassabov, S. R., 61 Kastan, M. B., 111, 112, 115, 117, 118 Katakai, T., 319, 323 Katayama, T., 18 Kato, T., 206, 248, 267, 281 Kato, T., Jr., 267 Katori, N., 217, 219 Kaufmann, S. H., 26 Kavakli, I. H., 77, 91, 93 Kavli, B., 10, 316, 317 Kawaguchi, N., 174, 180, 191 Kawamura, K., 266, 267 Kawano, M., 267 Kawashima, A., 53, 121 Kawate, T., 7, 9 Kay, S. A., 93, 94
350
AUTHOR INDEX
Kaylor, L., 311, 318, 322 Kayserili, H., 313, 317 Kazantsev, A., 55, 77, 90, 91, 94 Keating, K. M., 46 Keating, M. J., 317, 323 Kedar, P., 21 Keegan, K., 112 Keeney, S., 121, 299 Kelley, M. R., 23, 26 Kelly, R. C., 46 Kelly, U., 94 Kelly, V. P., 25 Kelman, Z., 150, 293 Kennedy, S., 324, 325 Kenny, M. K., 21 Kenter, A., 308, 312 Kenyon, C. J., 230 Keranen, S., 151 Kesti, T., 151 Khamlichi, A. A., 143, 183, 185, 311 Kharbanda, S., 118, 119 Khoung, C., 311, 312, 316, 318 Kiefer, J. R., 256 Kiener, A., 77, 82 Kikuchi, H., 116 Kikuchi, Y., 292 Kilbey, B. J., 180 Kim, J., 312 Kim, J. K., 211 Kim, K., 21 Kim, N., 139, 148 Kim, S. J., 21 Kim, S.-R., 206, 230, 232–235, 237, 239–241 Kim, S.-T., 51, 77–79, 81, 83–85, 87–89, 111, 112 Kim, Y. S., 93 Kimura, M., 186 Kingston, R. E., 61 Kinnucan, E., 291 Kinoshita, K., 182, 311, 313, 315, 317, 323 Kinoshita, T., 27 Kinoshita, Y., 27 Kinsohita, K., 317 Kinzler, K. W., 107 Kirchgessner, C., 308 Kirchhoff, T., 147, 149 Kirk-Bell, S., 207, 216 Kisker, C., 152, 155, 189, 220, 275 Kisselev, A. F., 122 Kitadokoro, K., 77, 79
Kitao, H., 119 Klapacz, J., 312, 318, 320 Klapper, W., 317, 323 Kleijer, W. J., 217, 219 Klein, H., 194 Klein, U., 143, 183, 185, 316 Klinman, N. R., 183, 321 Klungland, A., 21–25, 297 Kneitz, B., 322 Kobayashi, K., 9, 25, 94, 174, 180, 191 Kobayashi, S., 235, 236, 237, 239 Koffel-Schwartz, N., 248 Kofoid, E., 241 Kogoma, T., 281 Kohler, G., 308 Kohlhagen, G., 18 Kohn, K. W., 18 Koken, M. H., 297 Kokoska, R. J., 141, 153, 210, 211, 214, 233, 237, 238 Kolbanovskiy, A., 241, 242, 273 Kolchanov, N. A., 312, 318 Kolodner, R. D., 121, 155 Komori, A., 183, 321 Komori, H., 77, 79 Komori, T., 308 Komoro, K., 25 Kondo, K., 117 Kondo, N., 316 Kondratick, C. M., 155, 195, 206, 207, 217, 274 Kong, X. P., 252 Kooiman, P., 77 Kool, E. T., 215 Koonin, E. V., 51, 112, 212, 230, 232, 267–269, 289 Kornberg, A., 150, 158 Kornberg, R. D., 59 Korolev, S. V., 293 Kouchakdjian, M., 13 Kow, Y. W., 7, 9 Kowalczkowski, S. C., 194 Kowalski, J. C., 51 Koyoshi, S., 271 Kozak, M., 187 Kraemer, K. H., 44, 184, 221, 320 Krahn, J. M., 147 Krainer, A. R., 325 Kramarczuk, I. H., 210, 211, 273 Kramata, P., 177
AUTHOR INDEX
Krangel, M., 308 Krause, K., 317, 323 Krauss, G., 53 Krejci, L., 194 Krogan, N. J., 287 Krokan, H. E., 6, 10, 19, 25, 316, 317 Kroth, H., 210, 211, 239, 241, 273 Kruas, B. R., 187 Krzeminsky, J., 177, 210, 273 Kubota, Y., 24, 297 Kucherlapati, R., 322 Kucho, K., 75, 77, 91 Kufe, D., 23, 119 Kugelberg, E., 241 Kulaeva, O. I., 212, 230, 232 Kumagai, K., 267 Kumaki, S., 316, 317 Kume, H., 266 Kung, H. C., 55 Kunitomi, N., 174, 180, 182, 191 Kunkel, T. A., 19, 93, 139, 141, 147–149, 151–153, 157, 168, 173, 184, 190, 206, 208–212, 214, 221, 229, 233, 237, 238, 269, 271, 320 Kunz, B. A., 170, 192, 193 Kuo, C. F., 7 Kuppers, R., 323 Kuramitsu, S., 77, 79 Kuraoka, I., 122, 211 Kurihara, N., 121 Kurimasa, A., 111 Kuriyan, J., 243, 252 Kurosky, A., 7 Kusumoto, R., 60, 206, 207, 210, 211, 217, 269 Kuwana, T., 115, 116 Kuzminov, A., 281 Kwon, K., 14 Kycia, J. H., 14
L Labelle, M., 26 Lacroix-Triki, M., 27 Lagrazon, K., 286 Lahav, G., 125, 127 Lai, W. C., 267, 274 Lain de Lera, T., 149 Lalani, E., 143, 183, 185
351
Lam, K., 308 Lamarre, D., 23 Lamb, J., 185, 287 Lan, V. T. T., 168, 171 Landreau, C., 143 Lane, D. P., 21 Lane, W. S., 7, 9 Langenbacher, T., 79 Lansford, R., 308, 311, 312 Laokes, D., 174, 180, 181, 191, 194 Larimer, F. W., 168, 186, 206, 230 Larsen, E., 25 Lassus, P., 108 Laszo, A., 23 Latham, K. A., 7, 9 Lau, A. Y., 6, 9–11 Lau, P. J., 155 Lauder, S., 287 Laval, J., 7 Lavrik, O. I., 21 Lawrence, C. W., 152, 167–176, 179, 180, 185–191, 205, 206, 229, 232, 281 Lawrence, N. A., 143, 158 Layton, J. C., 240 Lazebnik, Y. A., 26, 108 Le, X. C., 9, 25 Lebecque, S. G., 312 Le Cam, E., 194 Lechler, T., 168 Le Deist, F., 308 Lee, C., 311, 319 Lee, J. H., 122, 211 Lee, J. W., 149 Lee, M. S., 16 Lee, P. L., 206, 241 Lee, P. W., 114 Lee, R. M., 325 Lee, S. H., 16, 19 Lee, S. K., 18 Lee, W. I., 316, 317 Leem, S. H., 149 Lees-Miller, S. P., 189 Legerski, R. J., 43, 53, 57 Lehmann, A. R., 57, 184, 207, 216, 217, 219–221, 243, 320 Lei, K., 119 Lemeur, M., 143, 308 Lemontt, J. F., 168–170, 188 Lengauer, C., 107
352
AUTHOR INDEX
Lenne-Samuel, N., 241–243 Leonard, G. A., 13 Le Page, F., 24, 25, 194 Lerenthal, Y., 123 Lesca, C., 27 Lescasse, R., 289 Leugers, S. L., 206 Levin, D. S., 21 Levine, A. J., 125, 127 Levine, A. S., 53, 206, 207, 212, 230, 232 Levine, M., 47 Levrero, M., 117, 118 Levy, D. L., 151, 155 Levy, Y., 313, 317 Li, B.-H., 56 Li, G., 189 Li, G. M., 120 Li, J., 22, 114 Li, L., 43, 53, 57 Li, X., 6, 13, 22, 113, 114 Li, Y., 155, 156, 194, 310 Li, Y. F., 83 Li, Z., 167, 168, 171–173, 183, 187, 190, 297, 310, 316, 321, 322 Liakopoulos, D., 292 Liao, W., 325 Liao, X., 291 Liberi, G., 298 Lick, K., 151 Lieber, M. R., 22, 149, 267, 308, 310–312 Lieberman, J., 26 Lim, D. S., 111, 112 Lim, S. E., 152 Lima, C. D., 292 Lin, C., 90, 91, 93, 94 Lin, D., 322 Lin, D. P., 121 Lin, J., 112, 122 Lin, J.-J., 51 Lin, P., 189 Lin, S. L., 289, 297 Lin, W., 168, 173, 186 Lin, Y. C., 113 Lindahl, T., 1, 2, 6, 16, 18, 21–25, 145, 147, 175, 182, 211, 297, 312, 316, 322 Lindsay, H. D., 110 Lindsey-Boltz, L. A., 43, 45, 57, 64, 110 Lindsley, J. E., 248 Ling, H., 152, 153, 212–215, 233, 234, 236–238, 244, 245
Linke, S. P., 108 Linn, S., 43, 45, 53, 57, 64, 121, 151, 155, 156, 217, 219 Lippard, S. J., 61 Lipps, G., 53 Lipton, L., 26 Little, J. W., 280 Liu, D., 91, 110 Liu, J., 87–90 Liu, M., 59 Liu, Q., 113 Liu, Y., 92, 113 Livingston, A. L., 16, 26 Livingston, D. M., 23, 112 Livneh, Z., 152, 168, 194, 206, 229, 239, 245, 249–252, 256, 281 Ljungquist, S., 18, 24 Lloyd, R. S., 7, 9, 16, 19, 210 Loakes, D., 174, 180, 182, 191 Loeb, L. A., 2, 18, 27, 158 Loechler, E. L., 148, 242 Loewith, R., 189 Logie, C., 61 Lohman, P. H., 168, 171, 207, 216 Lombardo, M. J., 206, 241 Longley, M. J., 148, 152, 156, 157 Lopes, M., 298 Lopez de Saro, F. J., 242, 243 Lopez-Fernandez, L. A., 147 Lopez-Garcia, J., 28 Lorch, Y., 59 Lord, J. M., 122 Losson, R., 28 Loukili, N., 289 Louneva, N., 177, 210, 273 Lovering, R., 285 Lowe, S. W., 108, 115 Lowery, D. M., 112 Lowndes, N. F., 112 Lozano, J. J., 289 Lu, A. L., 6, 13, 16, 19 Lu, C., 249 Lu, R., 7 Lu, X., 53 Lubratovich, M., 114 Lucchini, G., 155 Lucey, M. J., 28 Ludlow, C., 168 Lukas, C., 115 Lukas, J., 106, 107, 113–115, 122
AUTHOR INDEX
Luk-Paszyc, M. J., 155, 189, 220, 275 Luna, L., 9 Lundgren, K., 114 Luo, C., 114 Luo, G., 113, 122
M Ma, L., 289 Ma, Y., 308 MacCallum, R., 152, 155 MacDonald, J., 176, 179, 191 Macdougall, E., 25 MacFarlane, A. W., IV, 79 MacGinnitie, A. J., 324 MacGlashan, D., 47 Maciejewski, M. W., 19 Madhani, H. D., 287 Madsen, P., 324 Madura, K., 287 Maekawa, T., 60 Maenhaut, M. G., 230, 233, 237, 241 Maenhaut-Michel, G., 230, 240 Maga, G., 137 Magnac, C., 317 Mahajan, K. N., 149, 310 Mahajan, R., 291, 292 Maher, E. R., 26 Maher, V. M., 167, 168, 171–173, 187, 190, 220, 221, 297 Mahoney, W., 16, 19 Majka, J., 110 Mak, T., 26 Mak, T. W., 114 Makhov, A. M., 111 Maki, H., 18, 248 Malhotra, K., 77, 81, 85, 87, 88 Malik, S., 308 Malim, M. H., 325, 326 Malone, M. E., 6 Malynn, B., 310 Mangeat, B., 326 Manis, J. P., 308, 311, 312, 318, 322 Maniwa, Y., 110 Manke, I. A., 112 Mannino, J. L., 121 Mao, C., 256 Maor-Shoshani, A., 206, 239, 249, 250, 252, 256, 281
353
Marapaka, P., 7 Marcu, K. B., 317, 323 Marenstein, D. R., 19, 25 Margison, G. P., 25 Margosiak, S., 114 Margossian, L., 27 Margot, A., 194 Marin, M. C., 117 Marini, F., 139, 143, 148, 155, 183, 185 Market, E., 312, 316, 318 Markovina, S., 23 Marodi, L., 316, 317 Marot, D., 118 Marr, M. T., 57 Marriott, D., 287 Marsin, S., 24 Marth, J. D., 143 Martin, A., 313, 315, 316, 322, 323 Martin, S., 285 Martinez, A. C., 149 Martini, E. M., 299 Mas, P., 93 Maser, R. S., 112 Masson, M., 24 Masuda, K., 189 Masuda, Y., 155, 168, 173, 189, 220, 275 Masui, R., 77, 79 Masumura, K., 25 Masutani, C., 53, 60, 61, 141, 143, 153, 180, 189, 206–212, 215–217, 219, 220, 221, 233, 239, 269, 273–275, 321 Masuyama, S., 297 Mataga, N., 79, 84, 85 Matas, D., 24 Mathews, C. K., 208 Mathis, D., 143, 308 Matsuda, T., 153, 184, 208–212, 221, 269, 271, 320 Matsui, K., 206, 230, 232–235, 237, 239–241 Matsumoto, Y., 21 Matsunaga, T., 53, 121, 168, 171 Matsunga, T., 77, 87 Matsuoka, S., 112, 113 Matunis, M. J., 292 Matuschewski, K., 292 Maymon, M., 93 Maynard, J., 26 Mayne, L. V., 44, 57 Mazumder, A., 7, 18 McAuley-Hecht, K. E., 13
354
AUTHOR INDEX
McBride, K. M., 317 McCarthy, J., 317, 323 McCormick, J. J., 171, 220, 221, 297 McCulloch, S. D., 141, 153, 210, 211 McCullough, A. K., 7, 16 McDaniel, L. D., 57 McDonald, J. P., 143, 147, 148, 184, 194, 206–210, 212, 216, 221, 230, 232, 321 McDonald, W. H., 152 McElhinny, S. A., 153 McEntee, K., 242, 249 McFadden, G., 75 McGill, C. B., 184 McGrath, J. P., 286 McGregor, W. G., 167, 168, 171–173, 187, 190 McHenry, C. S., 141, 150 McKee, R. H., 169, 170, 185, 188 McKenna, S., 284, 285, 289–291, 297 McKenzie, G. J., 206, 241 McKeon, F., 117 McKeown, C. K., 24 McManus, T. P., 167, 168, 171–173, 187, 190 McMurray, C. T., 122 McNair, S., 220 McRee, D. E., 7 Mehta, A., 325 Meijer, M., 60, 189 Meira, L. B., 102 Melchers, F., 312 Melchior, F., 291, 292 Melino, G., 117, 118 Mellon, I., 44, 56 Melo, J., 102, 110, 113, 114 Memisoglu, A., 23 Mendelman, L. V., 172, 208 Mendelsohn, M., 308 Menissier-de Murcia, J., 24, 316 Mer, G., 112 Merlo, P., 118 Merson-Davies, L. A., 6 Messmer, B. T., 323 Metheny, L. J., 325 Metzger, S., 291, 292 Meyerhans, A., 325 Miao, G., 18, 23 Michael, D., 125 Michael, H., 26 Michaels, M. L., 7 Michel-Beyerle, M. E., 79
Michelson, R. J., 102, 109 Mihara, M., 115, 116 Mikami, Y., 9, 10 Miki, K., 77, 79 Miki, Y., 23 Millar, C. B., 25 Miller, H., 177, 210, 211 Miller, J. H., 14 Miller, J. K., 13 Milligan, D., 16 Mills, M., 122 Milstein, C., 308, 316, 320, 322 Milyavsky, M., 23, 24 Mimura, J., 266 Minko, I. G., 210 Minowa, O., 25 Mirzoeva, O. K., 122 Mishiro, S., 18 Missura, M., 53 Mitchell, B. S., 149, 310 Mitchell, D. L., 60, 189, 221 Mitomo, K., 23 Mitra, S., 6, 7, 9, 18–20, 23, 192 Mittal, S., 293 Mittelman, L., 123 Miura, A., 27 Miura, Y., 267 Miyamoto, Y., 94 Miyazaki, J., 9, 25, 94, 96 Mizukoshi, T., 61, 88, 212 Mizuta, R., 308 Mo, J., 61, 64, 91 Moch, C., 289 Mochan, T. A., 112 Mochida, S., 112 Mockler, T., 93 Modrich, P., 156 Moerschell, R. P., 180 Moggs, J. G., 53, 55, 60 Mol, C. D., 7, 10, 15, 19, 20, 110, 192 Moldovan, G. L., 155, 187, 219, 243, 274, 288, 292–294, 297, 298, 300 Molina, J. T., 21 Moll, U. M., 115, 116 Moller, S. G., 93 Mombaerts, P., 308 Moncalian, G., 122 Monden, Y., 25 Monick, M. M., 23 Moolenaar, G. F., 51
355
AUTHOR INDEX
Moore, D. H., 26 Moorthy, N. C., 23 Moraes, T. F., 284, 285, 289–291 Morales-Ruiz, T., 27 Moras, D., 232 Moreau, P. L., 248 Morelli, C., 168 Morgan, I. M., 220 Morgan, S. E., 118 Morgan, W. F., 122 Mori, K. J., 323 Mori, S., 113 Mori, T., 53, 121 Morikawa, K., 9, 10, 234, 235, 239 Morin, L., 289 Morioka, H., 53, 121 Morita, R. Y., 240 Moriya, M., 6, 177, 210, 211 Moriyama, K., 174, 180, 191 Morland, I., 9 Morrison, A., 168 Morrison, D. K., 119 Morrison, J. R., 323 Morrow, J. S., 112, 114 Moshous, D., 308 Mossmann, H., 143 Moustacchi, E., 168–170 Moyal, L., 123 Mu, D., 43, 53, 55, 59 Mueller, M., 93 Muijtens, M., 9, 25, 94 Mukhopadhyay, D., 324, 325 Mullen, G. P., 19 Mullenders, L. H., 44, 57, 216, 217 Muller, J. G., 6 Muller, S., 24 Muller, W., 316 Mungall, A. J., 168 Murakumo, Y., 155, 168, 180, 187, 189, 220, 275 Muramatsu, M., 182, 311, 313, 315, 317, 323 Murante, R. S., 167, 171–173, 190 Murli, S., 250 Murphy, M., 111 Murre, C., 323 Murthy, K. G., 23 Musacchio, A., 155 Muto, T., 313, 317 Muzi-Falconi, M., 298 Myers, P., 114
Myers, T. W., 173, 190 Myrnes, B., 6 Myung, K., 155
N Nadal, M., 289 Naderi, S., 108, 116 Nadji, S., 81, 82 Naegeli, H., 53, 63 Nagaoka, H., 317, 323 Nagawawa, H., 186 Nagendran, V., 316, 317 Nairn, R. S., 43 Najarian, M. T., 27, 170, 193 Naka, K., 114 Nakabeppu, Y., 25 Nakada, S., 119 Nakagawara, A., 267 Nakai, S., 25 Nakamura, H., 88 Nakanishi, M., 114 Nakata, A., 248, 281 Nakatani, Y., 122 Nakatsuru, Y., 266 Nakayama, K., 23 Nambu, Y., 319, 323 Nanjangud, G., 323 Naoi, Y., 168, 171, 186 Napolitano, R., 239, 241, 242, 250 Nash, H. M., 7, 9 Nash, R. A., 24, 297 Natarajan, A. T., 44, 57 Navaratnam, N., 323, 324 Navarro, S., 267 Navas, T. A., 155 Nealon, K., 21 Nebel, S., 118 Nebert, D. W., 265 Negishi, H., 116 Negishi, K., 174, 180–182, 191, 194 Negrini, M., 168 Nehme, A., 118 Nelson, J. L., 206 Nelson, J. R., 167, 171–176, 179, 188–191, 205, 232 Neuberger, M. S., 143, 157, 182, 183, 185, 194, 312, 315, 316, 319, 320, 322, 325, 326
356
AUTHOR INDEX
Neumeister, P., 323 Neuwald, A. F., 112 Newmeyer, D. D., 115, 116 Ng, H. H., 15 Ng, J. M. Y., 53 Ng, W. O., 77 Nguyen, A., 112 Nguyen, B., 114 Nguyen, D., 157, 168 Nguyen, T. D., 60 Niall, H. D., 283 Nicholl, I. D., 21 Nichols, A. F., 53 Nicholson, D. W., 26 Nick McElhinny, S. A., 149, 310 Nicolas, A., 194 Niedergang, C., 24 Nielsen, H., 25 Nikaido, O., 53, 77, 87, 121 Nilsen, H., 16, 25, 182, 312, 322 Nimnual, A., 120, 122 Ninio, J., 47 Nishijima, M., 267 Nishimura, S., 25 Nishishita, T., 18 Nissen, K. A., 60 Nisson, P. E., 168, 169, 171, 187 Niwa, H., 94, 96 Noda, T., 25 Nohmi, T., 25, 152, 168, 206, 229, 230, 232–235, 239–242, 248 Noll, D. M., 13 Nomura, T., 77, 87 Nonoyama, S., 316, 317 Nookala, R. K., 247 Norbury, C., 152, 155 Norman, D. P., 9, 11, 12 Noskov, V. N., 174, 180, 191, 194 Notarangelo, L. D., 313, 317 Novina, C. D., 26 Nowicka, A. M., 173–175, 179, 188, 191 Nuber, U., 285 Numata, S.-I., 155, 168, 187, 189, 275 Nunoshiba, T., 192, 193 Nurse, P., 155 Nussenzweig, A., 308, 312 Nussenzweig, M. C., 308, 312, 317, 318, 320 Nyberg, K. A., 102, 109
O Oakeley, E. J., 27 O’Brien, P. J., 11, 17, 45 O’Brien, T., 169, 170 Ocampo, M. T., 25 Ochs, H. D., 316, 317 Ochs, R. L., 112 O’Connell, M., 112 O’Donnell, M., 139, 150, 153, 206, 221, 235–237, 239, 241–243, 245, 249–252, 256, 273, 281 Oettinger, M. A., 308 Offer, H., 23, 24 Ogata, E., 18 Ogawa, O., 271 Ogi, T., 206, 266, 267, 269, 271, 273 O’Grady, P. I., 191 Ogura, Y., 189, 275 O’Handley, S. F., 7 Ohashi, E., 180, 189, 206, 210, 211, 220, 239, 269, 271, 273, 275 Ohkuma, Y., 60 Ohmae, M., 189 Ohmori, H., 152, 168, 177, 180, 189, 206, 210–212, 220, 229, 230, 233, 237, 239, 241, 266, 267, 269, 271, 273–275 Ohtsuka, E., 9, 10 Okada, A., 308 Okada, H., 114 Okada, T., 168, 185, 271 Okada, Y., 114 Okamoto, T., 53 Okamura, T., 79, 84, 85 Okazaki, I. M., 315, 323 Okazaki, T., 18 Okey, A. B., 265 Okumoto, D. S., 44, 56 Okuto, H., 174, 180, 191, 194 Olieric, V., 243, 245, 246 Oliver, J., 24 Ollis, D. L., 139 Olmsted, E. A., 143, 158 Olsson, C., 311 Olsson, M., 6 Oltz, E., 308, 310 Ong, P., 53 Onrust, R., 252 Ookuri, T., 186 Opitz-Araya, X., 108
AUTHOR INDEX
Opperman, T., 250 Oppezzo, P., 317 Orban, P. C., 143 O’Reilly, N. J., 285 Oren, M., 115–117, 125 Orkin, S., 310 Orntoft, T. F., 26 O’Rourke, E. J., 14, 17 Orren, D. K., 47, 49, 51 Oryhon, J., 293 Osborn, A. J., 110 O’Shea, C., 64 Osley, M. A., 287, 299 Ossovskaya, V. S., 189 Otevrel, T., 310 Otoshi, E., 87, 89 Otsuka, C., 174, 180–182, 191, 194 Ouellette, L. M., 220 O-Wang, J., 266, 267 Owen, B. A., 122 ¨ zer, Z., 81 O ¨ zgu¨r, S., 91, 93 O Ozkaynak, E., 283
P Paciotti, V., 155 Paciucci, R., 289 Page´s, V., 153, 155 Pakrasi, H. B., 77 Pal, B. C., 6 Pan, P. Y., 18, 23 Panaitescu, L., 171 Panayotou, G., 10 Pancoska, P., 115, 116 Pandey, P., 118 Pannicke, U., 308 Pao, A., 265 Papadimitriou, K., 110 Papaioannou, V., 308 Papavasiliou, F. N., 312, 316, 318 Pappin, D. J., 112, 285 Parikh, S. S., 9, 10, 17, 19 Park, C.-H., 53 Park, H. J., 81, 82 Park, H. W., 77, 78, 81 Park, J.-S., 57 Park, M. S., 21 Parker, A., 16, 19
357
Parlanti, E., 21 Parraga, M., 147 Parris, T., 121 Parsons, J. L., 26 Parsons, S. H., 26 Partch, C. L., 94 Parwaresch, R., 317, 323 Pascucci, B., 21 Pasqualucci, L., 323 Pastink, A., 168, 171 Pastushok, L., 284, 285, 289–291, 297 Pata, J. D., 153, 212, 213, 233, 234 Patel, D. J., 13, 63, 323 Paterson, M. C., 207, 216 Patil, C., 308 Paull, T. T., 122, 308 Paulovich, A. G., 189 Paunesku, T., 293 Pavletich, N. P., 285, 291 Pavlov, Y. I., 151, 174, 180, 184, 191, 221, 320 Payelle-Brogard, B., 317 Payne, G., 77, 79, 83–85 Paz-Elizur, T., 245 Pearl, L. H., 10, 245, 247, 252 Pedersen, L. C., 147 Pediconi, N., 118 Pelanda, R., 308, 312 Pellicioli, A., 155, 298 Pennell, R. I., 27 Pepper, E. D., 206, 240 Perricaudet, M., 118 Perrin, L., 326 Perry, J. R., 168, 186, 206, 230 Persinger, J., 61 Peters, A., 308, 311, 312, 318 Petersen-Mahrt, S. K., 157, 315, 319, 322, 325 Peterson, C. A., 53, 61 Peterson, C. L., 291 Petes, T. D., 291 Petit, C., 52, 94 Petrenko, O., 115, 116 Petrini, J. H., 106, 112, 122 Petruska, J., 172, 208, 221, 312, 318–320 Petti, A. A., 93 Pfander, B., 155, 187, 219, 243, 274, 288, 292–294, 297, 298, 300 Pfeifer, G. P., 265 Pham, P., 152, 153, 156, 212–214, 221, 233, 235–237, 239, 242, 249–252, 255, 256, 281, 312, 316–320
358
AUTHOR INDEX
Philippe, N., 308 Philipsen, A., 317, 323 Phillips, A. M., 51 Phillips, B., 195, 219, 274 Phillips, R. K., 26 Phoenix, F., 28 Phung, Q. H., 316, 322 Picard, D., 118 Picher, A. J., 147, 149 Pickart, C. M., 183–185, 286, 287, 289, 290 Pierce, A. J., 156 Piersen, C. E., 7, 19 Pietrokovski, S., 51 Pigatto, F., 26 Pillaire, M. J., 27 Pinaud, E., 311, 312, 316, 318 Pisani, F. M., 233–235, 239 Plante, D. T., 94 Plebani, A., 313, 316, 317 Plevani, P., 155, 298 Plosky, B. S., 143, 153, 212, 214, 221, 233, 234, 243, 321 Poch, O., 232 Podust, V., 156 Poirier, G. G., 23, 26 Polanowska, J., 122 Pomerantz, R. J., 326 Pommier, Y., 18, 26 Ponamarev, M. V., 157 Ponten, I., 239, 241, 273 Poon, K., 274 Popoff, I. J., 119 Popoff, S. C., 18 Porter, A. C., 108, 116 Posnick, L. M., 27, 170, 193 Potapova, O., 119, 237 Pourquier, P., 18 Pouryazdanparast, P., 241 Pouyet, J., 23 Povirk, L. F., 149 Pradhan, D., 112, 114 Prakash, L., 18, 45, 152, 155, 168, 169, 172–178, 185, 190–195, 206–212, 214, 215, 217, 219, 229, 233, 234, 243, 271–274, 285, 287, 288, 291, 293, 294, 297, 298 Prakash, S., 18, 45, 152, 155, 168, 172–178, 185, 190–192, 194, 195, 206–212, 214, 215, 217, 219, 229,
233, 234, 243, 271–274, 285, 287, 288, 291, 293, 294, 297, 298 Prasad, R., 19, 21, 147, 148, 208 Preston, B. D., 2, 18, 143, 158 Price, D. H., 59 Prince, M. A., 7 Pritsch, O., 317 Prives, C., 23, 24, 117 Protic, M., 293 Ptak, C., 289–291 Pu, H., 27 Puri, P. L., 118 Putnam, C. D., 9, 17, 102, 109 Pyrowolakis, G., 155, 187, 219, 243, 274, 288, 292–294, 297, 298, 300
Q Qiao, Y., 230, 239 Qin, X. F., 308, 312 Qiu, C., 15 Qiu, J., 21, 22 Quah, S.-K., 170 Quong, M. W., 323
R Raams, A., 217, 219 Rabinovich, N., 212, 230, 232 Rada, C., 182, 312, 316, 320, 322 Radic, M., 312 Radicella, J. P., 14, 17, 19, 24 Radman, M., 206, 207 Rahman, D., 112, 285 Rai, K. R., 323 Rajagopalan, K. V., 77 Rajagopalan, M., 242, 249 Rajewsky, K., 143, 183, 185, 271, 308, 312, 316 Rajpal, D. K., 175, 176, 190, 206 Ramiro, A. R., 317, 318, 320 Ramotor, D., 192, 193 Ramsden, D. A., 149, 153, 310 Randall, S. K., 212, 230, 232 Randrianarison, V., 118 Rangarajan, S., 156 Rangel, R., 317, 323 Rapic-Otrin, V., 53, 207, 208, 216
359
AUTHOR INDEX
Rasio, D., 155, 168, 187, 189 Rasmussen, H. H., 324 Rasmussen, L. J., 27, 170, 193 Raspaglio, G., 21 Rathbun, G., 308, 310 Rathi, A., 121 Rattray, A. J., 184 Raya, A., 267 Raynaud-Messina, B., 27 Reagan, M. S., 57, 59 Reardon, J. T., 43, 48, 52, 53, 55, 56, 63, 64, 81 Reback, P. B., 287 Rebollo, J. E., 248 Rechkoblit, O., 153, 173, 177, 178, 210, 273 Recht, J., 287, 299 Reddy, M. S., 194 Register, J., 114 Reha’k, M., 23 Reichlin, A., 308, 312 Reina-San-Martin, B., 317 Reinberg, D., 53, 59 Reinbolt, J., 243, 245, 246 Ren, Y., 81, 82 Reppert, S. M., 93, 94 Reuven, N. B., 206, 249–252, 281 Revert, F., 267 Revy, P., 313, 316, 317 Reynaud, C. A., 143, 147, 157, 221, 310, 320, 321 Reynolds, P., 297 Riabowol, K., 189 Richardson, J. A., 267, 274 Richmond, E., 291 Rickman, D. W., 94 Rieger, R. A., 7, 9, 14 Roberts, C. J., 168 Roberts, J. D., 173, 190 Roberts, J. W., 57 Roberts, R., 241 Roberts, R. J., 81, 88 Roberts, V. A., 75, 77, 91 Robertson, K. A., 26 Robins, P., 25, 211 Robson, C. N., 19 Robzyk, K., 287, 299 Roche, H. R., 170 Roches, H., 192, 193 Rodriguez, M. S., 112, 147, 292 Roe, S. M., 245, 247, 252
Roegner-Maniscalco, V., 248 Roest, H. P., 297 Rogers, S. G., 18 Rogers-Fani, P., 311 Rognes, T., 9 Rogozin, I. B., 143, 184, 212, 221, 312, 318, 320, 321 Roldan-Arjona, T., 27 Rolink, A., 312 Rolli, V., 24 Rolseth, V., 9 Romeijn, R. J., 168, 171, 185–187 Ropp, P. A., 139, 148, 149 Rosbash, M., 93, 94 Rosenberg, S. M., 206, 241 Rosenfeld, N., 125, 127 Rosette, C., 119 Rosewell, I., 25, 143, 183, 185 Rossi, O., 21 Roth, J. R., 241 Roth, T., 155, 168, 187, 189 Rothman, P., 311 Rotter, V., 23, 24 Rouse, J., 110 Roush, A. A., 206, 207 Roy, G., 23 Roy, R., 7, 23 Rudolph, F. B., 325 Ruetsch, N., 312 Ruhland, A., 169 Ruiz, J. F., 147, 149 Rupert, C. S., 79 Rupp, D., 43, 49 Rupp, W. D., 280 Russell, L. D., 267, 274 Russell, P., 152 Ryan, K., 114 Ryo, H., 77, 87
S Sabariegos, R., 147 Sabatino, R. D., 173, 190 Sablitzky, F., 311 Saijo, M., 122 Saito, S., 114 Saitoh, S., 152 Sakai, D., 60
360
AUTHOR INDEX
Sakai, T., 114 Sakai, W., 168, 171, 186 Sakamoto, A., 168, 171 Sakiyama, S., 266, 267 Sakumi, K., 25 Sakuraba, Y., 297 Sale, J. E., 168, 183, 185, 194, 315, 325 Salles, B., 27, 280 Salmon, M., 122 Samaranayake, M., 312, 318, 320 Sampson, D. A., 292 Sampson, J. R., 26 Samson, L. D., 6, 9–11, 23, 25, 27, 170, 193, 316 Sanadai, S., 174, 180, 191, 194 Sanal, O., 308, 313, 316, 317 Sancar, A., 43–45, 47–49, 51–53, 55–57, 59, 61, 63, 64, 73, 75, 77–79, 81–85, 87–94, 96, 110, 111, 121 Sancar, G. B., 52, 77, 79, 81–83, 87 Sanchez, A., 7 Sanchez, Y., 155 Sanchez-Pulido, L., 289 Sancho, E., 289 Sandell, L. L., 108 Saniger, M. L., 147 Sankaranand, V. S., 311, 313, 323–325 Sansom, O. J., 25 Sanz, F., 289 Sarasin, A., 25, 194, 217, 219, 220, 320 Sarbassova, D., 112 Sarker, A. H., 7, 9, 25 Sarkissian, T., 114 Sarrot-Reynauld, F., 316, 317 Sartorelli, V., 118 Sasaki, S., 116 Sata, K., 77, 87 Saurin, A. J., 285 Saus, J., 267 Saveliev, S., 255, 281 Savva, R., 10 Sawada, J., 122 Sawchuk, D., 308 Sawt, W., 310 Saxon, A., 311 Sayegh, C. E., 323 Sayer, J. M., 153, 177, 210, 211, 239, 241, 273 Schaffer, A., 183 Schar, P., 24, 28, 297 Scharer, O. D., 7, 9, 10
Scharff, M. D., 312, 313, 315–318, 322, 323 Scharz, K., 310 Schatz, D., 308, 316 Scheel-Toellner, D., 122 Scheffner, M., 285, 292 Schenten, D., 143, 271 Scherer, S. J., 121 Scheuermann, R. H., 248 Schiestl, R. H., 194, 298 Schlesinger, D. H., 283 Schmeits, J. L., 155 Schneider, J., 287 Schnitzlein, W. M., 75 Schonthal, A. H., 108, 116 Schrader, C. E., 322 Schreiber, V., 24 Schrock, R. D. III, 7, 9 Schuffert, A., 139, 148 Schuler, M., 115, 116 Schultz, L., 118 Schultz, M. C., 23 Schultz, R. A., 57, 267–269 Schwartz, D. C., 291 Schwartz, M. A., 119 Schwartz, M. F., 112, 114 Schwarz, K., 308 Schwarz, S., 27 Schwarz, S. E., 292 Scott, J., 323, 324 Scully, R., 112 Seawell, P. C., 7 Sedgwick, B., 6 Seeberg, E., 9, 17, 25, 193, 316 Seed, B., 310 Sehested, M., 112 Seidl, K., 310, 311 Seimiya, M., 267 Seitz, E. M., 281 Seki, M., 139, 148 Seki, S., 7, 9, 23, 25 Sekiguchi, J., 310 Sekiguchi, M., 25, 150, 248 Selby, C. P., 44, 47, 51, 52, 56, 57, 59, 91, 94, 96 Selfridge, J., 25 Sengupta, S., 117 Seo, K. Y., 242 Serwe, M., 311 Seufert, W., 292 Shafer, B. K., 184
AUTHOR INDEX
Shalitin, D., 91, 93, 94 Shall, S., 24 Shannon, K. E., 114 Sharief, F. S., 139, 148 Shaw, J., 26 Shaw, S. J., 94 Shcherbakova, P. V., 151 Sheehy, A. M., 325, 326 Shen, B., 21, 22, 110 Shen, X., 206, 221, 235, 239, 241, 242, 249, 250, 252, 255, 256, 273, 281 Sheng, W., 291 Sherman, F., 180, 286 Shi, Q., 51 Shibata, T., 77, 79 Shibue, T., 116 Shibutani, S., 6, 13, 239, 273, 274 Shikazono, N., 168, 171 Shilatifard, A., 287 Shiloh, Y., 110, 111, 123, 125 Shimizu, A., 319, 323 Shimizu, M., 233–235, 239 Shimizu, Y., 53, 266 Shimizu-Nishikawa, K., 186 Shimodaira, H., 121 Shinagawa, H., 248, 281 Shinkai, Y., 182, 271, 273, 308, 311, 313 Shinohara, A., 119 Shinoura, Y., 206, 248 Shiromoto, T., 9, 25 Shishikura, T., 267 Shivji, M. K., 143, 183, 185 Shock, D. D., 139, 208 Shoham, G., 14 Shulman, M. J., 323 Shyu, A.-B., 268 Sibghat-Ullah, 52 Siciliano, M. J., 9 Sidorkina, O. M., 7 Sieber, O. M., 26 Siede, W., 113, 141, 205–207, 265 Siegmann, M., 27 Sigal, A., 125, 127 Silbergleit, A., 185 Silvian, L. F., 152, 212–214, 233 Simhadri, S., 177 Simon, J. A., 168 Simpson, L. J., 168, 183, 185 Singer, B., 25 Singh, G., 61
Singh, K. K., 16, 19 Singh, M., 143, 158 Singhal, R. K., 21, 186 Sinha, N., 81, 82, 248 Skaliter, R., 245 Skjelbred, C. F., 25 Skuse, G. R., 325 Slayton, W. B., 158 Slechta, E. S., 241 Slupphaug, G., 10, 19, 25, 316, 317 Smerdon, M. J., 16, 59, 60, 112 Smerdon, S. J., 114, 139 Smeyne, R. J., 26 Smith, B. T., 237, 250 Smith, C. A., 7, 44, 56, 77, 87, 88 Smith, G., 308 Smith, H. C., 325 Smithies, O., 94 Smulson, M. E., 26 So, A. G., 173, 190 Sobol, R. W., 21 Sofuni, T., 230, 233, 237, 241 Sohail, A., 15, 312, 318, 320 Sonada, E., 308, 312 Song, F., 17 Songyang, Z., 112 Sonneveld, E., 185 Sonoda, E., 168, 185, 271 Sossou, M., 24 Soulier, J., 112 Soustelle, C., 194 Sowden, M. P., 325 Spadari, S., 137 Spangrude, G. J., 158 Spence, J., 286 Spielmann, H. P., 63 Spira, A. I., 18 Spitz, D. R., 23 Spivak, G., 44, 56 Spoonde, A. Y., 26 Spooner, E., 7, 9 Spyracopoulos, L., 289–291 Srinivasan, V., 75 Srivastava, D. K., 21, 148 Staden, R., 320 Staknis, D., 93 Stall, A., 308 Stamato, T., 310 Stamp, G., 25, 143, 183, 185 Stanewsky, R., 93, 94
361
362
AUTHOR INDEX
Stanley, R. J., 79 Stapleton, M. A., 51 Stark, G. R., 108, 116 Stary, A., 217, 219, 220, 320 Stasiak, A., 251, 252 Stavnezer, J., 311, 322 Stavropoulos, P., 318, 320 Stefanini, M., 57 Steinacher, R., 28 Steinborn, G., 206, 248 Steitz, T. A., 139, 153, 212, 213, 233, 234 Stelter, P., 187, 220, 243, 274, 294, 295, 297, 298 Stephen, S. Z., 322 Stern, D. F., 112–114, 155 Stewart, G. S., 112 Stewart, J., 243 Stewart, V., 308 Stivers, J. T., 2, 6, 14, 16, 17 Stoiber, D., 116 Stoica, B. A., 26 St Onge, R. P., 110 Storb, U., 182, 308, 311, 312, 318 Storck, S., 147 Stracker, T. H., 122 Strahl, B. D., 61 Strathern, J. N., 170, 184 Strauss, B. S., 241 Strominger, J., 308 Stubbe, J., 7 Stucki, M., 21, 112 Studwell-Vaughan, P. S., 242 Stukenberg, P. T., 242 Stumpf, J. D., 206 Suarez, M., 206, 207 Sugai, M., 311, 313, 319, 323 Sugasawa, K., 53, 60 Sugaya, E., 186 Sugino, A., 149 Suh, H., 308, 312 Sullivan, M. J., 108, 116 Sumii, M., 168, 173 Sun, B., 7, 9 Sun, G., 119 Sun, Q., 51 Sun, X., 23 Sun, Y., 310 Sun, Z., 112, 114 Sung, P., 119, 185, 194, 287, 288 Sunkara, S., 9
Sunnerhagen, P., 110 Suo, Z., 234, 236, 237 Suski, C., 139 Sutton, M. D., 250 Suzuki, H., 114 Suzuki, N., 239, 273, 274 Sved, J., 14 Svoboda, D. L., 43, 55 Swann, P. F., 19 Sweasy, J. B., 248 Syvaoja, J. E., 151 Szankasi, P., 168 Szyklo, T. E., 114
T Ta, V.-T., 317 Taccioli, G., 308, 310 Tada, Y., 266 Tagawa, M., 266, 267 Tainer, J. A., 7, 9, 10, 15, 17, 19–22, 75, 77, 91, 110, 192 Takahashi, J. S., 94 Takahashi, M., 168, 173, 189, 275 Takahashi, Y., 266, 292 Takai, H., 114 Takano, H., 25 Takano, R., 94 Takao, M., 9, 25, 94 Takaoka, A., 116 Takata, M., 168, 183, 185, 194, 271, 315 Takeda, S., 168, 183, 185, 194, 271, 315 Takemori, H., 77, 87 Takenaga, K., 267 Takeshita, M., 6, 194 Takio, K., 206, 207, 217 Tamada, T., 77, 79 Tamai, K., 112, 113 Tan, C.-K., 173, 190 Tanaka, A., 168, 171 Tanaka, H., 110 Tanaka, K., 122, 271, 273 Tang, J., 64, 119, 122 Tang, M., 206, 221, 235, 239, 242, 249, 250, 252, 256, 265, 281 Tang, R. H., 92 Taniguchi, T., 116 Tarone, R., 322 Tashiro, J., 311, 317
363
AUTHOR INDEX
Tateishi, S., 168, 185, 271, 297 Taylor, A. M., 112 Taylor, E. R., 220 Taylor, I. A., 112 Taylor, J. M., 323 Taylor, J.-S., 7, 77, 81, 82, 87–90, 153, 173, 175, 176, 178, 190, 210, 221, 235, 239, 242, 249, 250, 256, 273 Taylor, R., 21 Taylor, S., 323 Taylor, W. R., 108, 116 Tchaiko, P., 322 Tchou, J., 7 Teebor, G. W., 19, 25 Tempczyk-Russell, A., 114 Teng, B. B., 323, 325 Terada, T., 316 te Riele, H., 316 Terrados, G., 149 Texido, G., 316 Tezcan, I., 308, 313, 317 Thayer, M. M., 7, 9 Theis, K., 152 Thelander, L., 152 Thelen, M. P., 110 Therese, S. M., 94 Thiry, S., 27 Thoma, F., 60 Thomas, D. C., 173, 190 Thomas, H. J., 26 Thompson, C., 94, 96 Thompson, C. B., 157 Thompson, C. L., 91, 94 Thompson, L. H., 23, 24, 55 Thomson, J. B., 13, 292 Thomson, T. M., 289 Thornberry, N. A., 26 Thresher, R. J., 51, 91, 94, 96 Tian, M., 311, 312, 316, 318 Tini, M., 28 Tissier, A., 147, 148, 184, 194, 206–210, 216, 221 Tjian, R., 47 Tobias, J. W., 287 Toczyski, D. P., 102, 108, 109, 110, 113, 114 Toda, T., 152, 155 Todo, T., 75, 77, 87–91, 94, 96, 168, 206, 212, 229 Toh, E. A., 292 Toh, H., 75, 77, 87, 91
Toh-e, A., 292 Tokushige, H., 217, 219 Tomer, G., 249, 250, 256 Tomicic, M., 19, 23, 26 Tominaga, Y., 25 Tomkinson, A. E., 21, 149 Tomlinson, I. P., 26 Tommerup, N., 324 Tompkins, J. D., 206 Tonegawa, S., 308 Tong, A., 287 Torchia, J., 28 Tordo, N., 232 Torpey, L. E., 172 Torres-Ramos, C. A., 243, 293, 294 Toth, E. A., 152, 212–214, 233 Tournier, C., 120, 122 Trincao, J., 152, 212, 214, 233 Tripathy, D. K., 75 Tritt, R., 26 Troelstra, C., 57 Trono, D., 326 Trowsdale, J., 285 Trucco, C., 24 Truong, T., 119 Tsai, K. Y., 117 Tseng, H. M., 149 Tsui, C., 290 Tsuji, M., 230, 239 Tsunasawa, S., 180 Tsuzuki, T., 25 Tsvetkov, L. M., 113, 114 Tucker, J. D., 24 Turelli, P., 326 Turley, H., 26 Turner, J., 206, 249, 281 Turner, T. K., 120, 122
U Udell, C. M., 110 Ueda, R., 212 Uematsu, N., 212 Ueng, L. M., 18 Ugazio, A. G., 313, 317 Ulbright, T. M., 26 Ullah, A., 312, 318, 320 Ulrich, H. D., 155, 187, 220, 243, 274, 289, 291, 294, 295, 297–299
364
AUTHOR INDEX
Um, S. J., 28 Unk, I., 155, 175, 191, 192, 195, 219, 274 ¨ nsal-Kacmaz, K., 43, 45, 57, 64, 111 U Ura, K., 61 Urich, T., 312 Usuda, S., 18 Usui, T., 122 Utsugisawa, T., 119 Uziel, T., 123
V Vagas, E., 91 Vaillancourt, J. P., 26 Vaisman, A., 184, 194, 208–211, 214, 221, 233 Valencia, A., 289 Valentine, M. R., 235–237, 239 Vande Berg, B. J., 21, 139 VanDemark, A. P., 285, 290 van de Putte, P., 51 van der Horst, G. T. J., 9, 25, 94, 316 van der Spek, P. J., 53 Vanderwiele, D., 191 Van de Velde, J., 77 Van Gelder, R. N., 93, 94 van Gent, D. C., 168, 185, 308 van Gool, A., 57 van Gurp, C. G., 297 van Hoffen, A., 44, 57 van Houten, B., 43, 51, 63, 148 van Kesteren, M., 51 van Klaveren, J., 297 Van Komen, S., 194 van Leenen, D., 94 van Roijen, J. H., 297 van Rossum-Fikkert, S., 51 Van Sloun, P. P. H., 168, 185–187 van Zeeland, A. A., 44, 57 Varghese, A., 308 Varlet, I., 185 Varley, J. M., 114 Varshavsky, A., 283, 286, 287 Vartanian, J. P., 325, 326 Vasconcelos, Y., 317 Vassylyev, D. G., 9, 10 Veaute, X., 194, 248 Velasco-Miguel, S., 143, 267, 271, 274 Venclovas, C., 110 Venema, J., 44, 57
Venere, M., 112 Venezia, N. D., 118 Venkatasubrahmanyam, S., 287 Verbeek, S., 316 Verdine, G. L., 4, 7, 9–15, 17, 25 Vereault, A., 16 Verhoeven, E. E. A., 51 Verkaik, N. S., 168, 185 Verkerk, A., 94 Verkoczy, L. K., 183, 321 Verly, W. G., 7 Vermeulen, W., 57 Vermey, M., 297 Verselis, S. J., 114 Vidal, A. E., 19, 24, 216, 217 Vidanes, G., 113 Vijay-Kumar, S., 286 Vila, M. R., 289 Villalobo, E., 289 Villemain, J., 194 Viskochil, D., 325 Visse, R., 51 Vitaterna, M. H., 94 Vlatkovic, N., 155, 156 Vo, A., 311 Vogelstein, B., 107 Vojta, P. J., 139, 148 Volker, M., 216, 217 Volkmer, E., 110 von Borstel, R. C., 170 von Hippel, P. H., 46 Vorum, H., 324 Vousden, K. H., 117 Vreeburg, J. T., 297 Vuillier, F., 317, 320 Vuong, B. Q., 119, 122
W Wada, Y., 168, 171, 186 Wager-Smith, K., 93, 94 Wagner, J., 206, 219, 232–235, 237, 239–243, 245, 246, 249, 252 Wagner, R., 206 Wahl, G. M., 108 Wahrer, D. C., 114 Wain-Hobson, S., 325, 326 Wakasugi, M., 53, 55, 121 Wakeham, A., 114
AUTHOR INDEX
Walker, D. R., 51 Walker, G. C., 141, 168, 205, 206, 229, 230, 237, 248, 250, 265 Walker, L. J., 19 Wallace, J. D., 26 Wallace, S. S., 7, 9, 274 Walsh, C., 77, 82 Walworth, N., 113 Wang, B., 112 Wang, D., 61 Wang, F., 18, 23 Wang, G., 26, 291 Wang, H. G., 23 Wang, J., 310, 317, 323 Wang, J. Y., 108, 116–119, 121 Wang, M., 175, 292 Wang, P., 285, 291 Wang, R., 119 Wang, R. P., 77, 90, 91, 94 Wang, S. W., 152, 155 Wang, T. S., 110, 243, 275 Wang, X., 43, 108, 116, 207, 216 Wang, X.-D., 167, 168, 171–173, 187, 190 Wang, Y., 57, 121 Wang, Y. C., 221 Wang, Y. Q., 267 Wang, Z., 61, 147, 149, 151, 153, 168, 173, 175–178, 186, 190, 206, 209–211, 229, 273 Warren, A. J., 43 Washburn, R. S., 57 Washington, M. T., 172, 173, 175, 176, 190, 206, 208–211, 215, 234, 271–273 Watanabe, M., 114 Watanabe, Y., 60 Waters, T. R., 10, 19 Watkins, J. F., 288 Watson, J. D., 158 Watson, W. P., 13 Weaver, D. R., 93, 94 Webb, B. L., 281 Wedekind, J. E., 325 Weeda, G., 25, 316 Wei, C. S.-J., 177 Wei, K., 322 Wei, S. J., 23 Wei, Y. F., 77, 90, 91, 94, 289, 297 Weichselbaum, R., 118, 119 Weill, J. C., 143, 147, 157, 221, 310, 320, 321
Weinert, T. A., 102, 109 Weinfeld, M., 9, 25 Weiser, B., 206 Weis-Garcia, F., 308 Weiss, B., 2, 7, 18 Weissman, A. M., 285 Weitz, C. J., 93 Weller, S., 320 Werling, U., 121 Werner, R. M., 17 West, C. M., 108 West, M. G., 23 Westcott, S. L., 112 Weston, C. R., 119 Whitaker, L. L., 118 White, C. L., 143, 271 White, C. L. III, 267, 274 Wibley, J. E., 10 Wie, Y.-F., 168 Wiederhold, L. R., 23 Wierda, W. G., 317, 323 Wiesendanger, M., 322 Wilker, E., 114 Wilkinson, W. J., 286 Willer, D. O., 75 Williams, B. L., 114 Williams, B. R., 106, 122 Williams, G. T., 26, 182, 312, 322 Williams, K. R., 46 Wilson, D. M. III, 7 Wilson, S. H., 16, 19, 21, 24, 139, 143, 147, 148, 208 Wilson, T. E., 149 Winfried, E., 322 Wing, J., 220, 243 Winter, D. B., 184, 221, 316, 320, 322 Withka, J., 7 Witkin, E. M., 44, 56, 248 Wittenberg, C., 151 Wittschieben, B. ., 64 Wittschieben, J., 143, 183, 185 Wolberger, C., 290 Wolffe, A. P., 59 Wolgemuth, D. J., 207, 216 Woloschak, G. E., 293 Wong, E., 316 Wong, I., 13 Wong, J. A., 114 Woo, C., 322 Woo, R. A., 114
365
366
AUTHOR INDEX
Wood, A., 287 Wood, C. J., 323 Wood, L. D., 118 Wood, P., 316, 317 Wood, R. D., 22, 23, 24, 43, 53, 55, 64, 139, 143, 145, 147, 148, 183, 185, 211, 237 Woodgate, R., 143, 147, 148, 152, 153, 156, 168, 176, 179, 184, 191, 194, 205–217, 221, 229, 230, 232–239, 241–245, 248–252, 255, 256, 273, 281, 321 Woodring, P. J., 118 Workman, J. L., 61 Worthington, E. N., 77, 91, 93 Worthylake, D. K., 285, 288 Wright, C. L., 324 Wright, M., 27 Wright, P. M., 6, 13 Wu, K.-J., 59 Wu, M., 63 Wu, P., 15 Wu, X., 22, 61, 147, 149, 153, 168, 173, 175, 176, 186, 190, 206, 209, 210, 273 Wu, Y. J., 91, 92 Wu-Baer, F., 286 Wuerffel, R. A., 308, 312 Wyatt, M. D., 6, 9–11, 25 Wynshaw-Boris, A., 118
X Xanthoudakis, S., 18, 23, 26 Xiao, W., 27, 168, 280, 281, 284, 285, 288–293, 297–299 Xie, Z., 149 Xin, H., 168, 173, 186, 206 Xing, D., 7, 9 Xing, J. Z., 9, 25 Xu, B., 111, 112 Xu, J., 120, 122 Xu, L., 311 Xu, N., 268 Xu, P., 25 Xu, X., 113, 114 Xu, Y., 26, 118, 285 Xu, Z., 26 Xuong, N. G., 139 Xu-Welliver, M., 24
Y Yaffe, M. B., 112, 114 Yaffe, M. P., 286 Yagi, H., 153 Yagi, T., 168, 185 Yakovlev, A. G., 26 Yamada, A., 60, 206, 207, 215–217 Yamada, M., 206, 230, 232–235, 237, 239–241 Yamada, S., 182, 311, 313, 323 Yamaguchi-Iwai, Y., 185 Yamaizumi, M., 168, 185, 217, 219, 271, 297 Yamamoto, K., 23, 87 Yamamoto, Y., 230, 233, 237, 241 Yamanaka, S., 323 Yamashita, Y. M., 271 Yan, S., 63 Yanagida, M., 112 Yanai, H., 116 Yang, A., 117 Yang, B., 326 Yang, D., 206 Yang, D. D., 120, 122 Yang, H. Q., 92, 117, 118 Yang, I. Y., 210, 211 Yang, I.-Y., 177 Yang, K., 121 Yang, S. H., 18 Yang, W., 17, 152, 153, 212–215, 233, 234, 236–238, 244, 245 Yang, X. P., 21 Yao, M. S., 122 Yaspo, M.-L., 143, 183, 185 Yasuda, H., 292 Yasuda, S., 267 Yasui, A., 9, 25, 77, 79, 94 Yates, J. R., 152 Yeiser, B., 240 Yel, L., 316, 317 Yelent, B., 21, 22 Yin, J., 242 Yoder, B. L., 293, 294 Yodoi, J., 23 Yokoi, M., 206, 207, 217 Yokota, Y., 319, 323 Yokoyama, S., 77, 79 Yonei, S., 9, 25 Yoshida, A., 26 Yoshida, K., 23
AUTHOR INDEX
Yoshihara, K., 26 Yoshikawa, K., 311, 315, 323 Yoshioka-Yamashita, A., 121 Yoshizawa, I., 274 You, H. J., 2 Young, D., 189 Young, F., 308, 311 Young, M., 243 Yu, K., 311, 312 Yu, L., 267 Yu, S. L., 18, 215 Yu, S.-L., 177, 190 Yu, X., 93, 112 Yuan, F., 149, 168, 173, 175, 186, 206, 209, 210, 273 Yuan, W., 286 Yuan, Z. M., 118, 119 Yuasa, M., 206, 207, 217 Yudkovsky, N., 61 Yurieva, O., 243
Z Zabkiewicz, J., 25 Zacharias, W., 171 Zaika, A., 115, 116 Zajc, B., 177 Zakian, V. A., 108 Zan, H., 183, 321 Zawel, L., 53 Zeng, X., 184, 208, 216, 221, 316, 320 Zeugner, A., 93 Zgaga, Z., 194 Zhan, Q., 121 Zhang, B., 61
Zhang, C., 326 Zhang, D., 26 Zhang, H., 171, 326 Zhang, J., 311 Zhang, K., 81, 82, 311 Zhang, Q. M., 9, 25 Zhang, X., 15, 208 Zhang, Y., 149, 153, 168, 173, 175, 177, 178, 186, 206, 209, 210, 273, 287 Zhao, G. Y., 168, 185 Zhao, S., 77, 87–91, 93, 94 Zhao, X., 77, 79, 87–91, 94 Zhao, Y., 243 Zharkov, D. O., 7, 9, 14 Zheng, H., 43 Zheng, N., 285, 291 Zheng, Y., 27 Zhou, B., 212, 213 Zhou, B. B., 104, 110, 113, 114 Zhou, B. L., 153, 233, 234 Zhou, J., 24, 155, 189, 220, 275 Zhou, T., 149 Zhou, Z., 155 Zhu, B., 27 Zhu, C., 23, 310 Zhu, J., 118 Zhu, J.-K., 27 Zhu, Y., 289 Zicha, D., 216, 217 Zimmer, D., 310 Zimmerman, E., 26 Zinkel, R. A., 122 Ziqiang, L., 322 Zou, L., 110, 111 Zou, Y., 51 Zurer, I., 23, 24
367
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SUBJECT INDEX
A Abasic site, 4, 7, 8, 18 adenine removal in, 193 bypass of, 175, 181, 191–192, 193, 206, 239, 249 dAMP insertion opposite, 192 dCMP insertion opposite, 192–193 deoxyuracil removal in, 193 enzymes replicating, 175 lesions and residues of, 174–175 mutagenesis induced by, 237 mutations induced by residues of, 192 Pol IV bypass of, 239 Pol bypass of, 175, 181, 193 residues of, 174–175, 192 Rev1p bypass of, 175, 181, 191–192 UDG uracil action and, 18 Action spectrum, 85 Activation-induced cytosine deaminase (AID), 157 activity separation of, 316–317 antibody diversification and, 315 B-cell transfection of, 315 biochemical aspects of, 318–319 cancer and overexpression of, 323 cancer avoidance and SHM regulation of, 323 CLL expression of, 317 cryptic activity of, 318 CSR and activity separation of, 316–317 CSR initiation by, 317–318 CSR targeted by strand-breaks initiated by, 311–312 deamination dependent on, 318, 319–320 deficiency, 313 discovery of, 313 DNA action of, 312, 318–319 DNA breakpoints and, 312
domain structures of, 324, 325 E. coli, 317 expression regulation of, 323 gene conversion and, 315 immune specific processes and, 313 mRNA and, 315, 323–324 mutant, double expression in, 316 mutation generation and, 319 mutations, two phase, 314, 321 overexpression of, 323 RNA substrate for in vivo, 315–316 sequence specificity mechanisms of, 327 SHM and, 313–316, 323 SHM and activity separation of, 316–317 SHM and DNA action of, 318 SHM initiation by, 317–318 SHM mutations and, 183, 314, 320 SHM spectra stimulated by, 327 SHM transcription and, 312, 314 spliced forms of, 317 substrate, 327 wild-type, 317 AhR. See Arylhydrocarbon receptor AID. See Activation-induced cytosine deaminase Antibody AID and diversification of, 315 CSR, 326–327 diversity of, 308, 315 functional, 308 mutagenesis and diversity of, 308 SHM conversion of low-affinity to highaffinity, 326 AP. See Apurinic/apyrimidinic APE. See Apurinic/apyrimidinic endonuclease Apobec proteins, 323–326 antiretroviral activity of, 325–326 Apobec 1, 324–325 discovery of, 326
369
370
SUBJECT INDEX
Apobec proteins (cont.) DNA deaminases, 324 E. coli overexpression of, 325 mRNA tumor suppresser editing and, 325 retrovirus action of, 325–326 RNA deaminases, 324 Apurinic/apyrimidinic (AP), 2 BER initiation and, 145 NEIL proteins containing proteins like, 25 Apurinic/apyrimidinic endonuclease (APE), 2, 18–19, 20 cancer expression of, 26 DNA glycosylases and action of, 19 inactivation of, 26 redox activity of, 19, 20 structural biology of, 19, 20 Arylhydrocarbon receptor (AhR), 265–266 Ataxia telangiectasia-like disease (ATLD), 122 ATLD. See Ataxia telangiectasia-like disease Autonomous pathway, 106, 123, 124–127 cell death and, 125 checkpoint, 106, 124–125 p53 in, 125
B B cell GC formation in, 310 MMR, 314, 322 V(D)J recombination and maturation of, 310 Base excision DNA repair (BER), 1–28 cancer and, 27 chromatin substrate, 16 complexity of, 23 damage types repaired by, 5–7 disease and role of, 24–27 DNA glycosylase initiated, 145 E. coli gap filling during, 145 enzyme role, 27–28 enzymes, downstream, 18–23 initiation of, 2 knockout mice and, 24–27 long-patch, 147 mammalian, 23–27 MMR pathways overlapping with, 322 p53 and, 24 PARP in, 21
plant development and enzymes of, 27–28 Pol and synthesis of, 147 Pol for, 142–148 reaction, specialized and Pol function, 148 SHM and, 314, 321–322 subpathways, 143–145 synthesis, 147 uracil removal from DNA by, 316 XRCC1 binding to, 24 yeast, 147 BER. See Base excision DNA repair BRCT. See Breast cancer C-terminal domain Breast cancer C-terminal (BRCT) domain, 111–113 Burkitt’s lymphoma, 221
C Cancer, 24. See also Breast cancer C-terminal domain AID overexpression and, 323 APE expression in, 26 BER and, 27 Chk2 kinases predisposition to, 114 CS and, 45 DNA damage adaption and, 109 DNA damage resulting in, 63 DNA repair and, 101–102 genotoxic stress cell response in therapy for, 102 mismatched repair proteins and mutation rate of, 120 p53 and, 102, 115, 117 Pol and, 158 Pol and, 210–211 Pol overexpression and, 267 SHM/AID expression and avoidance of, 323 skin, 45, 158 TLS and, 168 UV-induced skin, 158 Cell cycle apoptosis and, 108 checkpoint prolongation, 107–108 checkpoints on, 104, 105, 106, 112, 140, 155, 299–300 DNA damage initiating checkpoints on, 104, 105, 106, 112
SUBJECT INDEX
eukaryotic cells and checkpoints on, 299–300 mismatched repair proteins, mutated and, 120–121 p53 withdrawal from, 116 Pol and checkpoints on, 140, 155 transcriptional repression/genes of, 108 Cell death adaptation-induced necrotic, 105, 109 AP and, 106, 125 apoptotic, 108, 115–120 catastrophic, 105, 109 clonogenic survival v. mitotic, 107–108 DNA damage response and effectors of apoptosis, 115–120 DNA damage-induced apoptotic, 116–117 fibroblasts and DNA damage induced apoptotic, 119 genotoxic stress induced, 105, 108 mitotic, 106, 107–108 necrotic, 105, 108, 109 p53-dependent, 106, 108, 116, 125, 127 passive, 107 premature senescence, 108 tyrosine kinases activated apoptotic, 118 UV-induced, 122 Chromatin ATP-dependent remodeling complexes of, 61 DNA transcription factors in, 61 protein effect of, 61 repair, 59–62 UV-induced damage distributed within, 59–60 Chronic lymphoid leukemia (CLL), 317 Circadian clock cryptochromes and regulation of, 94 function of, 94 mammalian, 94 UV light effect amelioration with, 91 Cis-syn cyclobutane dimers (CPD), 205 Pol and lesion of, 215 RAD30 bypass of, 207 UV induced, 205 clamp E. coli polymerase interaction with, 242–243, 244 mutagenesis and, 242, 248–249 Pol binding to, 242–247 Pol IV modulation by, 234–235
371
Pol V lesion bypass and, 252 TLS and, 242 Class-switch recombination (CSR), 308, 312 AID activity separation for, 316–317 AID initiation of, 317–318 AID required for, 313–315 AID-initiated strand-break targeted, 311–312 antibody, 326–327 defective, 313 domain, constant, 310–311 mechanism of, 311 MMR deficiencies and, 322 occurrence of, 309, 310–311 Pol in, 320 region specific process, 311 transcription, 309, 311–312 CLL. See Chronic lymphoid leukemia Cockayne Syndrome (CS), 44–45 Cognate clock proteins, 93 CPD. See Cis-syn cyclobutane dimers Cryptochrome, 73–96 biological world findings of, 75, 76 V. cholerae, 96 circadian clock regulation with, 94 circadian photoreception and, 94 clock function of, 94–96 cognate clock proteins bound to, 93 cytoplasm combinatorial heterodimers in cytoplasm and clock function of, 96 definition of, 73 discovery of, 77 DNA binding, 91–92 Drosophila, 93, 94 expression of, 91, 94 family, 74 function, 92–96 growth and development in plants regulated by, 91 heterologously expressed, 91 human identified, 90, 91, 93 isolation, 91 light-independent function, 94 mammalian expression of, 94 nucleic acid binding, 93 as photoreceptor, blue light, 90, 94 photoreceptor function of, 92–93 photosensory function of, 93–94, 95 phylogenetic trees of, 74–75, 76
372
SUBJECT INDEX
Cryptochrome (cont.) plant identified, 90 structure, 91–92 Synecocytis, 91 CS. See Cockayne Syndrome CSA proteins, 57 CSB proteins, 57, 59 CSR. See Class-switch recombination Cyclobutane pyrimidine dimer, 271
D Damaged DNA-binding protein (DDB), 53 DDB. See Damaged DNA-binding protein 50 -deoxyribosephosphate (50 -dRP), 2, 5, 208 DinB. See DNA damage inducible DNA AID action and, 318–319 AID action and breakpoints of, 312 backbone cleavage of, 2 base pairs distinguished by abnormal protein of, 63 binding, cooperative of, 46–47 chromatin and transcription factors of, 61 covalent linkage formed by, 266 cryptochrome binding of, 91–92 fidelity, 279 histone interactions in, 61 initiator protein in, 63 lesion/protein binding and, 53 lesions’ interference with, 2 MutY bound to, 16 nucleosome-bound, 16 nucleotide incorporation into, 239 PCNA and metabolism of, 293 Pol catalytic activities and, 219 Pol catalytic activities and, 219 Pol crystal structure and, 214 Pol tolerance of geometric distortions of, 214 Pol synthesis to short region, 209 Pol IV and, 234 protein binding of, 46–47 protein phosphorylation and initiation of response to, 111 SHM and AID action with, 318 substrates for duplex, 15–16
T:G mismatches in, 14–15 uracil residues in, 6 Y family Pol and replication of, 229 DNA bases alkylation of, 6 canonical, 6 hydrolytic deamination of aminecontaining, 5–6 oxidation of, 6 ultraviolet light-induced damage to, 6–7 DNA binding ATP-dependent, 47 DDB protein, 64 kinetic proofreading, 47–48 molecular mechanism of, 47 protein, 121–122 DNA damage abasic residues as, 174–175 adaption to, 105, 108–109, 125 adaptor proteins and signaling complexes in, 111–113 adjacent damaged bases converted to dinucleotide adduct, 74, 87 AP and repair activation of, 124 apoptosis induced by, 115–120 arrest response to, 64, 106, 107 avoidance process of, 281 biological responses to, 104–109 cancer and adaption to, 109 cell-cycle checkpoints and sensor of, 188–189 cell-cycle checkpoints initiated on, 104, 105, 106, 112 cellular effects, protection from of, 220–221 cellular response to, 102 detection specificity and, 52 DNA glycosylase recognition of, 10–16 DNA lesion level as cell response to, 102 DNA lesions and signaling in, 120–123 DNA replication and, 152 effector kinases and responses to, 113–114 eukaryote recovery and, 300 eukaryotic cell cycle arrest and, 64 fibroblasts and apoptosis induced by, 119 genome, 307 glycosylase recognition of, 10–15 irreversibility, 107–108 mechanism of, 114–115
SUBJECT INDEX
mitosis blocked by, 106, 107 mm2 mutant and, 284, 289 molecular components for initiating response to, 109–115 necrosis induced by adaptation to, 105, 109 NER elimination of, 43, 44 9-1-1 complex, 110 objectives in field of response to, 127–128 p53 activated by, 106, 107 p53 and response of, 115–117 6-4 photolyase repair of, 88 photolyase repair of UV-induced, 73, 74 PIKK family of protein kinases and, 110–111 Pol bypass of UV-induced, 273 Pol for replicating, 145, 152–155 Pol and, 273 Pol integration in responses to, 189–190 protein kinases activated by UV-induced, 111 proteins complexes induced by, 111 proteins recognizing, 53 PRR mutants and, 288 RecA and cellular response to, 248 RecA as sensor of, 281, 282 recognition, 43, 45–48 repair pathways, 141–142 response coordination, temporal to, 104–109 response coordination to, 101–128 responses, 102–120, 123–127, 189–190, 248 early, 107 immediate-early, 104–106 late, 107 results of, 63 REV1p and tolerance of, 188 S phase, 152 SAPK activated by, 119 search process in, 15–16 sensor, 53, 110, 123, 188–189 signal transduction, 103, 109–110 signaling and repair proteins in, 120–123 signaling mechanism, 104, 105, 120–123 SOS Pol response after, 280–281 S-phase in response to, 106 temporal coordination of responses to, 123–127 thermodynamic destabilization by, 63
373
tolerance, 188, 279, 280 transcribed strand repair, 56, 57 tyrosine kinases and response of, 118 UV-induced, 111 in vitro repair, 61 DNA damage inducible (dinB), 230 biochemical properties of, 232–239 mutagenesis pathway dependent on, 237 phenotype, 230 Pol activity of, 233 Pol IV as gene product of, 230–248 Pol IV encoded by, 232–234, 239 UmuC-like protein sequence homology shared with, 230–232 DNA glycosylases, 2, 4, 7–17 AAG structural family of, 9 activity, 18 adenine, 26 amine nucleophile of, 4, 5, 7, 14 APE action and, 19 BER initiated by, 145 bifuctional, 2, 4, 5, 7, 16, 18, 23 catalysis of, 16–17 cytosine recognition in, 13 damage recognition with, 10–16 end-product inhibited, 16 HhH-GPD superfamily of, 9 intrahelical recognition and removal mode of, 11 lesions acted on by, 15 mechanic classes of, 7–9 monofunctional, 2, 7, 18 MUG, 10 MutM structural family of, 9, 11–13 MutY recognition complex for, 13, 14, 15, 16 oxidative lesions repair with bifunctional, 23 plant development and, 27 recognition complexes for, 4, 10–15 reducing agent in bifunctional, 4, 7–8, 12 structure of, 8, 9 substrate binding, 16 substrate recognition, 14 T:G mismatches acted on by, 14–15 thymine removal from, 10 UDG catalysis of, 17 UDG structural family of, 9, 11–12 uracil removal from, 10
374
SUBJECT INDEX
DNA lesions, 123 detection, 120 DNA damage cell response and level of, 102 DNA damage signaling and, 120–123 DNA replication through, 205 6-4 photolyase as UV light-induced, 86 Pol cognate, 273–274 Pol substrates and, 176–177 removal, 43 repair, 56 sensor, 123 signal transduction pathway and, 109–110 UV light-induced, 86 DNA nucleobases, 1–2 DNA polymerase (Pol), 19 8-oxoG lesion bypass by, 210 Apobec proteins and deaminases of, 324 BER, 142–148, 145–147 Burkitt’s lymphoma and, 221 cancer and, 158 catalytic subunits, 139, 141 clamp binding by, 242–247 crystal structure of Y family, 234 dinB and activity of, 233 diversity, 139 DNA, 2–3 DNA damage replication with, 145, 152–155 DNA mismatched repair, 156 DNA repair and, 137 DNA replication and, 137, 150–151 DNA replication restart and, 155–156 dRP lyase activity, 138, 147 dysfunction, 157–158 E. coli, 137–139, 242–243, 244 error-prone, 145, 221, 229, 273, 312, 314 families, 137–139, 140, 142 functions of, 137–158 homologous recombination and, 155–156 human, 137–139, 148–149, 157 human health and mutations in, 157 immune system, 156–157 interstrand cross-link repair, 148–149 lesion bypass and switch of replicative and specialized, 255–257 lesion bypass by, 153, 154, 205, 255–257 NER, 143
NHEJ double-strand break, 149 nucleotide transfer reaction and Y family, 233–234 PAD and Y family, 214 PCNA function while bound to, 293 PCNA modifications and recruit of, 187 Pol and Pol aligned to, 212, 213 Pol and Y family, 269 Pol and REV1p in, 172 polymerization of, 139, 156 RAD30 protein and, 207 repair, 2–3 replicative, 219, 232 Rev and Y family, 220 Saccharomyces cerevisiae, 137–139 SHM transcription and error-prone, 312, 314 sister chromatid cohesion, 151–152 SOS, 242 structure/composition, 139–141 subdomains of catalytic subunits in, 139, 144 substrate preferences, 139–141 switch model, 249 TLS, 144, 146, 152–153, 249 TLS in vivo participation in, 241 UV-induced DNA damage bypass by error-prone, 273 in vitro measure of, 270 Y family, 144, 153, 206, 208, 214, 220, 221, 229, 232, 237–238, 269 yeast strain encoded, 151 DNA polymerase V (Pol V), 248–257 biochemical properties, 250–255 cellular, 230 clamp and lesion bypass by, 252 E. coli, 229–257 lesion bypass by, 250, 252–255 RecA protein filament contact with, 252, 254 studies, 250–251 TLS patch production by, 256 UmuDC protein and, 248–257 DNA polymerase IV (Pol IV) abasic site bypass by, 239 biochemical properties of, 232–239 cellular, 230 clamp modulation of, 234–235 dinB gene, 230–248 DNA and, 234
SUBJECT INDEX
DNA primer template interaction with, 245, 247 DNA replication by, 239 dNTP-stabilized misalignment and, 237, 238 E. coli, 229–257, 240–241 E. coli and induction of, 240 extension, mismatched by, 236 fidelity of, 235–237 incorporation, mismatched by, 236 induction of, 240 intracellular level regulation of, 240 lesion bypass by, 273–274 levels of, 240–241 little finger domain of, 245 metabolism and, 240 misinsertion capacities of, 235 mutagenesis and, 235, 241–242 mutagenesis, lesion-induced and, 237 mutational spectrum and overexpression of, 237 nucleotide incorporation catalyzed by, 233, 235 pattern of, 233, 234 replication intermediate access regulation to, 242–247 replisome, 240 subdomains of, 243, 244, 245 TLS and, 240, 241–242 in vivo expression level of, 239–240 in vivo functions, 239–242 DNA polymerase (Pol ), 205–221 active site surface area of, 214 base insertion accuracy and, 211 biochemical properties of, 207–210 cancer and, 210–211 catalytic core structure of, 212–215 cell function of, 216 CPD lesion and, 215 crystal structure of, 214 DNA and crystal structure of, 214 DNA geometric distortion tolerance by, 214 DNA replication by, 215 Drosophila, 212 error-prone, 207 extension, mismatched of, 208–210 fidelity and crystal structure of, 214 finger domains of, 213, 214, 215, 234 foci localization of, 213, 217
375
insertion by, 210 lesion bypass by, 273 little finger domain of, 213, 214, 215, 234 localization of, 215–217 mouse, 212, 215 mutational specificity of, 314, 320 nuclei with foci containing, 216–217 nucleotide incorporation in, 210, 211–212 palm domain of, 212–214, 215 PCNA interactions with, 219–220 Pol and, 212 Pol switch and, 219–220 Pol, Y family aligned to, 212, 213 regulation of, 215–217 Rev interactions with, 219–220 template-binding complex formation and, 215 three-dimensional structure of, 212, 213 thumb domain of, 213, 214, 215 TLS by, 210–212, 217, 220 truncated, 217–219 UV irradiation and cell, 216 XP variants and mutations in, 217–219 DNA polymerase (Pol ), 205–221 biochemical properties of, 207–210 cell function of, 216 DNA lesion miscoding potential and, 211 DNA short region synthesis of, 209 Drosophila, 212 duplication, 212 error-prone, 207 extension, mismatched of, 208–210 fidelity, 208 foci localization of, 213, 217 insertion by, 210 lesion bypass by, 273 localization of, 215–217 mispairs formed by, 208, 209 mouse, 212 nuclear, 217 nuclei with foci containing, 216–217 nucleotide incorporation in, 208, 210, 211–212 PCNA interactions with, 219–220 Pol and, 212 Pol switch and, 219–220 Pol, Y family aligned to, 212, 213 regulation, 215–217 Rev interactions with, 219–220 SHM and, 221, 314, 321
376
SUBJECT INDEX
DNA polymerase (Pol ) (cont.) TLS by, 210–212 UV irradiation and cell, 216 DNA polymerase IV (Pol IV), 229–257 DNA polymerase (Pol ) C2HC zinc cluster sequence of, 271 cancer and overexpression of, 267 C-terminal, 269, 271 DNA damage and, 273 DNA lesions, cognate for, 273–274 DNA repair and, 267 enzymatic properties of, 271–274 enzyme activities of, 269–270 eukaroytic homologs of, 270, 271 expression regulation of, 265–275 extension, 271–273 gene structure of, 265–271 human, 265–269 insertion, 271–273 length, 268, 269, 270 lesion bypass by, 273, 274 lesion substrates, 265–275 mouse, 265–269 NLS in, 269 N-terminal, 269, 270 p53 inactivation and overexpression of, 267 PCNA preference by, 275 Pol, Y family and, 269 promoter region of mouse, 266 protein interactions, 275 protein structure, 265–271 Rev1p and, 275 splicing variants in, 269–270 testis expression of, 267 TLS in vitro by, 271 in vivo by, 274–275 UV sensitivity of, 271–272 DNA polymerase V (Pol V) E. coli, 229–257 TLS, 281 DNA polymerase (Pol ) abasic site bypass by, 175, 181, 184, 193 B-family polymerase, 172 bypass frequency of mutant deficient, 179–180 cellular functions of, 167–195 dCMP insertion, 173–174, 179–180, 182 deoxycytidyl transferase activity, 174
DNA damage response integration of, 189–190 DNA lesions and substrates for, 176–177 double-strand break homologous repair by, 185 double-strand break repair and, 184–186 E. coli bypass by, 191 embryonic lethality and, 185 enzymatic studies with, 172–178 enzyme structure and, 195 eukaryote function and, 167 extension, 177 fidelity, 190–191 function, 190 genetic analysis, 178–182 G-template specific, 174 insertion and mutation frequency of, 194–195 insertion, lesion site of, 190 insertion step, 177–178 lesion bypass frequencies, 177, 178–179 lesion-containing template in vitro studies of, 174–178 mutagenesis induced by, 170, 176 mutation, 179–180, 194–195 nucleotide incorporation fidelity of, 173 nucleotide insertion, 178, 179, 191 oligonucleotide template studies of, 174, 180 6-4 photolyase bypass by, 179 properties of, 172–174 protein interaction and, 186–190 regulation of, 186–190 replication modes and, 187 Revp and activity of, 172, 173–174 SHM and, 182–184 SHM caused by inaccurate, 182 subunits of, 172 terminal mismatch extension efficiencies, 172–173 TLS by, 167, 171, 185 wild-type lesion bypass frequencies, 179 XP and defects in, 206 DNA repair, 46, 60–61, 298 cancer and, 101–102 cell cycle checkpoints and activation of, 103 cellular, 101 importance of, 101–102 mismatched, 156
377
SUBJECT INDEX
naked, 60–61 pathways, 141–142 Pol and, 137, 141–150, 156 Pol and, 267 process, 279 PRR and, 279 quantum yield in photolyase, 84 UV light effect amelioration with, 91 DNA repair polymerase (dRP), 19–23 long-patch repair, 3, 19–23 PARP and reaction of, 21 Pol and, 138, 147, 148 short-patch repair, 3, 19–23 DNA replication, 150–151, 298 base modifications and, 248 DNA damage and, 152 fork regression and, 155–156 homologous recombination and, 156 immune system development and, 156 lesion block of, 248 mitochondrial, 145, 152 PCNA clamp and, 110 Pol, 142, 150–151 Pol , 215 Pol IV, 239 process, 279 stalling of, 256 strand, 140, 142, 151 UDG and, 25 DNA-dependent protein kinases (DNA-PK), 118 DNA-PK. See DNA-dependent protein kinases dNTP. See Nucleoside triphosphate Drosophila cryptochrome, 93, 94 Pol , 212 Pol , 212 dRP. See DNA repair polymerase 50 -dRP. See 50 -deoxyribosephosphate
E E. coli AID, 317 Apobec overexpression in, 325 BER and gap filling of, 145 Cho, 63
clamp interaction with Pol of, 242–243, 244 DNA damage tolerance in, 280 dual incisions in, 49, 50 excinuclease, 49–51, 52 excision nuclease, 49–51, 55–56 molecular matchmakers, 47 NER, 43–65, 49–52, 143 photolyase, 78, 86, 87 6-4 photolyase, 87 Pol IV, 229–257, 240–241 Pol IV induction and, 240 Pol recognized in, 137–139, 153 Pol V, 229–257 Pol bypass of, 191 RecA protein in, 243, 299 repair pathway, 148 replicative machinery in, 256 transcription-coupled repair, 56–57, 58 transcription-independent repair in, 50 E2 proteins, 283–286 Effector kinases, 113–114 cancer predisposition of Chk2, 114 Chk1, 113–114, 123–124 Chk2, 114, 123–124 DNA damage response and, 113–114 FHA domain contained in Chk2, 114 knockout of Chk1, 113–114 knockout of Chk2, 114
F FHA. See Fork-head associated domain Fork-head associated (FHA) domain, 111–112 Chk2 kinase containing domain of, 114 interactions, 112–113
G G ! A hypermutation, 325–326 GASP. See Growth advantage in stationary phase GC. See Germinal centers Genetics, 1, 102 Genotoxic stress apoptosis activation by, 108 cell death induced by, 105, 108
378
SUBJECT INDEX
Genotoxic stress (cont.) cellular response to, 102, 103 p53 family member caused, 117–118 Germinal centers (GC), 310 Goodpasture antigen binding protein (GPBP), 267 GPBP. See Goodpasture antigen binding protein Growth advantage in stationary phase (GASP), 240
H 8-HDF. See 8-hydroxy7, 8-didemethyl-5deazariboflavin High fidelity (Hi Fi), 312, 314 HIGM. See Hyper-IgM syndrome HIV, 325–326 Human transcription termination factor 2 (TTF2), 59 8-hydroxy7, 8-didemethyl-5-deazariboflavin (8–HDF), 77, 79 Hyper-IgM syndrome (HIGM), 313
I ICL. See Interstrand cross links Ig. See Immunoglobulin gene Immunoglobulin gene (Ig), 156 high-affinity, 182 SHM induction in, 312 variant conversion of, 307 Integrative surveillance (IS), 106, 123–124, 126 lesion increase and, 123 regulatory hub of, 123 Interstrand cross links (ICL), 148–149 Ionizing radiation (IR), 116 ATM kinase activation with, 122–123 mutagenesis induced by, 169 p53 and, 127 IS. See Integrative surveillance
K Kinetic proofreading, 47–48, 52
L Lesion bypass oligonucleotide, 174, 180–181 Pol, 153, 154, 205, 250, 252–257, 273 replicative and specialized switch during, 255–257 Pol , 273 Pol , 273 Pol , 273 Pol specialized in, 255 Pol V, 250, 252–255 Pol frequencies of, 177, 178–179 RecA, 251 Rev1p frequencies of, 178–179 Lesions, 2 8-oxoG, 210 abasic residues and, 174–175 alkylated bases, 17 DNA glycosylases acting on, 15 DNA glycosylases, bifunctional repair of oxidative, 23 DNA replication blocked by, 248 excision of, 59 IS and increase of, 123 mutagenesis induced by, 237–239 NER of, 59, 63, 121 nontranscribed, 58 nucleobase, 2, 3 oxidative, 22–23, 24 oxoG, 6, 210 Pol IV and mutagenesis induced by, 237–239 Pol in vitro studies of templates containing, 174–178 Pol insertion and, 190 propanodeoxyguanosine, 177 Rev1p in vitro studies of templates containing, 174–178 Rev1p insertion and, 190 UV induced, 63, 91, 121–122 in vitro assembly of nucleosomes containing site-specific, 61 in vitro repair of oxidative, 24
379
SUBJECT INDEX
M MagI. See 3-methyladenine glycosylase I Maltose binding protein (MBP), 249 MEF. See Mouse embryo fibroblasts Methenyltetrahydrofolate (MTHF), 73, 75, 77, 78 5-methylcytosine, 14–15 Mitosis-promoting factor (MPF), 104, 106 MMR. See Postreplication mismatched repair Molecular matchmaker, 47 E. coli, 47 enzyme system use of, 52 Mouse embryo fibroblasts (MEF), 119–120 MPF. See Mitosis-promoting factor mRNA polymerase, 268 AID and, 315, 323–324 Apobec-1 and tumor suppresser editing by, 325 Apobec-1 as editing enzyme of, 323–324 editing, 315, 325 tissue expression of, 269 tumor suppresser editing by, 325 Mutagenesis abasic site induction of, 237 AID-induced, 323 antibody diversity and, 307 clamp and, 242, 248–249 dinB-dependent pathway of, 237 genetic requirements of induced, 248–250 IR induction of, 169 lesion-induced, 237–239 Pol IV and, 235, 237, 241–242 Pol induction of, 170, 176 RecA protein and, 248–249 SOS Pol and, 242 tumorigenesis and AID-induced, 323 UV light induction of, 168, 176, 237
N NEIL proteins, 9, 25 NER. See Nucleotide excision repair NHEJ. See Nonhomologous end-joining Nimejin Breakage Syndrome, 122 NLS. See Nuclear localization signal Nonhomologous end-joining (NHEJ), 149 Nuclear localization signal (NLS), 269
Nucleobases, 1–2, 3 Nucleoside triphosphate (dNTP), 212 Nucleotide excision repair (NER), 2, 43–65 damage recognition specificity with, 46 defects in, 43–44 DNA damage recognition and, 43 dual incisions in, 43, 48, 49, 50 E. coli, 43–65, 143 human, 43–65 human cell factors of, 63 kinetic proofreading, 47–48 lesion removal with, 63 mechanisms of, 48–56 Pol for, 143 repair factors in human, 49, 52–53 resynthesis, 43 SHM and, 316 SNF and, 61, 62 specificity of human, 56 steps of, 43 subpathways, 143 substrate range, 55 SWI and, 61, 62 transcription coupled, 56 transcription stimulated, 44 transcription-independent, 50, 54 UV sensitivity and, 43–44 UV-induced lesions and, 121 in vitro, 64 in vivo, 64
O OxoG. See 8-oxoguanine 8-oxoguanine (oxoG), 6 adenine paired with, 15 MutM binding to, 12–13 MutY recognition mode, 13, 14 Pol bypass of lesion, 210 Watson-Crick face of, 12
P p53, 102 activation, 106, 107, 127, 267 AP and, 125 apoptosis dependent on, 108, 116 BER and, 24
380
SUBJECT INDEX
p53 (cont.) cancer and, 115, 117 cell death and, 127 cells deficient in, 116 DNA damage activation of, 106, 107 DNA damage response and, 115–117 domain, 115 genes regulated by family of, 117 identified, 115–116 IR and, 127 mutations, 265 negative feedback regulation of, 125–127 PAH and mutations of, 265 phosphorylation, 115 Pol overexpression and inactivation of, 267 regulation, 125–127 related proteins, 117–118 self-regulatory loop, 127 transcription encoded factors and, 117 PAD. See Polymerase-associated domain PAH. See Polycyclic aromatic hydrocarbons PARP. See Poly ADP-ribose polymerase PCNA. See Proliferating cell nuclear antigen PEO. See Progressive external ophthalmoplegia Photolyase, 73–96 A. nidulans, 74 action spectrum and, 85–86 aromaticity loss and, 82 backbone distortion and, 81 binding free energy and positively charged groove in, 83 biological world findings of, 75, 76 canonical pyrimidines split by dimer radicals and, 84 catalysis, 79, 80, 83–86 crystal structure of decamer duplex and, 81, 82 cycloreversion catalyzed by, 83 Dewar valence isomer photorepair with, 90 dimer pyrimidine moieties in substrate binding by, 82 dimer splitting and, 84 dinucleotide flipping and, 83 discovery of, 77 E. coli, 78, 86 enzyme/substrate binding, 79, 83 family, 74 induced-fit mechanism and, 81
kinetic constants for reaction of, 79 monomeric proteins, 77 phylogenetic trees, 74–75, 76 prokaryotic organisms possessing, 75 quantum yield in, 84–85 quantum yield of DNA repair by, 84 reaction mechanism of, 79–86 structure, 77–79 substrate binding, 79, 81–83 substrate binding specificity of, 81, 83 T. thermophilus, 74 three-dimensional diffusion substrate binding, 81 vertebrates with, 75 in vitro, 79 in vivo, 79 6-4 Photolyase, 86–90 base-flipping mechanism, 88 binding, 87–88 catalysis, 88–90 DNA repair with, 88 E. coli, 87 enzyme-substrate complex formation and, 88 flavin to substrate electron transfer and, 89–90 photoantenna, 87 photolyases v., 90 photoproduct flipped out by enzyme bound DNA and, 88–89 Pol bypass of, 179 Rev1p bypass of, 179 three-dimensional diffusion and, 87 Photoreceptors, blue-light autophosphorylating kinase activities of, 93 cryptochrome as, 73–96, 90, 92–93 photolyase as, 73–96 plant identified, 90 Phylogenetics, 74–77 Pol. See DNA polymerase Pol . See DNA polymerase Pol . See DNA polymerase Pol IV. See DNA polymerase IV Pol . See DNA polymerase Pol V. See DNA polymerase V Pol . See DNA polymerase Poly ADP-ribose polymerase (PARP), 21, 23–24, 26
381
SUBJECT INDEX
Polycyclic aromatic hydrocarbons (PAH), 265, 266 Polymerase-associated domain (PAD), 214 Postreplication mismatched repair (MMR) B cell, 314, 322 BER pathways overlapping with, 322 CSR and deficiencies of, 322 SHM and, 314, 321–322 Postreplication repair (PRR), 279–300 DNA damage and mutants of, 288 error-free and error-prone, 298 eukaryote, 281–283, 295–297 function, 283 future directions for studies of, 297 Mms2-Ubc13-Rad5 and, 288–291 pathways, 281–283 PCNA and, 292–295, 300 PCNA covalent modifications and, 292–295 prokaryotes, 280–281 protein conjugation in, 286–292 Rad6-Rad-18 and, 286–288 RecA in, 280–281 reconstitution, 297 Ub conjugation in, 291 Ubc9-Sizl and, 291–292 Progressive external ophthalmoplegia (PEO), 157 Proliferating cell nuclear antigen (PCNA), 219 covalent modification of, 294, 295, 297–298 desumoylation, 300 DNA catalytic activities increased by, 219 DNA metabolism and, 293 eukaryotic organism conservation by, 293, 297 eukaryotic replication machinery and, 292–293 Lys63 polyubiquitination of, 298, 299 modification, 280 monoubiquitinated, 220 Pol bound, 293 Pol interactions with, 219 Pol preference of, 275 PRR and, 292–295, 300 PRR via covalent modifications of, 292–295 Rad6 -Rad18 complex targets, 288 SUMO conjugation to, 298
SUMO modification of, 292, 293, 297–298 sumoylation of, 293–295, 298 TLS interactions with, 274 Ub and covalent modification of, 297–298 Protein kinases. See also DNA-dependent protein kinases ATM, 111 ATR, 111 JNK, 119–120 p38, 119–120 PIKK family of, 110–111, 119, 123–124 stress-activated, 119–120 PRR. See Postreplication repair Pyrimidine dimer, 6–7
Q Quantum yield, 84–85 DNA repair by photolyase, 84 Fo¨rster radiationless energy transfer mechanism efficiency and, 84–85 interchromophore distance efficiency and, 85 photoantenna/catalytic cofactor energy transfer, 84
R RAD5 homologs, 295–296, 298–299 RAD30 genes CPD bypass by, 207 identification of, 206–207 Pol activity of, 207 postreplication repair pathway and, 207 template-dependent reaction and, 207 UV light and disruption of, 207 XP and, 207 Reactive oxygen species (ROS), 6 RecA protein DNA damage sensor, 281, 282 DNA damage tolerance in, 280, 281 E. coli, 299 filament, stabilized of, 252 lesion bypass and, 251 mutagenesis and, 248–249
382
SUBJECT INDEX
RecA protein (cont.) Pol V contact with filament of, 252, 254 targets, 300 Recombination signal sequences (RSS), 308 Repair proteins cancer cell mutations and mismatched, 120 cell cycle and mutated mismatched, 120–121 DNA damage signaling and, 120–123 mismatch, 120–121 MRE11–RAD50 complex, 122–123 mutated mismatched, 120–121 Rev proteins (Revp) abasic site bypass by, 175, 181, 191–192, 194 B-family polymerase, 172 cellular functions of, 167–195 chromosome aberration frequencies in, 185 competitive binding to, 189 dCMP insertion, 173–174, 178, 179–180, 182, 194, 220 deoxycytidyl transferase activity of, 174, 188 DNA damage tolerance and, 188 domain structure of, 168, 169 double-strand break repair and, 184–186 double-strand break-induced mutations of, 184 double-strand breaks and disruption of, 185–186 enzymatic studies with, 172–178 enzyme structure and, 195 enzymes recruiting, 186 eukaryote function and, 167 functions of, 170–171, 173–174, 190 functions of yeast, 173–174 genetic analysis of, 178–182 G-template specific DNA polymerase, 174 homology, 206 insertion and mutation frequency of, 194–195 insertion, lesion site of, 190 lesion bypass frequencies, 178–179 lesion-containing template in vitro studies of, 174–178 mutation, 169–170, 184, 188, 194–195
nucleotide insertion by, 179 oligonucleotide template studies of, 174, 180 6-4 photolyase bypass by, 179 Pol and, 275 Pol motifs in, 172 production levels, 187 properties, 172–174 protein interaction and regulation of, 186–190 regulation, 186–190 replication and mutant, 188 SHM and, 182–184, 183 subunits, 172 terminal mismatch extension efficiencies, 172–173 tetrahydrofuran bypass frequencies of, 181–182 TLS, in vivo and, 275 transcript level increase in, 186–187 transferase activity, 194 wild-type lesion bypass frequencies of, 179 Revp. See Rev proteins RNA polymerase, 57 AID, in vivo and substrate of, 315–316 Apobec proteins and deaminases of, 324 recognition by proxy with, 57 SHM and, 183 RSS. See Recombination signal sequences
S SAPK. See Stress activated protein kinases SCID. See Severe Combined Immune Deficiency SCN. See Suprachiasmatic nuclei Severe Combined Immune Deficiency (SCID), 111 SHM. See Somatic hypermutation Small ubiquitin-like modifier (SUMO), 280, 291 PCNA and conjugation of, 298 PCNA covalent modification and, 297–298 PCNA modification with, 292, 293 Ub charged surface and, 292
383
SUBJECT INDEX
SNF, 61, 62 Somatic hypermutation (SHM), 157, 313–323 AID activated mutation process, 314, 320 AID activity separation for, 316–317 AID and, 183, 312, 313–317, 320 AID initiation of, 317–318 AID required for, 313–315 AID stimulated spectra of, 327 AID/DNA action during, 318 antibody conversion from low-affinity to high-affinity with, 326 antibody diversity and, 307–312 antigen binding and, 307–308, 309, 310 antigen recognition and, 307–308 B-cell line frequency of, 182 biochemical perspective of, 326–327 cancer avoidance and AID expression regulation in, 323 definition of, 182 Hi Fi and, 312, 314 Ig induction of, 312 mutation process, 314, 320 mutational panacea, 307–327 NER pathway and, 316 Pol error-prone and, 312, 314 inaccurate produced, 182 Pol and mechanism of, 221 Pol and, 221 Pol and, 221 Pol and frequency of, 314, 321 Pol in, 182–183, 182–184, 221, 312, 314, 320–321 Pol mRNA levels during, 183 Rev1p in, 182–184, 183 transcript levels during, 183 transcription required by, 311–312, 313 transition mutations of, 318–319 two-phase process of, 314, 319–321 Stress activated protein kinases (SAPK), 119–120 DNA damage activated, 119 gene expression regulated by, 119 JNK as, 119–120 p38 as, 119–120 SUMO. See Small ubiquitin-like modifier Sumoylation, 279–300 PCNA, 293–295 PCNA conjugation and, 298
strategy of, 292 Ub and, 294–295 Suprachiasmatic nuclei (SCN), 94 SWI, 61, 62 Synecocytis, 91
T T. thermophilus, 74 TCR. See Transcription-coupled repair TDG. See Thymine DNA glycosylases TdT. See Terminal deoxynucleotidyl transferase Terminal deoxynucleotidyl transferase (TdT), 156–157, 308–310 3-methyladenine glycosylase I (MagI), 13–14 30 -untranslated region (30 -UTR), 268–269 30 -UTR. See 30 -untranslated region Thymine DNA glycosylases (TDG), 28 TLS. See Trans-lesion Synthesis Transcription-coupled receptors, 56–59 E. coli repair, 58 human cell repair, 60 Transcription-coupled repair (TCR), 64 Transcription-repair coupling factors (TRCFs), 44, 56–57, 60 Translesion Synthesis (TLS), 152, 205 cancer and, 168 clamp and, 242 (mis)incorporation, 205 patch, 256 PCNA interactions with, 274 Pol, 144, 146, 152–153, 155, 167, 171, 178, 185, 241, 249, 255–257 replicative and specialized switch during, 255–257 Pol , 210–212, 217, 220 Pol , 210–212 Pol in vivo participation in, 241 Pol IV and, 237, 240, 241–242 Pol in vitro, 271 Pol in vivo, 274–275 Pol V-mediated, 252, 281 Pol-clamp interaction with, 144, 155 Rev1p, 167, 178, 275 Rev1p and in vivo, 275 SOS Pol and, 242
384
SUBJECT INDEX
Translesion Synthesis (TLS) (cont.) steps of, 205 successful, 255 UmuC protein and, 249 TRCF. See Transcription-repair coupling factors Trichothiodystrophy (TTD), 45 TTD. See Trichothiodystrophy Tyrosine kinases, Abl, 118–119 apoptosis activated by, 118 cell adhesion and activation of, 119 DDB and, 122 nuclear, 118
U Ub. See Ubiquitination Ubiquitination (Ub), 279–300 cellular influence of, 283 E2 binding to, 285, 286 histone attachment to chains of poly, 287 Lys-63 conjugation of, 284, 290 mono-, 294–295 N-end rule of Rad6 encoded, 287 PCNA and poly-, 298–299 PCNA covalent modification and, 297–298 poly-, 284, 287, 289–290, 298–299 process of, 283, 284 PRR and conjugation of, 291 Rad6 encoded, 286, 287 structure, 286 SUMO charged surface and, 292 sumoylation and, 294–295 surface-exposed lysine residues and, 286 UDG. See Uracil-DNA glycosylase Ultraviolet (UV) light apoptosis induced by, 122 cancer induced by, 158 chromatin distributed damaged induced by, 59–60 circadian clock and ameliorating harmful effects of, 91 CPD induced by, 205 DNA repair and ameliorating harmful effects of, 91 lesions induced by, 63, 121–122 mutagenesis induced by, 168, 176, 237 NER and mutations of, 45 NER and sensitivity to, 43–44, 45, 121
NER correction of lesions induced by, 121 6-4 photolyase as DNA lesion induced by, 86 photolyase repair of DNA damaged by, 73, 74 Pol and irradiation of, 216 Pol and irradiation of, 216 Pol sensitivity to, 271–272 Pol and mutagenesis induced by, 176 RAD30 disruption and, 207 XP and mutation spectra of, 221 UmuC genes, 281 UmuD genes, 281 Uracil-DNA glycosylase (UDG) DNA glycosylase, 9, 11–12, 17 DNA replication and, 25 immune system and, 25 uracil action of, 18 UV. See Ultraviolet
V V(D)J recombination, 309 antigen recognition and binding and, 307–308 B cell maturation and, 310 initiation of, 308 RSS and, 308 T-cell receptor, 308 TdT and, 308–310
X Xeroderma pimentatosum (XP), 44, 158 mutations in, 217, 218 Pol defects and, 206 Pol mutations and variations of, 217–219 Pol , nuclear and, 217 RAD30 genes and, 207 SHM in, 320 UV mutation spectra in, 221 XP. See Xeroderma pimentatosum X-ray repair cross-complementing protein 1 (XRCC1), 24 XRCC1. See X-ray repair crosscomplementing protein 1