Dna Repair, Genetic Instability, And Cancer 9789812706782, 9789812700148

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DNA REPAIR, GENETIC INSTABILITY, AND CANCER

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DNA REPAIR, GENETIC INSTABILITY, AND CANCER

EDITORS

QINGYI WEI LEI LI

The University of Texas M.D. Anderson Cancer Center, Houston, USA

DAVID J CHEN

The University of Texas Southwestern Medical Center, Dallas, USA

World Scientific NEW HERSEY . LONDON . SINGAPORE . BEIJING . SHANGHAI . HONG KONG . TAIPEI . CHENNAI

Published by World Scientific Publishing Co. Pte. Ltd. 5 Toh Tuck Link, Singapore 596224 USA office: 27 Warren Street, Suite 401-402, Hackensack, NJ 07601 UK office: 57 Shelton Street, Covent Garden, London WC2H 9HE

Library of Congress Cataloging-in-Publication Data DNA repair, genetic instability, and cancer / [edited by] Qingyi Wei, Lei Li, David J. Chen. p. cm. Includes bibliographical references and index. ISBN-13 978-981-270-014-8 -- ISBN-10 981-270-014-5 1. Neoplasms--etiology. 2. Neoplasms--genetics. 3. DNA Damage--genetics. 4. DNA Repair--genetics. 5. DNA Repair-Deficiency Disorders. 6. Genetic Predisposition to Disease. I. Wei, Qingyi. II. Li, Lei. III. Chen, David. IV. Title. QH467.D164 2006 616.99'4042--dc22

2006050159

British Library Cataloguing-in-Publication Data A catalogue record for this book is available from the British Library.

Copyright © 2007 by World Scientific Publishing Co. Pte. Ltd. All rights reserved. This book, or parts thereof, may not be reproduced in any form or by any means, electronic or mechanical, including photocopying, recording or any information storage and retrieval system now known or to be invented, without written permission from the Publisher.

For photocopying of material in this volume, please pay a copying fee through the Copyright Clearance Center, Inc., 222 Rosewood Drive, Danvers, MA 01923, USA. In this case permission to photocopy is not required from the publisher.

Typeset by Stallion Press Email: [email protected]

Printed in Singapore.

SC - DNA Repair.pmd

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In memory of Dr. Lawrence (Larry) Grossman, one of the great pioneers and leaders in the field of DNA repair, who passed away on January 13, 2006

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Preface

Human DNA is constantly bombarded by endogenous (e.g. reactive oxygen species) and exogenous (UV, ionizing radiation and reactive chemicals) carcinogens. To ensure an accurate passage of genetic information onto daughter cells, cells have evolved elaborate surveillance systems and various DNA repair mechanisms that respond to the harmful stimuli and prevent damaged DNA from being converted to heritable mutations. Over the past 30 years, major frameworks have been established for major DNA repair pathways, including base-excision repair (BER), nucleotide excision repair (NER), mismatch repair (MMR), homologous recombination (HR), and non-homologous end-joining (NHEJ). Since apurinic/apyrimidinic (AP) endonuclease and uracilDNA glycosylase (UDG), the two enzymes involved in BER were discovered in Escherichia coli in the early 1970s, over 20 proteins have been identified as the core and accessory proteins of BER that primarily targets alkylated, deaminated, and oxidized bases, with a certain degree of substrate overlap with other pathways. In contrast, the NER pathway is much more versatile, and is a predominant mechanism protecting cells from UV- and chemical-induced bulky DNA lesions that are often mutagenic. To date, more than 30 genes have been identified that participate in NER. MMR, on the other hand, is an important genome caretaker system. It ensures genomic stability by correcting mismatches generated during DNA replication and vii

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recombination, suppressing homologous recombination, and triggering apoptosis of cells with severe DNA damage. In response to DNA double-strand breaks, the most dangerous lesions, HR may be used to repair the damage. HR plays critical roles in mitotic cells in repairing DNA double-strand breaks and interstrand crosslinks and in restarting replication forks blocked by DNA lesions produced by both reactive intermediates of normal cellular metabolism, exogenous chemicals, and radiation. However, NHEJ is the predominant repair pathway for removing DNA double-strand breaks in mammalian cells. To survive from lesions that block DNA replication, cells have also evolved a pathway that allows for damage tolerance or lesion bypass with high or low fidelity. A key question is how the cell cycle checkpoint machinery detects and signals the presence of damaged DNA that is embedded in millions to billions of normal base pairs. Partial answers come from recent structural and functional studies that reveal atomic details of DNA repair protein and nucleic acid interactions. The hallmark of cancer is genomic instability that may be initiated from DNA damage and faulty DNA repair systems. The pathogenesis of cancer, which is frequently an environmentally induced disease, reflects the outcome of disrupted balances among diverse biological systems, including those that govern cell growth and proliferation, signal transduction, DNA damage and repair, cell cycle checkpoint and control, and apoptosis in response to environmental insults. Yet, each individual is genetically unique and his or her responses to environmental risk factors or hazards are also unique. The role of DNA repair in the etiology of cancer has been well illustrated in several hereditary syndromes, in which an inherited defect in DNA repair and related biological processes is associated with extraordinarily high incidence of cancer. For example, patients with xeroderma pigmentosum (XP) have germline mutations in NER genes and have more than 100-fold increased risk of UV-induced skin cancers; patients with hereditary non-polyposis colon cancer (HNPCC) have a defect in MMR due to germline mutations; and patients with Fanconi anemia (FA) appear to be sensitive

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to agents that cause DNA-crosslinks and have 500-fold increased risk of developing squamous cell carcinomas of the head and neck. However, associations between inherited DNA repair defect and risk of cancer have not always been apparent in the general population. In the past 10 years, there has been a growing body of literature that begins to address this important research question at the population level. More recently, the discovery of single nucleotide polymorphisms (SNPs) in DNA repair genes has inspired a wave of association studies, some of which established a genetic basis for a suboptimal repair phenotype in the general population. These findings provide a rationale that by genetic screening for functional SNPs, it may be feasible to identify at-risk populations who can be targeted for primary prevention of cancer that has an etiology of genetically determined variation in DNA repair. To achieve the goal of eradicating cancer, it is paramount to understand the underlying molecular mechanisms for the maintenance of genetic stability. This book provides a snapshot of our current understanding of DNA damage repair and recent advances in the research of DNA repair, genetic instability and cancer. Qingyi Wei Lei Li David Chen

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Contributors

Sandeep Burma, PhD Assistant Professor Division of Molecular Radiation Biology Department of Radiation Oncology University of Texas Southwestern Medical Center 5801 Forest Park Road Dallas, TX 75390-9187 [email protected] Benjamin Chen, PhD Assistant Professor Division of Molecular Radiation Biology Department of Radiation Oncology University of Texas Southwestern Medical Center 5801 Forest Park Road Dallas, TX 75390-9187 [email protected]

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Contributors

David J. Chen, PhD Professor and Director Division of Molecular Radiation Biology Department of Radiation Oncology University of Texas Southwestern Medical Center 5801 Forest Park Road Dallas, TX 75390-9187 [email protected] Junjie Chen, PhD Professor Department of Therapeutic Radiology Hunter Bldg. Room 213C Yale University School of Medicine 333 Cedar Street, P.O. Box 208040 New Haven CT 06520-8040 [email protected] Rong Guo, PhD Post-Doctoral Fellow Lab of Genetics Genome Stability and Chromatin Remodeling Section National Institute on Aging/NIH 333 Cassell Drive, TRIAD Building RM. 3000 Baltimore, MD 21224 Tel.: 410-558-8489 Fax : 410-558-8331 [email protected]

contributors

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Contributors

Bo Hang, MD, PhD Staff Scientist Department of Molecular Biology Life Sciences Division Lawrence Berkeley National Laboratory University of California Berkeley, CA 94720 [email protected] Zhibin Hu, MD, PhD Post-Doctoral Fellow Department of Epidemiology The University of Texas M. D. Anderson Cancer Center 1515 Holcombe Blvd. Houston, Texas 77030 [email protected] Maxwell P. Lee, PhD Investigator Lab Population Genetics (HNC7Z35) National Cancer Institute Building 41, Room D702C 41 Library Drive Room D702 Bethesda, MD 20892-5060 [email protected] Guo-Min Li, PhD Professor Departments of Toxicology and Pathology University of Kentucky Medical Center 125 Health Sciences Research Building 800 Rose Street Lexington, KY 40536 [email protected]

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Contributors

Lei Li, PhD Associate Professor Department of Experimental Radiation Oncology The University of Texas M. D. Anderson Cancer Center 1515 Holcombe Blvd. Houston, Texas 77030 [email protected] Jac A. Nickoloff, PhD Professor and Chairman Department of Molecular Genetics and Microbiology CRF 127 University of New Mexico HSC 915 Camino de Salud NE Albuquerque, NM 87131 [email protected] Binghui Shen, PhD Professor and Director Department of Radiation Biology City of Hope National Medical Center and Beckman Research Institute 1500 East Duarte Road Duarte, CA 91010 [email protected]

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Contributors

Zhiyuan Shen, PhD Associate Professor and Chief Division of Radiation Cancer Biology Department of Radiation Oncology Robert Wood Johnson Medical School Cancer Institute of New Jersey University of Medicine and Dentistry of New Jersey 195 Little Albany St New Brunswick, NJ 08903-2681 [email protected] Purnima Singe, PhD Research Fellow Department of Radiation Biology City of Hope National Medical Center and Beckman Research Institute 1500 East Duarte Road Duarte, CA 91010 [email protected] Li-E Wang, MD Instructor Department of Epidemiology The University of Texas M. D. Anderson Cancer Center 1515 Holcombe Blvd. Houston, Texas 77030 [email protected]

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Contributors

Weidong Wang, PhD Senior Investigator Genome Instability and Chromatin Remodeling Section National Institute on Aging/NIH 333 Cassell Drive, TRIAD Building RM. 3000 Baltimore, MD 21224 [email protected] Qingyi Wei, MD, PhD Professor Department of Epidemiology — 1365 The University of Texas M. D. Anderson Cancer Center 1515 Holcombe Blvd. Houston, Texas 77030 [email protected] Jamie Wood Department of Therapeutic Radiology Hunter Bldg. Room 210 Yale University School of Medicine 333 Cedar Street, P.O. Box 208040 New Haven CT 06520-8040 [email protected]

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Wei Xiao, PhD Professor and Head Department of Microbiology and Immunology College of Medicine University of Saskatchewan 107 Wiggins Road Saskatoon, SK, S7N 5E5 Canada [email protected] Fang Xu Professor Department of Biology Ningxia Medical College 692 Sheng-Li Road Yinchun, Ningxia 750004, China [email protected] Wei Yang, PhD Senior Investigator Laboratory of Molecular Biology Molecular Structure Section (HNK6C5) NIDDKD/NIH Building 5, Room B107 5 Memorial Dr. Bethesda, MD 20892 [email protected]

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Yingnian Yu, MD Professor Department of Pathophysiology Center of Environmental Genomics Zhejiang University School of Medicine 353 Yan’an Road Hangzhou, Zhejiang 310031, China [email protected]

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Contents

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Preface

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Contributors

1

Chapter 1

DNA Damage Sensing and Signaling Jamie L. Wood and Junjie Chen

Chapter 2

Base Excision Repair Bo Hang

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Chapter 3

Nucleotide Excision Repair Lei Li

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Chapter 4

DNA Mismatch Repair: Biological Functions and Molecular Mechanisms Guo-Min Li

Chapter 5

Chapter 6

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Mammalian Homologous Recombination Repair and Cancer Intervention Zhiyuan Shen and Jac A. Nickoloff

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Role of Non-Homologous End Joining in the Repair of DNA Double-Strand Breaks Sandeep Burma, Benjamin Chen and David J. Chen

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Chapter 7

Chapter 8

Chapter 9

Chapter 10

Chapter 11

Chapter 12

Index

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A DNA-damage Response Network of Fanconi Anemia and BRCA Proteins Rong Guo and Weidong Wang

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Structure and Function of DNA Repair Proteins: Lesion Recognition Wei Yang

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DNA Damage Tolerance and Translesion Synthesis Fang Xu, Yingnian Yu and Wei Xiao

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Nucleases in DNA Repair, Replication and Recombination: Flap Endonuclease-1 as a Paradigm Purnima Singh and Binghui Shen Integrative Genomics and Epigenomics: Application in Cancer Research Maxwell P. Lee Molecular Epidemiology of DNA Repair and Cancer Susceptibility — A Review of Population-Based Studies Zhibin Hu, Li-E Wang and Qingyi Wei

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CHAPTER 1

DNA Damage Sensing and Signaling Jamie L. Wood and Junjie Chen∗

ABSTRACT Mechanisms have evolved to help protect our genetic material from endogenous and exogenous damage that are vital for an organism’s survival. This ability to recognize damaged DNA and simultaneously regulate cell cycle progression and DNA repair is critical for genomic stability, and defects in these pathways are hallmarks of cancer. The DNA damage response pathway is a multi-component signal transduction network that consists of a multitude of proteins whose functions are still under investigation. This review will focus on the current findings in the field including how the DNA damage response influences cancer predisposition and formation.

1. INTRODUCTION Safeguarding an organism’s genome from endogenous (reactive oxygen species, abnormal replication intermediates) and exogenous (UV and ionizing radiation, reactive chemicals) sources of DNA damage insures timely and accurate passage of genetic information onto daughter cells. Cells have evolved an elaborate surveillance system to respond to these harmful stimuli. These surveillance pathways are called cell cycle checkpoints, which are activated upon detection of DNA damage, thus allowing the repair of the genetic ∗ Corresponding

author. Guggenheim 1306, Division of Oncology Research, Mayo Clinic, 200 First Street S.W., Rochester, MN 55905. Tel.: 507-538-1545; Fax: 507-284-3906; E-mail: ∗ [email protected] 1

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lesions. Cell cycle checkpoints are critical for detecting damaged DNA and coupling progression through the cell cycle with DNA repair, and if the damage is too great, they can also trigger cell death. Increasing evidence has shown that defects within these checkpoints can lead to genetic instability, a hallmark of cancer cells.1 Indeed, proteins involved in the DNA damage response and DNA repair, when mutated, cause genetic diseases which predispose individuals to cancer. BRCA1 mutations account for over 50% of familial breast cancer cases, and mutations in ataxia-telangeictasia-mutated (ATM) cause ataxia-telangeictasia (A-T) syndrome, resulting in sensitivity to DNA damaging agents as well as cancer. Recent studies have also provided evidence that the DNA damage response is an anti-cancer barrier, which must be overcome in early neoplastic lesions for tumorigenesis to occur.2,3 Accurate and timely sensing and repair of DNA lesions is necessary for the cell to survive, and underscores the importance of sensing and signaling DNA damage. This review will focus on the mammalian DNA damage response pathway highlighting recent advances in the field. 2. THE DNA DAMAGE CHECKPOINT The DNA damage response is similar to other signal transduction pathways in that it has several different components that act in concert to activate the checkpoint. It has been suggested that these components consist of sensors, signal transducers, and effectors and recently, to that list has been added mediators.4 In the course of the cell cycle there are several checkpoints that can be activated in response to DNA damage; the G1/S checkpoint, the intra-S phase checkpoint, and the G2/M checkpoint. Canonically, the DNA damage response is a signal transduction cascade triggered by a series of phosphorylation-dependent events, which activate proteins involved in transducing the DNA damage signal (IR or stalled replication) to different effector proteins which evoke specific effects, namely halting cell cycle progression, activating DNA repair mechanisms and transcription, and triggering apoptosis (Fig. 1). Mediators are placed between the sensors and

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DNA Damage Sensing and Signaling

Fig. 1 Activation of the DNA damage response pathway in response to ionizing radiation and replication stress.

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signal transducers as their function is to help bring these proteins together to activate the response. However, the DNA damage response is not a linear pathway; DNA damage proteins can also act both upstream and downstream of their respective functions and maintain crosstalk with other proteins. Instead of a linear pathway as previously thought, there is an intricate network of proteins that function independently and dependently to coordinate the response to DNA damage and maintain genomic stability. We will begin to understand this process by first dissecting the definition of sensor proteins in this pathway. 3. RECOGNITION OF DNA DAMAGE It is important that DNA molecules, once damaged, must be recognized to initiate subsequent checkpoint responses. An “official” checkpoint-specific damage sensor is still unknown, though several candidate proteins have been implicated. Genetic systems such as yeast have provided elegant tools to dissect the pathway (Table 1); however, debate still rages as to what the original DNA damage sensor in mammals is. Nevertheless, we will discuss these proteins in their context as sensors: ATM/ATR, the MRN complex, Rad17 and the Rad9-Hus1-Rad1 complex. Ataxia-telangeictasia mutated (ATM) was first cloned in 1995 as the causative gene for ataxia-telangeictasia (A-T) disorder, which is characterized by cerebellar degeneration, immune system defects and cancer predisposition.5,6 From this phenotype, ATM has been deduced to be important for the cellular response to double-strand breaks (DSBs); however, the mouse KO is still viable.7 ATM is a member of the phophatidylinositol-3-kinase-like family of serine/threonine protein kinases (PIKKs). ATM phosphorylates its substrates on so called SQ/TQ motifs, which is on serine/threonines followed by a glutamine. Like other PIKKs, ATM contains a FAT domain, a phosphoinositide-3,4-kinase (PI3K) domain and a FAT carboxy-terminal (FAT-C) domain. ATM is a nuclear protein that is activated in response to DSBs and possibly chromatin alterations, which has been elegantly shown to dissociate from an inactive

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Table 1.

5

From Yeast to Man: Conservation of the DNA Damage Pathway

Function

Mammals

S. cerevisiae

S. pombe

PI3K-Kinases

ATM ATR

Tel1 Mec1

Tel1 Rad3

Replication Factor C-like

Rad17 Rfc2-5

Rad24 Rfc2-5

Rad17 Rfc3

PCNA-like

Rad9 Hus1 Rad1

Ddc1 Mec3 Rad17

Rad9 Hus1 Rad1

BRCA1 MDC1 53BP1 TopBP1 Claspin

Rad9 ? ? Dpb11 Mrc1

Crb2/Rph9 ? ? Cut5 Mrc1

Chk1 Chk2

Chk1 Rad53

Chk1 Cds1

Cdc25A, Cdc25C

Cdc25

Cdc25

p53

?

?

Sensors

Mediators BRCT containing

Transducers/Effectors Ser/Thr Kinase Phosphatase Transcription Factor

dimer into active monomers through auto-phosphorylation on serine 1981.8 These active ATM monomers then relocalize to sites of DSBs forming nuclear foci, a common hallmark of most DNA damage responsive proteins.9 Once at the site of the DSB, ATM phosphorylates several substrates, including H2AX, p53, Chk2, NBS1 and BRCA1. Phosphorylated H2AX (γH2AX) foci form rapidly at the site and within megabase regions surrounding the DSB, and are thought to be an essential and powerful mechanism for amplifying the damage signal through recruitment of transducer and repair proteins.10 ATM has been shown to bind to damaged chromatin,11 further enhancing its role as the sensor protein. However, work recently from Stephen Jackson and colleagues showed that ATM recruitment to DSBs is mediated by NBS1 through a conserved Cterminal motif that binds to ATM.12 This C-terminal motif is also

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found in Ku80 and ATRIP, and was demonstrated to be also responsible for the recruitment of DNA-PK and ATR, respectively. ATM is still the apical kinase that is responsible for the initiation of the DNA damage response to DSBs; however, its function as solely a sensor remains unclear. Ataxia-telangeictasia-mutated and Rad3 Related (ATR) was first cloned as the human homolog to fission yeast rad3 and was found to be a member of the PIKK family. ATR shares similar structural similarity with ATM, as well as its function as a serine/threonine kinase; however, ATR is an essential gene as null mutations are embryonic lethal and cells derived from these mice are not viable.13 Recently a point mutation in the ATR gene was discovered as the cause of an autosomal recessive disorder known as Seckel syndrome, which shares a similar phenotype with Nijmegen breakage syndrome.14 While ATM is activated in response to DSBs, ATR is activated upon replication stress. During the S-phase, ATR plays a pivotal role in insuring faithful replication, and becomes rapidly activated and binds to chromatin in response to UV, hyperoxia and replication inhibitors.15–17 How ATR became activated was recently addressed18 through the hypothesis that RPAcoated single-stranded DNA (ssDNA) is a signaling intermediate that is bound by ATR interacting protein (ATRIP), which in turn recruits ATR. Recent data suggest this theory is correct as a C-terminal motif in ATRIP is responsible for the recruitment of ATR in response to replication stress.12 Therefore, RPA:ssDNA has been postulated to be responsible for activating ATR and is the “signal” required to trigger ATRdependent checkpoint events. Next is the highly conserved MRN complex which contains three core proteins: Mre11, Rad50 and Nbs1 that function together to bind to DSBs, recruit ATM, and initiate DNA repair.19 The MRN complex possesses both 3 -5 -exonuclease and single strand endonuclease activities, DNA binding ability, as well as limited DNA unwinding activity.20 Electron microscopy and scanning force microscopy have shown that the MRN complex appears as a bipolar structure with a globular head and two long tails. The head consists of two RAD50 ATPase domains (one at the N-terminus, the other at the

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C-terminus of Rad50) which are brought together by a flexible hinge region at the center of the protein which contains a CXXC “zinchook” that can bind and dimerize with the “zinc-hook” of another Rad50 molecule.21 Mre11 binds to Rad50 and possesses DNA binding activity which, with the tethering ability of Rad50, can form a flexible tether that can bridge DNA molecules. Nbs1 is the last partner of this important triad, and is thought to stabilize the complex through its interactions with γH2AX at the site of the DSB, recruit ATM, and participate in ATM-dependent signaling events. Due to its endonuclease and exonuclease activity, the MRN complex has also been postulated to be involved in the early stages of DSB repair by binding to DNA ends and processing them for other repair factors. The MRN complex proteins are essential, as mouse knockouts for each protein is lethal, and in humans hypomorphic mutations in these three proteins cause cancer susceptibility syndromes; Ataxia-Telangeictasia-like disorder (Mre11 and Rad50), and Nijmegens breakage syndrome (Nbs1).22,23 Patients with these disorders share many similarities with AT patients, presenting with mental retardation, microcephaly, immunodeficiency and predisposition to lymphoid malignancies. Cells derived from these patients display radiation sensitivity, radiation resistant DNA synthesis and chromosome fragility. Both of these disorders recapitulate the AT phenotype and further couple the MRN complex to the function of ATM. Indeed, the MRN complex has been shown to be required for ATM-dependent signaling events, and Nbs1 itself is phosphorylated by ATM on Ser343, which is required for the activation of the S-phase checkpoint.24 Recently Nbs1 has been shown to have a critical role in ATM recruitment; a C-terminal motif of Nbs1 is required for ATM recruitment to DSBs and has also been shown to promote phosphorylation of ATM substrates such as Chk2, SMC1 and FANCD2, which can further facilitate checkpoint activation and repair.12,25–28 Thus, instead of acting as a substrate and downstream effector of ATM, the MRN complex is now believed to also function upstream of ATM and may serve as a sensor for DSBs. Finally, there is the Rad17-RFC and the Rad9-Hus1-Rad1 (9-1-1) complex. In yeast, early genetic screens discovered several mutants

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that were sensitive to DNA damage, and were called radiation sensitive or Rad genes. Of these, Rad17, Rad9, Hus1 and Rad1 have been characterized in mammalian and yeast systems and form clamp and clamp-loader complexes similar to that of proliferating cell nuclear antigen (PCNA) and the replication factor complex (RFC). PCNA is a sliding clamp that is loaded onto DNA at sites of ongoing replication and acts as a scaffold to tether other replication proteins. RFC is the clamp-loader responsible for loading PCNA onto DNA. Specifically, Rad17 has been purified and shows similarity with the RFC, with Rad17 replacing the large subunit, forming a clamp loader complex. Human Rad9, Hus1 and Rad1 from a doughnut-shaped trimeric clamp that has been shown to associate with Rad17. More importantly, these proteins have been shown to bind to chromatin after genotoxic stress, and the loading of this so-called 9-1-1 complex onto damaged DNA is Rad17 dependent, paralleling the function of RFC/PCNA in regular replication.29 This has caused several ideas as to Rad17 and the 9-1-1 complex as a potential DNA damage sensor, as Rad17 loading of the 9-1-1 complex is activated in response to DNA damage. However, recent studies suggest that ATR and the activation of ATR-dependent signaling pathways can occur independently of Rad17 and the 9-1-1 complex, suggesting that these Rad proteins may not be the sole sensors of replication stress. 4. ACTIVATION OF THE DNA DAMAGE RESPONSE Once DNA damage is sensed and the apical kinases ATM and ATR are activated, they phosphorylate several substrates that initiate a signaling cascade to activate the DNA damage response. These substrates are the so-called mediator and/or transducer proteins: BRCA1, MDC1 and 53BP1 and the kinases Chk1 and Chk2. BRCA1, MDC1 and 53BP1 are mediators in the checkpoint pathway that perform myriad functions to both activate and maintain the damage checkpoint through protein-protein interactions that recruit ATM/ATR substrates to the site of the damaged DNA. Chk1 and Chk2 are phosphorylated and activated by ATM/ATR and transduce the original damage signal and further amplify the

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DNA damage response, ensuring activation of surveillance pathways responsible for taking care of the genetic damage. The breast cancer susceptibility protein 1 (BRCA1) was originally cloned 15 years ago to the long arm of chromosome 17.30 BRCA1 was found to be a tumor suppressor, as women carrying a mutation in BRCA1 need to lose the wild-type allele for cancer formation. BRCA1 mutations are most commonly found in familial breast and ovarian cancer, with rare mutations found in sporadic breast and ovarian cancer. BRCA1 has two well conserved domains: an N-terminal RING domain and at its C-terminus tandem repeats termed the BRCT domain. Cell lines derived from BRCA1 tumors display sensitivity to DNA damaging agents, defective cell cycle checkpoints and HR repair.31 BRCA1 knockout mice are embryonic lethal, and ES cells derived from these mice display gross chromosomal rearrangements and defective DNA repair.32,33 BRCA1, through its RING domain, specifically interacts with BARD1 via its RING domain and forms a heterodimer that has E3 ubiquitin ligase activity. BRCA1/BARD1 ligase activity has been demonstrated in vitro and in vivo, and catalyzes primarily monoubiquination and polyubiquitination at K6.34,35 However, the importance of ubiquitination by BRCA1/BARD1 and its substrates still remains elusive. The BRCT domain has recently been demonstrated to be a phosphor-peptide binding motif by our group and others.36,37 The BRCT domain interacts with BACH1, a member of the DEAH helicase family, specifically through a phosphor-serine residue (Ser990) that when mutated, abolishes the BRCA1-BACH1 interaction, and disrupts the G2/M checkpoint.36 Mutations in breast cancer patients in the BRCT domain have been discovered that disrupt this interaction, pointing to the importance of BRCA1 BRCT domain interacting proteins in maintenance of genomic stability. It is believed that through its BRCT domain, BRCA1 can interact with multiple binding partners and thus participates in DNA damage response. While the RING domain of BRCA1 is also important for BRCA1 function, the molecular mechanisms still remain to be resolved. Our lab and others have discovered MDC1 as an adaptor protein in the DNA damage response pathway.38–40 MDC1 contains an

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N-terminal forkhead-associated (FHA) domain, a phosphothreonine/serine binding motif that is often found in proteins involved in cell cycle, DNA repair and mRNA processing.41 MDC1 contains a region of 14 conserved repeats (aa1124-1697) that specifically interacts with DNA-dependent protein kinase (DNA-PK), and regulates its autophosphorylation in response to DNA damage, and subsequent DNA repair. Residing in the C-terminus of MDC1 is the tandem BRCT domain, which interacts with γH2AX.38 MDC1 interacts with all three members of the MRN complex, SMC1, Chk2, BRCA1 and ATM. Previous studies using MDC1 small interfering RNA (siRNA) in cell lines have bolstered the mediator function of MDC1 as defects in the intra-S phase checkpoint, ATM recruitment, and DNA damaged induced apoptosis is impaired in MDC1 depleted cells.38–40 Recently, our lab, using a MDC1 deficient mouse model, has shown that MDC1 is a bridging molecule between H2AX and ATM. These three proteins form a positive feedback loop following DNA damage. MDC1 binds to γH2AX through its BRCT domain and interacts with activated ATM through its FHA domain. Because of these two protein-protein interactions, MDC1 brings ATM in the proximity of H2AX for further H2AX phosphorylation and subsequent accumulation of active ATM at the sites of DSBs (Lou, in press 2006). This loop acts to amplify ATM-dependent signaling events upon DNA damage and facilitates the ATM-dependent phosphorylation events since many ATM substrates are also shown to accumulate at the sites of DSBs in a MDC1/H2AX-dependent manner. P53 Binding Protein 1 (53BP1) was originally identified in a yeast two hybrid screen for proteins that bound to the DNA-binding region of p53. Similar to MDC1 and BRCA1, 53BP1 contains Cterminal BRCT domains, and a Tudor domain in the middle of the protein. 53BP1 was thought to be the homolog of yeast DNA damage checkpoint Rad9, due to sequence homology. Upon DNA damage, 53BP1 is phosphorylated by both ATM/ATR and relocalizes to the damaged site forming foci.42 The retention and accumulation of 53BP1 at DSBs requires γH2AX, and possibly HDAC4.43,44 Studies utilizing 53BP1-deficient cell lines and 53BP1 siRNA described a defect in the intra S-phase and G2/M checkpoint at low dose of

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ionizing radiation, pointing to a role of 53BP1 in cell cycle checkpoint control.45 53BP1 deficient mice are viable, but show increased genomic instability and tumor incidence, which suggests that 53BP1 is a bona fide tumor suppressor. This notion is further supported by recent findings from our lab indicating that 53BP1 is a haploinsufficient tumor suppressor as loss of one 53BP1 allele compromised genomic stability and DNA repair.43,46 Checkpoint Kinase 1 (Chk1) is a critical messenger of checkpoint control and is activated in response to diverse genotoxic insults. Together with another checkpoint kinase Chk2, it regulates fundamental cellular functions such as DNA replication, cell cycle progression, chromatin restructuring, and apoptosis.1 Chk1 was first discovered in fission yeast through its ability to complement a cold sensitive cdc2 mutant. Disruption of fission yeast Chk1 rendered the cells sensitive to UV, bypassing the normal G2/M delay and the cells entering mitosis despite damaged DNA.47 Unlike yeast, murine Chk1 is essential for development as Chk1 deficient embryos died between embryonic days, 3.5 and 7.5, and Chk1−/− ES cells underwent apoptosis.48 Chk1 is activated in response to DNA damage or replication stress by the ATR and ATM kinases. Chk1 has been demonstrated to be activated upon phosphorylation of two conserved serine residues, Ser317 and Ser 345.49 Chk1 also requires other factors for efficient activation; these adaptor proteins presumably function to recruit Chk1 to ATR or ATM.50 Chk1 monitors Sphase progression and G2/M transition by regulating the Cdc25 phosphatase. Cdc25 is crucial for removing the inhibitory phosphorylation on Thr14 and Tyr15 on the cyclin-dependent kinase (cdk), allowing it to become active.51 In yeast both Chk1 and Cds1 phosphorylate Cdc25 in the presence of DNA damage or replication block, resulting in the inhibition of Cdc25 activity and cell cycle arrest. Human Chk1 has similar functions, i.e. functions mainly mediated through regulation of the human homologues of Cdc25. Chk1-dependent phosphorylation of Cdc25A controls Cdc25A protein levels tightly during normal interphase and also following DNA damage or replication block,1 and thus regulates cell cycle progression.

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Another similar Ser/Thr kinase that also transduces the damage signal to its substrates is Checkpoint Kinase 2 (Chk2). Chk2 is primarily activated by ATM and is important for checkpoint control and apoptosis. Chk2 is the mammalian homolog of budding yeast Rad53 and fission yeast Cds1. In response to DNA damage, Chk2 is phosphorylated and activated by ATM. ATM phosphorylates Chk2 on Thr68, allowing Chk2 to dissociate from inactive dimers to active monomers.52 This site is not only critical for Chk2 activation but is a docking site for MDC1’s FHA domain, as well as Chk2’s own FHA domain.40 Chk2 also autophosphorylates two residues in its activation loop, which is required for full activity, and in its C-terminus S516, which is critical for the induction of apoptosis.53 As a terminal kinase in the DNA damage signaling pathway, Chk2 phosphorylates several substrates, including BRCA1, Plk1, E2F1, p53 and Cdc25A, affecting the cell cycle, apoptosis and DNA repair. Consistent with Chk2’s involvement in several important cellular processes, Chk2 has been postulated as a tumor suppressor. Mutations in Chk2 have been found in a subset of Li-Fraumeni syndrome patients, a syndrome with early childhood onset of tumors that has been mostly linked to p53 mutations.54 In hereditary breast cancer, a mutation causing the truncation of Chk2 (1100delC) has been found.55 Despite this evidence, Chk2 null mice are viable, healthy, and are not cancer-prone.56,57 Of note, these Chk2 deficient mice were extremely resistant to radiation-induced apoptosis, and have given rise to the hypothesis that Chk2 inhibitors may serve as chemopreventative agents. 5. ACTIVATION OF CELL CYCLE CHECKPOINTS Cell cycle checkpoints are the defense mechanism that is initiated by the DNA damage response. Checkpoints allow the cell time to repair by putting the brakes on the cell cycle progression. During this interphase there are specific checkpoints, named the G1/S, intra-S phase, replication, and G2/M checkpoint. We will begin with the important step that starts a cell’s fate down the path of mitosis, the transition from G1 to S phase.

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5.1. G1/S Phase Checkpoint Primarily the G1/S checkpoint is required to ensure the fidelity of the genome before it undergoes replication. The major effector of this checkpoint is p53, which is activated upon DNA damage by the ATM and Chk2 kinases. Normally p53 is targeted for degradation via the ubiquitin/proteasome pathway by the action of HDM2, an E3 ligase that binds, ubiquitinates and promotes p53 destruction.58 However, in response to DNA damage, p53 is phosphorylated on Ser20 by Chk2, which disrupts the HDM2-p53 interaction and stabilizes p53.59 P53 also undergoes phosphorylation by ATM and ATR in response to IR and UV, respectively, on Ser15, which increases p53’s transcriptional activity. Once active, p53 activates the transcription of several genes, one of which being p21Cip1/WAF1, a cyclindependent kinase (CDK) inhibitor.60 P21 can bind to CDK2/Cyclin A/E and inhibit its activity, which results in cell cycle arrest in the G1/S restriction point.61 Therefore, a pathway from DNA damage to ATM/Chk2 to p53/p21 ensures G1/S checkpoint control following DNA damage. 5.2. S-phase Checkpoint The signal for DNA damage during the S-phase can be DNA lesions (DSBs) or impaired DNA synthesis, leading to stalled replication forks. As mentioned before, ATM is activated by DSBs, while ATR becomes active upon replication stress. This leads to two appreciable checkpoints during the S-phase, the intra-S phase checkpoint mediated by ATM, and the replication checkpoint mediated by ATR. Despite this, they both have a common effect, in that they both function to block the firing of origins to delay DNA synthesis allowing for repair. The intra-S checkpoint occurs independently of stalled replication forks. In response to IR, ATM is activated and phosphorylates Chk1, which is the main transducer in this pathway. Chk1 phosphorylates Cdc25A, a serine/threonine phosphatase, which is required for the dephosphorylation of the inhibitory phospho-T14Y15 residues of Cdk2. Cdc25A is phosphorylated by Chk1 on four different residues (Ser123, 178, 278, 292), which

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ultimately promotes its degradation via recognition by the F-box protein β-TrcP.62,63 This leads to cell cycle arrest until the damage is repaired. There is also a parallel pathway that depends on SMC1 for the intra-S-phase checkpoint. SMC1 or structural maintenance of chromosomes 1 is a cohesion protein important for holding the sister chromatids together after replication. SMC1 has been proven to be an important target of ATM in response to IR. SMC1 is phosphorylated by ATM on two residues (Ser957 and Ser966), that when mutated, cause defective intra-S-phase checkpoint and radiosensitization in cells and in knockin mice.27,64 The molecular mechanism of how SMC1 works in the intra-S-phase checkpoint remains to be determined. SMC1 is not the only protein that has an effect on the intra-S-phase checkpoint; several mediator proteins also play a role. Due to their specific protein-protein interaction motifs, it is thought they act as molecular scaffolds or bridges, thereby recruiting or facilitating downstream targets for phosphorylation by apical kinases at the damage sites. NBS1 and BRCA1 are phosphorylated by ATM in response to IR, and these phosphorylations are important for checkpoint activation as well as SMC1 phosphorylation.24,65,66 53BP1 has also been shown to function during the intra-S checkpoint through its ability to affect BRCA1 and Chk2 phosphorylation and foci formation, but recent studies have shown that 53BP1-deficient cells have only a slight or moderate defect in the intra-S-phase checkpoint,43 suggesting the existence of multiple intra-S phase checkpoint pathways. Indeed, another mediator protein MDC1 also functions in the intra-S-phase checkpoint, as it regulates BRCA1 and SMC1 phosphorylation, as well as interacts and regulates the foci formation of NBS1 and BRCA1.38–40 Most importantly, depletion of MDC1 results in RDS in response to IR.38–40 Mediator proteins such as the ones just mentioned have become important in our understanding of the regulation and activation of the intra-S-phase checkpoint. As mentioned above, the replication checkpoint is also activated in the S-phase upon replication stress. While blocking origin firing, this checkpoint also promotes stabilization of stalled replication forks and restarting of replication following DNA repair. In response to replications stress, ATR is activated and together with

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its regulatory subunit, ATRIP, bind to RPA coated single-stranded DNA.18 Independently, the Rad17 clamp loader recruits and loads the 9-1-1 complex onto chromatin. Additionally, Claspin, a recent protein identified in human and Xenopus, associates with the stalled replication forks.67 Claspin, in response to replication stress, participates in the replication checkpoint by playing a role in Chk1 activation.68 It is known that the binding of ATR/ATRIP and the 9-1-1 complex occurs independently; however, it seems that they work in concert to phosphorylate and activate Chk1.29 Together, the recruitment of these proteins allows for the stabilization of the replication forks and full activation of Chk1, which in turn, halts origin firing and further replication through targeting Cdc25A for degradation. 5.3. G2/M Checkpoint The G2/M checkpoint prevents cells from initiating mitosis when they are exposed to DNA damage during G2, or if they harbor unrepaired damage from G1 or the S-phase and they progress into G2. The critical target of the G2/M checkpoint is inhibition of the Cyclin B/Cdk1 complexes, which occurs through inhibition of Cdc25C and also degradation of Cdc25A.69–71 Cdc25C, like Cdc25A, is a phosphatase whose job is to remove the inhibitory phosphorylation on Thr14 and Tyr15 of Cdk1, resulting in its activation. In response to damage, ATM/ATR are activated and phosphorylate both Chk1 and Chk2, although Chk1 has been shown to be predominant in G2 checkpoint activation. Chk1-deficient cells have defective G2/M checkpoint; however, Chk2 may only play a supporting role as Chk2deficient cells still maintain an intact G2/M checkpoint.56,71 Chk1 targets Cdc25A for degradation as previously discussed, and phosphorylates Cdc25C on a 14-3-3 binding site, which promotes its export and sequestration to the cytoplasm. ATM and ATR both function during the G2/M checkpoint. Based on knockout mice studies, it appears that ATM and ATR cooperate in activating early G2 arrest, but ATR is required for late G2 arrest.72 Mediator proteins again assert their abilities for checkpoint activation, since BRCA1, MDC1,

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and to a lesser extent 53BP1, are also required for G2/M checkpoint activation.40,45,73 6. THE DNA DAMAGE RESPONSE AND CANCER Maintaining genomic stability is ultimately what cells are supposed to do when they encounter genotoxic stress. Proteins involved in the DNA damage and cell cycle checkpoint response have been shown to be the gatekeepers of the genome, and when lost or mutated can cause genomic instability, a hallmark of cancer cells. Several of these proteins involved in DNA damage response are bona fide tumor suppressors, that when mutated cause inherited cancer susceptibility syndromes (Table 2). Recently, two papers have shown in elegant detail the importance of the DNA damage response pathway in the formation of cancer. Bartkova et al. and Gorgoulis et al. demonstrate that DNAdamage pathways are activated during uncontrolled DNA replication in cells with overexpression of oncogenes.2,3 The activation of normal DNA damage response in these precancerous cells should arrest cell cycle progression and/or activate apoptosis. The authors postulate that this in turn might cause a selection pressure to inactivate proteins involved in the DNA damage response so that full-blown cancer cells can emerge in the absence of these normal cell cycle brakes. Thus, a model may be that in early precancerous lesions the DNA damage response is activated and many of these Table 2.

Inherited Cancer Susceptibility Syndromes

Syndrome

Gene

Nature of Mutation

Cancer Phenotype

Ataxia-Telangeictasia (A-T)

ATM

Null and Hypomorphic

Lymphoid tumors, T-cell Leukemias

Ataxia-Telangeictasia-like Disorder (ATLD)

Mre11

Hypomorphic

None Reported

Nijmegens-Breakage Syndrome (NBS)

Nbs1

Hypomorphic

B-Cell Lymphomas, Leukemias

ATR

Hypomorphic

None Reported

Seckel Syndrome Familial Breast and Ovarian Cancer

BRCA1 Missense, Frameshift Breast and Ovarian Cancer

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lesions will not progress into malignant tumors. However, in some of these precancerous cells, the normal DNA damage response may be inactivated due to random mutations, thus leading to the further accumulation of genetic instability and eventually tumorigenesis. More work investigating these hypotheses begs attention as we can further understand how cancer develops.

7. SUMMARY In conclusion, the DNA damage response is an intricate network of signaling pathways whose sole purpose is to protect the genome. Deregulation of this response can lead to genomic instability and tumorigenesis, as evidenced by inherited cancer susceptibility syndromes associated with these proteins, and their inactivation or deregulation in cancer cells. Current and future research directed at understanding these pathways can lead to the elucidation of tumorigenesis and possible therapeutic applications that can ameliorate the looming threat that cancer is across the world.

ACKNOWLEDGMENTS We would like to thank all the members of the Chen lab for discussion and help, especially Z. Lou and Irene Ward for their suggestions and reading of the manuscript. JC is a recipient of the DOD breast cancer research Era of Hope scholar award.

References 1. Bartek J, Lukas J. (2003) Chk1 and Chk2 kinases in checkpoint control and cancer. Cancer Cell 3: 421–429. 2. Bartkova J, Horejsi Z, Koed K, et al. (2005) DNA damage response as a candidate anti-cancer barrier in early human tumorigenesis. Nature 434: 864–870. 3. Gorgoulis VG, Vassiliou LV, Karakaidos P, et al. (2005) Activation of the DNA damage checkpoint and genomic instability in human precancerous lesions. Nature 434: 907–913.

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4. Sancar A, Lindsey-Boltz LA, Unsal-Kacmaz K, Linn S. (2004) Molecular mechanisms of mammalian DNA repair and the DNA damage checkpoints. Annu Rev Biochem 73: 39–85. 5. Shiloh Y. (2003) ATM and related protein kinases: safeguarding genome integrity. Nat Rev Cancer 3: 155–168. 6. Lavin MF, Shiloh Y. (1997) The genetic defect in ataxia-telangiectasia. Annu Rev Immunol 15: 177–202. 7. Xu Y, Ashley T, Brainerd EE, et al. (1996) Targeted disruption of ATM leads to growth retardation, chromosomal fragmentation during meiosis, immune defects, and thymic lymphoma. Genes Dev 10: 2411–2422. 8. Bakkenist CJ, Kastan MB. (2003) DNA damage activates ATM through intermolecular autophosphorylation and dimer dissociation. Nature 421: 499–506. 9. Bartek J, Lukas C, Lukas J. (2004) Checking on DNA damage in S phase. Nat Rev Mol Cell Biol 5: 792–804. 10. Celeste A, Petersen S, Romanienko PJ, et al. (2002) Genomic instability in mice lacking histone H2AX. Science 296: 922–927. 11. Andegeko Y, Moyal L, Mittelman L, et al. (2001) Nuclear retention of ATM at sites of DNA double strand breaks. J Biol Chem 276: 38224– 38230. 12. Falck J, Coates J, Jackson SP. (2005) Conserved modes of recruitment of ATM, ATR and DNA-PKcs to sites of DNA damage. Nature 434: 605–611. 13. Brown EJ, Baltimore D. (2000) ATR disruption leads to chromosomal fragmentation and early embryonic lethality. Genes Dev 14: 397–402. 14. O’Driscoll M, Ruiz-Perez VL, Woods CG. (2003) A splicing mutation affecting expression of ataxia-telangiectasia and Rad3-related protein (ATR) results in Seckel syndrome. Nat Genet 33: 497–501. 15. Unsal-Kacmaz K, Makhov AM, Griffith JD, Sancar A. (2002) Preferential binding of ATR protein to UV-damaged DNA. Proc Natl Acad Sci USA 99: 6673–6678. 16. Das KC, Dashnamoorthy R. (2004) Hyperoxia activates the ATR-Chk1 pathway and phosphorylates p53 at multiple sites. Am J Physiol Lung Cell Mol Physiol 286: L87–97. 17. Guo Z, Kumagai A, Wang SX, Dunphy WG. (2000) Requirement for Atr in phosphorylation of Chk1 and cell cycle regulation in response to DNA replication blocks and UV-damaged DNA in Xenopus egg extracts. Genes Dev 14: 2745–2756. 18. Zou L, Elledge SJ. (2003) Sensing DNA damage through ATRIP recognition of RPA-ssDNA complexes. Science 300: 1542–1548. 19. Petrini JH, Stracker TH. (2003) The cellular response to DNA doublestrand breaks: defining the sensors and mediators. Trends Cell Biol 13: 458–462.

FA

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19

20. Trujillo KM, Yuan SS, Lee EY, Sung P. (1998) Nuclease activities in a complex of human recombination and DNA repair factors Rad50, Mre11, and p95. J Biol Chem 273: 21447–21450. 21. Hopfner KP, Craig L, Moncalian G, et al. (2002) The Rad50 zinc-hook is a structure joining Mre11 complexes in DNA recombination and repair. Nature 418: 562–566. 22. Stewart GS, Maser RS, Stankovic T, et al. (1999) The DNA double-strand break repair gene hMRE11 is mutated in individuals with an ataxiatelangiectasia-like disorder. Cell 99: 577–587. 23. Carney JP, Maser RS, Olivares H, et al. (1998) The hMre11/hRad50 protein complex and Nijmegen breakage syndrome: linkage of doublestrand break repair to the cellular DNA damage response. Cell 93: 477–486. 24. Lim DS, Kim ST, Xu B, et al. (2000) ATM phosphorylates p95/nbs1 in an S-phase checkpoint pathway. Nature 404: 613–617. 25. Buscemi G, Savio C, Zannini L, et al. (2001) Chk2 activation dependence on Nbs1 after DNA damage. Mol Cell Biol 21: 5214–5222. 26. Horejsi Z, Falck J, Bakkenist CJ, et al. (2004) Distinct functional domains of Nbs1 modulate the timing and magnitude of ATM activation after low doses of ionizing radiation. Oncogene 23: 3122–3127. 27. Yazdi PT, Wang Y, Zhao S, et al. (2002) SMC1 is a downstream effector in the ATM/NBS1 branch of the human S-phase checkpoint. Genes Dev 16: 571–582. 28. Nakanishi K, Taniguchi T, Ranganathan V, et al. (2002) Interaction of FANCD2 and NBS1 in the DNA damage response. Nat Cell Biol 4: 913–920. 29. Zou L, Cortez D, Elledge SJ. (2002) Regulation of ATR substrate selection by Rad17-dependent loading of Rad9 complexes onto chromatin. Genes Dev 16: 198–208. 30. Narod SA, Foulkes WD. (2004) BRCA1 and BRCA2: 1994 and beyond. Nat Rev Cancer 4: 665–676. 31. Scully R, Ganesan S, Vlasakova K, et al. (1999) Genetic analysis of BRCA1 function in a defined tumor cell line. Mol Cell 4: 1093–1099. 32. Shen SX, Weaver Z, Xu X, et al. (1998) A targeted disruption of the murine Brca1 gene causes gamma-irradiation hypersensitivity and genetic instability. Oncogene 17: 3115–3124. 33. Moynahan ME, Chiu JW, Koller BH, Jasin M. (1999) Brca1 controls homology-directed DNA repair. Mol Cell 4: 511–518. 34. Morris JR, Solomon E. (2004) BRCA1 : BARD1 induces the formation of conjugated ubiquitin structures, dependent on K6 of ubiquitin, in cells during DNA replication and repair. Hum Mol Genet 13: 807–817. 35. Wu-Baer F, Lagrazon K, Yuan W, Baer R. (2003) The BRCA1/ BARD1 heterodimer assembles polyubiquitin chains through an

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September 14, 2006 14:50

20

36. 37.

38. 39. 40.

41.

42.

43.

44.

45.

46.

47. 48.

49.

50.

WSPC/SPI-B414: DNA Repair, Genetic Instability & Cancer

ch01

Jamie L. Wood and Junjie Chen

unconventional linkage involving lysine residue K6 of ubiquitin. J Biol Chem 278: 34743–34746. Yu X, Chini CC, He M, et al. (2003) The BRCT domain is a phosphoprotein binding domain. Science 302: 639–642. Manke IA, Lowery DM, Nguyen A, Yaffe MB. (2003) BRCT repeats as phosphopeptide-binding modules involved in protein targeting. Science 302: 636–639. Stewart GS, Wang B, Bignell CR, et al. (2003) MDC1 is a mediator of the mammalian DNA damage checkpoint. Nature 421: 961–966. Goldberg M, Stucki M, Falck J, et al. (2003) MDC1 is required for the intra-S-phase DNA damage checkpoint. Nature 421: 952–956. Lou Z, Chini CC, Minter-Dykhouse K, Chen J. (2003) Mediator of DNA damage checkpoint protein 1 regulates BRCA1 localization and phosphorylation in DNAdamage checkpoint control. J Biol Chem 278: 13599– 13602. Durocher D, Taylor IA, Sarbassova D, et al. (2000) The molecular basis of FHA domain:phosphopeptide binding specificity and implications for phospho-dependent signaling mechanisms. Mol Cell 6: 1169–1182. Rappold I, Iwabuchi K, Date T, Chen J. (2001) Tumor suppressor p53 binding protein 1 (53BP1) is involved in DNA damage-signaling pathways. J Cell Biol 153: 613–620. Ward IM, Minn K, Jorda KG, Chen J. (2003) Accumulation of checkpoint protein 53BP1 at DNA breaks involves its binding to phosphorylated histone H2AX. J Biol Chem 278: 19579–19582. Kao GD, McKenna WG, Guenther MG, et al. (2003) Histone deacetylase 4 interacts with 53BP1 to mediate the DNA damage response. J Cell Biol 160: 1017–1027. Fernandez-Capetillo O, Chen HT, Celeste A, et al. (2002) DNA damageinduced G2-M checkpoint activation by histone H2AX and 53BP1. Nat Cell Biol 4: 993–997. Ward IM, Difilippantonio S, Minn K, et al. (2005) 53BP1 cooperates with p53 and functions as a haploinsufficient tumor suppressor in mice. Mol Cell Biol 25: 10079–10086. Walworth N, Davey S, Beach D. (1993) Fission yeast chk1 protein kinase links the rad checkpoint pathway to cdc2. Nature 363: 368–371. Takai H, Tominaga K, Motoyama N, et al. (2000) Aberrant cell cycle checkpoint function and early embryonic death in Chk1( − / − ) mice. Genes Dev 14: 1439–1447. Zhao H, Piwnica-Worms H. (2001) ATR-mediated checkpoint pathways regulate phosphorylation and activation of human Chk1. Mol Cell Biol 21: 4129–4139. Chen Y, Sanchez Y. (2004) Chk1 in the DNA damage response: conserved roles from yeasts to mammals. DNA Repair (Amst) 3: 1025–1032.

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51. Donzelli M, Draetta GF. (2003) Regulating mammalian checkpoints through Cdc25 inactivation. EMBO Rep 4: 671–677. 52. Ahn JY, Li X, Davis HL, Canman CE. (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. 53. Wu X, Chen J. (2003) Autophosphorylation of checkpoint kinase 2 at serine 516 is required for radiation-induced apoptosis. J Biol Chem 278: 36163–36168. 54. Bell DW, Varley JM, Szydlo TE, et al. (1999) Heterozygous germ line hCHK2 mutations in Li-Fraumeni syndrome. Science 286: 2528–2531. 55. Meijers-Heijboer H, van den Ouweland A, Klijn J, et al. (2002) Lowpenetrance susceptibility to breast cancer due to CHEK2(*)1100delC in noncarriers of BRCA1 or BRCA2 mutations. Nat Genet 31: 55–59. 56. Hirao A, Cheung A, Duncan G, et al. (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. 57. Takai H, Naka K, Okada Y, et al. (2002) Chk2-deficient mice exhibit radioresistance and defective p53-mediated transcription. Embo J 21: 5195–5205. 58. Levav-Cohen Y, Haupt S, Haupt Y. (2005) Mdm2 in growth signaling and cancer. Growth Factors 23: 183–192. 59. Chehab NH, Malikzay A, Stavridi ES, Halazonetis TD. (1999) Phosphorylation of Ser-20 mediates stabilization of human p53 in response to DNA damage. Proc Natl Acad Sci USA 96: 13777–13782. 60. Vousden KH, Lu X. (2002) Live or let die: the cell’s response to p53. Nat Rev Cancer 2: 594–604. 61. Xiong Y, Hannon GJ, Zhang H, et al. (1993) p21 is a universal inhibitor of cyclin kinases. Nature 366: 701–704. 62. Sorensen CS, Syljuasen RG, Falck J, et al. (2003) Chk1 regulates the S phase checkpoint by coupling the physiological turnover and ionizing radiation-induced accelerated proteolysis of Cdc25A. Cancer Cell 3: 247–258. 63. Busino L, Donzelli M, Chiesa M, et al. (2003) Degradation of Cdc25A by beta-TrCP during S phase and in response to DNA damage. Nature 426: 87–91. 64. Kitagawa R, Bakkenist CJ, McKinnon PJ, Kastan MB. (2004) Phosphorylation of SMC1 is a critical downstream event in the ATM-NBS1-BRCA1 pathway. Genes Dev 18: 1423–1438. 65. Gatei M, Scott SP, Filippovitch I, et al. (2000) Role for ATM in DNA damage-induced phosphorylation of BRCA1. Cancer Res 60: 3299–3304.

FA

September 14, 2006 14:50

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66. Kim ST, Xu B, Kastan MB. (2002) Involvement of the cohesin protein, Smc1, in Atm-dependent and independent responses to DNA damage. Genes Dev 16: 560–570. 67. Chini CC, Chen J. (2004) Claspin, a regulator of Chk1 in DNA replication stress pathway. DNA Repair (Amst) 3: 1033–1037. 68. Chini CC, Chen J. (2003) Human claspin is required for replication checkpoint control. J Biol Chem 278: 30057–30062. 69. Matsuoka S, Huang M, Elledge SJ. (1998) Linkage of ATM to cell cycle regulation by the Chk2 protein kinase. Science 282: 1893–1897. 70. Sanchez Y, Wong C, Thoma RS, et al. (1997) Conservation of the Chk1 checkpoint pathway in mammals: linkage of DNA damage to Cdk regulation through Cdc25. Science 277: 1497–1501. 71. Zhao H, Watkins JL, Piwnica-Worms H. (2002) Disruption of the checkpoint kinase 1/cell division cycle 25A pathway abrogates ionizing radiation-induced S and G2 checkpoints. Proc Natl Acad Sci USA 99: 14795–14800. 72. Brown EJ, Baltimore D. (2003) Essential and dispensable roles of ATR in cell cycle arrest and genome maintenance. Genes Dev 17: 615–628. 73. Yarden RI, Pardo-Reoyo S, Sgagias M, et al. (2002) BRCA1 regulates the G2/M checkpoint by activating Chk1 kinase upon DNA damage. Nat Genet 30: 285–289.

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CHAPTER 2

Base Excision Repair Bo Hang∗

ABSTRACT Base damage or hydrolytic decay originating from endogenous processes or caused by environmental compounds constantly occurs in genomic DNA. Base excision repair (BER) is the major repair mechanism that removes oxidized, alkylated, deaminated bases as well as apurinic/apyrimidinic (AP) sites. BER is initiated by specific and versatile DNA glycosylases, many of which have redundant functions. The AP site generated by a DNA glycosylase is processed by a stepwise process that involves AP endonuclease, AP lyase, DNA polymerase and DNA ligase. Mammalian BER can operate via distinct subpathways. Both the biochemistry and structural biology of the BER components have progressed rapidly over the last decade. Recent biochemical studies are also beginning to identify an intricate network of protein-protein interactions that may play an important role in mediating concerted steps in BER and in coordinating BER with other DNA metabolic pathways. Gene knockout animal models of the majority of the BER enzymes have been generated or attempted, which provided significant insights into their biological role in the protection of genomic stability. The importance of BER in humans is shown by the findings that defects in the BER enzymes can result in increased risk for cancer, neurodegenerative and other disorders. This is best illustrated by the recent discovery that biallelic germline mutations in DNA glycosylase MYH lead to increased risk for colorectal cancer.

∗ Department of Molecular Biology, Life Sciences Division, Lawrence Berkeley National Laboratory, University of California, Berkeley, California 94720, USA. Tel.: (510) 495-2537; Fax: (510) 486-6488. E-mail: [email protected]

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1. INTRODUCTION The very first enzymes involved in base excision repair (BER) were discovered from Escherichia coli in the early 1970s, two of which are the apurinic/apyrimidinic (AP) endonuclease1 and the uracil-DNA glycosylase (UDG).2 During the last 30 years, over 20 proteins have been identified as the core and accessory proteins that participate in BER. This pathway, characterized by free base release, is now recognized as a major excision mechanism that repairs a broad range of modified or inappropriate DNA bases formed endogenously or exogenously, thus playing a crucial rule in maintaining genomic integrity. The pathway details of BER from bacteria to mammals, which have been well characterized, involve several coordinated steps.3–5 Although the latter steps differ significantly between bacteria and mammalian cells, BER is usually initiated by a DNA glycosylase that recognizes a base damage and cleaves the N-glycosyl bond between the base and the sugar moiety (damage-specific step). The resulting AP site is repaired and the DNA strand integrity is restored by a stepwise process (damage-general) that involves AP endonuclease, AP lyase, DNA polymerase and DNA ligase. This basic BER mechanism is complicated by the presence of distinct subpathways in mammalian systems. Biochemical evidence indicates that there is also an intricate network of protein-protein interactions involving numerous factors inside and outside of BER, which, along with post-translational modifications, is thought to play a key role in coordination of BER functions as well as in cross-link of BER with other cellular damage response processes.6,7 BER primarily targets relatively small base lesions, including oxidized, alkylated and deaminated bases. Many of these toxic and/or mutagenic lesions are produced through endogenous processes such as oxidation stress and lipid peroxidation. Recently, a great deal of interest has been focused on repair of such endogenous base damage since they impose constant threats to the integrity of the genome.8,9 Redundant glycosylase activities are often found for specific base lesions, suggesting the biological importance for their efficient removal. Another crucial role of BER is to repair AP sites,

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resulting from DNA glycosylase action or from spontaneous and chemically induced hydrolysis. AP sites are a major threat to genetic stability, due to their abundance in the cell and their mutagenic or lethal property. BER appears to be crucial for all organisms as evidenced by its preservation throughout phylogeny. BER has been shown to be important in counteracting the increased spontaneous mutation rates in bacteria and yeast.10 In animal models, BER is shown to be essential for development and viability. Gene knockout mice that lack key components of BER, e.g. the major AP endonuclease (APE1, also termed HAP1, APEX1 and REF-1) and DNA polymerase β (POLβ), are embryonically lethal.11 In humans, data are beginning to emerge that BER deficiency can be an important contributing factor of cancer susceptibility, as demonstrated by the recently revealed causal link between the inherited gene mutations of the human adenine-DNA glycosylase (MYH) and the increased risk for colorectal cancer.12,13 BER is also associated with other disorders such as neurodegenerative diseases14 and immune system imbalance.15,16 Although there are still many questions that remain unknown, considerable progress has been made in the last decade in understanding the specificity and mechanism as well as biological functions of many BER proteins. This has been greatly facilitated by the major advances in discovery of new enzymes through proteomics, determination of high resolution structures of repair proteins and damaged DNA, revealing of novel protein-protein interactions and protein cellular localization, construction of gene mutant models, and analysis of human BER capacity such as genetic polymorphisms. The details of each of these areas have been reviewed in many previous articles, and this chapter mainly gives an overview, with emphasis on the biochemistry of mammalian DNA glycosylases and AP endonucleases and their biological relevance in the prevention of genetic instability. 2. THE BER PATHWAY The BER pathway can be divided into five sequential steps: (1) excision of a modified or altered base; (2) cleavage of the resulting AP

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site; (3) processing of the blocking termini at the break; (4) repair synthesis; and (5) ligation of the broken strands (Fig. 1).3–5 This process can be reconstituted in vitro with a minimal requirement of four purified enzymes, indicating the relative simplicity of the basic operative mechanism of this pathway, as compared with the other excision repair pathway, NER. The first BER step is catalyzed by damage-specific DNA glycosylases. Based on whether they have an associated AP lyase activity, two types of glycosylases are present with different catalytic mechanisms. Monofunctional DNA glycosylases simply hydrolyze the N-C1 glycosyl bond between the target base and the sugar, generating an AP site. Human enzymes of this type include methylpurineDNA glycosylase (MPG, also termed AAG, APNG and ANPG), P

P

A T

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C P

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G C P

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T A

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C G P

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POL β XRCC1

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POL β and/or POL δ/ε PCNA , RFC, RPA

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OH P P

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APE 1 or POL β OH

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AP lyase A T

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Monofunctional glycosylase

Bifunctional glycosylase P

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P OL β

P OL β

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L IG1 P

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FEN1 P

L IG 3/XRCC1 P

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OH P P

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P OH

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Short-patch

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BER

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Long-patch

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BER

Fig. 1 BER subpathways in mammalian cells. The roles of core enzymes and accessory proteins are listed in Table 1.

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uracil-DNA glycosylase (UNG), single-strand selective monofunctional uracil-DNA glycosylase (SMUG1), thymine-DNA glycosylase (TDG), and MYH. As for bifunctional DNA glycosylases, after excising the base, they further cleave the phosphodiester bond 3 to an AP site with an AP lyase activity. This type of glycosylases can be divided into two classes, depending on their mode of action in processing the AP site. One class, as exemplified by human endonuclease III (NTH1) and 8-oxoG glycosylase 1 (OGG1), cleaves the AP site through β elimination, yielding a 3 -phospho-αβ-unsaturated aldehyde. While enzymes in the other class, such as endonuclease VIII-like (NEIL) homologs, carry out the so-called β,δ-elimination by further cleaving the deoxyribose residue, leaving a 3 -phosphate at the strand break. Biochemical evidence has revealed at least two major subpathways in BER, the short-patch (single nucleotide) and the long-patch (more than one nucleotide), differing in their repair patch sizes and some participating proteins (Fig. 1). An AP site produced by a monofunctional glycosylase is processed by an AP endonuclease, such as APE1, which cleaves the phosphodiester bond 5 to the AP site, yielding a 3 -hydroxyl and 5 -deoxyribose-5-phosphate (5 -dRp) residue. APE1 also uses its 3 -dRpase activity to process the product produced by a bifunctional glycosylase, i.e. a 3 -cleaved AP site, by excising the 3 -terminal 4-hydroxypentenal residue, thus providing a suitable primer for subsequent DNA polymerase-mediated reaction. Recently, using hNEIL1, an APE1-independent BER mechanism was reported in human cells,17 in which the removal of the 3 -phosphate terminus generated by NEIL1 is dependent on polynucleotide kinase 3 -phosphatase (PNKP), instead of APE1. PNKP possesses both 5 -kinase and 3 -phosphatase activities and is the primary 3 -phosphatase activity in mammalian cells. This mechanism could provide an alternative pathway, for certain DNA glycosylases like NEIL1, to the traditional subpathways in BER. In the short-patch BER, the gap is filled through templatedirected synthesis by POLβ which can also use its dRpase activity to remove any remaining 5 -dRp moiety at the strand break. The nick is sealed by the complex of DNA ligase 3 (LIG3) and X-ray

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cross-complementation protein 1 (XRCC1). In the long-patch BER, POLδ or ε as well as POLβ can catalyze the repair patch synthesis. Together with proliferating cellular nuclear antigen (PCNA) and replication factor C (RFC), a polymerase displaces the 5 dRp-containing strand and synthesizes a segment of DNA 2–12 nucleotides long. The resulting flap-like structure is removed by the structure-specific flap endonuclease 1 (FEN1) and the nick sealed by LIG1. It is noteworthy that POLι, one of the Y-family translesion DNA polymerases, was also demonstrated to have a 5 -dRpase activity and to be able to replace POLβ as an efficient short gap-filling polymerase in vitro.18 However, given the unique base incorporation specificity of POLι, its role in BER, if it exists, is likely limited to certain special circumstances.18 Both short- and long-patch pathways have been reconstituted in studies using DNA glycosylases and downstream proteins (e.g. Refs. 19 and 20). There is also limited evidence showing the existence of long-patch BER in vivo.21 It is still not clear how the cell determines which subpathway to be used, but the nature of the base substrates and the glycosylases involved, the types of termini at AP sites, and the specific protein-protein interactions may all be relevant determinants.17,22 It has been suggested that the long-patch BER repairs reduced or oxidized AP sites since they are not efficient substrates for the dRpase activity of POLβ. The functions of both the core and accessory proteins involved in BER are summarized in Table 1.

3. DNA GLYCOSYLASES 3.1. Overview In general, the bacterial DNA glycosylases were among the first to be discovered, followed by their corresponding activities sought in higher species. Many human DNA glycosylases have also been cloned, by taking advantage of functional complementation with their E. coli or yeast homologs or by sequence search through the genome database. For example, using a proteomics strategy, three

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Table 1.

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The Human Proteins and their Functions in the BER Pathway Core Proteins

Protein Name and Synonyms

Gene Name

Function in BER

N-methylpurine-DNA glycosylase (MPG, AAG, ANPG, APNG)

MPG

Major glycosylase for alkylated bases, hypoxanthine, and etheno bases

Uracil-DNA glycosylase (UNG)

UNG

Major glycosylase for removal of U

Single-strand selective monofunctional UDG (SMUG1)

SMUG1

Major backup activity for UNG; 5-HmU-DNA glycosylase

Methyl-binding domain glycosylase 4 (MBD4, MED1)

MBD4

Glycosylase activity for U:G or T:G in U/TpG sequence

Thymine-DNA glycosylase (TDG)

TDG

U:G and T:G glycosylase; εC-DNA glycosylase

8-oxoG-DNA glycosylase (OGG1)

OGG1

Major glycosylase for 8-oxoG:C and Fapy-G

Endonuclease III (NTHL1, NTH1)

NTHL1

Major activity for oxidized pyrimidines; Tg-DNA glycosylase

MutY homolog DNA glycosylase (MYH)

MYH

Major glycosylase for excising adenosine from 8-oxoG:A

Endonuclease VIII-like 1 (NEIL1)

NEIL1

Activity for oxidized purine and pyrimidine bases, possibly the major backup activity for NTH1

Endonuclease VIII-like 2 (NEIL2)

NEIL2

Glycosylase activity mainly for cytosine base damage

Endonuclease VIII-like 3 (NEIL3)

NEIL3

Glycosylase activity for fragmented or oxidized bases

AP endonuclease (APE1, HAP1, APEX1, REF-1)

APEX1

Major AP endonuclease; Removal of 3 -blocking groups

AP endonuclease 2 (APE2)

APEX2

Minor AP endonuclease

Polynucleotide kinase phosphatase (PNKP)

PNKP

Removal of 3 -phosphate damage; Involved in APE1-independent BER

Polymerase β (POLβ)

POLβ

Major gap-filling DNA polymerase; Also dRpase activity (Continued)

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Table 1.

(Continued)

Protein Name and Synonyms

Gene Name

Function in BER

Polymerase δ (POLδ)

POL δ

DNA polymerase involved in long patch-BER

Polymerase ε (POLε)

POL ε

DNA polymerase involved in long patch-BER

Flap endonuclease 1 (FEN1)

FEN1

Major activity for removing 5 -overhanging flaps in long-patch BER

DNA ligase 1 (LIG1)

LIG1

DNA ligation

DNA ligase 3 (LIG3)

LIG3

DNA ligation (LIG3/XRCC1)

Accessory Proteins Proliferating cellular nuclear antigen (PCNA)

PCNA

“Sliding clamp” for POLs δ and ε and interacts with many BER proteins

Replication factor C (RFC)

RFC

A “clamp loader” that loads PCNA onto DNA

Replication protein A (RPA)

RPA

A single-strand DNA binding protein that interacts and stimulates later steps of long-patch BER

X-ray repair cross complementation group 1 (XRCC1)

XRCC1

A scaffold protein that interacts with many BER proteins

Poly(ADP-ribose) polymerase 1 (PARP1)

PARP1

Catalyzing poly(ADP-ribosyl) action and stimulating strand displacement synthesis

orthologs of E. coli endonuclease VIII (Nei), NEIL1, NEIL2 and NEIL3, have recently been identified from human cells.23 Although BER is preserved throughout phylogeny, many glycosylases are not ubiquitously distributed. An illustration of this is the unique presence of SMUG1 in higher species and the lack of Mug in Saccharomyces cerevisiae. Several DNA glycosylases are present as isoforms in both the nucleus and mitochondria, as a result of alternative splicing of their mRNA.24 For instance, two alternatively spliced isoforms are found

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for UNG, UNG1 (mitochondrial form) and UNG2 (nuclear form), which contain the respective subcellular targeting signals in their regions.25 Likewise, the transcripts for OGG1, NTH and MYH are also alternatively spliced. A full, functional BER pathway is present in mitochondria, at least for oxidative DNA damage,26 which is at higher levels in mitochondrial DNA. The repair specificity of BER depends on the recognition specificity of each of the individual DNA glycosylases. Although it is still poorly understood as to how DNA glycosylases detect a base damage in DNA, they are expected to scan a vast amount of bases in the genome before initiating BER. A glycosylase exhibits its substrate specificity by recognizing a subset of DNA base lesions. For some glycosylases, a primary substrate could be described, while for others, multiple functions of similar importance may be the case. Some DNA glycosylases, UNGs in particular, have very narrow substrate specificity. However, most glycosylases, such as MPG, NTH1 and NEIL1, are able to excise a relatively broad range of modified bases.23,27,28 DNA glycosylases are, in general, specific for either purine- or pyrimidine-derived base lesions, with some of them exhibiting activities for both types of lesions. For example, for the two families of glycosylases that remove oxidized bases, OGG1/Fpg and NTH1/Nth, the former primarily recognizes oxidized purines bases, while the latter mainly acts on oxidized pyrimidines.28,29 A big challenge in understanding the BER function is to clarify the in vivo role of individual DNA glycosylases. This aspect will be discussed in detail later. Cellular glycosylase activities could be modulated by multiple factors, including gene expression, protein stability, protein-protein interactions, post-translational modifications, and the presence of activators or inhibitors. In vivo, a plethora of DNA lesions can be formed by a given compound or exposure, and often no single lesion can be solely responsible for the observed deleterious biological endpoints. Therefore, complex repair mechanisms involving multiple glycosylases or repair pathways are often required for a given exposure. It is known that, for repairing ionizing radiation-induced oxidative DNA lesions, several DNA glycosylases are responsible,28 such as OGG1, NTH1 and NEILs

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with backup from both NER and MMR. This would ensure free or low levels of highly mutagenic lesions in the genome. Substrate overlap between different glycosylases is another common theme in BER, which may signify the biological importance of the target lesions and the necessity for redundancy of repair. For example, several mammalian glycosylases can excise the uracil residue from DNA: UNG, SMUG1, MBD4, TDG and NEIL1/NEIL2. In cells, these enzymes are likely to play distinct or specialized roles in uracil removal. The primary role of UNG2 may be replicationassociated, excising uracil misinserted opposite adenine during DNA synthesis.30 SMUG1 has been suggested to target uracil from deamination of cytosine.31 Both MBD4 and TDG may preferentially remove uracil generated at CpG sequences.32,33 NEIL1 also possesses excision activity for mismatched uracil, especially in the U:C context.34 Understanding substrate specificity and catalytic mechanism of DNA glycosylases has been greatly facilitated by structural biology approaches.35–38 To date, high resolution structures of numerous glycosylases of various sources, or their complexes with substrate DNA, have been determined. Based on these structures, mutant proteins have commonly been engineered. The experimental structural data also provide a framework for computational studies such as molecular dynamics simulations, which permit test of both substrates or non-substrates (e.g. Refs. 39 and 40). Distinct superfamilies of DNA glycosylases have been designated according to their structural similarity; however, the members within a superfamily could have very divergent sequences. For example, a number of DNA glycosylases, including AlkA, MutY, Nth and OGG1, contain the helix-hairpin-helix (HhH) motif, and belong to the HhH superfamily.3 Another set of this is the uracil-DNA glycosylase superfamily which includes UNG (Ung) TDG (Mug) and SMUG1, all of which possess the same fold and have evolved from a common ancestor.41 Within each superfamily, members usually exhibit smaller variations in their binding site structure or catalytic elements, which, in turn, constitute the molecular basis for their different substrate specificities.

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The structural biology of Udg and Mug is often compared. Udg (and UNG) has an exquisite substrate specificity for uracil in both ds and ssDNA, but not for thymine, while Mug (and TDG) posesses a broad substrate range, including both uracil and thymine when paired with guanine. Despite their low sequence homology (∼ 10%), the two proteins share a common fold (Fig. 2) and interact with DNA

Fig. 2 Top: Comparison of E. coli Mug (left) and HSV-1 Udg (right) folds. The secondary structure homology between Mug and Udg is 77% as identified by SSAP (highlighted in cyan); residues contributing to the binding pockets are in red. Bound sulphate ions in native crystals are colored yellow. Bottom: Comparison of binding pockets and catalytic residues in Mug and Udg. Residues forming the side of the pyrimidine pocket, including the catalytic Asp or Asn, are colored red; the Phe residues that provide the aromatic wall of the pocket are in magenta, and the residue forming the base of the pocket is in blue. The residues in the motif carrying the catalytic His/Asn and the Leu implicated in flipping are colored green. (Reprinted with permission from Elsevier, Copyright 1998.43 )

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substrates through conserved motifs.42–44 The differences in their active site architecture are proposed to be responsible for the divergent substrate specificity between these two enzymes. As shown in Fig. 2, in the active site of HSV1 Udg, a “barrier” residue (Y90) is present with its side chain sterically excluding the 5-methyl group of thymine. The tight binding to uracil by Udg is a result of specific hydrogen-bonding interactions involving N147 and bound uracil, which, in contrast, are unable to be formed for cytosine. The Mug active site is structurally similar to that in Udgs, but it is also larger with differences in certain key amino acid residues, turning it into a non-specific pyrimidine binding pocket. For example, the equivalent “barrier” residue Y90 is replaced by a sterically smaller glycine, which allows thymine binding.39,43 There are also no specific contacts with bound bases. The Mug binding pocket can also readily accommodate 3,N4 -ethenocytosine (εC) in its hydrophobic space at the bottom. εC is a different type of substrate for Mug/TDG enzymes, as compared to U(T):G mismatches.45,46 Based on the structural and biochemical data, the mechanisms of substrate binding and excision are proposed differently for different types of DNA glycosylases. In general, monofunctional glycosylases use a base flipping mechanism and DNA intercalation strategy to dock a target base into the active site pocket and to position a nucleophile, e.g. an activated water molecule as shown in both Mug and Udg (Fig. 2), for in-line attack on the scissile bond. The catalytic mechanism usually involves nucleophilic attack on the sugar C1 carbon of the modified base to destabilize and cleave the N-glycosyl bond. Bifunctional DNA glycosylases utilize an amine nucleophile, such as a deprotonated Lys, from their own structure. The nucleophile forms a covalent intermediate (Schiff base) with the deoxyribose and displaces the base, which is followed by transformations of the deoxyribose, resulting in strand cleavage on the 3 -side of the modified base and dissociation of the covalent intermediate. Such a proposed mechanism is supported by the borohydride-trapping experiments, which showed that only bifunctional glycosylases are covalently trapped to their substrate DNA.47

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3.2. Excision of Alkylated Bases Alkylating agents consist of a large variety of environmental chemicals and therapeutic drugs, some of which are potent mutagens and/or carcinogens.48 They are also continuously produced in vivo. These agents react with the nucleophilic centers of DNA bases, such as oxygen and nitrogen moieties, to form different classes of base damage. The main chemical mechanism of DNA alkylation is a SN 2 and SN 1 type reaction, in which an alkylating agent reacts with the electron-rich regions of the base residue. The N7 of guanine and N3 of adenine are the most reactive sites. Methylation may occur through the ubiquitous cellular methyl donor, S-adenosylmethionine (SAM).49 Alkylated bases are either mutagenic or toxic with few exceptions such as 7-methylguanine (7-meG). 3-methyladenine (3-meA) can block DNA synthesis, thus a potentially lethal adduct. These bases and many others caused by Nmethylation or ethylation are primarily removed by the MPG family of glycosylases. Bifunctional alkylating agents are also capable of forming exocyclic adducts. Among the most common ones are etheno adducts of dA, dC and dG.48 When examined in various systems, all the exocyclic adducts tested are mutagenic,50 owing to the blocking of the Watson-Crick hydrogen bonding area of a base by the exocyclic ring. Many 5-membered and some 6-membered exocyclic ring adducts are excised by two families of monofunctional glycosylases, the MPG/AlkA and TDG/Mug proteins.27 Similar activities on exocyclic adducts have also been found for other glycosylases, including SMUG1, MBD4 and NEIL1; however, the biochemical details are not fully characterized.

3.2.1. MPG In E. coli, two 3-meA-DNA glycosylases have been identified and designated Tag and AlkA. Tag is a constitutive enzyme with strict substrate specificity toward 3-meA, while AlkA has a broader substrate range than Tag and is inducible by alkylation adaptation.

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Despite the fact that they are functional homologs, MPG and AlkA belong to different structural families and have distinct catalytic mechanisms. However, the substrate specificities of the two enzymes are similar, broad and appear to be ever-growing. Independent of enzyme sources, these enzymes repair a number of alkylated bases, including N7-alkylguanine, N3-alkyladenine, O2 alkylthymine and O2 -alkylcytosine, and N3 and N7 derivatives formed by chloroethyl ethyl sulfide and mustard compounds.51 Over the years, their substrate ranges have gradually been expanded to a variety of other lesions, including 1,N6 -ethenoadenine (εA), 1,N2 -εG, N2 ,3-εG, 1,N6 -Hm-εA, 1,N6 -ethanoadenine (EA), N2 ,3-EG, 5-formyluracil (5-FoU), 8-oxoG, fragmented thymine, and all three deaminated purine bases, hypoxanthine, xanthine, and oxanine.3,27 Structural studies show that their permissive active sites are the basis for recognizing such a chemically diverse group of damaged bases,52 and that MPG can discriminate against the bases that either have a 6-amino group (adenine) or a 2-amino group (guanine), leading to a low level of their excision. Although the biological implications of repair of many of these substrates are not understood, the importance of these enzymes in bacterial and budding yeast in protecting against alkylating agents has been well documented.3,4 However, the role of MPG in mammalian systems is shown in complex patterns as reported in different studies. It is surprising that Mpg-knockout mice did not show any overt phenotypic abnormalities or significant increase in the spontaneous mutation rate,53,54 although increased mutations were observed when exposed to methyl methanesulfonate. When these mice were challenged with vinyl or ethyl carbamate which induces etheno adducts, levels of εA became significantly higher and persisted longer in the DNA, indicating the cellular removal of εA by MPG.55,56 In fact, using cell-free extracts from tissues of the Mpg−/− mice, MPG is the primary glycosylase for εA.53,57 It is puzzling, though, that the increased levels of these highly mutagenic adducts in liver and lung DNA of Mpg−/− mice were not paralleled by increased tumorigenesis in these organs. Interestingly, overexpression of MPG has led to deleterious effects, which can be attributed to

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the accumulation of highly mutagenic/cytotoxic AP sites as a result of the increased removal of alkylation damage (imbalanced BER).58,59 3.3. Excision of Deaminated Bases and Base Mismatches Uracil is one of the most abundant base lesions occurring in genomic DNA. Approximately 100 uracil residues are expected to form per human cell per day, which result from hydrolytic deamination of cytosine, yielding U:G mismatches.60 Such a process occurs more rapidly in ssDNA than in dsDNA. Uracils also occur in DNA by misincorporation of dUMP during DNA replication, generating U:A base pairs. Uracil in U:G pair would be mutagenic by causing a G:C to A:T transition. In contrast, a U:A pair is not miscoding, but may affect specific binding of other nuclear proteins to DNA. As noted above, a number of mammalian UDG activities have been identified and each may play a distinct cellular role in the removal of uracil. T:G mismatches can originate from spontaneous deamination of 5-meC in the most commonly methylated sites in the mammalian genome, 5 -CpG. Methylation at the CpG sites is an important regulator of gene transcription, and is involved in human carcinogenesis. T:G could contribute to as much as 30% of all human germline point mutations.61 Although the exact nature of the in vivo mechanism(s) that prevents this to happen is far from clear, how the cell carries out active demethylation and repairs deaminated base lesions has been a topic of considerable interest. MBD4 and TDG are two enzymes that are involved in the repair of these CpG-located base lesions. 3.3.1. UNG As the first discovered DNA glycosylase,2 it is probably the most extensively studied glycosylase. UNG is a ubiquitous enzyme and its primary role is removal of uracil residues from both dsDNA and ssDNA.62 UNG has a high turnover number on these substrates. UNG seems to be the primary glycosylase responsible for the post-replicative removal of uracil misincorporated opposite adenine as it localizes at replication forks and associates with replication

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proteins PCNA and RPA.30 UNG also excises 5-fluorouracil (5-FU) and 5-hydroxyuracil (5-OHU). When hUNG was tested for activity against γ-irradiated DNA, three cytosine products were excised, which are 5-OHU, isodialuric acid and alloxan.63 The end result is that all the UNG substrates are uracil analogs. This unusual narrow substrate specificity replies on its rigid preformed binding pocket that accommodates uracil but excludes other bases through steric hindrance and hydrogen bonding interactions, as described above. UNGs from a variety of sources can be inactivated by a small peptide called uracil-DNA glycosylase inhibitor (Ugi), encoded by the bacteriophage PBS2.64 Ugi inhibits these enzymes by forming essentially irreversible UNG·Ugi complexes. Activity of other glycosylases such as Mug/TDG is not inhibited by Ugi. UNG plays an important role in the immune system. More specifically, the Ung-knockout mice exhibit an altered pattern of somatic hypermutation and reduced class switch recombination of immunoglobulin genes.15 These mechanisms may account for the increased risk of B-cell lymphomas in these mice in the later stage65 (see Section 6.1). In humans, UNG deficiency is associated with life threatening hyper-IgM syndrome because of the similar alterations.16 3.3.2. SMUG1 This enzyme was isolated in 1999 as a second UDG activity, by using the “proteomics” approach.66 Two properties of this protein may be important for understanding its potential physiological role. One is that SMUG1 is able to efficiently excise uracil from ssDNA in vitro, although its primary substrate is dsDNA.67 Therefore, it may serve as a backup for UNG2. The other unique feature is that SMUG1 has not been found in lower species such as bacteria and yeast, suggesting that it may have evolved in higher organisms to counteract mutations that arise from cytosine deamination in DNA. A 5-hydroxymethyluracil (5-HmU)-DNA glycosylase activity was earlier identified in mammalian cells68 and later found to reside in SMUG1.69 5-HmU is a major thymine-derived DNA lesion

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produced by ionizing radiation and endogenous oxidative stress. SMUG1 has also been reported to excise several uracil derivatives with an oxidized substituent at C-5, including 5-OHU and 5-FoU.70 Despite the lack of significant sequence homology, SMUG1 shares common structural folds and active site structure with the functionally related UNG and TDG.67 3.3.3. MBD4 MBD4 (also MED1) was recently identified by using two different screening approaches.71,72 It contains two important domains: the N-terminal region is the methyl-CpG binding domain that preferentially binds to 5-meC:G or T:G mismatches, and the C-terminal glycosylase domain that is much more active in vitro on uracil or thymine at deaminated CpG or 5-meCpG sites.73,74 The MBD4’s unusual sequence context preference suggests that it may have evolved to process base lesions associated with methylated CpG sites. Indeed, Mbd4-knockout mice exhibit an increased frequency of CpG to TpG mutations and an increased risk of tumor formation in the gastrointestinal tract when crossed with mice bearing a defective adenomatous polyposis coli (Apc) gene.75,76 Other substrates identified for MBD4 are 5-FoU paired with G, T from T:O6 -meG, and εC from εC:G.77 MBD4 interacts with human MMR protein MLH1,72 which may link the pathways of BER and MMR. The recombinant MBD4 proteins from human and chicken were shown in one study to excise 5-meC from 5-meC:G pairs in the hemimethylated DNA.78 However, an independent study showed that human MBD4 had no 5-meC glycosylase activity toward either hemimethylated or fully methylated substrates.73 The reason for such a discrepancy is not clear. Similar searches for demethylating glycosylase activity in hTDG have also led to conflicting results regarding whether TDG has a 5-meC-DNA glycosylase activity.79–81 It should be noted that a partially purified human 5-meC glycosylase activity is active for fully methylated CpG sites,82 but its identity has yet to be determined. Apparently more studies are needed to clarify these issues.

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3.3.4. TDG Human TDG was initially purified and cloned as T:G(U:G)-DNA glycosylase.83,84 This enzyme and its E. coli homolog Mug were later found to have an efficient glycosylase activity toward εC,45,46 a highly mutagenic lesion formed by carcinogens such as vinyl chloride and lipid peroxidation products. Therefore, TDG/Mug are considered enzymes that act on both mismatch and alkylation damage. In fact, the uracil and εC excision activity are the only two efficient ones in all thymine-DNA glycosylases tested so far.81 The TDG/Mug enzymes have broad substrate spectra.27 TDG may play a specialized role in removing thymine and uracil arising from the CpG sites, since it exhibits the highest catalytic efficiency for those lesions located in these sequences.33 TDG also excises thymine from T:O6 -meG, T:2-amino-6 (methylamino)purine, as well as 5-FU, 5-HmU, 5-bromouracil (5-BrU) and thymine glycol (Tg) paired with guanine.33,81,85 TDG/Mug are able to remove several εC analogues, including εC, 8-Hm-εC, and ethanocytosine (EC) but only Mug removes 1,N2 -εG.27,86 Structural work on E. coli Mug and human TDG has provided insight into the similar substrate specificity and catalytic mechanism of the two enzymes, which is based on the strong conservation of tertiary structures as well as key residues between their common catalytic core domain.39,43,87 As described in Section. 5.1, TDG interacts with a number of proteins inside and outside BER. Its unusual interaction with transcription factors or co-factors suggests that it may be involved in gene regulation. TDG is actually shown to be indispensable for early embryonic development. 3.4. Excision of Oxidized and Fragmented Bases Oxidative DNA damage can be caused by reactive oxygen species (ROS), including superoxide radical (•O− 2 ), hydrogen peroxide (H2 O2 ), and hydroxyl radical (OH). They are formed either through endogenous metabolisms or by environmental sources such as ionizing radiation. A variety of DNA lesions in this class are primarily processed by BER, including various oxidized bases, oxidized AP

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sites, and single strand breaks.88–90 Among them, 8-oxoG, as one of the most frequent endogenous base lesions, has gained much attention in recent years. It is highly mutagenic by forming adenine mismatches (8-oxoG:A) that cause G:C to T:A transversions, which is a common somatic mutation associated with several types of human cancer. BER is the major mechanism for removing 8-oxoG from the genome. The prototype of such repair is shown in E. coli “GO system,” in which three enzymes act in concert to protect against the mutagenic effect of 8-oxoG.91 Formamidopyrimidine-DNA glycosylase (Fpg/MutM) excises 8-oxoG opposite cytosine in DNA, while MutY removes adenine from the 8-oxoG:A pair. In the latter case, when an 8-oxoG:C pair is restored, Fpg/MutM can further excise 8-oxoG. The third enzyme, MutT, removes oxidatively damaged dGTP from the nucleotide pool with its 8-oxo-dGTPase activity. Mammalian homologs of these three enzymes are OGG1, MYH and MTH, respectively, and they process 8-oxoG in a similar manner.28,88 With the addition of recently discovered NEIL proteins that also act on oxidized bases, it is evident that repair of oxidative lesions are mediated by multiple DNA glycosylases that have different specificities. In principle, purine-derived lesions are primarily removed by OGG1 and NEIL1, and pyrimidine lesions are excised by NTH1, NEIL1/NEIL2. Other glycosylases such as MPG and SMUG1 can also excise several oxidized base lesions, as mentioned above. 3.4.1. OGG1 E. coli Fpg and hOGG1 have similar substrate specificity but share no amino acid sequence homology. Although both are bifunctional glycosylases, Fpg possesses an intrinsic β,δ-lyase activity, whereas OGG1 has only a β-lyase activity. Many alternative splice variants for the OGG1 gene have been reported, and splicing at the C-terminal region forms two groups of variants, α-OGG1 and β-OGG1.90 The Nterminus of this gene contains a localization signal, which directs the transfer of β-OGG1 to mitochondria. Human OGG1 gene is located in chromosome 3p25, which is frequently lost in many types of cancer.

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OGG1 primarily excises oxidative and ring-fragmented purine derivatives, including 8-oxoG, Fapy-G and 8-OH-A when paired with cytosine in DNA.88,90 Unlike Fpg, it does not act on Fapy-A. OGG1 is the major mammalian excision activity for 8-oxoG opposite cytosine, as shown by the experiments using cell extracts from Ogg1knockout mice.92 Inactivation of this enzyme in these animals causes accumulation of 8-oxoG in the genome93,94 as well as increased risk of tumor formation in aging mice,94 as discussed below. 3.4.2. NTH1 In E. coli, Nth is the prototype enzyme that recognizes oxidized and fragmented pyrimidines and its homologs are highly conserved among species. This family of enzymes has broad substrate specificity.28,88 The principal substrates for human NTH1 are oxidized pyrimidines that include Tg, urea, 5,6-DHU, 5-OHU and 5-FoU. NTH1 also excises Fapy-G and Fapy-A.28,88 Data based on cell-free extracts from Nth1−/− mice show that this enzyme is the major excision activity for oxidized pyrimidines.34 There are backup activities, mainly for Tg:G, in tissues of these mice,95 which may explain why these animals did not exhibit overt phenotypic abnormalities. One backup glycosylase in Nth-knockout mice is NEIL1,34 which has similar substrate specificity as NTH1. 3.4.3. NEILs E. coli Nei, a member of the Fpg/Nei family, efficiently excises a number of oxidized pyrimidine bases, and shares significant substrate specificity with Nth and Fpg.96,97 Recently, three human orthologs of the nei gene, designated NEIL1, NEIL2 and NEIL3, were identified.98–100 NEIL1 and NEIL2 have been cloned and expressed, both of which are bifunctional glycosylases that exhibit a β,δ-elimination mode. Both enzymes use an N-terminal Pro as the active site. hNEIL1 is a structural and functional homolog of E. coli Nei, excising a wide range of oxidized purines and pyrimidines such as Tg, 5-OHU, 5-OHC, 5-FoU, 5-HmU, 5,6-DHU, 5,6-DHT,

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Fapy-G and Fapy-A.98–101 The 8-oxoG activity of NEIL1, primarily toward 8-oxoG:C, is very weak relative to other substrates. NEIL2 appears to have a narrower substrate specificity, mainly excising cytosine-derived oxidative lesions, with its primary activity against 5-OHU.98,99 It also recognizes the 8-oxoG in DNA bubble structures.102 Another difference between these two enzymes is that the expression level of NEIL1 increases in the S-phase, whereas NEIL2 expression is not cell cycle dependent. In a study using RNAi, embryonic stem cells deficient in NEIL1 are hypersensitive to ionizing radiation,103 indicating the importance of this enzyme in cellular defense of oxidative DNA damage. 3.4.4. MYH E. coli remove adenine-DNA glycosylase (MutY), and its mammalian homolog MYH are the major glycosylases that remove adenine from A:8-oxoG mismatches, thereby preventing G:C to T:A transversions. MutY/MYH are unique among DNA glycosylases inasmuch as they recognize a mismatch between 8-oxoG and a normal adenine but do not remove the damaged base.104–106 Besides A:8-oxoG, these enzymes also excise adenine from both A:G and A:C mismatches with less efficiency.104 hMYH physically interacts and co-localizes with proteins involved in DNA replication foci, suggesting a role of this protein in replication-coupled repair.107,108 hMYH has been a focus of research for the last few years since it was the first DNA glycosylase shown to be associated with genetic predisposition for cancer as a result of inherited biallelic mutations in its gene12,13 (see Section 6.2).

4. AP ENDONUCLEASES AP sites arise from spontaneous depurination/depyrimidination or from DNA glycosylase reaction as repair intermediates. The spontaneous depurination, as measured by Lindahl109 under in vitro conditions and extrapolated to a cellular genome, could produce AP

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sites at an estimated rate of ∼10,000 per day per cell. The occurrence of such AP sites in mammalian genomic DNA represents one of the most frequent and important endogenous lesions. AP sites are potentially mutagenic due to its noncoding nature or lethal by blocking DNA replication and transcription. Processing of AP sites in bacteria, yeast and mammalian cells has been extensively studied.110–112 Two 5 -AP endonucleases are present in E. coli, exonuclease III (Exo III) and endonuclease IV (Endo IV). Exo III and APE1 are homologous and are the major AP endonuclease activity in E. coli and human cells, respectively. Recently, APE2 was identified from human cells with a weak AP endonuclease activity113,114 and partly in association with PCNA in the nucleus.114 APE2 is also found in the mitochondria. The in vivo role of APE2 is not known. As described before, APE1 functions in both short- and longbatch BER by hydrolyzing AP sites (AP endonuclease activity) and processing 3 -blocking groups (3 -repair diesterase and phosphatase activity). APE1 also processes uncleaved modified bases such as exocyclic para-benzetheno adducts115 or oxidized bases, e.g. 5,6-DHU and 5-6-DHT,116 but such an action does not lead to BER since no base release takes place. Besides repair activities, this multifunctional protein also has several repair-unrelated functions, such as acting as a redox factor (REF-1) that is involved in transcription factor regulation.117 Its REF-1 and DNA repair activities can, in part, be distinguished biochemically.118 Although stable APE1 mutant cell lines have not been isolated, down-regulation of APE1 using RNAi in several human cell types activated apoptosis, which was correlated with accumulation of abasic DNAdamage.119 Disruption of APE1 in mice results in embryonic lethality,120,121 while APE1 heterozygous mice had a significantly elevated spontaneous mutation frequency.122 These findings indicate that APE1 plays a significant role in counteracting spontaneous mutagenesis. Interestingly, AP endonuclease-deficient yeast or E. coli stains are not lethal, but sensitive to alkylating or oxidizing agents that produce glycosylase substrates which, in turn, become the source of endogenous AP sites.112 This is supported by

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the observation that spontaneous mutation was decreased in the double mutant lacking both the major AP endonuclease (APN1) and 3-meA-DNA glycosylase 1 (MAG1) in yeast.123 Recent studies have shown that single nucleotide polymorphisms (SNPs) are present in APE1 identified in the human population.124 Four of seven variants in the repair domain of hAPE1 exhibit reduced AP endonuclease activity. However, the association of these variants with increased cancer risk has yet to be determined. 5. REGULATION OF BER FUNCTIONS Regulation of BER functions can occur at both the gene and protein levels. Recently, a great amount of data on protein-protein interactions have been reported, which have enhanced our understanding with regard to what governs the BER sequential reactions and how biological systems modulate and coordinate BER in damage response. 5.1. Protein-Protein Interactions Single BER reactions performed in vitro generally do not require protein complex formation or co-factors. However, recent biochemical studies have shown that, in order for BER enzymes to perform sequential and efficient reactions, protein-protein interactions are required at each major step.6,7 This has led to the “hand-off” or “passing the baton” model that explains the mechanism for coupling of these steps in BER.125,126 In addition to enhancing the efficiency of the pathway, such interactions may be important for avoiding cellular exposure to toxic or highly reactive intermediates, such as AP sites and single-strand breaks. BER could also be coordinated with the other major repair pathways such as NER and MMR. In addition, BER has been shown to link with transcription, replication and cell cycle regulation. Figure 3 highlights the current knowledge of such a complex network of protein-protein interactions in BER. Many protein-protein interactions can directly modulate the catalytic activity of the BER enzymes, for instance, the stimulation of

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Fig. 3 Summary of a network of identified protein-protein interactions in BER. Solid and dashed lines indicate physical and functional interactions, respectively. Solid lines without arrows means physical interaction without functional effect. Arrows indicate functional effects of one protein on the other: closed arrow, stimulation; open arrow; inhibition; closed arrow with a rectangle, both stimulation and inhibition reported). APTX, ataxia-related protein; E6, papillomavirus E6 protein; ERα, estrogen receptor α; YB-1, Y box-binding protein; WRN: Warner syndrome protein. Courtesy of Fan and Wilson.7

glycosylase activities by AP endonucleases.6,7 In the past few years, many researchers have investigated both the functional and physical interactions between these two types of enzymes. The first reported case is the enhancement of hUNG activity by hAPE.127 From the combined crystallographic and biochemical data, it was proposed that the mechanism for such enhancement is the forced release of hUNG by APE1 from its product AP site. Later studies have shown that similar “coupling” occurs with many other glycosylases.6 Protein-protein interactions may play an important role in protein recruitment and reaction coordination in BER. For instance, human XRCC1, a scaffold protein required in BER, interacts with and stimulates multiple human DNA glycosylases such as OGG1, NTH1, NEIL1/NEIL2 and MPG.6,7 It also interacts or forms complexes with downstream proteins at each of the major steps, including APE1, POL β, PARP1 and LIG3. These data suggest that XRCC1 is recruited to sites of DNA damage by glycosylases and then

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coordinates subsequent BER steps and possibly modulates the activity of the enzymes involved. Many biochemical studies have shown that BER and NER or MMR are closely related via multiple protein-protein interactions.6,7 DNA glycosylases have been found to interact with specific NER or MMR proteins. For example, MPG and TDG were found to interact with NER components, the RAD23 proteins128 or XPC-23B,129 respectively. In addition, BER and NER or MMR could cooperate through interactions mediated by accessory proteins such as PCNA and RPA. Both proteins are involved in the latter replication steps of the three repair pathways. BER proteins are also found to be functional partners with numerous interactive proteins involved in other important cellular processes. For example, human TDG interacts with several transcription factors or cofactors, including transcriptional coactivator CBP, retinoic acid receptor (RAR), retinoid X receptor (RXR), thyroid transcription factor-1 (TTF-1), and estrogen receptor α (ER α).6,7 These data suggest that TDG is not only a repair enzyme, but could also function as a co-regulator in gene expression. 5.2. Protein Post-translational Modifications Although data are limited, post-translational modifications of BER proteins seem to play an important role in regulation of enzyme activities and protein-protein interactions. Several types of modifications have been reported for the BER proteins, including phosphorylation, acetylation, sumoylation and nitrosylation.7 Some of these modifications cause functional consequences such as altered enzymatic activity or protein-protein interaction. Human OGG1, for example, has been shown to be phosphorylated, both in vitro and in vivo, with enhanced 8-oxoG activity.130,131 Another illustration is the modification of human TDG by small ubiquitin-like modifiers (SUMOs), which can lead to a dramatic conformational change of TDG that facilitates its dissociation from the product AP site and coordinates the transfer of the AP site to APE1. Therefore, sumoylation of TDG potentiates the APE1 stimulatory effect on the TDG

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activity.132,133 Recently, a crystal structure of the central region of TDG conjugated to SUMO-1 was solved,87 providing a structural basis for such mechanism.

6. RELEVANCE TO CANCER SUSCEPTIBILITY 6.1. Animal Models Although the properties of many BER proteins have been well characterized, their biological role in suppressing tumorigenesis is still largely unknown. In the last 10 years, development in targeted gene deletions of the major BER components has provided an excellent tool to study: (1) the biological importance of specific repair enzymes as well as specific DNA lesions; and (2) the backup repair mechanisms. Interestingly, gene deletion of BER proteins in the damage general step, such as APE1, POLβ, FEN1, LIG1 and XRCC1, all resulted in embryonic lethality in mice.11 One explanation for this is that these proteins are critically involved in pathways of cellular development, in addition to repair. The other reason could be that disruption of these latter steps in BER results in intolerable levels of toxic and mutagenic intermediates such as AP sites and dRp moieties. In contrast, numerous viable DNA glycosylase deficient mouse models have been generated since the first glycosylase knockout mice Mpg−/− was reported in later 90s.53,54 Based on genetic studies in E. coli and yeast, it was earlier expected that these knockout models would demonstrate elevated mutation rate as well as increased cancer risk. Although, as expected, higher levels of base lesions or mutation rates were usually observed in cells, it was somewhat surprising that many animals such as Mpg−/− and Nth1−/− were viable, fertile, and showing no overt phenotypic abnormalities such as increased tumor incidence.134 Existence of back-up repair, either from a different glycosylase(s) or an alternative repair pathway(s), was considered to be the reason. In addition, translesion DNA polymerases could also bypass lethal or promutagenic base lesions. Nevertheless, these models provided a genetic approach for

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defining enzyme specificity and backup activity in cells (e.g. Refs. 34 and 57). Later efforts on gene knockout have actually revealed that certain glycosylase knockout mice are prone to tumorigenesis. The first evidence of association of a glycosylase deficiency with spontaneous malignancy was from the Ung−/− mice.65 In addition to an increased level of uracil in their genome, these mice developed B-cell lymphomas after 18 months of living. Mbd4−/− mice, when combined with a defect in the adenomatous polyposis coli (Apc) gene, also exhibited increased incidents of gastrointestinal cancers with CpG to TpG mutations in the Apc gene, confirming that MBD4 functions in correcting mutations at CpG sites.75,76 Spontaneously developed lung cancers have also been obtained in aging Ogg1−/− mice.94 The double knockout Myh−/− /Ogg1−/− mice exhibited an increased incidence of tumors, predominantly those from the lung, small intestine and ovary.135,136 In these studies, in addition to accumulation of 8-oxoG, G:C: to A:T transversions were found in 75% of the lung tumors at an activating hot spot in K-ras, codon 12, but none in adjacent normal tissues. Malignant lung tumors were also increased in the Myh, Ogg1, Msh2 triple gene knockout mice.135 Msh2 is a mismatch repair gene and is also involved in the repair of 8-oxoG. In general, double or triple knockouts are more powerful in generating phenotypic abnormalities since these genes may act synergistically in response to certain types of DNA damage. These findings also indicate that unrepaired oxidized DNA purines, such as 8-oxoG, can play a causative role in tumorigenesis. 6.2. BER and Human Cancer Risk Although mutations in BER repair genes and low activity of BER proteins have been found in tumors, until recently, no inherited deficiency in the BER pathway was found to be causally linked to any human disorders. Single nucleotide polymorphisms (SNPs) of DNA repair genes have been known to cause reduced repair activities, which could be associated with a risk of developing cancer. A significant advance

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in understanding of the biological role of BER is the recent discovery of the association of SNPs in the MYH gene with human colorectal adenomas and carcinomas.12,13 MAP (MYH-associated polyposis) is designated to describe a syndrome associated with biallelic-inherited mutations of the human MYH gene, which predisposes the patients to colorectal tumors.137 These tumors usually have a significant excess of somatic G:C to T:A mutations in the APC and K-ras genes. The former is a key gatekeeper gene that is thought to play a crucial role in the regulation of cell proliferation of the colon. This finding is consistent with the established role of MYH as a major glycosylase activity for the removal of adenines misincorporated opposite 8-oxoG from the genome. The two most common SNPs in MYH are Y165C and G382D.12 In vitro studies of the equivalent E. coli MutY mutations, Y82C and G253D, show significantly reduced MutY activity toward A:8-oxoG.138 Unlike MYH, available data on similar causal links for other human glycosylases are rather inconclusive. In the case of hOGG1, its deficiency could also be a risk factor for various types of cancer, as suggested by a number of functional and epidemiological studies.139 Elevated levels of 8-oxoG have been observed in tissues/cells from several types of human cancer. Reduced 8-oxoG activity in peripheral blood cells has been associated with increased risk of lung cancer.140,141 As for gene mutations, a number of studies have shown that the OGG1 polymorphism may be associated with various malignancies. For example, case-control studies suggest that S326C, the most common OGG1 variant, may be a risk factor for a variety of human cancers, including those from lung, esophagus, stomach and colon.142 However, a study showed that S326C was not associated with altered OGG1 activity in human cells.143 It is apparent that more studies are needed to clarify the importance and mechanism of OGG1 and its individual variants in cancer susceptibility. Among many SNPs identified from XRCC1, several (R194W, R280H, R399Q) have been extensively studied and found to be associated with an increased risk for a number of tumor types.11 XRCC1 serves as a coordinator of BER and interacts with many BER partners (see Section 5.1). Therefore, polymorphism studies have been

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focused on the potential impact of a SNP, the interactive functions of XPCC1. R194W is located in the linker region between POLβ and PARP1 interaction domains and R399Q resides in the BRCT-I interaction domain. The R399Q variant has been shown to be associated with a number of functional changes such as reduced DNA repair capacity, increased DNA damage and higher p53 mutations.144–146 This polymorphism is associated with both an increased and a reduced risk for different types of tumors.

7. PERSPECTIVES Future work in the field of BER will continue to focus on understanding of both molecular mechanisms and biological roles of BER. Combined efforts and powerful synergy from molecular and structural biology, biochemistry, epidemiology, medicine and other allied fields should generate new and comprehensive insight into those unresolved issues related to BER. In addition to a continuous search for mechanisms by which glycosylases locate specific base damage from a million-fold excess of normal bases, studies on BER repair specificity and its related implications will still be the subject of interest. Future developments would include exploring new factors required for substrate recognition; probing biologically critical DNA substrates; predicting new substrates and designing enzyme inhibitors; and obtaining more structural details of enzyme-substrate complexes as well as of enzyme-enzyme interactions. All of these would help us to understand how the BER enzymes operate in biological systems and can be used as tools for biomedical applications. One of the recent advances in BER is the illustration of an intricate network of protein-protein interactions, which is primarily built upon in vitro studies. It is imperative to verify these interactions or links in vivo as well as to elucidate their biological consequences. These studies, together with those on damage response at higher levels, should provide novel insights into networked responses to crucial DNA base damage, especially the endogenous DNA lesions.

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There seems to be a feeling of excitement in this field as we are beginning to uncover the role of BER in pathogenesis of diseases such as cancer. Much more work is obviously needed to develop effective screening approaches or larger scale evaluation of BERrelated cancer risks for human subjects. In addition, several clinical areas such as detection of BER repair capacity in diagnosis, inhibition of repair enzymes for therapeutic purposes, and medical intervention of DNA repair deficiencies would be actively explored in the future. ACKNOWLEDGMENTS This work was supported by NIH grant CA72079 and was administrated by the Lawrence Berkeley National Laboratory under the Department of Energy contract DE-AC03-76SF00098. Abbreviations 3-meA 5,6-DHT 5,6-DHU 5-FU 5-FoU 5-HmU 5-OHU 5-meC 8-oxoG AP APE AlkA BER ε E Endo III Endo IV Exo III FEN1

3-methyladenine 5,6-dihydrothymine 5,6-dihydrouracil 5-fluorouracil 5-formyluracil 5-hydroxymethyluracil 5-hydroxyuracil 5-methylcytosine 7,8-dihydro-8-oxoguanine apurinic/apyrimidinic AP endonuclease E. coli 3-methyladenine-DNA glycosylase II base excision repair etheno ethano endonuclease III endonuclease IV exonuclease III flap endonuclease 1

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Fpg Hm LIG MBD4 MPG MYH Mug MutY NEIL NTH1 Nei OGG1 PARP1 PCNA PNKP POL RFC RPA SMUG1 SNP TDG Tg UNG XRCC1 dRpase

formamidopyrimidine-DNA glycosylase hydroxymethyl ligase methyl-binding domain glycosylase 4 methylpurine-DNA glycosylase MutY homolog mismatch-specific uracil-DNA glycosylase mismatch adenine-DNA glycosylase endonuclease VIII-like endonuclease Ill-like 1 endonuclease VIII 8-oxoG-DNA glycosylase 1 poly(ADP-ribose) polymerase 1 proliferating cell nuclear antigen polynucleotide kinase 3’-phosphatase polymerase replication factor C replication protein A single-strand selective monofunctional uracil-DNA glycosylase single nucleotide polymorphism thymine-DNA glycosylase thymine glycol uracil-DNA glycosylase X-ray cross-complementation protein 1 deoxyribophosphodiesterase

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References 1. Verly WG, Paquette Y. (1972) An endonuclease for depurinated DNA in Escherichia coli B. Can J Biochem 50: 217–224. 2. Lindahl T. (1974) An N-glycosidase from Escherichia coli that releases free uracil from DNA containing deaminated cytosine residues. Proc Natl Acad Sci USA 71: 3649–3653. 3. Friedberg EC, Walker GC, Siede W, et al. (2005) DNA Repair and Mutagenesis, 2nd ed., ASM Press., Washington, D.C. 4. Memisoglu A, Samson L. (2000) Base excision repair in yeast and mammals. Mutat Res 451: 39–51. 5. Wood RD, Mitchell M, Lindahl T. (2005) Human DNA repair genes. Mutat Res 577: 275–283. 6. Hang B, Singer B. (2003) Protein-protein interactions involving DNA glycosylases. Chem Res Toxicol 16: 1181–1195. 7. Fan J, Wilson DM. (2005) Protein-protein interactions and posttranslational modifications in mammalian base excision repair. Free Radic Biol Med 38: 1121–1138. 8. Jackson AL, Loeb LA. (2001) The contribution of endogenous sources of DNA damage to the multiple mutations in cancer. Mutat Res 477: 7–21. 9. Marnett LJ, Plastaras JP. (2001) Endogenous DNA damage and mutation. Trends in Genetics 17: 214–221. 10. Glassner BJ, Posnick LM, Samson LD. (1998) The influence of DNA glycosylases on spontaneous mutation. Mutat Res 400: 33–44. 11. Fortini P, Pascucci B, Parlanti E, et al. (2003) The base excision repair: mechanisms and its relevance for cancer susceptibility. Biochimie 85: 1053–1071. 12. Al-Tassan N, Chmiel NH, Maynard, et al. (2002) Inherited variants of MYH associated with somatic G:C → T:A mutations in colorectal tumors. Nat Genet 30: 227–232. 13. Sieber OM, Lipton L, Sieber OM, et al. (2003) Multiple colorectal adenomas, classic adenomatous polyposis, and germ-line mutations in MYH. N Engl J Med 348: 791–799. 14. Taroni F, DiDonato S. (2004) Pathways to motor incoordination: the inherited ataxias. Nat Rev Neurosci 5: 641–655. 15. Rada C , Williams GT, Nilsen H, et al. (2002) Immunoglobulin isotype switching is inhibited and somatic hypermutation perturbed in UNGdeficient mice. Curr Biol 12: 1748–1755. 16. Imai K, Slupphaug G, Lee WI, et al. (2003) Human uracil-DNA glycosylase deficiency associated with profoundly impaired immunoglobulin class-switch recombination. Nat Immunol 4: 1023–1028.

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55

17. Wiederhold L, Leppard JB, Kedar P, et al. (2004) AP endonucleaseindependent DNA base excision repair in human cells. Mol Cell 15: 209–220. 18. Bebenek K, Tissier A, Frank EG, et al. (2001) 5 -Deoxyribose phosphate lyase activity of human DNA polymerase iota in vitro. Science 291: 2156–2159. 19. Kubota Y, Nash RA, Klungland A, et al. (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. 20. Pascucci B, Maga G, Hubscher U, et al. (2002) Reconstitution of the base excision repair pathway for 7,8-dihydro-8-oxoguanine with purified human proteins. Nucl Acids Res 30: 2124–2130. 21. Sattler U, Frit P, Salles B, Calsou P. (2003) Long-patch DNA repair synthesis during base excision repair in mammalian cells. EMBO Rep 4: 363–367. 22. Dogliotti E, Fortini P, Pascucci B, Parlanti E. (2001) Multiple pathways for DNA base excision repair. The mechanism of switching among multiple BER pathways. Prog Nucl Acid Res Mol Biol 68: 1–28. 23. Hazra TK, Izumi T, Kow YW, 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. 24. Dianov GL, Souza-Pinto N, Nyaga SG, et al. (2001) Base excision repair in nuclear and mitochondrial DNA. Prog Nucl Acid Res Mol Biol 68: 285–297. 25. Otterlei M, Haug T, Nagelhus TA, et al. (1998) Nuclear and mitochondrial splice forms of human uracil DNA glycosylase contain a complex nuclear localization signal and a strong classical mitochondrial localization signal, respectively. Nucl Acids Res 26: 4611–4617. 26. Stuart JA, Mayard S, Hashiguchi K, et al. (2005) Localization of mitochondrial DNA base excision repair to an inner membrane-associated particulate fraction. Nucl Acids Res 33: 3722–3732. 27. Hang B. (2004) Repair of exocyclic DNA adducts: rings of complexity. Bioessays 26: 1195–1208. 28. Dizdaroglu M. (2005) Base-excision repair of oxidative DNA damage by DNA glycosylases. Mutat Res 591: 45–59. 29. Wallace SS. (1994) DNA damages processed by base excision repair: biological consequences. Int J Radiat Biol 66: 579–589. 30. Otterlei M, Warbrick E, Nagelhus TA, et al. (1999) Post-replicative base excision repair in replication foci. EMBO J 18: 3834–3844.

FA3

November 14, 2006 12:56

56

WSPC/SPI-B414: DNA Repair, Genetic Instability & Cancer

ch02

Bo Hang

31. Nilsen H, Haushalter KA, Robins P, et al. (2001) Excision of deaminated cytosine from the vertebrate genome: role of the SMUG1 uracilDNA glycosylase. EMBO J 20: 4278–4286. 32. Petronzelli F, Riccio A, Markham GD, et al. (2000) Investigation of the substrate spectrum of the human mismatch-specific DNA Nglycosylase MED1 (MBD4): fundamental role of the catalytic domain. J Cell Physiol 185: 473–480. 33. Waters TR, Swann PF. (1998) Thymine-DNA glycosylase and G to A transition mutations at CpG sites. Mutat Res 462: 137–147. 34. Takao M, Kanno S, Kobayashi K, et al. (2002) A back-up glycosylase in Nth1 knock-out mice is a functional Nei (endonuclease VIII) homologue. J Biol Chem 277: 42205–42213. 35. Parikh SS, Mol CD, Hosfield DJ, Tainer JA. (1999) Envisioning the molecular choreography of DNAbase excision repair. Curr Opin Struct Biol 9: 37–47. 36. Pearl LH. (2000) Structure and function in the uracil-DNA glycosylase superfamily. Mutat Res 460: 165–181. 37. Hollis T, Lau A, Ellenberger T. (2001) Crystallizing thoughts about DNA base excision repair. Prog Nucleic Acid Res Mol Biol 68: 305–314. 38. Fromme JC, Banerjee A, Verdine GL. (2004) DNA glycosylase recognition and catalysis. Curr Opin Struct Biol 14: 43–49. 39. Barrett TE, Scharer OD, Savva R, et al. (1999) Crystal structure of a thwarted mismatch glycosylase DNA repair complex. EMBO J 18: 6599–6609. 40. Guliaev AB, Hang B, Singer B. (2002) Structural insights by molecular dynamics simulations into differential repair efficiency for ethanoA versus etheno-A adducts by the human alkylpurine-DNA Nglycosylase. Nucl Acids Res 30: 3778–3787. 41. Aravind L, Koonin EV. (2000) The alpha/beta fold uracil DNA glycosylases: a common origin with diverse fates. Genome Biol 1: 1–8. 42. Savva R, McAuley-Hecht K, Brown T, Pearl L. (1995) The structural basis of specific base-excision repair by uracil-DNA glycosylase. Nature 373: 487–493. 43. Barrett TE, Savva R, Panayotou G, et al. (1998) Crystal structure of a G:T/U mismatch-specific DNA glycosylase: mismatch recognition by complementary-strand interactions. Cell 92: 117–129. 44. Scharer OD, Jiricny J. (2001) Recent progress in the biology, chemistry and structural biology of DNA glycosylases. Bioessays 23: 270–281. 45. Hang B, Medina M, Fraenkel-Conrat H, Singer B. (1998) A 55-kDa protein isolated from human cells shows DNA glycosylase activity toward 3,N4 -ethenocytosine and the G/T mismatch. Proc Natl Acad Sci USA 95: 13561–13566.

FA3

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WSPC/SPI-B414: DNA Repair, Genetic Instability & Cancer

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57

46. Saparbaev M, Laval J. (1998) 3,N4 -ethenocytosine, a highly mutagenic adduct, is a primary substrate for Escherichia coli double-stranded uracil-DNAglycosylase and human mismatch-specific thymine-DNA glycosylase. Proc Natl Acad Sci USA 95: 8508–8513. 47. Sun B, Latham KA, Dodson ML, Lloyd RS. (1995) Studies on the catalytic mechanism of five DNAglycosylases. Probing for enzyme-DNA imino intermediates. J Biol Chem 270: 19501–19508. 48. Singer B, Grunberger D. (1983) Molecular Biology of Mutagens and Carcinogens, Plenum Press, New York. 49. Rydberg B, Lindahl T. (1982) Nonenzymatic methylation of DNA by the intracellular methyl group donor S-adenosyl-L-methionine is a potentially mutagenic reaction. EMBO J 1: 211–216. 50. Barbin A. (2000) Etheno-adduct-forming chemicals: from mutagenicity testing to tumor mutation spectra. Mutat Res 462: 55–69. 51. Singer B, Hang B. (1997) What structural features determine repair enzyme specificity and mechanism in chemically modified DNA? Chem Res Toxicol 10: 713–732. 52. Lau AY, Wyatt MD, Glassner BJ, Samson LD, 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. 53. Engelward BP, Weeda G, Wyatt MD, et al. (1997) Base excision repair deficient mice lacking the Aag alkyladenine DNA glycosylase. Proc Natl Acad Sci USA 94: 13087–13092. 54. Elder RH, Jansen JG, Weeks RJ, et al. (1998) Alkylpurine-DNA-Nglycosylase knockout mice show increased susceptibility to induction of mutations by methyl methanesulfonate. Mol Cell Biol 18: 5828–5837. 55. Barbin A, Wang R, O’Connor PJ, Elder RH. (2003) Increased formation and persistence of 1,N6 -ethenoadenine in DNA is not associated with higher susceptibility to carcinogenesis in alkylpurine-DNA-Nglycosylase knockout mice treated with vinyl carbamate. Cancer Res 63: 7699–7703. 56. Ham AJ, Engelward BP, Koc H, et al. (2004) New immunoaffinity-LCMS/MS methodology reveals that Aag null mice are deficient in their ability to clear 1,N6 -etheno-deoxyadenosine DNA lesions from lung and liver in vivo. DNA Repair 3: 257–265. 57. Hang B, Singer B, Margison GP, Elder RH. (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.

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58

WSPC/SPI-B414: DNA Repair, Genetic Instability & Cancer

ch02

Bo Hang

58. Posnick LM, Samson LD. (1999) Imbalanced base excision repair increases spontaneous mutation and alkylation sensitivity in Escherichia coli. J Bacteriol 181: 6763–6771. 59. Calleja F, Jansen JG, Vrieling H, et al. (1999) Modulation of the toxic and mutagenic effects induced by methyl methanesulfonate in Chinese hamster ovary cells by overexpression of the rat N-alkylpurine-DNA glycosylase. Mutat Res 425: 185–194. 60. Lindahl T. (1993) Instability and decay of the primary structure of DNA. Nature 362: 709–715. 61. Bird A. (2002) DNA methylation patterns and epigenetic memory. Genes Dev 16: 6–21. 62. Kavli B, Sundheim O, Akbari M, et al. (2002) hUNG2 is the major repair enzyme for removal of uracil from U:A matches, U:G mismatches, and U in single-stranded DNA, with hSMUG1 as a broad specificity backup. J Biol Chem 277: 39926–39936. 63. Dizdaroglu M, Karakaya A, Jaruga P, et al. (1996) Novel activities of human uracil DNA N-glycosylase for cytosine-derived products of oxidative DNA damage. Nucleic Acids Res 24: 418–422. 64. Wang Z, Mosbaugh DW. (1989) Uracil-DNA glycosylase inhibitor gene of bacteriophage PBS2 encodes a binding protein specific for uracil-DNA glycosylase. J Biol Chem 264: 1163–1171. 65. Nilsen H, Stamp G, Andersen S, et al. (2003) Gene-targeted mice lacking the Ung uracil-DNAglycosylase develop B-cell lymphomas. Oncogene 22: 5381–5386. 66. Haushalter KA, Todd Stukenberg MW, Kirschner MW, Verdine GL. (1999) Identification of a new uracil-DNA glycosylase family by expression cloning using synthetic inhibitors. Curr Biol 9: 174–185. 67. Wibley JE, Waters TR, Haushalter K, et al. (2003) Structure and specificity of the vertebrate anti-mutator uracil-DNA glycosylase SMUG1. Mol Cell 11: 1647–1659. 68. Hollstein MC, Brooks P, Linn S, Ames BN. (1984) Hydroxymethyluracil DNA glycosylase in mammalian cells. Proc Natl Acad Sci USA 81: 4003–4007. 69. Boorstein RJ, Cummings A Jr, Marenstein DR, et al. (2001) Definitive identification of mammalian 5-hydroxymethyluracil DNA Nglycosylase activity as SMUG1. J Biol Chem 276: 41991–41997. 70. Masaoka A, Matsubara M, Tanaka T, et al. (2003) Repair roles of hSMUG1 assessed by damage specificity and cellular activity. Nucl Acids Res Suppl 3: 263–264. 71. Hendrich B, Bird A. (1998) Identification and characterization of a family of mammalian methyl-CpG binding proteins. Mol Cell Biol 18: 6538–6547.

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59

72. Bellacosa A, Cicchillitti L, Schepis F, et al. (1999) MED1, a novel human methyl-CpG-binding endonuclease, interacts with DNA mismatch repair protein MLH1. Proc Natl Acad Sci USA 96: 3969–3974. 73. Hendrich B, Hardeland U, Ng HH, et al. (1999) The thymine glycosylase MBD4 can bind to the product of deamination at methylated CpG sites. Nature 401: 301–304. 74. Petronzelli F, Riccio A, Markham GD, et al. (2000) Investigation of the substrate spectrum of the human mismatch-specific DNA Nglycosylase MED1 (MBD4): fundamental role of the catalytic domain. J Cell Physiol 185: 473–480. 75. Wong E, Yang K, Kuraguchi M, et al. (2002) Mbd4 inactivation increases C to T transition mutations and promotes gastrointestinal tumor formation. Proc Natl Acad Sci USA 99: 14937–14942. 76. Millar CB, Guy J, Sansom OJ, et al. (2002) Enhanced CpG mutability and tumorigenesis in MBD4-deficient mice. Science 297: 403–405. 77. Parsons BL. (2003) MED1: a central molecule for maintenance of genome integrity and response to DNA damage. Proc Natl Acad Sci USA 100: 14601–14602. 78. Zhu B, Zheng Y, Angliker H, et al. (2000) 5-Methylcytosine DNA glycosylase activity is also present in the human MBD4 (G/T mismatch glycosylase) and in a related avian sequence. Nucl Acids Res 28: 4157– 4165. 79. Zhu B, Zheng Y, Hess D, et al. (2000) 5-methylcytosine-DNA glycosylase activity is present in a cloned G/T mismatch DNA glycosylase associated with the chicken embryo DNA demethylation complex. Proc Natl Acad Sci USA 97: 5135–5139. 80. Sibghat-Ullah, Gallinari P, Xu YZ, et al. (1996) Base analog and neighboring base effects on substrate specificity of recombinant human G:T mismatch-specific thymine DNA-glycosylase. Biochemistry 35: 12926– 12932. 81. Hardeland U, Bentele M, Jiricny J, Schar P. (2003) The versatile thymine DNA-glycosylase: a comparative characterization of the human, Drosophila and fission yeast orthologs. Nucleic Acids Res 31: 2261–2271. 82. Vairapandi M, Liebermann DA, Hoffman B, Duker NJ. (2000) Human DNA-demethylating activity: a glycosylase associated with RNA and PCNA. J Cell Biochem 79: 249–260. 83. Neddermann P, Jiricny J. (1994) Efficient removal of uracil from G.U mispairs by the mismatch-specific thymine DNA glycosylase from HeLa cells. Proc Natl Acad Sci USA 91: 1642–1646. 84. Neddermann P, Gallinari P, Lettieri T, et al. (1996) Cloning and expression of human G/T mismatch-specific thymine-DNA glycosylase. J Biol Chem 271: 12767–12774.

FA3

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60

WSPC/SPI-B414: DNA Repair, Genetic Instability & Cancer

ch02

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85. Yoon JH, Iwai S, O’Connor TR, Pfeifer GP. (2003) Human thymine DNA glycosylase (TDG) and methyl-CpG-binding protein 4 (MBD4) excise thymine glycol (Tg) from a Tg:G mispair. Nucl Acids Res 31: 5399–5404. 86. Saparbaev M, Langouet S, Privezentzev CV, et al. (2002) 1,N2 ethenoguanine, a mutagenic DNA adduct, is a primary substrate of Escherichia coli mismatch-specific uracil-DNA glycosylase and human alkylpurine-DNA-N-glycosylase. J Biol Chem 277: 26987– 26993. 87. Baba D, Maita N, Jee JG, et al. (2005) Crystal structure of thymine DNA glycosylase conjugated to SUMO-1. Nature 435: 979–982. 88. Ide H, Kotera M. (2004) Human DNA glycosylases involved in the repair of oxidatively damaged DNA. Biol Pharm Bull 27: 480–485. 89. Boiteux S, Gellon L, Guibourt N. (2002) Repair of 8-oxoguanine in Saccharomyces cerevisiae: interplay of DNA repair and replication mechanisms. Free Radic Biol Med 32: 1244–1253. 90. Boiteux S, Radicella JP. (2000) The human OGG1 gene: structure, functions, and its implication in the process of carcinogenesis. Arch Biochem Biophys 377: 1–8. 91. Michaels ML, Tchou J, Grollman AP, Miller JH. (1992) A repair system for 8-oxo-7,8-dihydrodeoxyguanine. Biochemistry 31: 10964– 10968. 92. Klungland A, Rosewell I, Hollenbach S, et al. (1999) Accumulation of premutagenic DNA lesions in mice defective in removal of oxidative base damage. Proc Natl Acad Sci USA 96: 13300–13305. 93. Minowa O, Arai T, Hirano M, et al. (2000) Mmh/Ogg1 gene inactivation results in accumulation of 8-hydroxyguanine in mice. Proc Natl Acad Sci USA 97: 4156–4161. 94. Sakumi K, Tominaga Y, Furuichi M, et al. (2003) Ogg1 knockoutassociated lung tumorigenesis and its suppression by Mth1 gene disruption. Cancer Res 63: 902–905. 95. Ocampo MT, Chaung W, Marenstein DR, et al. (2002) Targeted deletion of mNth1 reveals a novel DNA repair enzyme activity. Mol Cell Biol 22: 6111–6121. 96. Melamede RJ, Hatahet Z, Kow YW, et al. (1994) Isolation and characterization of endonuclease VIII from Escherichia coli. Biochemistry 33: 1255–1264. 97. Jiang D, Hatahet Z, Melamede RJ, et al. (1997) Characterization of Escherichia coli endonuclease VIII. J Biol Chem 272: 32230– 32239. 98. Hazra TK, Kow YW, Hatahet Z, et al. (2002) Identification and characterization of a novel human DNA glycosylase for repair of cytosinederived lesions. J Biol Chem 277: 30417–30420.

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61

99. Hazra TK, Izumi T, Boldogh I, et al. (2002) Identification and characterization of a human DNA glycosylase for repair of modified bases in oxidatively damaged DNA. Proc Natl Acad Sci USA 6: 3523–3528. 100. Bandaru V, Sunkara S, Wallace SS, Bond JP. (2002) A novel human glycosylase that removes oxidative DNA damage and is homologous to Escherichia coli endonuclease VIII. Nucl Acids Res 30: 4926–4936. 101. Zhang QM, Yonekura S, Takao M, et al. (2005) DNA glycosylase activities for thymine residues oxidized in the methyl group are functions of the hNEIL1 and hNTH1 enzymes in human cells. DNA Repair 4: 71–79. 102. Dou H, Mitra S, Hazra TK. (2003) Repair of oxidized bases in DNA bubble structures by human DNA glycosylases NEIL1 and NEIL2. J Biol Chem 278: 49679–49684. 103. Rosenquist TA, Zaika E, Fernandes AS, et al. (2003) The novel DNA glycosylase, NEIL1, protects mammalian cells from radiationmediated cell death. DNA Repair 2: 581–591. 104. Slupska MM, Luther WM, Chiang JH, et al. (1999) Functional expression of hMYH, a human homolog of the Escherichia coli MutY protein. J Bacteriol 181: 6210–6213. 105. David SS, Williams SD. (1998) Chemistry of glycosylases and endonucleases involved in base excision repair. Chem Rev 98: 1221–1261. 106. Cheadle JP, Sampson JR. (2003) Exposing the MYH about base excision repair and human inherited disease. Hum Mol Genet 12: 159–165. 107. Parker A, Gu Y, Mahoney W, et al. (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. 108. Boldogh I, Milligan D, Soog Lee M, et al. (2001) hMYH cell cycledependent expression, subcellular localization and association with replication foci: evidence suggesting replication-coupled repair of adenine:8-oxoguanine mispairs. Nucl Acids Res 29: 2802–2809. 109. Lindahl T. (1979) DNA glycosylases, endonucleases for apurinic/ apyrimidinic sites and base excision repair. Prog Nucleic Acid Res Mol Biol 22: 135–192. 110. Lindahl T. (1990) Repair of intrinsic DNA lesions. Mutat Res 238: 305–311. 111. Wilson DM 3rd, Barsky D. (2001) The major human abasic endonuclease: formation, consequences and repair of abasic lesions in DNA. Mutat Res 485: 283–307. 112. Boiteux S, Guillet M. (2004) Abasic sites in DNA: repair and biological consequences in Saccharomyces cerevisiae. DNA Repair 3: 1–12. 113. Hadi MZ, Wilson, DM 3rd , (2000) Second human protein with homology to the Escherichia coli abasic endonuclease exonuclease III. Environ Mol Mutagen 36: 312–324.

FA3

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62

WSPC/SPI-B414: DNA Repair, Genetic Instability & Cancer

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114. D. Tsuchimoto, Sakai Y, Sakumi K, Nishioka K, Sasaki M, Fujiwara T, Nakabeppu Y. (2001) Human APE2 protein is mostly localized in the nuclei and to some extent in the mitochondria, while nuclear APE2 is partly associated with proliferating cell nuclear antigen. Nucl Acids Res 29: 2349–2360. 115. Hang B, Chenna A, Fraenkel-Conrat H, Singer B. (1996) An unusual mechanism for the major human apurinic/apyrimidinic (AP) endonuclease involving 5 cleavage of DNA containing a benzenederived exocyclic adduct in the absence of an AP site. Proc Natl Acad Sci USA 93: 13737–13741. 116. Gros L, Ishchenko AA, Ide H, et al. (2004) The major human AP endonuclease (Ape1) is involved in the nucleotide incision repair pathway. Nucl Acids Res 32: 73–81. 117. Xanthoudakis S, Miao G, Wang F, et al. (1992) Redox activation of Fos-Jun DNA binding activity is mediated by a DNA repair enzyme. EMBO J 11: 3323–3335. 118. Xanthoudakis S, Curran T. (1992) Identification and characterization of Ref-1, a nuclear protein that facilitates AP-1 DNA-binding activity. EMBO J 11: 653–665. 119. Fung H, Demple B. (2005) A vital role for Ape1/Ref1 protein in repairing spontaneous DNA damage in human cells. Mol Cell 17: 463–470. 120. Xanthoudakis S, Smeyne RJ, Wallace JD, 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. 121. Ludwig DL, MacInnes MA, Takiguchi Y, et al. (1998) A murine AP-endonuclease gene-targeted deficiency with post-implantation embryonic progression and ionizing radiation sensitivity. Mutat Res 409: 17–29. 122. Huamani J, McMahan CA, Herbert DC, et al. (2004) Spontaneous mutagenesis is enhanced in Apex heterozygous mice. Mol Cell Biol 24: 8145–8153. 123. Xiao W, Samson L. (1993) In vivo evidence for endogenous DNA alkylation damage as a source of spontaneous mutation in eukaryotic cells. Proc Natl Acad Sci USA 90: 2117–2121. 124. Hadi MZ, Coleman MA, Fidelis K, et al. (2000) Functional characterization of Ape1 variants identified in the human population. Nucl Acids Res 28: 3871–3879. 125. Wilson SH, Kunkel TA. (2000) Passing the baton in base excision repair. Nat Struct Biol 7: 176–178. 126. Mol CD, Izumi T, Mitra S, Tainer JA. (2000) DNA-bound structures and mutants reveal abasic DNA binding by APE1 and DNA repair coordination. Nature 403: 451–456.

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127. Parikh SS, Mol CD, Slupphaug G, et al. (1998) Base excision repair initiation revealed by crystal structures and binding kinetics of human uracil-DNA glycosylase with DNA. EMBO J 17: 5214– 5226. 128. Miao F, Bouziane M, Dammann R, et al. (2000) 3-Methyladenine-DNA glycosylase (MPG protein) interacts with human RAD23 proteins. J Biol Chem 275: 28433–28438. 129. Shimizu Y, Iwai S, Hanaoka F, Sugasawa K. (2003) Xeroderma pigmentosum group C protein interacts physically and functionally with thymine DNA glycosylase. EMBO J 22: 164–173. 130. Dantzer F, Luna L, Bjoras M, Seeberg E. (2002) Human OGG1 undergoes serine phosphorylation and associates with the nuclear matrix and mitotic chromatin in vivo Nucl Acids Res 30: 2349–2357. 131. Hu J, Imam SZ, Hashiguchi K, et al. (2005) Phosphorylation of human oxoguanine DNA glycosylase (α-OGG1) modulates its function. Nucl Acids Res 33: 3271–3282. 132. Hardeland U, Steinacher R, Jiricny J, Schar P. (2002) Modification of the human thymine-DNA glycosylase by ubiquitin-like proteins facilitates enzymatic turnover. EMBO J 21: 1456–1464. 133. Steinacher R, Schar P. (2005) Functionality of human thymine DNA glycosylase requires SUMO-regulated changes in protein conformation. Curr Biol 15: 616–623. 134. Parsons JL, Elder RH. (2003) DNA N-glycosylase deficient mice: a tale of redundancy. Mutat Res 531: 165–175. 135. Xie Y, Yang H, Cunanan C, et al. (2004) Deficiencies in mouse Myh and Ogg1 result in tumor predisposition and G to T mutations in codon 12 of the K-ras oncogene in lung tumors. Cancer Res 64: 3096–3102. 136. Russo MT, De Luca G, Degan P, et al. (2004) Accumulation of the oxidative base lesion 8-hydroxyguanine in DNA of tumor-prone mice defective in both the Myh and Ogg1 DNA glycosylases. Cancer Res 64: 4411–4414. 137. Sampson JR, Jones S, Dolwani S, Cheadle JP. (2005) MutYH (MYH) and colorectal cancer. Biochem Soc Trans 33: 679–683. 138. Chmiel NH, Livingston AL, David SS. (2003) Insight into the functional consequences of inherited variants of the hMYH adenine glycosylase associated with colorectal cancer: complementation assays with hMYH variants and pre-steady-state kinetics of the corresponding mutated E. coli enzymes. J Mol Biol 327: 431–443. 139. Weiss JM, Goode EL, Ladiges WC, Ulrich CM. (2005) Polymorphic variation in hOGG1 and risk of cancer: a review of the functional and epidemiologic literature. Mol Carcinog 42: 127–141.

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140. Paz-Elizur T, Krupsky M, Blumenstein S, et al. (2003) DNA repair activity for oxidative damage and risk of lung cancer. J Natl Cancer Inst 95: 1312–1319. 141. Gackowski D, Speina E, Zielinska M, et al. (2003) Products of oxidative DNA damage and repair as possible biomarkers of susceptibility to lung cancer. Cancer Res 63: 4899–4902. 142. Nohmi T, Kim SR, Yamada M. (2005) Modulation of oxidative mutagenesis and carcinogenesis by polymorphic forms of human DNA repair enzymes. Mutat Res 591: 60–73. 143. Janssen K, Schlink K, Gotte W, et al. (2001) DNA repair activity of 8oxoguanine DNA glycosylase 1 (OGG1) in human lymphocytes is not dependent on genetic polymorphism Ser326/Cys326. Mutat Res 486: 207–216. 144. Lunn RM, Langlois RG, Hsieh LL, et al. (1999) XRCC1 polymorphisms: effects on aflatoxin B1-DNA adducts and glycophorin A variant frequency. Cancer Res 59: 2557–2561. 145. Duell EJ, Wiencke JK, Cheng, et al. (2000) Polymorphisms in the DNA repair genes XRCC1 and ERCC2 and biomarkers of DNA damage in human blood mononuclear cells. Carcinogenesis 21: 965–971. 146. Hsieh LL, Chien HT, Chen IH, et al. (2003) The XRCC1 399Gln polymorphism and the frequency of p53 mutations in Taiwanese oral squamous cell carcinomas. Cancer Epidemiol Biomarkers Prev 12: 439–443.

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CHAPTER 3

Nucleotide Excision Repair Lei Li∗

ABSTRACT One of the major challenges for the earliest forms of life was to protect the cellular DNA from the highly mutagenic solar ultraviolet and from exposure to environmental mutagens. The nucleotide excision repair (NER) pathway is responsible for the removal of a large variety of lesions, including those induced by ultraviolet light and bulky adduct-forming chemicals. The NER mechanism consists of two major steps. The incision step involves introduction of dual incisions flanking the DNA lesion and subsequent removal of the damage-containing oligonucleotide. The next repair synthesis step restores the damaged site by filling in the gap created by the incision step. Humans with NER deficiencies suffer from the DNA repair syndrome, xeroderma pigmentosum (XP) with manifestations of both acute sunlight sensitivity and profoundly elevated occurrence of skin cancer. XP is a classical example of the direct relationship between genomic instability and cancer development. This chapter highlights the molecular mechanism of NER and its impact on human cancers.

1. INTRODUCTION The great majority of living organisms depend on the sun for their survival; they are sustained by solar energy, directly or indirectly. However, all living organisms coming in contact with sunlight as a part of their life routine must survive the harmful consequences of solar ultraviolet (UV) exposure. ∗ Associate Professor, Department of Experimental Radiology Oncology, The University of Texas M. D. Anderson Cancer Center, 1515 Holcombe Blvd., Houston, Texas 77030, USA. Tel.: 713-792-2514; Fax: 713-794-5369; E-mail: [email protected]

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The nucleotide excision repair (NER) pathway is the predominant mechanism protecting cells from UV-induced DNA lesions. In humans, UV-induced DNA lesions, if not removed, can result in either mutational or deleterious consequences, severely compromising the integrity of the genome and profoundly elevating cancer predisposition. Genetic defects in genes participating in the NER process give rise to the autosomal recessive disorder xeroderma pigmentosum, which is among the most compelling human disease models illustrating the relationship between genomic instability and cancer development. It is perhaps also involved in cancer etiology in the general population (see more details in Chapter 12). The NER pathway is extremely versatile. DNA lesions repaired by NER are not limited to those inflicted by UV light. Certain chemicals that react with the base components of nucleotide residues in DNA produce a form of DNA lesions termed bulky adducts, which often lead to distortion of the normal DNA helix structure. The majority of these DNA-modifying chemicals are known genotoxic carcinogens. The NER pathway plays a major role in removing bulky adduct lesions. Overall, it is estimated that more than 70% of environmentally-induced DNA alterations are repaired by the NER pathway. The mammalian NER mechanism has been thoroughly investigated since the early 1990s and is among the bestunderstood DNA repair pathways. The rapid success in elucidating the mammalian NER pathway was made possible by the large body of knowledge accumulated from studies of UV damage repair in bacteria during the prior decades. 2. BASIC MECHANISM OF NER Generally, excision repair mechanisms take advantage of the inherent redundancy of the DNA double helix. When a nucleotide is altered on one strand of the double helix, the uncompromised complementary strand can be used as the template for restoring the damaged site in an error-free fashion. At the DNA level, the NER process is accomplished in five steps as defined by the structural changes on the DNA (Fig. 1). Upon recognition of a DNA lesion, in this case a

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Fig. 1

Nucleotide excision repair at the DNA level.

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UV-induced thymidine dimer, DNA damage-binding activities are required to assemble a protein complex that enacts the dual incision process. In step 1, the double helix is unwound at the site of the lesion to create a bubble structure through the action of a DNA helicase. In step 2, the first incision takes place approximately six bases downstream of the lesion. This step requires an endonucleolytic activity. In step 3, a second incision takes place approximately 22 bases upstream of the lesion and it requires another endonuclease. This leaves the damage-containing oligonucleotide (on average, 28 bases long) attached to the genome only through its base-paring hydrogen bond; it is no longer covalently linked to the chromosomal DNA. In step 4, the damage-containing oligonucleotide is removed by a helicase, resulting in a gap repair intermediate. In step 5, the gap is filled by DNAsynthesis factors using the undamaged complementary strand as the template, completing the error-free repair process. 3. PROTEIN FACTORS INVOLVED IN NUCLEOTIDE EXCISION REPAIR (FIG. 2) 3.1. Lesion Recognition There are two NER subpathways distinguished by how a lesion is recognized. The global genomic repair (GGR) pathway locates DNA lesions through damage-recognizing proteins and repairs them genome-wide indiscriminately. In contrast, the transcriptioncoupled repair (TCR) pathway senses lesions indirectly via DNA damage that has stalled transcription complex. Hence, only lesions located in actively transcribed regions of the genome are repaired by the TCR mechanism.1 Given that only a small portion of the genome is occupied by coding sequences and that an even smaller portion is being actively transcribed at any given time, the proportion of lesion removed by the TCR pathway is expected to be proportionally small. Nevertheless, TCR appears to be capable of repairing DNA damages much faster than GCR.1 This important feature of TCR allows the more critical regions of the genome to receive preferential protection. Once a lesion is recognized, the remaining steps in NER are identical for GGR and TCR.

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Fig. 2

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Molecular mechanism of nucleotide excision repair.

In the GGR subpathway the XPC protein, which forms a complex with the hHR23A or hHR23B protein, serves as the major damage recognition factor.2,3 The XPC-hHR23B complex binds to both singlestranded and double-strand DNA, but prefers lesion-containing

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DNA.4–6 Current understanding is that XPC-hHR23B recognizes the brief-strand opening around the lesions that result from lesionmediated helix distortion. Biochemical evidence indicates that the presence of XPC-hHR23B greatly enhances the in vitro processing of most monoadduct lesions.7,8 It should be noted that other proteins, such as XPE (also called DDB1), may be required for the efficient recognition of a subset of lesions such as UV-induced cyclobutane pyrimidine dimers.9 XPE is a subunit of the heterodimeric DDB (DNA damage binding) complex identifiable by the gel-shift analysis with a UV-treated DNA probe and cell extracts.10,11 The human gene encoding the DDB2 subunits carries inactivating mutations in XPE patients.12 Upon UV exposure, the DDB complex becomes tightly associated with chromatin.13 Through proteomics analyses, the DDB complex was found to be part of a supercomplex, including cullin 4A, Roc1, and COP9 signalosome, suggesting that the function of this supercomplex may be involved in ubiquitination as an E3 ligase.14 Interestingly, XPC was identified as a substrate of the DDB-containing E3 ligase. Ubiquitinated XPC was shown to have enhanced lesion binding.15 A current model holds that the DDB complex may provide the initial recognition of UV lesions through its strong association with chromatin and its high affinity toward (6–4) photoproducts.13,16,17 The loading of the DDB complex onto the damage site then recruits XPC and enables its ubiquitination. With the enhanced DNA lesion binding, the ubiquitinated XPC displaces the DDB complex and initiates the assembly of the incision complex. However, XPE is not an essential factor for NER in vitro, presumably because XPC, in its unmodified form, exhibits sufficient DNA lesion binding activity toward a variety of lesions to support the in vitro NER reaction. Virtually all DNA lesions obstruct the elongation of RNA polymerase II transcription, if they are located on the transcribed strand. Because the stalled RNA transcription serves as a strong signal of the presence of DNA lesions, the XPC-hHR23B complex is not required for transcription-stalling lesions. This is consistent with the experimental observation that deficiencies in the XPC gene do not

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affect TCR function.18 Two proteins specifically required for TCR, CSA and CSB are involved in the recruitment of downstream incision factors, including the TFIIH complex, as discussed in the next section. The observed direct interaction between CSA and TFIIH may be a critical link between TCR damage recognition and the subsequent incision step.19 3.2. Introduction of Dual Incisions The NER incision process begins with the helix unwinding at the site of the lesion. The helicase activity required for helix unwinding is provided by the TFIIH complex. The TFIIH complex consists of nine subunits, two of which (XPB and XPD) exhibit DNA helicase activities with opposite polarities. In transcription, TFIIH is an initiation factor for Pol II-dependent RNA transcription.20–22 In NER, TFIIH is necessary for both GGR and TCR.23 TFIIH is most likely recruited to the lesion in the TCR subpathway, as mentioned in the preceding section, through interaction with the CSA protein.19 In the GGR subpathway, the recruitment of TFIIH to the lesion is accomplished by its association with XPC.23 The main function of the TFIIH complex at the initial stage of the incision is to create single-stranded regions (a bubble structure) at the site of the lesion. This step is necessary because both the NER incision nucleases require a single-strand-double-strand junction to introduce the nicks. Biochemical studies show that the single-strand opening generated by TFIIH during transcription is around 10–13 bases long.24 However, the incision nicks in the NER reaction are, on average, 28 bases apart. Therefore, it is clear that in the context of NER, TFIIH can create a more extensive opening. It is also possible that the first incision, made by XPG (discussed below), provides a relief in helix tension, enabling TFIIH to carry out additional unwinding. The role of TFIIH in NER incision does not appear to be limited to its catalytic helicase activities. Using substrates with artificial openings around the site of the lesion, in vitro studies showed that TFIIH was still indispensable for the subsequent incision despite the

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pre-existing single-strand opening.25 Hence, TFIIH may have additional functions following the formation of the bubble structure. One such function could be to recruit the first incision nuclease, XPG, as supported by the observation that TFIIH interacts directly with XPG.8 The dual incisions flanking the DNA lesion are introduced sequentially. The first incision takes place approximately 6 bases 3 to the damaged nucleotide and is introduced by the XPG protein.26 XPG is a structure-specific endonuclease that cleaves at the junction of single-stranded and double-stranded DNA. XPG is a member of the Fen-1 (Flap Endo Nuclease-1) family of endonucleases,27 which cleave protruding single-stranded DNA with 3 → 5 polarity at the junctions of duplex and unpaired DNA. The participation of XPG is an absolute prerequisite for the second incision.28 In addition to its endonucleolytic activity, XPG seems to exhibit structural functions. A catalytically defective XPG mutant was found to be capable of supporting the second incision, indicating that the presence of XPG may allow the assembly of subsequent protein complex necessary for the second incision.29 The second incision step of NER involves three key factors — XPA, RPA, and ERCC1-XPF. The XPA protein carries a zinc-finger motif, and its DNA-damage-binding activity was once believed to be the lesion-recognition component of NER, prior to the demonstration that XPC is the lesion recognition protein.30 A key characteristic of XPA is its interaction with both ERCC1,31,32 a subunit of the endonuclease responsible for the 5 -incision, and replication factor A (RPA), a single-stranded DNA-binding protein.33,34 The complex formed between XPA and RPA exhibits a higher affinity towards damaged single-stranded DNA than does XPA alone. This activity of the XPA-RPA complex is now considered a damage-verification step in which the presence of a lesion is confirmed before incisions are introduced. The single-stranded DNA-binding activity of this complex may also stabilize the helix opening for the impending incisions. The interaction between XPA and ERCC1, on the other hand, may represent direct recruitment that guides the 5 -incision activity to the site of the lesion.

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The single-stranded DNA-binding activity of RPA does not appear to be its only contribution to the incision process. RPA also interacts with both of the endonucleases involved in NER. When RPA binds to the undamaged strand in the helix opening, its 3 oriented side binds ERCC1-XPF, whereas its 5 -oriented side binds XPG. These interactions have been presumed to provide strand specificity of incision placement, which requires that incisions be made in the damaged strand.35 The second incision, which takes place approximately 22 bases at 5 to the lesion, is carried out by ERCC1-XPF, a heterodimeric endonuclease.36,37 Similar to XPG, ERCC1-XPF is a structure-specific endonuclease that cleaves the single strand–double strand junction. Its 5 → 3 polarity, however, is opposite to the polarity of XPG.36,38 ERCC1 appears to be the catalytic component of the ERCC1-XPF complex, at least in part because ERCC1 contains a double helixhairpin-helix (HhH) motif that is shared by other structure-specific endonucleases.36 In vivo, ERCC1 and XPF are mutually dependent upon each other for their stability. In cells lacking XPF, the level of ERCC1 is significantly reduced, and in cells lacking ERCC1, the level of XPF is significantly reduced.36,39 It has been speculated that formation of the complex allows ERCC1 and XPF to mask their otherwise proteolytically labile domains, which may facilitate degradation when these proteins are present as monomers.36 One unique feature that distinguishes ERCC1 and XPF mutants from other NER mutants is their profound sensitivity toward DNA interstrand crosslinking agents. Since interstrand crosslinks covalently join the two strands of the double helix and result in the damage of both strands, the processing of this type of lesion most likely involves a homologous recombination with an undamaged homologous sequence. It has been shown that the ECRR1-XPF complex is involved in the removal of protruding nonhomologous flaps present in recombination intermediates.40,41 Thus, the junction-specific endonuclease activity of ERCC1-XPF is likely to be required in both NER and homologous recombinational DNA repair pathways.

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When both the 3 - and 5 -incisions have been made, the damagecontaining oligonucleotide is no longer covalently attached to chromosomal DNA. It is unclear whether the base-pairing hydrogen bonds are sufficient to keep this oligonucleotide attached to the chromosome. Perhaps the helicase activity of TFIIH can act to facilitate the detachment of the excised oligonucleotide. In vitro, the excised oligonucleotide is released in the absence of DNA synthesis,25,28 suggesting that the formation of the gap intermediate structure does not require DNA synthesis. 3.3. Repair Synthesis Upon the release of the damage-containing oligonucleotide, the intermediate gap structure needs to be resynthesized to fully restore the sequence around the damaged site. Biochemical analysis suggests that the gap synthesis step could be uncoupled from the incision step.25,28 The only factor in common between these two steps is RPA, which most likely remains bound to the undamaged strand to provide protection and to facilitate subsequent replication. Because most DNA synthesis factors are essential for cell survival and hence few mammalian genetic models are available, most of the repair synthesis components involved in NER were identified based on biochemical studies with cellular extracts. Essentially, NER repair synthesis is largely reminiscent of replicative DNA synthesis in terms of its required components. In vitro studies with chemical inhibitors and inactivating antibodies suggest that the NER gap intermediate is filled in by either DNA polymerase delta or epsilon.42–44 Replication factor C binds to the primer ends and leads to loading of the PCNA homotrimer, which forms a ring-shaped clamp capable of recruiting DNA Pol delta or Pol epsilon to initiate the gap synthesis.45–47 This paradigm of DNA synthesis is firmly established for the simple extension of primed DNA templates, which includes the NER gap intermediate. The concluding step of NER is ligation of the 5 -end of the resynthesized patch to the original sequence. Most likely, DNA ligase I is responsible for this step, as ligase I has been found to interact with PCNA.48

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4. GENETICS OF NER AND THE ESSENTIAL ROLE OF NER IN PREVENTING TUMORIGENESIS Defects in the components of the NER mechanism give rise to the autosomal recessive disease xeroderma pigmentosum (XP), first described in 1874 by Hebra and Kaposi. The term “xeroderma pigmentosum” refers to the dry, pigmented skin characteristic of this disease (Fig. 3). XP is commonly characterized by photosensitivity, pigmentary changes, premature skin aging and malignant tumor development. These manifestations are due to a cellular hypersensitivity to UV radiation resulting from defects in NER. Since NER function can be hampered by mutations in any of the NER incision factors, seven complementation groups, XPA-XPG, corresponding to the defects in the gene products of the XPA-XPG genes, have been described. Each XP patient can be assigned to a particular XP complementation group through mutation analysis or complementation studies. Different XP groups have different occurrence rates. The XPA and XPC groups are the most common, while the XPE and XPF groups are relatively rare. Generally, the frequency of XP is one case per 250,000 individuals. The Japanese

A

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Fig. 3 Manifestations of xeroderma pigmentosum. A. The patient’s skin is characteristically dry with alternating hyper- and hypopigmentation. A basal cell carcinoma is visible beside the left eye. B. The degree of sunlight exposure directly affects the severity of the skin symptoms. Skin areas normally covered by cloth are much less affected. (Photographs courtesy of Dr. Tianwen Gao, Department of Dermatology, Xijing Hospital, Xian, China, with consent from patient guardians.)

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population has the highest XP occurrence, with one case per 40,000 individuals. Individuals with XP develop multiple cutaneous neoplasms at a very young age. XP patients younger than 20 years have a 1,000-fold increase in the incidence of melanoma and other types of skin cancer. The mean patient age at diagnosis of skin cancer is eight years in patients with XP compared to 60 years in the healthy population. Metastatic malignant melanoma and squamous cell carcinoma are the two primary causes of death in patients with XP.49,50 The extremely high predisposition to cancer and the early age of the onset of cancer in XP patients profoundly demonstrate the importance of the NER mechanism in maintaining the integrity of the genome and in preventing tumorigenesis. In addition to the naturally occurring human XP mutants, a series of lab-generated mutant Chinese hamster ovarian cell lines were created through the large-scale screening of UV sensitivity in mutagenized cells in culture. This series of cell lines (i.e. excision repair cross complementing, ERCC mutants 1 through 11) covers all XP genes except for the XPA gene.51 The ERCC mutants have been extremely useful in the isolation of NER genes, particularly in the 5 incision enzyme ERCC1, since no naturally occurring human ERCC1 mutant exists. Mouse knockout models have been established for most NER genes involved in the incision step.52 The cellular phenotype and cancer-proneness of these mice are largely similar to patient manifestations. In addition to skin disorders and cancer manifestations, XP is frequently accompanied by ocular and congenital abnormalities. Ocular problems occur in nearly 80% of individuals with XP, and the initial problems noted are photophobia and conjunctivitis. Congenital problems, such as microcephaly, spasticity, ataxia and dwarfism, are seen in nearly 20% of patients with XP, and the severity of these problems is proportional to the sensitivity of XP fibroblasts to UV radiation. Mechanistic links between NER deficiencies and the developmental abnormalities common in individuals with XP remain to be established.

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Cockayne syndrome (CS), another autosomal recessive disorder, is caused by mutations in the TCR factors CSA and CSB. Similar to patients with XP, CS patients are very sensitive to UV exposure and exhibit developmental abnormalities. Interestingly, cancer incidence is not elevated in patients with CS. A commonly accepted reason is that lesions unrepaired due to a TCR defect will eventually be repaired by the unaffected GGR pathway, which is able to limit the mutagenic consequence. However, most CS patients die of neurologic degeneration at a young age due to causes unrelated to cancer that may develop at a later age. A third disease associated with defects in the NER factors is tricothiodystrophy (TTD). This rare disorder is characterized by brittle hair, dry skin, dysmorphic face, mental retardation and a noticeable degree of photosensitivity, but no increased incidence of cancer. The mutations responsible for TTD occur in a specific region of the XPB and XPD genes that encode DNA helicases involved in transcription. Other mutations in these genes cause XP or CS. At least one subgroup of TTD patients (TTD-A) was found to have a defect in the TFB5 gene, which encodes a component of the TFIIH complex.53 5. APPROACHES IN STUDYING NER A number of genetic, biochemical and cell biology approaches have been applied to the study of the NER pathway. These approaches are vital in the identification of NER genes and in the elucidation of the molecular mechanisms underlying the NER process. Detailed historical accounts of the field of DNA repair can be found in Correcting the Blueprint of Life by Errol Friedberg.54 Functional cloning or expression cloning was the most effective tool in the identification of most NER genes. Almost all the NER genes were isolated by this method. In brief, the basic principle of functional cloning of the NER genes involves introducing a library of genomic or cDNA clones into XP mutant cells and using their UV hypersensitivity to select cells that acquire resistance to UV exposure. Presumably, cells surviving the UV selection would harbor a wild-type genomic sequence or cDNA of the deficient

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gene. Genomic DNA, expression cDNA libraries and cosmid-based genomic libraries were successfully used for this purpose. The unscheduled DNA synthesis (UDS ) assay was the first approach to provide direct evidence of UV-induced repair synthesis in mammalian cells. Under unperturbed conditions, only cells undergoing replicative synthesis (S-phase cells) are able to incorporate labeled nucleotide substrate, such as tritium-labeled thymidine. When unperturbed cells are pulse-labeled with [3 H]-thymidine, fixed, and exposed to photographic emulsion, the resulting autoradiograph will show only densely labeled nuclei of S-phase cells, whereas cells outside S-phase have undetectable amounts of [3 H]thymidine incorporation. In contrast, when cells are exposed to UV light prior to [3 H]-thymidine labeling, the nuclei of non-S-phase cells are dotted with silver grains, indicating UV-induced DNA repair synthesis distinct from replicative DNA synthesis.55 Using the UDS approach, James Cleaver tested fibroblast cells isolated from XP patients and discovered that these cells were severely defective in UV-induced UDS.56 This finding not only established the cellular feature of XP patients, but also provided the first link between DNA repair and cancer predisposition in humans. DNA repair synthesis was used to develop a biochemical assay for the measurement of NER activity in vitro. In this cell-free system, whole cell extracts or nuclear extracts are used to support repair synthesis resulting from the processing of damaged plasmid substrates. When extracts are prepared from cells proficient in NER, the damaged plasmids are efficiently labeled through the incorporation of radioactive nucleotide substrate added to the reaction. When extracts are prepared from XP cells defective in the incision process, the level of labeling in the damage plasmids is greatly reduced.57 The DNA repair synthesis assay has proven extremely powerful in testing the in vitro complementation of XP mutant cells lines and the eventual reconstitution of the NER reaction.58 The incision assay has played a critical role in elucidating the mechanism of NER in mammalian cells, in particular how NER occurs at the DNA level. In the incision assay, a chemically defined lesion is placed on a selected nucleotide residue. An adjacent

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nucleotide on the same strand is also labeled, so that the incision of the damage-containing oligonucleotide can be detected by resolving the DNA substrate on denaturing PAGE and visualized through autoradiography. The incision assay allowed the 5 - and 3 -NER incision sites to be located precisely, and revealed that the average length of the excised lesion-containing oligonucleotide is around 28 bases in mammals.59,60 The incision assay was also used in reconstituting the NER reaction with recombinant and purified NER factors.8 A number of other approaches, although not specific to the NER mechanism, have played vital roles in the elucidation of the NER pathway. Somatic cell fusion was used successfully in establishing the XP complementation groups because the fusion of cells with mutations in different XP genes results in normal NER function through cross-complementation. The clonogenic survival assay against UV irradiation is routinely used to measure cellular NER activity. Finally, the host-cell reactivation assay using a plasmid with known damage induced by UV or chemicals that cause bulky adducts was developed to measure the NER capacity of the test cells, and its application has been mostly in population-based studies of cancer susceptibility (described in Chapter 12). 6. PERSPECTIVES Research since the beginning of the 1990s has made NER one of the best-defined DNA repair mechanisms, both in vitro and in vivo. However, the next challenge is to elucidate the role of NER in the context of the overall defense network against DNA damage and to decipher the regulatory mechanisms governing the NER process. How NER interfaces with the chromatin remodeling machinery may become a new frontier in NER studies. Chromatin structure restricts the interaction of DNA with most nuclear factors. The creation of accessible DNA in the context of chromatin is a key regulatory step in virtually all DNA metabolisms, including DNA repair. While the TCR subpathway of NER can benefit from prior relaxation of the chromatin structure initiated by transcription,

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the GGR subpathway of NER has to repair lesions in both relaxed and condensed chromatin. The XPE(DDB1)-CUL4DDB2 complex was recently found to be involved in histone H2A monoubiquitination in response to UV damage, and the loss of monoubiquitinated histone H2A was associated with decreased activity of the GGR subpathway.61 This observation implies that XPE function is involved in histone modification, which facilitates the initiation of NER. The overall impact of chromatin remodeling on the efficiency of the NER pathway requires further understanding of the remodeling process and how it interacts with the NER mechanism. Cells respond to the formation of DNA lesions by activating cell cycle checkpoints (described in Chapter 1) to arrest cell cycle progression. A key question regarding checkpoint activation is how DNA lesions are detected by checkpoint components. In general, different DNA repair pathways possess unique damage recognition components specific to their specialized lesions. For cell cycle checkpoints, however, it is unlikely that damage sensing is accomplished by lesion-binding proteins specific to the checkpoint. It is generally accepted that cell cycle checkpoints detect DNA replication forks stalled by encountering lesions or DNA repair intermediates common to various types of DNA damage. In mammalian cells, damage induced by UV irradiation is sensed primarily by the ATR branch of the damage and replication checkpoint. Biochemical studies suggested that single-stranded DNA is detected by the ATR-dependent checkpoint.62 Since the repair of UV lesions create a gap intermediate with an approximately 28-bp single-stranded region, the NER mechanism has the potential to provide the activating signal for the checkpoint. Such a notion will likely evolve into a testable model in the checkpoint damage detection mechanism. In summary, the NER pathway, which is an outstanding example of the relationship between genomic instability and cancer development, still holds many unanswered questions. Continual study of this important repair pathway will shed light on the overall cellular defense mechanisms for the maintenance of genomic integrity.

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ACKNOWLEDGMENTS The author wishes to thank members of the Li laboratory at The University of Texas M. D. Anderson Cancer Center for their suggestions and helpful discussions. The author receives grant supports (CA97175, CA91029, and CA76172) from the NIH/National Cancer Institute (U.S.).

References 1. Mellon I, Spivak G, Hanawalt PC. (1987) Selective removal of transcription-blocking DNA damage from the transcribed strand of the mammalian DHFR gene. Cell 51: 241–249. 2. Sugasawa K, Okamoto T, Shimizu Y, et al. (2001) A multistep damage recognition mechanism for global genomic nucleotide excision repair. Genes Dev 15: 507–521. 3. Li L, Lu X, Peterson C, Legerski R. (1997) XPC interacts with both HHR23B and HHR23A in vivo. Mutat Res 383: 197–203. 4. Masutani C, Sugasawa K, Yanagisawa J, et al. (1994) Purification and cloning of a nucleotide excision repair complex involving the xeroderma pigmentosum group C protein and a human homologue of yeast RAD23. EMBO J 13: 1831–1843. 5. Shivji MK, Eker AP, Wood RD. (1994) DNA repair defect in xeroderma pigmentosum group C and complementing factor from HeLa cells. J Biol Chem 269: 22749–22757. 6. Reardon JT, Mu D, Sancar A. (1996) Overproduction, purification, and characterization of the XPC subunit of the human DNA repair excision nuclease. J Biol Chem 271: 19451–19456. 7. Aboussekhra A, Biggerstaff M, Shivji MK, et al. (1995) Mammalian DNA nucleotide excision repair reconstituted with purified protein components. Cell 80: 859–868. 8. Mu D, Park CH, Matsunaga T, et al. (1995) Reconstitution of human DNA repair excision nuclease in a highly defined system. J Biol Chem 270: 2415–2418. 9. Hwang BJ, Chu G. (1993) Purification and characterization of a human protein that binds to damaged DNA. Biochemistry 32: 1657–1666. 10. Chu G, Chang E. (1988) Xeroderma pigmentosum group E cells lack a nuclear factor that binds to damaged DNA. Science 242: 564–567. 11. Keeney S, Wein H, Linn S. (1992) Biochemical heterogeneity in xeroderma pigmentosum complementation group E. Mutat Res 273: 49–56. 12. Nichols AF, Ong P, Linn S. (1996) Mutations specific to the xeroderma pigmentosum group E Ddb- phenotype. J Biol Chem 271: 24317–24320.

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13. Otrin VR, McLenigan M, Takao M, et al. (1997) Translocation of a UVdamaged DNA binding protein into a tight association with chromatin after treatment of mammalian cells with UV light. J Cell Sci 110 ( Pt 10): 1159–1168. 14. Groisman R, Polanowska J, Kuraoka I, et al. (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. 15. Sugasawa K, Okuda Y, Saijo M, et al. (2005) UV-induced ubiquitylation of XPC protein mediated by UV-DDB-ubiquitin ligase complex. Cell 121: 387–400. 16. Fujiwara Y, Masutani C, Hanaoka F, Iwai S. (1997) Detection, purification and characterization of a protein that binds the (6-4) photoproductcontaining DNA in HeLa cells. Nucl Acids Symp Ser 277–278. 17. Fujiwara Y, Masutani C, Mizukoshi T, et al. (1999) Characterization of DNA recognition by the human UV-damaged DNA-binding protein. J Biol Chem 274: 20027–20033. 18. van Hoffen A, Venema J, Meschini R, et al. (1995) Transcription-coupled repair removes both cyclobutane pyrimidine dimers and 6-4 photoproducts with equal efficiency and in a sequential way from transcribed DNA in xeroderma pigmentosum group C fibroblasts. EMBO J 14: 360–367. 19. Henning KA, Li L, Iyer N, et al. (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. 20. Conaway RC, Conaway JW. (1989) An RNApolymerase II transcription factor has an associated DNA-dependent ATPase (dATPase) activity strongly stimulated by the TATA region of promoters. PNAS 86: 7356– 7360. 21. Feaver WJ, Svejstrup JQ, Henry NL, Kornberg RD. (1994) Relationship of CDK-activating kinase and RNA polymerase II CTD kinase TFIIH/TFIIK. Cell 79: 1103–1109. 22. Gerard M, Fischer L, Moncollin V, et al. (1991) Purification and interaction properties of the human RNA polymerase B(II) general transcription factor BTF2. J Biol Chem 266: 20940–20945. 23. Drapkin R, Reardon JT, Ansari A, et al. (1994) Dual role of TFIIH in DNA excision repair and in transcription by RNA polymerase II. Nature 368: 769–772. 24. Holstege F, van der Vliet P, Timmers H. (1996) Opening of an RNA polymerase II promoter occurs in two distinct steps and requires the basal transcription factors IIE and IIH. EMBO J 15: 1666–1677.

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25. Mu D, Sancar A. (1997) Model for XPC-independent transcriptioncoupled repair of pyrimidine dimers in humans. J Biol Chem 272: 7570– 7573. 26. O’Donovan A, Davies AA, Moggs JG, et al. (1994) XPG endonuclease makes the 3 incision in human DNA nucleotide excision repair. Nature 371: 432–435. 27. Shen B, Singh P, Liu R, et al. (2005) Multiple but dissectible functions of FEN-1 nucleases in nucleic acid processing, genome stability and diseases. Bioessays 27: 717–729. 28. Mu D, Hsu DS, Sancar A. (1996) Reaction mechanism of human DNA repair excision nuclease. J Biol Chem 271: 8285–8294. 29. Wakasugi M, Reardon JT, Sancar A. (1997) The non-catalytic function of XPG protein during dual incision in human nucleotide excision repair. J Biol Chem 272: 16030–16034. 30. Jones CJ, Wood RD. (1993) Preferential binding of the xeroderma pigmentosum group A complementing protein to damaged DNA. Biochemistry 32: 12096–12104. 31. Li L, Elledge S, Peterson C, et al. (1994) Specific association between the human DNA repair proteins XPA and ERCC1. PNAS 91: 5012–5016. 32. Park C, Sancar A. (1994) Formation of a ternary complex by human XPA, ERCC1, and ERCC4(XPF) excision repair proteins. PNAS 91: 5017–5021. 33. He Z, Henricksen LA, Wold MS, Ingles CJ. (1995) RPA involvement in the damage-recognition and incision steps of nucleotide excision repair. Nature 374: 566–569. 34. Li L, Lu X, Peterson C, Legerski R. (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. 35. de Laat WL, Appeldoorn E, Sugasawa K, et al. (1998) DNA-binding polarity of human replication protein A positions nucleases in nucleotide excision repair. Genes Dev 12: 2598–2609. 36. Sijbers AM, de Laat WL, Ariza RR, et al. (1996) Xeroderma pigmentosum group F caused by a defect in a structure-specific DNA repair endonuclease. Cell 86: 811–822. 37. Bessho T, Sancar A, Thompson LH, Thelen MP. (1997) Reconstitution of human excision nuclease with recombinant XPF-ERCC1 complex. J Biol Chem 272: 3833–3837. 38. Matsunaga T, Mu D, Park C-H, et al. (1995) Human DNA repair excision nuclease. J Biol Chem 270: 20862–20869. 39. Yagi T, Wood RD, Takebe H. (1997) A low content of ERCC1 and a 120 kDa protein is a frequent feature of group F xeroderma pigmentosum fibroblast cells. Mutagenesis 12: 41–44.

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40. Sargent RG, Brenneman MA, Wilson JH. (1997) Repair of site-specific double-strand breaks in a mammalian chromosome by homologous and illegitimate recombination. Mol Cell Biol 17: 267–277. 41. Adair GM, Rolig RL, Moore-Faver D, et al. (2000) Role of ERCC1 in removal of long non-homologous tails during targeted homologous recombination. EMBO J 19: 5552–5561. 42. Coverley D, Kenny MK, Lane DP, Wood RD. (1992) Arole for the human single-stranded DNA binding protein HSSB/RPA in an early stage of nucleotide excision repair. Nucl Acids Res 20: 3873–3880. 43. Hunting DJ, Gowans BJ, Dresler SL. (1991) DNA polymerase delta mediates excision repair in growing cells damaged with ultraviolet radiation. Biochem Cell Biol 69: 303–308. 44. Dresler SL, Gowans BJ, Robinson-Hill RM, Hunting DJ. (1988) Involvement of DNA polymerase delta in DNA repair synthesis in human fibroblasts at late times after ultraviolet irradiation. Biochemistry 27: 6379–6383. 45. Shivji MK, Podust VN, Hubscher U, Wood RD. (1995) Nucleotide excision repair DNA synthesis by DNA polymerase epsilon in the presence of PCNA, RFC, and RPA. Biochemistry 34: 5011–5017. 46. Shivji KK, Kenny MK, Wood RD. (1992) Proliferating cell nuclear antigen is required for DNA excision repair. Cell 69: 367–374. 47. Wood RD, Shivji MK. (1997) Which DNA polymerases are used for DNA-repair in eukaryotes? Carcinogenesis 18: 605–610. 48. Montecucco A, Rossi R, Levin DS, Gary R, et al. (1998) DNA ligase I is recruited to sites of DNA replication by an interaction with proliferating cell nuclear antigen: identification of a common targeting mechanism for the assembly of replication factories. EMBO J 17: 3786–3795. 49. Kraemer KH, Lee MM, Scotto J. (1987) Xeroderma pigmentosum. Cutaneous, ocular, and neurologic abnormalities in 830 published cases. Arch Dermatol 123: 241–250. 50. Kraemer KH, Lee MM, Scotto J. (1982) Early onset of skin and oral cavity neoplasms in xeroderma pigmentosum. Lancet 1: 56–57. 51. Busch DB, Cleaver JE, Glaser DA. (1980) Large-scale isolation of UVsensitive clones of CHO cells. Somatic Cell Genet 6: 407–418. 52. Friedberg EC, Meira LB. (2006) Database of mouse strains carrying targeted mutations in genes affecting biological responses to DNA damage Version 7. DNA Repair (Amst) 5: 189–209. 53. Ranish JA, Hahn S, Lu Y, Yi EC, et al. (2004) Identification of TFB5, a new component of general transcription and DNA repair factor IIH. Nat Genet 36: 707–713. 54. Friedberg EC. (1997) Correcting the Blueprint of Life: An Historical Account of the Discovery of DNA Repair Mechanisms. Cold Spring Harbor Laboratory Press ISBN: 0879695072.

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55. Djordjevic B, Tolmach LJ. (1967) Responses of synchronous populations of HeLa cells to ultraviolet irradiation at selected stages of the generation cycle. Radiat Res 32: 327–346. 56. Cleaver JE. (1968) Defective repair replication of DNA in xeroderma pigmentosum. Nature 218: 652–656. 57. Wood RD, Robins P, Lindahl T. (1988) Complementation of the xeroderma pigmentosum DNA repair defect in cell-free extracts. Cell 53: 97–106. 58. Aboussekhra A, Biggerstaff M, Shivji MKK, Vilpo JA, et al. (1995) Mammalian DNAnucleotide excision repair reconstituted with purified protein components. Cell 80: 859–868. 59. Huang JC, Svoboda DL, Reardon JT, Sancar A. (1992) Human nucleotide excision nuclease removes thymine dimers from DNA by incising the 22nd phosphodiester bond 5 and the 6th phosphodiester bond 3 to the photodimer. Proc Natl Acad Sci USA 89: 3664–3668. 60. Svoboda DL, Taylor JS, Hearst JE, Sancar A. (1993) DNA repair by eukaryotic nucleotide excision nuclease. Removal of thymine dimer and psoralen monoadduct by HeLa cell-free extract and of thymine dimer by Xenopus laevis oocytes. J Biol Chem 268: 1931–1936. 61. Kapetanaki MG, Guerrero-Santoro J, Bisi DC, et al. (2006) The DDB1CUL4ADDB2 ubiquitin ligase is deficient in xeroderma pigmentosum group E and targets histone H2A at UV-damaged DNA sites. PNAS 103: 2588–2593. 62. Zou L, Elledge SJ. (2003) Sensing DNA damage through ATRIP recognition of RPA-ssDNA complexes. Science 300: 1542–1548.

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CHAPTER 4

DNA Mismatch Repair: Biological Functions and Molecular Mechanisms Guo-Min Li∗

ABSTRACT DNA mismatch repair (MMR) is an important genome caretaker system. It ensures genomic stability by correcting mismatches generated during DNA replication and recombination, suppressing homeologous recombination, and triggering apoptosis of cells with severe DNA damage. Protein components required for these reactions are highly conserved through evolution, and MutS-like and MutL-like proteins in mammalian cells are key players responsible for the initiation steps of both the strand-specific mismatch correction and the MMR-dependent apoptotic signaling. The inactivation of the genes encoding these activities leads to genome-wide instability, particularly in simple repetitive sequences, and the predisposition to certain types of cancer, including hereditary non-polyposis colorectal cancer.

1. INTRODUCTION Cancer is a disease of DNA as cancer cells are associated with numerous mutations in DNA.1 It is known that mutations can be induced by either exogenous or endogenous agents. Exogenously, environmental chemical (e.g. benzo[a]pyrene) and physical (e.g. ultraviolet light) agents can covalently modify DNA to change genetic information. The genetic information can also be altered endogenously ∗ Graduate

Center for Toxicology and Markey Cancer Center, University of Kentucky College of Medicine, Lexington, KY 40536, USA. Tel.: (859) 257-7053; Fax: (859) 323-1059; E-mail: [email protected] 87

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through errors in normal DNA metabolism, including DNA replication, recombination and repair. For example, nucleotide misincorporation can occur during DNA synthesis conducted by both replicative and translesion by-pass DNApolymerases.2 To safeguard the integrity of the genome, cells possess multiple mutation avoidance systems, one of which is the DNA mismatch repair (MMR) system. The importance of MMR in maintaining genomic stability was demonstrated in bacteria more than 30 years ago with the observation that defects in this pathway led to elevated levels of spontaneous mutability.3 Inactivation of MMR in human cells also results in genome-wide instability, including microsatellite sequences, and is associated with the development of both hereditary and sporadic cancer.4,5 The genome-maintenance function of MMR has been attributed to its ability to correct DNA mismatches generated during DNA replication and to block DNA recombinations occurring among divergent DNA sequences.5,6 However, recent studies indicate that the MMR system also contributes to genomic stability by mediating cell cycle checkpoints and/or programmed cell death in response to certain DNA damaging agents.7,8 The latter function of the MMR system eliminates severely damaged cells from growing, thereby preventing tumorigenesis. The rapid increase in knowledge about MMR in yeast and humans in the last decade has led to our current understanding of the molecular mechanisms of MMR in eukaryotic cells. The focus of this chapter will be on human MMR and its role in cancer avoidance. However, as an introduction to MMR in human cells, the Escherichia coli system will be discussed briefly. Readers interested in the details of the genetics and biochemistry of MMR in humans and other organisms are referred to a number of excellent reviews.2,4,5,9–12 2. THE MISMATCH REPAIR SYSTEM CORRECTS BIOSYNTHETIC ERRORS 2.1. DNA Mismatch Repair in Escherichia coli In 1989, the MMR system in Escherichia coli was reconstituted using all purified proteins,13 which include MutS, MutL, MutH, DNA

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helicase II (MutU/UvrD), four exonucleases (ExoI, ExoVII, ExoX, and RecJ), single-stranded DNA binding protein (SSB), DNA polymerase III holoenzyme and DNA ligase.2 Among these proteins, MutS, MutL and MutH are the three components that initiate the repair process and are specific to MMR. MutS is the mismatch recognition protein. It recognizes both base-base mismatches and small nucleotide insertion/deletion (ID) mispairs.5 MutS possesses an intrinsic ATPase activity, which is essential for the MMR function. Thanks to the elegant work done by the laboratories of Yang, Hsieh, and Sixma, several structures of MutS protein-DNA complexes have been determined by X-ray crystallography.14,15 These structures have revealed that MutS binds to a mismatch as a homodimer. Interestingly, the mismatch-binding site comprises the non-homologous domains from each protein monomer, indicating an asymmetric use of residues from each subunit. Hence, the MutS homodimer appears to bind to a mismatch as a virtual heterodimer, a characteristic adopted by eukaryotic MutS homologs (see below). Although MutL is absolutely required for MMR, its enzymatic activity in MMR has not been defined. MutL interacts physically with MutS to enhance mismatch recognition, and it is required for the recruitment and activation of MutH, an endonuclease that incises the daughter DNA strand to target MMR on the newly synthesized strand.5 Since MutL is also responsible for loading helicase II at the MMR initiation site, the protein is considered as a molecular matchmaker to facilitate the assembly of functional MMR protein complexes.5,16 Like MutS, MutL functions as a homodimer and possesses an ATPase activity.17 Mutations in the ATP binding domain lead to a dominant negative mutator phenotype both in vivo and in vitro. MutL mutant proteins that are defective in ATP hydrolysis, but proficient in ATP binding, can activate MutH but cannot stimulate MutH in response to a mismatch or MutS, suggesting that ATP hydrolysis by MutL is essential for mediating the activation of MutH by MutS.18 In E. coli, DNA is methylated at the N6 position of adenine residues in GATC sequences, but the newly replicated daughter

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strand is transiently unmethylated in these sequences. It is these hemimethylated GATC sequences that allow repair to be targeted at the newly synthesized daughter strand, where the incorrect base is located. The enzyme recognizing the hemimethylated GATC sequence is MutH, a protein that functions as a monomer and belonging to the family of type-II restriction endonucleases.19 Upon its recruitment and activation by MutS and MutL in the presence of ATP, MutH specifically incises the unmethylated daughter strand within the GATC sequence,5,18 providing the initiation site for mismatch-provoked excision. It is well accepted that repair initiation begins with the binding of a MutS homodimer to the mismatch. A methyl group that is either upstream (5 ) or downstream (3 ) of the incorrect base is located by an unknown mechanism involving a concerted action of the initiation factors MutS, MutL and MutH in the presence of ATP. Three models have been proposed to address how mismatch binding by MutS leads to the downstream DNA cleavage at the hemimethylated GATC site (reviewed in Refs. 2 and 20). The strand break created by MutH at a GATC site of the unmethylated strand serves as a starting point for the excision of the mispaired base. In the presence of MutL, helicase II loads at the nick and unwinds the duplex from the nick towards the mismatch, revealing a single-strand DNA region of the un-nicked strand to which SSB binds to prevent its attack by nucleases. Depending on the position of the strand break relative to the mismatch, ExoI or ExoX (3 → 5 exonuclease), or ExoVII or RecJ (5 → 3 exonucleases) excises the nicked strand from the nicked site (the GATC site) up to and slightly past the mismatch. The resulting single-stranded gap undergoes repair resynthesis and ligation by the DNA polymerase III holoenzyme, SSB and DNA ligase (see Fig. 1). The E. coli methyl-directed, MutHLS-dependent MMR pathway possesses the following features: (1) the repair is only targeted at the newly synthesized strand, where the incorrect base is positioned; (2) the repair is bi-directional, i.e. excision can proceed from the nick in either a 5 → 3 or 3 → 5 direction; and (3) the system has a fairly broad substrate specificity, being able to process both base-base

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Fig. 1 Schematic diagram of MMR in E. coli. MutS recognizes and binds to the mismatch, and the formation of the MutS-DNA complex leads to the recruitment of MutL and MutH proteins to the heteroduplex. In the presence of ATP, MutS, and MutL, MutH specifically nicks the non-methylated (newly synthesized) strand at a hemimethylated GATC site. UvrD and an exonuclease load to the strand break to remove mispaired base. The resulting ssDNA gap is coated by SSB and filled by DNA polymerase III holoenzyme. DNA ligase seals the nick.

mismatches and a variety of small insertion/deletion mispairs. All of these properties require functional MutS, MutL and MutH proteins. Because the mechanism of the bacterial MMR pathway appears to be well conserved through evolution, it serves as a model for the MMR process in eukaryotic cells. 2.2. Mismatch Repair in Human Cells Human cells possess an MMR pathway that is similar to the E. coli methyl-directed, MutHLS-dependent system. As shown in Table 1, these two pathways have many similarities. Like the E. coli pathway, the human system efficiently repairs both base-base mismatches and

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Table 1. E. coli (MutS)2

(MutL)2

MutH UvrD ExoI, ExoVII, ExoX, RecJ Pol III holoenzyme

SSB

MMR Components and their Functions Human

hMutSα (MSH2-MSH6)a hMutSβ (MSH2-MSH3) hMutLα (MLH1-PMS2)a hMutLβ (MLH1-PMS1) hMutLγ (MLH1-MLH3)

b

DNA mismatch/damage recognition

?b ?b ExoI Polδ

DNA re-synthesis

PCNA

Participating in repair initiation and facilitating DNA re-synthesis Protecting template DNA from degradation; stimulating excision in the presence of the mismatch, but facilitating excision termination upon mismatch removal; promoting DNA resynthesis when phosphorylated Facilitating mismatch-provoked excision Invovled in 3 nick-directed repair; facilitating PCNA loading Nick ligation

RPA

RFC

a

Function

Molecular matchmaker; mediating the termination of mismatch-provoked excision upon mismatch removal Strand discrimination Unwinding DNA helix Removing mispaired base

HMGB1

DNA Ligase

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DNA ligase I

Major components in cells. Not yet identified.

small ID mispairs. The strand discrimination signal in human cells appears not to involve methylation, but it does involve recognizing and targeting a strand containing a pre-existing strand break, suggesting that like the E. coli reaction, the strand-specific repair in human cells is also nick-directed. As in E. coli, the human MMR reaction is capable of bi-directional processing of a mispaired base.2,5 Because of similarities in repair properties (substrate specificity, component activities, and repair mechanism) between the human

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and the E. coli MMR systems, a search has been undertaken to identify human MMR components homologous to E. coli MMR proteins. Indeed, human homologs of the E. coli MutS and MutL proteins, two of the three known MMR-specific components in E. coli, have been identified in human cells. In addition, most of the proteins involved in the steps of excision and resynthesis have recently been identified, which include exonuclease activity EXO1, single-strand DNA binding protein RPA , proliferating cellular nuclear antigen (PCNA), DNA polymerase δ, and DNA ligase I.2,21 The discovery of the similarities between the E. coli and human MMR has greatly advanced our current understanding of the human MMR pathway. It is noteworthy that the E. coli MutS and MutL proteins are homodimers, but their human counterparts function as heterodimeric complexes. Among the three human MutS homologs (hMSH2, hMSH3, and hMSH6) involved in strand-specific MMR, hMSH2 interacts with hMSH6 or hMSH3 to form hMutSα or hMutSβ, respectively. Like the MutS protein in E. coli, both hMutSα and hMutSβ possess an intrinsic ATPase activity, which plays a critical role in MMR initiation. In addition, these heterodimers are responsible for mismatch recognition, with hMutSα preferentially recognizing base-base mismatches and ID mispairs of one or two nucleotides, and hMutSβ preferentially recognizing ID mispairs (Fig. 2). Four human MutL homologs, hMLH1, hMLH3, hPMS1 and hPMS2, have been identified. hMLH1 interacts with hPMS2, hPMS1 or hMLH3 to form three heterodimeric complexes, designated as hMutLα, hMutLβ or hMutLγ, respectively. While the function of hMutLβ is not clear at this time, hMutLα is required for MMR, and hMutLγ appears to be involved in meiosis.2 Like the MutL protein in E. coli, hMutLα possesses an ATPase activity, and defects in this activity result in the loss of MMR function. Recently, in a reconstituted MMR system, hMutLα has been shown to be responsible for terminating mismatch-provoked excision upon mismatch removal.21 The identification of PCNA and EXO1 as required MMR proteins was made by virtue of their ability to interact with MSH2 and MLH1 in both yeast and human cells (reviewed in Ref. 2). PCNA

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Fig. 2 Schematic diagram of MMR in human cells. The human MMR system can bi-directionally process all base-base mismatches and 1–16 nt ID mispairs, and the strand-specific repair is directed at a strand break. Except for mismatch recognition, where hMutSα and hMutSβ are required to distinguish specific substrates as indicated, activities required for the remaining steps of the reaction are believed to be essentially the same for the processing of base-base mismatches and ID mispairs.

is required for the initiation step of MMR, possibly by transferring MutSα/MutSβ to the mismatch site. In addition, PCNA is also involved in the step of DNA resynthesis. Both genetic and biochemical studies have implicated EXO1 in MMR. However, whether or not EXO1 is required for both 5 and 3 nick-directed excision is not clear. Studies by Modrich and colleagues indicate that the nuclease is responsible for removing mismatched bases from both the 5 → 3 and the 3 → 5 orientations,22 but a recent work from our laboratory suggests that EXO1 can only mediate 5 nick-directed MMR.21 Our observations appear to be consistent with the fact that: 1) exo1 null mutants in mice and yeast only display a partial mutator phenotype23 ; and 2) only a 5 → 3 exonuclease is associated with the purified protein.24 Based on what we know of the E. coli MMR, it is reasonable to believe that the human MMR reaction may require more than one exonuclease. Other protein components involved in MMR are single strand DNA binding protein RPA, replication factor C (RFC), high mobility group box 1 protein (HMGB1), and DNA polymerase δ (pol δ).

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The roles for RPA and pol δ have been well characterized, whereas the functions of RFC and HMGB1 are not so clear. RPA was found to stimulate mismatch-provoked excision, protect the ssDNA region generated during excision, and facilitate DNA resynthesis. Recently, we have shown that RPA binds to nicked heteroduplex DNA earlier than MutSα and MutLα, and that it becomes phosphorylated at a point where extensive excision has occurred and pol δ is recruited to the DNA substrate (Guo, Wong, and Li, unpublished results), indicating that RPA is involved in all stages of MMR and that RPA phosphorylation may modulate its functions during MMR. Indeed, we show that: (1) phosphorylation greatly reduces the DNA binding ability of RPA; and (2) unphosphorylated RPA is a much better stimulator for the excision reaction than phosphorylated RPA, but phosphrylated RPA is a much better facilitator for the resynthesis reaction than unphosphorylated RPA. These results are consistent with the fact that protection of the ssDNA region and the displacement of MutSα-MutLα complexes along the excision tract require a form of RPA with a strong DNA binding affinity, and the resynthesis reaction requires the bound RPA to be easily removable, thereby making the template DNA available for DNA synthesis by pol δ (see Fig. 3, Guo, Wong and Li, unpublished results). 3. DEFECTS IN MMR DEFICIENCY LEAD TO HUMAN CANCER Given the important function of MMR in correcting biosynthetic errors, the MMR system has long been thought to be a critical cellular mechanism preventing tumorigenesis and certain other human diseases. Indeed, defects in MMR have been known to be the genetic basis of hereditary non-polyposis colorectal cancer (HNPCC) and a subset of sporadic colon cancers. HNPCC, or Lynch syndrome, is a heritable autosomal dominant disease, which is defined by the presence of colorectal cancer in at least three family members in two successive generations, with one affected member having been diagnosed before the age of 50. HNPCC, which accounts for 4–13%

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Initiation 5’

Excision

Mispaired base

5’ Unphosphorylated RPA Phosphorylated RPA

Resynthesis

Excision machinery

5’ Polymerization machinery

5’

Fig. 3 Regulation of RPA functions by phosphorylation. Unphosphorylated heterotrimeric (70 kD, 34 kD, and 13 kD) RPA initially binds to nicked heteroduplex DNA to facilitate the assembly of the MMR initiation complex and to promote mismatch-provoked excision. The 34-kD subunit begins to be phosphorylated after extensive excision. RPA phosphorylation greatly reduces its DNA binding affinity and can be readily displaced by DNA polymerase δ making the DNA template available for resynthesis.

of all colorectal cancers, is one of the most common forms of neoplasia in Western populations. In addition to colon cancer, individuals from HNPCC families are at an increased risk of developing cancers of the endometrium, ovary, stomach, urinary tract, brain and other epithelial organs. Although HNPCC was suspected to be a heritable disease almost a century ago, the molecular pathogenesis of this disease was not established until 1993. The dramatical discovery is described below.

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3.1. Microsatellite Instability in HNPCC and Sporadic Colorectal Cancer The genetic linkage analysis of HNPCC kindreds revealed that HNPCC is associated with a disease locus at chromosome 2p15– 16.25 It was therefore thought that the genetic basis for HNPCC could be the loss of a tumor suppressor gene at this locus. A team led by Vogelstein and de la Chapelle employed microsatellite markers to determine if allelic losses occurred in the p15–16 region of the chromosome 2. This strategy was undertaken because tumor suppressor genes, if mutated, often undergo loss of heterozygosity, leading to allelic losses in the area of the disease locus. Surprisingly, no allelic losses were found in HNPCC kindreds; instead, insertion or deletion mutations at repetitive sequences were found in 11 of the 14 tumors examined.26 These unexpected mutations were evident in each di- and trinucleotide repeat (microsatellite) tested and were referred to as RER+ (replication error positive), which is now called microsatellite instability (MSI). In addition, these investigators also found a subset of sporadic colon cancers with a similar phenotype, but occurring at a much lower incidence (6 out of 46).26 At the same time, two other groups led by Perucho27 and Thibodeau28 independently reported microsatellite alterations in 12–15% of sporadic colon cancers. Taken together, these findings suggest that MSI in HNPCC and a subset of sporadic colorectal cancers is a genomewide phenomenon and may be caused by a common mechanism. Although the genetic basis of HNPCC remained unidentified at that time, these studies provided an important clue as to the mechanism of its action.

3.2. Loss of MMR Function Is the Genetic Basis of HNPCC 3.2.1. The linking of MMR defects with MSI tumors The identification of MSI in colorectal cancers received a great deal of attention from cancer investigators as well as from geneticists and biochemists working in the field of DNA MMR, because the mutational fingerprint of HNPCC tumors is similar to that found

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in MMR-deficient cells. At that time, the following points had been established: 1) loss of MMR function leads to genome-wide basebase substitutions as well as frameshift mutations; 2) MMR proteins recognize and process ID mispairs; and 3) repetitive dinucleotide sequences undergo frequent contractions or expansions in MMRdeficient bacterial cells. Therefore, the hypothesis was made that the genetic defects in HNPCC involve a loss of MMR function. Several groups tested this hypothesis using different approaches. First, Petes and colleagues examined the stability of poly(GT) tracts in yeast strains with either a single or double knockout of MSH2, MLH1, or PMS1. All mutants (both single and double) exhibited elevated levels (100- to 700-fold) of tract instability involving the insertion or deletion of 2–4 repeated units.29 This study strongly supported an association of MMR defects with the HNPCC syndrome. Second, Kolodner and co-workers, and Vogelstein and coworkers independently searched for human MMR homolog genes and determined their association with HNPCC kindreds. Both groups reported the cloning of the hMSH2 gene using PCR products of degenerate primers derived from two highly conserved regions of the known bacterial MutS and yeast MSH proteins, located the gene on the p arm of chromosome 2, and identified germ-line mutations of hMSH2 in HNPCC families.30,31 Third, Modrich and co-workers,32 and Kunkel and co-workers33 took a biochemical approach and examined the MMR-proficiency of tumor cells derived from HNPCC and sporadic colorectal cancers with MSI. Both laboratories demonstrated that the cell extracts of these tumor cells are completely defective in the repair of base-base and ID mispairs. These in vitro biochemical studies provided definitive evidence that supports the hypothesis that MMR defects are the genetic basis of HNPCC. 3.2.2. Germline mutations of MMR genes in HNPCC Immediately following the mapping of the first HNPCC-linked locus to chromosome 2p, Lindblom et al.34 identified a second locus that is linked to HNPCC predisposition at p21–23 of chromosome 3. In a remarkably short period of time after the cloning of hMSH2,

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three human MutL homolog genes (hMLH1, hPMS1, and hPMS2) were cloned. Liskay and co-workers35 identified and mapped the hMLH1 gene to the second HNPCC locus, and missense germline mutations in hMLH1 were found in a family with a history of HNPCC. At the same time, Vogelstein and co-workers searched a human cDNA database and also identified the hMLH1 gene.36 In addition, they reported two additional human MutL homolog genes, hPMS1 and hPMS2, which are located on the q and p arms of chromosome 2 and 7, respectively.37 Germ-line mutations of each human MutL homolog were found in HNPCC kindreds, with defects in hMLH1 present in the majority of the HNPCC cases.36,37 Since the initial identification of HNPCC-linked genes, HNPCC kindreds have been extensively screened for mutations in each of these genes. It is now clear that mutations of hMSH2 and hMLH1 account for the vast majority of all HNPCC kindreds tested while mutations in hPMS1, hPMS2 and MSH6 are rare. No germ-line mutations in hMSH3 have yet been identified in HNPCC patients.2,4,5 The observed distribution of mutations of these genes in HNPCC is consistent with the relative importance of their functional roles in MMR, as judged by the fact that the protein products of hMSH2 and hMLH1 are obligatory components of all MMR-associated hMutS and hMutL heterodimers known to date. A fourth human MutL homolog gene, hMLH3, has been identified,38 but its involvement in the tumorigenesis of HNPCC is controversial. A controversy also exists in terms of whether or not germ-line mutations of hEXOI are linked to HNPCC. Nevertheless, recombinant hEXOI proteins carrying these alterations have been shown to either lack the exonuclease activity or display a reduced capacity to interact with hMSH239 ; mice defective in EXOI exhibit partial genomic instability, and are partially defective in strandspecific MMR and susceptible to cancer.23 3.3. Restoration of MMR to Tumor Cells with MSI by MMR Gene Products The most convincing evidence that the HNPCC syndrome is caused by MMR defects may come from the biochemical studies of this

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disease. Biochemical assays of extracts prepared from a number of cell lines that were derived from HNPCC and sporadic tumors with MSI have clearly demonstrated that these cells are deficient in strand-specific MMR. Further characterization of these cell lines has defined at least two in vitro complementation groups, which led to the isolation of hMutLα40 and hMutSα.41 Purified hMutLα or hMutSα restores strand-specific MMR to nuclear extracts derived from tumor cell lines that are defective in hMLH1/hPMS2 or hMSH2/hMSH6, respectively. Strong evidence supporting the concept that MMR genes are crucial to genomic stability was also provided by chromosome or gene transfer experiments.42–45 Boland and colleagues reported that the transfer of human chromosome 3 carrying the wild-type hMLH1 gene to an hMLH1-deficient colorectal tumor cell line restores MMR to the cell line.42 Similarly, human chromosome 2 containing both the hMSH2 and hMSH6 genes can complement MMR defects in hMSH2- and hMSH6-deficient tumor cell lines.43,44 Restoration of MMR to cell lines defective in hPMS2-, hMLH1- or hMSH6 has also been achieved by the introduction of the corresponding genes into these lines. Most strikingly, the transfected genes or chromosomes also stabilize simple repetitive sequence in the host cells. These studies further confirmed that the MMR system plays an essential role in the maintenance of genomic stability and showed the potential of gene therapy in the treatment of HNPCC. 3.4. Microsatellite Instability and MMR Deficiency in Non-Colorectal Cancers The identification of MSI in HNPCC patients in 1993 led to the elucidation of the molecular pathogenesis of this disease. Since then, a great body of work has been published, demonstrating that MSI is also associated with a wide variety of non-HNPCC and noncolonic tumors (for detailed reviews see Refs. 46 and 47). These tumors include endometrial, ovarian, gastric, cervical, breast, skin, lung, glioma, prostate, bladder, leukemia and lymphoma. Because

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these studies were carried out using different microsatellite markers and employing different numbers of samples, it is not surprising that the observed mutation rates varied from study to study, and, in some cases, were not in agreement with one another. To standardize MSI studies internationally, the Bethesda guidelines were developed, which suggest that at least five loci should be used in MSI studies; instability in one of the five loci will be scored as “MSI-L” and instability in two or more loci will be scored as “MSI-H”.47 Based on the Bethesda guidelines, non-colorectal tumors also exhibit the MSI-H and MSI-L phenotypes.47 Many sporadic endometrial and gastric tumors, lung cancers and lymphomas display a high level of MSI in many markers (reviewed in Ref. 20). Some tumors demonstrated greater instability in one marker than another. In this regard, tumors with MSI can be divided into two groups: one that displays elevated instability at mono- and di-nucleotide markers and, to a lesser degree, at larger repeat markers, and a second group that displays elevated instability only at specific larger repeat markers, such as tri- and tetranucleotide repeats. Endometrial and gastric tumors usually belong to the first group, while bladder, lung, head and neck cancers belong to the second group.47 The presence of MSI in sporadic non-colonic tumors stimulated a search for somatic mutations in MMR genes, particularly the hMSH2 and hMLH1 genes in these tumors. Although somatic mutations in each of the MMR genes in sporadic cancer and cell lines with MSI have been documented, the major mechanism underlying MMR deficiency in sporadic cancers was shown to be epigenetic silencing, i.e. promoter hypermethylation of MMR genes, especially the hMLH1 gene (see below for details). Biochemical studies showed that MSI-cell lines derived from sporadic leukemia, endometrial, ovarian, prostate, and bladder cancers were defective in strand-specific MMR.40,41,48 These findings suggest that the loss of MMR function is a likely cause of non-colonic sporadic cancer with MSI, although other mechanisms may also be involved in causing the MSI mutator phenotype.

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3.5. Mice with MMR Gene Knockouts Predisposed to Cancers To understand the relationship between MMR defects and tumorigenesis in HNPCC, mice with a knockout mutation in each MMR gene have been developed (reviewed in Refs. 2, 9, 23, 49). Although most of the knockout mice displayed a typical mutator phenotype (e.g. exhibiting MSI) and a predisposition to develop cancer, it is surprising that none of these MMR-deficient mice developed colon cancer as in HNPCC. Instead, a significant number of these animals developed lymphomas, particularly those deficient in MSH2, MLH1 and PMS2. In addition, these knockout animals also developed gastrointestinal tumors, skin neoplasms and/or sarcomas.2,9,23,49 The MSH2-deficient mice developed normally, and both male and female mice were fertile50,51 ; however, although there seemed to be no distinguishable differences between wild-type mice and heterozygous MSH2+/− mice, the homozygous MSH2−/− mice had a much shorter lifespan, with 50% dying at the age of six months. The MSH3−/− mice, although defective in the repair of small insertion/deletion mispairs, exhibited a tumor susceptibility phenotype that is similar to wild-type mice.52 The lack of a significant cancer phenotype in the MSH3-knockout mice provides an explanation as to why germ-line mutations of the human MSH3 gene have not been identified in HNPCC patients. The MSH6−/− mice displayed a tumor spectrum similar to MSH2−/− mice and usually developed tumors within their first year of life.50,53 The MSH6−/− mice did not show the typical MSI phenotype that is detected in MSH2−/− mice. Interestingly, the mutations and pathology observed in the MSH6−/− mice seemed to be similar to that observed in atypical HNPCC cases with hMSH6 mutations, which were characterized by a late cancer onset (> 60 years of age) and low rates of MSI.54 Mice defective in MutL homologs (MLH) share many of the MSH −/− characteristics in terms of cancer spectrum and genomic instability. However, a striking feature that is unique to MLH mutant mice (except PMS1 mutants) is that they are infertile.55–57 It has been shown that while both male and female mice defective in MLH1 or

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MLH3 were sterile,55–57 only male PMS2-knockouts were sterile.58 Unlike all of the MMR-deficient mice described above, mice deficient in the PMS1 gene showed no instability in repeat sequences except for a small mutation rate in mononucleotide repeats.49 Most strikingly, the PMS1-knockout mice did not develop any tumors.49 Mice defective in EXO1 behaved like the MLH1−/− mice, as they developed lymphoma, exhibited MSI, and were infertile.23 In summary, although MMR-deficient mice do not develop colon cancer and there are variations among these knockouts in terms of cancer spectra and mutator phenotypes (see Table 2, and Ref. 9), these studies certainly support the view that MMR defects lead to genomic instability and eventually to cancer, as originally proposed based on studies of the HNPCC syndrome. In addition, the phenotypes of the individual MMR gene knockouts are basically consistent with the role of their gene products in MMR, as characterized by previous biochemical and genetic studies.

Table 2.

Phenotypes of MMR-deficient Knockout Mice

Gene

MSI

MSH2

Yes

MSH3 MSH6

Yes Low instability in dinucleotide repeats Yes

MLH1

PMS1 PMS2 MLH3 EXO1

Mononucleotide repeats only Yes Yes Yes

Tumor Lymphoma, GI, skin, and other tumors GI tumors Lymphoma, GI and other tumors Lymphoma, GI, skin, and other tumors None Lymphoma and sarcoma Not available Lymphoma

Fertility

Reference

Yes

(50,51)

Yes Yes

(52,73) (52,53,73)

No

(55,56)

Yes

(49)

Male only

(49,58)

No No

(57) (23)

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3.6. Epigenetic Silencing of MMR Gene Expression Leads to Sporadic Cancers As described above, mutations in MMR genes that account for the hypermutable phenotype are associated with the HNPCC syndrome and a subset of sporadic colorectal cancers with MSI. However, in a significant fraction of sporadic colon tumors displaying MSI, no identifiable mutations have been found in known MMR genes, suggesting that a different mechanism may be involved in causing MSI in these cases. Surprisingly, the search for this mechanism has linked these tumors again to MMR defects. This time, however, an epigenetic factor, methylation, is responsible for suppressing the expression of MMR genes. Kane et al.59 demonstrated that hypermethylation of the hMLH1 promoter was correlated with a lack of hMLH1 expression in several sporadic colon tumors and cell lines that were free of mutations in the hMLH1 gene. Therefore, hypermethylation is probably a common mode of MMR gene inactivation in sporadic cancer.59 Since then, hypermethylation of the hMLH1gene has been extensively studied (reviewed in Ref. 60). According to the Bethesda guidelines,47 sporadic tumors can be classified into three types based on their MSI status in five sets of microsatellite markers: microsatellite stable (MSS, instability observed in none of the five markers), low-frequency MSI (MSI-L, instability observed in one of the five markers), and highfrequency MSI (MSI-H, instability observed in two or more markers). It has been reported that more than 95% of MSI-H tumors are due to the loss of expression of hMLH1,61 and interestingly, almost all MSI-H tumors that do not have a detectable mutation within the hMLH1 gene demonstrate hypermethylation in the hMLH1 promoter. Recently, hypermethylation of the hMLH1 promoter has been shown in an HNPCC patient, who does not have germ-line mutations in any of the known MMR genes.62 In contrast, hypermethylation of the hMSH2 gene is rarely observed in tumors with MSI. To determine the nature of the hypermethylation of the hMLH1 promoter in these MSI tumors, two independent research groups, led by Markowitz and Herman, treated several tumor cell lines with aberrant hMLH1 expression due to hypermethylation in the

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hMLH1 promoter with the demethylating agent 5-aza-deoxycytidine (reviewed in Refs. 60 and 63). This treatment successfully restored hMLH1 protein expression in all the tumor cells that lacked hMLH1 expression due to a methylated hMLH1 promoter, and the extracts derived from 5-aza-deoxycytidine-treated cells were capable of performing strand-specific MMR. These experiments indicate that in addition to genetic defects, epigenetic modification of MMR genes can also result in a mutator phenotype. 3.7. MMR Deficiency and Inactivation of Genes Critical for Cellular Growth Control and Genomic Stability Despite evidence described above that MMR genes function like tumor suppressor genes, the MMR pathway is a mutation avoidance system or a “caretaker” system.64 It is anticipated that the loss of MMR function will affect the stability of many other genes, including critical “gatekeeper” genes (e.g. tumor suppressor genes) and “caretaker genes” (e.g. DNA repair genes). Because of technical limitations, it is impossible at this time to assess the impact of MMRdeficiency on a genome-wide basis and to identify all the mutations that accumulate as a result of MMR-deficiency. However, using MSI analysis, it is possible to readily detect frameshift mutations in genes that contain simple repeat sequences within their coding regions, which in most cases lead to truncated proteins. Markowitz et al.65 reported that mutations in the type II transforming growth factor-β receptor (TGF-β RII) gene were associated with sporadic colorectal cancer cells defective in MMR. These mutations were all frameshift mutations and occur either in a six-bp GTGTGT repeat or in an (A)10 mononucleotide repeat.65 In each case, the frameshift mutation resulted in a mutant form of the TGFβ RII. Subsequent studies demonstrated that frameshift mutations of simple repeat tracts in the TGF-β RII gene were common in colorectal tumors with MSI. Similar mutations of TGF-β RII have also been observed in many types of MSI cancer, including gastric cancer, glioma, uterine cervical cancer, squamous carcinoma of the head and neck, ulcerative colitis-associated neoplasm and sporadic cecum

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cancer. It is known that TGF-β RII is required for transduction of the TGF-β growth inhibitory signal to suppress epithelial cell growth. The loss of the TGF-β RII function in tumors with MSI represents a crucial mechanism by which cells may escape from growth control. The targeted mutations in the simple repeated sequences of the TGFβ RII gene may be characteristic of the genome-wide alterations that are expected in MMR-deficient cells. In addition to TGF-β RII, somatic frameshift mutations of mononucleotide runs have been documented in several genes that are critical for cellular growth in tumors with MSI. These genes include the apoptosis gene Bax, insulin-like growth factor 2 receptor IGF2-R, transcription factor E2F-4, tumor suppressor genes APC and PTEN, and DNA repair genes hMSH3, hMSH6, Mre11, and MBD4/MED1. All of these genes are crucial either for cellular growth control or for genomic stability; and the inactivation of any of these genes could be a key mechanism by which tumors with MSI become neoplastic. Therefore, the potential impact of the loss of the tumor suppressor function of MMR is not only relevant to HNPCC, but to virtually all types of cancer. 4. THE MMR SYSTEM MEDIATES DNA DAMAGE SIGNALING 4.1. MMR Deficiency and Drug Resistance While MMR is well known for its role in correcting biosynthetic errors, other important roles for MMR proteins have also been identified, one of which is to mediate cell cycle arrests and programmed cell death (or apoptosis) of cells with heavily damaged DNA (for a review see Ref. 7). The identification of the involvement of the MMR system in DNA damage signaling developed from research on how chemical and physical DNA damaging agents cause cell death. Treatment of cells with chemical DNA-damaging agents such as the alkylating agents N-methyl-N’-nitro-N-nitrosoguanidine (MNNG), temozolomide, or procarbazine leads to increased cell death. These cytotoxic agents are therefore often used in chemotherapy to destroy rapidly growing tumor cells. It has been found almost

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universally that while cells that are proficient in MMR are sensitive to these agents, cells that are deficient in MMR are more resistant to killing by these agents. The phenomenon, which was initially observed in E. coli MMR mutants in the early 1980s, also applies to human cells. In fact, the first human MMR mutant cell line, MTI, was derived from the TK6 lymphoblastoid cell line under the selection of a high dose of MNNG. The MT1 cell line was subsequently found to harbor a mutation in hMSH6 and to be defective in strandspecific MMR. MMR-deficiency can be acquired following treatment with the alkylating agent MNNG, whereas MMR-deficient cell lines derived from HNPCC and MSI tumors are also resistant to alkylating agents. For example, an hMLH1-defective colorectal tumor cell line is resistant to killing by MNNG, but it becomes sensitive to the agent when it receives a wild-type copy of hMLH1 from the transfer of chromosome 3. Similarly, cells defective in other MMR genes, e.g. MSH2 and PMS2, also confer resistance to alkylating agents. Drug resistance has also been found to be associated with changes in the expression profiles of MMR genes, which results in a loss of MMR function. For example, treatment of the HL60 leukemia cells with methotrexate (MTX), a frequently used chemotherapeutic drug for cancer, induces over-expression of the hMSH3 gene. The primary target for MTX is dihydrofolate reductase (DHFR), a key enzyme that catalyzes the reduction of dihydrofolate to tetrahydrofolate in a reaction essential for nucleotide metabolism. It has been well documented that human tumor cells acquire resistance to MTX as a result of the amplification of a chromosomal region containing the DHFR gene, leading to an elevated expression of DHFR. Interestingly, the DHFR gene shares a promoter with a second gene (now known as hMSH3) that is transcribed in the opposite direction. Instead of enhancing MMR activity, over-expression of hMSH3 renders the MTX-treated cells to be defective in MMR, which is believed to be responsible for the MTX-resistant phenotype observed in HL60 and other tumor cells. This is because both hMSH3 and hMSH6 interact with hMSH2 to form hMutSα and hMutSβ, respectively, and over-expression of hMSH3 allows the protein to capture almost all cellular hMSH2 so that little hMutSα can be formed. At the same

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time, uncomplexed hMSH6 is subject to degradation. In normal conditions, the cellular hMutSα:hMutSβ ratio is ∼ 10:1, and cells with no hMutSα or a lower ratio of hMutSα:hMutSβ are hypermutable. This is because hMutSα can repair both base-base and ID mispairs, but hMutSβ could only process the ID mispairs. These observations clearly indicate that drug resistance is closely associated with MMR deficiency caused either by genetic mutations or by changes in the expression profiles of MMR genes. 4.2. MMR Proteins Promote DNA Damage-Induced Cell Cycle Arrest and Apoptosis The distinct responses to DNA-damaging agents between MMRproficient and MMR-deficient cells have stimulated research on how the cell responds to DNA damage. Studies from the laboratories of Richard Boland and Josef Jiricny have shown that cells proficient in MMR underwent growth arrest at the G2-phase of the cell cycle after treatment with MNNG or 6-thioguanine (6-TG), but the G2-phase arrest was not observed in cells deficient in MMR under the same treatment. A recent study by Brown, Baskaran, and their colleagues has indicated that the MMR system is also required for the activation of the S-phase checkpoint in response to ionizing radiation.66 In addition, cytotoxicity of DNA damaging agents is associated with apoptosis, and the apoptotic response only occurs in MMR-proficient cells.7 Recent studies have indicated that both p53 and the related protein p73 are implicated in MMR-dependent apoptosis. Upon treating the cells with DNA damaging agents, phosphorylation of p53 and/or p73 has been noted in MMR-proficient cells, but not in cells defective in either hMutSα or hMutLα.67,68 ATM, ATR, and/or c-Abl appear to be the kinases that phosphorylate these proteins during the damage response.68,69 Physical interactions between MMR proteins (e.g. hMutSα and hMutLα) and proteins involved in the DNA damage-signaling network (e.g. ATM and p73) have been recently identified.66,70 These observations indicate that MMR-dependent apoptosis in response to DNA damage involves a signaling cascade.

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4.3. MMR-Mediated Apoptosis Eliminates Potentially Tumorigenic Cells The molecular events involved in the MMR-dependent apoptotic response have not yet been established. However, increasing evidence suggests that the apoptotic signaling is initiated by MMR proteins. Two models have been proposed to account for MMRmediated apoptosis. One model proposes that a repetitive attempt by MMR to remove a DNA adduct in the template DNA strand causes cell death. DNA adducts in the template strand can pair with appropriate bases or lead to misincorporations during DNA replication. hMutSα, along with hMutLα, recognizes these unusual base pairs as “mismatches” and provokes a strand-specific MMR reaction. However, because MMR only targets the newly synthesized strand, adducts in the template strand cannot be removed, and thus the unusual base pairs reform upon DNA resynthesis during repair. As a result, the repair cycle may be perpetually reinitiated. Such a futile repair cycle may signal cells to switch on the apoptotic machinery. A recent study from the Jiricny laboratory appears to support this model, where they observed the formation of ssDNA gaps in wild-type cells treated with MNNG. A second model suggests that the death signal could come from the binding of hMutSα/hMutLα to DNA adducts in the replication fork and/or the unwound DNA helix. These protein-adduct complexes may block DNA transactions such as replication, transcription, and repair, and could be recognized as a signal for cell cycle arrest and apoptosis. Both models provide an explanation for the selective growth advantage of cells that are defective in MMR. Strong support for these models is provided by the fact that MutS and its eukaryotic homologs are capable of recognizing a variety of DNAadducts caused by DNAdamaging agents (reviewed in Ref. 7), such as MNNG, cisplatin, environmental chemical carcinogens, oxidative free radicals, and ultraviolet (UV) light. As described above, the repair function of the MMR pathway prevents mutations from building up in the genome by correcting mispairs. The apoptotic function of the pathway, however, maintains genomic stability by promoting apoptosis of cells with severely damaged DNA. Normally, base excision repair (see Chapter 2 for

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more details) and nucleotide excision repair (see Chapter 3 for more details) pathways are responsible for the repair of DNA damage induced by physical and chemical agents. However, when these pathways are not available or there is too much damage to be repaired, the genome is in danger of accumulating a large number of mutations that are tumorigenic. Therefore, eliminating these damaged cells from the body would be beneficial. The MMR system is capable of activating the apoptotic machinery to eliminate these pre-tumorigenic cells from growing by promoting apoptosis. The inability of this system to commit damaged cells for apoptosis is thought to be a molecular basis for cancer development.7,71

5. ROLE OF MMR IN OTHER DNA METABOLIC PATHWAYS It has been known for many years that MMR plays a role in modulating the efficiency and fidelity of DNA recombination, both mitotic and meiotic.5,6 Both genetic and biochemical studies have indicated that MutS-like and MutL-like proteins suppress mitotic recombination between similar, but non-identical (homeologous) DNA sequences. The inactivation of MMR in both E. coli and mammalians leads to an increased frequency of homeologous recombination. In vitro studies have revealed that MutS and MutL proteins inhibit DNAstrand exchange between two divergent sequences, most likely by binding to the mismatches generated during strand exchange. These observations suggest that the in vivo recombination event may be aborted at the stage of branch migration because of the detection of mismatches by MMR proteins. It is also possible that the reaction goes through, but the mismatches created are corrected, eliminating possible sequence changes created during recombination. The demonstration for the involvement of MMR components in meiotic recombination came from yeast and knockout mouse studies.5,9 While mitotic recombination uses the same MMR components that are used for the repair reaction, meiotic recombination appears to involve a different set of MMR proteins. For example,

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gene products of MSH2 and MSH6, subunits of MutSα, are critical components for repair and mitotic recombination, but they appear not to be involved in meiotic recombination, as MSH2−/− and MSH6−/− mutants exhibit no obvious defects in meiosis. However, mice lacking the MutL homolog gene MLH1, PMS2 or MLH3 are sterile because of abnormal meiotic recombination.9 MMR proteins are important for antibody class-switch recombination and somatic hypermutation, processes that occur during antigen-stimulated B-cell differentiation and increase the effectiveness of the humoral immune response. B-cells from mice defective in MSH2, MSH6, MLH1, PMS2 or EXO1 have reduced abilities to undergo class-switch recombination. It is known that class switch recombination is initiated by activation-induced cytidine deaminase (AID), which converts the cytosine to uracil and generates a G:U base pair. Since the AID-generated G:U mismatches can be recognized and processed by the MMR system, it is possible that the EXO1-catalyzed excision event to remove the G:U mismatch by the MMR system can occur simultaneously on both strands at strand breaks created by uracil DNA glycosylases, leading to double strand breaks. The generation of double strand breaks promotes class switch recombination.72 6. CONCLUSION AND PERSPECTIVES The seminal discovery of MMR defects in HNPCC and sporadic cancers with MSI in 1993 has greatly stimulated investigations on MMR and its role in preventing cancer. Almost all human homologs of the E. coli MMR components have now been identified and characterized by both genetic and biochemical approaches. It is now known that MMR maintains genomic stability through correcting biosynthetic errors, modulating DNA recombination, and mediating cell cycle arrests and apoptosis. While the first two functions ensure genomic stability by either removing mispairs or preventing the formation of mispairs, the apoptotic function of the system, however, is to eliminate genetically damaged and pre-tumorigenic cells from the body. Therefore, the loss of MMR function, by either

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genetic mutation or epigenetic modification, will lead to a mutator phenotype and predispose a cell to carcinogenesis. While MMR-mediated apoptosis is important for tumor suppression, the drug resistance property of MMR-deficient cells raises concerns for cancer chemotherapy. First, certain widely used clinical drugs, such as temozolomide, procarbazine, and cisplatin, are expected to be harmful for patients with tumors caused by MMR defects. This is because these drugs will preferentially kill the patient’s normal proliferating tissue via drug-induced apoptosis, rather than the tissue of the tumor. Therefore, chemotherapy should be used with caution for tumors with MMR defects. Second, since some tumor cells can acquire MMR deficiency upon exposure to certain drugs, the use of these drugs in clinical practice may lead to secondary cancers that are characterized by MMR defects. In light of these problems, how can cancer chemotherapy be improved? Several strategies are apparent. First, since the restoration of MMR to MMR-deficient tumor cells by gene transfer can restore their sensitivity to drugs, an improved therapy for MMR-deficient cancer could potentially include treatments that restore the tumors’ MMR function, e.g. by gene or chromosomal transfer technology, prior to drug application. Thus, the MMR-restored tumor should be sensitive to regular chemotherapy. In addition, the development of drugs that could specifically kill MMR-deficient cells but not MMR-proficient cells could lead to progress in future cancer chemotherapy. The genomic maintenance capability of MMR underscores the importance of the MMR pathway in cancer biology and molecular medicine. Despite rapid advances in the field of MMR in the past decade, the molecular mechanisms by which the MMR pathway suppresses homologous recombination, and mediates DNA damage-induced apoptosis are still not fully understood. Understanding these questions will open new strategies for cancertargeted drug design. ACKNOWLEDGMENTS Work in the author’s laboratory was supported by grants from National Institutes of Health (GM072756 and ES013193) and the

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Kentucky Lung Cancer Research Program. The author is the JamesGardner Endowed Chair in Cancer Research. References 1. Loeb LA, Loeb KR, and Anderson JP. (2003) Multiple mutations and cancer. Proc Natl Acad Sci USA 100: 776–781. 2. Kunkel TA, Erie DA. (2005) DNA mismatch repair. Annu Rev Biochem 74: 681–710. 3. Tiraby JG, Fox MS. (1973) Marker discrimination in transformation and mutation of pneumococcus. Proc Natl Acad Sci USA 70: 3541–3545. 4. Kolodner RD, Marsischky GT. (1999) Eukaryotic DNAmismatch repair. Curr Opin Genet Dev 9: 89–96. 5. Modrich P, Lahue R. (1996) Mismatch repair in replication fidelity, genetic recombination, and cancer biology. Annu Rev Biochem 65: 101– 133. 6. Harfe BD, Jinks-Robertson S. (2000) DNA mismatch repair and genetic instability. Annu Rev Genet 34: 359–399. 7. Li GM. (1999) The role of mismatch repair in DNA damage-induced apoptosis. Oncol Res 11: 393–400. 8. Stojic L, Brun R, Jiricny J. (2004) Mismatch repair and DNA damage signalling. DNA Repair (Amst) 3: 1091–1101. 9. Buermeyer AB, Deschenes SM, Baker SM, Liskay RM. (1999) Mammalian DNA mismatch repair. Annu Rev Genet 33: 533–564. 10. Jiricny J. (1998) Replication errors: cha(lle)nging the genome. EMBO J 17: 6427–6436. 11. Yang W. (2000) Structure and function of mismatch repair proteins. Mutat Res 460: 245–256. 12. Schofield MJ, Hsieh P. (2003) DNA mismatch repair: molecular mechanisms and biological function. Annu Rev Microbiol 57: 579–608. 13. Lahue RS, Au KG, Modrich P. (1989) DNA mismatch correction in a defined system. Science 245: 160–164. 14. Obmolova G, Ban C, Hsieh P, Yang W. (2000) Crystal structures of mismatch repair protein MutS and its complex with a substrate DNA. Nature 407: 703–710. 15. Lamers MH, Perrakis A, Enzlin JH, et al. (2000) The crystal structure of DNA mismatch repair protein MutS binding to a G x T mismatch. Nature 407: 711–717. 16. Sancar A, Hearst JE. (1993) Molecular matchmakers. Science 259: 1415– 1420. 17. Ban C, Yang W. (1998) Crystal structure and ATPase activity of MutL: implications for DNA repair and mutagenesis. Cell 95: 541–552.

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18. Junop MS, Obmolova G, Rausch K, et al. (2001) Composite active site of an ABC ATPase: MutS uses ATP to verify mismatch recognition and authorize DNA repair. Mol Cell 7: 1–12. 19. Ban C, Yang W. (1998) Structural basis for MutH activation in E. coli mismatch repair and relationship of MutH to restriction endonucleases. EMBO J 17: 1526–1534. 20. Li GM. (2003) DNA mismatch repair and cancer. Front Biosci 8: d997– 1017. 21. Zhang Y, Yuan F, Presnell SR, et al. (2005) Reconstitution of 5 -directed human mismatch repair in a purified system. Cell 122: 693–705. 22. Dzantiev L, Constantin N, Genschel J, et al. (2004) A defined human system that supports bidirectional mismatch-provoked excision. Mol Cell 15: 31–41. 23. Wei K, Clark AB, Wong E, et al. (2003) Inactivation of exonuclease 1 in mice results in DNA mismatch repair defects, increased cancer susceptibility, and male and female sterility. Genes Dev 17: 603–614. 24. Lee BI, Wilson DM, III. (1999) The RAD2 domain of human exonuclease 1 exhibits 5 to 3 exonuclease and flap structure-specific endonuclease activities. J Biol Chem 274: 37763–37769. 25. Peltomaki P, Lothe RA, Aaltonen LA, et al. (1993) Microsatellite instability is associated with tumors that characterize the hereditary nonpolyposis colorectal carcinoma syndrome. Cancer Res 53: 5853–5855. 26. Aaltonen LA, Peltomaki P, Leach FS, et al. (1993) Clues to the pathogenesis of familial colorectal cancer. Science 260: 812–816. 27. Ionov Y, Peinado MA, Malkhosyan S, et al. (1993) Ubiquitous somatic mutations in simple repeated sequences reveal a new mechanism for colonic carcinogenesis. Nature 363: 558–561. 28. Thibodeau SN, Bren G, Schaid D. (1993) Microsatellite instability in cancer of the proximal colon. Science 260: 816–819. 29. Strand M, Prolla TA, Liskay RM, Petes TD. (1993) Destabilization of tracts of simple repetitive DNA in yeast by mutations affecting DNA mismatch repair. Nature 365: 274–276. 30. Fishel R, Lescoe MK, Rao MR, et al. (1993) The human mutator gene homolog MSH2 and its association with hereditary nonpolyposis colon cancer. Cell 75: 1027–1038. 31. Leach FS, Nicolaides NC, Papadopoulos N, et al. (1993) Mutations of a mutS homolog in hereditary nonpolyposis colorectal cancer. Cell 75: 1215–1225. 32. Parsons R, Li GM, Longley MJ, et al. (1993) Hypermutability and mismatch repair deficiency in RER+ tumor cells. Cell 75: 1227–1236. 33. Umar A, Boyer JC, Thomas DC, et al. (1994) Defective mismatch repair in extracts of colorectal and endometrial cancer cell lines exhibiting microsatellite instability. J Biol Chem 269: 14367–14370.

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34. Lindblom A, Tannergard P, Werelius B, Nordenskjold M. (1993) Genetic mapping of a second locus predisposing to hereditary non-polyposis colon cancer. Nat Genet 5: 279–282. 35. Bronner CE, Baker SM, Morrison PT, et al. (1994) Mutation in the DNA mismatch repair gene homologue hMLH1 is associated with hereditary nonpolyposis colon cancer. Nature 368: 258–261. 36. Papadopoulos N, Nicolaides NC, Wei YF, et al. (1994) Mutation of a mutL homolog in hereditary colon cancer. Science 263: 1625–1629. 37. Nicolaides NC, Papadopoulos N, Liu B, et al. (1994) Mutations of two PMS homologues in hereditary nonpolyposis colon cancer. Nature 371: 75–80. 38. Lipkin SM, Wang V, Jacoby R, et al. (2000) MLH3: a DNA mismatch repair gene associated with mammalian microsatellite instability. Nat Genet 24: 27–35. 39. Sun X, Zheng L, Shen B. (2002) Functional alterations of human exonuclease 1 mutants identified in atypical hereditary nonpolyposis colorectal cancer syndrome. Cancer Res 62: 6026–6030. 40. Li GM, Modrich P. (1995) Restoration of mismatch repair to nuclear extracts of H6 colorectal tumor cells by a heterodimer of human MutL homologs. Proc Natl Acad Sci USA 92: 1950–1954. 41. Drummond JT, Li GM, Longley MJ, Modrich P. (1995) Isolation of an hMSH2-p160 heterodimer that restores DNA mismatch repair to tumor cells. Science 268: 1909–1912. 42. Koi M, Umar A, Chauhan DP, et al. (1994) Human chromosome 3 corrects mismatch repair deficiency and microsatellite instability and reduces N-methyl-N’-nitro-N-nitrosoguanidine tolerance in colon tumor cells with homozygous hMLH1 mutation. Cancer Res 54: 4308– 4312. 43. Tindall KR, Glaab WE, Umar A, et al. (1998) Complementation of mismatch repair gene defects by chromosome transfer. Mutat Res 402: 15–22. 44. Watanabe Y, Haugen-Strano A, Umar A, et al. (2000) Complementation of an hMSH2 defect in human colorectal carcinoma cells by human chromosome 2 transfer. Mol Carcinog 29: 37–49. 45. Buermeyer AB, Wilson-Van Patten C, Baker SM, Liskay RM. (1999) The human MLH1 cDNA complements DNA mismatch repair defects in Mlh1-deficient mouse embryonic fibroblasts. Cancer Res 59: 538–541. 46. Eshleman JR, Markowitz SD. (1995) Microsatellite instability in inherited and sporadic neoplasms. Curr Opin Oncol 7: 83–89. 47. Boland CR, Thibodeau SN, Hamilton SR, et al. (1998) A National Cancer Institute Workshop on Microsatellite Instability for cancer detection and familial predisposition: development of international criteria

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49.

50.

51. 52.

53. 54. 55.

56. 57. 58.

59.

60. 61.

62.

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for the determination of microsatellite instability in colorectal cancer. Cancer Res 58: 5248–5257. Gu L, Cline-Brown B, Zhang F, et al. (2002) Mismatch repair deficiency in hematological malignancies with microsatellite instability. Oncogene 21: 5758–5764. Prolla TA, Baker SM, Harris AC, et al. (1998) Tumor susceptibility and spontaneous mutation in mice deficient in Mlh1, Pms1 and Pms2 DNA mismatch repair. Nat Genet 18: 276–279. de Wind N, Dekker M, Berns A, et al. (1995) Inactivation of the mouse Msh2 gene results in mismatch repair deficiency, methylation tolerance, hyper-recombination, and predisposition to cancer. Cell 82: 321–330. Reitmair AH, Schmits R, Ewel A, et al. (1995) MSH2 deficient mice are viable and susceptible to lymphoid tumors. Nat Genet 11: 64–70. Edelmann W, Umar A, Yang K, et al. (2000) The DNA mismatch repair genes Msh3 and Msh6 cooperate in intestinal tumor suppression. Cancer Res 60: 803–807. Edelmann W, Yang K, Umar A, et al. (1997) Mutation in the mismatch repair gene Msh6 causes cancer susceptibility. Cell 91: 467–477. Kolodner RD, Tytell JD, Schmeits JL, et al. (1999) Germ-line msh6 mutations in colorectal cancer families. Cancer Res 59: 5068–5074. Baker SM, Plug AW, Prolla TA, et al. (1996) Involvement of mouse Mlh1 in DNA mismatch repair and meiotic crossing over. Nat Genet 13: 336–342. Edelmann W, Cohen PE, Kane M, et al. (1996) Meiotic pachytene arrest in MLH1-deficient mice. Cell 85: 1125–1134. Lipkin SM, Moens PB, Wang V, et al. (2002) Meiotic arrest and aneuploidy in MLH3-deficient mice. Nat Genet 31: 385–390. Baker SM, Bronner CE, Zhang L, et al. (1995) Male mice defective in the DNA mismatch repair gene PMS2 exhibit abnormal chromosome synapsis in meiosis. Cell 82: 309–319. Kane MF, Loda M, Gaida GM, et al. (1997) Methylation of the hMLH1 promoter correlates with lack of expression of hMLH1 in sporadic colon tumors and mismatch repair-defective human tumor cell lines. Cancer Res 57: 808–811. Grady WM, Markowitz SD. (2002) Genetic and epigenetic alterations in colon cancer. Annu Rev Genomics Hum Genet 3: 101–128. Thibodeau SN, French AJ, Cunningham JM, et al. (1998) Microsatellite instability in colorectal cancer: different mutator phenotypes and the principal involvement of hMLH1. Cancer Res 58: 1713–1718. Gazzoli I, Loda M, Garber J, et al. (2002) A hereditary nonpolyposis colorectal carcinoma case associated with hypermethylation of the MLH1

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66. 67.

68.

69.

70.

71.

72.

73.

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gene in normal tissue and loss of heterozygosity of the unmethylated allele in the resulting microsatellite instability-high tumor. Cancer Res 62: 3925–3928. Herman JG, Baylin SB. (2003) Gene silencing in cancer in association with promoter hypermethylation. N Engl J Med 349: 2042–2054. Kinzler KW, Vogelstein B. (1997) Cancer-susceptibility genes. Gatekeepers and caretakers. Nature 386: 761, 763. Markowitz S, Wang J, Myeroff L, et al. (1995) Inactivation of the type II TGF-beta receptor in colon cancer cells with microsatellite instability. Science 268: 1336–1338. Brown KD, Rathi A, Kamath R, et al. (2003) The mismatch repair system is required for S-phase checkpoint activation. Nat Genet 33: 80–84. Duckett DR, Drummond JT, Murchie AI, et al. (1996) Human MutSalpha recognizes damaged DNA base pairs containing O6methylguanine, O4-methylthymine, or the cisplatin-d(GpG) adduct. Proc Natl Acad Sci USA 93: 6443–6447. Gong JG, Costanzo A, Yang HQ, et al. (1999) The tyrosine kinase c-Abl regulates p73 in apoptotic response to cisplatin-induced DNA damage. Nature 399: 806–809. Stojic L, Mojas N, Cejka P, et al. (2004) Mismatch repair-dependent G2 checkpoint induced by low doses of SN1 type methylating agents requires the ATR kinase. Genes Dev 18: 1331–1344. Shimodaira H, Yoshioka-Yamashita A, Kolodner RD, Wang JY. (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. Fishel R. (2001) The selection for mismatch repair defects in hereditary nonpolyposis colorectal cancer: revising the mutator hypothesis. Cancer Res 61: 7369–7374. Stavnezer J, Schrader CE. (2006) Mismatch repair converts AIDinstigated nicks to double-strand breaks for antibody class-switch recombination. Trends Genet 22: 23–28. de Wind N, Dekker M, Claij N, et al. (1999) HNPCC-like cancer predisposition in mice through simultaneous loss of Msh3 and Msh6 mismatch-repair protein functions. Nat Genet 23: 359–362.

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CHAPTER 5

Mammalian Homologous Recombination Repair and Cancer Intervention Zhiyuan Shen∗,†,‡ and Jac A. Nickoloff †

ABSTRACT Homologous recombination (HR) is highly relevant to cancer intervention for two major reasons. First, defects or deregulation in HR systems are major contributing factors in tumorigenesis. HR is important for the accurate repair of DNA double-strand breaks and DNA interstrand crosslinks, and it restarts stalled or collapsed replication forks. During these processes, HR restores the gross structure of DNA with minimum probability of mutation. HR is also tightly co-ordinated with mitosis and cell cycle checkpoint regulation, to ensure that cells with excessive DNA damage have minimum chance of being propagated. Thus, HR plays pivotal roles in maintaining genome stability and preventing tumorigenesis. Although HR is generally an error-free DNA repair process, hyper-recombination, hypo-recombination and altered HR outcomes can all lead to genomic instability. Thus HR can be mutagenic and contribute to genomic instability when it is not properly regulated. Second, the status of HR in cancer cells may be used as a marker for tailoring individual cancer therapy and improving prognosis evaluation. Current studies are focused on the stepwise DNA structural changes that occur during HR, and on the identification and characterization of proteins involved in HR. There are ∗ Corresponding

author. of Molecular Genetics and Microbiology, University of New Mexico School of Medicine, Albuquerque, New Mexico 87131, USA. ‡ Division of Radiation Cancer Biology, Dept. of Radiation Oncology, UMDNJ-Robert Wood Johnson Medical School, The Cancer Institute of New Jersey, 195 Little Albany St, New Brunswick, NJ 08903-2681, USA. Tel.: 732-235-6101; E-mail: [email protected] † Department

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two categories of HR proteins: those with enzymatic activity that directly promote the chemical reactions during HR, and accessory proteins that regulate HR enzymes and co-ordinate HR reactions with other important cellular processes. It is important to fully identify and characterize the proteins of both categories as they may serve as targets for cancer therapy, or as markers for diagnosis, prognosis and treatment monitoring. In addition, a detailed picture of HR protein structure and function may provide new opportunities for cancer intervention through improvements in our understanding of cellular responses to therapeutic DNA damage.

1. INTRODUCTION Homologous recombination (HR) involves the exchange of DNA sequence information between homologous regions in a genome. Information exchange can be reciprocal, resulting in a crossover, or non-reciprocal, which is termed gene conversion. A fraction of gene conversions are associated with crossovers (Fig. 1). During gene conversion, sequence information is transferred from a donor to a recipient duplex. If the interacting sequences are heterozygous, this information transfer results in the loss of heterozygosity (LOH). Crossovers change linkage relationships among markers flanking the crossover point, but there is no net loss or gain of information. However, when crossovers occur between homologous chromosomes in G2 cells, chromosome segregation results in large-scale LOH of all markers centromere-distal to the crossover

Fig. 1

Major types of HR products.

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point in 50% of subsequent mitotic divisions.1 In addition to homologous chromosomes, interactions can occur between sister chromatids in the S and G2 phases of the cell cycle, and between regions of shared homology within a chromosome or on non-homologous chromosomes. A crossover between sister chromatids is called a sister chromatid exchange (SCE). Most SCEs have no genetic consequence because sister chromatids are usually identical. However, SCEs between repeated elements can occur in an “unequal” manner, resulting in repeat expansion and contraction in daughter cells. Other potentially deleterious or lethal rearrangements result when crossovers occur between inverted repeats, which cause sequence inversion, or between repeats on non-homologous chromosomes, which causes translocations.1 A well-understood function of HR in mammals is the creation of genetic diversity in meiosis.2 During gametogenesis, recombination between the two homologous chromosomes (one from each parent) increases genetic diversity in the subsequent generation. In recent years it has become increasingly clear that HR plays critical roles in the repair of DNA damage in mitotic cells, particularly DNA double-strand breaks (DSBs) and interstrand crosslinks, and in restarting replication forks blocked by DNA lesions produced by the reactive intermediates of normal cellular metabolism, and by exogenous chemicals and radiation. In these mitotic roles, HR promotes cell survival and preserves genome stability. Tumorigenesis is a multi-step process that often requires the inactivation of tumor suppressors and/or activation of oncogenes. These genetic changes are normally quite rare, but can occur far more frequently in cells with defects in one or more genome stabilizing functions. Genomic instability originates from four major sources: error-prone or mutagenic repair of DNA damage, infidelity of DNA replication, chromosome segregation errors during mitosis, and cell cycle checkpoint defects. HR is a form of relatively error-free DNA repair, and it promotes accurate DNA replication. Some HR proteins promote accurate chromosome segregation and/or interface with the cell cycle checkpoint machinery. For these reasons, HR roles in maintaining genome stability are important factors in cancer

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etiology. In addition, HR also directly modulates the cellular sensitivity to DNA damage produced by chemo- and radiotherapeutics during cancer treatment. Thus, HR proteins are increasingly used as tumor markers for diagnosis, in the design of customized cancer therapy regimens, and as drug targets to enhance tumor sensitivity to therapeutic agents. In this chapter, the authors discuss HR mechanisms and proteins in the context of cancer intervention. Readers are referred to recent reviews for additional perspectives on HR mechanisms, genetic consequences and relationships to disease.3–5 2. DNA DOUBLE-STRAND BREAK REPAIR BY HR Cellular DNA is under constant attack from endogenous agents such as reactive oxygen species produced during normal cellular metabolism, and exogenous agents such as radiation and environmental carcinogens. This damage must be repaired to ensure cell survival, but repair can occur by “error-prone” or “error-free” mechanisms — both types restore the gross DNA structure but error-prone repair has a greater risk of introducing mutations. DSBs are a highly lethal form of DNA damage that are produced directly by ionizing radiation, reactive oxidative species and endonucleases. DSBs can also be produced indirectly from the processing of other types of DNA damage, such as when a replication fork encounters a single-strand break, or from nucleotide excision repair (NER) of interstrand crosslinks. There are two major DSB repair pathways in mammalian cells: non-homologous end joining (NHEJ) and HR. During NHEJ, the broken DNA ends are processed by nucleases, aligned based on very limited or no homology at the ends, and then ligated to rejoin the broken ends (see Chapter 7). As a result, NHEJ often results in the loss (and sometimes gain) of sequence information at the DSB site, and two ends from different chromosomes can be rejoined causing chromosome translocations. Therefore, although NHEJ is efficient and promotes cell survival, it is an error-prone process. In contrast, HR uses homologous DNA as a template, usually a sister chromatid or a homologous chromosome, and is therefore more accurate than NHEJ. However, as noted above, HR can result

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in localized or large-scale LOH, and other genome rearrangements, particularly when associated with crossovers. Thus, a proper balance of HR is needed to allow the accurate repair of DNA damage while minimizing the risks of deleterious, HR-mediated genome rearrangements. Several mechanisms have evolved to achieve this balance. For example, HR is upregulated during the S and G2 phases, when sister chromatids are available.6 Sister chromatids are used preferentially as repair templates,7 and HR between diverged sequences is suppressed, in part by the mismatch repair system.8 There is active suppression of crossovers, particularly between nonhomologous chromosomes.9,10 Thus, genome instability is increased by defects in HR proteins, which shift DSB repair toward error-prone NHEJ, and by defects in HR regulation, which can alter HR frequency and/or outcome. DSB repair by HR in eukaryotes is thought to occur through a coordinated, multi-step mechanism that has been largely defined with yeast systems. Because most HR proteins are conserved from yeast to man, these steps, described below and diagrammed in Fig. 2, are likely to be conserved as well. 2.1. End-Processing and RAD51 Filament Formation Once a DSB is created, a number of proteins are recruited to the site, including the MRE11/RAD50/NBS1 (MRN) complex. MRN regulates a poorly understood pathway that resects ends to produce long, 3 -single-stranded DNA (ssDNA) tails (Fig. 2B). MRE11 has both endonuclease and 3 –5 exonuclease activity. Its exonuclease activity is the incorrect polarity for strand resection, suggesting that the endonuclease may process ends in conjunction with a helicase. In yeast, Exo1 has a role in end-processing, but other nucleases must be involved because processing still occurs in the absence of both Mre11 and Exo1, albeit more slowly.11 ssDNA is initially coated with replication protein A (RPA), which is subsequently replaced by RAD51 with the assistance of “mediator” proteins comprising five RAD51 paralogs (XRCC2, XRCC3, RAD51B, RAD51C, and RAD51D), RAD52 and BRCA2.

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Fig. 2 DNA double-strand break induced homologous recombination (see text for details).

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2.2. Homology Search and Strand Exchange The RAD51 nucleoprotein filament is capable of searching the genome for homologous sequences and catalyzing the strandexchange reaction, forming a D-loop structure (Fig. 2C). There is little known about how initial pairing of the nucleoprotein filament with a donor duplex converts to a normal Watson-Crick base pairing between the invading strand and the complementary strand of the donor duplex. This probably requires the dissociation of RAD51 from the ssDNA and may be catalyzed by Srs2 in yeast.11 However, mammalian Srs2 homologs have not been identified, and this function may be carried out in mammalian cells by the BRC repeats of BRCA2.12 2.3. Repair Synthesis Once the initial strand exchange is completed, the 3 -end of the invading strand is extended by DNA polymerase past the original site of the DSB (Fig. 2D). The extended end can dissociate and anneal to the other resected end; this single-end invasion mechanism is called synthesis-dependent strand annealing (SDSA) (Fig. 2E, F). Because SDSA does not create full Holliday junctions, crossovers are unlikely and this mechanism may be important for crossover suppression. Alternatively, if both ends invade the donor duplex, and repair synthesis is primed by both ends, a second strand switch, and a ligation, produces a covalently linked double Holliday junction intermediate. The branch migration of Holliday junctions creates or destroys heteroduplex DNA (hDNA) (Fig. 2G, H, I). 2.4. Resolution of Holliday Junctions The double Holliday junction intermediate can be resolved in several ways. Each junction can be resolved by a cleavage/rejoining reaction in one of two senses. If both junctions are resolved in the same sense, a non-crossover product results, while resolution in different senses produces a crossover (Fig. 2J, K, N). The double Holliday junctions may also resolve by converging into one another (not shown); this

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“dissolves” the Holliday junctions but produces linked molecules (hemi-catenane) that can be resolved by a type I topoisomerase.13 The convergence (or “reverse branch migration”) of Holliday junctions prevents crossovers and is thought to be an ATP-dependent process promoted by RecQ helicases such as Sgs1 in yeast and BLM in human cells,10,14 which interact with Top3/Top3α. However, recent data in yeast suggest that Sgs1 suppression of crossovers is ATP-independent,15 suggesting that Holliday junction convergence may involve other factors or occur passively by a “random walk.” 2.5. Resolution of Mismatches When donor and recipient duplexes are not identical, sequence differences in hDNA create mismatches that are substrates for the mismatch repair machinery. Normally, mismatches are repaired using the donor duplex as template, resulting in conversion of the recipient. This bias is probably a consequence of mismatch repair excising DNA from broken ends (in this case, the 3 invading strand) until it reaches the mismatch, followed by resynthesis.16 The mismatches remaining in hDNA after the Holliday junctions are resolved may be repaired by the mismatch repair system (Fig. 2L, M, O, and P), perhaps without the strand bias of the earlier mismatch repair events. Because gene conversion is a result of mismatch repair in hDNA, the length of the converted region (LOH) depends on the extent of hDNA and the direction of mismatch repair with hDNA. 2.6. DSB Repair by Single-Strand Annealing (SSA) Unlike the SDSA and double-Holliday junction models above, SSA is a non-conservative HR process that can repair a DSB between, or within linked, direct repeats (Fig. 3). With this arrangement, end-resection reveals complementary single strands in each repeat, which can anneal to form a single repeat. The other repeat and intervening sequences are lost, hence SSA is a non-conservative, mutagenic HR process. SSA creates 3 -flaps that are removed by the ERCC1/ERCC4 structure specific endonuclease in mammalian cells

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Single-strand anneal model (see text for details).

(Rad1/Rad10 in yeast).17,18 Because SSA requires end-processing to reveal ssDNA in both repeats, SSA is most efficient when repeats are close together.19 SSA can occur between unlinked repeats, leading to translocations, but this is not common because it requires DSBs in both repeats. 3. RESTART OF STALLED OR COLLAPSED REPLICATION FORKS BY HR The faithful transmission of DNA from parent to daughter cells is critical for genome stability. This requires high-fidelity replication of DNA once per cell cycle, and the proper segregation of

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chromosomes to daughter cells. Several mechanisms contribute to accurate genome transmission, including proofreading by DNA polymerases, repair of mismatches missed during proofreading, mitotic checkpoints that monitor chromosome segregation, and replication licensing that ensures a single round of DNA replication per cell cycle. Failure of the latter system leads to “endoreduplication” and polyploidy.20,21 In recent years another major mechanism to ensure replication fidelity has emerged, namely, the restart of blocked replication forks. Mammalian genomes have large amounts of non-coding, repetitive DNA, and a significant fraction of coding sequences are silent in any given cell. Lesions in these silent regions do not affect cellular function, but may stall or block replication forks. Such blocks may cause the DNA polymerase complex to dissociate from the fork, and if a stalled fork does not restart in a timely manner, it may collapse to a DSB and result in cell death. On the other hand, restarting a stalled replication fork can be mutagenic if an error-prone polymerase is recruited to perform translesion synthesis. A variety of models have been proposed to explain how HR processes may be involved in restarting blocked replication forks.3,22–28 Two representative models are shown in Figs. 4 and 5. 3.1. Restart of Stalled Replication Forks A replication fork that encounters a blocking lesion on one strand may continue DNA synthesis of the other strand using the intact strand as a template (Fig. 4A, B). However, this extension is limited due to the coupling between leading- and lagging-strand synthesis and eventually the replication fork stalls. Fork regression, which is analogous to strand invasion/branch migration, produces a 4-way junction akin to a Holliday junction (Fig. 4C, D). This separates the daughter strands from their original templates, and allows the blocked strand to extend past the position of the blocking lesion using the other newly synthesized strand as a template. The regressed fork, often described as a “chicken foot” structure, has been observed by electron microscopy,29 and can reverse direction

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Fig. 4

Restart of stalled replication fork .

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Fig. 5 Restart of collapsed replication fork with a one-ended DSB (see text for details).

to restore the replication fork (Fig. 4G). Alternatively, if the regressed fork cannot be reversed, the 4-way junction can be resolved much like a Holliday junction to restore the normal fork structure (Fig. 4H, K). In either case, the original lesion is bypassed and can be repaired later using the daughter strand as a template. Thus, this mechanism of replication fork restart is formally a lesion bypass process. These mechanisms allow relatively error-free lesion bypass, unlike direct lesion bypass by error-prone translesion DNA polymerases30 (Chapter 10). In HR-defective cells, blocked or stalled replication forks are shunted to the error-prone pathway. Thus, replication fork restart is another way that HR helps in maintaining genome stability.

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3.2. Restart of Collapsed Replication Forks When a replication fork encounters a block, such as a single-strand break (SSB) in a template strand, replication stops at the lesion, but can continue on the undamaged template. This may produce ssDNA regions (daughter strand gaps) that are bound by RPA, triggering checkpoint arrest (Fig. 5A, B). DNA synthesis and ligation restore one full duplex, leaving a single-ended DSB (Fig. 5C). This end can invade the intact duplex to reform the replication fork (Fig. 5D– G). When HR is defective, this pathway is not available, and cells may be forced to use other mechanisms, such as NHEJ, to rejoin the broken end to another molecule. It is possible that synthesis on the other daughter strand can continue beyond the single-strand break using the undamaged parental strand as the template (Fig. 5H). This intermediate can be processed by the damage bypass mechanism shown in Fig. 4B to restart replication. In the model shown in Fig. 5, the SSB is on the leading strand template. When SSBs occur on the lagging strand template, leading strand synthesis can bypass the lesion, and the fork can be restored by a fork regression mechanism, as shown in Fig. 4.

4. REPAIR OF INTERSTRAND CROSSLINKS BY HR Certain genotoxins such as mitomycin C and cisplatin have two reactive groups and can link opposite strands of dsDNA, creating interstrand crosslinks (ICLs). ICLs block both transcription and DNA replication. ICL repair may involve multiple repair pathways, including nucleotide excision repair (NER), HR and translesion synthesis.31 A characteristic feature of HR-defective cells is the marked hypersensitivity to agents that create ICLs. Because such agents are widely used in cancer chemotherapy, understanding the cellular responses to ICL damage, and the roles of HR and other pathways in ICL repair, has significant implications for cancer treatment. Several models for ICL repair have been proposed, and one example is shown in Fig. 6. In this model, ICL removal is initiated by the NER system. Dual incisions on a single strand flanking the

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Fig. 6

Replication independent repair of interstrand crosslinks.

ICL, coupled with a helicase activity, create a single-stranded gap opposite the damaged site. With single-strand damage, such gaps are readily filled by DNA polymerase, but the remaining damage in the template inhibits accurate repair synthesis. The incision reaction does generate a free 3 -end, which, however, can invade a homologous donor region and prime repair synthesis, ultimately creating

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Holliday junctions (Fig. 6D) that can branch migrate and thus allows the remaining damaged strand to be repaired by NER (Fig. 6E). Resolution of the double Holliday junctions separates the donor and recipient duplexes, and the genetic information at the ICL site is recovered from the donor duplex. In this model, repair is independent of replication and has a low risk of generating mutations. In a second model (Fig. 7), ICL repair initiates when a replication fork is blocked by the damage. As in the above, NER nucleases create single-strand incisions flanking the lesion (Fig. 7B). Then, translesion synthesis may re-fill the single-strand gap (Fig. 7C), allowing a second round of NER nuclease cleavage that produces a one-ended DSB (collapsed fork; Fig. 7D, E), which can be restarted as shown in Fig. 5B–G. Alternatively, if the other nascent strand is extended beyond the ICL site, the fork regression pathway (Figs. 7F–K) can bypass the ICL for later repair by BER or NER. Because these pathways involve translesion synthesis, there is a significant risk of mutation. Nevertheless, the cell restarts the fork allowing the cell to survive. The role of translesion synthesis and its coupling with HR in the repair of ICL are supported by recent reports.32,33 5. ENZYMATIC AND REGULATORY PROTEINS INVOLVED IN HR Because of the complexity of HR reactions and the dynamic interactions between HR and DNA metabolic and repair processes, a large number of proteins are involved, including enzymes that catalyze the chemical reactions, and accessory proteins that either regulate the activity of the enzymatic proteins or act at the interface between HR and other cellular processes. The key reactions in HR are the search for homology and strand exchange, catalyzed by the RecA protein in Escherichia coli and its homolog, RAD51 in eukaryotes. DMC1 is related to RAD51 and functions in meiosis. In yeast, accessory proteins or “mediators” include the RAD51 paralogs RAD55 and RAD57 (which form a heterodimer), and RAD54. In the early stages of HR, these proteins assist in RAD51 filament formation, which involves the replacement of RPA with RAD51 on

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Fig. 7

Replication coupled repair of interstrand crosslinks.

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single-stranded DNA tails. In higher eukaryotes there are two paralog complexes, RAD51C-XRCC3, and RAD51B-RAD51C-RAD51DXRCC2. The critical roles of RAD51 and paralogs in HR have been recently reviewed.34,35 Other enzymes, such as DNA helicases, nucleases, polymerases, topoisomerases and DNA ligase, participate in other steps, including the extension of the invading strand, branch migration, the resolution of Holliday junctions, and the ligation of DNA ends. It is clear that these enzymes have the most direct role in HR. However, their activities are tightly regulated so that HR is coordinated with the cell cycle and other repair processes. For example, in mammalian cells RAD51 activity is regulated by several accessory proteins, such as BRCA2, RAD52, RAD54 and perhaps BRCA1. Because HR has critical roles in restarting blocked replication forks, it is essential for the viability of higher eukaryotic cells with their large genomes. Thus, the inactivation of RAD51 is cell-lethal in higher eukaryotes.36 In contrast, the defects in accessory proteins or the partial inactivation of enzymatic HR proteins may not completely eliminate HR reactions; instead they may affect the accuracy or timing of HR reactions, which can result in genome instability and cancer development. In the following sections, we focus on the role of BRCA2 in HR regulation and describe the strategies for identifying other HR regulatory proteins. 5.1. Regulation of RAD51 Activity by BRCA2 In higher eukaryotes, RAD51 activity is regulated by the product of the breast cancer susceptibility gene, BRCA2. BRCA2 was originally implicated as a HR-regulatory protein by its interaction with RAD51,37–42 and subsequent studies confirmed a direct role for BRCA2 and its interaction with RAD51 in HR.12,43–45 Mutations that affect the BRCA2-RAD51 interaction result in genome instability and promote tumorigenesis.42,46–48 Human BRCA2 encodes a large protein of 3418 amino acids, with no significant homology to other human proteins. The overall homology between human and mouse BRCA2 is moderate (∼ 59%),

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but there are five highly conserved regions.49,50 A series of eight BRC repeats encoded by exon 11 are critical for BRCA2 and RAD51 interaction. Although the overall sequence in exon 11 shows only a moderate homology between mouse and human, the BRC repeats are conserved.51 All BRC repeats have been shown to interact with RAD51 but BRCs 1–4, 7 and 8 show the highest avidity.38–41,46 Overexpression of the BRC domains disrupts the normal BRCA2/RAD51 interaction, rendering cells hypersensitive to DNA damage, and leading to the loss of G2/M checkpoint regulation after DNA damage.47,52 Structural analysis further suggested a role for the BRC and RAD51 interaction in facilitating RAD51 filament formation.53,54 In addition, the C-terminal domain of BRCA2 (aa3267-3311 in human, and aa3196-3232 in mouse) coded by exon 27, also interacts with RAD5149,50 and plays a key role in regulating HR (see below). This region of mouse BRCA2 has 72% amino acid identity with human BRCA2. BRCA2 has been proposed to promote RAD51 filament formation by facilitating RPA removal from ssDNA. Proximate to the BRCA2 C-terminus, the longest conserved region is encoded by exons 14–24 (aa2479-3157 in human, aa2400-3075 in mouse).49 The crystal structure of this region revealed three ssDNA binding regions, termed oligonucleotide binding (OB) domains OB1–3. The entire conserved region is called the BRCA2-DNA binding domain (BRCA2-DBD). The OB domains resemble the ssDNA binding fold of RPA and may assist in the removal of RPA from ssDNA (by molecular mimicry) and thus promote RAD51 filament formation.55 Overexpression of individual OB domains impairs HR (M Brenneman, LS de la Pena, K Bystol, J LoPiccolo, K Ortega, J Fan, ZS, and JAN, manuscript submitted). 5.2. Linking HR to other Cellular Processes via RAD51-interacting Proteins Because RAD51 plays a central role in strand exchange during HR, RAD51 interacting proteins may co-ordinate HR with other cellular processes. One such protein is BRCA2, which may have other

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RAD51-independent functions in tumor suppression. For example, the N-terminal 100 aa of BRCA2 encodes a transcriptional regulation domain.56 Deletions in this region have been identified in some BRCA2-associated breast cancers.57 A candidate histone acetylation domain was identified next to the transcription regulation domain that may regulate the transcription activation.58 The BRCA2 transcriptional domain interacts with EMSY, which is implicated in many forms of human cancer.59,60 These studies suggest that the transcription regulation domain is an essential component of BRCA2 tumor suppressor activity. The BRCA2-DBD region interacts with DNA and with several proteins, including DSS1, hBubR1, filamin-A (ABP-280) and BCCIP.55,61–66 DSS1 (Deleted in Split-hand and Split-foot) interacts with BRCA2 aa2472-2957.64 DSS1 has been implicated in HR and BRCA2 stability.55,67–69 The yeast DSS1 homologue (also called SEM1) plays a role in cell growth and differentiation.70 BubR1 interacts with BRCA2 aa2867-3176 and affects BRCA2 phosphorylation status.61 BubR1 and its yeast homologues contain a kinase domain, and participate in mitotic checkpoint control.71 The BRCA2-DBD also interacts with the actin binding protein filamin-A/ABP280, and the recently identified protein BCCIP.62,66 Filamin-A is an actin crosslink protein72 that participates in remodeling the cytoskeleton structure in response to extracellular signals.73 BCCIP, also called Tok1, is an evolutionarily conserved protein that interacts with BRCA2 and the p21/Waf1/Cip1 cell cycle control protein,74 and controls both HR and the cell cycle.62,63,65,74–76 The human BRCA2 exon 27-RAD51 interaction is regulated by the phosphorylation of serine 3291 by a cyclin-dependent kinase that allows the co-ordination of this interaction with the cell cycle.77 BRCA2 has also been shown to function directly in cytokinesis and cell cycle regulation.78–80 Because BRCA2 is involved in transcription regulation, cell cycle regulation and cytokinesis, and interacts with proteins with diverse functions, it is likely that it plays a critical role in linking HR to other cellular processes. Other multifunctional RAD51-associated proteins (such as BRCA1 and p53) are also expected to play a role in linking HR to other cellular processes.

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5.3. Strategies to Identify Proteins that Regulate HR Due to the complexity of HR, a large protein network is expected to be involved in HR reactions and regulation. In the last few years, many new HR proteins have been discovered in mammalian cells, and it is likely that more will be found in the future. The identification of new HR proteins has been facilitated by combining genetic, sequence, protein interaction and functional analyses. In yeast, HR is the major repair mechanism for DSB repair, and HR defects confer severe sensitivity to ionizing radiation. Historically, the genes in the yeast RAD52 epistasis group (RAD51, RAD52, RAD54, RAD55 and RAD57) were identified by analyzing DNA damage-sensitive mutants. Similar genetic approaches applied to mammalian mutants sensitive to DNA damage led to the identification of XRCC2 and XRCC3. Sequence similarities with known yeast HR genes revealed mammalian HR genes, including human RAD52 and RAD51.81,82 HR-regulating proteins in mammalian cells have been identified based on their interactions with known HR proteins, including BRCA2, p53, SUMO-1/UBL1, UBC9/UBE2I and BRCA1.50,83–87 In recent years, nuclear focus formation by HR proteins, determined by indirect immunofluorescence, has been widely used as an indicator of HR function. By analyzing co-localization with RAD51 or other known HR proteins, additional HR regulating proteins can be identified. Haaf et al.88 showed that RAD51 forms nuclear foci in response to DNA damage, and it was later shown that DSB repair proteins form nuclear foci only in damaged areas of the nucleus.89 RAD51, RAD52 and BRCA2 are normally dispersed, but they redistribute to form foci upon DNA damage.46,88,90,91 Disruption of focus formation by dominant negative regulators sensitizes cells to DNA damage.46,47,52,92 Moreover, DNA damage-induced foci of HR proteins are present at ssDNA regions.92 Therefore, the formation of nuclear foci by HR proteins in response to DNA damage provides a convenient, visual assay that can give insight into in vivo function. Thus, proteins found to regulate focus formation of known HR proteins, and proteins that co-localize with known HR proteins, are

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good candidates for regulation of HR. This strategy has facilitated the identification or confirmation of additional HR regulatory proteins. For example, genetic evidence indicated that BRCA1 regulates HR,93 and this was supported by a study showing that BRCA1 affects RAD51 focus formation.94 6. ROLES OF HR IN MITOSIS AND CELL CYCLE CHECKPOINT CONTROL As outlined above, primary HR roles are to repair DNA damage and restart blocked replication forks, and these are critical for replication fidelity and the accurate repair of DNA damage. However, HR defects often result in mitotic abnormalities and alterations in cell cycle progression, suggesting at least an indirect role of HR in these processes. 6.1. HR Regulation of Mitosis The inactivation of several HR proteins has been shown to result in several mitotic abnormalities affecting centrosome duplication, chromosome segregation and G2/M checkpoint activation. These problems may lead to mitotic catastrophe or cell division errors. For example, the inactivation of XRCC3 not only reduces HR efficiency and alters HR outcomes, it also leads to endoreduplication and mitotic errors.95–102 Although the underlying causes of these abnormalities are not fully understood, several scenarios may be considered. First, the delay in the completion of DNA replication as a result of impaired HR in S phase, may result in an excessive level of HR intermediates (such as joint molecules) linking homologous chromosomes or sister chromatids. This may cause chromosome segregation errors in mitosis, such as those observed in XRCC3 defective cells,100 which are consistent with the proposed role for the RAD51C-XRCC3 complex in the late HR stages.95,103 Second, in parallel with the DNA replication cycle is a centrosome duplication cycle.104,105 Centrosome duplication is largely programmed before the start of the S phase, and is normally synchronized with

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the DNA replication cycle.106,107 If replication is delayed due to an excessive level of stalled replication forks , the replication and centrosome cycles may become uncoordinated, leading to centrosome amplification or fragmentation.108 HR regulatory proteins may also help maintain synchrony between the replication and centrosome cycles since centrosome amplification is apparent in BRCA1- and BRCA2-defective cells.109–113

6.2. Interface between HR and Cell Cycle Regulation Cell cycle checkpoints help maintain genome stability by restricting cells with damaged DNA from entering the next cell cycle phase, thus reducing the chance of their permanently altering the genome. For example, when cellular DNA is damaged in G1 phase, the G1/S checkpoint is activated to delay the progression into S phase. This reduces the chance of replication forks encountering DNA damage that leads to replication errors, and thus helps maintain genome stability and prevent cancer.114–118 When replication is not complete, or DNA damage is not fully repaired in S phase, cells become blocked at the G2/M border (G2/M checkpoint). The failure of this response allows cells to enter mitosis prematurely, resulting in a higher risk of mitotic errors, including anueploidy and polyploidy — the hallmarks of highly destabilized genomes common in cancer cells. The interface between HR and cell cycle control is apparent from a variety of studies. For example, the expression and distribution of some HR proteins, including RAD51, are cell cycle-dependent,119,120 HR activity is regulated during the cell cycle, peaking in the S and G2 phases when sister chromatids are available as repair templates,6 and HR defects may activate cell cycle checkpoints. A key example of the tight interface between HR and cell cycle control is demonstrated by the RAD51 and p53 interaction. Both RAD51 and BRCA2 interact with p53.40,84,87 These interactions appear to be important for both p53 inhibition of HR,84,121–124 and for RAD51 and BRCA2 inhibition of p53 transactivation, which is important for cell cycle regulation.40 When there is an excessive level of stalled replication forks , p53 is

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recruited to these sites to stabilize the forks, and this sequesters p53 away from its transactivation sites.125,126 7. HR AND CANCER INTERVENTION Effective cancer intervention includes prevention, early and accurate diagnosis, treatments tailored to patients’ specific needs, and precise prognostic evaluation. A better understanding of DNA repair in general, and HR in particular, may have a strong impact on cancer intervention on at least two fronts. On one hand, HR defects cause genome instability and therefore promote tumorigenesis, and thus clarifying HR roles in genome stabilization may provide opportunities for effective prevention and early diagnosis. On the other hand, HR is a critical aspect of cellular responses to DNA damage, and thus HR proteins and pathways may be useful as tumor markers and as targets for cancer therapy. 7.1. HR and Cancer Etiology Although cancer is fundamentally a state of unregulated cell growth, as many as 5–7 mutations are required to convert a normal cell to a malignant tumor cell through the deregulation of the growth control network. It is extremely unlikely for a single cell to acquire the required mutations unless its genome is destabilized, hence cancer initiation and progression can be considered a disease of genomic instability. HR protects the genome by promoting the relatively accurate repair of spontaneous DSBs and ensuring replication fidelity, and also influences mitotic chromosome segregation and cell cycle regulation.24,27,35,127–130 Defects in many HR genes have been implicated in a variety of human diseases, including cancer.5 There are several ways that HR defects may contribute to cancer etiology. A reduction in HR activity forces cells to use more errorprone pathways to repair DSBs or restart blocked replication forks. This mechanism is illustrated by the cancer predisposition associated with BRCA1 and BRCA2 mutations. BRCA1 and BRCA2 are both involved in HR via interactions with RAD51.35,131–136 Defects

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in these proteins reduce HR efficiency and accuracy resulting in genome instability. Nearly all carriers with germ-line mutations in BRCA1 or BRCA2 eventually develop cancer. Although HR is a fairly accurate DNA repair mechanism, it is not without risk of genetic change. Optimally, HR involves interactions between sister chromatids or between allelic loci on homologous chromosomes, as these pose the lowest risk of genetic alteration to the damaged locus. However, gene conversion between heterozygous alleles results in the localized loss of heterozygosity. This can, for example, replace a wild type tumor suppressor allele with a mutant allele, resulting in the loss of tumor suppressor function. HR between regions with significant sequence differences, so-called “homeologous” recombination is normally suppressed, at least in part by the mismatch repair machinery.137 A breakdown in this suppression increases homeologous recombination, and depending on the configuration of the interacting regions and HR outcome, can lead to gene inactivation (i.e. by conversion of a wild type gene from a pseudogene repair template), deletions, inversions, and translocations. Defects in RAD51-dependent strand exchange can shunt repair toward SSA, increasing the chance of deletions in repeated regions.138 The suppression of HR-mediated deletions by restricting interactions between diverged repeats is critical for the stability of the human genome, which has a large amount of repetitive DNA, including Alu, LINE and SINE elements. Alu elements are ∼ 300 bp in length, share significant homology to each other, are widely dispersed, constitute > 10% of the human genome,139,140 and are a major source of HR-dependent genome instability.141,142 HR between Alu repeats has been identified as the cause of deletions and duplications in the BRCA1 gene.143–146 7.2. HR as a Target in Cancer Treatment As discussed in Sections 2–4, HR has critical roles in DSB and ICL repair, and in restarting blocked and collapsed replication forks. HR defects typically render cells moderately sensitive to ionizing radiation, but highly sensitive to ICL agents such as mitomycin C

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and cisplatin. It has been suggested that cancers with HR defects will be more sensitive to therapeutic DNA damaging agents, especially those that create ICLs. Therefore, HR status may be a valuable marker to guide the selection of therapeutic DNA damaging agents. In a similar way, it may be possible to inhibit HR in tumor cells to increase their sensitivity to ICL agents. This strategy poses risks to the surrounding normal tissue because HR is an essential process in normal cells, and the non-specific reduction of HR may have serious side effects, including cell death or induced genome instability that could lead to secondary tumors. One way to circumvent this problem is to explore the functional redundancy of certain HR proteins. For example, although RAD52 is essential for RAD51-dependent HR in yeast, the inactivation of RAD52 in mammalian cells has little effect on HR or cellular sensitivity to DNA damage.147 However, the inactivation of both RAD52 and XRCC3 causes chromosome breaks and cell lethality, similar to the inactivation of RAD51.148 Therefore, it may be possible to target RAD52 in XRCC3-deficient cancer cells, killing the cancer cells but sparing the wild type cells. Like RAD52 in yeast, BRCA2 promotes RAD51-dependent HR. Yeast lacks BRCA2, and it is possible that BRCA2 has taken over certain RAD52 functions in mammalian cells. Thus, mammalian RAD52 may be redundant with BRCA2, and if this is the case, inhibiting RAD52 may kill BRCA2deficient cells, but not normal cells. In addition to redundant protein functions, there is also some promise in targeting redundant DNA repair pathways. A recent example of this strategy is the specific killing of BRCA2-defective cells, which are HR-deficient, by inhibiting poly(ADP-ribose) polymerase, which acts in base excision repair.149,150 8. CONCLUSIONS AND PERSPECTIVES The roles of HR in DNA repair, DNA replication, genome stabilization and tumor suppression are established in broad outline, but many details remain to be worked out. Addressing the following questions may provide new opportunities for cancer intervention.

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8.1. How Do Cells Select NHEJ or HR to Repair DSBs? DSBs can be repaired by NHEJ and HR, but it is unclear how cells choose which pathway to use. A simple model is that the two pathways compete for DSBs, but it is becoming clear that this pathway choice is regulated. For example, HR activity increases in S and G2 phases, while NHEJ has high activity in G1 phase. DSBs can occur in transcriptionally active or silent genes, or in non-coding sequences, and these contexts may affect the selection of HR or NHEJ. Ku and RAD52 each bind to DNA ends and play roles in NHEJ and HR, respectively, and it has been suggested that affinities to different structures at DNA ends might vary and thereby influence the choice between NHEJ and HR.151 An understanding of the specifics of DSB repair pathway choice may prove valuable in tailoring cancer therapy in various tumor types or tumor grades.

8.2. How Important is Unregulated HR in Tumorigenesis? HR has been characterized as an accurate repair pathway that maintains genome stability, but HR has associated risks of genome rearrangement, particularly when donor selection is not stringent. Clearly, HR levels must be balanced to optimize genome stability. Over-expression of HR proteins has been observed in some tumors,152–156 but the full scope of this problem is not known. Characterizing tumors by gene array or proteomic approaches will help identify novel HR tumor markers, and this information may guide therapy decisions in the future.

8.3. Are HR Regulatory Proteins Good Targets for Cancer Therapy? Defects in HR enzymes and HR regulatory proteins cause cancer, and may also sensitize cells to certain therapeutic modalities. While HR defects may be useful markers to guide DNA damage-based cancer therapy, this is generally true for HR enzymes directly involved in repair. HR regulatory proteins, on the other hand, may have

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other functions, i.e. in cell cycle checkpoint control or other processes related to cell death. Defects in these proteins may reduce HR but increase survival, perhaps with mutagenic consequences. This would be a potent means to cause further genomic instability and potentially enhance tumor progression toward a more malignant state. Therefore, a comprehensive understanding of potential risks and benefits is required when selecting HR regulatory proteins as targets for cancer therapy. 8.4. How Does HR Interface with the Damage Response Network and Other Repair Pathways? HR is tightly co-ordinated with cell cycle control, signal transduction in response to DNA damage, gene expression, cell death, and other cellular functions. HR co-operates, or is redundant with other repair pathways, such as mismatch repair, nucleotide excision repair, and base excision repair. An understanding of these interactions at the molecular level will provide new opportunities for combination therapy by the oncologist. The recent demonstration that BRCA2deficient cells are hypersensitive to drugs that block base excision repair is the latest,149,150 but likely not the last, example of the power of DNA repair pathway analysis in the battle against cancer. ACKNOWLEDGMENTS We apologize to the many authors whose work could not be cited due to space limitation. The homologous recombination-related research in the authors’ labs is supported by National Institute of Health grants ES08353 (ZS), CA115488 (ZS), CA100862 (JN), CA77693 (JN), and CA118357 (JN). References 1. Nickoloff JA. (2002) In: Bertino JR. (ed), Encyclopedia of Cancer, 2nd edn, Elsevier Science (USA), San Diego, Vol. 4, pp. 49–59. 2. Gerton JL, Hawley RS. (2005) Homologous chromosome interactions in meiosis: diversity amidst conservation. Nat Rev Genet 6: 477–487.

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3. Helleday T. (2003) Pathways for mitotic homologous recombination in mammalian cells. Mutat Res 532: 103–115. 4. West SC. (2003) Molecular views of recombination proteins and their control. Nat Rev Mol Cell Biol 4: 435–445. 5. Thompson LH, Schild D. (2002) Recombinational DNA repair and human disease. Mutat Res 509: 49–78. 6. Hinz JM, Yamada NA, Salazar EP, et al. (2005) Influence of doublestrandbreak repair pathways on radiosensitivity throughout the cell cycle in CHO cells. DNA Repair (Amst) 4: 782–792. 7. Johnson RD, Jasin M. (2000) Sister chromatid gene conversion is a prominent double-strand break repair pathway in mammalian cells. EMBO J 19: 3398–3407. 8. de Wind N, Dekker M, Berns A, et al. (1995) Inactivation of the mouse Msh2 gene results in mismatch repair deficiency, methylation tolerance, hyper-recombination and predisposition to cancer. Cell 82: 321–330. 9. Richardson C, Moynahan ME, Jasin M. (1998) Double-strand break repair by interchromosomal recombination: suppression of chromosomal translocations. Genes Dev 12: 3831–3842. 10. Ira G, Malkova A, Liberi G, et al. (2003) Srs2 and Sgs1-Top3 suppress crossovers during double-strand break repair in yeast. Cell 115: 401–411. 11. Krogh BO, Symington LS. (2004) Recombination proteins in yeast. Annu Rev Genet 38: 233–271. 12. Davies AA, Masson JY, McLlwraith MJ, et al. (2001) Role of BRCA2 in control of the RAD51 recombination and DNA repair protein. Mol. Cell 7: 273–282. 13. Wu L, Hickson ID. (2002) The Bloom’s syndrome helicase stimulates the activity of human topoisomerase IIIalpha. Nucl Acids Res 30: 4823–4829. 14. Karow JK, Constantinou A, Li JL, et al. (2000) The Bloom’s syndrome gene product promotes branch migration of Holliday junctions. Proc Natl Acad Sci USA 97: 6504–6508. 15. Lo Y-C, Paffett KS, Amit O, et al. (2006) Sgs1 regulates gene conversion tract lengths and crossovers independently of its helicase activity. Mol Cell Biol 26: 4086–4094 (in press). 16. Weng YS, Nickoloff JA. (1998) Evidence for independent mismatch repair processing on opposite sides of a double-strand break in Saccharomyces cerevisiae. Genetics 148: 59–70. 17. Brookman KW, Lamerdin JE, Thelen MP, et al. (1996) ERCC4 (XPF) encodes a human nucleotide excision-repair protein with eukaryotic recombination homologs. Mol Cell Biol 16: 6553–6562.

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18. Ivanov EL, Haber JE. (1995) RAD1 and RAD10, but not other excisionrepair genes, are required for double-strand break-induced recombination in Saccharomyces cerevisiae. Mol Cell Biol 15: 2245–2251. 19. Schildkraut E, Miller CA, Nickoloff JA. (2005) Gene conversion and deletion frequencies during double-strand break repair in human cells are controlled by the distance between direct repeats. Nucl Acids Res 33: 1574–1580. 20. Grafi G. (1998) Cell cycle regulation of DNA replication: the endoreduplication perspective. Exp Cell Res 244: 372–378. 21. Blow JJ, Dutta A. (2005) Preventing re-replication of chromosomal DNA. Nat Rev Mol Cell Biol 6: 476–486. 22. Carr AM. (2002) Checking that replication breakdown is not terminal. Science 297: 557–558. 23. Michel B, Grompone G, Flores MJ, Bidnenko V. (2004) Multiple pathways process stalled replication forks. Proc Natl Acad Sci USA 101: 12783–12788. 24. Michel B, Flores MJ, Viguera E, et al. (2001) Rescue of arrested replication forks by homologous recombination. Proc Natl Acad Sci USA 98: 8181–8188. 25. Sutton MD, Walker GC. (2001) Managing DNA polymerases: coordinating DNA replication, DNA repair, and DNA recombination. Proc Natl Acad Sci USA 98: 8342–8349. 26. Thompson LH, Hinz JM, Yamada NA, Jones NJ. (2005) How Fanconi anemia proteins promote the four Rs: replication, recombination, repair, and recovery. Environ Mol Mutagen 45: 128–142. 27. Kuzminov A. (2001) DNA replication meets genetic exchange: chromosomal damage and its repair by homologous recombination. Proc Natl Acad Sci USA 98: 8461–8468. 28. Saleh-Gohari N, Bryant HE, Schultz N, et al. (2005) Spontaneous homologous recombination is induced by collapsed replication forks that are caused by endogenous DNA single-strand breaks. Mol Cell Biol, 25: 7158–7169. 29. Sogo JM, Lopes M, Foiani M. (2002) Fork reversal and ssDNA accumulation at stalled replication forks owing to checkpoint defects. Science 297: 599–602. 30. Friedberg EC. (2005) Suffering in silence: the tolerance of DNA damage. Nat Rev Mol Cell Biol 6: 943–953. 31. Dronkert ML, Kanaar R. (2001) Repair of DNA interstrand crosslinks. Mutat Res DNA Repair 486: 217–247. 32. Albertella MR, Green CM, Lehmann AR, O’Connor MJ. (2005) A role for polymerase eta in the cellular tolerance to cisplatin-induced damage. Cancer Res 65: 9799–9806.

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33. McLlwraith MJ, Vaisman A, Liu Y, et al. (2005) Human DNA polymerase eta promotes DNA synthesis from strand invasion intermediates of homologous recombination. Mol Cell 20: 783–792. 34. Kawabata M, Kawabata T, Nishibori M. (2005) Role of recA/RAD51 family proteins in mammals. Acta Med Okayama 59: 1–9. 35. Thompson LH, Schild D. (2001) Homologous recombinational repair of DNA ensures mammalian chromosome stability. Mutat Res 477: 131–153. 36. Sonoda E, Sasaki MS, Buerstedde JM, et al. (1998) RAD51-deficient vertebrate cells accumulate chromosomal breaks prior to cell death. EMBO J 17: 598–608. 37. Chen J, Silver DP, Walpita D, et al. (1998) Stable interaction between the products of the BRCA1 and BRCA2 tumor suppressor genes in mitotic and meiotic cells. Mol Cell 2: 317–328. 38. Chen PL, Chen CF, Chen Y, et al. (1998) The BRC repeats in BRCA2 are critical for RAD51 binding and resistance to methyl methanesulfonate treatment. Proc Natl Acad Sci USA 95: 5287–5292. 39. Katagiri T, Saito H, Shinohara A, et al. (1998) Multiple possible sites of BRCA2 interacting with DNA repair protein RAD51. Genes Chromosomes Cancer 21: 217–222. 40. Marmorstein LY, Ouchi T, Aaronson SA. (1998) The BRCA2 gene product functionally interacts with p53 and RAD51. Proc Natl Acad Sci USA 95: 13869–13874. 41. Wong AK, Pero R, Ormonde PA, et al. (1997) RAD51 interacts with the evolutionarily conserved BRC motifs in the human breast cancer susceptibility gene BRCA2. J Biol Chem 272: 31941–31944. 42. Yuan SS, Lee SY, Chen G, et al. (1999) BRCA2 is required for ionizing radiation-induced assembly of RAD51 complex in vivo. Cancer Res 59: 3547–3551. 43. Larminat F, Germanier M, Papouli E, Defais M. (2002) Deficiency in BRCA2 leads to increase in non-conservative homologous recombination. Oncogene 21: 5188–5192. 44. Moynahan ME, Pierce AJ, Jasin M. (2001) BRCA2 is required for homology-directed repair of chromosomal breaks. Mol Cell 7: 263–272. 45. Xia F, Taghian DG, DeFrank JS, et al. (2001) Deficiency of human BRCA2 leads to impaired homologous recombination but maintains normal non-homologous end joining. Proc Natl Acad Sci USA 98: 8644–8649. 46. Chen G, Yuan SS, Liu W, et al. (1999) Radiation-induced assembly of RAD51 and RAD52 recombination complex requires ATM and c-Abl. J Biol Chem 274: 12748–12752.

FA

September 14, 2006 16:1

WSPC/SPI-B414: DNA Repair, Genetic Instability & Cancer

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47. Chen JJ, Silver D, Cantor S, et al. (1999) BRCA1, BRCA2, and RAD51 operate in a common DNA damage response pathway. Cancer Res 59: 1752s–1756s. 48. Yuan R, Fan S, Wang JA, et al. (1999) Co-ordinate alterations in the expression of BRCA1, BRCA2, p300, and RAD51 in response to genotoxic and other stresses in human prostate cancer cells. Prostate 40: 37–49. 49. Sharan SK, Bradley A. (1997) Murine BRCA2: sequence, map position, and expression pattern. Genomics 40: 234–241. 50. Sharan SK, Morimatsu M, Albrecht U, et al. (1997) Embryonic lethality and radiation hypersensitivity mediated by RAD51 in mice lacking BRCA2. Nature 386: 804–810. 51. Bignell G, Micklem G, Stratton MR, et al. (1997) The BRC repeats are conserved in mammalian BRCA2 proteins. Hum Mol Genet 6: 53–58. 52. Chen CF, Chen PL, Zhong Q, et al. (1999) Expression of BRC repeats in breast cancer cells disrupts the BRCA2-RAD51 complex and leads to radiation hypersensitivity and loss of G(2)/M checkpoint control. J Biol Chem 274: 32931–32935. 53. Galkin VE, Esashi F, Yu X, et al. (2005) BRCA2 BRC motifs bind RAD51DNA filaments. Proc Natl Acad Sci USA 102: 8537–8542. 54. Pellegrini L, Yu DS, Lo T, et al. (2002) Insights into DNA recombination from the structure of a RAD51-BRCA2 complex. Nature 420: 287–293. 55. Yang H, Jeffrey PD, Miller J, et al. (2002) BRCA2 function in DNA binding and recombination from a BRCA2-DSS1-ssDNA structure. Science 297: 1837–1848. 56. Milner J, Ponder B, Hughes-Davies L, et al. (1997) Transcriptional activation functions in BRCA2. Nature 386: 772–773. 57. Nordling M, Karlsson P, Wahlstrom J, et al. (1998) A large deletion disrupts the exon 3 transcription activation domain of the BRCA2 gene in a breast/ovarian cancer family. Cancer Res 58: 1372–1375. 58. Siddique H, Zou JP, Rao VN, Reddy ES. (1998) The BRCA2 is a histone acetyltransferase. Oncogene 16: 2283–2285. 59. Haber DA. (2003) The BRCA2-EMSY connection: implications for breast and ovarian tumorigenesis. Cell 115: 507–508. 60. Hughes-Davies L, Huntsman D, Ruas M, et al. (2003) EMSY links the BRCA2 pathway to sporadic breast and ovarian cancer. Cell 115: 523–535. 61. Futamura M, Arakawa H, Matsuda K, et al. (2000) Potential role of BRCA2 in a mitotic checkpoint after phosphorylation by hBUBR1. Cancer Res 60: 1531–1535. 62. Liu J, Yuan Y, Huan J, Shen Z. (2001) Inhibition of breast and brain cancer cell growth by BCCIPalpha, an evolutionarily conserved nuclear protein that interacts with BRCA2. Oncogene 20: 336–345.

FA

September 14, 2006 16:1

150

WSPC/SPI-B414: DNA Repair, Genetic Instability & Cancer

ch05

Zhiyuan Shen and Jac A. Nickoloff

63. Lu H, Guo X, Meng X, et al. (2005) The BRCA2-interacting protein BCCIP functions in RAD51 and BRCA2 focus formation and homologous recombinational repair. Mol Cell Biol 25: 1949–1957. 64. Marston NJ, Richards WJ, Hughes D, et al. (1999) Interaction between the product of the breast cancer susceptibility gene BRCA2 and DSS1, a protein functionally conserved from yeast to mammals. Mol Cell Biol 19: 4633–4642. 65. Meng X, Yuan Y, Maestas A, Shen Z. (2004) Recovery from DNA damage-induced G2 arrest requires actin-binding protein filamina/actin-binding protein 280. J Biol Chem 279: 6098–6105. 66. Yuan Y, Shen Z. (2001) Interaction with BRCA2 suggests a role for filamin-1 (hsFLNa) in DNA damage response. J Biol Chem 276: 48318–48324. 67. Gudmundsdottir K, Lord CJ, Witt E, et al. (2004) DSS1 is required for RAD51 focus formation and genomic stability in mammalian cells. EMBO Rep 5: 989-993. 68. Kojic M, Yang H, Kostrub CF, et al. (2003) The BRCA2-interacting protein DSS1 is vital for DNA repair, recombination, and genome stability in Ustilago maydis. Mol Cell 12: 1043–1049. 69. Li J, Zou C, Bai Y, et al. (2005) DSS1 is required for the stability of BRCA2. Oncogene 25: 1186–1194. 70. Reinman M, Jantti J, Alfthan K, et al. (2003) Functional inactivation of the conserved Sem1p in yeast by intrabodies. Yeast 20: 1071–1084. 71. Harris L, Davenport J, Neale G, Goorha R. (2005) The mitotic checkpoint gene BubR1 has two distinct functions in mitosis. Exp Cell Res 308: 85–100. 72. Gorlin JB, Yamin R, Egan S, et al. (1990) Human endothelial actinbinding protein (ABP-280, non-muscle filamin): a molecular leaf spring. J Cell Biol 111: 1089–1105. 73. Stossel TP, Condeelis J, Cooley L, et al. (2001) Filamins as integrators of cell mechanics and signalling. Nat Rev Mol Cell Biol 2: 138–145. 74. Ono T, Kitaura H, Ugai H, et al. (2000) TOK-1, a novel p21Cip1-binding protein that co-operatively enhances p21-dependent inhibitory activity toward CDK2 kinase. J Biol Chem 275: 31145–31154. 75. Meng X, Liu J, Shen Z. (2004) Inhibition of G1 to S cell cycle progression by BCCIP beta. Cell Cycle 3: 343–348. 76. Meng X, Lu H, Shen Z. (2004) BCCIP functions through p53 to regulate the expression of p21Waf1/Cip1. Cell Cycle 3: 1457–1462. 77. Esashi F, Christ N, Gannon J, et al. (2005) CDK-dependent phosphorylation of BRCA2 as a regulatory mechanism for recombinational repair. Nature 434: 598–604.

FA

September 14, 2006 16:1

WSPC/SPI-B414: DNA Repair, Genetic Instability & Cancer

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ch05

151

78. Daniels MJ, Wang Y, Lee M, Venkitaraman AR. (2004) Abnormal cytokinesis in cells deficient in the breast cancer susceptibility protein BRCA2. Science 306: 876–879. 79. Kerr P, Ashworth A. (2001) New complexities for BRCA1 and BRCA2. Curr Biol 11: R668–676. 80. Marmorstein LY, Kinev AV, Chan GK, et al. (2001) A human BRCA2 complex containing a structural DNA binding component influences cell cycle progression. Cell 104: 247–257. 81. Shen Z, Denison K, Lobb R, et al. (1995) The human and mouse homologs of the yeast RAD52 gene: cDNA cloning, sequence analysis, assignment to human chromosome 12p12.2-p13, and mRNA expression in mouse tissues. Genomics 25: 199–206. 82. Shinohara A, Ogawa H, Matsuda Y, et al. (1993) Cloning of human, mouse and fission yeast recombination genes homologous to RAD51 and recA. Nat Genet 4: 239–243. 83. Kovalenko OV, Plug AW, Haaf T, et al. (1996) Mammalian ubiquitinconjugating enzyme Ubc9 interacts with RAD51 recombination protein and localizes in synaptonemal complexes. Proc Natl Acad Sci USA 93: 2958–2963. 84. Linke SP, Sengupta S, Khabie N, et al. (2003) p53 interacts with hRAD51 and hRAD54, and directly modulates homologous recombination. Cancer Res 63: 2596–2605. 85. Shen Z, Pardington-Purtymun PE, Comeaux JC, et al. (1996) UBL1, a human ubiquitin-like protein associating with human RAD51/RAD52 proteins. Genomics 36: 271–279. 86. Shen Z, Pardington-Purtymun PE, Comeaux JC, et al. (1996) Associations of UBE2I with RAD52, UBL1, p53, and RAD51 proteins in a yeast two-hybrid system. Genomics 37: 183–186. 87. Sturzbecher HW, Donzelmann B, Henning W, et al. (1996) p53 is linked directly to homologous recombination processes via RAD51/RecA protein interaction. EMBO J 15: 1992–2002. 88. Haaf T, Golub EI, Reddy G, et al. (1995) Nuclear foci of mammalian RAD51 recombination protein in somatic cells after DNA damage and its localization in synaptonemal complexes. Proc Natl Acad Sci USA 92: 2298–2302. 89. Nelms BE, Maser RS, MacKay JF, et al. (1998) In situ visualization of DNA double-strand break repair in human fibroblasts. Science 280: 590–592. 90. Liu Y, Li M, Lee EY, Maizels N. (1999) Localization and dynamic relocalization of mammalian RAD52 during the cell cycle and in response to DNA damage. Curr Biol 9: 975–978.

FA

September 14, 2006 16:1

152

WSPC/SPI-B414: DNA Repair, Genetic Instability & Cancer

ch05

Zhiyuan Shen and Jac A. Nickoloff

91. Liu Y, Maizels N. (2000) Co-ordinated response of mammalian RAD51 and RAD52 to DNA damage. EMBO Rep 1: 85–90. 92. Raderschall E, Golub EI, Haaf T. (1999) Nuclear foci of mammalian recombination proteins are located at single-stranded DNA regions formed after DNA damage. Proc Natl Acad Sci USA 96: 1921–1926. 93. Moynahan ME, Chiu JW, Koller BH, Jasin M. (1999) BRCA1 controls homology-directed DNA repair. Mol Cell 4: 511–518. 94. Bhattacharyya A, Ear US, Koller BH, et al. (2000) The breast cancer susceptibility gene BRCA1 is required for subnuclear assembly of RAD51 and survival following treatment with the DNA crosslinking agent cisplatin. J Biol Chem 275: 23899–23903. 95. Brenneman MA, Wagener BM, Miller CA, et al. (2002) XRCC3 controls the fidelity of homologous recombination: roles for XRCC3 in late stages of recombination. Mol Cell 10: 387–395. 96. Brenneman MA, Weiss AE, Nickoloff JA, Chen DJ. (2000) XRCC3 is required for efficient repair of chromosome breaks by homologous recombination. Mutat Res 459: 89–97. 97. Cui X, Brenneman M, Meyne J, et al. (1999) The XRCC2 and XRCC3 repair genes are required for chromosome stability in mammalian cells. Mutat Res 434: 75–88. 98. Daboussi F, Thacker J, Lopez BS. (2005) Genetic interactions between RAD51 and its paralogues for centrosome fragmentation and ploidy control, independently of the sensitivity to genotoxic stresses. Oncogene 24: 3691–3696. 99. Griffin CS. (2002) Aneuploidy, centrosome activity and chromosome instability in cells deficient in homologous recombination repair. Mutat Res 504: 149–155. 100. Griffin CS, Simpson PJ, Wilson CR, Thacker J. (2000) Mammalian recombination-repair genes XRCC2 and XRCC3 promote correct chromosome segregation. Nat Cell Biol 2: 757–761. 101. Henry-Mowatt J, Jackson D, Masson JY, et al. (2003) XRCC3 and RAD51 modulate replication fork progression on damaged vertebrate chromosomes. Mol Cell 11: 1109–1117. 102. Yamada NA, Hinz JM, Kopf VL, et al. (2004) XRCC3 ATPase activity is required for normal XRCC3-RAD51C complex dynamics and homologous recombination. J Biol Chem 279: 23250–23254. 103. Liu YL, Masson JY, Shah R, et al. (2004) RAD51C is required for Holliday junction processing in mammalian cells. Science 303: 243–246. 104. Doxsey S, McCollum D, Theurkauf W. (2005) Centrosomes in cellular regulation. Annu Rev Cell Dev Biol 21: 411–434. 105. Doxsey S, Zimmerman W, Mikule K. (2005) Centrosome control of the cell cycle. Trends Cell Biol 15: 303–311.

FA

September 14, 2006 16:1

WSPC/SPI-B414: DNA Repair, Genetic Instability & Cancer

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106. Arlot-Bonnemains Y, Prigent C. (2002) Cell cycle. A trigger for centrosome duplication. Science 295: 455–456. 107. Balczon R. (2000) Centrosome replication in somatic cells: the significance of G1 phase. Curr Top Dev Biol 49: 251–266. 108. Hut HM, Lemstra W, Blaauw EH, et al. (2003) Centrosomes split in the presence of impaired DNA integrity during mitosis. Mol Biol Cell 14: 1993–2004. 109. Deng CX. (2002) Roles of BRCA1 in centrosome duplication. Oncogene 21: 6222–6227. 110. Hsu LC, White RL. (1998) BRCA1 is associated with the centrosome during mitosis. Proc Natl Acad Sci USA 95: 12983–12988. 111. Hut HM, Rembacz KP, van Waarde MA, et al. (2005) Dysfunctional BRCA1 is only indirectly linked to multiple centrosomes. Oncogene 24: 7619–7623. 112. Tutt A, Gabriel A, Bertwistle D, et al. (1999) Absence of BRCA2 causes genome instability by chromosome breakage and loss associated with centrosome amplification. Curr Biol 9: 1107–1110. 113. Xu X, Weaver Z, Linke SP, et al. (1999) Centrosome amplification and a defective G2-M cell cycle checkpoint induce genetic instability in BRCA1 exon 11 isoform-deficient cells. Mol Cell 3: 389–395. 114. Almasan A, Linke SP, Paulson TG, et al. (1995) Genetic instability as a consequence of inappropriate entry into and progression through S-phase. Cancer Metastasis Rev 14: 59–73. 115. Bartek J, Lukas J. (2001) Mammalian G1- and S-phase checkpoints in response to DNA damage. Curr Opin Cell Biol 13: 738–747. 116. Elledge SJ. (1996) Cell cycle checkpoints: preventing an identity crisis. Science 274: 1664–1672. 117. Nojima H. (1997) Cell cycle checkpoints, chromosome stability and the progression of cancer. Hum Cell 10: 221–230. 118. Zhou BB, Elledge SJ. (2000) The DNA damage response: putting checkpoints in perspective. Nature 408: 433–439. 119. Chen FQ, Nastasi A, Shen ZY, et al. (1997) Cell cycle-dependent protein expression of mammalian homologs of yeast DNA double-strand break repair genes RAD51 and RAD52. Mutat Res 384: 205–211. 120. Yamamoto A, Taki T, Yagi H, et al. (1996) Cell cycle-dependent expression of the mouse RAD51 gene in proliferating cells. Mol Gen Genet 251: 1–12. 121. Mekeel KL, Tang W, Kachnic LA, et al. (1997) Inactivation of p53 results in high rates of homologous recombination. Oncogene 14: 1847–1857. 122. Romanova LY, Willers H, Blagosklonny MV, Powell SN. (2004) The interaction of p53 with replication protein A mediates suppression of homologous recombination. Oncogene 23: 9025–9033 .

FA

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123. Yoon D, Wang Y, Stapleford K, et al. (2004) p53 inhibits strand exchange and replication fork regression promoted by human RAD51. J Mol Biol 336: 639–654. 124. Yun S, Lie ACC, Porter AC. (2004) Discriminatory suppression of homologous recombination by p53. Nucl Acids Res 32: 6479–6489. 125. Gottifredi V, Shieh S, Taya Y, Prives C. (2001) p53 accumulates but is functionally impaired when DNA synthesis is blocked. Proc Natl Acad Sci USA 98: 1036–1041. 126. Takimoto R, El-Deiry WS. (2001) DNA replication blockade impairs p53-transactivation. Proc Natl Acad Sci USA 98: 781–783. 127. Courcelle J, Hanawalt PC (2001) Participation of recombination proteins in the rescue of arrested replication forks in UV-irradiated Escherichia coli need not involve recombination. Proc Natl Acad Sci USA 98: 8196–8202. 128. Kuzminov A. (1995) Collapse and repair of replication forks in Escherichia coli. Mol Microbiol 16: 373–384. 129. Saintigny Y, Delacote F, Vares G, et al. (2001) Characterization of homologous recombination induced by replication inhibition in mammalian cells. EMBO J 20: 3861–3870. 130. Thompson LH, Schild D. (1999) The contribution of homologous recombination in preserving genome integrity in mammalian cells. Biochimie 81: 87–105. 131. Jasin M. (2002) Homologous repair of DNA damage and tumorigenesis: the BRCA connection. Oncogene 21: 8981–8993. 132. Johnson RD, Jasin M. (2001) Double-strand-break-induced homologous recombination in mammalian cells. Biochem Soc Trans 29: 196–201. 133. Orelli BJ, Bishop DK. (2001) BRCA2 and homologous recombination. Breast Cancer Res 3: 294–298. 134. Shivji MK, Venkitaraman AR. (2004) DNA recombination, chromosomal stability and carcinogenesis: insights into the role of BRCA2. DNA Repair (Amst) 3: 835–843. 135. Venkitaraman AR. (2002) Cancer susceptibility and the functions of BRCA1 and BRCA2. Cell 108: 171–182. 136. Venkitaraman AR. (2001) Functions of BRCA1 and BRCA2 in the biological response to DNA damage. J Cell Sci 114: 3591–3598. 137. Radman M. (1989) Mismatch repair and the fidelity of genetic recombination. Genome 31: 68–73. 138. Tutt A, Bertwistle D, Valentine J, et al. (2001) Mutation in BRCA2 stimulates error-prone homology-directed repair of DNA double-strand breaks occurring between repeated sequences. EMBO J 20: 4704–4716.

FA

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139. Batzer MA, Deininger PL. (2002) Alu repeats and human genomic diversity. Nat Rev Genet 3: 370–379. 140. Schmid CW. (1996) Alu: structure, origin, evolution, significance and function of one-tenth of human DNA. Prog Nucl Acid Res Mol Biol 53: 283–319. 141. Deininger PL, Batzer MA. (1999) Alu repeats and human disease. Mol Genet Metab 67: 183–193. 142. Kolomietz E, Meyn MS, Pandita A, Squire JA. (2002) The role of Alu repeat clusters as mediators of recurrent chromosomal aberrations in tumors. Genes Chromosomes Cancer 35: 97–112. 143. Montagna M, Santacatterina M, Torri A, et al. (1999) Identification of a 3-kb Alu-mediated BRCA1 gene rearrangement in two breast/ovarian cancer families. Oncogene 18: 4160–4165. 144. Puget N, Sinilnikova OM, Stoppa-Lyonnet D, et al. (1999) An Alumediated 6-kb duplication in the BRCA1 gene: a new founder mutation? Am J Hum Genet 64: 300–302. 145. Rohlfs EM, Puget N, Graham ML, et al. (2000) An Alu-mediated 7.1-kb deletion of BRCA1 exons 8 and 9 in breast and ovarian cancer families that results in alternative splicing of exon 10. Genes Chromosomes Cancer 28: 300–307. 146. Swensen J, Hoffman M, Skolnick MH, Neuhausen SL. (1997) Identification of a 14-kb deletion involving the promoter region of BRCA1 in a breast cancer family. Hum Mol Genet 6: 1513–1517. 147. Yamaguchi-Iwai Y, Sonoda E, Buerstedde JM, et al. (1998) Homologous recombination, but not DNA repair, is reduced in vertebrate cells deficient in RAD52. Mol Cell Biol 18: 6430–6435. 148. Fujimori A, Tachiiri S, Sonoda E, et al. (2001) RAD52 partially substitutes for the RAD51 paralog XRCC3 in maintaining chromosomal integrity in vertebrate cells. EMBO J 20: 5513–5520. 149. Bryant HE, Schultz N, Thomas HD, et al. (2005) Specific killing of BRCA2-deficient tumors with inhibitors of poly(ADP-ribose) polymerase. Nature 434: 913–917. 150. Farmer H, McCabe N, Lord CJ, et al. (2005) Targeting the DNA repair defect in BRCA mutant cells as a therapeutic strategy. Nature 434: 917–921. 151. Ristic D, Modesti M, Kanaar R, Wyman C. (2003) RAD52 and Ku bind to different DNA structures produced early in double-strand break repair. Nucl Acids Res 31: 5229–5237. 152. Barlund M, Monni O, Kononen J, et al. (2000) Multiple genes at 17q23 undergo amplification and over-expression in breast cancer. Cancer Res 60: 5340–5344.

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153. Han HY, Bearss DJ, Browne LW, et al. (2002) Identification of differentially expressed genes in pancreatic cancer cells using cDNA microarray. Cancer Res 62: 2890–2896. 154. Maacke H, Jost K, Opitz S, et al. (2000) DNA repair and recombination factor RAD51 is over-expressed in human pancreatic adenocarcinoma. Oncogene 19: 2791–2795. 155. Maacke H, Opitz S, Jost K, et al. (2000) Over-expression of wild-type RAD51 correlates with histological grading of invasive ductal breast cancer. Int J Cancer 88: 907–913. 156. Wu GJ, Sinclair CS, Paape J, et al. (2000) 17q23 amplifications in breast cancer involve the PAT1, RAD51C, PS6K, and SIGMA1B genes. Cancer Res 60: 5371–5375.

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CHAPTER 6

Role of Non-Homologous End Joining in the Repair of DNA Double-Strand Breaks Sandeep Burma, Benjamin Chen and David J. Chen∗

ABSTRACT Of the various types of DNA damage that can occur within the mammalian cell nucleus, the DNA double-strand break (DSB) is perhaps the most dangerous. DSBs are induced by ionizing radiation, chemotherapeutic drugs, as well as by the byproducts of cellular metabolism. Understanding exactly how a mammalian cell responds to and repairs a DSB is very important because, on one hand, DSBs cause cancer while, on the other hand, DSBs are induced by chemotherapeutic agents to treat the disease. Of the two main mechanisms by which cells can repair DSBs, NHEJ (non-homologous end joining) and HR (homologous repair), NHEJ is the predominant repair pathway in mammalian cells. In this chapter we describe the main steps in the NHEJ pathway of repair highlighting major recent discoveries, and also provide a perspective on the link between defective NHEJ and human disease.

1. INTRODUCTION The integrity of the human genome is constantly threatened by internal as well as external factors with the propensity to cause DNA ∗ Corresponding

author. Professor and Director, Division of Molecular Radiation Biology, Department of Radiation Oncology, University of Texas Southwestern Medical Center, Dallas, TX 75390-9187. Tel.: 214648-5597; Fax: 214-648-5995; E-mail: [email protected] 157

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damage. Of the various types of DNA damage that can occur within the mammalian cell nucleus, the DNA double-strand break (DSB) is perhaps the most dangerous. DSBs are induced by ionizing radiation (X-rays or γ-rays) as well as by radiomimetic drugs used for chemotherapy. Moreover, the byproducts of cellular metabolism (reactive oxygen species or ROS) can also induce these breaks. In addition, programmed DSBs are generated during regulated cellular processes such as V(D)J recombination. Estimates put the number of endogenous DSBs anywhere between 10–100 per nucleus per day. It is, therefore, very important that the cells have in place a machinery that is poised to respond promptly to these breaks. The importance of such a repair mechanism is underscored by the fact that defects in the proteins responding to DSBs result in cancer in human patients as well as in mouse models. In this chapter we will describe the predominant DSB repair pathway in mammalian cells, NHEJ (nonhomologous end joining), with the objective of providing an up-todate understanding of this pathway and its implications in human disease.

1.1. DNA Double-Strand Breaks and Cancer Understanding exactly how a mammalian cell responds to and repairs a DSB is very important from the standpoint of cancer biology because on the one hand, DSBs cause cancer, while on the other hand, DSBs are also induced by chemotherapeutic agents to treat the disease.1 A direct link between DSBs and cancer has been surmised by researchers based upon the fact that many cancer-predisposition syndromes in humans are caused by mutations in DSB-responsive genes; the same link is also observed in mouse models knocked out for these genes.2,3 The link between DSBs and cancer was concretized by seminal findings from two independent groups published in 2005.4,5 In the simplest terms, the researchers observed the induction of DSBs in the early stages of tumorigenesis and the concomitant activation of damage-responsive proteins which led, in turn, to apoptosis (which

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eliminated these pre-cancerous cells). Interestingly, in progressive stages of cancer, while the DSBs persisted, the proteins responding to the breaks (such as p53) became mutated, thereby lowering the barriers (such as apoptosis or senescence) to full blown carcinogenesis. 1.2. How Do DNA Double-Strand Breaks Arise? DSBs are typically induced by intrinsic sources such as the byproducts of cellular metabolism or by extrinsic sources such as x-rays or γ-rays. Both stimuli ultimately result in the generation of free oxygen radicals which can break the phosphodiester bonds in the DNA backbone; two such breaks on opposite strands of DNA, when present close enough to each other, result in a DSB.6 Other agents that perturb the DNA structure or inhibit processes involving DNA such as replication can also generate DSBs in a secondary manner. Finally, defined DSBs are also generated (and then rejoined) in a programmed manner to facilitate normal cellular processes such as V(D)J recombination which is necessary for generating antibody and T cell receptor diversity.7

2. PATHWAYS TO DSB REPAIR The two main mechanisms by which cells can repair DSBs are NHEJ (non-homologous end joining)8 and HR (homologous repair). During NHEJ, the two broken ends of DNA are simply pieced together, sometimes after limited processing of the DNA ends, resulting in quick, but error-prone, repair. HR is a more accurate method of repair as information is copied from an intact homologous DNA duplex; however, as HR requires the presence of an intact sister chromatid, this method of repair can only operate in the S/G2 phases of the cell cycle in mammalian cells.9 While HR appears to be the predominant mechanism of repair in yeast, NHEJ may be the main pathway of repair in mammalian cells, especially in the G1 phase of the cell cycle.

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2.1. The Non-Homologous End Joining (NHEJ) Repair Pathway NHEJ is a very simple albeit somewhat erroneous repair pathway that, at the very least, requires the DNA-end binding Ku complex (which initially recognizes the DNA break), a protein kinase DNA-PKcs (which signals the presence of a break and activates repair proteins at the break), potential DNA-end processing enzymes (which make the DNA ends compatible for ligation), and the XRCC4-Ligase IV complex (which re-ligates the broken DNA ends). A detailed description of the individual steps in NHEJ and the proteins involved follows. These steps are outlined in Fig. 1.

Ionizing radiation

I. Binding of Ku to DNA ends

II. Recruitment of DNA-PKcs

DNA-PKcs and ATM-mediated phosphorylation of DNA-PKcs

III. Recruitment of end-processing enzymes: Wrn, Artemis, Tdp1, PNK, etc.

DNA-PKcs-mediated phosphorylation of repair proteins

IV. Recruitment of Lig4/XRCC4/XLF complex

V. DNA end ligation

Dissociation of repair complex

Fig. 1

Steps in the NHEJ pathway for the repair of DSBs.

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2.2. The DNA-Dependent Protein Kinase The DNA-dependent protein kinase (DNA-PK) is a key player in the NHEJ pathway of DSB repair, and has additional functions in mammalian cells including telomere maintenance and the induction of apoptosis.10 DNA-PK consists of an approximately 470-kDa catalytic subunit (DNA-PKcs) and a DNA-end binding component, Ku.11 DNA-PKcs, by virtue of its C-terminal kinase domain, belongs to a family of phosphatidylinositol 3-kinase-like protein kinases (PIKKs), other members of which include the ATM and ATR kinases.12 It is important to mention here that the ATM/ATR kinases implement another important aspect of the cellular response to DSBs, namely the enforcement of cell cycle checkpoints that would give the cell enough time to carry out repair (for more details, see Chapter 1). Interestingly, it has been revealed recently that ATM, while primarily enforcing cell cycle checkpoints, might also be involved, to a lesser extent, in DNA repair, which is discussed later in this chapter.13,14 Ku is a heterodimer of two proteins of approximately 70 and 80 kDa which are termed Ku70 and Ku80, respectively.15 Ku forms a ring-like structure; it first binds to the DNA end, recruits DNA-PKcs and then stabilizes its binding to DNA.16 DNA-PKcs is a serine/threonine kinase with a specificity for S/TQ sites.17 The recruitment of DNA-PKcs to DNA breaks by Ku results in the activation of its kinase function such that it can now phosphorylate other proteins as well as itself. The recruitment of DNA-PKcs to DNA breaks can be visualized within the nucleus by immunostaining irradiated cells with antibodies recognizing the active (phosphorylated) form of DNA-PKcs.18,19 In such stainings, DNA-PKcs forms distinct, punctate “foci” at the sites of DSBs. Indeed, the recruitment of DNA-PKcs to breaks can occur in a matter of seconds as revealed by the imaging of live cells with a single laser-generated DNA damage spot (our unpublished results). DNA-PKcs can tether the broken DNA ends together in a synaptic complex containing two DNA-PKcs molecules; therefore, one function of DNA-PK in DSB repair may be to bridge the broken ends to facilitate rejoining.20 The other function of DNA-PK is to

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recruit and, perhaps, activate the proteins involved in DNA endprocessing and ligation (see below). Interestingly, it was recently shown that DNA-PK interacts specifically with MDC1 (mediator of DNAdamage checkpoint protein 1), and that this interaction, though not required for the initial recruitment of DNA-PK to a break, may serve to enhance the local concentration of the repair enzyme at the sites of DNA damage.21 2.3. The DNA Ligase IV-XRCC4 Complex The co-ordinated assembly of Ku and DNA-PKcs on DNA ends is followed by the recruitment of the DNA ligase IV-XRCC4 complex that is responsible for the rejoining step.22 This complex lies at the center of the NHEJ pathway and is present in all eukaryotes including yeast (which lacks DNA-PKcs but not Ku). Ligase IV exists in a tight complex with XRCC4; the latter stabilizes Ligase IV and stimulates its DNA ligation activity.23 IR-induced phosphorylation of XRCC4 is dependent on DNA-PK in vivo, and the phosphorylation of XRCC4 stimulates the ligation activity of XRCC4-DNA ligase IV in vitro.24,25 We would like to point out here that though XRCC4Ligase IV plays such an important role in the repair of DSBs, it is completely dependent on DNA-PKcs/Ku for its targeting to breaks, and thus functions exclusively in the NHEJ pathway of repair. The importance of Ligase IV in DNA repair is bolstered by the fact that human patients with inherited hypomorphic mutations in Ligase IV are radiosensitive and are impaired in DSB repair.26 In 2006, a potential third partner of the XRCC4/Ligase IV complex was identified by two independent groups.27,28 Named XLF/Cernunnos, this 33 kDa protein interacts with XRCC4/Ligase IV and promotes NHEJ by regulating the activity of this complex. Consequently, human patients with mutations in this protein display radiation sensitivity and impaired V(D)J recombination 2.4. DNA End Processing May Be Required Prior to Repair In addition to the NHEJ core components, several accessory factors have been implicated in the end-processing step of the NHEJ repair

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pathway. End-processing is particularly important for the repair of IR-induced DSBs which may otherwise be unligatable with gaps and end-blocking groups, e.g. 3 -phosphates or phosphoglycolate (PG), and 5 -hydroxyl.29 The 3 -end phosphates can be effectively removed by the polynucleotide kinase/phosphatase (PNKP). In addition, the kinase activity of PNKP could phosphorylate the 5 -hydroxyl ends to facilitate the final ligation or end-joining.30 The removal of 3 PGs may require both apurinic/apyrimidinic endonuclease (APE1) and tyrosyl DNA phosphodiesterase (Tdp1). APE1 is the major PGs activity in human cell extract,31 and is capable of removing PGs from the 3 -blunt or recessed ends, but not from the 3 -protruding ends.32 Tdp1, in contrast, is the only enzyme capable of removing the glycolate from the 3 -protruding ends, leaving the 3 -phosphates to be processed by PNKP.33 Alternatively, the end blocking groups can also be removed by nucleolytic cleavage. The Werner syndrome protein (Wrn) is an ATP-dependent RecQ helicase which also possesses 3 –5 -exonuclease activity.34 Wrn interacts with both Ku35,36 and DNA-PKcs, and is phosphorylated by DNA-PKcs.37 The 3 – 5 -exonuclease activity of Wrn may be required for the removal of 3 -Ps or PGs. Artemis, a 5 –3 -exonuclease, which has also been identified to interact with DNA-PKcs, is also phosphorylated by this kinase.38 Upon complex formation and DNA-PKcs-mediated phosphorylation, Artemis acquires endonucleolytic activity capable of opening the hairpin loop during V(D)J recombination as well as removing both 5 and 3 protruded ends for NEHJ repair.38 Recent evidence also suggested that Artemis could be the downstream target of ATM. Artemis-deficient cells, similar to A-T cells, display a subtle but persistent defect in DSB repair upon IR. Approximately 10% of radiation-induced DSBs are repaired with slow kinetics, probably due to the complexity of damage or “dirty” DSBs, and Artemis/ATM-dependent end-processing activity is likely to be required for repairing the subset complex DSBs.39 Besides processing the ends, there are gaps at DSBs that need to be filled in prior to end ligation. Three members of the PolX family, Polµ, Polλ and terminal deoxynucleotidyl transferase (TdT), all containing the BRCT domain, have been linked to NHEJ repair. TdT is capable of adding

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random nucleotide at the coding end during V(D)J recombination in a template-independent manner.40 Polµ forms a stable complex with XRCC4/LigIV and could catalyze accurate gap filling in the presence of Ku and XRCC4/LigIV.41 In addition, it is interesting that the protein level of Polµ increases after IR, further suggesting a role for Polµ in DSB repair.41 Polλ, like Polµ, also forms a stable complex with XRCC4/LigIV, and its DNA synthesis activity increases upon its interaction with XRCC4/LigaseIV complex.42 2.5. The Kinase Activity of DNA-PKcs and Autophosphorylation DNA-PKcs is a serine/threonine kinase whose kinase activity is critical to DSB repair. This is clear from the fact that DNA-PKcs inactivated by point mutations in its kinase domain fails to complement the DSB-repair defect of DNA-PKcs-deficient cells.43 Although DNA-PK phosphorylates a large number of substrates in vitro, relevant in vivo targets have been rather hard to find. Potential in vivo substrates include XRCC424 and Artemis37 ; but these links have not been solidified by the identification of the actual phosphorylation sites, in vivo demonstration of phosphorylation, and site-directed mutagenesis of the phosphorylation sites. Recently, it became clear that one of the most important substrates of DNA-PK is its catalytic subunit itself. A total of seven potential phosphorylation sites on DNA-PKcs were reported based upon the mass spectrometry of DNA-PKcs autophosphorylated in vitro.18,44 Of these sites, six are clustered in a 38-amino acid stretch in a central region of DNA-PKcs (amino acids 2609 to 2647). Of these sites, at least three have definitely been demonstrated to be phosphorylated in vivo upon irradiation, these phosphorylations are important for NHEJ as mutation of these sites impairs DSB repair and results in cellular radiosensitivity.18,44,45 Interestingly, although it was originally assumed that these sites must represent autophosphorylation sites as they were identified by the analysis of DNA-PKcs autophosphorylated in vitro, it became clear later on that the ATM kinase can contribute to phosphorylation at these sites (our unpublished

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results). This strengthens the emerging concept that ATM, apart from implementing cell cycle checkpoints, also impinges on DNA repair.13,14 Interestingly, these sites are also phosphorylated by the ATR kinase in reponse to UV-induced DNA damage or upon the induction of replication blocks (our unpublished results), which raises the intriguing possibility that DNA-PKcs functions not just in DSB repair but also in the resolution of other types of DNA lesions or structures. Unlike the S/TQ cluster, a site that is not phosphorylated at all by ATM/ATR kinases and that represents a true autophosphorylation site on DNA-PKcs, was identified by the mass spectrometric analysis of endogenous DNA-PKcs from irradiated cells. Such a method can prove to be instrumental in revealing novel phosphorylation sites that are functionally relevant in vivo. The phosphorylation site identified by this method, S2056, is distinct from the S/TQ cluster in that its phosphorylation upon IR is totally dependent upon DNA-PK; also the phosphorylation pattern (temporal and dose-dependent) is different from that of the other identified sites.19 As with the S/TQ cluster, mutation of this site results in radiation sensitivity and abrogated repair. However, one can surmise that the role of S2056 in repair may be different from the S/TQ cluster, or that it may respond to different lesions and these molecular details still need to be worked out. Interestingly, it was recently reported that S2056 lies within a second cluster of potential phosphorylation sites.46 Bona fide as well as potential phosphorylation sites on DNAPKcs are schematically represented in Fig. 2. 2.6. Consequences of Phosphorylation The phosphorylation of DNA-PKcs at the identified sites is obviously of importance as mutations at these sites impair DSB repair and result in radiation sensitivity.18,19,44,45 However the molecular mechanisms underlying the role of phosphorylation in repair have yet to be worked out. Theoretically, DNA-PKcs phosphorylation could lead to DNA-PK complex dissociation, conformational change in DNA-PKcs to facilitate subsequent repair steps, the creation of

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S2023 S2029 ?? S2041 S2053 S2056*

T2609* S2612 T2620 S2624 T2638* T2647*

S3205 FAT

LZ

S3821

S4026 T4102

PI3K

Caspase 3 cleavage

Fig. 2 Schematic diagram depicting phosphorylation sites on DNA-PKcs. Most DNA-PKcs phosphorylation sites have been identified by mass spectrometry of in vitro autophosphorylated DNA-PKcs except S2056 which was identified by the analysis of DNA-PKcs from irradiated HeLa cells. In addition, several serine residues adjacent to S2056 were also reported to be the potential phosphorylation sites based on mutagenesis analysis (indicated by ??). Among all the sites, phosphorylation in vivo was detected only at S2056, T2609, T2638, and T2647 (indicated by *). While in vivo S2056 phosphorylation is clearly due to DNA-PKcs autophosphorylation, preliminary evidence suggests that phosphorylations at T2609, T2638 and T2647 in vivo are also mediated ATM and ATR kinases.

docking sites for other damage-responsive proteins, or the targeting of DNA-PKcs for degradation. The autophosphorylation of purified DNA-PKcs in vitro results in the disruption of the DNA-PK complex and the loss of kinase activity.47,48 Therefore, it is postulated that a similar dissociation of the DNA-PK complex at a break could serve to make space for other factors that may be needed for subsequent repair events. Also, phospho-DNA-PKcs co-localizes with a number of damage-responsive proteins with BRCT (BRCA1 carboxylterminal) domains.18,21 As the BRCT domain was reported to be a phospho-serine/threonine binding module,49–51 the creation of docking sites is a very plausible consequence of DNA-PKcs phosphorylation. 3. REGULATION OF NHEJ IN A CELL CYCLE-DEPENDENT MANNER It has been proposed that NHEJ and HR may be differentially regulated throughout the cell cycle with NHEJ playing a major role in the G1/early S phases and HR playing a major role in the late S/G2 phases when an optimally positioned sister chromatid is available.52 Indeed, DNA-PKcs-deficient V3 cells are less sensitive to IR in the S phase of the cell cycle as compared to G1, indicating that HR may

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partly compensate for NHEJ-deficiency in S phase cells.19 It has been reported that DNA-PK kinase activity is high during the G1 phase and low during the S phase and, consequently, DNA-PK-deficient cells are significantly impaired in DSB repair only during the G1 phase.52 We recently demonstrated that the IR-induced phosphorylation of DNA-PKcs and focus formation, while quite robust in G1 cells, is significantly attenuated in S phase cells, indicating that DNAPK may function primarily in G1 while HR takes over during S/G2.19 Spontaneous HR is increased in DNA-PK-defective cells indicating that the two pathways may compete (either passively or actively) with each other for access to DSBs.53,54 In contrast to the attenuated DNA-PK response in irradiated S phase cells, DSBs generated when a DNA replication fork encounters a damaged template result in robust DNA-PKcs phosphorylation and focus formation.19 Such breaks also induce DNA-PK-dependent phosphorylation of histone H2AX confirming that an active form of DNA-PK is localized at the sites of replication-associated breaks.55 This indicates that in S phase cells, replication-associated breaks, unlike IR-induced breaks, may be preferentially repaired by DNA-PK. 3.1. Dephosphorylation of DNA-PK The phosphorylation of DNA-PKcs is a transient event lasting from 8–24 hours, and it can be generally assumed that once repair is complete, DNA-PKcs may have to dephosphorylate to facilitate subsequent events. As an extension of this line of reasoning, it may also be possible that the dephosphorylation of DNA-PKcs may actually be required for repair to progress to completion. These questions can only be resolved by the identification of phosphatases that impinge on the latter steps in the DNA damage response. A first step in this direction was taken by the identification of protein phosphatase 5 (PP5) as a DNA-PKcs interacting protein that specifically dephosphorylates the catalytic subunit on two sites (T2609 and S2056).56 The dephosphorylation of DNAPKcs by PP5 seems to be of functional significance to NHEJ as the over-expression of dominant negative PP5 in mammalian cells

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lead to increased radiation sensitivity. The role of protein phosphatases in DNA damage responses via their actions on DNAPKcs, ATM, H2AX, and Chk1 is an emerging area of research.57–59 Future research should shed more light on the exact consequences of DNA-PKcs dephosphorylation in NHEJ. 3.2. DSB Repair in the Context of Chromatin Our DNA is tightly wrapped around histones to form chromatin.60 This fact has to be taken into account while we try to understand the complexities of DSB repair within a living cell. Logically, one can envisage that the interaction of DNA with histones may actually serve as an impediment to repair. This concept is well established in the field of transcription where chromatin-remodeling complexes and histone-modifying activities are required to alleviate the inhibition of transcription due to the compaction of chromatin. The importance of the chromatin structure in DSB repair is just beginning to be understood. In this context it is important to discuss one of the earliest events that occur at the site of a break, namely the phosphorylation of a histone subtype H2AX. Histone H2AX is a histone H2A variant that is rapidly phosphorylated in its unique C-terminal tail when DSBs are introduced into mammalian cells.61 This phosphorylation is extensive (spanning about 100 kilobases), and occurs within seconds at the sites of DSBs, resulting in discrete γ-H2AX (phosphorylated-H2AX) foci at the DNA damage sites. DNA-PK, ATM, and ATR have all been implicated in H2AX phosphorylation in mammalian cells.62,63 The phosphorylation of yeast H2A at serine 129 (homologous to serine 139 of mammalian H2AX) causes chromatin decondensation and is required for efficient DNA doublestrand break repair.64 A barrage of recent papers demonstrate that phosphorylated yeast H2A serves to recruit histone modifying and chromatin remodeling activities to DSBs, thereby facilitating DSB repair.65–69 It is conceivable, therefore, that similar activities would have to be recruited to DSBs in mammalian cells to allow the decondensation of chromatin fibers, thereby making the breaks accessible to repair factors. Indeed, it was recently shown that the access

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of RAD51 (a protein that facilitates homologous recombination) to DNA breaks in mammalian cells is facilitated by chromatin unwinding due to histone acetylation.70 Interestingly, γH2AX has been shown to be dephosphorylated by a protein phosphate PP2A; this dephosphorylation step has also been proposed to be important for DSB repair.58,59 4. ANIMAL MODELS AND PATIENTS WITH DEFECTIVE DNA REPAIR Knockout mouse models that are defective in DNA-PKcs, Ku70, Ku80, or Artemis are defective in DSB repair and are radiationsensitive, while the absence of XRCC4 and Ligase IV in mice leads to embryonic lethality due to neuronal apoptosis.71–76 In animal models, inactivation of NHEJ components leads to increased genomic instability, while inactivation of Ku leads to cancer predisposition. Given the important role of the DNA damage response in safeguarding genomic integrity, it is not surprising that defects in these pathways lead to several human disorders with pleiotropic clinical features. In humans no defects have been correlated with a DNA-PKcs deficiency, perhaps due to lethality, but polymorphisms in Ku70 and XRCC4 have been linked to an increased risk of breast cancer.77 A hypomorphic mutation in the Ligase IV gene leads to ligase IV syndrome with developmental and growth delays, and lymphoid tumors in two out of six patients studied.78,79 Mutations in Artemis, that provides the hairpin opening activity in V(D)J recombination and which might be involved in the processing of complex breaks during NHEJ, lead to RS-SCID (severe combined immunodeficiency with sensitivity to ionizing radiation).38,80 Finally, deficiencies in ATM, the main protein that implements cell cycle checkpoints and also impinges on DSB repair, result in Ataxia Telangiectasia, a progressive neurodegenerative, cancer-predisposition syndrome.81 5. FUTURE PERSPECTIVES While we already have a good understanding of the steps involved in NHEJ, a greater understanding of this and other repair processes

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is needed at the molecular level, especially how modifications such as phosphorylations activate or fine-tune repair responses. This is important in light of the numerous disorders associated with defective DNA-damage responses in humans. Also, a better understanding of these processes will lead to the development of effective radiosensitisers that block repair, thereby allowing lower doses to be used for radio- or chemotherapy in the treatment of cancer. ACKNOWLEDGMENTS Our research is supported by grants from NIH (CA50519, CA86936, PO1-CA92584), Department of Defense BCRP (DAMD17-02-1-0439), NASA (NNA05CM04G, NNJ05HD36G) to DJC and a grant from NASA (NNA05CS97G) to SB. REFERENCES 1. Kastan MB, Bartek J. (2004) Cell-cycle checkpoints and cancer. Nature 432: 316–323. 2. Khanna KK, Jackson SP. (2001) DNA double-strand breaks: signaling, repair and the cancer connection. Nat Genet 27: 247–254. 3. Pierce AJ, Stark JM, Araujo FD, et al. (2001) Double-strand breaks and tumorigenesis. Trends Cell Biol 11: S52–59. 4. Bartkova J, Horejsi Z, Koed K, et al. (2005) DNA damage response as a candidate anti-cancer barrier in early human tumorigenesis. Nature 434: 864–870. 5. Gorgoulis VG, Vassiliou LV, Karakaidos P, et al. (2005) Activation of the DNA damage checkpoint and genomic instability in human precancerous lesions. Nature 434: 907–913. 6. Cadet J, Douki T, Gasparutto D, Ravanat JL. (2003) Oxidative damage to DNA: formation, measurement and biochemical features. Mutat Res 531: 5–23. 7. Gellert M. (2002) V(D)J recombination: RAG proteins, repair factors, and regulation. Annu Rev Biochem 71: 101–132. 8. Lieber MR, Ma Y, Pannicke U, Schwarz K. (2004) The mechanism of vertebrate nonhomologous DNA end joining and its role in V(D)J recombination. DNA Repair (Amst) 3: 817–826. 9. Thompson LH, Schild D. (2002) Recombinational DNA repair and human disease. Mutat Res 509: 49–78. 10. Burma S, Chen DJ. (2004) Role of DNA-PK in the cellular response to DNA double-strand breaks. DNA Repair (Amst) 3: 909–918.

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11. Smith GC, Jackson SP. (1999) The DNA-dependent protein kinase. Genes Dev 13: 916–934. 12. Shiloh Y. (2003) ATM and related protein kinases: safeguarding genome integrity. Nat Rev Cancer 3: 155–168. 13. Lobrich M, Jeggo PA. (2005) The two edges of the ATM sword: cooperation between repair and checkpoint functions. Radiother Oncol 76: 112–118. 14. Lobrich M, Jeggo PA. (2005) Harmonising the response to DSBs: a new string in the ATM bow. DNA Repair (Amst) 4: 749–759. 15. Doherty AJ, Jackson SP. (2001) DNA repair: how Ku makes ends meet. Curr Biol 11: R920–924. 16. Walker JR, Corpina RA, Goldberg J. (2001) Structure of the Ku heterodimer bound to DNA and its implications for double-strand break repair. Nature 412: 607–614. 17. Kim ST, Lim DS, Canman CE, Kastan MB. (1999) Substrate specificities and identification of putative substrates of ATM kinase family members. J Biol Chem 274: 37538–37543. 18. Chan DW, Chen BP, Prithivirajsingh S, et al. (2002) Autophosphorylation of the DNA-dependent protein kinase catalytic subunit is required for rejoining of DNA double-strand breaks. Genes Dev 16: 2333–2338. 19. Chen BP, Chan DW, Kobayashi J, et al. (2005) Cell cycle dependence of DNA-dependent protein kinase phosphorylation in response to DNA double-strand breaks. J Biol Chem 280: 14709–14715. 20. Cary RB, Peterson SR, Wang J, et al. (1997) DNA looping by Ku and the DNA-dependent protein kinase. Proc Natl Acad Sci USA 94: 4267–4272. 21. Lou Z, Chen BP, Asaithamby A, et al. (2004) MDC1 regulates DNAPK autophosphorylation in response to DNA damage. J Biol Chem 279: 46359–46362. 22. Martin IV, MacNeill SA. (2002) ATP-dependent DNA ligases. Genome Biol 3: REVIEWS3005. 23. Calsou P, Delteil C, Frit P, et al. (2003) Co-ordinated assembly of Ku and p460 subunits of the DNA-dependent protein kinase on DNA ends is necessary for XRCC4-ligase IV recruitment. J Mol Biol 326: 93–103. 24. Matsumoto Y, Suzuki N, Namba N, et al. (2000) Cleavage and phosphorylation of XRCC4 protein induced by X-irradiation. FEBS Lett 478: 67–71. 25. Leber R, Wise TW, Mizuta R, Meek K. (1998) The XRCC4 gene product is a target for, and interacts with, the DNA-dependent protein kinase. J Biol Chem 273: 1794–1801. 26. Riballo E, Critchlow SE, Teo SH, et al. (1999) Identification of a defect in DNA ligase IV in a radiosensitive leukemia patient. Curr Biol 9: 699–702.

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27. Ahnesorg P, Smith P, Jackson, SP. (2006) XLF interacts with the XRCC4DNA ligase IV complex to promote DNA nonhomologous end-joining. Cell 124: 301–313. 28. Buck D, Malivert L, de Chasseval R, et al. (2006) Cernunnos, a novel nonhomologous end-joining factor, is mutated in human immunodeficiency with microcephaly. Cell 124: 287–299. 29. Ward JF. (1988) DNA damage produced by ionizing radiation in mammalian cells: identities, mechanisms of formation, and reparability. Prog Nucl Acid Res Mol Biol 35: 95–125. 30. Chappell C, Hanakahi LA, Karimi-Busheri F, et al. (2002) Involvement of human polynucleotide kinase in double-strand break repair by nonhomologous end joining. EMBO J 21: 2827–2832. 31. Parsons JL, Dianova II, Dianov GL. (2004) APE1 is the major 3 phosphoglycolate activity in human cell extracts. Nucl Acids Res 32: 3531–3536. 32. Suh D, Wilson DM, III, Povirk LF. (1997) 3 -phosphodiesterase activity of human apurinic/apyrimidinic endonuclease at DNA double-strand break ends. Nucl Acids Res 25: 2495–2500. 33. Inamdar KV, Pouliot JJ, Zhou T, et al. (2002) Conversion of phosphoglycolate to phosphate termini on 3 overhangs of DNA double-strand breaks by the human tyrosyl-DNA phosphodiesterase hTdp1. J Biol Chem 277: 27162–27168. 34. Bohr VA, Cooper M, Orren D, et al. (2000) Werner syndrome protein: biochemical properties and functional interactions. Exp Gerontol 35: 695–702. 35. Li B, Comai L. (2001) Requirements for the nucleolytic processing of DNA ends by the Werner syndrome protein-Ku70/80 complex. J Biol Chem 276: 9896–9902. 36. Orren DK, Machwe A, Karmakar P, et al. (2001) A functional interaction of Ku with Werner exonuclease facilitates digestion of damaged DNA. Nucl Acids Res 29: 1926–1934. 37. Yannone SM, Roy S, Chan DW, et al. (2001) Werner syndrome protein is regulated and phosphorylated by DNA-dependent protein kinase. J Biol Chem 276: 38242–38248. 38. Ma Y, Pannicke U, Schwarz K, Lieber MR. (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. 39. Riballo E, Kuhne M, Rief N, et al. (2004) A pathway of double-strand break rejoining dependent upon ATM, Artemis, and proteins locating to gamma-H2AX foci. Mol Cell 16: 715–724.

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40. Gauss GH, Lieber MR. (1996) Mechanistic constraints on diversity in human V(D)J recombination. Mol Cell Biol 16: 258–269. 41. Mahajan KN, McElhinny N, Mitchell BS, Ramsden DA. (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. 42. Fan W, Wu X. (2004) DNA polymerase lambda can elongate on DNA substrates mimicking non-homologous end joining and interact with XRCC4-ligase IV complex. Biochem Biophys Res Commun 323: 1328–1333. 43. Kurimasa A, Kumano S, Boubnov NV, et al. (1999) Requirement for the kinase activity of human DNA-dependent protein kinase catalytic subunit in DNA strand break rejoining. Mol Cell Biol 19: 3877–3884. 44. Douglas P, Sapkota GP, Morrice N, et al. (2002) Identification of in vitro and in vivo phosphorylation sites in the catalytic subunit of the DNAdependent protein kinase. Biochem J 368: 243–251. 45. Ding Q, Reddy YV, Wang W, et al. (2003) Autophosphorylation of the catalytic subunit of the DNA-dependent protein kinase is required for efficient end processing during DNA double-strand break repair. Mol Cell Biol 23: 5836–5848. 46. Cui X, Yu Y, Gupta S, et al. (2005) Autophosphorylation of DNAdependent protein kinase regulates DNA end processing and may also alter double-strand break repair pathway choice. Mol Cell Biol 25: 10842–10852. 47. Chan DW, Lees-Miller SP. (1996) The DNA-dependent protein kinase is inactivated by autophosphorylation of the catalytic subunit. J Biol Chem 271: 8936–8941. 48. Merkle D, Douglas P, Moorhead GB, et al. (2002) The DNA-dependent protein kinase interacts with DNA to form a protein-DNA complex that is disrupted by phosphorylation. Biochemistry 41: 12706–12714. 49. Manke IA, Lowery DM, Nguyen A, Yaffe MB. (2003) BRCT repeats as phosphopeptide-binding modules involved in protein targeting. Science 302: 636–639. 50. Rodriguez M, Yu X, Chen J, Songyang Z. (2003) Phosphopeptide binding specificities of BRCT domains. J Biol Chem 278: 52914–52918. 51. Yu X, Chini CC, He M, et al. (2003) The BRCT domain is a phosphoprotein binding domain. Science 302: 639–642. 52. Hendrickson EA. (1997) Cell-cycle regulation of mammalian DNA double-strand-break repair. Am J Hum Genet 61: 795–800. 53. Allen C, Kurimasa A, Brenneman MA, et al. (2002) DNA-dependent protein kinase suppresses double-strand break-induced and spontaneous homologous recombination. Proc Natl Acad Sci USA 99: 3758–3763.

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54. Fukushima T, Takata M, Morrison C, et al. (2001) Genetic analysis of the DNA-dependent protein kinase reveals an inhibitory role of Ku in late S-G2 phase DNA double-strand break repair. J Biol Chem 276: 44413–44418. 55. Furuta T, Takemura H, Liao ZY, et al. (2003) Phosphorylation of histone H2AX and activation of Mre11, RAD50, and Nbs1 in response to replication-dependent DNA double-strand breaks induced by mammalian DNA topoisomerase I cleavage complexes. J Biol Chem 278: 20303–20312. 56. Wechsler T, Chen BP, Harper R, et al. (2004) DNA-PKcs function regulated specifically by protein phosphatase 5. Proc Natl Acad Sci USA 101: 1247–1252. 57. Bakkenist CJ, Kastan MB. (2004) Phosphatases join kinases in DNAdamage response pathways. Trends Cell Biol 14: 339–341. 58. Chowdhury D, Keogh MC, Ishii H, et al. (2005) Gamma-H2AX dephosphorylation by protein phosphatase 2A facilitates DNA double-strand break repair. Mol Cell 20: 801–809. 59. Keogh MC, Kim JA, Downey M, et al. (2006) A phosphatase complex that dephosphorylates gamma-H2AX regulates DNA damage checkpoint recovery. Nature 439: 497–501. 60. Luger K, Mader AW, Richmond RK, et al. (1997) Crystal structure of the nucleosome core particle at 2.8 A resolution. Nature 389: 251–260. 61. Rogakou EP, Pilch DR, Orr AH, et al. (1998) DNA double-stranded breaks induce histone H2AX phosphorylation on serine 139. J Biol Chem 273: 5858–5868. 62. Burma S, Chen BP, Murphy M, et al. (2001) ATM phosphorylates histone H2AX in response to DNA double-strand breaks. J Biol Chem 276: 42462–42467. 63. Fernandez-Capetillo O, Lee A, Nussenzweig M, Nussenzweig A. (2004) H2AX: the histone guardian of the genome. DNA Repair (Amst) 3: 959–967. 64. Downs JA, Lowndes NF, Jackson SP. (2000) A role for Saccharomyces cerevisiae histone H2A in DNA repair. Nature 408: 1001–1004. 65. Bird AW, Yu DY, Pray-Grant MG, et al. (2002) Acetylation of histone H4 by Esa1 is required for DNA double-strand break repair. Nature 419: 411–415. 66. Chai B, Huang J, Cairns BR, Laurent BC. (2005) Distinct roles for the RSC and Swi/Snf ATP-dependent chromatin remodelers in DNA double-strand break repair. Genes Dev 19: 1656–1661. 67. Downs JA, Allard S, Jobin-Robitaille O, et al. (2004) Binding of chromatin-modifying activities to phosphorylated histone H2A at DNA damage sites. Mol Cell 16: 979–990.

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68. Morrison AJ, Highland J, Krogan NJ, et al. (2004) INO80 and gammaH2AX interaction links ATP-dependent chromatin remodeling to DNA damage repair. Cell 119: 767–775. 69. van Attikum H, Fritsch O, Hohn B, Gasser SM. (2004) Recruitment of the INO80 complex by H2A phosphorylation links ATP-dependent chromatin remodeling with DNA double-strand break repair. Cell 119: 777–788. 70. Murr R, Loizou JI, Yang YG, et al. (2006) Histone acetylation by TrrapTip60 modulates loading of repair proteins and repair of DNA doublestrand breaks. Nat Cell Biol 8: 91–99. 71. Frank KM, Sekiguchi JM, Seidl KJ, et al. (1998) Late embryonic lethality and impaired V(D)J recombination in mice lacking DNA ligase IV. Nature 396: 173–177. 72. Gao Y, Sun Y, Frank KM, et al. (1998) A critical role for DNA end-joining proteins in both lymphogenesis and neurogenesis. Cell 95: 891–902. 73. Kurimasa A, Ouyang H, Dong LJ, et al. (1999) Catalytic subunit of DNA-dependent protein kinase: impact on lymphocyte development and tumorigenesis. Proc Natl Acad Sci USA 96: 1403–1408. 74. Nussenzweig A, Chen C, da Costa Soares V, et al. (1996) Requirement for Ku80 in growth and immunoglobulin V(D)J recombination. Nature 382: 551–555. 75. Ouyang H, Nussenzweig A, Kurimasa A, et al. (1997) Ku70 is required for DNA repair but not for T cell antigen receptor gene recombination in vivo. J Exp Med 186: 921–929. 76. Rooney S, Sekiguchi J, Zhu C, et al. (2002) Leaky Scid phenotype associated with defective V(D)J coding end processing in Artemis-deficient mice. Mol Cell 10: 1379–1390. 77. Fu YP, Yu JC, Cheng TC, et al. (2003) Breast cancer risk associated with genotypic polymorphism of the nonhomologous end-joining genes: a multigenic study on cancer susceptibility. Cancer Res 63: 2440–2446. 78. O’Driscoll M, Cerosaletti KM, Girard PM, et al. (2001) DNA ligase IV mutations identified in patients exhibiting developmental delay and immunodeficiency. Mol Cell 8: 1175–1185. 79. O’Driscoll M, Gennery AR, Seidel J, et al. (2004) An overview of three new disorders associated with genetic instability: LIG4 syndrome, RS-SCID and ATR-Seckel syndrome. DNA Repair (Amst) 3: 1227–1235. 80. Moshous D, Callebaut I, de Chasseval R, et al. (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. 81. Lavin MF, Shiloh Y. (1996) Ataxia-telangiectasia: a multifaceted genetic disorder associated with defective signal transduction. Curr Opin Immunol 8: 459–464.

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CHAPTER 7

A DNA-damage Response Network of Fanconi Anemia and BRCA Proteins Rong Guo and Weidong Wang∗

ABSTRACT Fanconi anemia (FA) is a rare genetic disorder characterized by congenital abnormalities, bone marrow failure, chromosomal instability and cancer predisposition. The disease has often been considered as a model for studying the repair of interstrand DNA crosslinks (ICLs) because the hallmark feature of FA is cellular hypersensitivity to drugs that cause ICLs. To date, 11 FA genes have been discovered, and their products function in the same DNA damage response network as the breast cancer susceptibility gene products, BRCA1 and BRCA2. One FA protein, FANCD1, is identical to BRCA2, whereas another one, FANCJ is the same as BRIP1, a DNA helicase interacting with BRCA1. A key component of this network is a multisubunit complex, termed FA core complex, which contains at least eight FA proteins, including a ubiquitin ligase (FANCL) and a DNA translocase (FANCM), and is required for the monoubiquitination of FANCD2 in response to DNA damage. Studies of FA have provided novel insights into the mechanisms of genome maintenance and DNA repair.

1. INTRODUCTION In 1967, a Swiss pediatrician, Guido Fanconi, first described Fanconi anemia (FA) in a familial form of aplastic anemia in three brothers ∗ Corresponding

author. Laboratory of Genetics, National Institute on Aging, National Institutes of Health, 333 Cassell Drive, TRIAD center Room 3000, Baltimore, Maryland 21224, USA. Tel.: 410-558-8334; Fax: 410-558-8331; E-mail: [email protected] 177

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of short stature who had increased skin pigmentation. Since then, the disease has been found in all races and ethnic groups, with a prevalence of less than 1 per 100,000 in the general population.1 Although the incidence of FA is rare, the frequency of normal people who carry FA mutations is much higher (about 1 in 300). The disease is characterized by diverse clinical symptoms that often include developmental defects affecting the skeleton (absence of thumbs), kidneys, heart, and other organs. The patients frequently develop life-threatening bone-marrow failure, and are at high risk of developing cancer, including acute myeloblastic leukemia, and head and neck squamous cell carcinomas. FA used to be considered an autosomal recessive disease, in which patients, both male or female, inherited defective gene alleles from both parents. It is now known that one of the FA genes, FANCB, is localized on the X-chromosome, so that in the families of FANCB patients, the disease is transmitted as X-linked: only male members of the family are affected, and they inherit their defective gene only from their mother.2 FA is a genomic instability disease characterized by the increased levels of spontaneous and drug-induced chromosome breakage, and the formation of radial chromosomes (triradial and quadriradial). A higher rate of sister-chromatid exchange has also been reported, particularly in chicken DT40 cells in which FA genes are inactivated. The types of chromosomal abnormalities in FA cells closely resemble those in Bloom syndrome,1 a disease that also features genomic instability and cancer predisposition. In fact, at least eight FA proteins were purified in the same complex with BLM,3 the helicase mutated in Bloom syndrome. Thus, FAproteins, like BLM, are important parts of the cellular machinery that guards the genome, and studies of these proteins should provide an insight into the mechanism of genome maintenance. The hallmark of FA cells is cellular hypersensitivity to crosslinking agents, such as mitomycin C (MMC), diepoxybutane (DEB), and cisplatin. This hypersensitivity provides the basis for a diagnostic test. DNA crosslinking agents lead to covalent bonds between interor intra-strand. The interstrand crosslinks (ICLs) effectively prevent

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the separation of DNA strands, and therefore block critical cellular processes such as DNAreplication, transcription and recombination. These crosslinks must be repaired or bypassed for cells to survive. Organisms have developed different strategies to deal with ICLs.4 In bacteria, the major pathway includes the combined action of nucleotide excision repair and homologous recombination, whereas a minor pathway involves translesion synthesis. In eukaryotes, particularly mammals, these pathways are less understood. The ICL drugs are widely used in chemotherapy for cancer. Understanding the mechanism of their action and repair is thus important for the optimal use of these drugs. FA has been considered an important genetic model for the study of ICL repair because the disease is known to be caused by mutations in many genes (at least 12 now) that are likely to affect various steps of this pathway. The identification of these gene products should therefore reveal the different steps in the pathway. So far, 11 FA gene products have been identified; they indeed appear to participate in a DNA repair network with breast cancer susceptibility gene products BRCA1 and BRCA2. This network of proteins promotes homologous recombination and translesion synthesis pathways to repair the DNA lesions. 2. MANY FA GENE PRODUCTS LACK RECOGNIZABLE DOMAINS AND ARE CONSERVED ONLY IN HIGHER EUKARYOTES FA is genetically heterogeneous, and has been classified into complementation groups by somatic cell hybridization studies using sensitivity to MMC as the assay.1,5 The genes for these groups have been identified by different approaches, including complementation with cDNA libraries, positional cloning, candidate gene screening and protein association. FANCC, FANCA, FANCG, FANCF and FANCE (in the chronological order of their identification) were the first five genes to be identified by functional complementation and positional cloning (Table 1 and Fig. 1). Each of these five FA proteins interacts with the others by yeast two-hybrid-assays

FA

A

66 1.6

Chromosome Location

Protein Product (KD)

Requirement of D2 Ubiquitination

Conservation

16q24.3

163

phosphorylated following DNA damage

+

V

Xp22.31

95

nuclear localization sequence

+

V

10

9q22.3

63

3

13q12-13

380

D2

3

3p25.3

155, 162

2.5

6p21-22

60

+

V

BRC repeats,

−

V

OB domain monoubiquitinated

+

V, insect, worm

+

V

+

V

+

V

F

2

11p15

42

G (XRCC9)

9

9p13

68

TPR repeats, phosphorylated following DNA damage

I

1.5

—

+

V

J (BACH1, BRIP1)

1.5

17q22

140

Helicase activity

−

V, yeast

L (PHF9)

A > C.50,51 These observations imply that Polι must have very specialized cellular functions, one of which could be the somatic hypermutation (SHM) during immunoglobulin development, which is required for antibody diversity. In contrast to its extremely promiscuous behavior on template T, Polι is very accurate at either undamaged or even some adducted template A. This strong template-dependent bias of fidelity is explained by the unique Hoogsteen base-pairing of Polι,52 which provides an elegant mechanism for promoting replication through minor-groove purine adducts that interfere with replication. This along with other observations would support a postulation that Polι was derived from the gene duplication of Polη, which primarily acts on thymine damage, and evolved to protect us from the mutations at damaged adenosines. Interestingly, the biochemical properties of purified Drosophila Polι are more like Polη in its ability to traverse CPD accurately and it does not display the promiscuous behavior on template T.53 4.4.5. Polκ Several groups consistently reported that mammalian Polκ cannot bypass CPD in vitro. However, DinB1-deleted mouse cells are moderately UV-sensitive. It turned out that Polκ can efficiently extend G and A placed opposite the 3 -T of a T-T CPD, after nucleotide incorporation opposite O6 -methylguanine and 8-oxoguanine by Polδ, or from the base mispairs in undamaged DNA, suggesting that Polκ may be an extender of damaged or mispaired bases.54 On the other hand, Polκ can incorporate opposite an abasic site preferentially with A; however, further efficient extension requires T as the next template base, which results in a –1 deletion. This observation indicates that after A incorporation opposite the abasic site, Polκ misaligns the primer terminal A with the next template T, and thus slips the abasic site during subsequent DNA synthesis.55 One striking observation is that Polκ can bypass a variety of benzo[a]pyrene (B[a]P) diol

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epoxide (BPDE) stereoisomers such as N 2 -dG (dG-N 2 -BPDE),56 the most potent known carcinogenic compounds produced by industrial and cigarette smoke, with much higher efficiency and accuracy than Polη and Polι, by inserting the correct C opposite the bulky lesion. While the above observation suggests that Polκ plays an important role in suppressing B[a]P-induced mutagenesis in humans, it would be difficult to propose that all mammals coevolved to respond to environmental pollutions which occur primarily in the past two centuries. Furthermore, the bacterial Polκ homolog Pol IV is also capable of error-free bypass of the dG-N 2 BPDE DNA adduct in E. coli cells, despite the fact that they do not have an enzymatic system (cytochrome P-450) to activate B[a]P into BPDE. It has been suggested that thymine glycol and some estrogenderived DNA adducts may serve as natural substrates for Polκ, since Polκ can bypass these lesions with reasonable efficiency in vitro.54 The substrate specificity of specialized DNA polymerases, especially the highly mutable ones such as Polι and Polκ, suggests that their genes may be under strict regulation. Limited studies indicate that XPV, POLI and POLK genes are ubiquitously expressed in all tissues with a small increase in proliferating tissues such as the testis. In addition, several studies indicate that mammalian POLK can be induced by specific DNA damaging agents such as 3-methylcholanthrene, a compound similar to B[a]P, and to UV and doxorubicin, both of which produce thymine glycol in addition to other lesions.54 Human and mouse POLK promoters contain xenobiotic responsive elements (XREs) which are known to be binding sites for the AhR-Arnt transcriptional activator. The AhR (arylhydrocarbon receptor) is activated by polycyclic aromatic hydrocarbons (PAHs), among which B[a]P is the most extensively studied. This regulatory mechanism indicates that POLK is induced in response to specific DNA damaging agents that produce adducts bypassed by Polκ. The DinB-Polκ homolog is found in Schizosaccharomyces pombe and Caenorhabditis elegans, but not in S. cerevisiae or Drosophila melanogaster. The substrate specificity and biological functions of Polκ in fission yeast and nematode remain to be determined.

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4.5. X-Family DNA Polymerases X-family DNA polymerases also participate in lesion bypass in a broader sense. Mammalian Polβ, which is involved in shortpatch base excision repair, also possesses a 5 -deoxyribose phosphate (dRP) lyase activity (see Chapter 3). Mammalian cells also contain three other X-family polymerases that are involved in nonhomologous end joining (NHEJ; see Chapter 7), namely, a templateindependent terminal deoxynucleotidyl transferase (TdT) and the template-dependent Polµ and Polλ. These three members constitute a gradient of template dependence to handle double-strand DNA ends with partial complementary overhangs to promote accuracy in NHEJ.57 In an in vitro DNA synthesis assay, Polµ predominantly generates –1 deletions at dinucleotide repeats and promotes microhomology search and pairing between the primer and template strands, resulting in frameshift mutations.58 4.6. DNA Polymerase Switch during TLS Knowing the limited specificity and processivity of Y-family DNA polymerases, several questions may arise. Firstly, how is the replicative DNA polymerase displaced at a replication fork fork blocked by DNA damage? Secondly, how are TLS polymerases recruited to the damage site? Thirdly, how are the inserter and extender coordinated to bypass the given lesion? Finally, if a DNA lesion (e.g. an abasic site) can be bypassed by several TLS polymerases, how are they selected in cells? The coordination of TLS polymerases in lesion bypass was first reported by Drs. Prakash and their colleagues in 2000, in which Polι and Polζ acted sequentially to bypass a variety of lesions tested, whereas neither enzyme was able to complete the lesion bypass.50 It was soon reported that Polη, Polι and Polκ contain putative PCNAbinding motifs and physically interact with PCNA. Furthermore, the catalytic activities of these enzymes are stimulated in the presence of PCNA, indicating a crucial role for PCNA in the targeting of TLS polymerases to the stalled replication fork.7 Consistent with the discovery of the covalent modification of PCNA by Ub and

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SUMO,26 and the genetic evidence that monoubiquitylated PCNA promotes TLS,27 it was recently reported that Polη preferentially interacts with monoubiquitylated PCNA,28,29 providing an underlying mechanism for a polymerase switch in response to DNA damage. It was recently reported that ubiquitylated PCNA activates Polι and Rev1 activities,59 and that the Ub-binding domains found in Y-family polymerases play a critical role in regulating TLS.60 While switching from a replicative polymerase to Y-family polymerases is probably mediated by PCNA modification, the noncatalytic activity of Rev1 as described before appears to play a central role in polymerase switching during TLS, possibly from insertion to extension. Supporting the above notion are several recent reports that Rev1 interacts with Polη, Polκ, Polι, Polλ and the Rev7 subunit of Polζ 7 ; that all these interactions require only the C-terminal 100 amino acids of Rev1; and that the binding of Polκ to Rev1 can be competed by increasing the amounts of Rev7.61 It should be noted that Rev1 from S. cerevisiae or C. elegans does not interact with Polη in a yeast two-hybrid assay and their C-terminal regions are not conserved with mammalian Rev1, although budding yeast Rev1 does interact with Rev7. The polymerase switch model is further supported by the colocalization of TLS polymerases and PCNA following DNA damage (primarily using UV). The discrete DNA damage-induced nuclear foci can be observed after a mild detergent treatment of cultured mammalian cells, which presumably represent large TLS protein complexes at stalled replication forks. The Polη foci are co-localized with those of Polι,62 as well as Rev1.63 Furthermore, UV-induced Polη foci also co-localize with those of PCNA bound on chromatin,29 and Polη nuclear foci formation is abolished in cells with Rad18 deletion.28 These observations collectively suggest that monoubiquitylated PCNA by Rad6-Rad18 is essential for TLS complex assembly. Bacteria such as E. coli do not contain Polζ homologs; hence, Pol IV and Pol V appear to perform both insertion and extension reactions. Nevertheless, polymerase switching between a replicative polymerase and a TLS polymerase also requires a β sliding-clamp,

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a functional equivalent to eukaryotic PCNA. One report64 indicates that the Pol III holoenzyme arrests immediately proceeding the damaged base, causing the β clamp to dissociate from the replicative machinery and to interact with Pol V, which catalyzes a short patch (1–60 bases) of DNA synthesis in the presence of RecA. Another report65 suggests that different subunits of the β clamp dimer can bind to Pol III and Pol IV at the same time. When Pol III is stalled at the replication fork, Pol IV takes control, and rapidly switches back after lesion bypass. The polymerase switch in this case does not involve the dissociation of either polymerase. It is yet unclear whether the difference between the above two reports is due to the use of different TLS polymerases, or the experimental conditions. 4.7. Untargeted Mutagenesis A mutation that occurs at a site where there is a premutagenic DNA lesion is called a (lesion) targeted mutation, whereas mutations found at sites unrelated to DNA lesions are referred to as untargeted mutations (UTMs). One example of UTM is the increase in the mutagenesis of unirradiated phage λ when infecting an E. coli host that has been previously UV irradiated — a phenomenon known as Weigle mutagenesis. Genetic analysis and an in vitro reconstructive study demonstrate that both chromosomal and phage UTMs are attributable to the SOS response and the activities of Pol IV and Pol V, as well as other proteins required for TLS. Nevertheless, the critical roles of Y-family polymerases in UTM are demonstrated by the observation that dinB overexpression is sufficient to dramatically increase both targeted and untargeted mutagenesis in the absence of DNA damage and an SOS response.66 Little is known about UTM in mammalian cells. By using an experimental system to monitor mutagenesis of a target gene in a plasmid transfected into mammalian host cells previously exposed to the chemical mutagen N-methyl-N  -nitro-N-nitrosoguanidine (MNNG), the increase of UTM that was observed exhibits a distinct mutational spectrum with predominant transversion mutations and a preference for hot spots.67 It appears that a decrease in replication

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fidelity rather than a compromised mismatch repair is responsible for MNNG-induced UTM, and that Polζ,68 as well as possibly other Y-family polymerases, plays a role in UTM. 4.8. Somatic Hypermutation B lymphocytes undergo the additional alterations induced by antigen stimulation. Activated B lymphocytes proliferate vigorously in the germinal centers of lymphoid follicles, where two different types of genetic alterations, somatic hypermutation (SHM) and class-switch recombination (CSR), take place. SHM introduces point mutations in the V-region exons of immunoglobulin light and heavy chains, and B cells expressing mutated immunoglobulin with a high affinity are selected, resulting in affinity maturation. SHM introduces mainly nucleotide substitutions, and occasionally insertions and deletions, into target sequences at a rate of approximately 10−3 per base pair per generation, or a frequency of as much as 5% in vivo. Fifty percent to 60% of SHM base substitutions are transitions, exhibiting a strong bias for transitions over transversions. In a two-phase SHM model,69 DNA lesions are introduced by activationinduced cytidine deaminase (AID) in Phase I via its direct DNA deamination in single-stranded DNA, within the transcribed region, or via its RNA-editing activity (AID is a homolog of an RNA editing enzyme APOBEC1), which recognizes a putative mRNA precursor and converts it to mRNA encoding an endonuclease. The cytidine deaminase activity is targeted to RGYW/WRCY hot spots (where R is A or G, Y is C or T, W is A or T, and the position of the mutated nucleotide within the motif is underlined). In this phase of SHM, the duplex containing the U:G mismatch has two possible fates: replication which will fix the C/G→T/A transition mutation; or deglycosylation by a uracil glycosylase which will create an abasic site which undergoes further mutagenic repair. Mutations may be introduced opposite the abasic site by error-prone polymerases. In Phase II SHM, mutations occur at adjacent A and T, which involve mismatch repair as well as error-prone polymerases. Mice deficient in Msh2, Msh6 or Exo1 all showed a shift of the SHM spectrum towards

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more mutations at G and C, and fewer mutations at A and T. It was suggested that the G:U mismatch is recognized by the Msh2–Msh6 heterodimer, leading to Exo1-mediated excision and a repair patch that is re-synthesized by TLS DNA polymerases. Either PCNA modification or an uncharacterized factor was postulated to recruit TLS DNA polymerases to the Msh2–Msh6-recognized sites.70 It was found that mice and B cell lines with low levels of Rev3 mRNA had reduced SHM frequencies at all base pairs and, in the mouse study, an impaired affinity maturation of B cells. XPV cells have a significantly reduced mutational frequency at Ig V regions, but this reduction is restricted to A:T base pairs. Targeted inactivation of the RAD30B/POLI gene in the Burkitt’s lymphoma BL2 cell line abolishes SHM,71 suggesting that Polι is absolutely required for SHM. BL2 only displays hypermutations at G:C base pairs that are dependent on AID and Polι, suggesting that the AID-generated uracil in BL2 is processed by this polymerase. In contrast, the 129derived strains of mice, which have a spontaneous missense mutation in the gene encoding Polι, undergo normal SHM, both in terms of frequency and mutational pattern.72 Rev1-deficient DT40 chicken cells display a markedly reduced level of non-templated immunoglobulin gene mutation, indicating a defect in TLS. On the other hand, mice deficient in Polµ, λ, κ and β undergo normal SHM, suggesting that these polymerases are not required for SHM. Thus, it appears that SHM depends largely on specific Y-family DNA polymerases, perhaps because they tend to generate base substitutions rather than insertions and deletions. The fact that B cells deficient in Polζ, η and ι do not have identical phenotypes argues against a strictly compensation model. For example, deficiency in Polι impacts G:C base pair mutation; deficiency in Polη impacts mutations from A:T base pairs; whereas deficiency in Polζ causes a reduction of mutations at all base pairs. Therefore, the data for SHM are best explained by a model wherein a mutasome complex of TLS DNA polymerases is recruited to the uracilcontaining Ig locus and Polη mutates A:T base pairs with a slight strand bias, Polι mutates G:C base pairs and Polζ extends from mismatches generated by both Polη and Polι.73

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5. TLS AND CANCER Specialized TLS DNA polymerases are required to bypass DNA damage lesions that would otherwise cause replication arrest and cell death. A high fidelity lesion bypass would protect cells from genome instability and would be expected to prevent carcinogenesis. However, when operating on non-canonical templates, such as undamaged DNA or on non-cognate lesions, these polymerases are found to exhibit a considerably reduced fidelity, causing increased mutation rates. Experimental overexpression of TLS polymerases can also lead to an increased mutation rate, indicating that the deregulation of TLS polymerases may contribute to carcinogenesis. Xeroderma pigmentosum variant (XPV) is arguably the best case that links TLS to cancer. XPV patients, who represent approximately 20% of all patients with XP, have no abnormalities in NER but are defective in TLS of TT dimers. The POLH gene is located on chromosome 6p21.1, spanning 40 kb of DNA with 11 exons and an open reading frame of 2139 bp. More than 25 alleles of POLH have been identified in the cell lines derived from XPV patients, including base pair substitutions, small insertions and deletions resulting in frameshift and premature termination. Most of these mutations abolish or severely affect in vitro TLS activities while others may interfere with nuclear localization or nuclear foci formation after UV irradiation. In addition to the mutations identified from XPV patients, six single nucleotide polymorphisms (SNPs) with amino acid substitutions have been deposited in the NCBI database. The biochemical property, biological significance and cancer epidemiological studies of these SNPs have not been reported to date. SNPs of REV1 and POLI are suggested to be associated with lung cancer risk. An SNP of Phe257Ser of Rev1 was associated with squamous cell carcinoma risk and an SNP of Thr706Ala of Polι was associated with adenocarcinoma and squamous cell carcinoma risk, particularly in individuals above than 61 years of age.74 A study of 11 human lung cancer cell lines found a common polymorphism of Thr706Ala in six cell lines. Furthermore, mouse PolI was identified as a candidate for the pulmonary adenoma resistance 2 (Par2) gene,75

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which plays a role in the resistance to urethane-induced lung adenoma/adenocarcinoma. A few studies have looked at the correlation between cancer and the altered expression of TLS genes. One reported POLK overexpression in 21 of the 29 matched specimens of non-small cell lung cancer.76 In addition, five matched specimens exhibited elevated POLK expression in both tumor and control tissues, whereas only one non-tumor tissue expressed a higher level of POLK than its tumor counterpart. Since Southern analysis did not detect POLK gene amplification, the elevated POLK expression is likely attributable to deregulated transcription. Another recent study reported an overexpression of specialized polymerases in more than 45% of the 68 tumor samples,77 demonstrating a greater than two-fold enhanced expression of at least one specialized polymerase. Of particular note is the finding that Polβ was overexpressed at both the mRNA and protein levels in approximately one-third of all the tumor types studied, with overexpression being particularly frequent in the uterus, ovary, prostate and stomach samples. Polλ and Polι were found to be overexpressed to a significant extent in a range of tumor types, albeit less frequently than Polβ. On the other hand, Polκ was also found to be rarely overexpressed in tumors but commonly underexpressed in many samples. For example, in a study of 131 self-paired cancerous and non-tumor samples, including 23 lung cancers, 49 stomach cancers and 59 colorectal cancers, Polκ, η, ι and ζ are all significantly downregulated in human lung, stomach, and colorectal cancers, with the only exception of Polη in colorectal cancers,78 suggesting that these enzymes are probably not closely associated with the elevated mutations in human cancer. The significance of the altered expression of TLS polymerases in tumorigenesis requires further investigations. 6. MAMMALIAN ERROR-FREE DDT? Although it is not yet clear whether mammalian cells contain an error-free DDT pathway like their yeast counterpart, several pieces of evidence support its existence. Firstly, with the only exception

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Table 2. Genes Involved in Yeast DDT/PRR and Their Human Homologs S. cerevisiae RAD5 RAD6 RAD18 UBC13 MMS2 POL30 POL32

Humana SMARCA3/HLTF HHR6A/UBE2A HHR6B/UBE2B RAD18 hUBC13/UBE2N hMMS2/UBE2V2 UEV1/UBE2V1 PCNA POLD3

Chromosomal Locationb 3q25.1–q26.1 Xq24–q25 5q23–q31 3p25–p24 12q22 8q11.21 20q13.2 20p12 11q14

a The

human genes on the list are the most closely related sequence homologs to the corresponding yeast genes. In the case of RAD5, SHPRH has been indicated to be a human RAD5 orthologue (K. Myung, personal communication), while the functional relationship of SMARCA3/HLTF with RAD5 is currently unknown. b Chromosomal locations of human genes are from http://www. ncbi.nlm.nih.gov/mapview.

of RAD5, mammalian genes homologous to yeast error-free DDT (Table 2) have been identified and characterized. In particular, human and mouse MMS2 and UBC13 genes have been isolated and demonstrated to functionally complement the corresponding yeast mutants.79,80 The yeast and human Ubc13-Mms2 complex structure and biochemical activities are also highly conserved.13,16,17 Secondly, the antisense suppression of MMS2 in cultured human cells results in an increase in UV-induced mutagenesis and completely abolishes the UV-induced gene conversion events,81 which is reminiscent of the yeast mms2 mutant.14 Thirdly, Ubc13 and Mms2 form nuclear foci after DNA damage; these foci are co-localized with newly synthesized DNA,82 as well as PCNA (P Andersen and Xiao, data not shown). Finally, the ablation of Mms2 or Ubc13 activity by interference RNA resulted in a significant increase in spontaneous Rad51 nuclear foci,82 which is indicative of endogenous double-strand breaks, possibly due to collapsed replication forks. However, despite the efforts from several laboratories, polyubiquitylated PCNA has not been detected.28,29 Nevertheless, the experimental inhibition of Lys63-linked ubiquitylation in cultured human cells resulted in

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compromised DNA repair/lesion bypass phenotypes, reminiscent of the yeast mms2 mutant,83 and further supporting an error-free mode of DDT in mammals. If error-free DDT does exist and operates in mammals as in budding yeast, it would play a crucial role in protecting cells from genomic instability and carcinogenesis. The implication of mammalian DDT on public health is yet to be appreciated. Acknowledgments We wish to thank Dr. B Wouters for the unpublished results, Dr. L Huang for helpful discussion and Michelle Hanna for proofreading the manuscript. This work was supported by the Chinese Natural Science Foundation operating grants to FX and YNY, and the Canadian Institutes of Health Research operating grants to WX. References 1. Broomfield S, Hryciw T, Xiao W. (2001) DNA postreplication repair and mutagenesis in Saccharomyces cerevisiae. Mutat Res 486: 167–184. 2. Barbour L, 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. 3. Pastushok L, Xiao W. (2004) DNA postreplication repair modulated by ubiquitination and sumoylation. Adv Protein Chem 69: 279–306. 4. Woodgate R. (1999) A plethora of lesion-replicating DNA polymerases. Genes Dev 13: 2191–2195. 5. Wang Z. (2001) Translesion synthesis by the UmuC family of DNA polymerases. Mutat Res 486: 59–70. 6. Prakash S, Prakash L. (2002) Translesion DNA synthesis in eukaryotes: a one- or two-polymerase affair. Genes Dev 16: 1872–1883. 7. Friedberg EC, Lehmann AR, Fuchs RP. (2005) Trading places: how do DNA polymerases switch during translesion DNA synthesis? Mol Cell 18: 499–505. 8. Kuzminov A. (1999) Recombinational repair of DNA damage in Escherichia coli and bacteriophage λ. Microbiol Mol Biol Rev 63: 751–813. 9. Pham P, Seitz EM, Saveliev S, et al. (2002) Two distinct modes of RecA action are required for DNA polymerase V-catalyzed translesion synthesis. Proc Natl Acad Sci USA 99: 11061–11066.

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10. Pickart CM. (2001) Mechanisms underlying ubiquitination. Annu Rev Biochem 70: 503–533. 11. Pickart CM. (2001) Ubiquitin enters the new millennium. Mol Cell 8: 499–504. 12. Hofmann RM, Pickart CM. (1999) Noncanonical MMS2-encoded ubiquitin-conjugating enzyme functions in assembly of novel polyubiquitin chains for DNA repair. Cell 96: 645–653. 13. McKenna S, Spyracopoulos L, Moraes T, et al. (2001) Noncovalent interaction between ubiquitin and the human DNA repair protein Mms2 is required for Ubc13-mediated polyubiquitination. J Biol Chem 276: 40120–40126. 14. Broomfield S, Chow BL, 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. 15. Sancho E, Vila MR, Sanchez-Pulido L, et al. (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. 16. VanDemark AP, Hofmann RM, Tsui C, et al. (2001) Molecular insights into polyubiquitin chain assembly: crystal structure of the Mms2/Ubc13 heterodimer. Cell 105 711–720. 17. Moraes TF, Edwards RA, McKenna S, et al. (2001) Crystal structure of the human ubiquitin conjugating enzyme complex, hMms2-hUbc13. Nat Struct Biol 8: 669–673. 18. Villalobo E, Morin L, Moch C, et al. (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. 19. Brown M, Zhu Y, Hemmingsen SM, Xiao W. (2002) Structural and functional conservation of error-free DNA postreplication repair in Schizosaccharomyces pombe. DNA Repair 1: 869–880. 20. Hochstrasser M. (2000) Evolution and function of ubiquitin-like protein-conjugation systems. Nat Cell Biol 2: E153–157. 21. Nelson JR, Lawrence CW, Hinkle DC. (1996) Thymine-thymine dimer bypass by yeast DNA polymerase ζ. Science 272: 1646–1649. 22. Nelson JR, Lawrence CW, Hinkle DC. (1996) Deoxycytidyl transferase activity of yeast REV1 protein. Nature 382: 729–731. 23. Brusky J, Zhu Y, 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. 24. Xiao W, Chow BL, Broomfield S, 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.

FA

November 2, 2006 9:1

262

WSPC/SPI-B414: DNA Repair, Genetic Instability & Cancer

ch09

Fang Xu, Yingnian Yu and Wei Xiao

25. Ulrich HD, Jentsch S. (2000) Two RING finger proteins mediate cooperation between ubiquitin-conjugating enzymes in DNA repair. EMBO J 19: 3388–3397. 26. Hoege C, Pfander B, Moldovan GL, et al. (2002) RAD6-dependent DNA repair is linked to modification of PCNA by ubiquitin and SUMO. Nature 419: 135–141. 27. Stelter P, Ulrich HD. (2003) Control of spontaneous and damageinduced mutagenesis by SUMO and ubiquitin conjugation. Nature 425: 188–191. 28. Watanabe K, Tateishi S, Kawasuji M, et al. (2004) Rad18 guides polη to replication stalling sites through physical interaction and PCNA monoubiquitination. EMBO J 23: 3886–3896. 29. Kannouche PL, Wing J, Lehmann AR. (2004) Interaction of human DNA polymerase η with monoubiquitinated PCNA: a possible mechanism for the polymerase switch in response to DNA damage. Mol Cell 14: 491–500. 30. Haracska L, Kondratick CM, Unk I, et al. (2001) Interaction with PCNA is essential for yeast DNA polymerase η function. Mol Cell 8: 407–415. 31. Haracska L, Torres-Ramos CA, Johnson RE, et al. (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. 32. Pfander B, Moldovan GL, Sacher M, et al. (2005) SUMO-modified PCNA recruits Srs2 to prevent recombination during S phase. Nature 436: 428–433. 33. Papouli E, Chen S, Davies AA, et al. (2005) Crosstalk between SUMO and ubiquitin on PCNA is mediated by recruitment of the helicase Srs2p. Mol Cell 19: 123–133. 34. Xiao W, Chow BL, Fontanie T, et al. (1999) Genetic interactions between error-prone and error-free postreplication repair pathways in Saccharomyces cerevisiae. Mutat Res 435: 1–11. 35. Seki M, Masutani C, Yang LW, et al. (2004) High-efficiency bypass of DNA damage by human DNA polymerase Q. EMBO J 23: 4484–4494. 36. Lawrence CW. (2004) Cellular functions of DNA polymerase ζ and Rev1 protein. Adv Protein Chem 69: 167–203. 37. Wagner J, Gruz P, Kim SR, et al. (1999) The dinB gene encodes a novel E. coli DNA polymerase, DNA Pol IV, involved in mutagenesis. Mol Cell 4: 281–286. 38. Fuchs RP, Fujii S, Wagner J. (2004) Properties and functions of Escherichia coli: Pol IV and Pol V. Adv Protein Chem 69: 229–264. 39. Tang M, Bruck I, Eritja R, et al. (1998) Biochemical basis of SOS-induced mutagenesis in Escherichia coli: reconstitution of in vitro lesion bypass

FA

November 2, 2006 9:1

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40.

41.

42.

43.

44.

45.

46.

47. 48.

49.

50.

51. 52.

53.

ch09

263

dependent on the UmuD’2C mutagenic complex and RecA protein. Proc Natl Acad Sci USA 95: 9755–9760. Tang M, Shen X, Frank EG, et al. (1999) UmuD’(2)C is an error-prone DNA polymerase, Escherichia coli Pol V. Proc Natl Acad Sci USA 96: 8919–8924. Nair DT, Johnson RE, Prakash L, et al. (2005) Rev1 employs a novel mechanism of DNA synthesis using a protein template. Science 309: 2219–2222. Otsuka C, Kunitomi N, Iwai S, et al. (2005) Roles of the polymerase and BRCT domains of Rev1 protein in translesion DNA synthesis in yeast in vivo. Mutat Res 578: 79–87. McDonald JP, Levine AS, 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. Johnson RE, Prakash S, Prakash L. (1999) Efficient bypass of a thymine-thymine dimer by yeast DNA polymerase, Polη. Science 283: 1001–1004. Johnson RE, Kondratick CM, Prakash S, Prakash L. (1999) hRAD30 mutations in the variant form of xeroderma pigmentosum. Science 285: 263–265. Masutani C, Kusumoto R, Yamada A, et al. (1999) The XPV (xeroderma pigmentosum variant) gene encodes human DNA polymerase η. Nature 399: 700–704. Johnson RE, Washington MT, Prakash S, Prakash L. (2000) Fidelity of human DNA polymerase η. J Biol Chem 275: 7447–7450. Washington MT, Prakash L, Prakash S. (2001) Yeast DNA polymerase η utilizes an induced-fit mechanism of nucleotide incorporation. Cell 107: 917–927. Zhang Y, Yuan F, Wu X, Wang Z. (2000) Preferential incorporation of G opposite template T by the low-fidelity human DNA polymerase ι. Mol Cell Biol 20: 7099–7108. Johnson RE, Washington MT, Haracska L, et al. (2000) Eukaryotic polymerases ι and ζ act sequentially to bypass DNA lesions. Nature 406: 1015–1019. Zhang Y, Yuan F, Wu X, et al. (2001) Response of human DNA polymerase ι to DNA lesions. Nucleic Acids Res 29: 928–935. Nair DT, Johnson RE, Prakash S, et al. (2004) Replication by human DNA polymerase-ι occurs by Hoogsteen base-pairing. Nature 430: 377–380. Ishikawa T, Uematsu N, Mizukoshi T, et al. (2001) Mutagenic and nonmutagenic bypass of DNA lesions by Drosophila DNA polymerases dpolη and dpolι. J Biol Chem 276: 15155–15163.

FA

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264

WSPC/SPI-B414: DNA Repair, Genetic Instability & Cancer

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54. Ohmori H, Ohashi E, Ogi T. (2004) Mammalian Pol κ: regulation of its expression and lesion substrates. Adv Protein Chem 69: 265–278. 55. Ohashi E, Ogi T, Kusumoto R, et al. (2000) Error-prone bypass of certain DNA lesions by the human DNA polymerase κ. Genes Dev 14: 1589– 1594. 56. Shen X, Sayer JM, Kroth H, et al. (2002) Efficiency and accuracy of SOSinduced DNA polymerases replicating benzo[a]pyrene-7,8-diol 9,10epoxide A and G adducts. J Biol Chem 277: 5265–5274. 57. McElhinny N, Havener JM, Garcia-Diaz M, et al. (2005) A gradient of template dependence defines distinct biological roles for family X polymerases in nonhomologous end joining. Mol Cell 19: 357–366. 58. Zhang Y, Wu X, Yuan F, et al. (2001) Highly frequent frameshift DNA synthesis by human DNA polymerase µ. Mol Cell Biol 21: 7995–8006. 59. Garg P, Burgers PM. (2005) Ubiquitinated proliferating cell nuclear antigen activates translesion DNA polymerases η and REV1. Proc Natl Acad Sci USA 102: 18361–18366. 60. Bienko M, Green CM, Crosetto N, et al. (2005) Ubiquitin-binding domains in Y-family polymerases regulate translesion synthesis. Science 310: 1821–1824. 61. Guo C, Fischhaber PL, Luk-Paszyc MJ, et al. (2003) Mouse Rev1 protein interacts with multiple DNApolymerases involved in translesion DNA synthesis. EMBO J 22: 6621–6630. 62. Kannouche P, Fernandez de Henestrosa AR, Coull B, et al. (2002) Localization of DNA polymerases η and ι to the replication machinery is tightly co-ordinated in human cells. EMBO J 21: 6246–6256. 63. Tissier A, Kannouche P, Reck MP, et al. (2004) Co-localization in replication foci and interaction of human Y-family members, DNA polymerase pol η and REVl protein. DNA Repair 3: 1503–1514. 64. Fujii S, Fuchs RP. (2004) Defining the position of the switches between replicative and bypass DNA polymerases. EMBO J 23: 4342–4352. 65. Indiani C, McInerney P, Georgescu R, et al. (2005) A sliding-clamp toolbelt binds high- and low-fidelity DNA polymerases simultaneously. Mol Cell 19: 805–815. 66. Tang M, Pham P, Shen X, et al. (2000) Roles of E. coli DNA polymerases IV and V in lesion-targeted and untargeted SOS mutagenesis. Nature 404: 1014–1018. 67. Zhang X, Yu Y, Chen X. (1994) Evidence for nontargeted mutagenesis in a monkey kidney cell line and analysis of its sequence specificity using a shuttle-vector plasmid. Mutat Res 323: 105–112. 68. Zhu F, Jin CX, Song T, et al. (2003) Response of human REV3 gene to gastric cancer inducing carcinogen N-methyl-N’-nitro-Nnitrosoguanidine and its role in mutagenesis. World J Gastroenterol 9: 888–893.

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69. Maizels N. (2005) Immunoglobulin gene diversification. Annu Rev Genet 39: 23–46. 70. Lee GS, Brandt VL, Roth DB. (2004) B cell development leads off with a base hit: dU:dG mismatches in class switching and hypermutation. Mol Cell 16: 505–508. 71. Faili A, Aoufouchi S, Flatter E, et al. (2002) Induction of somatic hypermutation in immunoglobulin genes is dependent on DNA polymerase ι. Nature 419: 944–947. 72. McDonald JP, Frank EG, Plosky BS, et al. (2003) 129-derived strains of mice are deficient in DNA polymerase ι and have normal immunoglobulin hypermutation. J Exp Med 198: 635–643. 73. Diaz M, Lawrence C. (2005) An update on the role of translesion synthesis DNA polymerases in Ig hypermutation. Trends Immunol 26: 215–220. 74. Sakiyama T, Kohno T, Mimaki S, et al. (2005) Association of amino acid substitution polymorphisms in DNA repair genes TP53, POLI, REV1 and LIG4 with lung cancer risk. Int J Cancer 114: 730–737. 75. Lee GH, Matsushita H. (2005) Genetic linkage between Polι deficiency and increased susceptibility to lung tumors in mice. Cancer Sci 96: 256– 259. 76. Wang J, Kawamura K, Tada Y, et al. (2001) DNA polymerase κ, implicated in spontaneous and DNA damage-induced mutagenesis, is overexpressed in lung cancer. Cancer Res 61: 5366–5369. 77. Albertella MR, Lau A, O’Connor MJ. (2005) The over-expression of specialized DNA polymerases in cancer. DNA Repair 4: 583–593. 78. Pan Q, Fang Y, Xu Y. (2005) Down-regulation of DNA polymerases κ, η, ι, and ζ in human lung, stomach, and colorectal cancers. Cancer Lett 217: 139–147. 79. Xiao W, Lin SL, Broomfield S, et al. (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. 80. Ashley C, Pastushok L, McKenna S, et al. (2002) Roles of mouse UBC13 in DNA postreplication repair and Lys63-linked ubiquitination. Gene 285: 183–191. 81. Li Z, Xiao W, McCormick JJ, Maher VM. (2002) Identification of a protein essential for a major pathway used by human cells to avoid UVinduced DNA damage. Proc Natl Acad Sci USA 99: 4459–4464. 82. Andersen PL, Zhou H, Pastushok L, et al. (2005) Distinct regulation of Ubc13 functions by the two ubiquitin-conjugating enzyme variants Mms2 and Uev1A. J Cell Biol 170: 745–755. 83. Chiu RK, Brun J, Ramaekers C, et al. (2006) Lysine 63-polyubiquitination guards against translesion synthesis-induced mutations. PLoS Genetics 2: 1070–1083.

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CHAPTER 10

Nucleases in DNA Repair, Replication and Recombination: Flap Endonuclease-1 as a Paradigm Purnima Singh and Binghui Shen∗

ABSTRACT The DNA of an organism suffers from external environmental stresses as well as internal insults and errors resulting from replication and recombination. There are multiple DNA repair pathways to maintain the integrity of the genome. DNA nucleases play a crucial role in mismatch repair, nucleotide excision repair, base excision repair and double-strand break repair. This chapter summarizes the role of nucleases in DNA repair, replication and recombination. One of such nucleases, flap endonuclease1 (FEN-1), is discussed in detail as a multifunctional and structure-specific nuclease involved in several nucleic acid processing pathways, including RNAprimer removal, long patch base excision repair, and the resolution of di- and tri-nucleotide repeat secondary structures and stalled DNA replication forks. The multiple functions of FEN-1 are regulated via several means, including the formation of complexes with different protein partners, nuclear localization in response to cell cycle or DNA damage, and post-translational modifications. Its functional deficiency is predicted to cause genetic diseases, including Huntington’s disease, myotonic dystrophy and cancers.

∗ Corresponding

author. Department of Radiation Biology, City of Hope National Medical Center and Beckman Research Institute, Duarte, CA 91010. Tel.: (626)-301-8879; Fax: (626)-301-8280; E-mail: [email protected] 267

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1. INTRODUCTION Organisms are constantly exposed to endogenous and exogenous agents that damage DNA. There are various forms of DNA damage, such as strand breaks, base modifications, crosslinks and mismatches. There are also multiple DNA repair pathways, including mismatch repair (MMR), nucleotide excision repair (NER), base excision repair, homologous recombination (HR) and non-homologous end joining (NHEJ), which have been described in detail in earlier chapters. Each of these repair pathways targets a specific type of damage and requires a number of proteins. Central to most DNA repair pathways is a nucleolytic step that is required for eliminating the damaged nucleotides. Nucleases cleave the phosphodiester bonds between a deoxyribose and a phosphate residue, thereby producing 5 -terminal phosphate and 3 -terminal hydroxyl groups. Exonucleases cleave either from the 5 - or the 3 -end, whereas endonucleases hydrolyze internal phosphodiester bonds. Nucleases act in a variety of structural frameworks, ranging from site-specific (e.g. abasic endonuclease) to structure-specific (e.g. members of the Rad2 nuclease family such as XPG, EXO1 and FEN-1) and non-specific (e.g. DNase I). The identification of proteins as nucleases is usually possible via primary sequence analysis due to common conserved domains, usually consisting of acidic and basic residues that form the active site. The active site residues coordinate catalytically essential divalent cations, such as Mg2+ or Mn2+ , that are important for water activation and stabilization of the intermediates of phosphodiester hydrolysis. Defects in the repair pathways or specific nucleases can lead to the accumulation of unrepaired DNA damage and unresolved structures. This results in an increased mutation rate and genomic instability, which can lead to cancer and other serious diseases. This chapter will briefly highlight the roles of various DNA nucleases in multiple mammalian DNA repair pathways (Table 1) and then discuss in detail about flap endonuclease-1 (FEN-1), a multifunctional and structure-specific nuclease involved in several nucleic acid processing pathways.

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Human DNA Repair Nucleases

Nuclease

Activity

Pathway

Ref.

EXO1 APE1 APE2 XPG XPF-(ERCC1) FEN-1

5 to 3 exo AP endo/3 to 5 exo AP endo/3 to 5 exo endo, 3 -incision endo, 5 -incision 5 to 3 exo, flap/ gap endo 3 to 5 exo

MMR BER BER NER NER BER, repl. (Okazaki)

(1–3) (4,5) (6,7) (8,9) (9,10) (11,12)

DSB repair, SSA, repl, rec.

(13)

endo 5 to 3 exo

repl. (restart) DSB repair

(14,15) (16,17)

3 to 5 3 to 5 3 to 5 3 to 5 3 to 5 endo 3 to 5 3 to 5

repl. (proofreading), MMR (?) repl. (proofreading), MMR (?) repl. of mt DNA (proofreading) unknown (proofreading?) unknown (proofreading?) repl. (Okazaki) BER unknown (proofreading?)

(18,19) (18,19) (19,20) (21,22) (21,22) (12,24) (25,26) (27)

MRE11(Rad50-NBS1) MUS81-(EME1) Artemis(DNA-PKcs) Pol δ Pol ε Pol γ TREX1 TREX2 DNA2 WRN p53

exo exo exo exo exo exo exo

2. NUCLEASES IN DNA REPAIR PATHWAYS 2.1. Exonuclease-1 (EXO1) in Mismatch Repair (MMR) The mammalian mismatch repair system functions in postreplicative genome surveillance to remove base mismatches and small insertion/deletion loops (IDLs). The Msh2-Msh6 complex recognizes single base mismatches and 1 bp IDL mutations, and the MSH2-MSH3 complex recognizes 1 bp as well as 2–4 bp IDLs. Upon mismatch or IDL recognition, the heterodimer Mlh1-Pms2 is recruited, and a nick is then made either 3 or 5 of the mismatch. The nuclease that incises the nascent DNA strand has yet to be identified in eukaryotic systems. When the strand break is located 5 to the mismatch, EXO1 degrades the nascent strand using its 5 to 3 exonuclease activity2 (Fig. 1). EXO1 is a member of the Rad2 family

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Fig. 1 Exonuclease 1 in MMR. EXO1 is a 5 - to 3 -exonuclease involved in a 5 excision of the damaged unmethylated strand. It can also possibly function as a 3 - to 5 -nuclease. In addition, the 3 -exonucleolytic activities of pol δ and ε may be involved in a 3 -excision.

of structure-specific nucleases. Nucleases involved in 3 to 5 excision, when the nick site is located 3 of the mismatch, have not been definitively identified, but DNA pol δ and ε and EXO1 may be involved. A recent report by Dzantiev et al.3 shows that EXO1 can act bidirectionally (5 to 3 hydrolysis directed by a 5 -strand break and 3 to 5 excision directed by a 3 -nick) and the polarity is modulated by other interacting proteins. In addition to its role in MMR, EXO1 also has a function in recombination and double strand break (DSB) repair.1

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2.2. AP Endonuclease (APE1), Flap Endonuclease-1(FEN-1) and Werner Syndrome Protein (Wrn) in Base Excision Repair (BER) BER is responsible for the repair of non–bulky lesions produced by the oxidation, alkylation or deamination of DNA bases. BER is initiated by a damage specific DNA glycosylase that cleaves the N-glycosylic bond between base and sugar, producing an apurinic/apyrimidinic (AP) site (Fig. 2). AP sites can also be a direct damage product. AP sites are processed by BER in essentially two different ways. An AP endonuclease can hydrolyze the DNA sugar–phosphate backbone 5 of an AP site, producing a 3 -terminal hydroxyl group and a 5 -dRP (5 -deoxyribose 5 -phosphate) moiety. The 5 -dRP moiety can be processed during short-patch BER by the dRPase activity of pol β, leading to a one-nucleotide gap. Subsequently, a single nucleotide is inserted by pol β and the remaining nick is sealed by a complex of DNA ligase III and XRCCI.4 Longpatch BER may be required in the presence of modified AP sites where the 5 -moiety cannot be removed by dRPase activity. After strand displacement by pol β and pol δ/pol ε, a flap structure is formed, which is cleaved by FEN-1 in a PCNA-dependant manner followed by synthesis and ligation by DNA ligase I. In human cells, APE1 is the major AP endonuclease. hAPE1 has a strong AP endonuclease activity, but it also exhibits other DNA repair activities: 3 - to 5 -exonuclease, phosphodiesterase, 3 -phosphatase and RNase H; however, it should be noted that these additional activities are much weaker than its AP endonuclease activity. Recently, it has been shown that APE1 also has a 3 -mismatch exonuclease activity, and thus might also be considered as a proofreading enzyme. Indeed, APE1 is a multifunctional enzyme involved in DNA repair, transcription regulation and oxidative signaling. A second AP endonuclease (hAPE2) has also been identified, but shows only weak AP endonuclease activity. Recent reports suggest that APE2 participates in both nuclear and mitochondrial BER, and also that nuclear APE2 functions in the PCNAdependent BER pathway.4–7

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Fig. 2 AP endonuclease, FEN-1 and Wrn in BER. APE1 hydrolyses 5 of an AP site. The resulting 5 -abasic terminus can be processed by pol β. After strand displacement, a flap structure is formed that can be cleaved by FEN-1 which has intrinsic 5 -FEN, 3 -EXO and GEN activities.

The Werner syndrome protein (WRN) has a 3 to 5 exonuclease activity. Recently, Harrigan et al.26 demonstrated that WRN participates in BER in vivo and is likely to promote the efficiency of pol β mediated BER via its exonuclease and helicase activities. They based their conclusions on the fact that WRN exonuclease removed 3 -mismatches and worked cooperatively with pol β on

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BER intermediates containing 3 -mismatches. They also demonstrated that the down-regulation of WRN expression resulted in an elevated cellular sensitivity to MMS and a greatly reduced LP BER. 2.3. Xeroderma Pigmentosum Complementation Group G (XPG) and F (XPF-ERCC1) in Nucleotide Excision Repair (NER) NER removes a variety of forms of DNA damage, including photoproducts induced by UV and other bulky lesions produced by various chemicals. NER consists of two sub-pathways: global genome repair (GGR), which removes the damage in the genome overall, and transcription-coupled repair (TCR), which repairs the transcribed strands of active genes. The main difference in GGR and TCR is the requirement of different factors during the initial recognition steps. A pre-incision complex is formed by XPC-TFIIH, XPA, RPA and the two structure-specific endonucleases, XPF-ERCC1 and XPG. TFIIH and two ATP-dependent helicases, XPB and XPD, are responsible for opening the DNA double helix around the lesion. A dual incision occurs by XPG and XPF-ERCC1, which cuts 3 and 5 to the lesion, respectively. In this way, the damage is released in a 24–32 nucleotide long oligonucleotide. The resulting gap is filled by pol δ/ε, and the remaining nick is sealed by ligase I (Fig. 3).28 The endonuclease XPG belongs to the Rad2 family of structurespecific nucleases. Members of this family contain an N-terminal and internal domain with highly conserved acidic amino acid residues that are essential for nuclease activity. In addition to the 3 -incision during NER, XPG has a structural function during NER and in the not well-characterized TCR BER. XPG helps to open the DNA around a lesion and to stabilize the damage recognition complex. XPG is also able to cleave several structures such as bubbles, splayed arms, stem loops and flaps.8,9 Besides, XPF-ERCC1 can also cleave 3 -flap structures and have an NER-dependent role in homologous recombination, single-strand annealing (SSA) and repair of

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Fig. 3 XPG and XPF in NER. After unwinding the DNA around the lesion, dual incision by XPF-ERCC1 (5 to the damage) and XPG (3 to the damage) releases the damage. The resulting gap is filled by pol δ/ε.

interstrand crosslinks. In addition, XPF-ERCC1 has a role in the formation of UV-induced chromosome exchanges, which indicates that XPF-ERCC1 functions in the bypass or repair of damage during replication.10

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2.4. MRE11(-Rad50-NBS1), Mus81(-Eme1) and Artemis(-DNAPKcs) in Double-Strand Break Repair Double-strand breaks can be repaired either by homologous recombinational repair (HR) or non-homologous end joining (NHEJ). HR uses a homologous DNA template and is highly accurate, whereas NHEJ rejoins the broken ends without using a template and is often accompanied by the loss of several nucleotides. During HR, DSBs are converted to 3 -single-stranded DNA tails, which are bound by RPA. The processing of DSBs probably requires MRE11-Rad50-NBS1 complex.13 MRE11 is a 3 - to 5 -exonuclease on DNA duplexes, and is also an endonuclease on ssDNA and hairpin structures.29 Rad52 interacts with RPA and promotes the binding of Rad51 to ssDNA, which may be stabilized by Rad51 paralogues. Subsequently, the Rad51 bound ssDNA invades a homologous molecule in a reaction stimulated by Rad54. After DNA synthesis and ligation, two Holliday junctions are formed and branch migration can occur. The Mus81-Eme1 heterodimer resolves the Holliday junction (Fig. 4). NHEJ is initiated by the binding of Ku70-Ku80 dimers to the DNA ends. In higher eukaryotes the DNA-PKcs is subsequently recruited. DNA-PKcs phosphorylates Artemis, thereby activating its endonucleolytic activity. The Artemis-DNA-PKcs complex cleaves the 5 - and 3 -overhangs and also nicks DNA hairpins near the tip.17,30 The 5 - and 3 -overhang cleavage is important for preparing dsDNA breaks for ligation by XRCC4/DNA ligase IV. DSBs that are not suitable for ligation may be processed by MRE11-Rad50-NBS1 and other nucleases such as FEN-1.31 In addition, the MRE11-Rad50NBS1 possibly acts as an exonuclease during SSA (Fig. 4). 2.5. Proofreading by 3
- to 5
-exonucleases In eukaryotic cells, DNA pol α, δ and ε are essential for the replication of nuclear DNA, and pol γ for the replication of mitochondrial DNA. Pol δ, Pol ε and Pol γ, but not Pol α, possess a 3 - to 5 -exonuclease activity for proofreading and the removal of misincorporated nucleotides. Pol δ has a function in DNA synthesis during BER, NER, MMR and recombination. Similar to pol δ, pol ε is

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Fig. 4 MRE11, Mus81 and Artemis in DSB Repair. The Mre11/Rad50/NBS1 complex processes the DNA termini of the double-strand break before the initiation of invasion by Rad51. A key intermediate in the homologous recombination is the Holliday intermediate. Resolvases such as Mus81-Eme1 cleave the Holliday junction to separate the two duplexes. In NHEJ DNA-PKcs stimulate the nuclease activity of Artemis, which cuts away protruding single-stranded regions at the DNA ends and creates double-stranded structures that are good ligase substrates.

likely to be involved in DNA synthesis during BER, NER, MMR and recombination. Like Pol α, Pol β and the translesion polymerases lack intrinsic proofreading activity. Several 3 - to 5 -exonucleases have been identified that may substitute for the missing proofreading function of some of the polymerases. TREX1 and its homologue TREX2, exhibit the major 3 - to 5 -exonuclease activity in human cells and process ssDNA and dsDNA. The TREX proteins can remove

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incorrect nucleotides inserted by these DNA polymerases to generate the paired 3 -termini necessary for continued DNA synthesis. The TREX1 protein has been demonstrated to function as an editing exonuclease for DNA pol β in a reconstituted base excision repair assay.32 The activity of the TREX proteins in conjunction with a helicase activity could provide the necessary components to facilitate the excision step in DNA mismatch repair in human cells, as has been indicated for EXO1 in bacteria. 3 - to 5 -exonuclease activities have also been discovered for WRN, p53 and MRE11. In complex with Ku70/Ku80, WRN efficiently digests ssDNA and dsDNA with mismatched nucleotides.33 Most human cancers have mutations in the conserved DNA binding and exonuclease domain of p53. It has been proposed that p53 could participate in DNA repair processes through its 3 - to 5 exonucleolytic activity.34 p53 has been shown to physically interact with pol α and found to strongly stimulate an in vitro reconstituted BER system, which is suggestive of its role as a proofreader for these polymerases. 3. FLAP ENDONUCLEASE-1 (FEN-1) FEN-1 is a structure-specific metallonuclease that is best known for its involvement in RNA primer removal and long patch base excision repair.11,35 FEN-1 recognizes and cleaves a typical DNA 5 -flap substrate generated by pol β or δ strand-displacement synthesis during okazaki fragment maturation. FEN-1 is involved in multiple pathways, some of which are seemingly contradictory (i.e. genome maintenance vs. apoptotic DNA fragmentation). To date, at least 20 proteins are known to interact with FEN-1, some forming distinct complexes that affect one or more FEN-1 activities presumably to direct FEN-1 to a particular DNA metabolic pathway. In addition, the protein localizes in the nucleus and undergoes several post-translational modifications to optimize its biochemical activities or to regulate its interaction with protein partners in response to environmental stress and cell cycle. Furthermore, there has been a rapid accumulation of knowledge regarding the structural biology

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of FEN-1 and the assignment of function(s) (i.e. DNA-binding vs. catalysis) to specific residues.36 The associated implications of its biochemistry for human disease have also been an intense area of discussion. Finding an inter-relationship among the various biochemical activities, metabolic pathways, and pathological consequences due to the functional loss of FEN-1 has significant implications for human health. 3.1. Multiple Biochemical Activities Involved in Distinct DNA Metabolic Pathways The 5 -flap endonuclease (FEN) and 5 - to 3 -exonuclease (EXO) activities were the first to be attributed to FEN-1 by the Lieber group37 and have been extensively reviewed recently.38 FEN-1 was shown to be an efficient 5 -flap endonuclease on DNA or RNA flaps, but it also cleaves pseudo-Y structures (a flap structure lacking the adjacent upstream primer) to a lesser extent.37,39 A double-flap structure, which has a 3 -single nucleotide (nt) tail in addition to the 5 -flap, was recently found to be the optimal and in vivo substrate for FEN-1 (Fig. 5A).40–43 Furthermore, FEN-1 was shown to cleave 5 -double-flap and 5 -flap DNA structures in the context of a nucleosome.44 FEN activity is considered crucial for the enzyme to fulfill its tasks in Okazaki fragment maturation and BER.11,35,38 The 5 - to 3 -exonuclease activity of FEN-1 cleaves nick, gap and 5 -end recessed dsDNA.37,39,45 In addition, FEN-1 can exonucleolytically cleave blunt-ended DNA, but with less efficiency. The role of the exonuclease activity is less obvious, but it has been implicated in Okazaki fragment maturation38 and apoptosis (Fig. 5B).46 Recently, two separate studies determined that FEN-1 has a gap endonuclease (GEN) activity.46,47 CRN-1, the FEN-1 homologue in C. elegans, was shown to cleave the template strand of a gapped DNA duplex resembling an intermediate DNA structure generated during apoptotic DNA fragmentation.46 Human FEN-1 (hFEN-1) was found to cleave the template strand of gapped DNA fork and bubble substrates that mimics a stalled DNA replication fork (Fig. 5A). This GEN activity is independent of its 5 -flap endonuclease activity in that it does not require a 5 -free ssDNA end for its activity.

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Fig. 5 Various biochemical activities of FEN-1. (A) Three categories of substrates with various conformations. FEN-1 possesses three different activities, including 5 -flap endonuclease, 5 -exonuclease and gap endonuclease activity. (B) A dynamic model for the FEN-1 in vitro activities from the 5 -flap removal to the double-strand DNA break. FEN-1 first cleaves the 5 -flap structure thereby generating a nick. Subsequently it removes a few nucleotides from the 5 -end and then cleaves the template strand resulting in a double-strand DNA break. The size and thickness of arrows represent the strength of the activities. The color of the arrows represents different activities: Red — flap endonuclease activity; blue — exonuclease activity; orange — gap endonuclease activity.47

Based on this biochemical evidence, a scenario can be proposed for the stepwise degradation of a 5 -flap DNA by FEN-1. It can be hypothesized that FEN-1 first cleaves the 5 -flap structure to generate a nick. With a nick in the DNA duplex, FEN-1 removes a few nucleotides from the 5 -end to produce a gap, which is then cleaved on the template strand resulting in a dsDNA break (Fig. 5B).

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Considering this scenario, FEN-1 would be harmful to cells if unregulated, and it has been shown that the over-expression of FEN-1 does result in an increase of dsDNA breaks after γ irradiation.48 The scenario in Fig. 5B may not occur normally in vivo due to the fact that the GEN activity of the FEN-1 is approximately 10-fold weaker than the FEN activity. In addition, the role of post-translational modifications and protein interaction partners in the regulation of the three FEN-1 activities to prevent this scenario should not be overlooked. For example, DNA ligase I may compete with FEN-1 for the nick site and possibly ligate the nick site before EXO activity can occur. In response to apoptotic stimuli, GEN and EXO activities of FEN1 can also be stimulated to promote apoptotic DNA fragmentation through interaction with endonuclease G (Endo G), which is normally found in mitochondria and thus, separated from FEN-1.7 3.2. Critical Structural Elements for Various Functions of FEN-1 Based on the crystal structure of FEN-1 homologues and mutational studies over the years, biochemical functions can be assigned to specific amino acid residues and motifs in FEN-1 proteins (Fig. 6A). 3.2.1. Catalysis Initial studies by Shen et al.49 led to the identification of seven conserved aspartic and glutamic acid residues in human FEN-1 (D34, D86, E158, E160, D179, D181 and D233), which when mutated, resulted in the complete loss of flap endonuclease activity. Recently, Qiu et al.36 identified two positively charged amino acid residues, K93 and R100, that are also important for catalysis. The mutation of each did not affect substrate binding of FEN-1, but completely abolished FEN activity. 3.2.2. Substrate binding A helical clamp is formed by a helix-loop-helix (HLH) motif located over the active center. The helical clamp contains a number of

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Fig. 6 Structure and function of FEN-1. (A) Functional motifs and critical residues of human FEN-1. The conserved acidic residues coordinate two Mg2+ , forming an active center for catalysis (dark blue). Seven clusters, (1)–(7), of positively charged residues important for substrate binding (red). Loops 1, 2, 4 and 7 interact with the upstream portion of the DNA substrate, whereas loops 5 and 6 including the H3TH motif interact with the downstream portion. Loop 3 located in the helical clamp region is responsible for the interaction with single-stranded DNA flap. Residues important for post-translational modification are shown in blue: phosphorylation sites (S187); acetylation sites (K354, K375, K377, and K380). The other two clusters of residues are responsible for PCNA interaction (PCNA, yellow) and nuclear localization (NLS, green).47 (B) Interaction interface of A. fulgidus FEN-1 and DNA substrate. (Reprinted from Cell 116: 39–50, 2004; Chapados et al. Structural basis for FEN-1 substrate specificity and PCNAmediated activation in DNA replication and repair, with permission from Elsevier).36 The DNA substrate is kinked by ∼90◦ . The H3TH motif interacts with the downstream DNA duplex while the helical clamp is responsible for the interaction with the single strand 5 -flap. Four loops (1, 2, 4 and 7) contribute to binding of the upstream DNA duplex. Additional amino acid residues form a hydrophobic wedge and pocket to stabilize the 3 -base pair and 3 -flap, respectively.36 Abbreviations: Nt — N terminus; Ct — C terminus; Nuclease — nuclease activity domain; PCNA — proliferating cell nuclear antigen; NLS — nuclear localization signal.

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positively charged and bulky amino acid residues on its inner side that are thought to contact the ssDNA flap of the substrates.50,51 In addition to the helical clamp, the Pyrococcus furiosus FEN-1 (pfFEN-1) crystal structure revealed a positively-charged groove containing the active center and a H3TH motif, which were proposed to mediate the binding of ssDNA and dsDNA portions of the flap substrates, respectively.50 The recent structure of Archaeoglobus fulgidus FEN-1 (afFEN-1) bound to DNA identified two additional HLH motifs that contact the upstream portion of the DNA flap substrate.36 Furthermore, biochemical and mutational analyses of Pyrococcus horikoshii FEN-1 (phFEN-1) revealed a total of five loop regions that are important for FEN-1 DNA binding.52 As summarized in Fig. 6A, the amino acid residues located in seven loop regions are critical for binding certain regions of flap DNA substrate. Loops 1, 2, 4 and 7 interact with the upstream portion of dsDNA substrate, whereas loops 5 and 6, which include the H3TH motif, interact with the dsDNA downstream portion. Loop 3, located in the helical clamp, is responsible for interaction with the single-stranded flap. These data are consistent with the structure of the afFEN-1-DNA complex (Fig. 6B).36 The downstream duplex portion of the flap substrate binds to FEN-1 using the H3TH domain and is kinked approximately 90◦ with respect to the upstream portion of the duplex. The 5 -ssDNA flap is postulated to lie in the groove created by the helical clamp.53,54 Furthermore, a single nucleotide 3 -flap on the upstream primer was shown to reside in a pocket formed by a conserved threonine and a backbone carbonyl moiety. In addition, the last base pair of the upstream duplex, which is adjacent to the single-nucleotide 3 -flap, stacks on a structure termed the hydrophobic wedge (Fig. 6B). Although the C-terminal amino acids present in the eukaryotic FEN-1 homologues are absent in the archaeal or eubacterial FEN-1 proteins, they may also be important for substrate binding in eukaryotic FEN-1s.55 The mutation of these conserved positively charged amino acid residues43 or the truncation of this region severely affects the enzyme activities of FEN-1.

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3.2.3. PCNA interaction The Lieber and Burgers groups were the first to report that human and yeast PCNA increases DNA polymerase processivity by acting as a molecular clamp. Via peptide mapping experiments and protein truncation analysis, a conserved region in the C terminus of the hFEN-1 nuclease (337 QGRLDDFFK345 ) has been shown to be the PCNA interaction motif.56,57 3.2.4. Post-translational modifications Because FEN-1 is involved in a number of biochemical processes, it is of considerable interest to determine how these multiple activities of FEN-1 are regulated. An obvious way to regulate FEN-1 is post-translational modification. For example, the transcriptional coactivator p300 has been shown in vitro and in vivo to acetylate four C-terminal lysines, Lys-354, Lys-375, Lys-77 and Lys-380 of FEN1,55 which were previously shown to be important for DNA binding (Fig. 6A).55 FEN-1 acetylation is induced by UV treatment of cells and correlated with decreased DNA binding.55 Interestingly, most of the lysines within the C-terminal tail are conserved in higher eukaryotes indicating that the acetylation of these residues might be an evolutionary conserved regulatory mechanism. Another identified FEN-1 post-translational modification is phosphorylation. Cyclin–dependent kinase (Cdk), Cdk1-Cyclin A, phosphorylates FEN-1 at Ser-187 in the late S phase. In vitro Cdk1cyclin A phosphorylation of FEN-1 Ser-187, which is located in the internal nuclease domain, has been shown to reduce the endo- and exo-nuclease activities of FEN-1 without affecting DNA binding. In addition, the phosphorylation of FEN-1 abolishes PCNA binding. These findings indicate that phosphorylation is a crucial cell-cycle regulatory factor of FEN-1 activity.43 3.2.5. Nuclear localization FEN-1 is required to migrate into the nucleus to function in eukaryotic cells. Primary sequence analysis of eukaryotic FEN-1s

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identified a bipartite C-terminal motif that is rich in positively charged amino acid residues as a putative nuclear localization signal (NLS) (Fig. 6A). Further studies using site-directed mutagenesis on the two clusters of positively charged residues demonstrated that the residues Lys-354, Arg-355, Lys-356, Lys-365, Lys-366 and Lys-367 are the critical elements of the FEN-1 NLS (Fig. 6A).58 In addition, this study also demonstrated that nuclear localization of FEN-1 is cell-cycle dependent and inducible upon the treatment of cells with DNA damaging agents.58 This important observation indicates that cell cycle and DNA damage triggers regulate the nuclear localization of FEN-1. 3.3. Interacting Partners of FEN-1 The type of proteins with which FEN-1 interacts is evidence of its pivotal role in several DNA metabolic pathways. To date, at least 20 different proteins involved in several pathways have been reported to interact with hFEN-1. Based on the known effects of these interactions on the biochemical activities of FEN-1, its interacting partners can be grouped into six classes (Table 2). The first group includes protein partners assisting FEN-1 in RNA primer removal during DNA replication. In eukaryotic cells, DNA synthesis is primed by the primase/polymerase α (primosome), which synthesizes the iRNA/DNA initiator primer.59 After primer synthesis, replication factor C (RFC) loads the homotrimeric ring-shaped protein known as PCNA onto dsDNA. PCNA acts as a sliding clamp molecular adaptor that localizes bound proteins to DNA. When the processive replisome encounters the downstream Okazaki fragment, a portion of the iRNA/DNA primer is displaced to form a 5 -flap structure, which needs to be removed prior to ligating the remaining DNA segments. FEN-1 interacts with PCNA, pol δ, replication protein A (RPA) and DNA ligase I for efficient Okazaki fragment processing.60 To date three models of 5 -flap removal have been proposed.12,23,39,61 The most recent and favored model involves the ssDNA binding protein RPA, the helicase/nuclease DNA2, and FEN-1. According to this model, RPAcoats the ssDNAflap generated

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Proteins Interacting With FEN-1

Proteins

Possible Functions

Organisms*

References

PCNA RPA RFC hnRNP DNA pol α DNA pol ε DNA2 WRN BLM DNA pol β APE-1 Rad9-Rad1-Hus1 Pol4 Dnl4/Lif1 Endo G HIV Integrase p300

DNA replication „ „ „ „ „ „ „ „ DNA repair „ „ „ „ Apoptosis HIV replication Acetylation

Sc, H Sc H H Sc ct Sc H H H H H Sc Sc Ce H/HIV H

(74) (75) (76) (77) (75) (78) (79) (62) (63) (65) (62) (69) (70) (70) (46) (71) (55)

*Initial work was done with protein homologues from different organisms: Sc — Saccharomyces cerevisiae; H — Humans; Ce — Caenorhabditis elegans; HIV — Human immunodeficiency virus type 1; ct — calf thymus.47

by pol δ-mediated strand displacement synthesis and recruits DNA2 and FEN-1 to the flap. Because RPA stimulates DNA2 and inhibits FEN-1, DNA2 cleaves a large portion of the flap such that RPA can no longer bind. The dissociation of RPA alleviates FEN-1 inhibition. Thus, FEN-1 cleaves the remaining 5 -flap to create a nick, which is then ligated. Other proposals suggested that the DNA2-mediated pathway is activated only when FEN-1 activity is impaired or low, such as during the formation of long flaps coated by RPA.12 FEN1 also interacts with members of the RecQ helicase family (WRN and BLM), and the possible physiological roles of these interactions with FEN-1 in DNA replication and replication fork rescue have been discussed.62,63 Moreover, phenotypes associated with DNA2 yeast mutants can be rescued by the expression of BLM or WRN, which stimulate FEN-1 activity.63,64 It is interesting to speculate based on these results that the complex formed between FEN-1 and

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either RecQ helicase is the functional homologue of DNA2. Furthermore, Imamura and Campbell have postulated that a helicase-like BLM may be necessary to facilitate strand displacement synthesis,64 which suggests that the RecQ helicases might coordinate the formation and FEN-1-mediated removal of 5 -flap structures during Okazaki fragment maturation. The second group of FEN-1 interacting proteins is DNA repair proteins. In mammalian cells, single-base lesions are repaired by “single-nucleotide base excision repair (BER)” or long patch BER. APE1 recognizes and cleaves 5 to the apurinic/apyrimidinic sites, thereby generating a nick on which pol β is loaded. If the lyase activity of pol β cannot remove the deoxylribosyl moiety after it has added the complementary nucleotide to the free 3 -hydroxyl, strand displacement synthesis occurs, thereby creating a small 5 -flap (2–10 nts) that must be cleaved by FEN-1 with subsequent ligation of the nick to complete the repair. The role of FEN-1 in long-patch BER is regulated and coordinated by its physical interaction with important BER components like pol β, APE-1 and PCNA.62,65,66 Furthermore, WRN has recently been shown in vitro to participate in long-patch BER by facilitating pol β strand-displacement synthesis via its helicase activity.67 Therefore, the WRN interaction with, and stimulation of, FEN-1 may also be important for an efficient 5 -flap removal in long-patch BER. A more recent proposal implicated the FEN-1/WRN complex in replication fork rescue through break-induced repair.47 FEN-1 was also shown to interact with the human Rad9-Rad1-Hus1 checkpoint complex (9-1-1 complex). The 9-1-1 complex, a heterotrimeric protein that is similar to the PCNA toroidal sliding-clamp, is induced by DNA-damage in a P53dependent manner.68 Although the 9-1-1 complex is unable to stimulate pol δ synthesis like PCNA, it has been shown to stimulate the FEN activity of FEN-1 on flap substrates.69 The role of this interaction in vivo remains to be shown, but because 9-1-1 accumulates at the sites of DNA damage, the interaction of FEN-1 with 9-1-1 is likely to be important in stimulating FEN-1 activity for DNA repair. Recently, scFEN-1 was shown to interact with Pol4 and Dnl4/Lif1, which are

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components of non-homologous end-joining (NHEJ) dsDNA break repair pathway.70 The third category which consists of apoptotic proteins is exemplified by endonuclease G (Endo G). Parrish et al.46 demonstrated that CRN-1, the C. elegans FEN-1 homologue, in association with CPS-6 (Endo G), mediates stepwise DNA degradation during apoptosis. This is the first example in a burgeoning field that is sure to lead to the identification of apoptotic FEN-1 interacting proteins across many species. The fourth category of FEN-1 interaction partners is involved in HIV replication. FEN-1 interacts with, and is stimulated by, HIV-1 integrase.71 Work has also shown that FEN-1 is seemingly involved in the processing of HIV DNA replication intermediates.72,73 The last two categories of interaction partners are those proteins that post-translationally modify FEN-1 through acetylation or phosphorylation. These proteins and their involvement in FEN-1 function have been discussed above in detail. In summary, although some of the above groups have merely one example so far, it will be of great interest to determine whether FEN-1 interacts with other proteins within these pathways, especially those involved in apoptosis and HIV replication. 3.4. Phenotypes and Biological Functions FEN-1 (RAD27) plays an important role in DNA metabolism, and the loss of it causes severe biological effects.80 In yeast, the deletion of RAD27 leads to temperature sensitivity, hypersensitivity to DNA-alkylating agents such as methyl methanesulfonate (MMS), strong mutator phenotype, and genome instability.66,74,80–86 The majority of the mutations display a typical duplication mutation spectrum resulting from the failure of RNA primer removal during lagging strand DNA synthesis,83 while the rest may result from its role in mismatch repair82 and/or long-patch BER.44 A hallmark of genomic instability is the expansion of variable nucleotide repeat sequences, which is the cause of genetic diseases like Huntington’s disease and myotonic dystrophy.87

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FEN-1 helps prevent trinucleotide repeat (TNR) expansion and contraction, as evidenced by the studies in S. cerevisiae. Phenotypically, the FEN-1 deletion mutant exhibits instability in short DNA repeats, microsatellites and minisatellites, and chromosome loss.66,74,80–86,88,89 Simple repeat sequences or the so-called “atrisk motif sequences (ARMS),” readily form secondary structures, thereby becoming inaccessible to FEN-1 nuclease since the enzyme is thought to require a free 5 -end.90 If this is the case in vivo, then the presence or absence of FEN-1 should not make a difference in the stability of repeat sequences in the genomes.88,91,92 However, the deletion of RAD27 and the presence of the ARMS sequences in the yeast chromosomes show an increase in tri-nucleotide repeat sequence instability by two orders of magnitude. Recent in vitro work from the Topal lab has shown that the delay in FEN-1 addition to a pol β reconstituted replication system results in expansions, while the simultaneous addition of FEN-1 and pol β showed little to no expansion. The expansions shown when FEN-1 addition is delayed are probably due to the ability of the flap structures within repeat sequences to convert into internal loops, which are resistant to FEN1 cleavage and ligatable. Therefore, any delay in flap cleavage due to a secondary structure formation within the flap could lead to an internal loop formation and thus expansion in vivo. Although internal loop structures are resistant to FEN-1 cleavage, work from the Bambara laboratory has suggested that the combined use of BLM helicase activity and FEN and EXO nuclease activities of FEN-1 can remove internal loop structures that can form in trinucleotide repeat sequences.69 In light of the newly discovered GEN activity of FEN-1, it is also possible that FEN-1 in complex with WRN may employ the EXO and GEN activities rather than FEN activity to remove such structures. The importance of FEN-1 in the prevention of oncogenesis has also been studied.93 In mice, a homozygous knockout of FEN-1 is known to be embryonic lethal, consistent with the observation that FEN-1 null mouse blastocysts are arrested in the S phase.94 FEN-1 heterozygous knockout mice are viable and appear to be free of disease. However, FEN-1 heterozygous knockout mice that

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are also heterozygous for the adenomatous polyposis coli (APC) gene develop adenocarcinomas which result in decreased animal survival.93 This result suggested that FEN-1 is a tumor suppressor gene.43 The role of FEN-1 in apoptosis was demonstrated by Parrish et al.46 in C. elegans. The reduction of Crn-1 (C. elegans FEN-1 homologue) activity by RNA interference resulted in resistance to DNA degradation in apoptotic cells in a manner similar to that displayed by cells lacking the mitochondrial endonuclease, CPS6/endonuclease G. From these observations, a model for CRN1/CPS-6 (FEN-1/Endo G)-mediated stepwise DNA degradation during apoptosis was proposed. According to the model, CRN-1 which localizes to nuclei can associate and cooperate with CPS-6 to promote stepwise DNA fragmentation, utilizing the endonuclease activity of CPS-6 and both the 5 –3 -exonuclease activity and a previously uncharacterized GEN activity of CRN-1. Recent evidence suggests that the human immunodeficiency virus type 1 (HIV-1) utilizes FEN-1 endonuclease activity to process its DNA during its replication cycle. The HIV-1 central DNA flap (CDF) is formed during the reverse transcription of HIV-1 RNA genome. The 99 nucleotide long CDF is a substrate for human FEN-1 regardless of the presence of a secondary structure within the CDF.73 Furthermore, HIV-1 integrase has been reported to interact with hFEN-1, and mutual stimulation of their respective activities has been observed. These results suggest that FEN-1 is responsible for processing of the CDF, a critical step in the HIV-1 life cycle. 4. NUCLEASES AND DISEASES As discussed above, DNA repair enzymes play a vital role in protecting the cells from the genotoxic effects of DNA damage. The clinical consequences of errors in DNA metabolism are apparent in several rare human disorders, e.g. xeroderma pigmentosum (XP) and Werner syndrome (WS). Although many of the causative genes of these conditions have been identified, their specific cellular mechanisms are not always clear.

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XPG (xeroderma pigmentosum group G) is a structure-specific endonuclease that makes an incision 3 to the DNA photoproducts and other helix distorting DNA adducts in NER. Defects in XPG (i.e. point mutations) can result in xeroderma pigmentosum (XP), which is characterized by extreme photosensitivity, skin abnormalities in sun-exposed areas, followed by a high incidence of skin cancer and frequent neurological abnormalities. Patients with large truncations in the XPG protein frequently have the features of Cockayne syndrome, which is characterized by growth retardation, mental deficiency and skeletal abnormalities.95 Recently, several polymorphisms have been identified in the XPG gene. These polymorphisms may affect the DNA repair capacity, thereby modulating cancer susceptibility.96,97 Germ-line mutations in the mismatch repair genes, hMSH2, hMSH6, hMLH1 and hPMS2, have been found in families with hereditary nonpolyposis colorectal cancer (HNPCC). However, some HNPCC families fail to exhibit mutations in the known MMR genes, suggesting that some other component of the MMR pathway may be responsible. Mutations in EXO1, an exonuclease known to cleave both 3 and 5 of the mismatch, have been found in patients with atypical HNPCC, suggesting a possible role of EXO1 as a cancer predisposing gene.98,99 In a recent report,100 a gene-environment interaction between polymorphism in APE1 and XRCC1 genes and cigarette smoking with regard to lung cancer risk was shown. The results suggest that APE1 Asp148Glu and XRCC1 Ard399Gln polymorphism might modify the risk of lung cancer attributable to cigarette smoke exposure. WRN is distinguished from other members of the family of RecQ helicases by the presence of an N-terminal 3 –5 -exonuclease domain. Werner syndrome (WS) is a rare autosomal recessive disorder characterized by features of premature ageing. Individuals with WS manifest the early onset of alopecia, cataracts, atherosclerosis and osteoporosis. Additionally they are predisposed to developing diabetes and cancer.25

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Nucleases are important in the understanding of human diseases, as they are the key enzymes in DNA replication, repair and recombination pathways. Some nucleases like FEN-1 are multifunctional with multiple roles. Genome stability is dependent on numerous DNA metabolic proteins, which ensure that replication, repair and recombination occur with high fidelity. Although most of the proteins involved in DNA dynamics and metabolism have been identified, how they are regulated and coordinated in these processes remain unclear. The functional or regulatory loss of DNA metabolism proteins has been linked to several human diseases, including cancers, premature ageing syndromes, Huntington’s disease, Friederich’s ataxia and myotonic dystrophy. Although the causes of certain DNA metabolic diseases have been identified, a detailed understanding of the molecular pathology is lacking, because many of these diseases are caused by defects in proteins that have multiple functions. Clearly, more studies are necessary to further understand precisely how the loss of multi-task nucleases is manifested in diseases. 5. SUMMARY/PERSPECTIVES The significance of FEN-1 nuclease cannot be overstated considering that it has evolved three distinct biochemical activities to carry out its multiple biological roles. FEN-1 nucleases are unique in their requirement for specific DNA structures rather than DNA sequence to target their three activities. In addition to the regulation of FEN-1 activity by a DNA substrate structure, post-translational modifications and protein-protein interactions increase the precision with which FEN-1 functions in vivo. As FEN-1 is required for genome stability it can therefore be considered a tumor suppressor gene. The onset of tumorigenesis is rooted in the accumulation of genetic aberrations, such as point mutations, frame shifts, chromosomal defects, microsatellite expansions and contractions. The sequential progression from normal to malignant cells is accompanied by the increase of such alterations in cells and continues within the heterogeneic

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tumor cell population during outgrowth. The mechanisms by which these errors accumulate are thought to be due in part to the damage acquired by the genes involved in genomic stability. This instability would result in further mutations to the regulatory genes. The involvement of FEN-1 in controlling genomic stability through multiple metabolic actions suggests that the functional loss of FEN-1 through mutation would facilitate further tumor mutagenesis. Polymorphisms are present throughout the human genome and are common in the human population. Single nucleotide polymorphisms (SNPs) in FEN-1 have been recently found in the human population (http://www.ncbi.nlm.nih.gov/SNP/snp_ref.cgi?locusId = 2237 & choose Rs = all). We believe that studying the effects of these SNPs on FEN-1 and other DNA repair, replication and recombination nucleases may correlate with the differences in individual susceptibilities to diseases. These systematic studies would provide a convincing model for diseases caused by various defects of the multi-functional nucleases. Acknowledgments The work in the Shen laboratory is supported by the grant R01CA073764 from the National Institutes of Health. References 1. Tsubouchi H, Ogawa H. (2000) Exo1 roles for repair of DNA doublestrand breaks and meiotic crossing over in Saccharomyces cerevisiae. Mol Biol Cell 11: 2221–2233. 2. Genschel J, Bazemore LR, Modrich P. (2002) Human exonuclease I is required for 5 and 3 mismatch repair. J Biol Chem 277: 13302–13311. 3. Dzantiev L, Constantin N, Genschel J, et al. (2004) A defined human system that supports bidirectional mismatch-provoked excision. Mol Cell 15: 31–41. 4. Evans AR, Limp-Foster M, Kelley MR. (2000) Going APE over ref-1. Mutat Res 461: 83–108. 5. Chou KM, Cheng YC. (2003) The exonuclease activity of human apurinic/apyrimidinic endonuclease (APE1). Biochemical properties

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6.

7.

8.

9. 10.

11.

12.

13.

14.

15.

16.

17.

ch10

293

and inhibition by the natural dinucleotide Gp4G. J Biol Chem 278: 18289–18296. Johnson RE, Torres-Ramos CA, Izumi T, et al. (1998) Identification of APN2, the Saccharomyces cerevisiae homolog of the major human AP endonuclease HAP1, and its role in the repair of abasic sites. Genes Dev 12: 3137–3143. Unk I, Haracska L, Prakash S, Prakash L. (2001) 3 -phosphodiesterase and 3 –5 exonuclease activities of yeast Apn2 protein and requirement of these activities for repair of oxidative DNA damage. Mol Cell Biol 21: 1656–1661. O’Donovan A, Davies AA, Moggs JG, et al. (1994) XPG endonuclease makes the 3 -incision in human DNA nucleotide excision repair. Nature 371: 432–435. Mu D, Hsu DS, Sancar A. (1996) Reaction mechanism of human DNA repair excision nuclease. J Biol Chem 271: 8285–8294. Matsunaga T, Mu D, Park CH, et al. (1995) Human DNA repair excision nuclease. Analysis of the roles of the subunits involved in dual incisions by using anti-XPG and anti-ERCC1 antibodies. J Biol Chem 270: 20862–20869. Lieber MR. (1997) The FEN-1 family of structure-specific nucleases in eukaryotic DNA replication, recombination and repair. Bioessays 19: 233–240. Ayyagari R, Gomes XV, Gordenin DA, Burgers PM. (2003) Okazaki fragment maturation in yeast. I. Distribution of functions between FEN1 AND DNA2. J Biol Chem 278: 1618–1625. D’Amours D, Jackson SP. (2002) The MRE11 complex: at the crossroads of DNA repair and checkpoint signalling. Nat Rev Mol Cell Biol 3: 317– 327. Fabre F, Chan A, Heyer WD, Gangloff S. (2002) Alternate pathways involving Sgs1/Top3, Mus81/ Mms4, and Srs2 prevent formation of toxic recombination intermediates from single-stranded gaps created by DNA replication. Proc Natl Acad Sci USA 99: 16887–16892. Whitby MC, Osman F, Dixon J. (2003) Cleavage of model replication forks by fission yeast Mus81-Eme1 and budding yeast Mus81-Mms4. J Biol Chem 278: 6928–6935. Moshous D, Callebaut I, de Chasseval R, et al. (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. Ma Y, Schwarz K, Lieber MR. (2005) The Artemis: DNA-PKcs endonuclease cleaves DNA loops, flaps, and gaps. DNA Repair (Amst) 4: 845– 851.

FA

November 2, 2006 9:2

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WSPC/SPI-B414: DNA Repair, Genetic Instability & Cancer

ch10

Purnima Singh and Binghui Shen

18. Tran HT, Gordenin DA, Resnick MA. (1999) The 3 -> 5 -exonucleases of DNA polymerases delta and epsilon and the 5 -> 3 -exonuclease Exo1 have major roles in postreplication mutation avoidance in Saccharomyces cerevisiae. Mol Cell Biol 19: 2000–2007. 19. Hubscher U, Maga G, Spadari S. (2002) Eukaryotic DNA polymerases. Annu Rev Biochem 71: 133–163. 20. Ropp PA, Copeland WC. (1996) Cloning and characterization of the human mitochondrial DNA polymerase, DNA polymerase gamma. Genomics 36: 449–458. 21. Mazur DJ, Perrino FW. (1999) Identification and expression of the TREX1 and TREX2 cDNA sequences encoding mammalian 3 -> 5 exonucleases. J Biol Chem 274: 19655–19660. 22. Mazur DJ, Perrino FW. (2001) Excision of 3 -termini by the Trex1 and TREX2 3 -> 5 -exonucleases. Characterization of the recombinant proteins. J Biol Chem 276: 17022–17029. 23. Bae SH, Bae KH, Kim JA, Seo YS. (2001) RPA governs endonuclease switching during processing of Okazaki fragments in eukaryotes. Nature 412: 456–461. 24. Bae SH, Choi E, Lee KH, et al. (1998) DNA2 of Saccharomyces cerevisiae possesses a single-stranded DNA-specific endonuclease activity that is able to act on double-stranded DNA in the presence of ATP. J Biol Chem 273: 26880–26890. 25. Huang S, Li B, Gray MD, et al. (1998) The premature ageing syndrome protein, WRN, is a 3 -> 5 -exonuclease. Nat Genet 20: 114–116. 26. Harrigan JA, Wilson DM, III, Prasad R, et al. (2006) The Werner syndrome protein operates in base excision repair and co-operates with DNA polymerase beta. Nucl Acids Res 34: 745–754. 27. Huang P. (1998) Excision of mismatched nucleotides from DNA: a potential mechanism for enhancing DNA replication fidelity by the wild-type p53 protein. Oncogene 17: 261–270. 28. Hanawalt PC. (1994) Transcription-coupled repair and human disease. Science 266: 1957–1958. 29. Paull TT, Gellert M. (2000) A mechanistic basis for MRE11-directed DNA joining at microhomologies. Proc Natl Acad Sci USA 97: 6409– 6414. 30. Ma Y, Pannicke U, Schwarz K, Lieber MR. (2002) Hairpin opening and overhang processing by an Artemis/DNA-dependent protein kinase complex in non-homologous end joining and V(D)J recombination. Cell 108: 781–794. 31. Wu X, Wilson TE, Lieber MR. (1999) A role for FEN-1 in nonhomologous DNA end joining: the order of strand annealing and nucleolytic processing events. Proc Natl Acad Sci USA 96: 1303–1308.

FA

November 2, 2006 9:2

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ch10

295

32. Hoss M, Robins P, Naven TJ, et al. (1999) A human DNA editing enzyme homologous to the Escherichia coli DnaQ/MutD protein. EMBO J 18: 3868–3875. 33. Li B, Navarro S, Kasahara N, Comai L. (2004) Identification and biochemical characterization of a Werner’s syndrome protein complex with Ku70/80 and poly(ADP-ribose) polymerase-1. J Biol Chem 279: 13659–13667. 34. Mummenbrauer T, Janus F, Muller B, et al. (1996) p53 protein exhibits 3 - to -5 exonuclease activity. Cell 85: 1089–1099. 35. Bambara RA, Murante RS, Henricksen LA. (1997) Enzymes and reactions at the eukaryotic DNA replication fork. J Biol Chem 272: 4647–4650. 36. Chapados BR, Hosfield DJ, Han S, et al. (2004) Structural basis for FEN-1 substrate specificity and PCNA-mediated activation in DNA replication and repair. Cell 116: 39–50. 37. Harrington JJ, Lieber MR. (1994) Functional domains within FEN-1 and RAD2 define a family of structure- specific endonucleases: implications for nucleotide excision repair. Genes Dev 8: 1344–1355. 38. Liu Y, Kao HI, Bambara RA. (2004) Flap endonuclease-1: a central component of DNA metabolism. Annu Rev Biochem 73: 589–615. 39. Murante RS, Huang L, Turchi JJ, Bambara RA. (1994) The calf 5 - to 3 -exonuclease is also an endonuclease with both activities dependent on primers annealed upstream of the point of cleavage. J Biol Chem 269: 1191–1196. 40. Kaiser MW, Lyamicheva N, Ma W, et al. (1999) A comparison of eubacterial and archaeal structure-specific 5 -exonucleases. J Biol Chem 274: 21387–21394. 41. Storici F, Henneke G, Ferrari E, et al. (2002) The flexible loop of human FEN1 endonuclease is required for flap cleavage during DNA replication and repair. EMBO J 21: 5930–5942. 42. Kao HI, Henricksen LA, Liu Y, Bambara RA. (2002) Cleavage specificity of Saccharomyces cerevisiae flap endonuclease-1 suggests a double-flap structure as the cellular substrate. J Biol Chem 277: 14379–14389. 43. Friedrich-Heineken E, Henneke G, Ferrari E, Hubscher U. (2003) The acetylatable lysines of human Fen1 are important for endo- and exonuclease activities. J Mol Biol 328: 73–84. 44. Huggins CF, Chafin DR, Aoyagi S, et al. (2002) Flap endonuclease-1 efficiently cleaves base excision repair and DNA replication intermediates assembled into nucleosomes. Mol Cell 10: 1201–1211. 45. Lindahl T. (1971) The action pattern of mammalian deoxyribonuclease IV. Eur J Biochem 18: 415–421.

FA

November 2, 2006 9:2

296

WSPC/SPI-B414: DNA Repair, Genetic Instability & Cancer

ch10

Purnima Singh and Binghui Shen

46. Parrish JZ, Yang C, Shen B, Xue D. (2003) CRN-1, a Caenorhabditis elegans FEN-1 homologue, co-operates with CPS-6/EndoG to promote apoptotic DNA degradation. EMBO J 22: 3451–3460. 47. Shen B, Singh P, Liu R, et al. (2005) Multiple but dissectible functions of FEN-1 nucleases in nucleic acid processing, genome stability and diseases. Bioessays 27: 717–729. 48. Vispe S, Ho EL, Yung TM, Satoh MS. (2003) Double-strand DNA break formation mediated by flap endonuclease-1. J Biol Chem 278: 35279– 35285. 49. Shen B, Nolan JP, Sklar LA, Park MS. (1997) Functional analysis of point mutations in human flap endonuclease-1 active site. Nucl Acids Res 25: 3332–3338. 50. Hosfield DJ, Frank G, Weng Y, et al. (1998) Newly discovered archaebacterial flap endonucleases show a structure-specific mechanism for DNA substrate binding and catalysis resembling human flap endonuclease-1. J Biol Chem 273: 27154–27161. 51. Hwang KY, Baek K, Kim HY, Cho Y. (1998) The crystal structure of flap endonuclease-1 from Methanococcus jannaschii. Nat Struct Biol 5: 707–713. 52. Matsui E, Musti KV, Abe J, et al. (2002) Molecular structure and novel DNA binding sites located in loops of flap endonuclease-1 from Pyrococcus horikoshii. J Biol Chem 277: 37840–37847. 53. Ceska TA, Sayers JR, Stier G, Suck D. (1996) A helical arch allowing single-stranded DNA to thread through T5 5 -exonuclease. Nature 382: 90–93. 54. Bornarth CJ, Ranalli TA, Henricksen LA, et al. (1999) Effect of flap modifications on human FEN1 cleavage. Biochemistry 38: 13347–13354. 55. Hasan S, Stucki M, Hassa PO, et al. (2001) Regulation of human flap endonuclease-1 activity by acetylation through the transcriptional coactivator p300. Mol Cell 7: 1221–1231. 56. Gary R, Ludwig DL, Cornelius HL, et al. (1997) The DNA repair endonuclease XPG binds to proliferating cell nuclear antigen (PCNA) and shares sequence elements with the PCNA-binding regions of FEN-1 and cyclin-dependent kinase inhibitor p21. J Biol Chem 272: 24522–24529. 57. Warbrick E, Lane DP, Glover DM, Cox LS. (1997) Homologous regions of Fen1 and p21Cip1 compete for binding to the same site on PCNA: a potential mechanism to co-ordinate DNA replication and repair. Oncogene 14: 2313–2321. 58. Chai Q, Qiu J, Chapados BR, Shen B. (2001) Archaeoglobus fulgidus RNase HII in DNA replication: enzymological functions and activity

FA

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59. 60.

61.

62.

63.

64.

65.

66.

67.

68.

69.

70.

71.

ch10

297

regulation via metal cofactors. Biochem Biophys Res Commun 286: 1073– 1081. Arezi B, Kuchta RD. (2000) Eukaryotic DNA primase. Trends Biochem Sci 25: 572–576. Kang HY, Choi E, Bae SH, et al. (2000) Genetic analyses of Schizosaccharomyces pombe DNA2(+) reveal that DNA2 plays an essential role in Okazaki fragment metabolism. Genetics 155: 1055–1067. Jin YH, Ayyagari R, Resnick MA, et al. (2003) Okazaki fragment maturation in yeast II. Cooperation between the polymerase and 3 -5 exonuclease activities of Pol delta in the creation of a ligatable nick. J Biol Chem 278: 1626–1633. Brosh RM, Jr., von Kobbe C, Sommers JA, et al. (2001) Werner syndrome protein interacts with human flap endonuclease-1 and stimulates its cleavage activity. EMBO J 20: 5791–5801. Sharma S, Otterlei M, Sommers JA, et al. (2004) WRN helicase and FEN-1 form a complex upon replication arrest and together process branch-migrating DNA structures associated with the replication fork. Mol Biol Cell 15: 734–750. Imamura O, Campbell JL. (2003) The human Bloom syndrome gene suppresses the DNA replication and repair defects of yeast DNA2 mutants. Proc Natl Acad Sci USA 100: 8193–8198. Prasad R, Dianov GL, Bohr VA, Wilson SH. (2000) FEN1 stimulation of DNA polymerase beta mediates an excision step in mammalian long-patch base excision repair. J Biol Chem 275: 4460–4466. Gary R, Kim K, Cornelius HL, et al. (1999) Proliferating cell nuclear antigen facilitates excision in long-patch base excision repair. J Biol Chem 274: 4354–4363. Ahn B, Harrigan JA, Indig FE, et al. (2004) Regulation of WRN helicase activity in human base excision repair. J Biol Chem 279: 53465–53474. Parrilla-Castellar ER, Arlander SJ, Karnitz L. (2004) Dial 9–1-1 for DNA damage: the Rad9-Hus1-Rad1 (9–1-1) clamp complex. DNA Repair (Amst) 3: 1009–1014. Wang W, Bambara RA. (2004) Human bloom protein stimulates flap endonuclease-1 activity by resolving DNA secondary structure. J Biol Chem 280: 5391–5399. Tseng HM, Tomkinson AE. (2004) Processing and joining of DNA ends co-ordinated by interactions among Dnl4/Lif1, Pol4 and FEN-1. J Biol Chem 279: 47580–47588. Faust EA, Triller H. (2002) Stimulation of human flap endonuclease1 by human immunodeficiency virus type 1 integrase: possible role

FA

November 2, 2006 9:2

298

72.

73.

74.

75.

76.

77.

78.

79.

80.

81.

82.

WSPC/SPI-B414: DNA Repair, Genetic Instability & Cancer

ch10

Purnima Singh and Binghui Shen

for flap endonuclease-1 in 5 -end processing of human immunodeficiency virus type 1 integration intermediates. J Biomed Sci 9: 273–287. Brin E, Yi J, Skalka AM, Leis J. (2000) Modeling the late steps in HIV1 retroviral integrase-catalyzed DNA integration. J Biol Chem 275: 39287–39295. Rumbaugh JA, Fuentes GM, Bambara RA. (1998) Processing of an HIV replication intermediate by the human DNA replication enzyme FEN1. J Biol Chem 273: 28740–28745. Barnes CJ, Wahl AF, Shen B, et al. (1996) Mechanism of tracking and cleavage of adduct-damaged DNA substrates by the mammalian 5 - to 3 -exonuclease/endonuclease RAD2 homologue 1 or flap endonuclease-1. J Biol Chem 271: 29624–29631. Biswas EE, Zhu FX, Biswas SB. (1997) Stimulation of RTH1 nuclease of the yeast Saccharomyces cerevisiae by replication protein A. Biochemistry 36: 5955–5962. Maga G, Villani G, Tillement V, et al. (2001) Okazaki fragment processing: modulation of the strand displacement activity of DNA polymerase delta by the concerted action of replication protein A, proliferating cell nuclear antigen, and flap endonuclease-1. Proc Natl Acad Sci USA 98: 14298–14303. Chai Q, Zheng L, Zhou M, et al. (2003) Interaction and stimulation of human FEN-1 nuclease activities by heterogeneous nuclear ribonucleoprotein A1 in alpha-segment processing during Okazaki fragment maturation. Biochemistry 42B: 15045–15052. Siegal G, Turchi JJ, Myers TW, Bambara RA. (1992) A 5 -3 -exonuclease functionally interacts with calf DNA polymerase epsilon. Proc Natl Acad Sci USA 89: 9377–9381. Budd ME, Campbell JL. (1997) Ayeast replicative helicase, DNA2 helicase, interacts with yeast FEN-1 nuclease in carrying out its essential function. Mol Cell Biol 17: 2136–2142. Reagan MS, Pittenger C, Siede W, Friedberg EC. (1995) Characterization of a mutant strain of Saccharomyces cerevisiae with a deletion of the RAD27 gene, a structural homolog of the RAD2 nucleotide excision repair gene. J Bacteriol 177: 364–371. Murray JM, Tavassoli M, al-Harithy R, et al. (1994) Structural and functional conservation of the human homolog of the Schizosaccharomyces pombe rad2 gene, which is required for chromosome segregation and recovery from DNA damage. Mol Cell Biol 14: 4878–4888. Johnson RE, Kovvali GK, Prakash L, Prakash S. (1995) Requirement of the yeast RTH1 5 -3 -exonuclease for the stability of simple repetitive DNA. Science 269: 238–240.

FA

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WSPC/SPI-B414: DNA Repair, Genetic Instability & Cancer

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83. Tishkoff DX, Filosi N, Gaida GM, Kolodner RD. (1997) A novel mutation avoidance mechanism dependent on S. cerevisiae RAD27 is distinct from DNA mismatch repair. Cell 88: 253–263. 84. Freudenreich CH, Kantrow SM, Zakian VA. (1998) Expansion and length-dependent fragility of CTG repeats in yeast. Science 279: 853– 856. 85. Schweitzer JK, Livingston DM. (1998) Expansions of CAG repeat tracts are frequent in a yeast mutant defective in Okazaki fragment maturation. Hum Mol Genet 7: 69–74. 86. Negritto MC, Qiu J, Ratay DO, et al. (2001) Novel function of Rad27 (FEN-1) in restricting short-sequence recombination. Mol Cell Biol 21: 2349–2358. 87. Pearson CE, Tam M, Wang YH, et al. (2002) Slipped-strand DNAs formed by long (CAG)*(CTG) repeats: slipped-out repeats and slipout junctions. Nucl Acids Res 30: 4534–4547. 88. Gordenin DA, Kunkel TA, Resnick MA. (1997) Repeat expansion — all in a flap? Nat Genet 16: 116–118. 89. Sato A, Kanai A, Itaya M, Tomita M. (2003) Co-operative regulation for Okazaki fragment processing by RNase HII and FEN-1 purified from a hyperthermophilic archaeon, Pyrococcus furiosus. Biochem Biophys Res Commun 309: 247–252. 90. Murante RS, Rust L, Bambara RA. (1995) Calf 5 -3 -exo-endonuclease must slide from a 5 end of the substrate to perform structure-specific cleavage. J Biol Chem 270: 30377–30383. 91. Spiro C, Pelletier R, Rolfsmeier ML, et al. (1999) Inhibition of FEN-1 processing by DNA secondary structure at trinucleotide repeats. Mol Cell 4: 1079–1085. 92. Henricksen LA, Tom S, Liu Y, Bambara RA. (2000) Inhibition of flap endonuclease-1 by flap secondary structure and relevance to repeat sequence expansion. J Biol Chem 275: 16420–16427. 93. Kucherlapati M, Yang K, Kuraguchi M, et al. (2002) Haploinsufficiency of flap endonuclease (Fen1) leads to rapid tumor progression. Proc Natl Acad Sci USA 99: 9924–9929. 94. Larsen E, Gran C, Saether BE, et al. (2003) Proliferation failure and gamma radiation sensitivity of Fen1 null mutant mice at the blastocyst stage. Mol Cell Biol 23: 5346–5353. 95. Nouspikel T, Lalle P, Leadon SA, et al. (1997) A common mutational pattern in Cockayne syndrome patients from xeroderma pigmentosum group G: implications for a second XPG function. Proc Natl Acad Sci USA 94: 3116–3121. 96. Emmert S, Slor H, Busch DB, et al. (2002) Relationship of neurologic degeneration to genotype in three xeroderma pigmentosum group G patients. J Invest Dermatol 118: 972–982.

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97. Jeon HS, Kim KM, Park SH, et al. (2003) Relationship between XPG codon 1104 polymorphism and risk of primary lung cancer. Carcinogenesis 24: 1677–1681. 98. Wu Y, Berends MJ, Post JG, et al. (2001) Germ-line mutations of EXO1 gene in patients with hereditary nonpolyposis colorectal cancer (HNPCC) and atypical HNPCC forms. Gastroenterology 120: 1580– 1587. 99. Sun X, Zheng L, Shen B. (2002) Functional alterations of human exonuclease 1 mutants identified in atypical hereditary nonpolyposis colorectal cancer syndrome. Cancer Res 62: 6026–6030. 100. Ito H, Matsuo K, Hamajima N, et al. (2004) Gene-environment interactions between the smoking habit and polymorphisms in the DNA repair genes, APE1 Asp148Glu and XRCC1 Arg399Gln, in Japanese lung cancer risk. Carcinogenesis 25: 1395–1401.

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CHAPTER 11

Integrative Genomics and Epigenomics: Application in Cancer Research Maxwell P. Lee∗

ABSTRACT Genome sequence information from human as well as model organisms together with the advancement in high-throughput technology have transformed traditional biology researches into a new era of large-scale systems biology. Systems biology aims to provide a system-level understanding of biology which includes genome organization, epigenome, transcriptome and proteome. Information from these “-omics” researches can be used to build biological and mathematical models to describe biological phenomena and to give predictive values. In this review, the author provides an overview of the recent progress made in genomics and epigenomics, including a summary of some of the recent studies in this field carried out by the author and colleagues.

1. INTRODUCTION There is no agreed simple definition of systems biology. Systems biology is generally considered as a field that aims to understand biology at a system-level. This usually requires the use of high-throughput technology to obtain measurements of biological phenomena on a large-scale. The most common platforms of highthroughput assays are the various forms of microarray technologies, ∗ Laboratory

of Population Genetics, National Cancer Institute, 41 Library Dr. D702C, Bethesda, MD20892, USA. Tel.: 301-435-1536; Fax: 301-402-9325; E-mail: [email protected] 301

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which define the fields of genomics, epigenomics, proteomics, etc. These “-omics” studies produce the components for the assembly of the systems. Systems biology aims to build predictive models using well established principles and methods in mathematics and engineering to understand biology. It is clear from this working definition that systems biology involves integrative researches in many fields, including biology, mathematics, statistics and engineering. It is not possible to cover the entire field of systems biology in a chapter. In this chapter, the author reviews some aspects of the systems biology researches that are related to genome and epigenome with a focus on integrated genetic and epigenetic studies, and presents some of our recent studies in the field.

2. EPIGENOMICS The application of genomics in epigenetic research defines a new field “epigenomics”.1–3 Epigenetics studies any molecular process that regulates gene expression without DNA sequence changes.4,5 Common epigenetic alterations include DNA methylation6 and post-translational modifications of histones.7 Genomic imprinting is a special case of the epigenetic phenomena in which epigenetic modifications differ between the two parental chromosomes, leading to a preferential expression of one parental allele.8–10 X-chromosome inactivation provides another case of epigenetic regulation of gene expression, resulting in a preferential silencing of one parental allele.11 Genomic imprinting plays important roles in hereditary and sporadic cancers. Beckwith-Wiedemann syndrome (BWS) is a hereditary cancer syndrome. Both genetic and epigenetic alterations in several imprinted genes contribute to the cancer and the over-growth phenotypes of BWS. The loss of imprinting (LOI) of IGF2 is the most common molecular event observed in human cancers.10 LOI of IGF2, which is associated with an increase in the gene expression of IGF2, a growth factor, is also coupled with loss of expression of CDKN1C, a cell cycle kinase inhibitor.

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Given the importance of genomic imprinting in cancer biology, we set out to identify the general imprinting signature by a computational analysis of DNA sequences.12 Approximately 50 imprinted genes were identified from the human and mouse genomes at the time when the author and colleagues, started the sequence analysis. The list of the imprinted genes can be found at the web site http://www.geneimprint.com/. Because the regulatory elements are usually conserved, they could be identified in chromosomal regions that are conserved between mouse and human. The author and colleagues identified 16 motifs. Based on the distribution of the motifs in 24 imprinted and 128 non-imprinted genes, a logistic regression model was developed to distinguish imprinted genes from non-imprinted genes. Their model correctly assigned 126 out of 128 non-imprinted genes and 23 out of 24 imprinted genes in the training dataset. The accuracy, sensitivity and specificity rates of the model are 98%, 96% and 98%, respectively. The methylation of CpG islands in the promoter regions of the tumor suppressor genes is often associated with human cancers. CpG islands are DNA fragments that are usually 200-bp to 1-kb in length, which have a CG content greater than 55% and the CpG(observed)/CpG(expected) greater than 0.65.13 The annotation of CpG islands for the entire human genome is available at UCSC (http://genome.ucsc.edu/). Imprinted genes are often associated with differentially methylated CpG islands. Restriction landmark genomic scanning (RLGS) was developed to analyze the methylation of DNA, and it has the capacity to identify thousands of DNA fragments modified by DNA methylation. RLGS was used to identify several imprinted genes, including an U2af binding protein14 and Grf1 encoding a RAS-specific guanine nucleotide exchange factor.15 Strichman-Almashanu et al. developed an assay to isolate methylated CpG islands with methylation-sensitive restriction enzyme digestion.16 Two of the 30 methylated CpG islands that they have identified are associated with imprinted genes. More recently, methylation analysis has shifted toward genomewide studies using high-throughput technology, including CpG

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island array,17,18 BAC clone array coupled with methylationsensitive Not I digestion,19 DNA microarray coupled with immunocapturing with the antibody targeting at cytosine methylation,20 Illumina’s BeadArray platform to analyze CpG site methylation,21 and tiling array for the promoter region methylation.22,23 Gene expression is controlled by distinct chromatin structures. This epigenetic information is largely controlled by specific histone modifications, which have been observed in the active or silent states of transcription at various loci, including immunoglobulin genes, imprinted genes, olfactory genes and genes subject to X inactivation.24 Many studies on specific histone modifications at tumor suppressor genes, immunoglobulin genes, imprinted genes, and genes associated with X inactivation have revealed a common regulatory mechanism of histone modifications on transcription regulation. Histone H3 lysine 4 methylation and lysine 9/14 acetylation were associated with active chromatin, whereas histone H3 lysine 9 and lysine 27 were associated with inactive chromatin. Most of these studies were done on selected loci and only a few recent studies were carried out to characterize chromatin on a genomewide level using ChIP-on-chip experiments, including genome tiling array for human chromosome 21 and 22 to probe histone H3 lysine 4 di- and tri-methylation and lysine 9/14 acetylation25 ; tiling array for probing RNA polymerase II preinitiation complex in the promoter regions26 ; cDNA array to probe histone lysine methylation27 ; and a sequence tag method called genome-wide mapping technique (GMAT) to probe lysine acetylation of histone H3.28 3. TRANSCRIPTOME AND GENE EXPRESSION REGULATION The completion of the human genome sequence coupled with the high-throughput genomics-based global gene expression profiling technologies enables the evaluation of entire cellular transcriptome in the human genome and the networks of interactions between the genome and genes. Affymetrix DNA oligonucleotide microarray and cDNA microarray are the two major platforms for analyzing

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global transcription and gene regulatory networks. There have been many comprehensive reviews on whole-genome gene expression profiling with microarray technology.29–31 I will discuss several studies in the context of integrating gene expression profiling with pathways. Cancer is a complex disease that results from mutations in the tumor suppressor genes, oncogene, and DNA repair genes as well as epigenetic alteration and global gene expression changes. Understanding the latent factors that reflect the coordinated gene expression pattern may provide a better basis for the diagnosis and treatment of human cancers. Using human primary mammary epithelial cell cultures (HMECs) to express a specific oncogene, Bild et al. developed a series of pathway signatures by evaluating gene expression profiles using Affymetrix U133 chips, analyzing the data with the principal component analysis.32 They found these oncogenic signatures to be robust, and are able to classify the subtypes of human cancers to predict their outcomes. Signature scores in cancer cell lines can also predict their sensitivity to therapeutic agents targeted at the pathway. The author and colleagues have been systematically studying the microarray gene expression data. The coordinated expression of genes in a single pathway is a common phenomenon. They have combined the microarray gene expression data with the biological knowledge of pathways, metabolic pathways (KEGG) and signal transduction pathways (BioCarta), to study the correlation of gene expression for genes in these pathways.33 They defined a coherence indicator, which is the ratio of the number of gene pairs that show a significant correlation in expression to the total number of gene pairs in the pathway, to measure the level of coordinated gene expression in the pathway. They found a high coherence indicator of KEGG pathways and a low coherence indicator of BioCarta pathways in the tumors, which suggest an increased metabolism rate in tumor cells and a disruption of some of the signal transduction pathways in cancers. To study the interaction between a transcription start site and a translational initiation site, the author and colleagues analyzed

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systematically the sequence in the promoter regions of the human genome.34 They have defined a novel core promoter subclass that is devoid of the ATG translation initiation codon within approximately 500 bp of the major transcription start site. This feature, which they termed an “ATG desert,” occurs frequently and non-randomly throughout the genome. The presence of the “ATG desert” element is necessary to suppress aberrant transcripts that may produce cryptic proteins that can be translated if ATG codon is present. This study is a good example of how cells integrate transcription and translation regulation via a sequence element. 4. ALLELE-SPECIFIC GENE EXPRESSION A biological role has been identified for differential allelic gene expression that is associated with X-chromosome inactivation and genomic imprinting. In both cases, gene expression is from one chromosome only, whereas transcription from the other chromosome is completely silenced. In other words, allelic gene expression is binary; it is either “on” or “off” for each chromosome (Fig. 1A). However, prior to the studies by the author and colleagues, no large-scale analysis of differential allelic expression of human genes had been carried out. Hence the role played by allelic gene expression variation in processes other than X-chromosome inactivation and genomic imprinting remains unanswered. The author and colleagues are particularly interested in the quantitative difference in allelic gene expression between the two chromosomes as depicted in Fig. 1B. In this case, both alleles are expressed, but to different degrees. To address the role of quantitative allelic difference in gene expression between the two chromosomes, they developed novel experimental strategies to study epigenetic regulation at the genome-wide level. The HuSNP microarray was designed to simultaneously type 1,494 SNPs in the human genome. To study gene expression variation globally, they reengineered the assay protocol and software of the Affymetrix HuSNP chip so that it measured allele-specific transcript variation.35 This is an example of integrating genotyping with gene expression

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B Active

Partial active

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Fig. 1 Epigenetic phenomena such as chromosome inactivation and genomic imprinting are illustrated in part A. T and C within the transcripts indicate the SNP used to evaluate allele-specific gene expression. H3K9-Me refers to lysine 9 methylation of histone H3, which serves as an example of inactive chromatin. H3-Ac refers to the acetylation of histone H3, which serves as an example of active chromatin. 5m C refers to DNA methylation at cytosine, which is often associated with inactive chromatin. The quantitative allelic difference in gene expression between the two chromosomes is illustrated in part B. Note that the T allele is transcribed 2-fold more than the C allele.

analysis. For the assay to be feasible, two prerequisites were necessary: 1) the SNPs on the chip had to be located within exons; and 2) the system had to measure allele-specific expression quantitatively. The author and colleagues determined that 1,063 of the SNPs were located within the exons. To address the second requirement, they developed a computational method to extract the fluorescent intensity for each probe from an Affymetrix data file and quantify the ratio of expression of the two alleles. To assess the precision of the system, they performed experiments in duplicate for both genomic DNA and polyA RNA from three fetuses. The correlation between these duplicated experiments was very high, with average Pearson correlation coefficients of 0.98 (p < 0.001) for genomic DNA and 0.95

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(p < 0.001) for RNA. They then performed genotyping and allelespecific gene expression in the kidney and liver taken from seven fetuses. Genotype cells were obtained using the Affymetrix MAS5 software, and quantitative allele-specific gene expression was computed using their novel methodology. To be included in their analysis, each SNP had to meet the following criteria: 1) at least one fetus was heterozygous for the SNP; 2) the SNP was among the 1,063 mapped within a transcribed region; and 3) the gene containing the SNP was expressed in the kidney or liver. They found that 603 SNPs met all the three criteria, and 326 (54%) of these showed preferential expression of one allele with > 2-fold difference in at least one sample; and for 170 genes, there was > 4-fold difference in expression between the two alleles. Four of these 170 genes were imprinted, (i.e. SNPRN, IPW, HTR2A and PEG3). Thus these findings validated their approach of adapting the HuSNP chip to study allele-specific gene expression. The majority of the genes that showed preferential expression of one allele were scattered on different chromosomes, suggesting that allele-specific expression was not limited to imprinted domains. Recent studies from many laboratories have confirmed these observations.36–39 Pant et al. using an oligonucleotide array, reported allelic variation in gene expression for 731 out of 1,389 genes.40 Studies from the author’s lab and others suggest that allelespecific gene expression regulation may play an important role in inter-individual steady state mRNA levels and the inter-individual phenotypic differences that result from variability in gene expression across populations. Allelic variation in gene expression could provide a quantitative representation of hypomorphic mutations. To gain an insight into the biological function of allelic variation in gene expression, the author and colleagues started to characterize allelic variation in different individuals, and in multiple tissues of the same individual.37 They found that allelic variation in an individual’s multiple tissues always preferentially expressed the same allele. The preferentially expressed allele, however, varied between different individuals, suggesting a dependence on genomic context.

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Both relative expression of the two alleles and the frequency of allelic variation in the samples also varied. To complement the experimental studies of allele-specific gene expression, the author and colleagues also developed computational methods to identify the genes that preferentially expressed one allele. Their lab developed two computational methods to identify differential allelic gene expression using EST sequences in UniGene. To model the allele-specific gene expression, it was assumed that each cDNA library represented an individual and all the libraries constituted a population. If both alleles of an SNP in a gene were present in a cDNA library, the individual was heterozygous for the SNP. If only allele A was detected in the cDNA library, the genotype for that individual could be either AAor AB. The probability of AAor AB genotype could be inferred using Bayes’ rule. Genotype frequencies, PAA , PAB , and PBB , were then estimated from individual library genotypes. The allele frequency in the population was calculated as QA = PAA + 0.5PAB and QB = 1 − QA . The expected heterozygote frequency, based on the Hardy-Weinberg equilibrium distribution, was calculated as QAB = 2QA QB . The observed heterozygous frequency PAB should be lower than the expected heterozygous frequency QAB for imprinted genes and genes with allelic variation in gene expression. This concept can be analyzed using Z-statistics. The author and colleagues have demonstrated that their approach is effective in identifying allelic variation in gene expression.41 The second method to identify allelic variation in gene expression is to analyze allele frequency in a heterozygous EST library using a binomial distribution. A significant deviation from 0.5 provided the evidence for allelic variation in gene expression. Using this in silico analysis, they identified 524 genes that displayed allelic variation in at least one cDNA library. Six genes were selected for validation and their experiment confirmed allelic variation in gene expression in all the six genes examined.37 The allelic variation in gene expression between the two chromosomes of the same individual (intra-individual allelic variation) is closely related to the inter-individual allelic variation in gene

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expression. DNA polymorphisms and variability in gene expression provide the genetic and epigenetic basis for human phenotypic variation. Studies in humans and other organisms suggest that the variation in transcript level accounts for the majority of the variation among species and across individuals within species.42–45 Gene expression variation is determined by genetic and epigenetic mechanisms. The effect of a regulatory SNP on differential gene expression can manifest in inter-individual gene expression variation,42,46–48 as well as in intra-individual allelic gene expression variation between the two chromosomes.35,39,49,50 An integrated analysis of gene expression, chromatin structure, and DNA methylation using SNP chips provides a unique opportunity for conducting integrated genetic and epigenetic studies of cancer etiology. 5. PERSPECTIVE Recently, NCI and NHGRI initiated The Cancer Genome Atlas (TCGA). The goals of TCGA are to develop a comprehensive catalog of genomic changes that occur in tumors and to understand how these changes relate to the biological processes in cancer. The genome characterization component of TCGA involves studies of chromosome rearrangements, chromosome copy number changes, epigenetics and gene expression. The studies discussed in this review article can contribute to the broader TCGA efforts. To gain a quantitative understanding of the biological processes from the “-omics” data, it is necessary to have robustly extract biological signals from inherently noisy microarray data. To achieve this goal, it will be necessary to develop analytic methods that can effectively reduce the complexity and noise in the “-omics” data. Some of the most promising methods include principal component analysis (PCA), factor analysis (FA), and independent component analysis (ICA), which can reduce the complexity and noise of the “-omics” data and enable the identification of latent factors underlying the observed patterns of gene expression, chromatin structure and DNA methylation. The success of the systems biology approach to understanding cancer etiology will require the development of

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mathematical models that can capture and predict the dynamic behavior of genetic and epigenetic events in normal and cancer cells.

ACKNOWLEDGMENTS The author wishes to thank Dr Howard Yang for the many stimulating discussions on the application of mathematics and statistics in systems biology. This research was supported by the Intramural Research Program of the NIH and the National Cancer Institute.

References 1. Beck S, Olek A, Walter J. (1999) From genomics to epigenomics: a loftier view of life. Nat Biotechnol 17: 1144. 2. Bjornsson HT, Cui H, Gius D, et al. (2004) The new field of epigenomics: implications for cancer and other common disease research. Cold Spring Harb Symp Quant Biol 69: 447–456. 3. Novik KL, Nimmrich I, Genc B, et al. (2002) Epigenomics: genome-wide study of methylation phenomena. Curr Issues Mol Biol 4: 111–128. 4. Feinberg AP. (2000) DNA methylation, genomic imprinting and cancer. Curr Top Microbiol Immunol 249: 87–99. 5. Jones PA, Baylin SB. (2002) The fundamental role of epigenetic events in cancer. Nat Rev Genet 3: 415–428. 6. Bird A, Macleod D. (2004) Reading the DNA methylation signal. Cold Spring Harb Symp Quant Biol 69: 113–118. 7. Jenuwein T, Allis CD. (2001) Translating the histone code. Science 293: 1074–1080. 8. Bartolomei MS, Zemel S, Tilghman SM. (1991) Parental imprinting of the mouse H19 gene. Nature 351: 153–155. 9. DeChiara TM, Robertson EJ, Efstratiadis A. (1991) Parental imprinting of the mouse insulin-like growth factor II gene. Cell 64: 849–859. 10. Rainier S, Johnson LA, Dobry CJ, et al. (1993) Relaxation of imprinted genes in human cancer. Nature 362: 747–749. 11. Lyon MF. (1961) Gene action in the X-chromosome of the mouse (Mus musculus L.). Nature 190: 372–373. 12. Wang Z, Fan H, Yang HH, et al. (2004) Comparative sequence analysis of imprinted genes between human and mouse to reveal imprinting signatures. Genomics 83: 395–401. 13. Takai D, Jones PA. (2002) Comprehensive analysis of CpG islands in human chromosomes 21 and 22. Proc Natl Acad Sci USA 99: 3740–3745.

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14. Hayashizaki Y, Shibata H, Hirotsune S, et al. (1994) Identification of an imprinted U2af binding protein related sequence on mouse chromosome 11 using the RLGS method. Nat Genet 6: 33–40. 15. Plass C, Shibata H, Kalcheva I, et al. (1996) Identification of Grf1 on mouse chromosome 9 as an imprinted gene by RLGS-M. Nat Genet 14: 106–109. 16. Strichman-Almashanu LZ, Lee RS, Onyango PO, et al. (2002) Agenomewide screen for normally methylated human CpG islands that can identify novel imprinted genes. Genome Res 12: 543–554. 17. Huang TH, Perry MR, Laux DE. (1999) Methylation profiling of CpG islands in human breast cancer cells. Hum Mol Genet 8: 459–470. 18. Wei SH, Chen CM, Strathdee G, et al. (2002) Methylation microarray analysis of late-stage ovarian carcinomas distinguishes progressionfree survival in patients and identifies candidate epigenetic markers. Clin Cancer Res 8: 2246–2252. 19. Ching TT, Maunakea AK, Jun P, et al. (2005) Epigenome analyses using BAC microarrays identify evolutionary conservation of tissue-specific methylation of SHANK3. Nat Genet 37: 645–651. 20. Weber M, Davies JJ, Wittig D, et al. (2005) Chromosome-wide and promoter-specific analyses identify sites of differential DNA methylation in normal and transformed human cells. Nat Genet 37: 853–862. 21. Bibikova M, Lin Z, Zhou L, et al. (2006) High-throughput DNA methylation profiling using universal bead arrays. Genome Res 16: 383–393. 22. Fukasawa M, Kimura M, Morita S, et al. (2006) Microarray analysis of promoter methylation in lung cancers. J Hum Genet 51: 368–374. 23. Hatada I, Fukasawa M, Kimura M, et al. (2006) Genome-wide profiling of promoter methylation in human. Oncogene 25: 3059–3064. 24. Feinberg AP. (2004) The epigenetics of cancer etiology. Semin Cancer Biol 14: 427–432. 25. Bernstein BE, Kamal M, Lindblad-Toh K, et al. (2005) Genomic maps and comparative analysis of histone modifications in human and mouse. Cell 120: 169–181. 26. Kim TH, Barrera LO, Zheng M, et al. (2005) A high-resolution map of active promoters in the human genome. Nature 436: 876–880. 27. Miao F, Natarajan R. (2005) Mapping global histone methylation patterns in the coding regions of human genes. Mol Cell Biol 25: 4650–4661. 28. Roh TY, Cuddapah S, Zhao K. (2005) Active chromatin domains are defined by acetylation islands revealed by genome-wide mapping. Genes Dev 19: 542–552. 29. Brentani RR, Carraro DM, Verjovski-Almeida S, et al. (2005) Gene expression arrays in cancer research: methods and applications. Crit Rev Oncol Hematol 54: 95–105.

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30. Ebert BL, Golub TR. (2004) Genomic approaches to hematologic malignancies. Blood 104: 923–932. 31. Simon R. (2005) Road map for developing and validating therapeutically relevant genomic classifiers. J Clin Oncol 23: 7332–7341. 32. Bild AH, Yao G, Chang JT, et al. (2006) Oncogenic pathway signatures in human cancers as a guide to targeted therapies. Nature 439: 353–357. 33. Yang HH, Hu Y, Buetow KH, Lee MP. (2004) A computational approach to measuring coherence of gene expression in pathways. Genomics 84: 211–217. 34. Lee MP, Howcroft K, Kotekar A, et al. (2005) ATG deserts define a novel core promoter subclass. Genome Res 15: 1189–1197. 35. Lo HS, Wang Z, Hu Y, et al. (2003) Allelic variation in gene expression is common in the human genome. Genome Res 13: 1855–1862. 36. Ge B, Gurd S, Gaudin T, et al. (2005) Survey of allelic expression using EST mining. Genome Res 15: 1584–1591. 37. Lin W, Yang HH, Lee MP. (2005) Allelic variation in gene expression identified through computational analysis of the dbEST database. Genomics 86: 518–527. 38. Pastinen T, Ge B, Gurd S, et al. (2005) Mapping common regulatory variants to human haplotypes. Hum Mol Genet 14: 3963–3971. 39. Pastinen T, Sladek R, Gurd S, et al. (2004) A survey of genetic and epigenetic variation affecting human gene expression. Physiol Genomics 16: 184–193. 40. Pant PV, Tao H, Beilharz EJ, et al. (2006) Analysis of allelic differential expression in human white blood cells. Genome Res 16: 331–339. 41. Yang HH, Hu Y, Edmonson M, et al. (2003) Computation method to identify differential allelic gene expression and novel imprinted genes. Bioinformatics 19: 952–955. 42. Cheung VG, Conlin LK, Weber TM, et al. (2003) Natural variation in human gene expression assessed in lymphoblastoid cells. Nat Genet 33: 422–425. 43. Enard W, Khaitovich P, Klose J, et al. (2002) Intra- and inter-specific variation in primate gene expression patterns. Science 296: 340–343. 44. Johnson NA, Porter AH. (2000) Rapid speciation via parallel, directional selection on regulatory genetic pathways. J Theor Biol 205: 527– 542. 45. Levine M, Tjian R. (2003) Transcription regulation and animal diversity. Nature 424: 147–151. 46. Brem RB, Yvert G, Clinton R, Kruglyak L. (2002) Genetic dissection of transcriptional regulation in budding yeast. Science 296: 752–755. 47. Schadt EE, Monks SA, Drake TA, et al. (2003) Genetics of gene expression surveyed in maize, mouse and man. Nature 422: 297–302.

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48. Yvert G, Brem RB, Whittle J, et al. (2003) Trans-acting regulatory variation in Saccharomyces cerevisiae and the role of transcription factors. Nat Genet 35: 57–64. 49. Cowles CR, Hirschhorn JN, Altshuler D, Lander ES. (2002) Detection of regulatory variation in mouse genes. Nat Genet 32: 432–437. 50. Yan H, Yuan W, Velculescu VE, et al. (2002) Allelic variation in human gene expression. Science 297: 1143.

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CHAPTER 12

Molecular Epidemiology of DNA Repair and Cancer Susceptibility — A Review of Population-based Studies Zhibin Hu, Li-E Wang and Qingyi Wei∗

ABSTRACT The role of DNA repair in the etiology of cancer has been well illustrated in several hereditary syndromes, in which an inherited defect in DNA repair is associated with an extraordinarily high incidence of cancer. However, such an association between an inherited DNA repair defect and the risk of cancer has not been apparent in the general population. In the past 10 years, there has been a growing body of literature that addressed this important research question at the population level. Several assays that measure DNA repair phenotypes (e.g. transfection-based host-cell reactivation, expression of genes and proteins, and cytogenetics, etc.) have been applied to population-based studies. It has been demonstrated that a suboptimal DNA repair capacity for removing DNA damage induced by ultraviolet light and benzo[a]pyrene diol epoxide, two-well known environmental carcinogens, is associated with an increased risk of skin cancer (e.g. melanoma and non-melanoma skin cancers) and tobaccorelated cancers (e.g. lung and head and neck cancers), respectively. More recently, the completion of the Human Genome Project and the initiation of the Environmental Genome Project have led to the discovery of numerous single nucleotide polymorphisms in DNA repair genes, followed by a

∗ Corresponding

author. Department of Epidemiology, The University of Texas M. D. Anderson Cancer Center, 1515 Holcombe Blvd, Houston, Texas 77030, USA. Tel.: 713-792-3020; Fax: 713-563-0999; E-mail: [email protected] 315

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wave of association studies, for cancer in particular. Published data appear to have established a genetic basis for a suboptimal repair phenotype in the general population and provide a rationale for future genetic screening for at-risk populations who can be targeted for the primary prevention of cancer.

1. INTRODUCTION DNA in humans is constantly assaulted by cellular metabolites and exogenous damaging agents. DNA lesions that are not properly repaired prior to cell division can be mutagenic, resulting in a precancerous cell. Fortunately, species of all living organisms developed a sophisticated DNA repair machinery that battles against genomic insults and maintains genomic integrity. More than 130 genes are known to be involved in various repair pathways for repairing different classes of DNA lesions, and this number is likely to increase as knowledge of the mechanisms of DNA repair increases over time.1,2 It has long been recognized that there is an inter-individual variability in the response to carcinogens in humans.3 This variability can be estimated by the cellular DNA repair capacity (DRC), an extent to which the cell meets the challenge from exogenous as well as endogenous environments. Defective DRC phenotype is often a rare, recessive trait that exists in some genetic syndromes, such as ataxia telangiectasia (AT), Fanconi anemia, and Bloom’s syndrome, which are characterized by both chromosomal instability and high risk of cancer. For example, xeroderma pigmentosum (XP), a disease caused by a deficiency in nucleotide excision repair (NER) is characterized by UV light sensitivity and high risk of skin cancer.4 Apart from these rare syndromes, individuals also differ widely in their capacity to repair DNA damage, and reduced DRC in the general population appeared to be associated with the risk of developing cancer.5,6 It is believed that the DRC phenotype is genetically determined, and emerging studies have shown that polymorphisms in DNA repair genes may modify the DRC phenotype and thus contribute to the variation of carcinogenesis in the general population,7,8 providing the rationale of using genetic markers to predict cancer risk for the purpose of identifying at-risk individuals for prevention.

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In this chapter, the authors provide an overview of the use of the markers for the DRC phenotype and genotypes in association studies. 2. DNA REPAIR PHENOTYPES AND CANCER SUSCEPTIBILITY A number of epidemiologic studies have been conducted to compare the measurements of the DNA repair phenotype between cancer cases and healthy control subjects and to assess the role of DNA repair in the development of human cancer. A variety of measures of the DNA repair phenotype were used in these studies, and the currently used assays do not detect specific aspects of repair but rather assess its global effects: 1) direct measures of repair kinetics using circulating lymphocytes, such as host-cell reactivation; 2) levels of carcinogen-DNA adducts in human tissues and of carcinogen-protein adducts in the blood; and 3) DNA repair gene and protein expression levels. 2.1. Host-Cell Reactivation Assay The DRC phenotype assay9,10 takes advantage of non-replicating transient expression plasmid that is damaged prior to transfection into the host cell rather than using direct damage to the genome. This technique minimizes cytotoxic effects of the damaging agents that might indirectly compromise the repair machinery of the cell and thus provides a direct measure of the host DRC.10 Specifically, in the assay, a damaged non-replicating recombinant plasmid harboring either a chloramphenicol acetyltransferase (pCMVcat) or luciferase (pCMVluc) reporter gene is introduced by transfection into cultured lymphocytes. The readout of the reporter gene expression is the reactivation outcome of the damage plasmid by the cellular repair mechanism, because the presence of only one unrepaired DNA lesion can block the transcription of the essential gene.11 Thus, the reactivated CAT or LUC enzyme activity is measured as a function of excision repair of the damaged gene, regardless of the type of report genes used.7

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There have been a number of studies using the DRC assay in principle to investigate the association between a suboptimal DRC and the risk of cancers, including cancers of the skin, lung, head and neck, and breast10,12–22 (See Table 1). In general, case patients were likely to have a suboptimal DRC and a high risk of cancers studied so far, compared to cancer-free controls. For example, in both the initial pilot study17 (51 patients and 56 frequency-matched controls) and a subsequent large hospital-based case-control study (316 each of lung cancer patients and controls) of lung cancer,18 it was noted that there was a five-fold variation in DRC of the cancer-free controls, but statistically significantly lower DRC was observed in the cases compared with the controls, and this difference was associated with a greater than two-fold increased risk of lung cancer. Furthermore, in a recent expanded study with a more-than-doubled sample size, the mean DRC ± SD for 764 lung cancer cases was 7.82% (±2.82), compared with 8.79% (±3.87) for the 677 controls (P < 0.001). The adjusted risk estimate associated with a suboptimal DRC (defined as below the control median) was 1.50 with a more stringent confidence interval (CI) from 1.2 to 1.9.8

Table 1. Cancer Skin BCC BCC BCC SCC/BCC Melanoma Lung

Head & neck Prostate Breast ∗ For

Studies on DRC and Cancer Risk

Case/Control

OR (95%CI)

88/123 86/87 49/68 280/177 132/145 312/324 51/56 316/316 467/488 55/61 140/96 69/79

2.3 (1.2–4.5)* 1.1 (0.9–1.3) 1.2 (0.5–2.7)** 3.8 (2.3–5.7)* 1.0 (0.6–1.7) 2.0 (1.5–2.8) 5.7 (2.1–15.7) 2.1 (1.5–3.0) 1.9 (1.4–2.4) 2.2 (1.0–4.8) 2.1 (1.2–3.9) 3.4 (1.2–9.8)

Reference Wei et al.10 Hall et al.12 D’Errico et al.13 Matta et al.14 Landi et al.15 Wei et al.16 Wei et al.17 Wei et al.18 Shen et al.19 Cheng et al.20 Hu et al.21 Shi et al.22

the low tertile DRC and others for the low-median DRC. based on published data.

∗∗ Recalculated

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To facilitate epidemiological studies seeking to investigate the association between DNA repair proficiency and cancer susceptibility, time, cost and repeatability are major concerns in the choice of one particular assay. However, the host-cell reactivation (HCR) DRC assay is currently deemed not to be amenable to large-scale population-based screening. More simplified DNA repair phenotype assays that can accommodate the needs for measuring specific repair pathways are critical for the development of the research field. A promising development of using mitomycin C as the damaging agent in the HCR assay may provide a novel DRC assay for measuring DNA crosslink or double-strand break repair pathways,23 which may not only be relevant in evaluating susceptibility to cancer but also useful in assessing therapeutic responses to chemo- and radiotherapies in cancer patients. 2.2. Induced DNA Adducts A relatively large variation is observed in the level of persistent DNA adducts in vivo.24,25 This variation could be a valid phenotypic marker for the joint effect of host metabolic activities and DNA repair in response to carcinogen exposure.26 A number of methods have been developed to measure DNA adducts in humans, among which, 32 P-postlabeling analysis constitutes a sensitive method that does not need a prior knowledge of the structures of adducts.27 Studies have shown that significantly elevated levels of DNA adducts were detected in the peripheral lung, bronchial epithelium, or bronchioalveolar lavage cells of smokers, for which a linear correlation between adduct levels and a daily or lifetime consumption of cigarettes has been established.28,29 Furthermore, the levels of adducts were negatively associated with the consumption of fruits and vegetables,30,31 possibly through a stimulation of DNA repair or a reduced level of oxidative stress.32 In a pilot study of in vitro induced BPDE-DNA adducts in cultured peripheral lymphocytes, lung cancer patients tended to accumulate higher levels of adducts than the controls, representing an association between

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levels of DNA adducts and high risk of lung cancer.26 Recently, a meta-analysis including a total of 691 cancer patients and 632 control subjects from five lung cancer studies, one oral cancer study, and one bladder cancer study found that cases had higher levels of DNA adducts than controls among current smokers and that current smokers with high levels of DNA adducts had a significantly increased risk of developing lung and bladder cancers.33 More recently, immunofluorescence using a polyclonal anti-BPDE-DNA primary antibody was used to quantify BPDE-DNA adducts as a measure of DRC. One study using this assay found that a deficient DRC contributed to the susceptibility to breast cancer, particularly in familial breast cancer.34 2.3. DNA Repair Gene RNA Expression The gene expression levels can be affected genetically and epigenetically, such as polymorphisms and the methylation status of the promoters, respectively. Several epidemiological studies have investigated the variation in gene expression levels using a multiple reverse transcriptase-polymerase chain reaction assay that allows investigators to measure the levels of several DNA repair gene transcripts relative to that of a ubiquitous housekeeping gene.35 This assay allowed flexibility in grouping the genes into one experiment involved in the same repair pathway, such as mismatch repair36 and NER.37–39 In a case-control study of the relative expression levels of five NER genes (ERCC1, XPB, XPG, CSB and XPC) in phytohemagglutinin-stimulated peripheral lymphocytes, a relative reduction of 12.2% and 12.5% in the baseline expression levels of XPG and CSB, respectively, was found in lung cancer cases compared with the controls, and this reduction was associated with a more than two-fold increased risk of lung cancer.38 In another similar study, the levels of ERCC1, XPB, XPG and CSB transcripts were found to be lower among the head and neck cancer cases than among the controls, and were also associated with a significantly increased cancer risk.39

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2.4. DNA Repair Protein Expression Levels However, the association studies using RNA expression may be biased, because the transcript level does not necessarily reflect the level of protein expression, and protein activities may only be initiated after phosphorylation. In a recent study, the expression of six core NER proteins (ERCC1, XPA, XPB, XPC, XPD and XPF) was measured in cultured lymphocytes and found to be associated with the risk of head and neck cancer.40 Specifically, compared with the controls, the cases had lower relative expression levels for all the NER proteins, particularly XPC and XPF, which were reduced by about 25% (P < 0.01).40 When the median expression levels of the NER proteins in the controls were used as the cutoff values, there was a significantly increased cancer risk associated with the low expression of XPA (nearly three-fold), XPC (two-fold), XPD (three-fold), and XPF (greater than five-fold), but not ERCC1 and XPG.40 In a multivariate logistic regression model that included all the covariates and NER proteins, however, only the low expression of XPF remained a significant risk factor (greater than an 11-fold risk).40 These results suggest that XPF may be a crucial rate-limiting factor in NER repair and that the reverse-protein microarray assay may be a useful tool for measuring the protein markers for susceptibility to cancer. 3. DNA REPAIR GENE POLYMORPHISMS AND CANCER SUSCEPTIBILITY Of specific interest for this section has been the selection of genetic variants of candidate DNA repair genes in association studies. To date, there are more than 10 million single nucleotide polymorphisms (SNPs) with a minor allele frequency > 1% that are estimated to exist in the genome.41 For the DNA repair genes, there are more than 130 genes involved in the four main repair pathways (Table 2), 68 of which are resequenced in the National Institute of Health Sciences (NIEHS) Environmental Genome Project (http://egp.gs.washington.edu/) (Table 3). Of the identified 11,507

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Table 2.

Genes in Human DNA Repair Pathways

Type

Genes Involved

Damage recognition checkpoint

> 30 genes: RAD1, RAD9, RAD17, Chk1, Chk2, ATM, ATR, BRCA1, BRCA2, P53, TP53BP1, MDC1…

Base excision repair

> 30 genes: DNA ligase (LIGI, LIGIII); DNA glycosylase (hOGG1, hMYH, hNTG1, MBD4, TDG, UNG); APE1, POLB, XRCC1, FEN1, RPA1–3, PCNA, PARP, DNA polymerase (POLE1–5, POLD1–2), RFC…

Nucleotide excision repair

> 30 genes: XPA, XPC, ERCC1, ERCC2, ERCC3, ERCC4, ERCC5, CSB/ERCC6, RPA1–3, RFC1–5, DDB1, DDB2, HHR23A, HHR23B, p34, p38, p41, p44, p62, PCNA, DNA polymerase (POLD1, POLD2, POLE1–4, POLH), LIGI …

Mismatch repair

> 10 genes: hMLH1, hMLH3, hPMS1, hPMS2, hMSH2, hMSH3, hMSH6/hGTBP…

Double strand break repair Non-homologous end joining

Homologous recombination

> 15 Genes: DNA-PKcs, KU70 (XRCC6), KU80 (XRCC5), MRE11, NBS1, RAD50, XRCC4, LIGIV… > 15 Genes: MRE11, NBS1, RAD50, RAD51, RAD52, RAD54, XRCC2, XRCC3, XRCC4, XRCC7, XRCC8, RAD51B, RAD51C, RAD51D…

variants, some are more likely to be a priori disease-causing ones than others. Although the significance of these variants is largely unknown, the implication is that variants that cause amino acid substitutions (i.e. non-synonymous) may have an impact on the function of the proteins and therefore on the efficiency of the DNA repair phenotype. Those variants that do not cause an amino acid change may also have an impact on the DNArepair phenotype, because they may lie in introns that regulate splicing; may cause mRNA instability; or may be linked to genetic changes in other unknown genes. Therefore, knowing the impact of these polymorphisms on gene functions is important for us to further understand their influence on disease outcomes.

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Table 3. Summary of Genetic Variants Identified in Genes in Different DNA Repair Pathways in the EGP Databases (http://egp.gs.washington.edu/) Pathway

Damage recognition checkpoint Base excision repair Nucleotide excision repair Mismatch repair Double strand break repair Total

No. of Genes in Pathway

No. of Genes Screened

No. of No. of Variants Alleles > 10% Identified Frequency∗

>30

11

2147

602

>30 >30

26 22

3061 3377

818 813

>10 >30

9 13

2001 2825

537 722

>130

68

11507

2934

*Genes with roles in more than one pathway are counted in all relevant pathways. The total represents the number of either unique genes and variants, not the sum of the column.

3.1. Study Design and SNP Selection 3.1.1. Linkage mapping and linkage disequilibrium (LD) mapping For the past two decades, the dominant study design for investigating the genetic basis of an inherited disease has been linkage analysis in families. The foremost prerequisite for successful linkage mapping is a set of families in which the disease phenotype is segregating and the correspondence between the genotype and phenotype is nearly one-to-one. Where complex inheritance or frequent phenocopies are abundant, LD mapping methods, depending on allelic associations, are expected to be more powerful, because each of the disease-predisposing alleles only contributes to some modest risks.42,43 Twin studies indicate that much of the familial aggregation of cancer results from inherited susceptibility.44 Highly penetrant mutations in known genes cannot account for much of the excess risk observed in the general population. For example, mutations in the known predisposition genes, including BRCA1 and BRCA2, account for only approximately 20% of the two-fold excess risk in breast cancer patients’ relatives.45 The remaining familial risk could also be due to high-penetrance mutations in as yet unidentified genes, and

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Summary of nsSNPs Identified in Genes in Different DNA Repair Pathways (MAF > 0.05)

Damage recognition checkpoint

BRCA1*

Wild Type

Variant

Variant Frequency

SIFT Predict

PolyPhen Predict Benign Benign Benign Benign Benign Benign Benign Benign

C A A A G G G C

T G G G C C A A

871 1038 1183 1613 185 353 412 1136

PRO [P] LEU [L] GLU [E] GLY [G] LYS [K] ARG [R] SER [S] GLY [G] GLU [E] GLN [Q] GLU [E] ASP [D] GLY [G] SER [S] GLN [Q] LYS [K]

0.43 0.32 0.33 0.33 0.37 0.45 0.41 0.46

Tolerant Damaging Tolerant Tolerant Tolerant Tolerant Tolerant Tolerant

MUTYH OGG1 TDG

1136410 3136820 10342 2307289 140693 3219489 1052133 4135113

T T G T G G C G

C G T C A C G A

762 148 273 342 346 335 326 199

VAL [V] ALA [A] ASP [D] GLU [E] ALA [A] SER [S] SER [S] PRO [P] GLU [E] LYS [K] GLN [Q] HIS [H] SER [S] CYS [C] GLY [G] SER [S]

0.19 0.46 0.07 0.06 0.07 0.41 0.32 0.09

Tolerant Tolerant Tolerant Damaging Tolerant Tolerant Tolerant Tolerant

XRCC1

2888805 1799782

G C

A T

367 194

VAL [V] MET [M] ARG [R] TRP [W]

0.09 0.12

Tolerant Tolerant

25489

G

A

280

ARG [R]

0.10

Tolerant

ADPRT APEX MBD4

ERCC2

HIS [H]

25487

G

A

399

ARG [R]

GLN [Q]

0.23

Tolerant

1799793 1052559

G A

A C

312 751

ASP [D] LYS [K]

ASN [N] GLN [Q]

0.24 0.22

Tolerant Tolerant

Benign Benign Benign Benign Benign Benign Benign Probably Damaging Benign Probably Damaging Probably Damaging Benign Benign Benign

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Nucleotide excision

799917 16941 16942 1799966 NBS1* 1805794 TP53BP1* 560191 689647 2602141

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Allele 1 Allele 2 Codon

Wild Type

Variant

Variant Frequency

SIFT Predict

PolyPhen Predict

Repair

ERCC4 ERCC5

2020955 17655

T G

C C

662 1104

SER [S] ASP [D]

PRO [P] HIS [H]

0.06 0.38

Tolerant Damaging

A G G C G A A T T C

399 1097 1213 1230 1413 119 173 249 499 939

GLY [G] ASP [D] MET [M] VAL [V] ARG [R] GLY [G] ARG [R] PRO [P] GLN [Q] ARG [R] ARG [R] HIS [H] SER [S] ASN [N] ALA [A] VAL [V] ALA [A] VAL [V] LYS [K] GLN [Q]

0.22 0.18 0.19 0.08 0.19 0.14 0.06 0.20 0.24 0.34

Tolerant Tolerant Tolerant Tolerant Tolerant Tolerant Tolerant Tolerant Tolerant Tolerant

Benign Possibly Damaging No prediction No prediction No prediction No prediction No prediction Benign Benign Benign Benign Benign

2228528 2228526 2228527 4253211 2228529 POLD1** 1726801 1726803 RAD23B 1805329 XPC 2228000 2228001

G A A G A G G C C A

735943 4149963

G C

A T

354 439

ARG [R] THR [T]

HIS [H] MET [M]

0.31 0.13

Tolerant Tolerant

4149965 1047840 1776148 9350

G G G C

A A A T

458 589 670 757

VAL [V] GLU [E] GLY [G] PRO [P]

MET [M] LYS [K] GLU [E] LEU [L]

0.12 0.31 0.32 0.31

Tolerant Tolerant Tolerant Damaging

1799977 144848 861539

A A C

G C T

219 372 241

ILE [I] VAL [V] ASN [N] HIS [H] THR [T] MET [M]

0.18 0.26 0.22

Tolerant Tolerant Tolerant

ERCC6

Mismatch repair

Double strand break repair

MLH1 BRCA2 XRCC3

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involved in double-strand break repair. involved in mismatch repair.

∗∗ Also

Benign Probably Damaging Benign Benign Benign Probably Damaging Benign Benign Benign

325

∗ Also

EXO1

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a polygenic mechanism appears to be more plausible.46 Under the polygenic model, efforts have been made to perform association or LD studies, where a large number of alleles, each conferring a small genotypic risk, can be combined additively or multiplicatively to confer a range of susceptibility in the general population.47,48 3.1.2. “Map-based” approach and “sequence-based” approach Currently, there are two competitive strategies of selecting SNPs in the association study design, i.e., the “map-based” approach and the “sequence-based” approach.49 The first, usually referred as indirect association, is to test a dense map of SNPs for disease association under the assumption that if a risk polymorphism exists, it will either be genotyped directly or be in strong LD with one of the typed SNPs. Because SNPs in LD seem to have a block-like structure,50 it has been suggested that a judiciously chosen subset of tagging SNPs can identify most of the genetic variation through haplotypes.51–53 This tagging-SNP approach has the advantage of scanning of an entire region of interest without having to rely on choosing an a priori candidate. With the advances in high-throughput technology and genome-wide association methods, this approach will be more practical and feasible, if not easily implemented. An alternate approach is the putative functional SNP analysis, based on candidate genes and sequences.54,55 In this strategy, genotyping focuses on SNPs in the coding regions of candidate genes or pathways, which alter or terminate an amino acid sequence, disrupt splice sites, and cause sequency changes in the promoter regions. This “sequence-based” approach has the advantage of maximizing inferences about biological plausibility and disease causality but is limited by the amount of currently available information on the functions of the SNPs in a specific disease process. 3.1.3. Functional consideration of candidate SNPs The identification of biologically plausible, disease-causing variants from among a large number of identified SNPs is crucial for the

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appropriate study design and interpretation of candidate SNPs in association studies. Inference about the functions of a genetic variant can be made using experimental systems, including in vitro systems and in vivo animal models. These include the effect of variants on the regulatory-region control of gene expression, RNA stability or degradation, protein structure, protein denaturing, and other measures of in vivo and in vitro control of protein levels.55,56 An alternative approach to selecting SNPs is the use of computational algorithms based on evolutionary and/or structural analyses. For example, two computational algorithms (the Sorting Intolerant from Tolerant, SIFT: http://blocks.fhcrc.org/sift/SIFT.html and the Polymorphism Phenotyping Algorithm, PolyPhen: http:// www.bork.embl-heidelberg. de/PolyPhen) are available in predicting the functional consequences of nsSNPs. SIFT is based on an evolutionary approach and Bayesian polygenetic analyses,57 while PolyPhen uses a structural-based approach.58 For example, using SIFT and PolyPhen, significant relationships between the detected odds ratios associated with cancer risk and the conservation levels and/or position-specific independent counts of the SNP affected amino acids were found for 46 nsSNPs in 39 different cancer-related genes from 166 molecular epidemiological studies, suggesting that these computational algorithms can optimize the ability to identify meaningful and reproducible SNP and disease associations.55,59 As shown in Table 3, the authors also used the SIFT and PolyPhen software to evaluate specific nsSNPs with a minor frequency allele (MAF) > 0.05 in the select DNA repair genes. The SIFT program predicts that BRCA1 E1038G, ERCC5 D1104H, EXO1 P757L and MBD4 S342P may have functional consequences, while the PolyPhen algorithm predicts the same for ERCC5 D1104H and EXO1 P757L, specifically for EXO1 T439M, TDG G199S, XRCC1 R194W and R280H. Among these, the NER gene ERCC5 D1104H polymorphism was suggested to be associated with lung and bladder cancer risk.60,61 The MMR gene EXO1 P757L and T439M polymorphisms were also suggested to be associated with the risk of colorectal cancer in a Japanese population.62 Furthermore, the two germ-line mutations R194W and R280H in the well-characterized BER geneXRCC1 were

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associated with cancer risk as suggested by a recent large metaanalysis.63 3.1.4. Allele frequencies for susceptibility loci Two polarized views have dominated much of the literature on the allelic frequencies of common diseases.64,65 A simple way of describing the relative importance of a locus from the standpoint of public health is to use the population attributable fraction (PAF). Common modest-risk alleles may account for a greater PAF in common diseases than do rare high-risk alleles; this is often referred to as the common-disease common-variant (CDCV) hypothesis,66 which provides an important impetus for the Haplotype Map Project and the genome-wide association scans.51–53 However, an extreme alternative to the CDCV hypothesis is the classical multiple rare-variant hypothesis, which hypothesizes that disease susceptibility is due to one or more distinct genetic variants in different individuals and that disease-susceptibility alleles have low population frequencies.67 The most neutral hypothesis states that the allelic spectrum of disease-causing variants is the same as the general spectrum of all genetic variants.68 Under this neutral model, the total contribution of the rare variants among the DNA-repair genes to the etiology of diseases would be about the same as the intermediate variants and slightly less than the common variants, according to their contributions to genetic variation.69 However, it is difficult to provide a sufficient statistical power to detect the moderate effects of the rare variants and haplotypes in association studies. 3.2. Studies on Polymorphisms of the Select DNA Repair Genes and Cancer Risk 3.2.1. ATM ATM is an important damage-sensoring protein that plays a central role in the DNA-damage checkpoint pathway. Mutations in ATM cause ataxia telangiectesia (AT), a genetic syndrome characterized by cerebellar degeneration, immunodeficiency, genome instability,

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clinical radiosensitivity and cancer predisposition.70 ATM exhibits significant sequence homology to the phosphoinositide 3-kinases,71 and ATM has protein kinase activity. Upon exposure of the cells to ionizing radiation, ATM phosphorylates many proteins, including Chk2, P53, NBS1 and BRCA1 (see Chapter 1 for more details). The ATM gene contains 397 SNPs, 67 of which have an MAF of more than 0.1; however, no common nsSNP was found in this gene as suggested by the NIEHS EGP databases (http://egp.gs.washington.edu/). Several epidemiological studies evaluated the association of germ-line mutations in ATM and breast cancer risk.72–74 It is reported that the frequency of two tightly linked intronic ATM polymorphisms IVS22-77 T>C and IVS48+238 C>G was associated with an increased breast cancer risk in Caucasian women.72 Consistently, another study suggested that the IVS21 + 1049 T>C, IVS34 + 60 G>A, and 3393 T>G polymorphisms in ATM were breast cancer susceptibility loci in an Asian population.73 However, the Nurses’ Health Study cohort provides no support for an association between common haplotypes of ATM and breast cancer risk.74 3.2.2. XRCC1 XRCC1 is a well-characterized scaffolding protein which participates in the BER pathway (see Chapter 2 for more details). Both biological and biochemical evidence indicates a direct role for XRCC1 in BER since it interacts with a complex of DNA repair proteins, including poly(ADP-ribose) polymerase (PARP), DNA ligase 3 (LIG3) and DNA polymerase-β. There are a total of eight nsSNPs in XRCC1, three of which are common (i.e. MAF > 0.05) and lead to amino acid substitutions in XRCC1 at codon 194 (base C to T, or amino acid Arg to Trp at exon 6); codon 280 (base G to A or amino acid Arg to His at exon 9); and codon 399 (base G to A or amino acid Arg to Gln at exon 10) (see http://egp.gs.washington.edu). The Arg399Gln polymorphism is located in the region of the BRCT-I interaction domain of XRCC1 within a poly(ADPribose) polymerase binding region,75 whereas the Arg194Trp and

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Arg280His variants occur in the newly identified PCNA binding region.76 The presence of the variant 399Gln allele has been shown to be associated with measurable reduced DRC as assessed by the persistence of DNA adducts78–79 , elevated levels of sister chromatid exchanges78,80 ; increased red blood cell glycophorin A77 ; P53 mutations81 ; and prolonged cell cycle delay.82 However, even though the BRCT-I domain is critical for XRCC1-dependent singlestrand break repair for the maintenance of genetic integrity, the Arg399Gln polymorphism in BRCT-I does not have a significant impact on this function,83 and negative functional studies were also reported.84–86 It is reported that individuals with the variant 194Trp allele had fewer bleomycin- or BPDE-induced chromosomal breaks than those with wild-type genotype87 ; however, other studies did not find a significant association of Arg194Trp with altered levels of DNA adducts77 or G2 cell cycle delay.82 A large number of molecular epidemiological studies have been conducted to evaluate the role of the XRCC1 polymorphisms in cancer risk. A recently published meta-analysis described a systematic review on a total of 38 published studies examining the relationship between XRCC1 nsSNPs and cancer risk.63 It was found that the variant genotypes of Arg194Trp and Arg280His, but not the Arg399Gln, were associated with a significantly altered cancer risk for all tumor types without between-study heterogeneity.63 By pooling of data from seven studies, it was found that the XRCC1 399Gln/Gln genotype was associated with an increased risk of tobacco-related cancers among light smokers but a decreased risk among heavy smokers88 ; therefore, a detailed analysis of effect of modification by environmental exposure and gene-environment interactions in single large studies was recommended. 3.2.3. XPD The 54.3-kb XPD gene codes for an evolutionarily conserved helicase, a subunit of TFIIH that is essential for transcription and NER (see Chapter 3 for more details). Mutations in XPD prevent the protein from interacting with p44, another subunit of TFIIH,89 and

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decrease helicase activity, resulting in a defect in NER.90 Several XPD polymorphisms in the coding regions with relatively high MAFs were recently identified,91 including two nsSNPs, G23592A (Asp312Asn) in exon 10 and A35931C (Lys751Gln) in exon 23. The Lys751Gln polymorphism, which is about 50 bases upstream from the poly (A) signal, has been suggested to be associated with higher levels of chromatid aberrations.92 It is reported that 312Asn and 751Gln variant genotypes were consistently associated with a lower DRC phenotype in two studies.7,8 However, it is also reported that 751Gln variant genotypes were associated with a higher DRC phenotype, and 312Asn variant genotypes93 were not associated with an altered DRC phenotype.92,93 Some case-control studies have been conducted in different ethnic populations to investigate the associations between XPD polymorphisms and the risk of cancers.94 However, the results from these molecular epidemiological studies are confusing rather than conclusive. In two recently published meta-analyses, an elevated lung cancer risk associated with XPD 312Asn and 751Gln variant alleles were confined to the fixed combination model but not in the random effect model,95,96 while in another study, XPD Lys751Gln polymorphism was found not to be associated with head and neck cancer risk in a multi-center study with 555 cases and 792 controls.88 In the prospective Nurses’ Health Study, the variant 751Gln and 312Asn genotypes were found to be associated with a decreased risk of melanoma and squamous cell carcinoma but not with basal cell carcinoma.97 Furthermore, the association of the 751Gln allele with melanoma risk was significantly modified by lifetime severe sunburns, cumulative sun exposure with a bathing suit and a constitutional susceptibility score.97 3.2.4. XRCC3 XRCC3 is located on chromosome 14q32.3 and structurally related to Rad51, participating in homologous recombinational repair of DNA double-strand breaks and crosslinks98,99 (also see Chapter 5 for more details). XRCC3-deficient cells do not form Rad51 foci after radiation

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damage; they exhibit genetic instability and increased sensitivity to DNA-damaging agents such as ultraviolet light.100 XRCC3 has an nsSNP of C→T substitution at position 18067 in exon 7 that results in an amino acid change (Thr241Met),91 which is thought to have possible functional relevance and/or the interaction with other proteins involved in DNA damage and repair. Previously, two small case-control studies reported that the XRCC3 Thr241Met polymorphism is associated with the risk of melanoma101 and bladder cancer79 ; this association was substantiated in a large case-control study of breast cancer.102 However, such associations were not supported by recent reports of both prospective and retrospective studies with relative large sample sizes.103–106 4. CONCLUSIONS AND PERSPECTIVES As summarized in Fig. 1, although some phenotypic studies were good at predicting cancer risk, they had relatively large variations in risk estimates because of the large assay variation in addition to the relatively small number of subjects included in the studies. On the other hand, recent meta-analyses suggest that most reported genotype-risk associations are not withstanding further testing, and these false positive associations are probably responsible for the bulk of the failure to replicate the associations between common variants and complex traits.107 Using the labor-intensive phenotypic assays as “intermediates,” the authors’ long-term goal is to identify genotypes that predict the phenotypes of suboptimal DRC, incurred adduct formation, and altered gene expression. With high-throughput genotyping methods becoming available, mature, and more affordable, genome-wide association approaches should be feasible in the years to come, which may provide formidable challenges in throughput-data processing and analysis. Candidate gene/SNP approaches will remain critical to confirm the causal relationships of specific SNPs or genes in the regions identified by genome-wide or LD-based approaches. Furthermore, knowledge of the functional significance of SNPs is key to understanding the biological basis of an epidemiological

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Head and Neck cancer: DRC (20) ERCC1 RNA (39) XPB/ERCC3 RNA (39) XPG/ERCC5 RNA (39) CSB/ERCC6 RNA (39) XPC RNA (39) ERCC1 Protein (40) XPA Protein (40) XPC Protein (40) XPD Protein (40) XPF Protein (40) XPG Protein (40)

15.7 7.9

7.3

Lung cancer: DRC (17) DRC (18) BPDE-DNA adducts (26) ERCC1 RNA (38) XPB/ERCC3 RNA (38) XPG/ERCC5 RNA (38) CSB/ERCC6 RNA (38) XPC RNA (38)

15.7 29.4

Cutaneous malignant melanoma: DRC (16)

0.0

1.0

2.0

3.0 4.0 Odds Ratio

5.0

6.0

7.0

Fig. 1

association in respect to the function to be determined before or after the association studies. Powerful statistical and computational methods need to be developed to model the relationship in genegene and gene-environment interactions, and disease susceptibility, especially for genome-wide association studies.

4.1. Statistical Power and Multiple Testing Significance thresholds in the order of P < 10−6 have been proposed for genome-wide association studies, owing to the need to allow for the very small prior probability that any given locus or region is truly associated with a disease.108 False positives can arise from statistical fluctuation and the widespread, inappropriate use of P values below 0.05 as a criterion for declaring a success. Ideally, reported estimates of statistical significance should reflect the likelihood of observing a particular P value, taking into account all of the multiple-tested

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hypotheses (a “frequentist” approach), or alternatively, the likelihood of a particular association being correct in light of the P value and the prior probability of the association being true (a “Bayesian” approach).109 Practically, it will be difficult to achieve “statistical significance” in a single genome-wide association study for the given P value. However, permutation analysis can provide an empirical pool of the observed data110 and false-discovery-rate analysis may also prove to be useful.111 In addition, ranking markers with functional relevance will always be an efficient way to find out a true association. 4.2. Ethnic Difference (Population Stratification) The simplest study design for assessing genotype-phenotype correlation is the traditional case-control approach. This design carries the strong assumption that any noted differences in allele frequencies may actually be related to the outcome measured without any unobserved confounding effects. However, allele frequencies are known to vary widely within and between populations because each population has a unique genetic background, and ancestral patterns of geographical migration, mating practices, reproductive expansions and bottlenecks, and stochastic variation all yield differences in allele frequencies. Besides the commonly used self-reported race as a controlling variable for ancestry, recently, the notion of using anonymous genetic markers as indicators of the amount of ethnic background diversity in the cases and controls was developed and recommended.112,113 In addition, the family-based matching designs and corresponding statistical methods still provide a powerful control tool for population stratification.114 4.3. Higher-order Gene-Gene and Gene-Environmental Interactions True heterogeneity (a failure to replicate an initial reported association) of observational studies between populations may exist

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because of modifier genes and environmental confounders. Identifying such modifiers is a principal challenge for the future, and helps resolve, in general, the genetics of complex diseases, including cancer, for which all the contributing loci might be thought of as “modifiers,” because no single locus of large effect exists. Multipletesting becomes an even more intractable problem when the ultimate utility of whole-genome association studies allows for gene-gene or gene-environment interactions. However, gene-gene interactions in the risk assessment, in general, are more biologically plausible between genes involved in a physical interaction, found in the same pathway or involved in the same regulatory network. It would be reasonable to test for genegene interactions between all pairs of non-synonymous cSNPs in physically interacting genes in any pathways. Such a strategy, of course, may be influenced by the current lack of knowledge about certain biological processes. An effort to reproduce the obtained gene-gene interactions could use experimental systems or computer simulations, with the latter requiring improved bioinformatics tools to expertly annotate the known pathways of physical interactions.115 ACKNOWLEDGMENTS We have attempted to cite as many references as possible and apologize if some appropriate references have been inadvertently omitted. This work was in part supported by the National Institutes of Health National Cancer Institute grants R01 CA 100264 (to QW) and the National Institute of Environmental Health Sciences grant R01 ES11740 (to QW). We thank Ms Monica Domingue for assistance in preparation of the manuscript. References 1. Sancar A. (1995) DNA repair in humans. Annu Rev Genet 29: 69–105. 2. Wood RD, Mitchell M, Sgouros J, Lindahl T. (2001) Human DNArepair genes. Science 291: 1284–1289. 3. Pero RW, Bryngelsson C, Mitelman F, et al. (1978) Inter-individual variation in the responses of cultured human lymphocytes to exposure

FA

October 4, 2006 15:45

336

4. 5.

6.

7.

8.

9.

10.

11.

12.

13.

14.

15.

16.

WSPC/SPI-B414: DNA Repair, Genetic Instability & Cancer

ch12

Zhibin Hu, Li-E Wang and Qingyi Wei

from DNA damaging chemical agents: inter-individual variation to carcinogen exposure. Mutat Res 53: 327–341. Friedberg EC, Walker GC, Siede W. (1995) DNA repair and mutagenesis. ASM Press, Washington (DC). Wei Q. (1998) Effect of aging on DNA repair and skin carcinogenesis: a mini-review of population-based studies. J Investig Dermatol Symp Proc 3: 19–22. Neumann AS, Sturgis EM, Wei Q. (2005) Nucleotide excision repair as a marker for susceptibility to tobacco-related cancers: a review of molecular epidemiological studies. Mol Carcinog 42: 65–92. Qiao Y, Spitz MR, Guo Z, et al. (2002) Rapid assessment of repair of ultraviolet DNA damage with a modified host-cell reactivation assay using a luciferase reporter gene and correlation with polymorphisms of DNA repair genes in normal human lymphocytes. Mutat Res 509: 165–174. Spitz MR, Wei Q, Dong Q, et al. (2003) Genetic susceptibility to lung cancer: the role of DNA damage and repair. Cancer Epidemiol Biomarkers Prev 12: 689–698. Athas WF, Hedayati MA, Matanoski GM, et al. (1991) Development and field-test validation of an assay for DNA repair in circulating human lymphocytes. Cancer Res 51: 5786–5793. Wei Q, Matanoski GM, Farmer ER, et al. (1993) DNA repair and aging in basal cell carcinoma: a molecular epidemiology study. Proc Natl Acad Sci USA 90: 1614–1618. Protic-Sabljic M, Kraemer KH. (1985) One pyrimidine dimer inactivates expression of a transfected gene in xeroderma pigmentosum cells. Proc Natl Acad Sci USA 82: 6622–6626. Hall J, English DR, Artuso M, et al. (1994) DNA repair capacity as a risk factor for non-melanocytic skin cancer — a molecular epidemiological study. Int J Cancer 58: 179–184. D’Errico M, Calcagnile A, Iavarone I, et al. (1999) Factors that influence the DNA repair capacity of normal and skin cancer-affected individuals. Cancer Epidemiol Biomarkers Prev 8: 553–559. Matta JL, Villa JL, Ramos JM, et al. (2003) DNA repair and nonmelanoma skin cancer in Puerto Rican populations. J Am Acad Dermatol 49: 433–439. Landi MT, Baccarelli A, Tarone RE, et al. (2002) DNA repair, dysplastic nevi, and sunlight sensitivity in the development of cutaneous malignant melanoma. J Natl Cancer Inst 94: 94–101. Wei Q, Lee JE, Gershenwald JE, et al. (2003) Repair of UV light-induced DNA damage and risk of cutaneous malignant melanoma. J Natl Cancer Inst 95: 308–315.

FA

October 4, 2006 15:45

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ch12

337

17. Wei Q, Cheng L, Hong WK, et al. (1996) Reduced DNA repair capacity in lung cancer patients. Cancer Res 56: 4103–4107. 18. Wei Q, Cheng L, Amos CI, et al. (2000) Repair of tobacco carcinogeninduced DNA adducts and lung cancer risk: a molecular epidemiologic study. J Natl Cancer Inst 92: 1764–1772. 19. Shen H, Spitz MR, Qiao Y, et al. (2003) Smoking, DNA repair capacity and risk of non-small cell lung cancer. Int J Cancer 107: 84–88. 20. Cheng L, Eicher SA, Guo Z, et al. (1998) Reduced DNA repair capacity in and neck cancer patients. Cancer Epidemiol Biomarkers Prev 7: 465– 468. 21. Hu JJ, Hall MC, Grossman L, et al. (2004) Deficient nucleotide excision repair capacity enhances human prostate cancer risk. Cancer Res 64: 1197–1201. 22. Shi Q, Wang LE, Bondy ML, et al. (2004) Reduced DNA repair of benzo[a]pyrene diol epoxide-induced adducts and common XPD polymorphisms in breast cancer patients. Carcinogenesis 25: 1695– 1700. 23. Ahn B, Kang D, Kim H, et al. (2004) Repair of mitomycin C crosslinked DNA in mammalian cells measured by a host cell reactivation assay. Mol Cells 18: 249–255. 24. Everson RB, Randerath E, Santella RM, et al. (1986) Detection of smoking-related covalent DNA adducts in human placenta. Science 231: 54–57. 25. Geneste O, Camus AM, Castegnaro M, et al. (1991) Comparison of pulmonary DNA adduct levels, measured by 32P-postlabelling and aryl hydrocarbon hydroxylase activity in lung parenchyma of smokers and ex-smokers. Carcinogenesis 12: 1301–1305. 26. Li D, Wang M, Cheng L, et al. (1996) In vitro induction of benzo(a)pyrene diol epoxide-DNA adducts in peripheral lymphocytes as a susceptibility marker for human lung cancer. Cancer Res 56: 3638–3641. 27. Phillips DH. (2002) Smoking-related DNA and protein adducts in human tissues. Carcinogenesis 23: 1979–2004. 28. Phillips DH, Hemminki K, Alhonen A, et al. (1988) Monitoring occupational exposure to carcinogens: detection by 32P-postlabelling of aromatic DNA adducts in white blood cells from iron foundry workers. Mutat Res 204: 531–541. 29. Phillips DH, Schoket B, Hewer A, et al. (1990) Influence of cigarette smoking on the levels of DNAadducts in human bronchial epithelium and white blood cells. Int J Cancer 46: 569–575. 30. Palli D, Vineis P, Russo A, et al. (2000) Diet, metabolic polymorphisms and dna adducts: the EPIC-Italy cross-sectional study. Int J Cancer 87: 444–451.

FA

October 4, 2006 15:45

338

WSPC/SPI-B414: DNA Repair, Genetic Instability & Cancer

ch12

Zhibin Hu, Li-E Wang and Qingyi Wei

31. Peluso M, Airoldi L, Magagnotti C, et al. (2000) White blood cell DNA adducts and fruit and vegetable consumption in bladder cancer. Carcinogenesis 21: 183–187. 32. Hu JJ, Roush GC, Berwick M, et al. (1996) Effects of dietary supplementation of alpha-tocopherol on plasma glutathione and DNArepair activities. Cancer Epidemiol Biomarkers Prev 5: 263–270. 33. Veglia F, Matullo G, Vineis P. (2003) Bulky DNA adducts and risk of cancer: a meta-analysis. Cancer Epidemiol Biomarkers Prev 12: 157–160. 34. Kennedy DO, Agrawal M, Shen J, et al. (2005) DNA repair capacity of lymphoblastoid cell lines from sisters discordant for breast cancer. J Natl Cancer Inst 97: 127–132 35. Wei Q, Xu X, Cheng L, et al. (1995) Simultaneous amplification of four DNA repair genes and beta-actin in human lymphocytes by multiplex reverse transcriptase-PCR. Cancer Res 55: 5025–5029. 36. Wei Q, Guan Y, Cheng L, et al. (1997) Expression of five selected human mismatch repair genes simultaneously detected in normal and cancer cell lines by a non-radioactive multiplex reverse transcriptionpolymerase chain reaction. Pathobiology 65: 293–300. 37. Cheng L, Guan Y, Li L, et al. (1999) Expression in normal human tissues of five nucleotide excision repair genes measured simultaneously by multiplex reverse transcription-polymerase chain reaction. Cancer Epidemiol Biomarkers Prev 8: 801–807. 38. Cheng L, Spitz MR, Hong WK, et al. (2000) Reduced expression levels of nucleotide excision repair genes in lung cancer: a case-control analysis. Carcinogenesis 21: 1527–1530. 39. Cheng L, Sturgis EM, Eicher SA, et al. (2002) Expression of nucleotide excision repair genes and the risk for squamous cell carcinoma of the head and neck. Cancer 94: 393–397. 40. Wei Q, Wang LE, Sturgis EM, et al. (2005) Expression of nucleotide excision repair proteins in lymphocytes as a marker of susceptibility to squamous cell carcinomas of the head and neck. Cancer Epidemiol Biomarkers Prev 14: 1961–1966. 41. Kruglyak L, Nickerson DA. (2001) Variation is the spice of life. Nat Genet 27: 234–236. 42. Risch N, Merikangas K. (1996) The future of genetic studies of complex human diseases. Science 273: 1516–1517. 43. Lander ES. (1996) The new genomics: global views of biology. Science 274: 536–539. 44. Lichtenstein P, Holm NV, Verkasalo PK, et al. (2000) Environmental and heritable factors in the causation of cancer — analyses of cohorts of twins from Sweden, Denmark, and Finland. N Engl J Med 343: 78–85.

FA

October 4, 2006 15:45

WSPC/SPI-B414: DNA Repair, Genetic Instability & Cancer

Molecular Epidemiology of DNA Repair and Cancer Susceptibility

ch12

339

45. Anglian Breast Cancer Study Group. (2000) Prevalence and penetrance of BRCA1 and BRCA2 mutations in a population-based series of breast cancer cases. Br J Cancer 83: 1301–1308. 46. Pharoah PD, Antoniou A, Bobrow M, et al. (2002) Polygenic susceptibility to breast cancer and implications for prevention. Nat Genet 31: 33–36. 47. Lander ES, Schork NJ. (1994) Genetic dissection of complex traits. Science 265: 2037–2048. 48. Risch NJ. (2000) Searching for genetic determinants in the new millennium. Nature 405: 847–856. 49. Peltonen L, McKusick VA. (2001) Genomics and medicine. Dissecting human disease in the postgenomic era. Science 291: 1224–1229. 50. Daly MJ, Rioux JD, Schaffner SF, et al. (2001) High-resolution haplotype structure in the human genome. Nat Genet 29: 229–232. 51. Patil N, Berno AJ, Hinds DA, et al. (2001) Blocks of limited haplotype diversity revealed by high-resolution scanning of human chromosome 21. Science 294: 1719–1723. 52. Gabriel SB, Schaffner SF, Nguyen H, et al. (2002) The structure of haplotype blocks in the human genome. Science 296: 2225–2229. 53. Altshuler D, Brooks LD, Chakravarti A, et al. (2005) A haplotype map of the human genome. Nature 437: 1299–1320. 54. Neale BM, Sham PC. (2004) The future of association studies: genebased analysis and replication. Am J Hum Genet 75: 353–362. 55. Rebbeck TR, Spitz M, Wu X. (2004) Assessing the function of genetic variants in candidate gene association studies. Nat Rev Genet 5: 589– 597. 56. Botstein D, Risch N. (2003) Discovering genotypes underlying human phenotypes: past successes for Mendelian disease, future approaches for complex disease. Nat Genet 33(Suppl): 228–237. 57. Ng PC, Henikoff S. (2001) Predicting deleterious amino acid substitutions. Genome Res 11: 863–874. 58. Ramensky V, Bork P, Sunyaev S. (2002) Human non-synonymous SNPs: server and survey. Nucl Acids Res 30: 3894–3900. 59. Zhu Y, Spitz MR, Amos CI, et al. (2004) An evolutionary perspective on single-nucleotide polymorphism screening in molecular cancer epidemiology. Cancer Res 64: 2251–2257. 60. Sanyal S, Festa F, Sakano S, et al. (2004) Polymorphisms in DNA repair and metabolic genes in bladder cancer. Carcinogenesis 25: 729–734. 61. Jeon HS, Kim KM, Park SH, et al. (2003) Relationship between XPG codon 1104 polymorphism and risk of primary lung cancer. Carcinogenesis 24: 1677–1681.

FA

October 4, 2006 15:45

340

WSPC/SPI-B414: DNA Repair, Genetic Instability & Cancer

ch12

Zhibin Hu, Li-E Wang and Qingyi Wei

62. Yamamoto H, Hanafusa H, Ouchida M, et al. (2005) Single nucleotide polymorphisms in the EXO1 gene and risk of colorectal cancer in a Japanese population. Carcinogenesis 26: 411–416. 63. Hu Z, Ma H, Chen F, et al. (2005) XRCC1 polymorphisms and cancer risk: a meta-analysis of 38 case-control studies. Cancer Epidemiol Biomarkers Prev 14: 1810–1818. 64. Pritchard JK, Cox NJ. (2002) The allelic architecture of human disease genes: common disease-common variant or not? Hum Mol Genet 11: 2417–2423. 65. Blangero J. (2004) Localization and identification of human quantitative trait loci: king harvest has surely come. Curr Opin Genet Dev 14: 233–240. 66. Weiss KM, Clark AG. (2002) Linkage disequilibrium and the mapping of complex human traits. Trends Genet 18: 19–24. 67. Smith DJ, Lusis AJ. (2002) The allelic structure of common disease. Hum Mol Genet 11: 2455–2461. 68. Wang WY, Pike N. (2004) The allelic spectra of common diseases may resemble the allelic spectrum of the full genome. Med Hypotheses 63: 748–751. 69. Frank SA. (2004) Genetic predisposition to cancer — insights from population genetics. Nat Rev Genet 5: 764–772. 70. Shiloh Y. (1997) Ataxia-telangiectasia and the Nijmegen breakage syndrome: related disorders but genes apart. Annu Rev Genet 31: 635–662. 71. Savitsky K, Bar-Shira A, Gilad S, et al. (1995) A single ataxia telangiectasia gene with a product similar to PI-3 kinase. Science 268: 1749–1753. 72. Angele S, Romestaing P, Moullan N, et al. (2003) ATM haplotypes and cellular response to DNA damage: association with breast cancer risk and clinical radiosensitivity. Cancer Res 63: 8717–8725. 73. Lee KM, Choi JY, Park SK, et al. (2005) Genetic polymorphisms of ataxia telangiectasia mutated and breast cancer risk. Cancer Epidemiol Biomarkers Prev 14: 821–825. 74. Tamimi RM, Hankinson SE, Spiegelman D, et al. (2004) Common ataxia telangiectasia mutated haplotypes and risk of breast cancer: a nested case-control study. Breast Cancer Res 6: R416-R422. 75. Masson M, Niedergang C, Schreiber V, et al. (1998) XRCC1 is specifically associated with poly(ADP-ribose) polymerase and negatively regulates its activity following DNA damage. Mol Cell Biol 18: 3563– 3571. 76. Fan J, Otterlei M, Wong HK, et al. (2004) XRCC1 co-localizes and physically interacts with PCNA. Nucleic Acids Res 32: 2193–2201.

FA

October 4, 2006 15:45

WSPC/SPI-B414: DNA Repair, Genetic Instability & Cancer

Molecular Epidemiology of DNA Repair and Cancer Susceptibility

ch12

341

77. Lunn RM, Langlois RG, Hsieh LL, et al. (1999) XRCC1 polymorphisms: effects on aflatoxin B1-DNA adducts and glycophorin A variant frequency. Cancer Res 59: 2557–2561. 78. Duell EJ, Wiencke JK, Cheng TJ, et al. (2000) Polymorphisms in the DNA repair genes XRCC1 and ERCC2 and biomarkers of DNA damage in human blood mononuclear cells. Carcinogenesis 21: 965–971. 79. Matullo G, Guarrera S, Carturan S, et al. (2001) DNA repair gene polymorphisms, bulky DNA adducts in white blood cells and bladder cancer in a case-control study. Int J Cancer 92: 562–567. 80. Lei YC, Hwang SJ, Chang CC, et al. (2002) Effects on sister chromatid exchange frequency of polymorphisms in DNA repair gene XRCC1 in smokers. Mutat Res 519: 93–101. 81. Hsieh LL, Chien HT, Chen IH, et al. (2003) The XRCC1 399Gln polymorphism and the frequency of p53 mutations in Taiwanese oral squamous cell carcinomas. Cancer Epidemiol Biomarkers Prev 12: 439–443. 82. Hu JJ, Smith TR, Miller MS, et al. (2001) Amino acid substitution variants of APE1 and XRCC1 genes associated with ionizing radiation sensitivity. Carcinogenesis 22: 917–922. 83. Taylor RM, Thistlethwaite A, Caldecott KW. (2002) Central role for the XRCC1 BRCT I domain in mammalian DNA single-strand break repair. Mol Cell Biol 22: 2556–2563. 84. Pastorelli R, Cerri A, Mezzetti M, et al. (2002) Effect of DNArepair gene polymorphisms on BPDE-DNA adducts in human lymphocytes. Int J Cancer 100: 9–13. 85. Gao WM, Romkes M, Day RD, et al. (2003) Association of the DNA repair gene XPD Asp312Asn polymorphism with p53 gene mutations in tobacco-related non-small cell lung cancer. Carcinogenesis 24: 1671– 1676. 86. Hou SM, Ryk C, Kannio A, et al. (2003) Influence of common XPD and XRCC1 variant alleles on p53 mutations in lung tumors. Environ Mol Mutagen 41: 37–42. 87. Wang Y, Spitz MR, Zhu Y, et al. (2003) From genotype to phenotype: correlating XRCC1 polymorphisms with mutagen sensitivity. DNA Repair 2: 901–908. 88. Huang WY, Olshan AF, Schwartz SM, et al. (2005) Selected genetic polymorphisms in MGMT, XRCC1, XPD, and XRCC3 and risk of head and neck cancer: a pooled analysis. Cancer Epidemiol Biomarkers Prev 14: 1747–1753. 89. Coin F, Marinoni JC, Rodolfo C, et al. (1998) Mutations in the XPD helicase gene result in XP and TTD phenotypes, preventing interaction between XPD and the p44 subunit of TFIIH. Nat Genet 20: 184–188.

FA

October 4, 2006 15:45

342

WSPC/SPI-B414: DNA Repair, Genetic Instability & Cancer

ch12

Zhibin Hu, Li-E Wang and Qingyi Wei

90. Winkler GS, Araujo SJ, Fiedler U, et al. (2000) TFIIH with inactive XPD helicase functions in transcription initiation but is defective in DNA repair. J Biol Chem 275: 4258–4266. 91. Shen MR, Jones IM, Mohrenweiser H. (1998) Nonconservative amino acid substitution variants exist at polymorphic frequency in DNA repair genes in healthy humans. Cancer Res 58: 604–608. 92. Lunn RM, Helzlsouer KJ, Parshad R, et al. (2000) XPD polymorphisms: effects on DNA repair proficiency. Carcinogenesis 21: 551–555. 93. Moller P, Knudsen LE, Frentz G, et al. (1998) Seasonal variation of DNA damage and repair in patients with non-melanoma skin cancer and referents with and without psoriasis. Mutat Res 407: 25–34. 94. Goode EL, Ulrich CM, Potter JD. (2002) Polymorphisms in DNArepair genes and associations with cancer risk. Cancer Epidemiol Biomarkers Prev 11: 1513–1530. 95. Hu Z, Wei Q, Wang X, et al. (2004) DNA repair gene XPD polymorphism and lung cancer risk: a meta-analysis. Lung Cancer 46: 1–10. 96. Benhamou S, Sarasin A. (2005) ERCC2 /XPD gene polymorphisms and lung cancer: a HuGE review. Am J Epidemiol 161: 1–14. 97. Han J, Colditz GA, Liu JS, et al. (2005) Genetic variation in XPD, sun exposure, and risk of skin cancer. Cancer Epidemiol Biomarkers Prev 14: 1539–1544. 98. Tebbs RS, Zhao Y, Tucker JD, et al. (1995) Correction of chromosomal instability and sensitivity to diverse mutagens by a cloned cDNA of the XRCC3 DNA repair gene. Proc Natl Acad Sci USA 92: 6354–6358. 99. Liu N, Lamerdin JE, Tebbs RS, et al. (1998) XRCC2 and XRCC3, new human Rad51-family members, promote chromosome stability and protect against DNA crosslinks and other damages. Mol Cell 1: 783– 793. 100. Masson JY, Stasiak AZ, Stasiak A, et al. (2001) Complex formation by the human RAD51C and XRCC3 recombination repair proteins. Proc Natl Acad Sci USA 98: 8440–8446. 101. Winsey SL, Haldar NA, Marsh HP, et al. (2000) A variant within the DNA repair gene XRCC3 is associated with the development of melanoma skin cancer. Cancer Res 60: 5612–5616. 102. Kuschel B, Auranen A, McBride S, et al. (2002) Variants in DNAdoublestrand break repair genes and breast cancer susceptibility. Hum Mol Genet 11: 1399–1407. 103. Auranen A, Song H, Waterfall C, et al. (2005) Polymorphisms in DNA repair genes and epithelial ovarian cancer risk. Int J Cancer 117: 611– 618. 104. Webb PM, Hopper JL, Newman B, et al. (2005) Double-strand break repair gene polymorphisms and risk of breast or ovarian cancer. Cancer Epidemiol Biomarkers Prev 14: 319–323.

FA

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WSPC/SPI-B414: DNA Repair, Genetic Instability & Cancer

Molecular Epidemiology of DNA Repair and Cancer Susceptibility

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343

105. Tranah GJ, Giovannucci E, Ma J, et al. (2004) XRCC2 and XRCC3 polymorphisms are not associated with risk of colorectal adenoma. Cancer Epidemiol Biomarkers Prev 13: 1090–1091. 106. Han J, Hankinson SE, Ranu H, et al. (2004) Polymorphisms in DNA double-strand break repair genes and breast cancer risk in the Nurses’ Health Study. Carcinogenesis 25: 189–195. 107. Ioannidis JP, Ntzani EE, Trikalinos TA, et al. (2001) Replication validity of genetic association studies. Nat Genet 29: 306–309. 108. Hirschhorn JN, Daly MJ. (2005) Genome-wide association studies for common diseases and complex traits. Nat Rev Genet 6: 95–108. 109. Newton-Cheh C, Hirschhorn JN. (2005) Genetic association studies of complex traits: design and analysis issues. Mutat Res 573: 54–69. 110. Doerge RW, Churchill GA. (1996) Permutation tests for multiple loci affecting a quantitative character. Genetics 142: 285–294. 111. Storey JD, Tibshirani R. (2003) Statistical significance for genome-wide studies. Proc Natl Acad Sci USA 100: 9440–9445. 112. Pritchard JK, Rosenberg NA. (1999) Use of unlinked genetic markers to detect population stratification in association studies. Am J Hum Genet 65: 220–228. 113. Bacanu SA, Devlin B, Roeder K. (2000) The power of genomic control. Am J Hum Genet 66: 1933–1944. 114. Jorde LB. (2000) Linkage disequilibrium and the search for complex disease genes. Genome Res 10: 1435–1444. 115. Mayo M. (2005) Learning Petri net models of non-linear gene interactions. Biosystems 82: 74–82.

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INDEX

N-methyl-N  -nitro-Nnitrosoguanidine (MNNG), 254 N2 ,3-ethanoguanine (N2 ,3-εG), 36 hMLH1, 99–101, 104, 105, 107 hMSH2, 98–101, 104 hMSH3, 99, 106, 107 hMSH6, 100, 102, 106, 107 hPMS2, 99, 100, 322 Apc, 39, 49 Caenorhabditis elegans, 251 MYH, 50 RAD6 pathway, 237 Schizosaccharomyces pombe, 251 “mediator” proteins, 123

α-OGG1, 41 β-elimination 27 β,δ-elimination, 27, 42 β-OGG1, 41 1,N2 -ethanoguanine (1,N2 -εG), 36 1,N6 -ethanoadenine (EA), 36 3,N4 -ethenocytosine (εC), 34 3-methyladenine (3-meA), 35 3-methylcholanthrene, 251 5 -deoxyribose-5-phosphate (5 -dRp), 27 5-fluorouracil (5-FU), 38 5-formyluracil (5-FoU), 36 5-hydroxymethyluracil (5-HmU), 38 5-hydroxymethyluracil (5-HmU)-DNA glycosylase, 38 5-hydroxyuracil (5-OHU), 38 5-meC-DNA glycosylase, 39 5-meCpG sites, 39 5,6-dihydrouracil (5,6-DHU), 42, 44, 52 5,6-dihydrothymine (5,6-DHT), 42, 52 7-methylguanine (7-meG), 35 7,8-dihydro-8-oxoguanine (8-oxoG), 27, 36, 41–43, 47, 49, 50, 52, 53 8-oxoG-DNA glycosylase 1 (OGG1), 27, 29, 32, 41 32 P-postlabeling, 319 (6-4) photoproducts, 249 N-glycosyl bond, 24, 34

A AAG, 26, 29 abasic site, 245, 247, 250, 252, 255 AlkA, 32, 35, 36, 40 alkylating agents, 35, 36, 40 alkylation adaptation, 35 alkylated bases, 35, 36 allele-specific gene expression, 306–309 alloxan, 38 alternative splicing, 30 345

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Index

346

Alu repeat, 142 amino acid substitution, 322, 329 animal models, 23, 25, 48 ANPG, 26, 29 AP endonuclease, 23–25, 27, 29, 43–46, 52, 204, 205, 209, 210, 222, 271, 272, 277 AP endonuclease 1 (APE1), 25, 27, 29, 44, 45, 47, 48 AP endonuclease 2 (APE2), 29, 44 APEX1, 25, 29 APNG, 26, 29 apurinic/apyrimidinic (AP) sites, 23 artemis, 160, 163, 164, 169, 275, 276 arylhydrocarbon receptor, 251 association study, 326 at-risk individual, 316 at-risk population, 316 ataxia telangiectasia, 316, 328 ataxia-telangeictasia-mutated (ATM), 2, 4–8, 10–15, 194–196 ATR, 4, 6, 8, 10, 11, 13–15, 183, 187, 194, 195

B B lymphocytes, 255 B[a]P, 250, 251 BACH1, 181, 191 back-up activity on repair, 48 bacteriophage T7 polymerase, 244 BARD1, 194 base excision repair (BER), 23, 24, 52, 109, 143, 145, 203, 205, 207, 213, 215, 216, 222, 224–227, 240, 252, 267, 268, 271, 277, 286, 322, 323 base flipping, 34 base modification, 243 base unstacking, 209, 214, 216, 221, 227

base-base mismatch, 89, 91, 93, 94 Bayesian approach, 334 Bayesian polygenetic analyses, 327 BCCIP, 137 benzetheno adducts, 44, benzo[a]pyrene (B[a]P) diol epoxide (BPDE), 251 benzo[a]pyrene, 315 BER subpathways, 26 between-study heterogeneity, 330 bifunctional glycosylases, 34, 41, 42 biological plausibility, 326 bladder cancer, 320, 327, 332 bleomycin, 330 BLM, 178, 192, 193, 198 block-like structure, 326 Bloom’s syndrome, 178, 316 BRC repeats, 181, 190 BRCA1, 5, 8–10, 12, 14, 15, 135, 137–142, 177, 179, 183, 189–194 BRCA2, 123, 125, 135–138, 140–143, 177, 179, 183, 189–194, 197 BRCT repeats, 191, 193 breast cancer, 177, 179, 189, 192, 194, 320, 323, 329, 332 BRIP1, 177, 191, 194 bronchial epithelium, 319 bronchioalveolar lavage cell, 319 budding yeast, 233, 237, 240, 243, 253, 260

C cancer predisposition, 329 cancer prevention, 141, 316 cancer risk, 316, 320, 321, 327, 328, 330–332 cancer susceptibility, 7, 9, 16, 17 candidate gene, 326, 332 carcinogen exposure, 319 carcinogenesis, 257, 260, 316 case-control study, 320, 332 cell cycle, 1, 2, 10–14, 16

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Index

cell cycle checkpoint, 119, 121, 139, 140, 145 cell cycle delay, 330 cell death, 257 cerebellar degeneration, 328 chemotherapy, 106, 112, 131, 158, 170, 179, 319 checkpoints, 1, 2, 9, 12, 13 chloramphenicol acetyltransferase, 317 chromosomal break, 330 chromosomal instability, 316 chromosome replication, 238, 243 class-switch recombination, 255 Cockayne syndrome, 77, 290, coding region, 326, 331 collapsed replication forks, 119, 130, 131, 142 colon cancer, 95, 96, 97, 103 common-disease common-variant hypothesis, 328 complementary overhang, 252 computational algorithms, 327 confidence interval, 318 CpG sites, 37, 39, 40, 49 crossover, 120, 121, 123, 125, 126 cutaneous malignant melanoma, 333 cyclobutane pyrimidine dimer, 249 cytotoxic effect, 317

D damage avoidance, 234 deaminated bases, 23, 24, 37 deamination of 5-methylcytosine, 52 deamination of thymine, 209 deoxyribophosphodiesterase (dRpase), 28, 29, 53, 271 deglycosylation, 255 deletion, 245, 250, 252, 253, 255–257 deoxycytidyl transferase, 238 deoxyribose phosphate, 252 de-ubiquitin, 195 disease causality, 326

index

347

disease-causing variant, 326, 328 DNA adduct, 319, 320, 330 DNA crosslink, 319 DNA damage, 1, 2, 4–6, 8–13, 15–17 DNA damage tolerance (DDT), 233, 234 DNA damaging agents, 332 DNA double-strand breaks (DSBs), 119, 121, 157–159 DNA glycosylase, 23–32, 34, 35, 37, 40, 41, 43–45, 47, 48 DNA helicase II, 89 DNA lesion recognition, 203 DNA lesions, 316 DNA ligase, 23, 24, 27, 30, 89–93, 329 DNA methylation, 302, 303, DNA polymerase, 23, 24, 28, 30, 88, 93, 94, 96, 233, 238, 240, 242–247, 249, 251, 252, 256, 257, 329 DNA polymerase III, 89–91 DNA post-replication repair (PRR), 233, 234 DNA repair, 2, 10, 12, 14, 49, 51, 52, 65, 73, 78–80, 106, 119, 121, 143, 161, 162, 165, 169, 179, 182, 186, 192, 193, 195, 203, 233, 234, 240, 241, 260, 267–269, 271, 277, 286, 289, 290, 292, 305, 315–317, 319–323, 328, DNA repair capacity, 316, 317, 320 DNA repair deficiency, 52 DNA repair machinery, 316 DNA repair phenotype, 317, 319, 322 DNA repair proficiency, 319 DNA replication, 11, 16, 80, 88, 109, 121, 128, 139, 140, 195, 206, 285, 287 DNA-damage checkpoint, 328 DNA-dependent protein kinase (DNA-PK), 161, 162, 164–168 double-strand break repair, 319 downstream, 4, 7, 14, 28, 46, 68, 71, 163, 181, 184 doxorubicin, 251 Drosophila melanogaster, 251

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348

E endonuclease, 181, 186, 255 endonuclease III (Nth), 31 endonuclease III-like 1 (NTH1), 27, 53 endonuclease IV (Nfo), 44 endonuclease VIII (NEI), 30 endonuclease VIII-like (NEIL), 27 endonuclease VIII-like 1 (NEIL1), 29 endonuclease VIII-like 2 (NEIL2), 29 endonuclease VIII-like 3 (NEIL3), 29 endoreduplication, 128, 139 Environmental Genome Project, 315, 321 environmental pollution, 251 enzyme activity, 317 epigenome, 301, 302 epigenetics, 302, 310 ERCC4, 186 Escherichia coli, 234 ethnic diversity, 334 eukaryotic mitochondrial polymerase γ, 244 eukaryotic organisms, 245, 248 excision, 24–26, 32, 34, 40, 42 exocyclic adducts, 35 exon, 329, 331, 332 exonuclease, 89–91, 93, 94, 99 exonuclease 1 (EXO1), 93, 94, 103, 111, 123, 255, 268, 269, 270, 277, 325, 327, exonuclease III (Xth), 44

F familial aggregation, 323 family history, 99 FANCA, 179, 181, 182, 184, 193, 195 FANCB, 178, 181, 185 FANCC, 179, 181, 188 FANCD1, 177, 186, 189, 190, 193, 194

FANCD2, 177, 181–185, 187–191, 193–198 FANCE, 179, 181 FANCF, 179, 181, 194 FANCG, 179, 181, 182, 188, 195 FANCI, 198 FANCJ, 177, 181, 183, 186, 189, 191–194, 198 FANCL, 177, 181, 185, 186, 188, 194 FANCM, 177, 181, 183, 185–187, 192, 193, 195, 198 Fanconi anemia, 177, 183, 189, 198, 316 Fanconi anemia core complex, 184 Fapy-A, 42, 43 Fapy-G, 29, 42, 43 fixed combination model, 331 flap endonuclease-1 (FEN-1), 28, 30, 52, 72, 267, 268, 271, 277, 278–280 formamidopyrimidine-DNA glycosylase, 41 fragmented thymine, 36 frequentist approach, 334 functional significance, 332 functional SNP, 326 functionality, 25

G gene conversion, 120, 126, 142 gene expression, 317, 320, 332 gene transcript, 320 gene-environment interaction, 330, 333–335 gene-gene interaction, 335 general population, 315, 316, 323 genetic polymorphism, 25 genetic markers, 334 genetic syndrome, 328 genetic variant, 328 genome, 317, 321, 328 genome-wide association, 326, 328, 332–334 genomic integrity, 316

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genomics, 301, 302 genotype-phenotype correlation, 334 germ-line mutation, 327, 329 glycophorin A, 330 GO system, 41

index

349

housekeeping gene, 320 Huntington’s disease, 267, 287, 291 hydrogen-bonding interactions, 34 hyper-IgM syndrome, 38 hypoxanthine, 29, 36

I H HAP1, 25, 29 haplotype, 326, 328, 329 Haplotype Map Project, 328 head and neck cancers, 315, 320, 321 Hef, 181, 186, 187 helicase, 177, 178, 181, 182, 186, 187, 191–193, 197, 198 helix-hairpin-helix (HhH) motif, 32 hereditary non-polyposis colorectal cancer (HNPCC), 87, 95–100, 102–104, 106, 107, 111, 290 heterogeneity, 334 high mobility group box 1 protein (HMGB1), 94 high-penetrance mutations, 323 high-throughput technology, 326 hMutLα, 92, 93, 100, 108, 109 hMutS, 99 hMutSα, 92–94, 100, 107–109 homeologous recombination, 87, 110, 142 homologous annealing, 73, 112, 119, 120, 124, 145, 169, 179, 183, 190, 193, 204, 235, 268, 273, 275, 276, 322, 331 homologous recombination (HR), 73, 119, 120, 124, 142, 179, 183, 190, 193, 235, 268, 273, 275, 276 homologous recombinational repair, 331 homologous template, 235 homotrimer, 240 Hoogsteen base-pairing, 250 hospital-based case-control study, 318 host cell reactivation assay, 317

immunodeficiency, 328 immunofluorescence, 320 individual variation, 50 inference, 326, 327 inherited susceptibility, 323 insertion, 245, 253, 255–257 insertion fidelity, 249 insertion deletion mispairs, 89, 91, 102 interaction domain, 329 interstrand crosslink (ICL), 119, 121, 122, 131 interstrand DNA crosslinks, 177, 196 intron, 322 ionizing radiation, 157, 158, 169, 329 isodialuric acid, 38

K kinetic proofreading, 213, 215, 216, 224 knockout mice, 25, 48, 49

L LD mapping, 323 lesion bypass, 233, 234, 242, 245, 248, 249, 252, 254, 257, 260 LIG1, 28, 30, 48 LIG3, 27, 30, 47 linear correlation, 319 linkage analysis, 323

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350

linkage disequilibrium, 323 long-patch BER, 28, 30 loss of heterozygosity (LOH), 120, 142 low fidelity polymerases, 235 luciferase, 317 lung cancer, 318–320

M map-based approach, 326 MDC1, 8–10, 12, 14, 15 MED1, 29, 39 meta-analysis, 320, 328, 330–332 methyl-binding domain glycosylase 4 (MBD4), 29 methylation, 320 methylpurine-DNA glycosylase (MDG), 26 microarray, 301, 304–306, 310 microsatellite instability (MSI), 97 minor allele frequency, 321 minor-groove purine adduct, 250 mismatch, 87, 89, 92–94, 111 mismatch adenine-DNA glycosylase (MutY), 43 mismatch-specific uracil-DNA glycosylase (Mug), 53 mismatch repair (MMR), 49, 87, 88, 91, 123, 126, 142, 145, 203, 206, 213, 223, 227, 240, 255, 267–269, 277, 287, 290, 320, 322, 323, 325 MMR, 204–207, 215, 216, 224–227 MMR-Mediated Apoptosis, 109, 112 molecular dynamics simulations, 32 molecular epidemiology, 315 mono-ubiquitylated substrate, 236 monofunctional glycosylases, 27, 34, 35 MPH1, 186, 187 Mre11, 6, 7, 16, 106, 123, 269, 275–277, 322 mRNA instability, 322 mRNA precursor, 255

Msh2 gene, 49 MTH, 41 multi-center study, 331 multifunctional protein, 238 multiple rare-variant hypothesis, 328 multiple reverse transcriptase-polymerase chain reaction, 320 multiple testing, 333, 335 multiplicative, 326 multivariate logistic regression, 321 Mus81, 186, 269, 275, 276 mutagenesis, 237, 238, 241, 243, 245–248, 251, 254, 259 mutagenic bypass, 238 mutation, 323, 328, 330 mutational hot spots, 244 mutation spectrum, 254 MutH, 88–91 MutL, 88–91, 93, 99, 110, 111 MutS, 88–91, 93, 98 MutT, 41 MutY homolog DNA glycosylase (MYH), 25, 29, 43, 50 MYH-associated polyposis (MAP), 50 mytomycin C, 319

N N-2-aminofluorene, 243 nascent template, 235 National Institutes of Environmental Health Sciences (NIEHS), 321, 329 NCBI database, 257 NER, 203–207, 210, 211, 216, 223–227 non-canonical templates, 257 non-cognate lesion, 257 non-homologous end joining (NHEJ), 122, 157–164, 166–169, 204, 268, 275, 287, 322 nuclear foci, 241, 253, 257, 259 nuclear localization, 257

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nucleases, 71, 72, 90, 94, 122, 123, 133, 135, 205, 215, 267–270, 273, 275, 276, 281, 283, 284, 288, 289, 290–292 nucleotide excision repair, 65–69, 110, 122, 131, 145, 179, 186, 203, 240, 249, 267, 268, 273, 316, 322 nucleotide incorporation, 235, 249, 250 Nurses’ Health Study cohort, 329

O O4 -methylthymine, 243 O6 -methylguanine, 243, 250 odds ratio, 327 Okazaki fragment maturation, 277, 278, 286 oral cancer, 320 oxanine, 36 oxidative damage, 41, 207, oxidative stress, 39, 319 oxidized bases, 29, 31, 40, 41, 44

P PCNA, 28, 30, 38, 44, 47 peripheral blood lymphocyte, 319, 320, permutation analysis, 334 phenotype, 237, 238, 244–246, 249, 256, 260 PHF9, 181, 185 phosphorylation, 321 phytohemagglutinin, 320 pilot study, 318, 319 plasmid, 317 Pol II, 244 Pol III, 244, 254 Pol IV, 235, 246, 247, 251, 253, 254 Pol V, 235, 246, 247, 253, 254 Polα, 243 Polβ, 25, 27–29, 48, 51, 252, 258

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351

Polδ, 28, 30, 240, 243, 244, 250 Polη, 233, 234, 241, 248–253, 256, 258 Polι, 28, 249–253, 256–258 Polκ, 249–253, 256, 258 Polλ, 252, 253, 256, 258 Polµ, 252, 256 Polν, 245 Polθ, 244, 245 Polε, 30, 240 Polζ, 238, 241, 245, 248, 252, 253, 255, 256, 258 poly(ADP-ribose) polymerase 1 (PARP1), 30 polygenic mechanism, 326 polygenic model, 326 polymerase, 329 polymerase binding region, 329 polymorphism, 310 polymorphism phenotyping algorithm, 327 polynucleotide kinase phosphatase (PNKP), 29 PolyPhen, 327 population attributable fraction, 328 population based screening, 319 population stratification, 334 population-based study, 315 proliferating cell nuclear antigen (PCNA), 93, 239 promoter, 320 promoter region, 326 proteasome, 236 protein adduct, 317 protein conjugation, 237 protein expression, 321 protein kinase activity, 329 protein post-translational modifications, 47 protein-protein interactions, 23–25, 28, 31, 45–47, 51, 246, 248 proteolysis, 236

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352

R RAD51, 123, 125, 133, 135, 136, 138–141, 143, 181, 190–192 RAD52, 123, 135, 138, 143, 144 RAD6 pathway, 239 radiosensitivity, 329 radiotherapy, 319 random effect model, 331 rate-limiting factor, 321 reactive oxygen species (ROS), 40 recessive trait, 316 recombinant plasmid, 317 REF-1, 29, 44 regulatory region, 327 repair kinetics, 317 repair synthesis, 26 replication-coupled repair, 43 repeatability, 319 replication arrest, 257 replication error, 247 replication factor C (RFC), 28, 30, 94 replication fork, 233, 235, 238, 252–254, 259 replication protein A (RPA), 30, 93–95 replication-blocking lesion, 233, 234 replicative machinery, 254 replicative polymerase, 253 reporter gene, 317 resynthesis, 90, 92–96, 109 reverse-protein microarray, 321 reversible binding, 226 risk assessment, 335 risk estimate, 318, 332 risk factor, 321

sequence homology, 329 sequence-based approach, 326 sequence variation, 219 short patch, 254 short-patch BER, 27 SIFT, 327 single nucleotide polymorphisms (SNPs), 45, 49, 257, 315 single-strand annealing (SSA), 126 single-strand selective monofunctional uracil-DNA glycosylase (SMUG1), 27 single-stranded DNA binding protein (SSB), 89 sister chromatid exchange, 233 skin cancer, 315, 316 sliding-clamp, 233, 240, 253 slippage, 244 smoker, 319, 320, 330 SNP, 323, 327 somatic hypermutation, 250, 255 sorting intolerant from tolerant, 327 SOS regulon, 235 SOS response, 235, 247, 254 splice site, 326 spontaneous depurination, 44 spontaneous mutagenesis, 239 sporulation, 238 ssDNA-dependent ATPase, 239 stalled replication fork, 128–130, 140 statistical power, 328, 333 subgroups, 77 substitution, 255–257 substrate overlap, 32 SUMO protein, 47 sumoylation, 47, 237, 240, 241 sunburns, 331 synergistic interaction, 239 systems biology, 301, 302, 310, 311

S T S-adenosylmethionine (SAM), sample size, 318, 332 Schiff base, 34 sequence fidelity, 234

35 T:G mismatches, 37, 39 tag, 35 tagging SNP, 326

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targeted gene deletion, 48 targeted mutation, 254 terminal deoxynucleotidyl transferase (TdT), 252 thymine–thymine dimer (TT), 248 thymine, 33, 34, 39, 40 thymine glycol (Tg), 40 thymine-DNA glycosylase (TDG), 27, 29 tobacco smoking, 330 transcription, 317, 330 transcription factors, 40, 47 transfection, 317 transient expression, 317 translesion bypass, 88, translesion DNA synthesis (TLS), 233, 234, 249 translesion efficiency, 248 translesion synthesis, 128, 131, 133 trinucleotide repeat expansion (TNR), 288

U U:A base pair, 37 U:G base pair, 111 Ubc13-Mms2 heterodimer, 237 ubiquitin (Ub), 182, 184, 188, 236 ubiquitination/ubiquitylation, 236 ubiquitin-conjugating enzyme (UBc), 236 ubiquitin-conjugating variant (UEv), 236 ubiquitin ligase, 177, 181–185, 193–195, 236 untargeted mutation, 254, UNG1 (mitochondrial form), 31 UNG2 (nuclear form), 31 upstream, 331 uracil, 32–34, 37–40, 49 uracil glycosylase, 255 uracil-DNA glycosylase (UNG), 27, 29

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353

uracil-DNA glycosylase inhibitor (Ugi), 38 USP1, 196 UV irradiation, 238, 257 UV light sensitivity, 316

W Watson-Crick base-pairing, 249 Werner syndrome, 163, 271, 272, 289, 290 wild-type genotype, 330

X X-ray cross-complementation protein 1 (XRCC1), 28 xanthine, 36 xenobiotic responsive elements (XREs), 251 xeroderma pigmentosum (XP), 65, 66, 75, 234, 248, 257, 273, 289, 290, 316 xeroderma pigmentosum complementation group F (XPF), 273 xeroderma pigmentosum complementation group G (XPG), 273, 290 xeroderma pigmentosum variant (XPV), 248, 257 XPF, 186

Y Y-family polymerases, 253–255

246, 248, 249,

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