DNA Replication and Mutation [1 ed.] 9781619424555, 9781613244906

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Copyright © 2012. Nova Science Publishers, Incorporated. All rights reserved. DNA Replication and Mutation, Nova Science Publishers, Incorporated, 2012. ProQuest Ebook Central,

Copyright © 2012. Nova Science Publishers, Incorporated. All rights reserved. DNA Replication and Mutation, Nova Science Publishers, Incorporated, 2012. ProQuest Ebook Central,

DNA AND RNA: PROPERTIES AND MODIFICATIONS, FUNCTIONS AND INTERACTIONS, RECOMBINATION AND APPLICATIONS

Copyright © 2012. Nova Science Publishers, Incorporated. All rights reserved.

DNA REPLICATION AND MUTATION

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DNA AND RNA: PROPERTIES AND MODIFICATIONS, FUNCTIONS AND INTERACTIONS, RECOMBINATION AND APPLICATIONS

Copyright © 2012. Nova Science Publishers, Incorporated. All rights reserved.

DNA REPLICATION AND MUTATION

RAYMOND P. LEITNER EDITOR

Nova Science Publishers, Inc. New York

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Copyright © 2012 by Nova Science Publishers, Inc. All rights reserved. No part of this book may be reproduced, stored in a retrieval system or transmitted in any form or by any means: electronic, electrostatic, magnetic, tape, mechanical photocopying, recording or otherwise without the written permission of the Publisher. For permission to use material from this book please contact us: Telephone 631-231-7269; Fax 631-231-8175 Web Site: http://www.novapublishers.com NOTICE TO THE READER The Publisher has taken reasonable care in the preparation of this book, but makes no expressed or implied warranty of any kind and assumes no responsibility for any errors or omissions. No liability is assumed for incidental or consequential damages in connection with or arising out of information contained in this book. The Publisher shall not be liable for any special, consequential, or exemplary damages resulting, in whole or in part, from the readers’ use of, or reliance upon, this material. Any parts of this book based on government reports are so indicated and copyright is claimed for those parts to the extent applicable to compilations of such works.

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Independent verification should be sought for any data, advice or recommendations contained in this book. In addition, no responsibility is assumed by the publisher for any injury and/or damage to persons or property arising from any methods, products, instructions, ideas or otherwise contained in this publication. This publication is designed to provide accurate and authoritative information with regard to the subject matter covered herein. It is sold with the clear understanding that the Publisher is not engaged in rendering legal or any other professional services. If legal or any other expert assistance is required, the services of a competent person should be sought. FROM A DECLARATION OF PARTICIPANTS JOINTLY ADOPTED BY A COMMITTEE OF THE AMERICAN BAR ASSOCIATION AND A COMMITTEE OF PUBLISHERS. Additional color graphics may be available in the e-book version of this book. Library of Congress Cataloging-in-Publication Data DNA replication and mutation / editor, Raymond P. Leitner. p. ; cm. Includes bibliographical references and index. ISBN:  (eBook) 1. DNA replication. I. Leitner, Raymond P. [DNLM: 1. DNA Replication. 2. DNA Damage. 3. Mutation. QU 475] QP624.D633 2011 572.8'645--dc23 2011013658

Published by Nova Science Publishers, Inc. † New York

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Contents vii 

Preface Chapter I

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Chapter II

Chapter III

Chapter IV

Connections between BLM and the Fanconi Anemia Pathway in the Repair of Replication Fork Damage Meghan Larin and John Peter McPherson  Mutagenic Potential of Methacrylates Used in Restorative Dentistry Janusz Blasiak, Ewelina Synowiec, Piotr Czarny,Elzbieta Pawlowska and Joanna Szczepanska  DNA Mutation of the Progranulin (GRN) Gene in Familial Frontotemporal Lobar Degeneration (FTLD): A Study of the Pathology of Nine Cases R. A. Armstrong  Effect of a Specific Mammalian DNA Polymerase α-Inhibitor, Dehydroaltenusin, on DNA Replication in Cultured Cells Takeshi Mizuno, Isoko Kuriyama,   Masaharu Takemura, Kengo Sakaguchi, Fumio Sugawara, Hiromi Yoshida and Yoshiyuki Mizushina 

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vi Chapter V

Contents Gel-Based Methods Using DNA-Binding Zinc(II) Complexes for the Detection of DNA Mutations Eiji Kinoshita, Emiko Kinoshita-Kikuta and Tohru Koike 

Chapter VI

DNA Mutations and Genetic Coding Jean-Luc Jestin 

Chapter VII

Nuclear DNA Replication in Trypanosomatid Protozoa M. S. da Silva, R. C. V. da Silveira, A. M. Perez, J. P. Monteiro, S. G. Calderano, J. P. da Cunha, M. C. Elias and M. I. N. Cano 

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Index

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Preface In this book, the authors present current research in the study of DNA replication and mutation, including connections between BLM and the Fanconi anemia pathway in the repair of replication fork damage; the mutagenic potential of methacrylates used in restorative dentistry; DNA mutation of the progranulin gene in familial frontotemporal lobar degeneration; effect of DNA polymerase ainhibitor dehydroaltenusin on DNA replication in cultured cells; gel-based methods of DNA-binding zinc II complexes in the detection of DNA mutations and DNA mutations and genetic coding. Chapter I - The authors DNA is continuously subjected to damage-causing endogenous and exogenous factors, resulting in lesions ranging from interstrand crosslinks to double strand breaks. These and other types of DNA damage encumber the S-phase of the cell cycle, as damaged template DNA can pose a problem for replication fork progression. Stalled replication forks can lead to fork collapse, causing additional DNA damage in the form of double strand breaks. Mammalian cells have developed checkpoint mechanisms that sense and repair DNA damage while maintaining replication fork stability. The process of replication fork restart is complex and involves multiple pathways depending on the lesion type. Initiation of these checkpoints results in cell cycle arrest and consequent stabilization of the replication fork in order to prevent collapse. Repair of damage is believed to occur via translesion synthesis (TLS) or homologous recombination, however the exact mechanism through which these pathways act to maintain genomic stability and accomplish repair of DNA lesions remains unknown. This chapter will highlight pathways required for attempt to describe and evaluate current research into the area of resumption of replication fork stalls in mammalian cells. In this reviewthe authors will focus on the role of helicases and endonucleases in the restart of damaged replication forks, with a particular

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emphasis on interplay between BLM helicase and components of the Fanconi anemia (FA) pathway of DNA repair. Chapter II - DNA damage is a prerequisite for DNA mutation, so the studies on DNA-damaging potential of environmental and medical chemicals are justified. Methacrylates are used in the polymer form as composite restorative materials in dentistry. However, the process of polymerization, led in situ, is always incomplete and the polymers can release monomers into the oral cavity and the pulp, from where they can migrate into the bloodstream reaching virtually all organs. The local concentration of the released monomers can be high enough to induce adverse biological effects. Genotoxicity of methacrylate monomers is of a special significance due to potential serious phenotypic consequences, including mutations and cancer, and a long latency period. In the present work, the authors investigated genotoxicity of a mixture of monomers of model methacrylate composite consisting of 45% 2-hydroxyethyl methacrylate and 55% bisphenol Adiglycidyl dimethacrylate (w/w) (HEMA/Bis-GMA) in human gingival fibroblast by measuring DNA damage and repair, apoptosis and cell cycle. The mixture displayed the ability to damage cellular DNA as measured by the alkaline and neutral comet assay indicating the presence of DNA single-strand breaks, alkalilabile sites and double-strand breaks in DNA. The ability of the composite to induce DNA double-strand breaks, the most serious DNA damage, was confirmed by pulsed field gel electrophoresis. This compound oxidatively modified the DNA bases, as checked by two DNA repair enzymes: endonuclease III and formamidopyrimidine-DNA glycosylase. HEMA/Bis-GMA induced apoptosis and disturbed the cell cycle, increasing the fraction of the cells in the G2/M checkpoint. The results obtained indicate that the HEMA/Bis-GMA model methacrylate composite, which can be considered as a representative for methacrylate-based dental restorations, displays a broad spectrum of genotoxicity and so – a high mutagenic potential. Therefore, new formulations of materials should be created with the emphasis on the decrease of their mutagenic potential. Chapter III - Frontotemporal lobar degeneration (FTLD) with transactive response (TAR) DNA-binding protein of 43kDa (TDP-43) proteinopathy (FTLDTDP) is a neurodegenerative disease characterized by variable neocortical and allocortical atrophy principally affecting the frontal and temporal lobes. Histologically, there is neuronal loss, microvacuolation in the superficial cortical laminae, and a reactive astrocytosis. A variety of TDP-43 immunoreactive changes are present in FTLD-TDP including neuronal cytoplasmic inclusions (NCI), neuronal intranuclear inclusions (NII), dystrophic neurites (DN) and, oligodendroglial inclusions (GI). Many cases of familial FTLD-TDP are caused by DNA mutations of the progranulin (GRN) gene. Hence, the density, spatial

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patterns, and laminar distribution of the pathological changes were studied in nine cases of FLTD-TDP with GRN mutation. The densities of NCI and DN were greater in cases caused by GRN mutation compared with sporadic cases. In cortical regions, the commonest spatial pattern exhibited by the TDP-43 immunoreactive lesions was the presence of clusters of inclusions regularly distributed parallel to the pia mater. In approximately 50% of cortical gyri, the NCI exhibited a peak of density in the upper cortical laminae while the GI were commonly distributed across all laminae. The distribution of the NII and DN was variable, the most common pattern being a peak of NII density in the lower cortical laminae and DN in the upper cortical laminae. These results suggest in FTLD-TDP caused by GRN mutation: 1) there are greater densities of NCI and DN than in sporadic cases of the disease, 2) there is degeneration of the corticocortical and cortico-hippocampal pathways, and 3) cortical degeneration occurs across the cortical laminae, the various TDP-43 immunoreactive inclusions often being distributed in different cortical laminae. Chapter IV - In a screening for selective inhibitors of eukaryotic DNA polymerase (pol) species, dehydroaltenusin extracted from a fungus (Alternaria tennuis) was found to be an inhibitor of DNA replicative pol α. This compound inhibited only mammalian pol α, and did not influence the activity of other DNA replicative pols such as pols δ and ε, but also showed no effect even on pol α activity from other vertebrate, fish, or plant species. Dehydroaltenusin also had no effect on other DNA metabolic enzymes tested. The inhibitory effect of dehydroaltenusin on mammalian pol α was dose-dependent with an IC50 value of 0.5 μM. This effect was 10-fold stronger than that of aphidicolin, a well-known potent inhibitor of eukaryotic DNA replicative pols α, δ and ε. The inhibitory mode of dehydroaltenusin for mammalian pol α activity was competitive with the DNA template-primer and non-competitive with the nucleotide substrate. This compound inhibited the proliferation of a human cervix carcinoma cell line, HeLa, with an LD50 value of 38.0 μM, by arresting cells at S-phase, and preventing the incorporation of thymidine into the cells. Dehydroaltenusin increased cyclin E and cyclin A levels, and induced cell apoptosis. Selective inhibitors of DNA replicative pol α, such as dehydroaltenusin, might provide novel markers for the development of anti-proliferative drugs. NIH3T3 cells that took up dehydroaltenusin by hypotonic shift, that is, the transient exposure of cultured cells in hypotonic buffer to small molecules that cannot penetrate cells, also showed inhibition of cell growth. At a low concentration (10 μM) of dehydroaltenusin, DNA replication was inhibited and several large foci of replication protein A (RPA) were found. Furthermore, when dehydroaltenusin

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was incubated with aphidicolin, RPA foci were not observed in cells. In summary, these findings suggest that dehydroaltenusin blocks DNA replication through pol α inhibition, and generates single-stranded DNA, resulted in uncoupling of leading strand and lagging strand synthesis. Dehydroaltenusin could be useful as a “molecule probe” for pol α research in DNA replication. The role of pol α function in DNA replication is discussed. Chapter V - Detection of single-nucleotide polymorphisms (SNPs) or mutations in the human genome can contribute to the establishment of genetic linkages, the diagnosis of inherited diseases, and the selection of rational remedies. Several procedures have been developed to facilitate high-throughput detection of SNPs and mutations, but these usually require expensive apparatus and the services of skillful analysts, making it difficult for most clinical researchers and physicians to obtain useful data on SNPs and mutations. The establishment of a reliable and cost-effective detection method that uses conventional laboratory equipment is therefore desirable, particularly in medical applications. Here, the authors introduce two methods based on polyacrylamide gel electrophoresis (PAGE) of DNA-binding zinc(II) complexes that are suitable for detecting mutations. The first method, known as Zn2+–cyclen/PAGE, is based on a difference in the mobility in PAGE of mutant DNA compared with that of nonmutated DNA of the same chain length. This difference in mobility results from binding of the Zn2+–cyclen complex (cyclen = 1,4,7,10tetraazacyclododecane) to thymine bases, which results in a decrease in the total charge and a local conformational change in the target DNA. The Zn2+– cyclen/PAGE method, when combined with polymerase chain reaction (PCR)based heteroduplexing, permits visualization of heteroduplex bands on PAGE gels and allows screening for SNPs and mutations. The second method is known as Zn2+–Phos-tag/PAGE. This is based on a difference in the mobility of a phosphorylated DNA fragment compared with that of its nonphosphorylated analogue containing an identical numbers of base pairs when they are subjected to PAGE on a phosphate-affinity gel containing an immobilized polyacrylamidebound dizinc(II) complex phosphate-binding tag molecule, Zn2+–Phos-tag {Phostag = 1,3-bis[bis(pyridin-2-ylmethyl)amino]propan-2-olate}. The Zn2+–Phostag/PAGE method, when combined with an allele-specific PCR method using a 1:1 mixture of 5'-phosphate-labeled and nonlabeled allele-specific primers, permits the separation of the less mobile 5'-phosphate-labeled PCR product from its more-mobile nonlabeled counterpart, and permits the determination of a genotype as heterozygotic or homozygotic. Here, the authors review some applications of gel-based methods based on zinc(II) complexes, and the authors

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compare their resolving power in separating mutant DNA with that of conventional gel-based electrophoresis techniques. Chapter VI - There is a need to understand how biological organization at one level translates into biological organization at the next level. Biology encompasses numerous levels from nanometers to meters, from atoms and macromolecules to organisms and populations of individuals. Here the authors shall consider the DNA mutations at the level of single nucleotides and their impact at the higher level of codons, sets of synonymous codons and their symmetries linked to tRNA-aminoacylation and degeneracy in the genetic code. The authors review will be restricted to DNA mutations, which are either singlebase deletions known to be highly deleterious within open reading frames or base substitutions, which are transitions (substitutions of a pyrimidine by another pyrimidine or substitutions of a purine by another purine) or transversions. Concepts such as coevolution, minimization of the effects of errors or metrics which found so far little use in the natural sciences, can be successfully applied to understand the connections between biological properties at these molecular and macromolecular levels. Chapter VII - The parasites belonging to the family Trypanosomatidae (order Kinetoplastida) are among the most primitive eukaryotes. Some trypanosomatids are the etiologic agents of neglected human pathologies such as South American and African trypanosomiasis and leishmaniasis. As a consequence of their ancient phylogenetic position, nuclear DNA replication in trypanosomatid protozoa shows conserved and non-conserved features. DNA replication in trypanosomatids initiates nearly simultaneously in the nucleus and in the genetic material of the single mitochondrion (or kinetoplast), suggesting that DNA synthesis is coordinately regulated in both organelles. In eukaryotes, nuclear DNA replication is preceded by assembly of the pre-replication complex, which is coordinated by the Origen Recognition Complex (ORC). However, in trypanosomatids, the prereplication complex differs from other eukaryotes and is similar to Archaea. All of these parasites contain only one protein that recognizes the replication origins and is found in the nucleus throughout the cell cycle, which suggests that it is not involved in the control of replication initiation. In the S phase, DNA replication starts at these origins and, in trypanosomes, occurs mainly at the nuclear periphery. In Leishmania spp., from the beginning up to mid S phase, replication sites are spread throughout the nuclear space to form subnuclear foci of active DNA replication. From mid-to-late S phase, replication is restricted to sites at the nuclear periphery. Few nuclear DNA polymerases have been described in trypanosomatid protozoa, although putative members of all polymerase families are found in their genomes. Structural and functional analyses indicate that most

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of these polymerases are highly conserved, with some of them being involved in polymerization and the repair of DNA damage. Although there are no descriptions of DNA polymerase δ in these protozoa, one of this protein´s partners, proliferating cell nuclear antigen (PCNA), is found in the nucleus throughout the cell cycle. Trypanosomatid PCNA forms distinct subnuclear foci in the S phase, whereas its distribution is more diffuse in the G2/M phase and in post-mitotic phase cells. This finding suggests that there may be phase-specific regulation of PCNA in the cell cycle. DNA replication in trypanosomatid telomeres is terminated by the action of telomerase. The biochemical properties of the trypanosomatid enzyme are conserved and resemble those described in other eukaryotes. Leishmania telomeres replicate late in S phase and at the beginning of G2 phase the chromosomes cluster at the nuclear periphery. Telomerase colocalizes with telomeres from the late S to G2 phases. These observations point to the existence of replication factories in trypanosomatids, the importance of which will be reviewed and discussed in this chapter.

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Chapter I

Connections between BLM and the Fanconi Anemia Pathway in the Repair of Replication Fork Damage

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Meghan Larin and John Peter McPherson∗ Department of Pharmacology and Toxicology, University of Toronto, Toronto, Ontario, Canada

Abstract Our DNA is continuously subjected to damage-causing endogenous and exogenous factors, resulting in lesions ranging from interstrand crosslinks to double strand breaks. These and other types of DNA damage encumber the S-phase of the cell cycle, as damaged template DNA can pose a problem for replication fork progression. Stalled replication forks can lead to fork collapse, causing additional DNA damage in the form of double strand breaks. Mammalian cells have developed checkpoint mechanisms that sense and repair DNA damage while maintaining replication fork stability. The process of replication fork restart is complex and involves multiple pathways depending on the lesion type. Initiation of these checkpoints results in cell ∗

Corresponding Author: [email protected].

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cycle arrest and consequent stabilization of the replication fork in order to prevent collapse. Repair of damage is believed to occur via translesion synthesis (TLS) or homologous recombination, however the exact mechanism through which these pathways act to maintain genomic stability and accomplish repair of DNA lesions remains unknown. This chapter will highlight pathways required for attempt to describe and evaluate current research into the area of resumption of replication fork stalls in mammalian cells. In this review we will focus on the role of helicases and endonucleases in the restart of damaged replication forks, with a particular emphasis on interplay between BLM helicase and components of the Fanconi anemia (FA) pathway of DNA repair.

Replication fork integrity must be continually maintained in order to ensure fidelity of DNA replication during cell division. This ideal objective is often jeopardized by the presence of lesions or structural abnormalities in DNA caused by various endogenous and exogenous sources. The nature of the lesion in question and its location on either the leading-strand or lagging-strand template generally dictates the severity of the impact on replication fork progression and the mode of repair employed. For example, cells are continually detecting and replacing damaged nucleotides or bases that can be converted into mutations or gaps in DNA sequence following replication. Such lesions on the leading-strand template might block the polymerase on the leading-strand but not on the laggingstrand. Other lesions such as inter-strand cross-links or pre-existing template gaps can block progression of replication forks. When replication-coupled repair is compromised, DNA becomes more susceptible to breakage, inappropriate recombination and genomic instability. Failure to properly restart the replication process can gravely impact cell survival and genomic stability, accordingly it is crucial that the various components of the cellular response to DNA damage that detect, signal and restore operational replication forks work properly at all times. Defects in this cellular response to replication fork damage have been linked to numerous human syndromes characterized by increased cancer incidence, developmental abnormalities and premature aging (Burtner and Kennedy, 2010; Lavin, 2008; Spry et al., 2007). Cells trigger repair of replication forks through signal-transduction cascades, collectively known as checkpoints, that sense the presence of replication stress or DNA damage and induce cell-cycle arrest and DNA repair (Falck, 2001, 2002; Heffernan, 2002; Schechter, 2004; Weiss, 2002). The phosphoinositide 3-kinaserelated kinases ataxia telangiectasia mutated (ATM) and ATM- and Rad3-related (ATR) initiate S-phase checkpoint responses in mammals. ATM is activated

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primarily by the presence of double-strand breaks following the recruitment of this kinase to the MRN complex, comprised of MRE11, RAD50 and NBS1 bound to the broken ends of DNA. ATR is activated by the presence of exposed single – stranded DNA coated with replication protein A at sites of stalled replication forks. This coated single-stranded DNA serves to recruit ATR-ATR-interacting protein (ATRIP) and RAD17, which then facilitates the establishment of the checkpoint clamp RAD9-RAD1-HUS1 which is subsequently phosphorylated by ATR and completes checkpoint activation. Downstream kinases CHK1 and CHK2 serve to broadcast the checkpoint signal throughout the cell, following activation by ATR and ATM respectively. Both checkpoint kinases ATM and ATR trigger an intra-S checkpoint that prevents the initiation of new replication forks. In addition, ATR and Chk1 kinase direct processes to stabilize stalled replication fork structures and ensure their repair by homologous recombinationdriven mechanisms. Replicative blocks need to be removed or bypassed in order for replication to resume. The processes required for repair are believed to utilize translesion polymerases and/or homologous recombination (Baynton, 2000; Goodman, 2000; Segurado, 2002) (Rothkamm, 2003). Translesion polymerases are specialized DNA polymerases recruited to stalled replication forks in order to continue DNA synthesis past lesions that normally would derail DNA synthesis by normal replicative polymerases. It is unclear how these translesion polymerases facilitate error-free repair, given that these enzymes generate misincorporation rates several-fold higher than that of normal polymerases. The repair of damaged replication forks by homologous recombination mechanisms bypasses the potential for mutagenesis inherent in translesion polymerases. It is believed that homologous recombination rectifies stalled replication forks through the generation of a four-stranded DNA intermediate known as a Holliday junction. Cleavage of this intermediate at the branch point of the junction appears to be necessary to facilitate restart of replication, especially when lesions impact both strands of the DNA helix (Cromei, 2000).

RecQ DNA Helicases The unwinding of DNA is required in several discrete processing steps that culminate in replication fork repair. An important family of helicases that function by unwinding complementary strands of DNA is known as the RecQ helicases, first identified in E. coli as RecQ (Umezu and Nakayama, 1993), and later in S.

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cerevisiae (SGS1; (Gangloff et al., 1994)) and S. pombe (RQH1; (Stewart et al., 1997)). Interestingly, subsequent studies in mammalian systems identified five RecQ helicases; Bloom’s (BLM), Werner’s (WRN), RecQL1, RecQL4, and RecQL5 (van Brabant et al., 2000). Mutations in the genes coding for BLM and WRN lead to Bloom syndrome and Werner syndrome respectively, whereas mutations in RecQL4 can result in either Rothmund-Thomson syndrome, RAPADILINO syndrome, or Baller–Gerold syndrome, depending on the nature of the mutation (Chu and Hickson, 2009). Although each of these helicases carries out a similar function, each member of this family is likely to possess a distinct function as the syndromes listed above are each considered distinct, linked only by the prevalence of genomic instability and increased cancer risk in patients. On a molecular level, the RecQ helicases are utilized in DNA repair and maintenance of genomic stability for both their pro- and anti-recombination activities, which is based on the premise that these proteins have an affinity for various HR intermediates and replication fork structures (Ouyang et al., 2008). Interactions have been identified between the different RecQ helicases as well as with other proteins involved in replication fork restart, indicating that the specific activities of these enzymes do not operate independently, but are choreographed to achieve a common goal.

BLM and Regulation of Homologous Replication Mutations in the gene BLM cause Bloom syndrome, an autosomal recessive disorder characterized by deficits in growth, immune system cellularity, fertility and tumour suppression (Bloom, 1954). On a molecular level, Bloom syndrome is characterised by a substantially increased number of sister chromatid exchanges and formation of unusual chromosomal structures, such as quadriradial structures formed through crossover events (German, 1965; van Brabant et al., 2000). The gene encoding BLM was located by searching candidate genes for the presence of a DExH box motif, a domain conserved amongst RecQ helicases in both bacteria and yeast (Ellis et al., 1995). Subsequent studies revealed that BLM possesses DNA-dependent ATPase activity and acts to unwind DNA in the 3’-5’ direction (Karow et al., 1997). Consistent with other RecQ helicases, BLM exhibits affinity for specific DNA substrates, including branched structures mimicking HJ, 3’tailed and forked duplex structures, bubble substrates, and G-quadruplexes (Mohaghegh et al., 2001). Perhaps one of the more interested characteristics of

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BLM is its ability to act as both a pro- and anti-recombination protein in the repair of stalled/ collapsed replication forks and in the maintenance of genome stability. Patients suffering from Bloom syndrome demonstrate increased levels of SCE, indicating that BLM must somehow act to protect the cell against inappropriate HR events. It has been shown that BLM levels peak during S/G2 phase, indicating that BLM is necessary during replication likely not only to promote strand separation, but also to prevent SCE (Wu, 2007). The anti-recombination and anticrossover roles of BLM are complex and still not well understood, however progress has been made over the past decade, and we are closer to understanding the role of BLM in the maintenance of genomic stability and DNA repair. Before considering the role of BLM in suppression of inappropriate homologous recombination, it is necessary to understand the stages of this repair process. In the case of a double-strand break, endonucleases remove nucleotides on the 3’-strand, forming ssDNA. The ssDNA strand is then coated by RPA, which prevents the strand from forming secondary structures. Next, RAD51 is recruited to the strand, displacing RPA and forming a RAD51 filament. The ssDNA filament then invades the sister chromatid with the purpose of seeking out a homologous sequence, where a D-loop structure can then be formed. From this step, referred to as the ‘synapsis’ step, the cell has three options for repair: classical DSB repair (DSBR), Synthesis-Dependent Strand Annealing or break induced repair (BIR) (Li and Heyer, 2008). To date, BLM has only been implicated in DSBR and SDSA, so only these outcomes will be discussed. For DSBR, the invading strand of the D-loop is extended using the sister chromatid. The other end of the DSB joins with the newly synthesised strand, forming a double Holliday junction structure. This intermediate can have multiple fates. One such outcome is branch migration, followed by dissolution of the Holliday junction. This process is directly mediated by BLM. Studies have demonstrated that BLM forms a complex in vivo with TOPOIII (Johnson et al., 2000; Wu et al., 2000a), and a similar interaction has also been observed in S. cerevisiae between Sgs1 and Top3 (Gangloff et al., 1994). Using a synthetic DHJ structure, Wu and colleagues demonstrated that the interaction between BLM and TOPOIIIa is required for the unwinding and dissolution of this structure (Wu and Hickson, 2003). Using an elegant model involving a radioactive labelled DHJ substrate, they demonstrated that BLM and TOPOIIIa mediate a non-crossover event following dissolution (Wu and Hickson, 2003). More recently, it has been observed that this complex also includes RecQ-mediated genomic instability 1 (RMI) which aids in the dissolution of double Holliday junctions (Raynard et al., 2006; Yin et al., 2005) and RMI2, which helps to stabilize and suppress sister chromatid exchange at this complex (Singh et al., 2008; Xu et al., 2008).

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A second fate for DHJ is resolution through the use of various endonucleases involved in DNA repair, resulting in either crossover or non-crossover products. While BLM is not directly involved in this process, as its main function is to prevent crossover events from occurring, interactions between BLM and endonucleases involved in resolution of HJ may implicate it in this process. An interaction between MUS81 and BLM has been identified (Shimura et al., 2008; Zhang et al., 2005). MUS81 acts together with EME1 as a structure-specific endonuclease that is believed to cleave structural DNA intermediates that arise during processing of replication fork damage (Osman, 2007). A direct interaction of MUS81 at the C-terminus of BLM has been observed - the same region at which RAD51 and TOPOIIIa are known to interact with BLM (Zhang et al., 2005). It was also observed that BLM enhanced the accumulation of MUS81 at stalled replication forks, and that addition of BLM enhanced the action of MUS81 on the resolution of nicked HJs and 3’-flap structures (Zhang et al., 2005). Zhang and associates postulated that BLM may function as a ‘platform’ for various proteins necessary for DNA repair. This is a possibility - however, considering BLM’s role in the prevention of crossover events and the role of Mus81 in resolving HJ through crossover events, it will be interesting to further explore the ramifications of this interaction. Following the synapsis step, another potential option for replication fork restart using HR is SDSA. In this mechanism, the invading strand of the D-loop elongates using the sister chromatid as template, however the loop is quickly dissolved, leading to a non-crossover event. This process of D-loop dissolution is mediated through another interaction between BLM and RAD51 (Wu et al., 2001). BLM was found to disrupt the RAD51 filament of the 3’-invading strand of the D-loop, dissolving this structure and facilitating SDSA (Bachrati et al., 2006). Several other lines of evidence implicate BLM in processes that govern replication fork restart. BLM has been shown to interact with p53, RAD51, PCNA and the BRCA1-associated genome surveillance complex at stalled replication forks (Sengupta, 2003; Wang, 2000). It is likely that BLM and other RecQ helicases work together with associated topoisomerases to relieve topological barriers that accumulate at DNA replication forks. Loss of this activity would be expected to result in the accumulation of abnormal replication intermediates that could jeopardize sister-chromatid disjunction. Indeed, recent studies in BLMdeficient cells have demonstrated the presence of ultrafine bridges of chromatin during anaphase. The appearance of these chromatin bridges has been attributed to impaired chromosome segregation events. Interestingly, BLM, TOP3a and FANCD2/I proteins (components of the FA pathway of DNA repair) have been

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shown to localize to these ultrafine bridges, suggestive of a functional connection between BLM and the FA pathways of repair (Chan, 2007; Chan, 2009). Recent research has identified several other examples of interaction between BLM and the Fanconi Anemia (FA) pathway, another DNA repair pathway directly linked to homologous repair of replication fork damage.

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Connections between FANCM of the Fanconi Anemia Pathway and BLM The Fanconi Anemia (FA) pathway is an area of much focus in the DNA damage response field. Defects in one of at least fifteen genes can result in this hereditary syndrome that is characterized by developmental defects, bone marrow failure, and genomic instability resulting in increased cancer risk (Moldovan, 2009; Wang, 2007). On a molecular level, cells from patients suffering from FA display sensitivity to cross-linking agents, indicating a defect in DNA repair pathways that respond to cross-links and other form of DNA damage that impede replication. The FA pathway currently consists of at least fifteen proteins, eight of which (FANCA, B, C, E, F, G, L, M) constitute the FA core complex. This core complex possesses E3 ubiquitin ligase activity and is responsible for ubiquitinating downstream FA proteins FANCD2 and FANCI in response to ICL damage (Moldovan, 2009). This ubiquitination event leads to recruitment of other proteins thought to be involved in homologous recombination-mediated repair. While each of these proteins is an integral part of the pathway, some have additional responsibilities and connections with other proteins or downstream processes. For example, both FANCM and FANCJ possess the ability to bind and restructure DNA and both proteins have been shown to interact directly with BLM (Deans and West, 2009; Suhasini et al., 2011). The FANCM protein has been the focus of many studies in recent years due to its unique characteristics when compared to other proteins in the FA complementation group, as well as its interacting partners. FANCM was first identified in S. cerevisiae as MPH1 (Scheller et al., 2000), followed by archaea (Komori et al., 2002) and later in mammals (Meetei et al., 2005). Mammalian FANCM contains domains conserved from its yeast and archaeal counterparts including Walker and DEAH motifs which place FANCM in the superfamily 2 (SF2) of helicases (Meetei et al., 2005; Whitby). Also, as is observed in both yeast and archaea, FANCM possesses an ERCC4-nuclease domain characteristic of the XPF/Mus81 family of structure-specific endonucleases (Meetei et al., 2005).

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Interestingly, while both the helicase and nuclease domains of FANCM orthologues in yeast and archaea are functional, the domains in mammalian FANCM are degenerate and therefore do not demonstrate the activity typically attributed to these domains (Meetei et al., 2005). Despite these observations, mammalian FANCM does act to remodel DNA as it was shown to demonstrate DNA-dependent ATPase activity (Meetei et al., 2005). Biochemical analysis of mammalian FANCM has yielded interesting results regarding its interacting protein partners. Much of this data has helped to shed light on the unique function of FANCM as an important component in the repair of replication forks, the FA repair pathway, and its potential role as a bridging protein between the FA core complex and BLM. Co- immunoprecipitation experiments revealed that FANCM associates with the FA core complex and the BLM pathway (Deans and West, 2009; Meetei et al., 2005). Further investigations revealed the identity of another FANCM binding partner; FAAP24, which forms a heterodimer with FANCM (Ciccia et al., 2007). The FAAP24 protein contains an inactive ERRC4 domain, similar to that of FANCM, and interacts directly with the FA core complex (Ciccia et al., 2007). The FAAP24 protein is critical to the function of FANCM regarding its role in the FA pathway, which is discussed in greater detail below. The FANCM/FAAP24 complex has more recently been found to interact with two additional proteins; MHF1-MHF2, which also contribute to the function of the FA core complex (Singh et al., 2010). The role of FANCM in the FA and DNA repair pathways is somewhat complicated and not well understood, however great strides in the understanding of the function of this protein have been made in the past few years. As previously mentioned, FANCM was found to directly associate with members of the FA core complex through immunoprecipitation assays (Meetei et al., 2005). Additional studies showed that FANCM is required for ubiquitination of FANCD2 in response to ICL damage (Xue et al., 2008), and must be present in a complex with FAAP24 for this to take place (Ciccia et al., 2007). Based on the significant role FANCM/FAAP24 plays in the function of the FA pathway, and the structural aspects of this protein, it was hypothesised that FANCM/FAAP24 is involved in DNA repair and replication restart through HR. Direct and stable interaction between FANCM/FAAP24 and chromatin indicate this protein complex is necessary for loading of the FA core complex onto DNA in response to cross-link damage, or other situations where this pathway is required (Kim et al., 2008). In addition, FANCM/FAAP24 interacts directly with ATR/CHK1 signalling components to stabilize CHK1 which serves to maintain a DNA damage response at stalled or collapsed replication forks (Collis et al., 2008; Luke-Glaser et al., 2010; Sobeck et al., 2009).

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Although FANCM/FAAP24 possesses domains for both helicase and endonuclease functions, this complex cannot perform either function (Meetei et al., 2005). As this complex has been implicated in the FA pathway and the DNA damage response, how exactly does it interact in this pathway? As previously mentioned, structural analysis of FANCM revealed that this protein does have DNA-dependent ATPase activity (Meetei et al., 2005), indicating it is likely able to metabolize DNA structures in some way. The chromatin-binding properties of FANCM prompted investigation of its potential to resolve various replication fork intermediate structures. FANCM/FAAP24 showed affinity for Holliday junction structures but was only able to resolve forms with homologous branches which allow the structure to be resolved without helicase (Xue et al., 2008). Such structures could be resolved by FANCM/FAAP24 in an ATP-dependent process, indicating that FANCM is indeed able to promote HJ branch migration (Gari et al., 2008b; Xue et al., 2008). The FANCM/FAAP24 complex has also been shown to dismantle D-loops and mediate fork regression in response to damage (Gari et al., 2008a) and has demonstrated affinity for ssDNA, splayed arm DNA, and 3’flaps (Whitby, 2010). FANCM has been proposed to be a molecular interface between processes mediated by the FA pathway and BLM. Although this interaction was identified during the first study into the function of mammalian FANCM (Meetei et al., 2005), research into the significance of the connection between these two proteins was only recently described (Deans and West, 2009). Initially, members of the FA core complex and BLM were first found to co-immunoprecipitate. BLM and TOPOIIIa, as well as RPA70, were found to co-immunoprecipitate with FANCA, G, C, E, and F, establishing a connection between these two pathways, termed the BRAFT (BLM, RPA, FANC, TOPOIIIa) complex (Meetei et al., 2003). More recent studies have however identified two domains in FANCM, known as MM1 and MM2. The MM1 motif interacts directly through the FA core complex through association with FANCF and also binds TOPOIIIa, whereas MM2 was found to interact with the BLM complex protein RMI1, possibly through association with its OB-fold domain (Deans and West, 2009). Perhaps one of the most interesting findings of this study is with respect to the co-operation between the FA pathway and BLM. First, the authors postulated that FANCM may potentially act as a bridge between the FA core complex and BLM. FANCF was immunoprecipitated from extracts of control cells or cells depleted of FANCM. Under these conditions, FANCF was able to pull down other members of the FA core complex which associated with FANCM, however no components of the BLM complex were found (Deans and West, 2009). The second interesting finding is that both FANCM and BLM are required for DNA cross-link repair,

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which has historically been linked mainly with the FA pathway. FANCM/FAAP24 has previously been shown to participate in DNA cross-link repair, with a particular requirement for this complex in RPA foci formation and phosphorylation following generation of cross-links. Furthermore, FAAP24 has been shown to bind to cross-linked DNA structures in vitro (Huang et al., 2010). In cells deficient for FANCM, the authors found it was not possible to rescue mitomycin-C sensitivity with FANCM MM1 or MM2 mutant constructs, suggesting that resistance to cross-link damage requires both FANCM and BLM (Deans and West, 2009). Similar to Bloom syndrome cells, FANCM deficient cells exhibit an increased propensity for sister chromatid exchange, suggestive of an analogous function for FANCM in suppressing recombination during homologous recombination-mediated repair of replication-associated damage (Deans and West, 2009). Taken together, these findings suggest that FANCM/FAAP24 plays a pivotal role in coordinating actions of the FA core complex with the helicase activity of BLM.

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Connections between FANCJ of the FA Pathway and BLM Another member of the FA pathway has recently been identified as a protein with the ability to interact with BLM. The FANCJ protein, (also referred to as BACH1 or BRIP1), is similar to FANCM in that it possesses the ability to interact with and mediate the resolution of various DNA structural intermediates. It is unique, however, in that FANCJ functions downstream of the FA core complex, associating instead with proteins involved in DSB repair such as BRCA1. The FANCJ protein is also hypothesised to participate in replication restart. This and other characteristics of this unique FA protein are discussed below. In humans, BACH1 was first identified through a GST pull-down assay using the BRCA1 C-Terminal (BRCT) tandem motifs, which are phosphopeptide binding domains necessary for the tumour suppressor of BRCA1 (Moynahan et al., 1999; Scully et al., 1999; Wu et al., 2000b; Zhong et al., 1999) (Cantor et al., 2001). Structural and biochemical analysis of BACH1 revealed the presence of an N-terminal DEAH domain, as well as DNA-dependent ATPase activity, indicative of SF2 helicases (Cantor et al., 2001). Interestingly, mutations in BACH1 were found in cases of early onset breast cancer where individuals demonstrated wild type BRCA1 and BRCA2 (Cantor et al., 2001; De Nicolo et al., 2008; Seal et al., 2006). Mapping of BACH1 in a human FA case proved that BACH1/BRIP1 was

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actually FANCJ (Levitus et al., 2005), establishing a role for this protein not only as a tumour suppressor gene based on its interactions with BRCA1, but also as a component of the FA pathway. Cells deficient in FANCJ and BRCA1 are sensitive to both cross-linking damage (Litman et al., 2005) and γ-irradiation (Kumaraswamy and Shiekhattar, 2007), pointing to a common deficiency in double-strand break repair. Mammalian cells deficient in either FANCJ or BRCA1 are still able to ubiquitinate FANCD2, indicating both FANCJ and BRCA1 lie downstream of the FA pathway (Litman et al., 2005). Given that FANCJ possesses a helicase domain and DNA-dependent ATPase activity, how does this protein function with respect to DNA damage? Kumaraswamy and colleagues determined through a series of experiments that the ATPase activity of FANCJ is controlled by phosphorylation and is most active during S-phase of the cell cycle, linking actvity of this enzyme to repair events required during DNA replication (Kumaraswamy and Shiekhattar, 2007). Interestingly, FANCJ is associated not only with BRCA1 and FA repair pathways, but also has recently been shown to physically interact with BLM and co-localize to subnuclear foci in the presence of replication fork damage (Suhasini et al., 2011). Interestingly, FANCJ is required for BLM protein stability, as BLM is degraded by a proteosome-mediated pathway in cells deficient in FANCJ. Both FANCJ and BLM helicase activities were found to cooperate in the unwinding of damaged DNA substrates, suggesting that both enzymes act together to facilitate repair of the same replication fork intermediates (Suhasini et al., 2011).

The Structure-Specific Endonuclease SLX4-SLX1 and Its Relationship to BLM and the FA Pathway Structures-specific endonucleases are believed to play a critical role in DNA repair through the selective cleavage of DNA intermediates that arise during replication fork processing, such as Holliday junctions. Whereas some members of this functional class (such as Mus81-Eme1) have been extensively researched, recently another structure-specific endonuclease known as SLX4 has come to light that appears not only to cleave DNA intermediates but also coordinate the activity of other endonucleases. Slx4 was first identified along with several other proteins encoded by SLX (Synthetic Lethal of unknown function) genes, that when mutated cause lethality to yeast cells deficient in Sgs1, the functional equivalent to BLM. Interestingly, Slx3 and Slx2 identified in the same study were

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subsequently identified as Mus81-Eme1. Slx4 and its catalytic partner Slx1 were found to act as a heterodimeric endonuclease that was initially shown to resolve 5’ flap structures. The human orthologue of yeast Slx4 (SLX4) was identified in 2009 (Fekairi et al., 2009; Munoz et al., 2009; Svendsen et al., 2009). Extensive studies outlined in these three papers identified several novel interactions and properties of human SLX4, some of which mirrored those previously deciphered in yeast, including its ability to cleave 5’ and 3’ flap structures and resolve Holliday junction substrates (Fekairi et al., 2009; Munoz et al., 2009; Svendsen et al., 2009). Interestingly, it was found that SLX1-SLX4 can act to resolve static HJs, a unique characteristic of this enzyme that sets it apart from Mus81-Eme1 (Svendsen et al., 2009). Although the results of these assays to determine the function of SLX1-SLX4 on various DNA structures are compelling, we cannot presently conclude these results mirror processes that occur in cells, given that the DNA substrates used are synthetic and based on a theoretical model of homologous recombination. Very recently, SLX4 research has jumped from in vitro model systems to in vivo disease relevance through new studies that place it in the FA pathway (Crossan et al., 2011; Kim et al., 2011; Stoepker et al., 2011). SLX4 knockout mice have been recently described that demonstrate several phenotypic anomalies including growth defects, cellular senescence, sensitivity to cross-linking agents, increased chromosomal instability, and developmental defects such as microphthalmia (Crossan et al., 2011). Significantly, the anomalies observed phenocopy those seen in Fanconi Anemia (FA). Remarkably, mutations in SLX4 (renamed FANCP) have recently been linked to patients afflicted with this disease (Kim et al., 2011; Stoepker et al., 2011), indicating that SLX4 participates in the FA pathway (Crossan et al., 2011; Stoepker et al., 2011) (Kim et al., 2011). Despite these exciting findings, the direct relationship between SLX4-SLX1 and other members of the FA pathway remains to be determined. Another interesting feature of SLX4-SLX1 is its ability to act as a bridge between itself and other structure-specific endonucleases (Fekairi et al., 2009; Munoz et al., 2009; Svendsen et al., 2009). For example, in addition to SLX1, SLX4 associates directly with Mus81-Eme1 and XPF-ERCC1, associations that appear to reflect cooperativity between these proteins, as the presence of SLX4SLX1 increases Mus81-Eme1 and XPF-ERCC1 activities. Svendson and colleagues proposed that SLX4-SLX1 acts to nick static Holliday junction substrates in order for MUS81-EME1 to cleave and resolve these structures. Taken together, this data implies the potential role of SLX4 as a protein scaffold, which may aid the spatial and temporal coordination of several steps in replication

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fork repair that involve cleavage of DNA intermediates by different endonucleases (Munoz et al., 2009). It is likely that SLX4-SLX1 not only physically interacts but also functionally interacts with other components of replication fork repair. One such example is the potential for functional redundancy between SLX4 and Bloom’s helicase (BLM). Either Slx4-Slx1 or Sgs-1-Top3 have to be present to ensure viability of yeast cells, indicative of some level of functional redundancy between these two complexes with regards to DNA repair mechanisms (Mullen et al., 2001)(Gangloff et al., 1994). Fricke and Brill proposed that the functional redundancy was linked to the ability of both complexes to act on converging replication forks, a situation that would cause replication fork stalling (Fricke, 2003). In general, they proposed that Sgs1-Top3 resolve stalled forks through decatenation, and thereby prevent crossover events. On the other hand, the endonuclease activity of Slx1-Slx4 can cleave at the 5’ side of the two converging replication forks, and lead to single-strand annealing and a crossover event. The absence of both proteins would, according to the results of the lethal screen, leave the cell without any means of repairing these converging forks, leading to cell death (Fricke, 2003).

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Conclusion Remarkable progress has recently been made not only in the identification of new helicase and endonuclease activities that participate in the repair of damaged replication forks, but also in understanding interactions between these enzymes that imply the coordinated activity of these proteins in the DNA damage response. Further studies will undoubtedly shed light on how the activity of each of these components is temporally and spatially regulated to facilitate an optimal repair event in the presence of replication-fork associated damage.

Acknowledgments Research conducted in the McPherson laboratory is supported by The Cancer Research Society Inc. and an Early Researcher Award from the Province of Ontario Ministry of Research and Innovation. Meghan Larin is supported by a Doctoral Award from the Canadian Institutes of Health Research/Fanconi Canada.

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References Bachrati, C.Z., Borts, R.H., and Hickson, I.D. (2006). Mobile D-loops are a preferred substrate for the Bloom's syndrome helicase. Nucleic Acids Res. 34, 2269-2279. Baynton, I., Fuchs, R.P. (2000). Lesions in DNA: hurdles for polymerases. Trends Biochem. Sci. 25, 74-79. Bloom, D. (1954). Congenital telangiectatic erythema resembling lupus erythematosus in dwarfs; probably a syndrome entity. AMA Am. J. Dis. Child 88, 754-758. Burtner, C.R., and Kennedy, B.K. (2010). Progeria syndromes and ageing: what is the connection? Nat. Rev. Mol. Cell Biol. 11, 567-578. Cantor, S.B., Bell, D.W., Ganesan, S., Kass, E.M., Drapkin, R., Grossman, S., Wahrer, D.C., Sgroi, D.C., Lane, W.S., Haber, D.A., et al. (2001). BACH1, a novel helicase-like protein, interacts directly with BRCA1 and contributes to its DNA repair function. Cell 105, 149-160. Chan, K.-L., North, P.S., Hickson, I.D. (2007). BLM is required for faithful chromosome segregation and its localization defines a class of ultrafine anaphase bridges. EMBO J. 26, 3397-3409. Chan, K.L., Palmai-Pallag, T., Ying, S., Hickson, I.D. (2009). Replication stress induces sister-chromatid bridging at fragile site loci in mitosis. Nature Cell Biol. 11, 753-760. Chu, W.K., and Hickson, I.D. (2009). RecQ helicases: multifunctional genome caretakers. Nat. Rev. Cancer 9, 644-654. Ciccia, A., Ling, C., Coulthard, R., Yan, Z., Xue, Y., Meetei, A.R., Laghmani el, H., Joenje, H., McDonald, N., de Winter, J.P., et al. (2007). Identification of FAAP24, a Fanconi anemia core complex protein that interacts with FANCM. Mol. Cell 25, 331-343. Collis, S.J., Ciccia, A., Deans, A.J., Horejsi, Z., Martin, J.S., Maslen, S.L., Skehel, J.M., Elledge, S.J., West, S.C., and Boulton, S.J. (2008). FANCM and FAAP24 function in ATR-mediated checkpoint signaling independently of the Fanconi anemia core complex. Mol. Cell 32, 313-324. Cromei, G.A., Leach, D.R. (2000). Control of crossing over. Molecular Cell 6, 815-826. Crossan, G.P., van der Weyden, L., Rosado, I.V., Langevin, F., Gaillard, P.H., McIntyre, R.E., Gallagher, F., Kettunen, M.I., Lewis, D.Y., Brindle, K., et al. (2011). Disruption of mouse Slx4, a regulator of structure-specific nucleases, phenocopies Fanconi anemia. Nat. Genet 43, 147-152.

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De Nicolo, A., Tancredi, M., Lombardi, G., Flemma, C.C., Barbuti, S., Di Cristofano, C., Sobhian, B., Bevilacqua, G., Drapkin, R., and Caligo, M.A. (2008). A novel breast cancer-associated BRIP1 (FANCJ/BACH1) germ-line mutation impairs protein stability and function. Clin. Cancer Res. 14, 46724680. Deans, A.J., and West, S.C. (2009). FANCM connects the genome instability disorders Bloom's Syndrome and Fanconi Anemia. Mol. Cell 36, 943-953. Ellis, N.A., Groden, J., Ye, T.Z., Straughen, J., Lennon, D.J., Ciocci, S., Proytcheva, M., and German, J. (1995). The Bloom's syndrome gene product is homologous to RecQ helicases. Cell 83, 655-666. Falck, J., Mailand, N., Syljuasen, R.G., Bartek, J., Lukas, J. (2001). The ATMchk2-cdc25A checkpoint pathway guards against radioresistant DNA synthesis. Nature 410, 842-847. Falck, J., Petrini, J.H.J., Williams, B.R., Lukas, J., and Bartek, J. (2002). The DNA damage-dependent intra-S phase checkpoint is regulated by parallel pathways. Nature Genet. 30, 290-294. Fekairi, S., Scaglione, S., Chahwan, C., Taylor, E.R., Tissier, A., Coulon, S., Dong, M.Q., Ruse, C., Yates, J.R., 3rd, Russell, P., et al. (2009). Human SLX4 is a Holliday junction resolvase subunit that binds multiple DNA repair/recombination endonucleases. Cell 138, 78-89. Fricke, W.M., Brill, S.J. (2003). Slx1-Slx4 is a second structure-specific endonuclease functionally redundant with Sgs1-Top3. Genes and Development 17, 1768-1778. Gangloff, S., McDonald, J.P., Bendixen, C., Arthur, L., and Rothstein, R. (1994). The yeast type I topoisomerase Top3 interacts with Sgs1, a DNA helicase homolog: a potential eukaryotic reverse gyrase. Mol. Cell Biol. 14, 83918398. Gari, K., Decaillet, C., Delannoy, M., Wu, L., and Constantinou, A. (2008a). Remodeling of DNA replication structures by the branch point translocase FANCM. Proc. Natl. Acad. Sci. U S A 105, 16107-16112. Gari, K., Decaillet, C., Stasiak, A.Z., Stasiak, A., and Constantinou, A. (2008b). The Fanconi anemia protein FANCM can promote branch migration of Holliday junctions and replication forks. Mol. Cell 29, 141-148. German, J., Arechibald, TR., Bloom, D. (1965). Chromosomal breakage in a rare and probably genetically determinded. Science 148, 506-506 Goodman, M.F., Tippin, B. (2000). The expanding polymerase universe. Nature Reviews Molecular and Cell Biology 1, 101-109. Heffernan, T.P., Simpson, D.A., Frank, A.R., Heinloth, A.N., Paules, R.S., Cordeiro-Stone, M., Kaufmann, W.K. (2002). An ATR- and Chk1-dependent

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S checkpoint inhbits replicon initiation following UVC-induced DNA damage. Mol. Cell Biol. 22, 8552-8561. Johnson, F.B., Lombard, D.B., Neff, N.F., Mastrangelo, M.A., Dewolf, W., Ellis, N.A., Marciniak, R.A., Yin, Y., Jaenisch, R., and Guarente, L. (2000). Association of the Bloom syndrome protein with topoisomerase IIIalpha in somatic and meiotic cells. Cancer Res. 60, 1162-1167. Karow, J.K., Chakraverty, R.K., and Hickson, I.D. (1997). The Bloom's syndrome gene product is a 3'-5' DNA helicase. J. Biol. Chem. 272, 30611-30614. Kim, J.M., Kee, Y., Gurtan, A., and D'Andrea, A.D. (2008). Cell cycle-dependent chromatin loading of the Fanconi anemia core complex by FANCM/FAAP24. Blood 111, 5215-5222. Kim, Y., Lach, F.P., Desetty, R., Hanenberg, H., Auerbach, A.D., and Smogorzewska, A. (2011). Mutations of the SLX4 gene in Fanconi anemia. Nat. Genet. 43, 142-146. Komori, K., Fujikane, R., Shinagawa, H., and Ishino, Y. (2002). Novel endonuclease in Archaea cleaving DNA with various branched structure. Genes Genet. Syst. 77, 227-241. Kumaraswamy, E., and Shiekhattar, R. (2007). Activation of BRCA1/BRCA2associated helicase BACH1 is required for timely progression through S phase. Mol. Cell Biol. 27, 6733-6741. Lavin, M.F. (2008). Ataxia-telangiectasia: from a rare disorder to a paradigm for cell signalling and cancer. Nat. Rev. Mol. Cell Biol. 9, 759-769. Levitus, M., Waisfisz, Q., Godthelp, B.C., de Vries, Y., Hussain, S., Wiegant, W.W., Elghalbzouri-Maghrani, E., Steltenpool, J., Rooimans, M.A., Pals, G., et al. (2005). The DNA helicase BRIP1 is defective in Fanconi anemia complementation group. J. Nat. Genet. 37, 934-935. Li, X., and Heyer, W.D. (2008). Homologous recombination in DNA repair and DNA damage tolerance. Cell Res. 18, 99-113. Litman, R., Peng, M., Jin, Z., Zhang, F., Zhang, J., Powell, S., Andreassen, P.R., and Cantor, S.B. (2005). BACH1 is critical for homologous recombination and appears to be the Fanconi anemia gene product FANCJ. Cancer Cell 8, 255-265. Luke-Glaser, S., Luke, B., Grossi, S., and Constantinou, A. (2010). FANCM regulates DNA chain elongation and is stabilized by S-phase checkpoint signalling. EMBO J. 29, 795-805. Meetei, A.R., Medhurst, A.L., Ling, C., Xue, Y., Singh, T.R., Bier, P., Steltenpool, J., Stone, S., Dokal, I., Mathew, C.G., et al. (2005). A human ortholog of archaeal DNA repair protein Hef is defective in Fanconi anemia complementation group M. Nat. Genet. 37, 958-963.

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Meetei, A.R., Sechi, S., Wallisch, M., Yang, D., Young, M.K., Joenje, H., Hoatlin, M.E., and Wang, W. (2003). A multiprotein nuclear complex connects Fanconi anemia and Bloom syndrome. Mol. Cell Biol. 23, 34173426. Mohaghegh, P., Karow, J.K., Brosh, R.M., Jr., Bohr, V.A., and Hickson, I.D. (2001). The Bloom's and Werner's syndrome proteins are DNA structurespecific helicases. Nucleic Acids Res. 29, 2843-2849. Moldovan, G.-L., D'Andrea, A.D. (2009). How the Fanconi anemia pathway guards the genome. Annu. Rev. Genet. 43, 223-249. Moynahan, M.E., Chiu, J.W., Koller, B.H., and Jasin, M. (1999). Brca1 controls homology-directed DNA repair. Mol. Cell 4, 511-518. Mullen, J.R., Kaliraman, V., Ibrahim, S.S., and Brill, S.J. (2001). Requirement for three novel protein complexes in the absence of the Sgs1 DNA helicase in Saccharomyces cerevisiae. Genetics 157, 103-118. Munoz, I.M., Hain, K., Declais, A.C., Gardiner, M., Toh, G.W., Sanchez-Pulido, L., Heuckmann, J.M., Toth, R., Macartney, T., Eppink, B., et al. (2009). Coordination of structure-specific nucleases by human SLX4/BTBD12 is required for DNA repair. Mol. Cell 35, 116-127. Osman, F., Whitby, M.C. (2007). Exploring the roles of Mus81-Eme1/MMS4 at perturbed replication forks. DNA repair 6, 1004-1017. Ouyang, K.J., Woo, L.L., and Ellis, N.A. (2008). Homologous recombination and maintenance of genome integrity: cancer and aging through the prism of human RecQ helicases. Mech. Ageing Dev. 129, 425-440. Raynard, S., Bussen, W., and Sung, P. (2006). A double Holliday junction dissolvasome comprising BLM, topoisomerase IIIalpha, and BLAP75. J. Biol. Chem. 281, 13861-13864. Rothkamm, K., Kruger, I., Thompson, L.H., Lobrich, M. (2003). Pathways of DNA double-strand break repair during the mammalian cell cycle. Mol. Cellular biology 23, 5706-5715. Schechter, D., Costanzo, V., Gautier, V. (2004). ATR and ATM regulate the timing of DNA replication origin firing. Nature Cell Biology 6, 648-655. Scheller, J., Schurer, A., Rudolph, C., Hettwer, S., and Kramer, W. (2000). MPH1, a yeast gene encoding a DEAH protein, plays a role in protection of the genome from spontaneous and chemically induced damage. Genetics 155, 1069-1081. Scully, R., Ganesan, S., Vlasakova, K., Chen, J., Socolovsky, M., and Livingston, D.M. (1999). Genetic analysis of BRCA1 function in a defined tumor cell line. Mol. Cell 4, 1093-1099.

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Seal, S., Thompson, D., Renwick, A., Elliott, A., Kelly, P., Barfoot, R., Chagtai, T., Jayatilake, H., Ahmed, M., Spanova, K., et al. (2006). Truncating mutations in the Fanconi anemia J gene BRIP1 are low-penetrance breast cancer susceptibility alleles. Nat. Genet. 38, 1239-1241. Segurado, M., Gomez, M., Antequera, F. (2002). Increased recombinational intermediates and homologous integration hot spots at DNA replication origins. Molecular Cell 10, 907-916. Sengupta, S., Linke, S.P., Pedeux, R., Yang, Q., Farnsworth, J., Garfield, S.H., Valerie, K., Shay, J.W., Ellis, N.A., Wasylyk, B., Harris, C.C. (2003). BLM helicase-dependent transport of p53 to sites of stalled DNA replication forks modulates homologous recombination. The EMBO J. 22, 1210-1222. Shimura, T., Torres, M.J., Martin, M.M., Rao, V.A., Pommier, Y., Katsura, M., Miyagawa, K., and Aladjem, M.I. (2008). Bloom's syndrome helicase and Mus81 are required to induce transient double-strand DNA breaks in response to DNA replication stress. J. Mol. Biol. 375, 1152-1164. Singh, T.R., Ali, A.M., Busygina, V., Raynard, S., Fan, Q., Du, C.H., Andreassen, P.R., Sung, P., and Meetei, A.R. (2008). BLAP18/RMI2, a novel OB-foldcontaining protein, is an essential component of the Bloom helicase-double Holliday junction dissolvasome. Genes Dev. 22, 2856-2868. Singh, T.R., Saro, D., Ali, A.M., Zheng, X.F., Du, C.H., Killen, M.W., Sachpatzidis, A., Wahengbam, K., Pierce, A.J., Xiong, Y., et al. (2010). MHF1-MHF2, a histone-fold-containing protein complex, participates in the Fanconi anemia pathway via FANCM. Mol. Cell 37, 879-886. Sobeck, A., Stone, S., Landais, I., de Graaf, B., and Hoatlin, M.E. (2009). The Fanconi anemia protein FANCM is controlled by FANCD2 and the ATR/ATM pathways. J. Biol. Chem. 284, 25560-25568. Spry, M., Scott, T., Pierce, H., and D'Orazio, J.A. (2007). DNA repair pathways and hereditary cancer susceptibility syndromes. Front Biosci. 12, 4191-4207. Stewart, E., Chapman, C.R., Al-Khodairy, F., Carr, A.M., and Enoch, T. (1997). rqh1+, a fission yeast gene related to the Bloom's and Werner's syndrome genes, is required for reversible S phase arrest. Embo. J. 16, 2682-2692. Stoepker, C., Hain, K., Schuster, B., Hilhorst-Hofstee, Y., Rooimans, M.A., Steltenpool, J., Oostra, A.B., Eirich, K., Korthof, E.T., Nieuwint, A.W., et al. (2011). SLX4, a coordinator of structure-specific endonucleases, is mutated in a new Fanconi anemia subtype. Nat. Genet. 43, 138-141. Suhasini, A.N., Rawtani, N.A., Wu, Y., Sommers, J.A., Sharma, S., Mosedale, G., North, P.S., Cantor, S.B., Hickson, I.D., and Brosh, R.M., Jr. (2011). Interaction between the helicases genetically linked to Fanconi anemia group J and Bloom's syndrome. The EMBO J.

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Svendsen, J.M., Smogorzewska, A., Sowa, M.E., O'Connell, B.C., Gygi, S.P., Elledge, S.J., and Harper, J.W. (2009). Mammalian BTBD12/SLX4 assembles a Holliday junction resolvase and is required for DNA repair. Cell 138, 63-77. Umezu, K., and Nakayama, H. (1993). RecQ DNA helicase of Escherichia coli. Characterization of the helix-unwinding activity with emphasis on the effect of single-stranded DNA-binding protein. J. Mol. Biol. 230, 1145-1150. van Brabant, A.J., Stan, R., and Ellis, N.A. (2000). DNA helicases, genomic instability, and human genetic disease. Annu. Rev. Genomics Hum. Genet 1, 409-459. Wang, W. (2007). Emergence of a DNA-damage response network consisting of Fanconi anaemia and BRCA proteins. Nature Reviews Genetics 8, 735-748. Wang, Y., Cortez, D., Yazdi, P., Neff, N., Elledge, S.J., Qin, J. (2000). BASC, a super complex of BRCA1-associated proteins involved in the recognition and repair of aberrant DNA structures. Genes and Development 14, 927-939. Weiss, R.S., Matsuoka, S., Elledge, S.J., and Leder, P. (2002). Hus1 acts upstream of Chk1 in a mammalian DNA damage response pathway. Current Biol. 12, 73-77. Whitby, M.C. The FANCM family of DNA helicases/translocases. DNA Repair (Amst) 9, 224-236. Whitby, M.C. (2010). The FANCM family of DNA helicases/translocases. DNA Repair (Amst) 9, 224-236. Wu, L. (2007). Role of the BLM helicase in replication fork management. DNA Repair (Amst) 6, 936-944. Wu, L., Davies, S.L., Levitt, N.C., and Hickson, I.D. (2001). Potential role for the BLM helicase in recombinational repair via a conserved interaction with RAD51. J. Biol. Chem. 276, 19375-19381. Wu, L., Davies, S.L., North, P.S., Goulaouic, H., Riou, J.F., Turley, H., Gatter, K.C., and Hickson, I.D. (2000a). The Bloom's syndrome gene product interacts with topoisomerase III. J. Biol. Chem. 275, 9636-9644. Wu, L., and Hickson, I.D. (2003). The Bloom's syndrome helicase suppresses crossing over during homologous recombination. Nature 426, 870-874. Wu, X., Petrini, J.H., Heine, W.F., Weaver, D.T., Livingston, D.M., and Chen, J. (2000b). Independence of R/M/N focus formation and the presence of intact BRCA1. Science 289, 11. Xu, D., Guo, R., Sobeck, A., Bachrati, C.Z., Yang, J., Enomoto, T., Brown, G.W., Hoatlin, M.E., Hickson, I.D., and Wang, W. (2008). RMI, a new OB-fold complex essential for Bloom syndrome protein to maintain genome stability. Genes Dev. 22, 2843-2855.

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Xue, Y., Li, Y., Guo, R., Ling, C., and Wang, W. (2008). FANCM of the Fanconi anemia core complex is required for both monoubiquitination and DNA repair. Hum. Mol. Genet. 17, 1641-1652. Yin, J., Sobeck, A., Xu, C., Meetei, A.R., Hoatlin, M., Li, L., and Wang, W. (2005). BLAP75, an essential component of Bloom's syndrome protein complexes that maintain genome integrity. Embo J. 24, 1465-1476. Zhang, R., Sengupta, S., Yang, Q., Linke, S.P., Yanaihara, N., Bradsher, J., Blais, V., McGowan, C.H., and Harris, C.C. (2005). BLM helicase facilitates Mus81 endonuclease activity in human cells. Cancer Res. 65, 2526-2531. Zhong, Q., Chen, C.F., Li, S., Chen, Y., Wang, C.C., Xiao, J., Chen, P.L., Sharp, Z.D., and Lee, W.H. (1999). Association of BRCA1 with the hRad50hMre11-p95 complex and the DNA damage response. Science 285, 747-750.

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Chapter II

Mutagenic Potential of Methacrylates Used in Restorative Dentistry

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Janusz Blasiak1,∗, Ewelina Synowiec1, Piotr Czarny1, Elzbieta Pawlowska2 and Joanna Szczepanska2 1

2

Department of Molecular Genetics, University of Lodz, Lodz, Poland Department of Pediatric Dentistry, Medical University of Lodz, Lodz, Poland

Abstract DNA damage is a prerequisite for DNA mutation, so the studies on DNA-damaging potential of environmental and medical chemicals are justified. Methacrylates are used in the polymer form as composite restorative materials in dentistry. However, the process of polymerization, led in situ, is always incomplete and the polymers can release monomers into the oral cavity and the pulp, from where they can migrate into the bloodstream reaching virtually all organs. The local concentration of the released monomers can be high enough to induce adverse biological effects. Genotoxicity of methacrylate monomers is of a special significance due to potential serious phenotypic consequences, including mutations and cancer, ∗

Corresponding Author: Department of Molecular Genetics, University of Lodz, Pomorska 141/143, 90-236 Lodz, Poland; e-mail [email protected], phone +48 42 635 43 34, fax +48 42 635 44 84.

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Janusz Blasiak, Ewelina Synowiec, Piotr Czarny et al. and a long latency period. In the present work, we investigated genotoxicity of a mixture of monomers of model methacrylate composite consisting of 45% 2-hydroxyethyl methacrylate and 55% bisphenol A-diglycidyl dimethacrylate (w/w) (HEMA/Bis-GMA) in human gingival fibroblast by measuring DNA damage and repair, apoptosis and cell cycle. The mixture displayed the ability to damage cellular DNA as measured by the alkaline and neutral comet assay indicating the presence of DNA single-strand breaks, alkali-labile sites and double-strand breaks in DNA. The ability of the composite to induce DNA double-strand breaks, the most serious DNA damage, was confirmed by pulsed field gel electrophoresis. This compound oxidatively modified the DNA bases, as checked by two DNA repair enzymes: endonuclease III and formamidopyrimidine-DNA glycosylase. HEMA/Bis-GMA induced apoptosis and disturbed the cell cycle, increasing the fraction of the cells in the G2/M checkpoint. The results obtained indicate that the HEMA/Bis-GMA model methacrylate composite, which can be considered as a representative for methacrylate-based dental restorations, displays a broad spectrum of genotoxicity and so – a high mutagenic potential. Therefore, new formulations of materials should be created with the emphasis on the decrease of their mutagenic potential.

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Keywords: DNA damage; DNA repair; mutation; methacrylates; dental composites; apoptosis; cell cycle; HEMA; Bis-GMA

Introduction Mutations in DNA are consequences of non-repaired or misrepaired DNA damages, which can be induced by a variety of physical and chemical agents. Therefore, DNA-damaging substances may be considered as potentially mutagenic. It is especially important when such substances are in permanent contact with the human organism, like many chemicals used in medicine as drugs and biomaterials applied in a variety of restorative and aesthetic procedures. The latter include dental composites, which are usually polymers of methacrylates and 2-hydroxyethyl methacrylate (HEMA), bisphenol A-diglycidyl dimethacrylate (Bis-GMA), urethane dimethacrylate (UDMA) and triethylene glycol dimethacrylate (TEGDMA) are monomers commonly used in dentistry and orthodontics in polymeric form]. The process of polymerization, which is led in situ, is always incomplete, resulting in the presence of free monomers in the oral cavity [1]. Moreover, unpolymeryzed monomers may migrate into the pulp, through the microchannels present in the dentin. Additionally, monomers can be

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Mutagenic Potential of Methacrylates Used in Restorative Dentistry

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released from polymer-based tooth restorations in the result of mechanical shearing and enzymatic degradation of polymers. Polymethacrylates may contain hydrolyzable ester groups at their surface and the action of esterases may release products of degradations of monomers into the oral cavity. Resulting concentrations of methacrylate monomers and products of their degradation in the pulp may be in the millimolar range, sufficiently high to induce adverse biological effects [2, 3]. The monomers may migrate with the bloodstream from the pulp into virtually any organ of the organism, where the concentration may be much lower than in the pulp. However, the release of monomers from the restoration may last for many years and may result in the accumulation of biological effects. Although methacrylate esters possess a similar spectrum and pattern of toxicity [4], their genotoxicity seems to be of a special interest for several reasons. First, genotoxic substances may induce DNA damage, which is a prerequisite to DNA mutations, which, in turn, may be crucial for cancer and severe genetic diseases. Second, genotoxic effects may not require a high concentration of a substance, because changes in one cell may result in its proliferation and clonal expansion of the changes. Third, genotoxic action may induce delayed phenotypic effects and if such action affects the germ cells, the effects will also be expressed in the next generation(s). The genotoxicity of methacrylates was shown in several studies, but few of them addressed a direct interaction with DNA [1, 5-14]. The common use of methacrylates, not only in restorative dentistry but in other branches of medicine and industry, along with the importance of DNA mutations in pathology, cause that they must meet a variety of regulatory mutagenicity testing requirements [15]. Mutagenicity testing has been performed on many methacrylates indicating that these compounds should be non-mutagenic in the whole animal [15]. Moreover, as a number of methacrylates are used in materials having contact with food, they must meet the requirements of food contact notification around the world. Methacrylates used in dentistry are of special concern, because they may be released permanently to the bloodstream. There are some conflicting results and relationships concerning mutagenicity and the DNA-damaging potential of methacrylate monomers. Bis-GMA is considered as one of the most both cyto- and genotoxic methacrylates used in dentistry and recently, it was shown to induce DNA double-strand breaks, with a much higher efficacy than UDMA, TEGDMA and HEMA [16]. Such a high cytototoxic and genotoxic potential of Bis-GMA was also shown in other studies [8, 17]. On the other hand, TEGDMA and UDMA were shown to induce point mutations and chromosomal aberrations in the Somatic Mutation and Recombination Test (SMART), whereas HEMA and Bis-GMA did not show any genotoxic properties in the same assay [18]. These authors obtained a similar relationship in their other

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Figure 1. Chemical structure of 2-hydroxyethyl methacrylate (HEMA) and bisphenol Adiglycidyl dimethacrylate (Bis-GMA).

work performed with two commercial dental bonding agents containing BisGMA, UDMA and HEMA [19]. Some dental methacrylates were shown positive in mammalian cell mutagenesis, as can be found in the Chemical Carcinogenesis Research Information System (http://toxnet.nlm.nih.gov/cgibin/sis/htmlgen? CCRIS). It is beyond the scope of this paper to discuss the relationship between mutagenicity and DNA-damaging potential of a chemical, but some solid evidence of such potential related to dental monomers should provoke research on the protection against it, despite negative results obtained in standard mutagenicity testing. The objective of this work was to assess the mutagenic potential expressed by DNA-damaging ability of methacrylate monomers contained in a model dental adhesive, composed of HEMA and Bis-GMA, 45/55% w/w ratio (HEMA/BisGMA), in human gingival fibroblasts (HGFs). The structure of the monomers is presented in Figure 1. To explore the mechanism underlying the DNA-damaging action of this model adhesive, the studies were assisted by investigation of DNA repair, cell viability, apoptosis and cell cycle.

Materials and Methods Chemicals HEMA, Bis-GMA, gradisol and RNase A, low melting point (LMP) and normal melting point (NMP) agarose, phosphate buffered saline (PBS), DAPI (4’,6-diamidino-2-phenylindole), dimethyl sulfoxide (DMSO), fetal bovine serum

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(FBS), MTT, lectin, penicillin, streptomycin, Bradford reagent were from were from Sigma Chemicals (St. Loius, MO, USA). Quantum 333 medium, Dulbecco’s phosphate buffered saline (DPBS), trypsin and EDTA were from PAA Laboratories GmbH (Cölbe, Germany). Methanol-free formaldehyde solution was from Thermo Fisher Scientific (Worcester, MA, USA). Plasmid DNA purification kit was provided by EURx (Gdansk, Poland). CleanCut agarose, proteinase K reaction buffer, wash buffer and Saccharomyces cerevisiae molecular marker were obtained from Bio-Rad (Hercules, CA, USA). Cell viability, apoptosis and cell cycle kits were purchased in BD Bioscences (San Jose, CA, USA). ApoAlert Caspase Colorimetric Assay Kit was bought from Clontech Laboratories Inc (Palo Alto, CA, USA). All other chemicals were of the highest commercial grade available.

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Cells Human gingival fibroblasts (HGFs) cell line was purchased from Provitro (Berlin, Germany). The cells were grown in Quantum 333 medium containing Lglutamine and supplemented with 1% antibiotic-antimycotic solution (10,000 Units/ml penicillin, 10 mg/ml streptomycin sulphate, 25 µg/ml amphotericin B) in 75 cm2 cell culture flasks to approximately 75-80% confluence and maintained in an incubator with 5% CO2 atmosphere at 100% humidity at 37°C. After reaching confluence, the cells were washed with Dulbecco’s phosphate buffered saline, detached from the flasks by a brief treatment with 0.05% trypsin-0.02% EDTA. Escherichia coli, strain DH5α cells with pUC19 plasmid, were grown in a LB broth at 37°C overnight.

Cell Treatment HEMA/Bis-GMA model composite was added from its solution in DMSO to the cells in their growth medium at final concentrations from the range 0.01-0.20 mM. The control cells received only growth medium and DMSO at 0.3%. DMSO at this concentration did not affect the processes under study (results not shown). To examine DNA damage, cell viability, apoptosis and cell cycle, the cells were incubated with HEMA/Bis-GMA for 6 h at 37°C. Each DNA damage experiment included a positive control, which was hydrogen peroxide at 20 µM for 15 min on ice [20]. H2O2 at this concentration produced a pronounced DNA damage, which

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Janusz Blasiak, Ewelina Synowiec, Piotr Czarny et al.

resulted in the tail DNA of 30-40%. Positive controls in the remaining experiments were included in the appropriate kits and are described below.

Cell Viability HGFs were washed three times with PBS and then diluted in PBS to concentration of 2.5 × 105 cells/ml. For preparation of dead cells (positive control), one sample was treated with 96% ethanol for 1 min. All samples were centrifuged and cell pellets were suspended in 100 µl of 0.5 µM calceinacetoxymethyl ester (cal AM)/10 µM propidium iodine (PI) in PBS. Cells were gently shaken and incubated with the dyes for 30 min at 37°C in a tissue culture incubator and then analyzed on a LSRII flow cytometer (Becton Dickinson, San Jose, USA) equipped with 488 nm laser excitation and BD FACS Diva software v 4.1.2. 5×104 cells were analyzed in each experiment repeated in triplicate.

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Plasmid Relaxation Assay pUC19 plasmids were isolated from DH5α Escherichia coli cells with Genematrix Plasmid Miniprep DNA Purification Kit (EURx, Gdansk, Poland) according to the manufacturer instruction. Plasmids were exposed to UV irradiation at 35 J/m2 (positive control) to check the migration of its multimeric forms (supercoiled, nicked circular and linear). UV irradiation induced strand breaks in DNA and caused the relaxation of supercoiled plasmid – one break is enough to relax one molecule of it. Structural differences between supercoiled, nicked circular and linear forms of the plasmid accounted for their different electrophoretic mobility. Plasmid samples at 250 ng/μl were subjected to 1% agarose gel electrophoresis carried out in TBE (Tris-Borate-EDTA) buffer. The gel was stained with ethidium bromide (0.5 mg/ml) and the DNA was visualized under ultraviolet light (302 nm), scanned by a CCD camera, and densitometry analysis was performed with the GeneTools by Syngene (Cambridge, UK) software. UV irradiation was performed at 4°C with UVC-6-12 lamp (NeoLab, Heidelberg, Germany) emitting UV light at 254 nm at a dose rate of 0.12 J m-2 s-1. The ability of HEMA and MAA to damage DNA was quantified by calculating the ratio of the open circular DNA to the total amount of DNA (R). The values for

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supercoiled DNA were multiplied by 1.66 to correct for the decreased intercalating ability of ethidium bromide [21].

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Comet Assay The comet assay was performed under alkaline conditions essentially according to the procedure of Singh et al. [22] with modifications [23] as described previously [24]. A freshly prepared suspension of HGFs in 0.75% LMP agarose dissolved in DPBS was spread onto microscope slides pre-coated with 0.5% NMP agarose. The cells were then lysed for 1 h at 4°C in a buffer consisting of 2.5 M NaCl, 100 mM EDTA, 1% Triton X-100, 10 mM Tris, pH 10. After lysis, the slides were placed in an electrophoresis unit, the DNA was allowed to unwind for 40 min in the electrophoretic solution consisting of 300 mM NaOH, 1 mM EDTA, pH > 13. Electrophoresis was conducted at 4°C (the temperature of the running buffer did not exceed 12°C) for 20 min at an electric field strength of 0.73 V/cm (29 mA). The slides were then neutralized with 0.4 M Tris, pH 7.5, stained with 2 μg/ml DAPI and covered with cover slips. To prevent additional DNA damage, all of the steps described above were conducted under dimmed light or in the dark. In the neutral version of the comet assay, electrophoresis was run in a buffer consisting of 100 mM Tris and 300 mM sodium acetate at pH adjusted to 9.0 by glacial acetic acid [25]. Electrophoresis was conducted for 60 min, after a 20 min equilibrium period, at electric field strength of 0.41 V/cm (50 mA) at 4°C. The slides were examined at 200× magnification in an Eclipse fluorescence microscope (Nikon, Tokyo, Japan) attached to a COHU 4910 video camera (Cohu, Inc., San Diego, CA) equipped with a UV filter block consisting of an excitation filter (359 nm) and barrier filter (461 nm) and connected to a personal computer-based image analysis system, Lucia-Comet v. 4.51 (Laboratory Imaging, Praha, Czech Republic). Fifty images were randomly selected from each sample and the comet tail DNA was measured. Two parallel tests with aliquots of the same sample of cells were performed for a total of 100 cells. Each experiment was repeated three times. The percentage of DNA in the tail (% tail DNA) was analyzed. It is positively correlated with the level of DNA breakage or/and alkali labile sites in the cell and is negatively correlated with the level of DNA crosslinks [20]. For the neutral version, this quantity correlates positively with DNA double-strand breaks. The mean value of the % tail DNA in a particular sample was taken as an index of the DNA damage in this sample.

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Oxidative Damage to DNA To assess the ability of HEMA/Bis-GMA to induce oxidative damage to DNA, we used endonuclease III (Endo III) and formamidopyrimidine-DNA glycosylase (Fpg), which are enzymes of the base excision DNA repair pathway. Endo III converts oxidized pyrimidines into strand breaks, which can be detected by the comet assay [26]. Fpg recognizes and removes 7,8-dihydro-8-oxoguanine (8-oxoguanine), the imidazole ring-opened purines 2,6-diamino-4-hydroxy-5formamidopyrimidine (Fapy-Gua) and 4,6-diamino-5-formamidopyrimidine (Fapy-Ade) as well as small amounts of 7,8-dihydro-8-oxo-adenine (8oxoadenine) [27]. These enzymes recognize also a variety of other modification to the DNA bases, which cannot be detected by the comet assay in its basic form. The removing of modified bases from DNA by the enzymes leads to apurinic or apyrimidinic sites, which are subsequently cleaved by their AP-lyase activity producing a gap in the DNA strand, which can be detected by the comet assay [23]. After the incubation with HEMA/Bis-GMA and lysis in 2.5 M NaCl, 0.1 mM Na2EDTA, 10 mM Tris-HCl, 1% Triton X-100, pH 10.0, for 1 h at 4°C, the slides from the comet assay were washed three times in the enzyme buffer: 40 mM HEPES-KOH, 0.1 M KCl, 0.5 mM EDTA, 0.2 mg/ml bovine serum albumin, pH 8.0 and drained, and the agarose was covered with 30 µl of the enzyme buffer or the enzyme at 1 µg/ml in buffer, sealed with a cover glass and incubated for 30 min at 37°C [28]. Further steps were as described in the Comet assay section. To check the ability of both enzymes in order to recognize the oxidative damage to DNA, we exposed HGFs to 20 μM hydrogen peroxide for 15 min on ice (positive control, results not shown). Since the enzymatic buffer induced DNA damage, the values of the %DNA in tail in the buffer were subtracted from the value for the enzyme with the buffer. To express oxidative modification to the DNA bases evoked by HEMA/Bis-GMA at a given concentration, %DNA in tail for the control (no HEMA/Bis-GMA) was subtracted from %DNA in tail for the mixture at this concentration.

DNA Double-Strand Breaks Assays We evaluated the ability of HEMA/Bis-GMA to induce DNA double-strand breaks (DSBs) by the neutral comet assay and pulsed field gel electrophoresis. Aliquots of 5×107 cells/ml were washed 3 × in PBS by centrifugation (1000 × g, 15 min, 4°C) and mixed with 2% CleanCut agarose to a final concentration of 0.75% at 50°C. The cells/agarose mixtures were transferred to plug molds. After

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solidifying, agarose plugs were lysed in Proteinase K Reaction Buffer with proteinase K (Bio-Rad) and were incubated overnight at 50°C. Thereafter, plugs were washed four times in a 1 × wash buffer (Bio-Rad) and stored at 4°C in this buffer. Washed plugs were inserted into the wells of a 1.0% agarose gel prepared in 0.5 × TBE buffer. The gel was then placed horizontally in a gel box of the contour-clamped homogeneous electric field gel electrophoresis apparatus CHEF II (Bio-Rad) with 120° angle between fields. Electrophoresis ran for 24 h at 6 V/cm and the time-pulse gradient was 1-500 s for 60 sec and 1500-1600 s for 120 sec at 14°C. Plugs containing the chromosomes of the yeast Saccharomyces cerevisiae were used as molecular weight standards. The gel was stained with ethidium bromide (0.5 mg/ml) and the plasmid DNA was visualized under ultraviolet light (302 nm), scanned by a CCD camera, and densitometry analysis was performed with the GeneTools by Syngene (Cambridge, UK) software. DSBs were measured as the fraction of activity released (FAR; ratio of DNA released in an electrophoretic lane and total DNA) [29].

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Apoptosis The BD Annexin V-FITC Apoptosis Detection Kit I was used to measure apoptosis. The kit contains Annexin V conjugated to the flurochrome FTIC. This complex displays a high affinity to the membrane phospholipid phosphatidylserine, which undergoes externalization in the early stages of apoptosis. Propidium iodine (PI) was used to distinguish early apoptotic cells from death cells resulted from late apoptosis or necrosis. Cells that were viable were Annexin V-FITC and PI negative, cells that were in early apoptosis were Annexin-FITC positive and PI negative and cells that were in late apoptosis or already dead were both Annexin-FITC and PI positive. After 6 h of incubation with HEMA/Bis-GMA, the cells were washed in cold PBS and re-suspended in 1 × binding buffer at 106 cells/ml. Aliquot of 100 μl (105 cells) was transferred to a 5 ml culture tube, 5 μl of Annexin V-FITC and 5 μl of PI were added, gently vortexed and incubated for 15 min at room temperature in the dark. Then, 400 μl of 1 × binding buffer was added to each tube and samples were analyzed by flow cytometry. Each experiment had negative, positive and unstained control samples. About 10,000 events were counted per sample. The apoptosis ratio was calculated as a percent of apoptotic cells in a sample.

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Cell Cycle The CycleTEST PLUS DNA Reagent Kit was used to determine the DNA index (DI) and cell-cycle phase distributions. Nuclei were isolated, stained with propidium iodine and afterward analyzed on the LSRII flow cytometer according to the manufacturer instruction. The DI was calculated by dividing the mean of the relative content of the exposed G0/G1 population by the mean of the control G0/G1 population. The suspension of cells was washed 3 times in the manufacturer-provided Buffer Solution, adjusted to a concentration of 106 cells/ml and then stained according to the manufacturer instruction. Results were analyzed by CellFIT software.

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Data Analysis The values in this study were expressed as mean ± S.E.M. from three experiments, i.e. the data from three experiments were pooled and the statistical parameters were calculated. The data obtained from cell viability was expressed as mean ± S.D. The Mann-Whitney test was used to determine differences between samples with distributions departing from normality. The differences between samples with the normal distribution were evaluated by applying the Student’s t-test. Data analysis was performed using SigmaStat software (v. 3.0.0, SPSS, Chicago, USA).

Results Cell Viability HEMA/Bis-GMA evoked a concentration-dependent decrease in the viability of HGFs with almost all cells dying at 0.5 and 1 mM (Figure 2). The EC50 value for this mixture was estimated to be 0.12 mM.

DNA Damage In Vitro HEMA/Bis-GMA did not introduce DNA breaks to isolated DNA, as evaluated by the plasmid relaxation assay, in which the ratio of the amount of

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open circular forms of plasmid DNA to the total amount of DNA was calculated (Table 1). The pUC19 plasmid used in our experiment was sensitive to DNAbreaking agents as checked by UV irradiation.

Figure 2. Viability of human gingival fibroblasts exposed for 6 h at 37°C to the mixture of methacrylates containing 45% 2-hydroxyethyl methacrylate and 55% bisphenol Adiglycidyl dimethacrylate (w/w) (HEMA/Bis-GMA) in dependence on Bis-GMA concentration. The viability was measured by flow cytometry with thiazole orange and propidium iodide. Displayed is the mean of three experiments of 5 × 104 measurements each, error bars denote standard deviation. The inset presents the viability at low concentrations of Bis-GMA. The radius of symbol is greater than the bars length for the highest concentrations of the mixture. *** – p < 0.001, ** – p < 0.01, p < 0.05 as compared with unexposed control.

Table 1. The ratio of the open circular DNA to the total amount of DNA (R) of the isolated pUC19 plasmid exposed for 6 h at 37°C to the mixture of methacrylates containing 45% 2-hydroxyethyl methacrylate and 55% bisphenol A-diglycidyl dimethacrylate (w/w) (HEMA/Bis-GMA) in dependence on Bis-GMA concentration. The values for supercoiled DNA were multiplied by 1.66 to correct for the decreased intercalating ability of ethidium bromide

***

Bis-GMA (mM)

R

0.00 0.05 0.10 0.20 UV (positive control)

0.102 0.097 0.091 0.106 0.430***

– p < 0.001.

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Figure 3. DNA damage in human gingival fibroblasts exposed for 6 h at 37°C to the mixture of methacrylates containing 45% 2-hydroxyethyl methacrylate and 55% bisphenol A-diglycidyl dimethacrylate (w/w) (HEMA/Bis-GMA) in dependence on Bis-GMA concentration. DNA damage was measured as percentage in the tail DNA in comets of alkaline (empty symbols) or neutral (filled symbols) version of the comet assay. The mean value for one hundred cells analyzed at each concentration in three independent experiments is displayed, error bars represent SEM, **p < 0.01, ***p < 0.001 as compared with unexposed controls.

DNA Damage in Human Gingival Fibroblasts Figure 3 displays the mean percentage tail DNA of human gingival fibroblasts exposed for 6 h to HEMA/Bis-GMA and analyzed by the comet assay in its alkaline and neutral versions. The composite increased tail DNA in a concentration-dependent manner (p < 0.001 at all concentrations in the alkaline version). The alkaline version detects single- and double-strand breaks as well as alkali labile sites.

DNA Double-Strand Breaks We employed the neutral comet assay to screen for DNA double-strand breaks (DSBs). We observed a significant increase in the tail DNA for all tested concentrations of HEMA/Bis-GMA (Figure 3). The positive results obtained in this assay were verified by using pulsed field gel electrophoresis that is considered as a more reliable method than neutral comet assay to do so. The results (Figure 4) confirm the ability of the compound to induce DSBs.

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Figure 4. Fraction of the activity released (FAR) in pulsed field gel electrophoresis of DNA from human gingival fibroblasts exposed for 6 h at 37°C to the mixture of methacrylates containing 45% 2-hydroxyethyl methacrylate and 55% bisphenol Adiglycidyl dimethacrylate (w/w) (HEMA/Bis-GMA) in dependence on Bis-GMA concentration. Presented are means of three independent experiments, error bars denote SD, * - p < 0.05, ** - p < 0.01, *** – p < 0.001 as compared with unexposed control.

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DNA Repair We analyzed the kinetics of DNA repair in human gingival fibroblasts after HEMA/Bis-GMA treatment at 0.1 mM Bis-GMA by measuring the extent of DNA damage in the cells immediately after exposure as well as 30, 60, 90 and 120 min thereafter with the alkaline comet assay (Figure 5). The cells exposed to 10 µM hydrogen peroxide (positive control) were able to recover within 45 min (results not shown). The cells exposed to HEMA/Bis-GMA removed damage to their DNA in a time-dependent manner and they removed more than 50% of the initial damage on 120-min incubation (p < 0.001). The kinetics of removing of DNA damage was almost exactly linear since 30 min of the repair incubation. The control cells displayed an elevated level of DNA damage after a 60-min incubation and later, which was probably due to the prolonged time the cells were out of incubator.

Oxidative Modifications to the DNA Bases Figure 6 presents the mean % tail DNA of human gingival fibroblasts exposed for 6 h at 37°C to HEMA/Bis-GMA at 0.05 or 0.10 mM Bis-GMA, lysed and post-treated with Endo III or Fpg, reduced by mean % tail DNA for cells

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without treatment with any enzyme and cells incubated only with an enzymatic buffer. As a result, we analyzed only these modifications to the DNA bases, which were not recognized in the non-modified, alkaline version of the comet assay. The cells exposed to HEMA/Bis-GMA and treated with either enzyme showed greater % tail DNA than those untreated with any enzyme (p < 0.01 at 0,05 mM BisGMA and p < 0.001 at 0.10 mM Bis-GMA for both enzymes). This indicates that oxidative modifications to the DNA bases play a role in the genotoxic action of HEMA/Bis-GMA. A significant DNA damage observed in the cells not exposed to the model composite was likely to arise from the endogenous oxidative damage to the DNA bases as well as from the oxidative damage to DNA introduced during processing of the cells. The cells exposed to 20 µM H2O2 displayed a significant extent of DNA bases modifications recognized by Endo III or Fpg (data not shown).

Figure 5. Time-course of DNA repair in human gingival fibroblasts exposed for 6 h at 37°C to the mixture of methacrylates containing 45% 2-hydroxyethyl methacrylate and 55% bisphenol A-diglycidyl dimethacrylate (w/w) (HEMA/Bis-GMA) (filled symbols). After the exposure, the cells were washed and incubated in a composite-free medium at 37°C. Also displayed is the extent of DNA damage in control cells not exposed to the composite (empty symbols). The number of cells analyzed in each time-interval was 100. The results are mean of three independent experiments. Error bars denote SEM, ** – p < 0.01, *** – p < 0.001 as compared with the initial DNA damage.

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Figure 6. Oxidative DNA base modifications evoked by the mixture of methacrylates containing 45% 2-hydroxyethyl methacrylate and 55% bisphenol A-diglycidyl dimethacrylate (w/w) (HEMA/Bis-GMA) during 6 h exposure at 37°C measured as percentage of DNA in the tail in the alkaline comet assay with endonuclease III (empty bars) and formamidopyrimidine-DNA glycosylase (filled bars) at 1 µM. The value of comet tail DNA in the presence of either enzyme was reduced by the value obtained in comet assay without any enzyme and the value for enzymatic buffer only. The number of cells analyzed for each sample was 100. The results are the mean of three independent experiments. Error bars denote SEM, ** – p < 0.01, *** – p < 0.001 as compared with the unexposed to the composite enzyme-treated control.

Apoptosis HEMA/Bis-GMA induced apoptosis in HGFs in a concentration-dependent manner (Figure 7). We observed about three times increase in the apoptosis ratio for the highest concentration of the mixture, corresponding to 2 mM Bis-GMA.

Cell Cycle In order to assess the influence of HEMA/Bis-GMA on the progression of the cell cycle of HGFs, we determined the DNA content in specific phases and check points of the cycle by flow cytometry (Figure 8). In general, the monomers of the model composite evoked an increase in the G0/G1 cell population, accompanied by a mild decrease in the S phase cell population and an increase in the G2/M population. The increase in the G2/M population at 0.15 mM HEMA/Bis-GMA was significant (p < 0.05).

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Figure 7. Apoptosis of human gingival fibroblasts exposed for 6 h at 37°C to the mixture of methacrylates containing 45% 2-hydroxyethyl methacrylate and 55% bisphenol Adiglycidyl dimethacrylate (w/w) (HEMA/Bis-GMA). Apoptosis was assessed by flow cytometry with Annexin V-FITC/propidium iodine. Displayed is the mean of three experiments of 5 × 104 measurements each, error bars denote standard error. The apoptosis was expressed as a ratio of the number of early and late apoptotic cells to the number of cells with no measurable apoptosis; *p < 0.05 as compared with unexposed control.

Figure 8. Cell cycle analysis in human gingival fibroblasts exposed for 6 h at 37°C to the mixture of methacrylates containing 45% 2-hydroxyethyl methacrylate and 55% bisphenol A-diglycidyl dimethacrylate (w/w) (HEMA/Bis-GMA) for 6 h at 37°C at 0,1 mM BisGMA. Percentage of cells in the G0/G1 (white bars), S (grey bars) and G2/M (dark grey bars) stage of the cell cycle after treatment with HEMA/Bis-GMA is presented. Data is expressed as means of three independent experiments, error bars denote SD, * – p < 0.05 as compared with unexposed control. The Bis-GMA axis is not linear.

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Discussion In the present work, we showed the DNA-damaging potential of methacrylate monomers present in the mixture of HEMA and Bis-GMA at 45%/55% (w/w) proportion. Such mixture is considered as a base of a model dental composite. The mixture decreased the viability of the human gingival fibroblasts and did not interact with an isolated DNA. Therefore, the interaction between the methacrylates and cellular components other than DNA may be important for their genotoxic effects. It is not easy to predict how such effects would be modulated by a complex interaction between cells within the organism. Therefore, we cannot precisely assess the phenotypic consequences of the observed genotoxic and cytotoxic effects for the organism. As human gingival fibroblasts are permanently dividing cells, non-repaired or misrepaired DNA damage may result in a mutation. Because the mixture of the methacrylates we employed in the present work is considered as a component of the model dental composite, we can speculate that methacrylate-based dental materials have a mutagenic potential for humans. This is of a special concern, because dental restorations are common and are usually carried for many years. The genotoxicity of methacrylate-based dental materials in vivo was shown by Di Pietro et al. [6]. It is difficult to assess the actual concentration of the methacrylate monomers which can be released from the restorations. However, a mutagenic effect of a chemical may be concentrationindependent, because a single DNA damage may result in a mutation which may lead to a cancer phenotype of a cell and its clonal expansion resulting in a tumor. However, the increase in concentration may be associated with the increase in the probability of a mutation. Recently, Koin et al showed the release of Bis-GMA and other products containing bisphenol A moiety from model dental composites [30]. In the present study, we used human gingival fibroblasts because they are the nearest target in the action of methacrylate monomers released from tooth restorations. Other targets include pulp cells, which can be reached through microchannels in the dentin, and endothelial cells in the oral cavity. However, micro-injuries generated in the oral cavity by food or tooth brushing, may create another route of migration of the monomers into the bloodstream. We observed a significant decrease in the viability of HGFs exposed to HEMA/Bis-GMA, which reached about 90% at 0.5 mM Bis-GMA (Figure 2). Recently, Urcan et al. measured EC50 for HEMA and Bis-GMA in HGFs to be 11.20 and 0.09 mM, respectively [16]. Therefore, the cytotoxicity of Bis-GMA is two order of magnitude greater than that of HEMA and we can assume that the

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cytotoxic effects exerted by HEMA/Bis-GMA were determined by Bis-GMA, if no special interaction occurred between the two compounds. We obtained a slightly greater value for EC50 for Bis-GMA than Urcan et al. [16]. However, they applied the XTT viability assay, with a 24 h exposure to HEMA, whereas the exposure time in our experiment was 6 h. If we assume a linear relationship between the viability and exposure time, which was roughly the case in our experiment, the results on viability obtained by Urcan et al. and us would be similar. Methacrylate monomers can reach millimolar concentrations at least in the pulp and that is why they are used at relatively high concentrations of up to 2.5 × 10-2 mol/L and above, in many in vitro studies [8, 12]. We have included the results of the plasmid relaxation experiment, although they are negative, because we consider them important for the study. Comparing the results of this experiment with the results of the comet assay we drew a conclusion that HEMA/Bis-GMA had to be activated by cellular components to interact with the cellular DNA. HEMA/Bis-GMA induced DNA damage, which resulted in a significant increase of the percentage of DNA in the tail of comets in the alkaline (pH > 13) version of the comet assay. This version enables detection of single- and double DNA strand breaks (SSBs and DSBs, respectively) as well as alkali labile sites. The results obtained in the neutral version, designated for detection of DSBs, suggest the ability of the chemical to induce such kinds of DNA damage. However, the neutral version of the comet assay cannot be considered as the most reliable method for assessing DSBs because SSBs may interfere with measuring the breaks in this assay. This interference occurs because the relaxation of DNA supercoils, which is essential for the picture of a comet, may occur at both neutral and alkaline pH. In other words, all positive cases in the neutral comet assay should be verified and this is the practice in our laboratory [31, 32]. Pulsed field gel electrophoresis is considered as a more consistent method for assessing DSBs than neutral comet assay [33]. Our analysis with PFGE confirmed the results obtained in the neutral comet assay. The ability of HEMA/Bis-GMA to induce DSBs is of a special concern because such DNA damage, if not repaired or misrepaired, may result in chromosome rearrangements and deletions, leading to fusion genes expressing oncogenic proteins [34]. Apart from chromosomal rearrangement, DSBs may result in the inactivation of tumor suppressor genes, activation of oncogenes and disturbing the structure of mutator genes, which may result in their aberrant expression, which, in turn, may support cancer transformation [35]. We showed that HEMA/Bis-GMA modified oxidatively the DNA bases of HGFs. The modified bases were directly detected by the DNA repair enzymes

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EndoIII and Fpg. However, the total oxidative damage to DNA was surely greater than we observed in the experiment with these enzymes, because they were not able to recognize all possible oxidative modifications to the DNA bases [36]. Moreover, a fraction of the total DNA damage observed in the alkaline comet assay might be also of oxidative origin. Furthermore, an oxidative damage to DNA may be localized in the sugar-phosphate backbone, where it may not be detected by Endo III or Fpg [37]. Our observations confirm our previous reports and other studies indicating pro-oxidative potential of methacrylate monomers [38-41]. The ability of HEMA/Bis-GMA to induce oxidative modifications to the DNA bases suggests that the general mechanism of cellular damage by methacrylate-based dental materials includes the oxidative action [5, 42]. The primary reaction of the cell to DNA damage is its repair, but if the DNA repair systems cannot handle the damage, they may require a prolonged repair time resulting from the activation of the G2/M or G1/S checkpoint or the cell may undergo apoptosis. Therefore, the ability of HEMA/Bis-GMA to generate DNA damage may also determine its ability to influence the cell cycle and induce apoptosis, which is in agreement with our previous studies and other investigations [40, 43-47]. In summary, HEMA and Bis-GMA, methacrylates used in the production of restorative materials, may induce pronounced genotoxic effects, including DNA damage, apoptosis and cell cycle disturbances. This provides the evidence on the genotoxic potential of these compounds.

Acknowledgments This work was supported by the Ministry of Science and Higher Education, grant number N N 403 188134, but the ministry was not involved in the conduct of the research and preparation of the manuscript. We thank Anna Luczynska for helping us in preparing the manuscript.

Conflict of Interest Statement The authors declare that there is not any conflict of interest.

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References

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[24] Blasiak, J., Gloc, E., Woźniak, K., Drzewoski, J., Zadrozny, M., Skorski, T. and Pertynski, T. (2003). Free radicals scavengers can differentially modulate the genotoxicity of amsacrine in normal and cancer cells. Mutat. Res., 535, 25-34. [25] Singh, N.P. and Stephens R.E. (1997). Microgel electrophoresis: sensitivity, mechanisms and DNA electrostretching. Mutat. Res., 383, 167-175. [26] Collins, A.R., Duthie, S.J. and Dobson, V.L. (1993). Direct enzymatic detection of endogenous base damage in human lymphocyte DNA. Carcinogenesis, 14, 1733-1735. [27] Krokan, H.E., Standal, R. and Slupphaug, G. (1997) DNA glycosylases in the base excision repair. Biochem. J., 325, 1-16. [28] Tudek, B., Van Zeeland, A.A., Kusmierek, J.T. and Laval, J. (1998). Activity of Escherichia coli DNA-glycosylases on DNA damaged by methylating and ethylating agents and influence of 3-substituted adenine derivatives. Mutat. Res., 407, 169-176. [29] Foray, N., Arlett, C.F. and Malaise, E.P. (1999). Underestimation of the small residual damage when measuring DNA double-strand breaks (DSB): is the repair of radiation-induced DSB complete? Int. J. Radiat. Biol., 75, 1589-1595. [30] Koin, P.J., Kilislioglu, A., Zhou, M., Drummond, J.L. and Hanley, L. (2008). Analysis of the degradation of a model dental composite. J. Dent. Res., 87, 661-665. [31] Nieborowska-Skorska, M., Stoklosa, T., Datta, M., Czechowska, A., Rink, L., Slupianek, A., Koptyra, M., Seferyncka, I., Krszyna, K., Blasiak, J. and Skorski, T. (2006). ATR-Chk1 axis protects BCR/ABL leukemia cells from the lethal effect of DNA double-strand breaks. Cell Cycle, 5, 994-1000. [32] Szaflik, J.P., Janik-Papis, K., Synowiec, E., Ksiazek, D., Zaras, M., Wozniak, K., Szaflik, J. and Blasiak, J. (2009). DNA damage and repair in age-related macular degeneration. Mutat. Res., 669, 169-176. [33] Slater, G.W. (2009) DNA gel electrophoresis: the reptation model(s). Electrophoresis, 30, 181-187. [34] Prensner, J.R. and Chinnaiyan, A.M. (2009). Oncogenic gene fusions in epithelial carcinomas. Curr. Opin. Genet. Dev., 19, 82-91. [35] Helleday, T., Lo, J., van Gent, D.C. and Engelward, B.P. (2007). DNA double-strand break repair: from mechanistic understanding to cancer treatment. DNA Repair, 16, 923-935. [36] Cadet, J., Bourdat, A.G., D’Ham, C., Duarte, V., Gasparutto, D., Romieu, A., Ravanat, J.L. (2000). Oxidative base damage to DNA: specificity of base excision repair enzymes. Mutat. Res, 462, 121-128.

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[37] Eker, A.P., Quayle, C., Chaves, I. and van der Horst, G.T. (2009). DNA repair in mammalian cells: Direct DNA damage reversal: elegant solutions for nasty problems. Cell. Mol. Life Sci., 66, 968-980. [38] Chang, M.-C., Chen, L.-I., Chan, C.-P., Lee, J.-J., Wang, T.-M., Yang, T.T., Lin, P.-S., Lin, H.-J., Chang, H.-H. and Jeng, J.-H. (2010). The role of reactive oxygen species and hemeoxygenase-1 expression in the cytotoxicity, cell cycle alteration and apoptosis of dental pulp cells induced by BisGMA. Biomaterials, 31, 8164-81171. [39] Pawłowska, E., Popławski, T., Książek, D., Szczepańska, J. and Blasiak, J. (2010). Genotoxicity and cytotoxicity of 2-hydroxyethyl methacrylate. Mutat. Res., 696, 122-129. [40] Spagnuolo, G., D’Antò, V., Valletta, R., Strisciuglio, C., Schmalz, G., Schweikl, H. and Rengo, S. (2008). Effect of 2-hydroxyethyl methacrylate on human pulp cell survival pathways ERK and AKT. J. Endod., 34, 684688. [41] Walther, U.I., Siagian, I.I., Walther, S.C., Reichl, F.X. and Hickel, R. (2004). Antioxidative vitamins decrease cytotoxicity of HEMA and TEGDMA in cultured cell lines. Arch. Oral. Biol., 49, 125-131. [42] Eckhardt, A., Gerstmayr, N., Hiller, K.A., Bolay, C., Waha, C., Spagnuolo, G., Camargo, C., Schmalz, G. and Schweikl, H. (2009). TEGDMA-induced oxidative DNA damage and activation of ATM and MAP kinases. Biomaterials, 30, 2006-2014.

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Chapter III

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DNA Mutation of the Progranulin (GRN) Gene in Familial Frontotemporal Lobar Degeneration (FTLD): A Study of the Pathology of Nine Cases R. A. Armstrong∗ Vision Sciences, Aston University, Birmingham, UK

Abstract Frontotemporal lobar degeneration (FTLD) with transactive response (TAR) DNA-binding protein of 43kDa (TDP-43) proteinopathy (FTLDTDP) is a neurodegenerative disease characterized by variable neocortical and allocortical atrophy principally affecting the frontal and temporal lobes. Histologically, there is neuronal loss, microvacuolation in the superficial cortical laminae, and a reactive astrocytosis. A variety of TDP-43 immunoreactive changes are present in FTLD-TDP including neuronal cytoplasmic inclusions (NCI), neuronal intranuclear inclusions (NII), dystrophic neurites (DN) and, oligodendroglial inclusions (GI). Many cases ∗

Corresponding Author: Dr. R.A. Armstrong, Vision Sciences, Aston University, Birmingham B4 7ET, UK. Tel: 0121-359-3611; Fax 0121-333-4220; EMail [email protected].

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R. A. Armstrong of familial FTLD-TDP are caused by DNA mutations of the progranulin (GRN) gene. Hence, the density, spatial patterns, and laminar distribution of the pathological changes were studied in nine cases of FLTD-TDP with GRN mutation. The densities of NCI and DN were greater in cases caused by GRN mutation compared with sporadic cases. In cortical regions, the commonest spatial pattern exhibited by the TDP-43 immunoreactive lesions was the presence of clusters of inclusions regularly distributed parallel to the pia mater. In approximately 50% of cortical gyri, the NCI exhibited a peak of density in the upper cortical laminae while the GI were commonly distributed across all laminae. The distribution of the NII and DN was variable, the most common pattern being a peak of NII density in the lower cortical laminae and DN in the upper cortical laminae. These results suggest in FTLD-TDP caused by GRN mutation: 1) there are greater densities of NCI and DN than in sporadic cases of the disease, 2) there is degeneration of the corticocortical and cortico-hippocampal pathways, and 3) cortical degeneration occurs across the cortical laminae, the various TDP-43 immunoreactive inclusions often being distributed in different cortical laminae.

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Keywords: Frontotemporal lobar degeneration with TDP-43 proteinopathy (FTLD-TDP), TAR DNA-binding protein (TDP-43), Progranulin (GRN) mutation, Spatial topography

Introduction Frontotemporal lobar degeneration (FTLD) is the second commonest form of cortical dementia of early-onset after Alzheimer’s disease (AD) (Tolnay and Probst, 2002; Josephs, 2008). The disorder is associated with a heterogeneous group of clinical syndromes including frontotemporal dementia (FTD), FTD with motor neuron disease (FTD/MND), progressive non-fluent aphasia (PNFA), semantic dementia (SD), and progressive apraxia (PAX) (Snowden et al., 2007). FTLD with transactive response (TAR) DNA-binding protein of 43kDa (TDP-43) proteinopathy (FTLD-TDP), previously called FTLD with ubiquitin positive inclusions (FTLD-U), is characterized by variable neocortical and allocortical atrophy principally affecting the frontal and temporal lobes. In addition, there is neuronal loss, microvacuolation in the superficial cortical laminae, and a reactive astrocytosis (Cairns et al., 2007a). A variety of TDP-43 immunoreactive lesions are present in FTLD-TDP including neuronal cytoplasmic inclusions (NCI), neuronal intranuclear inclusions (NII), oligodendroglial inclusions (GI), and dystrophic neurites (DN).

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Heterogeneity of FTLD-TDP There are several sources of pathological heterogeneity within FTLD-TDP. First, a number of genetic defects have been identified in these cases; many being caused by DNA mutations of the progranulin (GRN) gene (Baker et al., 2006; Cruts et al., 2006; Mukherjee et al., 2006; Mackenzie et al., 2006a; Behrens et al., 2007; Rademaker and Hutton, 2007). A less prevalent disorder, FTLD with valosin-containing protein (VCP) gene mutation (Forman et al., 2006), also has TDP-43 immunoreactive inclusions and recently, variants in the ubiquitin associated binding protein 1 (UBAP1) gene (Luty et al., 2008; Rollinson et al., 2009) were shown to have TDP-43 inclusions. FTLD caused by charged multivesicular body protein 2B gene mutations (CHMP2B), however, has ubiquinated but no TDP-43 immunoreactive inclusions (Van der Zee, 2007). Second, four or five pathological subtypes of FTLD-TDP have been proposed based on the predominant type of inclusion present as detected with anti-ubiquitin immunohistochemistry (IHC), and the distribution and density of the pathological changes in the cortex (Mackenzie et al., 2006b; Sampathu et al., 2006; Cairns et al., 2007b; Josephs, 2008; Mackenzie et al., 2009). Patterns of histology based solely on cortical pathology include the systems of Sampathu et al. (2006) and Neumann et al. (2007) whereas Mackenzie et al. (2006b) propose a system that includes both cortical and dentate gyrus inclusions. More recently, Josephs (2008) has proposed five subtypes of FTLD-TDP and Mackenzie et al. (2009) four subtypes plus a group containing unclassifiable cases. Third, FTLD can occur in combination with MND (FTLD-MND) and such cases are often associated with a more localized pattern of frontal lobe atrophy (Whitwell et al., 2006). Fourth, a proportion of FTLD-TDP cases have coexisting hippocampal sclerosis (HS) in which there is neuronal loss in the subiculum and sector CA1 of the hippocampus (Josephs et al., 2006; Amandir-Ortiz et al., 2007). A significant degree of Alzheimer’s disease (AD) pathology, viz., senile plaques (SP) and neurofibrillary tangles (NFT) are also present in some cases.

DNA Mutations The majority of GRN mutation cases come from two families, viz. hereditary dysphasic disinhibition dementia (HDDD) family HDDD1 (Behrens et al., 2007 and family HDDD2 (Mukherjee et al., 2006). Several different frame-shift and

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premature termination DNA mutations have been identified in these cases (Beck et al., 2008). For example, HDDD2 may be caused by a novel missense mutation (Ala-9Asp) within the signal peptide of GRN (Mukherjee et al., 2006). No abnormal accumulation of GRN protein occurs as a result of these mutations but there is abnormal accumulation of TDP-43. Abnormal protein products, such as TDP-43, may accumulate within the endoplasmic reticulum of the cell due to inefficient secretion or mutant RNA may have a lower expression within the cell at least in some mutants (Mukherjee et al., 2006). TDP-43 is a nuclear protein but in FTLD-TDP, TDP-43 is redistributed from the nucleus to the cytoplasm, is ubiquinated, hyperphosphorylated, and then cleaved to generate C-terminal fragments (Neumann et al., 2007). These fragments accumulate to form the NCI and NII which may cause cell death.

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Objectives Inclusions incorporating abnormal protein aggregates are a common histological feature of many neurodegenerative diseases; the majority characterized by tau (tauopathies) or α-synuclein (synucleinopathies) immunoreactivity (Goedert et al., 2001). In the neocortex of the tauopathies (Armstrong et al., 1998; 1999a), synucleinopathies (Armstrong et al., 1997a; 2004) and neuronal intermediate filament inclusion disease (NIFID) (Bigio et al., 2003, Cairns et al., 2003; Josephs et al., 2003; Armstrong and Cairns, 2006), the NCI occur in clusters which exhibit a regular periodicity parallel to the pia mater. suggesting the inclusions develop in relation to specific cortico-cortical and cortico-hippocampal projections. In addition, in many neurodegenerative disorders, the density of the pathological changes varies significantly across the cortex from pia mater to white matter (Armstrong et al., 1997b; 1999b; 2000). The laminar distribution of a pathological change may reflect degeneration of specific anatomical pathways that have their cells of origin or axon terminals within particular laminae (Armstrong and Slaven, 1994). Hence, the objectives of the present study were to determine in the frontal and temporal lobe in FTLD-TDP with GRN mutation: 1) the density and distribution of the TDP-43 immunoreactive inclusions, 2) the spatial patterns of the inclusions in the neocortex and hippocampus parallel to the tissue boundary, and 3) the laminar distribution of the inclusions across the neocortex.

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Table 1. Demographic features and gross brain weight of the nine cases of frontotemporal lobar dementia with TDP-43 proteinopathy (FTLD-TDP) caused by progranulin (GRN) gene mutation. Braak score is based on density and distribution of neurofibrillary tangles (M = male, F = female) Case

Gender

A B C D E F G H J

F F M F M M M F M

Age (yrs) 74 84 67 67 63 66 79 82 65

Onset (yrs) 68 69 52 58 57 55 71 73 52

Duration (yrs) 6 15 15 9 6 11 8 9 13

Brain weight (gm) 975 570 960 880 1080 1050 1150 800 1300

Braak score 0 4 0 2 1 1 6 -

Materials and Methods

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Cases Nine cases of familial FTLD-TDP (see Table 1) with GRN mutation were obtained from the Departments of Neurology and Pathology and Immunology, Washington University, School of Medicine, St. Louis, Mo., USA. The majority (N = 7) of the GRN cases come from the HDDD2 family and the remainder (N = 2) from the HDDD1 family. All cases exhibited frontotemporal lobar degeneration with neuronal loss, microvacuolation affecting the superficial cortical laminae, and reactive astrocytosis (Cairns et al., 2007b). None of the cases had coexisting motor neuron disease (FTLD-MND) (Kersaitis et al., 2006; Josephs et al., 2005) or HS. Braak stage of the cases was based on the density and distribution of neurofibrillary tangles (NFT) (Braak et al., 2006). Histological Methods After death, the consent of the next-of-kin was obtained for brain removal, following local Ethical Committee procedures and the 1995 Declaration of Helsinki (as modified in Edinburgh, 2000). Tissue blocks were taken from the frontal lobe at the level of the genu of the corpus callosum to study the middle frontal gyrus (MFG) and the temporal lobe at the level of the lateral geniculate

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Figure 1. Familial frontotemporal lobar degeneration caused by DNA mutation of the progranulin (GRN) gene with TDP-43 proteinopathy (FTLD-TDP). TDP-43 immunoreactive lesions in the frontal cortex showing neuronal cytoplasmic inclusion (NCI), neuronal intranuclear inclusions (NII), and a dystrophic neurite (DN) (TDP-43 immunohistochemistry, bar = 50μm).

body to study the inferior temporal gyrus (ITG), parahippocampal gyrus (PHG), CA1/2 sectors of the hippocampus, and dentate gyrus (DG). Tissue was fixed in 10% phosphate buffered formal-saline and embedded in paraffin wax. Immunohistochemistry (IHC) was performed on 4 to 10µm sections with a rabbit polyclonal antibody that recognizes TDP-43 epitopes (dilution 1:1000; ProteinTech Inc., Chicago, IL).

Morphometric Measurements In the MFG, ITG, and PHG of each case, histological features were counted along strips of tissue (1600 to 3200µm in length) located parallel to the pia mater, using 250 x 50µm sample fields arranged contiguously. The sample fields were located both in the upper and lower cortex, the short edge of the sample field being orientated parallel with the pia mater and aligned with guidelines marked on the slide. Between 32 and 64 sample fields were usually necessary to sample each

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region. In the hippocampus, the features were counted in the cornu ammonis (CA) in a region extending from the prosubiculum/CA boundary to the maximum point of curvature of the pyramidal layer before it extends to join the dentate fascia via CA3 and CA4. Hence, the region sampled encompassed approximately sectors CA1 and CA2, the short dimension of the contiguous sample field being aligned with the alveus. Very little pathology was observed to extend into CA3/4 in the cases studied and these areas were not sampled. NCI have been commonly observed in the DG fascia in FTLD-TDP (Mackenzie et al., 2006b; Woulfe et al., 2001; Kovari et al., 2004) and the sample field was aligned with the upper edge of the granule cell layer. The NCI (Figure 1) are rounded, spicular, or skein-like in shape (Yaguchi et al., 2004; Davidson et al., 2007), while the GI morphologically resemble the ‘coiled bodies’ reported in various tauopathies such as corticobasal degeneration (CBD), progressive supranuclear palsy (PSP), and argyrophilic grain disease (AGD). The NII (Figure 1) are lenticular or spindle-shaped (Pirici et al., 2006) and the DN (Figure 1) are characteristically long and contorted (Hatanpaa et al., 2008).

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Measurement of Density of Inclusions The data were used to calculate the density of each of the TDP-43 immunoreactive inclusions in each brain region. Data analysis was by analysis of variance (ANOVA) with subsequent comparisons between brain regions using Fisher’s ‘protected least significant difference’ (PLSD) as a post-hoc procedure (STATISTICA software, Statsoft Inc., 2300 East 14th St, Tulsa, Ok, 74104, USA). First, the densities of each TDP-43 immunoreactive inclusion in the upper cortical laminae of neocortical regions were compared with those in sectors CA1/2 and the DG using a one-way ANOVA. A similar analysis was then carried out but substituting densities in the lower cortical laminae. Second, densities of each inclusion were compared between the upper and lower cortex using twofactor, split-plot ANOVA with brain region as a main-plot factor and cortical laminae as the sub-plot factor.

Spatial Patterns of the Inclusions To determine the spatial patterns of the TDP-43 immunoreactive inclusions parallel to the tissue boundary, the data were analyzed by spatial pattern analysis (Armstrong, 1993a; 1997; 2006). This method uses the variance-mean ratio (V/M)

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of the data to determine whether the inclusions were distributed randomly (V/M = 1), regularly (V/M < 1), or were clustered (V/M > 1) along the strip of tissue studied. Counts of inclusions in adjacent sample fields were then added together successively to provide data for increasing field sizes, e.g., 50 x 250µm, 100 x 250µm, 200 x 250µm etc., up to a size limited by the length of the strip sampled. V/M is plotted against field size to determine first, whether the clusters of inclusions were regularly or randomly distributed and second, to estimate the mean cluster size parallel to the tissue boundary. A V/M peak indicated the presence of regularly spaced clusters (Armstrong, 1993a; 1997; 2006). The statistical significance of a peak was tested using the ‘t’ distribution.

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Laminar Distribution The distribution of the TDP-43 immunoreactive lesions from pia mater to white matter was studied using methods based on those of Duyckaerts et al. (1986). Five traverses from the pia mater to the edge of the white matter were located at random within each gyrus. All pathological changes were counted manually in 50 x 250µm sample fields arranged contiguously, the larger dimension of the field being located parallel with the surface of the pia mater. An eye-piece micrometer was used as the sample field and was moved down each traverse one step at a time from the pia mater to white matter. Histological features of the section were used to correctly position the field. Counts from the five traverses were averaged to study the laminar distribution of lesions within each cortical gyrus. No attempt was made to locate precisely the boundaries between individual cortical laminae in the FTLD-TDP cases. First, the degree of pathological change and neuronal loss made laminar identification difficult. Second, identification is especially difficult in the frontal cortex because it exhibits a heterotypical structure, i.e., six laminae cannot be clearly identified even when the cortex is fully developed. Instead, variations in lesion density with distance below the pia mater were analyzed using a curve fitting procedure (STATISTICA software, Statsoft Inc., 2300 East 14th St, Tulsa, Ok, 74104, USA) (Snedecor and Cochran, 1980). For each cortical area studied, polynomials of order 1, 2, 3 … n, can be fitted successively to the data and the addition of each extra term adds a further ‘bend’ to the curve. Hence, quadratic curves are parabolic, cubic curves are ‘S’ shaped and quartic curves have three ‘bends’ and may appear to be ‘double peaked’. With each fitted polynomial, the correlation coefficients (Pearson’s ‘r’), regression coefficients, standard errors (SE), values of t, and the residual mean

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square are obtained. From these statistics, a judgment can be made as to whether a polynomial of sufficiently high degree has been fitted to the data. Hence, at each stage, the reduction in the sums of squares (SS) is tested for significance as each term is added. The analysis is continued by fitting successively higher order polynomials until either a non-significant value of F is obtained or there is little gain in the explained variance. The distribution of the pathological changes in each cortical area was then classified according to whether a single (unimodal) or double (bimodal) peak of density was present. If the distribution was unimodal, a further classification was made according to whether the density peak was located in the upper laminae, in the middle of the profile, or in the lower laminae. If a bimodal distribution was present, the data were further classified according to whether the upper density peak was greater than, equal to, or less than the lower density peak.

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Density of TDP-43 Immunoreactive Inclusions The densities of the TDP-43-immunoreactive lesions in each brain region, averaged over the nine cases, are shown in Figure 2. Significant differences in density of NCI were observed between regions in the upper cortex (F = 4.87, P < 0.01). Densities were greater in the MFG compared with ITG, PHG, and CA1/2 and greater in the DG compared with the PHG. There were no significant differences between brain regions when lower cortex data were substituted in the analysis (F = 1.93, P > 0.05). Apart from the DG, in which no GI were recorded, there were no significant differences in GI density between regions when either upper cortex data (F = 0.61, P > 0.05) or lower cortex data (F = 1.54, P > 0.05) were analyzed. There were no significant differences in the densities of NII when upper cortex data were analyzed (F = 2.56, P > 0.05), but densities were greater in the MFG compared with the ITG and DG when lower cortex data were included. Excluding the DG, there were no significant differences in DN density between regions when either upper cortex data (F = 0.25, P > 0.05) or lower cortex data (F = 0.66, P > 0.05) were included in the analysis. The analyses also suggested that the densities of the NCI were greater in the upper laminae of the MTG only (brain region x laminae: F = 4.58, P < 0.05) but there were no differences in densities of the GI, NII, or DN in the upper and lower cortex.

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Figure 2. Mean densities (50 x 250μm field, error bars indicate standard errors) of the TDP-43 immunoreactive pathological features in the frontal and temporal lobe (MFG = Middle frontal gyrus, ITG = Inferior temporal gyrus, PHG = Parahippocampal gyrus, CA1/2 = Sectors CA/2 of the hippocampus, DG = Dentate gyrus), U = Upper cortex, L = Lower cortex) of cases of frontotemporal lobar dementia with TDP-43 proteinopathy (FTLD-TDP) caused by progranulin (GRN) gene mutation. Analysis of variance (ANOVA) (1-way): 1) Comparing upper cortex, CA1/2, and DG data: NCI, F = 4.87 (P < 0.01); GI, F = 0.61 (P > 0.05; NII, F = 2.56 (P > 0.05); DN F = 0.25 (P > 0.05); 2) Comparing lower cortex, CA1/2, and DG data: NCI, F = 1.93 (P > 0.05); GI = F = 1.54 (P > 0.05); NII, F = 3.06 (P < 0.05); DN F = 0.66 (P > 0.05); 2) Two-factor (repeated measures ANOVA); NCI Brain region F = 4.18 (P < 0.05), lamina F = 2.48 (P > 0.05), Interaction F = 4.58 (P < 0.05); GI Brain region F = 0.31 (P < 0.05), lamina F = 2.53 (P > 0.05), Interaction F = 0.47 (P > 0.05); NII Brain region F = 2.36 (P < 0.05), lamina F = 1.12 (P > 0.05), Interaction F = 1.82 (P > 0.05); DN Brain region F = 64 (P < 0.05), lamina F = 2.26 (P > 0.05), Interaction F = 0.48 (P > 0.05).

The data suggest there were quantitative differences in the density of the TDP-43 immunoreactive inclusions between brain regions. First, the greatest densities of NCI were observed in the MFG and DG, densities often being significantly lower in sectors CA1/2 of the hippocampus and in other cortical areas thus confirming the DG as a significant site of temporal lobe pathology in FTLD-TDP (Woulfe et al., 2001; Kovari et al., 2004). Second, densities of the NII were also greater in the frontal lobe especially in the lower cortex. The densities

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of the DN were variable but greater densities were present in the neocortex. The GI exhibited the lowest densities of the TDP-43 immunoreactive lesions recorded and apart from the DG, exhibited no regional preference. Differences in density of various pathological features have been observed between familial and sporadic cases of FTLD-TDP. For example, GRN cases often have consistently higher densities of NCI, DN, and vacuoles in the MFG compared with the sporadic cases. Previous studies report that DN were more frequent in the frontal cortex and less frequent in the DG in GRN cases (Hatanpaa et al., 2008). In addition, cases lacking GRN mutations may have a less severe pathology affecting the neocortex and striatum while NII are usually absent or infrequent (Mackenzie et al., 2006a). Hence, FTLD-TDP caused by GRN mutations may be a distinct subtype of FTLD-TDP (Mackenzie, 2007).

Spatial Patterns of TDP-43 Immunoreactive Inclusions

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Examples of the spatial patterns of the TDP-43 immunoreactive lesions in individual brain regions in Case A are shown in Figure 3.

Figure 3. Examples of the spatial patterns exhibited by TDP-43 immunoreactive lesions (NCI = neuronal cytoplasmic inclusions, NII = neuronal internuclear inclusions, DN = dystrophic neurites) in frontotemporal lobar degeneration with TDP-43 proteinopathy (FTLD-TDP) caused by progranulin (GRN) gene mutation.

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In the upper laminae of the frontal cortex, the NCI exhibited a V/M peak at a field size of 400μm suggesting the presence of clusters of NCI, 400μm in diameter, regularly distributed parallel to the pia mater. In the upper laminae of the PHG, the NII exhibited a V/M peak at a field size of 200μm suggesting the presence of clusters of NII regularly distributed parallel to the pia mater. By contrast, the DN in the upper laminae of the frontal cortex did not exhibit a pattern of regular clustering, the V/M ratio increasing with field size without reaching a peak suggesting the presence of large scale clustering of the DN, i.e., in which the clusters were greater than 1600μm. A summary of the spatial patterns observed in all regions and cases is shown in Table 2. In cortical areas, the commonest spatial pattern exhibited by the TDP-43 immunoreactive lesions was the presence of clusters regularly distributed parallel to the pia mater, this pattern being present in 11/19 (58%) analyses of NCI, 1/3 (33%) of GI, 11/31 (35%) of NII, and 8/18 (44%) of DN. Table 2. Frequency of different types of spatial patterns (R = random distribution, Reg = regular distribution) exhibited by the pathological inclusions (NCI = neuronal cytoplasmic inclusions, GI = oligodendroglial inclusions, NII = neuronal intranuclear inclusions (NII), DN = dystrophic neurites) in various brain regions (cerebral cortex, CA1/2 = CA sectors of hippocampus, DG = dentate gyrus) in nine cases of frontotemporal lobar dementia with TDP-43 proteinopathy (FTLD-TDP) caused by progranulin (GRN) gene mutation, N = number of brain regions analyzed Feature

Region

N

R

Reg

NCI

Cortex CA1/2 DG Cortex CA1/2 DG Cortex CA1/2 DG Cortex CA1/2 DG

19 2 6 3 1 0 31 3 3 18 0 1

5 1 1 2 0

GI

NII

DN

2 0 0 0 0

Regular Clustering 11 1 3 1 1

Large Clusters 1 0 2 0 0

12 1 1 4

6 0 0 1

11 2 2 8

2 0 0 5

0

0

1

0

Ch-square (χ2) contingency tables: Comparison of totals for cortical regions versus CA1/2 and DG; NCI (χ2 = 2.88, P > 0.05); GI (χ2 = 1.33, P > 0.05), NII (χ2 = 2.79, P > 0.05), DN (χ2= 1.17, P > 0.05).

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In areas not exhibiting regular clustering, lesions were either randomly or regularly distributed or present in significantly larger clusters. A random distribution of the NII was present in 12/31 (39%) of analyses. Although sample sizes were smaller, regularly spaced clustering of lesions was also present in sectors CA1/2 of the hippocampus and in the DG. Chi-square (χ2) contingency tests suggested there were no significant differences in the relative frequency of the different spatial patterns between cortical areas and the hippocampus. In many brain regions, the TDP-43 immunoreactive lesions occurred in clusters that exhibited a regular periodicity parallel to the tissue boundary. This type of spatial pattern is similar to that reported for NCI characterized by tau (Armstrong, 1993b; et al., 1998; 1999), α-synuclein (Armstrong et al., 2004), and α-internexin reactivity (Armstrong and Cairns, 2006). In the tauopathies and synucleinopathies, the spatial patterns of the NCI clusters within the cerebral cortex and hippocampus suggested that the inclusions were related to the cells of origin of specific cortico-cortical and corticohippocampal projections (De Lacoste and White, 1993; Hiorns et al., 1991; Delatour et al., 2004). The cells of origin of the cortico-cortical projections are clustered and occur in bands that are more or less regularly distributed along the cortical strip. Individual bands of cells, approximately 500-800μm in width, traverse the cortical laminae in columns (Hiorns et al., 1991). This pattern of cortical degeneration may also be present in cases of FTLD-TDP caused by GRN DNA mutation. In 33-58% of cortical regions, TDP-43 immunoreactive lesions were regularly distributed parallel to the pia mater consistent with their development in relation to these connections. However, the estimated width of the NCI, NII, or DN clusters approximated to the dimension of the cells of origin of the cortico-cortical projections in between 22% and 36% of cortical gyri. In the majority of cortical areas, the NCI, NII, and DN developed in smaller clusters, usually between 50 and 200μm in diameter; a size similar to the α-internexin immunoreactive NCI in NIFID (Armstrong and Cairns, 2006). Moreover, in a significant number of regions the NII were randomly distributed. Hence, in many areas TDP-43 pathology appears to affect a subset of cells associated with corticocortical pathways. In some cortical areas, however, the smaller clusters of TDP-43 lesions were aggregated into larger ‘superclusters’ and in a significant number of further regions, clusters larger than 800μm in diameter were present. This observation suggests that smaller clusters of the NCI, NII, and DN may evolve into larger clusters as the disease progresses (Armstrong, 1993b).

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Laminar Distribution of TDP-43 Immunoreactive Inclusions

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Examples of the laminar distribution of the NCI and NII in a single gyrus (Case A, MFG) are shown in Figure 4. The distribution of the NCI was fitted by a third-order polynomial (r = 0.55, P < 0.05) with a density peak in the upper laminae. By contrast, the distribution of the NII was fitted by a fourth-order polynomial (r = 0.60, P < 0.01) suggesting a bimodal distribution of the NII; the peaks of density in the upper and lower laminae having approximately equal magnitude. A summary of the laminar distributions shown by all TDP-43 immunoreactive inclusions over regions and cases is shown in Table 2. In approximately 50% of gyri studied, the density of the NCI was greatest in the upper laminae; four gyri having a unimodal and three a bimodal distribution. In a number of gyri, there was no significant difference in density with distance below the pia mater. In 7 gyri analyzed, the GI exhibited a peak of density in the upper laminae in two gyri, a peak in the lower laminae in one gyrus, and no significant

Figure 4. Examples of the laminar distribution of the neuronal cytoplasmic inclusions (NCI) and neuronal intranuclear inclusions (NII) in a single gyrus (Case A, MFG) in familial frontotemporal lobar degeneration with TDP-43 proteinopathy (FTLD-TDP) caused by progranulin (GRN) gene mutation. Curve fitting; NCI third order polynomial (r = 0.55, P < 0.05), NII fourth-order polynomial (r = 0.60, P < 0.01).

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variation in density across the cortex in four gyri. In 17 gyri analyzed, the NII exhibited a peak of density in the lower cortex in five gyri, while four gyri, a bimodal distribution was present; peaks of density being equal in the upper and lower cortex. In eight gyri, there was no significant variation in density of the NII across the cortex. The distribution of the DN was quite variable, the single most common being a peak of density in the upper cortex. In a proportion of gyri, the NII were distributed primarily in the lower laminae suggesting significant pathological change affecting the lower cortex. Consistent with this hypothesis, abnormally enlarged neurons and a significant gliosis was also present in the lower cortex. Pathological changes in the lower laminae in FTLD-TDP could be a consequence of the loss of cortical projections originating or terminating in laminae V/VI (De Lacoste and White, 1993). In many gyri, however, the data suggest even greater pathological changes affecting the upper cortical laminae. In a significant proportion of gyri, the NCI occurred either with a density peak in the upper cortex or in a bimodal distribution in which the upper density peak was greater than the lower. In addition in several gyri, vacuolation was most pronounced in the superficial cortical laminae. Hence, cortical degeneration affecting the frontal and temporal lobes occurs across all laminae in FTLD-TDP caused by GRN mutation. The greater degree of degeneration of the upper laminae suggests that it may precede that of the lower laminae; degeneration of the lower laminae occurring secondarily. Table 3. Frequency of the different types of laminar distribution of the pathological inclusions (NCI = Neuronal cytoplasmic inclusions, GI = Glial inclusions, NII = neuronal intranuclear inclusions, DN = dystrophic neurites) in gyri of the frontal and temporal cortex in nine cases frontotemporal lobar dementia with TDP-43 proteinopathy (FTLD-TDP) caused by progranulin (GRN) gene mutation (N = number of gyri analyzed. Data indicate the number of gyri in which a particular histological feature exhibited a single (unimodal) or double (bimodal) density peak in U = upper, M = middle, L = lower cortical laminae, NS = no significant change in density from pia mater to white matter)

Lesion NCI GI NII DN

Unimodal distribution N U M 15 4 0 7 2 0 17 0 0 14 5 1

L 1 1 5 2

Bimodal distribution U>L U=L L>U 3 2 0 0 0 0 0 4 0 2 1 0

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NS 5 4 8 3

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The data suggest a hypothesis to explain cortical degeneration in FTLD-TDP caused by GRN mutation. First, pathological changes initially affect the upper laminae of the frontal and temporal lobe resulting in the development of NCI and vacuolation. Second, loss of ascending projections leads to the abnormal enlargement of neuronal perikarya, neuronal loss, and gliosis primarily affecting the deeper cortical laminae V and VI accompanied by the appearance of NII and GI. Third, the pathology may spread between cortical areas via the feedforward and feedback cortco-cortical pathways.

Conclusion

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Many cases of familial FTLD-TDP are caused by DNA mutations of the GRN gene. These cases are characterized by greater densities of NCI and DN than in sporadic cases of FTLD-TDP. In addition, cortical degeneration occurs across the cortical laminae including a specific degeneration of the cortico-cortical and cortico-hippocampal pathways. The TDP-43 immunoreactive inclusions are often distributed in different cortical laminae.

Acknowledgments We thank clinical, genetic, pathology, and technical staff for making information and tissue samples available for this study and we thank the families of patients whose generosity made this research possible.

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Chapter IV

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Effect of a Specific Mammalian DNA Polymerase α-Inhibitor, Dehydroaltenusin, on DNA Replication in Cultured Cells Takeshi Mizuno1, Isoko Kuriyama2,Masaharu Takemura3, Kengo Sakaguchi4,5, Fumio Sugawara4,5, Hiromi Yoshida2,5 and Yoshiyuki Mizushina2,5,∗ Cellular Dynamics Laboratory, Advanced Science Institute, RIKEN, Wako, Saitama, Japan 2 Laboratory of Food & Nutritional Sciences, Department of Nutritional Science, Kobe-Gakuin University, Nishi-ku, Kobe, Hyogo, Japan 3 Graduate School of Mathematics and Science Education, Tokyo University of Science, Shinjuku-ku, Tokyo, Japan 4 Department of Applied Biological Science, Tokyo University of Science, Noda, Chiba, Japan 5 Cooperative Research Center of Life Sciences, Kobe-Gakuin University, Chuo-ku, Kobe, Japan 1

∗

Corresponding Author: Yoshiyuki Mizushina, Ph.D., Laboratory of Food and Nutritional Sciences, Department of Nutritional Science, Kobe-Gakuin University, 518 Arise, Ikawadani-cho, Nishi-ku, Kobe, Hyogo 651-2180, Japan; Tel.: +81-78-974-1551 (ext.3232); fax: +81-78-974-5689; E-mail address: [email protected] (Y. Mizushina).

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Abstract In a screening for selective inhibitors of eukaryotic DNA polymerase (pol) species, dehydroaltenusin extracted from a fungus (Alternaria tennuis) was found to be an inhibitor of DNA replicative pol α. This compound inhibited only mammalian pol α, and did not influence the activity of other DNA replicative pols such as pols δ and ε, but also showed no effect even on pol α activity from other vertebrate, fish, or plant species. Dehydroaltenusin also had no effect on other DNA metabolic enzymes tested. The inhibitory effect of dehydroaltenusin on mammalian pol α was dose-dependent with an IC50 value of 0.5 μM. This effect was 10-fold stronger than that of aphidicolin, a well-known potent inhibitor of eukaryotic DNA replicative pols α, δ and ε. The inhibitory mode of dehydroaltenusin for mammalian pol α activity was competitive with the DNA template-primer and noncompetitive with the nucleotide substrate. This compound inhibited the proliferation of a human cervix carcinoma cell line, HeLa, with an LD50 value of 38.0 μM, by arresting cells at S-phase, and preventing the incorporation of thymidine into the cells. Dehydroaltenusin increased cyclin E and cyclin A levels, and induced cell apoptosis. Selective inhibitors of DNA replicative pol α, such as dehydroaltenusin, might provide novel markers for the development of anti-proliferative drugs. NIH3T3 cells that took up dehydroaltenusin by hypotonic shift, that is, the transient exposure of cultured cells in hypotonic buffer to small molecules that cannot penetrate cells, also showed inhibition of cell growth. At a low concentration (10 μM) of dehydroaltenusin, DNA replication was inhibited and several large foci of replication protein A (RPA) were found. Furthermore, when dehydroaltenusin was incubated with aphidicolin, RPA foci were not observed in cells. In summary, these findings suggest that dehydroaltenusin blocks DNA replication through pol α inhibition, and generates singlestranded DNA, resulted in uncoupling of leading strand and lagging strand synthesis. Dehydroaltenusin could be useful as a “molecule probe” for pol α research in DNA replication. The role of pol α function in DNA replication is discussed.

Keywords: dehydroaltenusin, DNA polymerase (DNA-directed DNA polymerase (E.C. 2.7.7.7), pol) α, DNA replication, enzyme inhibitor, molecule probe, anti-cancer, DNA replication fork uncoupling

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Abbreviations pol TdT dTTP dNTP IC50 LD50 PCNA, RPA

DNA-directed DNA polymerase (EC 2.7.7.7) terminal deoxynucleotidyl transferase 2'-deoxythymidine 5'-triphosphate 2'-deoxyribonucleotside 5'-triphosphate 50 % Inhibitory concentration 50 % Lethal dose, proliferating cell nuclear antigen replication protein A

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1. Introduction DNA replication in eukaryotes is a key system to maintain cellular renewal [1], and DNA polymerases (pol, i.e., DNA-dependent DNA polymerase, E.C. 2.7.7.7) have important roles in DNA replication. Pol catalyzes the polymerization of deoxyribonucleotides alongside a DNA strand, which they "read" and use as a template [2]. The newly polymerized molecule is complementary to the template strand and identical to the template’s partner strand. Pol can add free nucleotides only to the 3’ end of the newly formed strand, meaning that elongation of the new strand occurs in a 5’ to 3’ direction. The human genome encodes at least 15 pols to conduct cellular DNA synthesis [3, 4]. Eukaryotic cells contain three DNA replicative pols (α, δ and ε), mitochondrial pol γ, and at least 12 non-replicative pols [β, ζ, η, θ, ι, κ, λ, μ, ν, terminal deoxynucleotidyl transferase (TdT) and REV1] [3-5]. Pols have a highly conserved structure, which means that their overall catalytic subunits vary, on the whole, very little among species. Conserved structures usually indicate important, irreplaceable functions of the cell, the maintenance of which provides evolutionary advantages. On the basis of sequence homology, eukaryotic pols can be divided into four main different families: A, B, X, and Y [6]. Family A includes mitochondrial pol γ, and pols θ and ν; and family B includes three DNA replicative pols (α, δ, and ε) and pol ζ. Family X contains pols β, λ, μ, and TdT; and family Y includes pols η, ι, and κ, and REV1. Because not all functions of eukaryotic pols have been fully elucidated, selective inhibitors of each pol are useful tools and molecular probes to distinguish pols and to clarify their biological functions [7]. For example, aphidicolin is a selective inhibitor of eukaryotic pols

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α, δ and ε, which are essential for DNA replication [8], and this inhibitor has been very useful for studying the DNA replication system [1]. Previously, however, there have been no reports of inhibitors capable of distinguishing among pols α, δ and ε. We therefore established an assay to detect pol inhibitors, and have been screening natural sources for inhibitors for 16 years; as a result, we found an interesting inhibitor that influenced only the activity of mammalian pol α. The agent, which was determined to be dehydroaltenusin [9], has been reported to be an inhibitor of myosin light chain kinase [10, 11]. In this review, the role of pol α function in DNA replication was revealed by using dehydroaltenusin, as compared with aphidicolin, as a “molecular probe” for pol α research.

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2. Production and Isolation of Dehydroaltenusin An assay method to detect pol inhibitors was established [12, 13] in which poly(dA)/oligo(dT)12-18 (A/T = 2/1) and 3H-labeled 2'-deoxythymidine 5'triphosphate (dTTP) were used as the DNA template-primer and nucleotide substrate, respectively. Purified mammalian pols α, β, γ, δ, ε, η, ι, κ, λ and TdT, which have high activity, were kind gifts from pol researchers around the world. We screened for pol inhibitors, and found a natural compound that inhibits mammalian pol α activity but not pol β activity present in a fungus (Alternaria tennuis) collected from fields in the vicinity of Noda city in Chiba prefecture, Japan [9]. Compounds were extracted from the mycelium of the fungus with CH2Cl2, and then purified by silica gel column and Sephadex LH-20 column chromatography. EI (Electron Impact) mass, negative FABHR (Fast Atom Bombardment High Resolution) mass, and 1H-, 13C- and DEPT (Distortionless Enhancement by Polarization Transfer) NMR spectroscopic analyses suggested that the inhibitor fraction was dehydroaltenusin. The chemical structure of dehydroaltenusin is shown in Figure 1A. We established a method for total chemical synthesis of dehydroaltenusin, and succeeded in completely synthesizing the compound [14-16]. The synthetic dehydroaltenusin had the same properties of pol α inhibition as the natural compound [14].

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3. Effect of Dehydroaltenusin on Pols and other DNA Metabolic Enzymes

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Dehydroaltenusin dose-dependently inhibited mammalian (i.e., calf and mouse) pol α, with 50% inhibition observed at a dose of 0.68 and 0.50 μM, respectively (Table 1), and almost complete inhibition at 4 μM. Aphidicolin (Figure 1B), a potent inhibitor of mammalian pol α, shows complete inhibition of mammalian pol α at 40 μM [17]; thus, the effect of dehydroaltenusin on this enzyme was almost 10-fold stronger than that of aphidicolin.

Figure 1. Chemical structure of dehydroaltenusin and aphidicolin. (A) Dehydroaltenusin, (B) Aphidicolin.

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Enzyme --- Mammalian DNA polymerases ---

IC50 value (μM) Dehydroaltenusin

Aphidicolin

Calf DNA polymerase α

0.68 ± 0.05

20.1 ± 0.8

Mouse the largest subunit of DNA polymerase α

0.50 ± 0.05

17.8 ± 0.8

Rat DNA polymerase β

64.0 ± 1.5

>100

Human DNA polymerase γ

>100

>100

Human DNA polymerase δ

>100

12.7 ± 0.5

Human DNA polymerase ε

>100

16.1 ± 0.6

Human DNA polymerase η

>100

>100

Mouse DNA polymerase ι

>100

>100

Human DNA polymerase κ

>100

>100

Human DNA polymerase λ Calf terminal deoxynucleotidyl transferase (TdT) --- Fish DNA polymerases ---

>100 >100

>100 >100

Cherry salmon DNA polymerase α

>100

28.3 ± 1.0

Cherry salmon DNA polymerase δ --- Plant DNA polymerases ---

>100

24.2 ± 0.9

Cauliflower DNA polymerase α --- Prokaryotic DNA polymerases --E. coli DNA polymerase I (Klenow fragment) Taq DNA polymerase T4 DNA polymerase --- Other DNA metabolic enzymes ---

>100

32 ± 1.2

>100 >100 >100

>100 >100 >100

Calf primase of DNA polymerase α Mouse primase (the smallest subunit of DNA polymerase α)

>100

>100

>100

>100

Human telomerase

88.0 ± 1.8

>100

HIV-1 reverse transcriptase

90.9 ± 2.0

>100

T7 RNA polymerase T4 polynucleotide kinase Bovine deoxyribonuclease I

89.4 ± 2.0

>100 >100 >100

>100 >100

One unit of pol activity is defined as the amount that catalyzes the incorporation of 1 nmol of deoxyribonucleotside triphosphate (i.e., dTTP) into synthetic DNA template-primer (i.e., poly(dA)/oligo(dT)12-18, A/T = 2/1) at 37 ºC in 60 min. Dehydroaltenusin or aphidicolin was incubated with each enzyme (0.05 units). Enzyme activity in the absence of the compounds was taken as 100 %. Data are shown as the means ± SE of three independent experiments.

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The inhibitory effect of dehydroaltenusin on calf pol α and the core domain (p110) of the largest subunit of mouse pol α was more than 100-fold stronger than that on rat pol β, and approximately 130-fold stronger than that on human telomerase, human immunodeficiency virus type-1 (HIV-1) reverse transcriptase and T7 RNA polymerase (Table 1). This compound had no inhibitory effect either on other mammalian pols such as A-family pols (human pol γ), B-family pols (human pols δ and ε), X-family pols (human pol λ and calf TdT) and Y-family pols (human pols η, κ and mouse pol κ), or on pol α or δ from a fish (cherry salmon); pol α from a higher plant (cauliflower); prokaryotic pols such as the Klenow fragment of E. coli pol I, Taq pol and T4 pol; or other DNA metabolic enzymes such as calf and mouse primase of pol α, T4 polynucleotide kinase and bovine deoxyribonuclease I. The IC50 values in Table 1 did not change when activated DNA (i.e., double-stranded DNA digested by deoxyribonuclease I) and four 2'-deoxyribonucleoside 5'-triphosphate (dNTPs) were used as the DNA template-primer and nucleotide substrate, respectively, instead of poly(dA)/oligo(dT)12-18 and dTTP. We would, however, like to emphasize here that dehydroaltenusin first intercalates into the DNA molecule as a substrate (i.e., DNA template-primer), and subsequently inhibits both activities indirectly through the induction of a conformational change in the DNA. This compound effected no thermal transition in melting temperature; thus, none of the dehydroaltenusin bound to the doublestranded DNA, suggesting that dehydroaltenusin must inhibit enzyme activity by interacting with pol α directly. We then investigated whether an excessive amount of nucleic acid [i.e., poly(rC)] or protein [i.e., bovine serum albumin (BSA)] would prevent the inhibitory effect of dehydroaltenusin to determine whether this effect results from non-specific adhesion to pol α or selective binding to specific sites on the protein. Poly(rC) and BSA had little or no influence on inhibition by dehydroaltenusin, suggesting that binding to pol α occurs selectively. These results suggested that dehydroaltenusin might be a selective inhibitor of mammalian pol α.

4. Mode of Pol α Inhibition by Dehydroaltenusin Next, to elucidate the mechanism of inhibition, the extent of inhibition as a function of the DNA template-primer or nucleotide substrate concentration was

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studied (Table 2). In this kinetic analysis, poly(dA)/oligo(dT)12-18 (A/T = 2/1) and dTTP were used as the DNA template-primer and nucleotide substrate, respectively. Double reciprocal plots of the results showed that the dehydroaltenusin-induced inhibition of calf pol α activity was competitive with the DNA template-primer and non-competitive with the nucleotide substrate. With respect to the DNA template-primer, the apparent maximum velocity (Vmax) was unchanged at 55.6 pmol/h, whereas a 140%, 220% and 768% increase in the Michaelis constant (Km) was observed in the presence of 0.25, 0.5 and 1 μM dehydroaltenusin, respectively. The Km for the nucleotide substrate was unchanged at 1.65 μM, whereas the Vmax for the nucleotide substrate decreased from 29.2 pmol/h with no dehydroaltenusin to 5.36 pmol/h in the presence of 1 μM dehydroaltenusin. The inhibition constant (Ki) value, obtained from Dixon plots, was found to be 0.23 μM and 0.18 μM for the DNA template-primer and nucleotide substrate, respectively. When activated DNA and four dNTPs were used as the DNA template-primer and nucleotide substrate, respectively, the inhibition of calf pol α by dehydroaltenusin was, again, competitive with the DNA template-primer and non-competitive with the nucleotide substrate. On the other hand, the inhibition of pol α by aphidicolin is uncompetitive with activated DNA as the DNA template-primer and competitive with the nucleotide substrate [17]. Moreover, aphidicolin inhibits pol α by competing with 2'-deoxycytidine 5'-triphosphate (dCTP) but not by competing with the other three dNTPs [9]. By contrast, the inhibition of pol α by dehydroaltenusin was noncompetitive with the four dNTPs. Table 2. Kinetic analysis of the inhibitory effects of dehydroaltenusin on pol α as a function of DNA template-primer dose and nucleotide substrate concentration Substrate

DNA template-primer a)

Nucleotide substrate a) b)

b)

Dehydroaltenusin (μM) 0 0.25 0.5 1 0 0.25 0.5 1

Km (μM) 7.8 10.9 17.2 59.9

1.65

Vmax (pmol/h)

Ki (μM)

Inhibitory mode

55.6

0.23

Competitive

29.2 14.9 10.2 5.36

0.18

Non-competitive

Poly(dA)/oligo(dT)12-18. dTTP.

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Thus, the mode of the inhibitory effect of dehydroaltenusin on pol α differs from that of aphidicolin. These results suggest that dehydroaltenusin directly binds to the DNA template-primer-binding site of pol α, but may bind to or interact with a site distinct from the nucleotide-substrate-binding site. Mammalian pol α is made up of four subunits, termed p180, p68, p54 and p46 [18-20]. The largest subunit, p180, and the smallest subunit, p46, have catalytic (DNA polymerization) and primase activity, respectively [21, 22]. The other subunits, p68 and p54, have no known enzyme activity. Both the DNAbinding site and the nucleotide substrate-binding site of pol α are located in the largest subunit, p180 [21, 22]. We further studied the interaction between dehydroaltenusin and the p180 largest subunit of pol α. From the results of both a gel mobility shift assay and surface plasmon resonance (SPR) analysis using Biacore 3000, dehydroaltenusin was suggested to interact directly with the p180 catalytic subunit of pol α, and not to interact with the other three subunits [9].

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5. Effect of Dehydroaltenusin on Human Cancer Cell Growth To determine the effects of dehydroaltenusin on cultured cancer cells, we tested its influence on four human cancer cell lines, A549 (lung cancer cell lines), BALL-1 (acute lymphoblastoid leukemia cells), HeLa (cervical carcinoma cells) and NUGC-3 (stomach cancer cells) [23]. Dehydroaltenusin efficiently inhibited all cell growth tested in a dose-dependent manner, and the LD50 value varied from 38.0 to 44.4 μM after 24 h of incubation (Table 3). Table 3. Inhibitory effect of dehydroaltenusin on the proliferation of human cancer cells Cell line A549 BALL-1 HeLa NUGC-3

Cell type Human lung cancer cell Human acute lymphoblastoid leukemia cell Human cervix cancer cell Human stomach cancer cell

IC50 value (μM) 44.4 ± 2.2 43.2 ± 2.0 38.0 ± 1.5 40.1 ± 1.7

Human cancer cells were incubated with dehydroaltenusin for 24 h. The cell viability was determined by MTT assay. Data are shown as the means ± SE of five independent experiments.

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HeLa, NUGC-3 and A-549 cells are adherent cell lines; however, the nonadherent cell line BALL-1 was suppressed with almost the same cell growth inhibitory results as the adherent cell lines. These results suggest that dehydroaltenusin may have a generalized observed effect on “human cancer cells”. The inhibitory effect on HeLa cells was strongest among the human cancer cell lines tested; therefore, we concentrated on the properties of HeLa cells in our subsequent studies.

6. Inhibitory Properties of ehydroaltenusin on Human Cancer Cell Growth

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To clarify the effect of dehydroaltenusin on cell-cycle regulation, we analyzed the cell cycle of HeLa using flow cytometry with DAPI staining [24]. When the HeLa cells were treated with dehydroaltenusin at 38.0 μM (= LD50 value), the distribution of cells in the cell cycle changed time dependently (Table 4). In the presence of dehydroaltenusin for 12 h, the percentage of cells in S-phase increased (from 39.1% to 50.1%) and the percentage of cells in G2/M-phase decreased (from 30.6% to 17.3%). The amount of G1-phase cells did not change during the incubation (from 30.2% to 32.6%). Table 4. Effect of dehydroaltenusin on the cell cycle

Incubation time (h)

Phase of the cell cycle (%) G1

S

G2/M

0

30.3 ± 1.5

39.1 ± 1.9

30.6 ± 1.6

6

27.3 ± 0.8

45.8 ± 2.5

26.9 ± 0.9

12

32.6 ± 1.7

50.1 ± 3.0

17.3 ± 0.5

Flow cytometric analysis of HeLa cells treated with 38.0 μM (= LD50 value) dehydroaltenusin. The DNA content of 8000 stained cells was analyzed by using a cell counter analyzer (Partec, CCA Model; Munster, Germany) with Multicycle 3.11 software (Phoenix Flow Systems, San Diego, CA). The cell cycle distribution was calculated as the percentage of cells that were in G1, S and G2/M-phases. Data are shown as the means ± SE of four independent experiments.

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We assessed whether the effect of dehydroaltenusin on the cell cycle was associated with the expression of cyclin proteins by Western blotting. As shown in Figure 2A, the amount of cyclin A and cyclin E, which are regulated in G1/Sphase [25-28], increased with dehydroaltenusin treatment, but the level of cyclin B, which is regulated in G2/M-phase [29], decreased significantly.

Figure 2. Effect of dehydroaltenusin on protein expression in HeLa cells. (A) Cyclin expression was analyzed by Western blotting. Cell extracts of the nuclear fraction were prepared from cells treated with 38.0 μM (= LD50 value) dehydroaltenusin. Cyclins E, A and B were detected with specific antibodies. (B) Effect of dehydroaltenusin on binding of the pol complex to chromatin. Shown is the Western blot analysis of pol α, pol ε and PCNA in the chromatin-bound fraction. Densitometric assay of the proteins was performed and the fold induction was calculated.

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Figure 3. Measurement of DNA, RNA and protein synthesis in HeLa cells incubated with dehydroaltenusin. 1 x 104 HeLa cells were incubated without (control, open square) or with 38.0 μM (= LD50 value) dehydroaltenusin (closed circle) starting from 0 h; and at 0.5 h, as probes for DNA, RNA and protein synthesis, [methyl-3H] thymidine, [5,6-3H] uridine and L-[4,5-3H] leucine (final 3 μCi, 4 μCi, 4 μCi, respectively) were added. These three metabolites were measured simultaneously. Panels A, B and C show the incorporation of thymidine, uridine and leucine, respectively. Data are shown as the means ± SE of five independent experiments.

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These results suggest that dehydroaltenusin induced S-phase arrest in human cancer cells. These results were more directly confirmed by an incorporation experiment (see below). Because dehydroaltenusin inhibited mammalian pol α activity in vitro [9], we hypothesized that the growth suppression caused by dehydroaltenusin was a result of its inhibition of pol α. To test this possibility, the amount of pol α, pol ε and proliferating cell nuclear antigen (PCNA) was determined by Western blots. After treatment with 38.0 μM of dehydroaltenusin at various times, protein expression levels were assessed (Figure 2B). The protein expression patterns were consistent with the cell cycle analysis data. The increase in the percentage of cells in S-phase was accompanied by accumulation of PCNA in the chromatin fraction; however, the amount of pol α and pol ε did not change during incubation with dehydroaltenusin. The effects of dehydroaltenusin on DNA, RNA and protein synthesis were examined further via an incorporation experiment. Figure 3A, 3B and 3C show the incorporation of [3H]-labeled thymidine, [3H]-uridine and [3H]-leucine into HeLa cells, respectively. Dehydroaltenusin inhibited only the incorporation of [3H]-thymidine into the cells. The incorporation of [3H]-thymidine was decreased to 45% of the control value after 6 h of incubation in the presence of 38.0 μM dehydroaltenusin (Figure 3A). Neither [3H]-uridine nor [3H]-leucine incorporation was affected by dehydroaltenusin (Figure 3B and 3C). These observations indicate that dehydroaltenusin must inhibit cell growth by blocking the S-phase of DNA replication, in other words, by inhibiting pol α activity in vivo. To examine whether the decrease in cell numbers caused by dehydroaltenusin (Table 3) was due to apoptosis, DNA fragmentation was analyzed by electrophoresis [24]. The formation of DNA ladders was dose-dependent in HeLa cells treated with 0 to 100 μM dehydroaltenusin for 24 h. DNA ladders appeared after treatment with 75 μM dehydroaltenusin, as shown in Figure 4. Therefore, both the inhibition of in vivo DNA synthesis and the apoptotic effect occurred in cells incubated with dehydroaltenusin at concentrations of more than 75 μM. DNA ladders were not evident during the initial 6 h but were apparent at 12 h and thereafter. This observation indicates that the inhibition of pol α activity by dehydroaltenusin has a strong apoptotic effect on human cancer cells. The effect of dehydroaltenusin must be due to a combination of growth arrest and cell death. This compound has also been shown to be a promising suppressor of solid tumor growth in an in vivo anti-tumor assay on nude mice bearing solid tumors of HeLa cells [30]; therefore, the block of DNA replication caused by pol α inhibition may lead to an anti-cancer effect.

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Figure 4. DNA fragmentation of dehydroaltenusin-treated HeLa cells on agarose gel electrophoresis. Lanes 1–5, HeLa cells (2 x 106) were incubated for 24 h with 0, 25, 50, 75 and 100 μM dehydroaltenusin, respectively. Total DNA was then extracted and analyzed by 1.4% agarose gel electrophoresis. DNA bands were stained with ethidium bromide, and visualized under UV light.

7. Effect of Dehydroaltenusin on DNA Replication in NIH3T3 Cells To monitor the effect of dehydroaltenusin on DNA replication in the mammalian cell nucleus, we next observed replication foci by immunofluorescent techniques. When we previously characterized mouse DNA replication proteins in a mouse embryonic fibroblast cell line, NIH 3T3, we found that several replication factors stained as foci in NIH3T3 cells after pre-extraction with CSK buffer containing 0.5% Triton X-100 detergent on ice 5 min [31]. These DNA replication factors include PCNA, pol α, DNA ligase I, and replication protein A (RPA). When human cancer cells were investigated with dehydroaltenusin, a large amount of dehydroaltenusin in the medium (LD50 = 38 µM) was essential to inhibit cell growth (Table 3). In the in vitro assay, however, a 50-fold smaller amount (IC50 = 0.68 µM) of dehydroaltenusin was enough to inhibit pol α activity (Table 1). We therefore examined another approach to incorporate dehydroaltenusin into cells.

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Figure 5. Effect of inhibitors on DNA replication in the nucleus of mouse NIH3T3 cells. (A) Immunofluorescent localization of chromatin-bound RPA. NIH3T3 cells were treated by hypotonic shift with 10 µM aphidicolin or 10 µM dehydroaltenusin. After 60 min, cells were pre-extracted with cytoskelton buffer containing 0.5% Triton X-100 for 5 min on ice and fixed with 3.7% formaldehyde. PCNA and the RPA-p34 subunit were detected by indirect immunofluorescence with anti-PCNA rabbit polyclonal antibody and anti-RPA mouse monoclonal antibody, respectively. The secondary antibodies for PCNA and the RPA-p34 subunit were Alexa488-conjugated anti-rabbit antibody (green color) and Alexa 594-conjugated anti-mouse antibody (red color), respectively. DNA was stained with Hoecst33258 (blue color). The white arrows indicate nuclei in which RPA foci were localized by dehydroaltenusin treatment. (B) Quantification of the percentage of PCNA and RPA in the nuclei of mouse NIH3T3 cells. PCNA and RPA were double-stained by anti-PCNA rabbit polyclonal antibody and anti-RPA mouse monoclonal antibody, respectively, and then the number of stained nuclei was counted.

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We considered whether hypotonic shift would be suitable for these experiments, and therefore we optimized hypotonic shift for NIH3T3 cells using fluorescein isothiocianate (FITC)-labeled 2'-deoxyuridine 5'-triphosphate (dUTP); this nucleotide was incorporated into the nucleus by hypotonic shift and could be observed as DNA replication foci [32, 33]. This result showed that we can incorporate chemicals efficiently into NIH3T3 cells by hypotonic shift, and t DNA replication was inhibited at a low concentration of dehydroaltenusin (20 µM). Unexpectedly, large RPA foci were observed in the cells treated with dehydroaltenusin, whereas aphidicolin incorporation did not result in any large RPA foci (Figure 5). We confirmed that the large RPA foci were dependent on the penetration of dehydroaltenusin. Interestingly, in these nuclei, the RPA foci did not co-localize with PCNA (Arrows in Figure 5A). In addition, treatment with a combination of dehydroaltenusin and aphidicolin did not result in large RPA foci (Figure 5B). We concluded that the large RPA foci were due to impairment in lagging strand synthesis as discussed below.

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8. Discussion Fifteen pols are encoded in the human genome. Among these pols, three (pols α, δ and ε) participate in chromosomal DNA replication. [34, 35]. Pol α is the only enzyme that is coupled with DNA primase, and therefore essential for the initiation of both leading strand synthesis and lagging strand synthesis. To understand the role of the different pols in chromosomal DNA replication, the inhibition of each individual pol is an important approach. However, pols α, δ and ε belong to the same family of B-type pols [6], and all of the B-type catalytic subunits have a similar amino acid motif [36]. Furthermore, because the catalytic subunits of pols α, δ and ε share the same structural domains, distinguishing among these individual pols has been difficult so far. Indeed, aphidicolin inhibits all three pols equally [9]. Hence, our discovery of dehydroaltenusin as a mammalian pol-α-specific inhibitor is an important step forward. Such novel chemicals might be useful tools for analysis of the cellular DNA replication mechanism. Interestingly, it has been reported in Saccharomyces cerevisiae that pol ε is primarily responsible for copying the leading-strand template, and pol δ is primarily responsible for copying the lagging-strand template. However, it remains elusive whether the three pols have the same functions in other eukaryotes including metazoans.

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In the cell nucleus, DNA replication is observed as discrete foci. Because more than 10 molecules of pols and PCNA are estimated to be located in the DNA replication foci, these foci are also called ‘DNA replication factories’. However, the strong replicative helicase candidate proteins minichromosome maintenance 2–7 (MCM2–7) are not co-localized with DNA replicative pols and PCNA in these foci, giving rise to what is known as the ‘MCM-paradox’ [37]. However, the uncoupling of DNA replicative helicase and replicative pols in mammalian cells has not been clearly studied so far. The uncoupling of leading-strand pols and lagging-strand pols has been characterized mainly in vitro with a bacterial plasmid and pol III system [38, 39], although Byun et al. have characterized the uncoupling of MCM helicase and pol activity using a Xenopus egg extract in vitro system [40]. In all of these in vitro experiments, single stranded-DNA was exposed. By contrast, in vivo experiments have not been carried out, particularly in mammalian cells. At least to our knowledge, the perturbation of lagging strand synthesis has not been investigated in mammalian cells. The aberrant RPA foci observed here is an unique novel phenotype in lagging strand inhibition. We speculate that the foci represent the uncoupling of leading strand synthesis and lagging strand synthesis caused by pol α inhibition due to dehydroaltenusin treatment (Figure 6A). Moreover, because some RPA foci did not co-localize with PCNA, formation of the complex of DNA replicative factors may collapse, resulting in inhibition of DNA elongation pols. By contrast, we can exclude the possibility that the aberrant RPA foci derived from an uncoupling of MCM helicase and DNA replicative pols caused by the inhibition of DNA replicative pols α, δ and ε using aphidicolin treatment (Figure 6B). Due to the impairment in lagging strand initiation, the collapse of elongation pols (only pol δ, and not pol ε) must be an interesting possibility. In eukaryotic cells, the fork protection complex stabilizes MCM helicase and these pols. Recently, highconcentration aphidicolin treatment was shown to cause disassembly of the elongation factor and large RPA foci in nucleus [41]. The relationship between the aberrant foci and the fork protection complex is unclear. A lack of specific inhibitors of MCM helicase and DNA replicative pol inhibitors has hampered the analysis of replication factories in the nucleus. Although an MCM helicasespecific inhibitor, helicanomysin, has been reported [42], unfortunately helicanomysin is difficult to use in immunofluorescence analysis owing to its strong self-fluorescence (unpublished result by T.M). The optimization of hypotonic shift to NIH3T3 cells can allow the introduction of bulky and hydrophobic chemicals into cells. Several techniques, such as microinjection, electroporation, stamporation [43], lipid-mediated transfection, pinocytosis, hypotonic shift, beads loading, and stretching cells [32],

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have also been developed for the introduction of small molecules into cells. Here, to introduce dehydroaltenusin into cells, we tried to optimize the uptake of small molecules into mouse NIH3T3 cells by hypotonic shift. NIH3T3 cells that took up dehydroaltenusin by hypotonic shift showed inhibition of cell growth, thereby indicating that hypotonic shift is a suitable method for incorporating small molecules into cells.

Figure 6. Models of DNA replication fork arrest due to inhibition of pol α. (A) Uncoupling of the leading and lagging strands by dehydroaltenusin treatment. (B) Uncoupling of DNA replicative pols and MCM helicase by aphidicolin treatment.

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Chemical knock down may be the best method for detection of cell viability. Genomic knock down is theoretically the best method; however, essential gene product is difficult to manipulate this target gene. Conditional knock downs of pol α have been developed. Temperature-sensitive mutants have been collected by Nurse et al. [44]. For cell cycle analysis, pols also identified their cdc collection. For example, in yeast, pols α and δ are cdc17 and cdc2, respectively. In mammalian cells, only pol α has a temperature-sensitive mutant, tsFT20, because pol α gene is localized on the X-chromosome. Genes for other pols are encoded on the somatic chromosome. Thus, in diploid cells, two alleles hamper the establishment of temperature-sensitive cell lines. Using this specific inhibitor, the effect of inhibition of pol α on DNA replication was monitored. The in situ detection of inhibition of the DNA replication fork will shed light on replication factories and whether pols δ and ε are lagging pol and leading pol, respectively, in the mammalian cell nucleus. We can only speculate about the efficiency of the incorporation of dehydroaltenusin into cells. Experiments with small interfering RNA (siRNA), and the development pol ε or pol δ specific inhibitors would help in future analyses.

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Conclusion Dehydroaltenusin did not influence the activity of mammalian pols δ and ε, which are other DNA replicative pols, but also showed no effect on pol α of another vertebrate, cherry salmon. Dehydroaltenusin was significantly effective in suppressing tumor growth in nude mice bearing solid tumors, because this compound has inhibitory activity against pol α and cytotoxicity against human cancer cells. Dehydroaltenusin is a type of antibiotic produced by a fungus and is chemically stable under in vivo conditions, indicating that it may be useful for analyzing the DNA replication system within cells, and for clinical use. Aphidicolin, once believed to be a pol α-specific inhibitor, is now known to also inhibit the activity of pols δ and ε [8]. No pol α inhibitors with such a limited action spectrum have been reported to date, and dehydroaltenusin will be a key agent for analyzing both the in vitro and in vivo functions of pol α in more detail. Specific inhibitors of each of the mammalian DNA replicative pol species (i.e., pols α, δ and ε) would be a useful tool with which to study DNA replication in vitro. Furthermore, the in vivo studies reviewed here suggest that these compounds may be potentially useful as chemotherapeutic drugs against cancer.

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Dehydroaltenusin will be developed as not only an agent of pol α study, but also as a pharmaceutical drug based on DNA replication inhibition.

Acknowledgments This work was supported in part by a Grant-in-aid for “Academic Frontier” Project for Private Universities: matching fund subsidy from MEXT (Ministry of Education, Culture, Sports, Science and Technology of Japan), 2006–2010 (H. Y. and Y. M.). Y. M. acknowledges a Grant-in-Aid for Young Scientists (A) (No. 19680031) from MEXT.

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DePamphilis, M.L. (1996). DNA replication in eukaryotic cells. Cold Spring Harbor Laboratory Press, in U.S.A. Kornberg, A. and Baker, T.A. (1992). Eukaryotic DNA polymerase, “DNA replication” 2nd edition. Freeman, WD, and Co., New York, Chapter 6, pp. 197-225. Hubscher, U., Maga, G. and Spadari, S. (2002). Eukaryotic DNA polymerases. Annu. Rev. Biochem., 71, 133-163. Friedberg, EC., Feaver, W.J. and Gerlach, V.L. (2000). The many faces of DNA polymerases: strategies for mutagenesis and for mutational avoidance. Proc. Natl. Acad. Sci. USA, 97, 5681-5683. Takata, K., Shimizu, T., Iwai, S. and Wood, R.D. (2006). Human DNA polymerase N (POLN) is a low fidelity enzyme capable of error-free bypass of 5S-thymine glycol. J. Biol. Chem., 281, 23445-23455. Bebenek, K. and Kunkel, T.A. (2004). DNA Repair and Replication, Advances in Protein Chemistry, Elsevier, San Diego. in: W. Yang, (Ed.), vol. 69, pp. 137-165. So, A.G. and Downey, K.M. (1992). Eukaryotic DNA replication. Crit. Rev. Biochem. Mol. Biol., 27, 129-155. Ikegami, S., Taguchi, T., Ohashi, M., Oguro, M., Nagano, H. and Mano, Y. (1978). Aphidicolin prevents mitotic cell division by interfering with the activity of DNA polymerase-α. Nature, 275, 458-460. Mizushina, Y., Kamisuki, S., Mizuno, T., Takemura, M., Asahara, H., Linn, S., Yamaguchi, T., Matsukage, A., Hanaoka, F., Yoshida, S., Saneyoshi, M.,

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Sugawara, F. and Sakaguchi, K. (2000). Dehydroaltenusin, a mammalian DNA polymerase α inhibitor. J. Biol. Chem., 275, 33957-33961. Thomas, R., Rogers, D. and Williams, D.J. (1971). J. Chem. Soc. D., 8, 393. Nakanishi, S., Toki, S., Saitoh, Y., Tsukuda, E., Kawahara, K., Ando, K. and Matsuda, Y. (1995). Isolation of myosin light chain kinase inhibitors from microorganisms: dehydroaltenusin, altenusin, atrovenetinone, and cyclooctasulfur. Biosci. Biotech. Biochem., 59, 1333-1335. Mizushina, Y., Tanaka, N., Yagi, H., Kurosawa, T., Onoue, M., Seto, H., Horie, T., Aoyagi, N., Yamaoka, M., Matsukage, A., Yoshida, S. and Sakaguchi, K. (1996). Fatty acids selectively inhibit eukaryotic DNA polymerase activities in vitro. Biochim. Biophys. Acta, 1308, 256-262. Mizushina, Y., Yoshida, S., Matsukage, A. and Sakaguchi, K. (1997). The inhibitory action of fatty acids on DNA polymerase β. Biochim. Biophys. Acta, 1336, 509-521. Takahashi, S., Kamisuki, S., Mizushina, Y., Sakaguchi, K., Sugawara, F. and Nakata, T. (2003). Total synthesis of dehydroaltenusin. Tetraherdon Lett., 44, 1875-1877. Kamisuki, S., Takahashi, S., Mizushina, Y., Sakaguchi, K., Nakata, T. and Sugawara, F. (2004). Precise structural elucidation of dehydroaltenusin, a specific inhibitor of mammalian DNA polymerase α. Bioorg. Med. Chem., 12, 5355-5359. Kuramochi, K., Fukudome, K., Kuriyama, I., Takeuchi, T., Sato, Y., Kamisuki, S., Tsubaki, K., Sugawara, F., Yoshida, H. and Mizushina, Y. (2009). Synthesis and structure-activity relationships of dehydroaltenusin derivatives as selective DNA polymerase alpha inhibitors. Bioorg. Med. Chem., 17, 7227-7238. Oguro, M., Suzuki-Hori, C., Nagano, H., Mano, Y. and Ikegami, S. (1979). The mode of inhibitory action by aphidicolin on eukaryotic DNA polymerase α. Eur. J. Biochem., 97, 603-607. Takada-Takayama, R., Tada, S., Hanaoka, F. and Ui, M. (1990). Peptide mapping of the four subunits of the mouse DNA polymerase -primase complex. Biochem. Biophys. Res. Commun., 170, 589-595. Suzuki, M., Enomoto, T., Masutani, C., Hanaoka, F., Yamada, M. and Ui, M. (1989). DNA primase-DNA polymerase α assembly from mouse FM3A cells: Purification of constituting enzymes, reconstitution, and analysis of RNA priming as coupled to DNA synthesis. J. Biol. Chem., 264, 1006510071. Miyazawa, H., Izumi, M., Tada, S., Takada, R., Masutani, M., Ui, M. and Hanaoka, F. (1993). Molecular cloning of the cDNAs for the four subunits

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Takeshi Mizuno, Isoko Kuriyama, Masaharu Takemura et al. of mouse DNA polymerase α-primase complex and their gene expression during cell proliferation and the cell cycle. J. Biol. Chem., 268, 8111-8122. Copeland, W.C. and Wang, T.S. (1991). Catalytic subunit of human DNA polymerase α overproduced from baculovirus-infected insect cells. Structural and enzymological characterization. J. Biol. Chem., 266, 2273922748. Copeland, W.C. and Wang, T.S. (1993). Enzymatic characterization of the individual mammalian primase subunits reveals a biphasic mechanism for initiation of DNA replication. J. Biol. Chem., 268, 26179-26189. Murakami-Nakai, C., Maeda, N., Yonezawa, Y., Kuriyama, I., Kamisuki, S., Takahashi, S., Sugawara, F., Yoshida, H., Sakaguchi, K. and Mizushina, Y. (2004). The effects of dehydroaltenusin, a novel mammalian DNA polymerase α inhibitor, on cell proliferation and cell cycle progression. Biochim. Biophys. Acta, 1674, 193-199. Kamisuki, S., Murakami, C., Ohta, K., Yoshida, H., Sugawara, F., Sakaguchi, K. and Mizushina, Y. (2002). Actions of derivatives of dehydroaltenusin, a new mammalian DNA polymerase α-specific inhibitor. Biochem. Pharmacol., 63, 421-427. Ohtsubo, M., Theodoras, A.M., Schumacher, J., Roberts, J.M. and Pagano, M. (1995). Human cyclin E, a nuclear protein essential for the G1-to-S phase transition. Mol. Cell Biol., 15, 2612-2624. Ohtsubo, M. and Roberts, J.M. (1993). Cyclin-dependent regulation of G1 in mammalian fibroblasts. Science, 259, 1908-1912. Fang, F. and Newport, J.W. (1991). Evidence that the G1-S and G2-M transitions are controlled by different cdc2 proteins in higher eukaryotes. Cell, 66, 731-742. Pagano, M., Pepperkok, R., Verde, F., Ansorge, W. and Draetta, G. (1992). Cyclin A is required at two points in the human cell cycle. EMBO J., 11, 961-971. Nurse, P. (1990). Universal control mechanism regulating onset of M-phase. Nature, 344, 503-508. Maeda, N., Kokai, Y., Ohtani, S., Sahara, H., Kuriyama, I., Kamisuki, S., Takahashi, S., Sakaguchi, K., Sugawara, F., Yoshida, H., Sato, N. and Mizushina, Y. (2007). Anti-tumor effects of dehydroaltenusin, a specific inhibitor of mammalian DNA polymerase α. Biochem. Biophys. Res. Commun., 352, 390-396. Izumi, M., Yanagi, K., Mizuno, T., Yokoi, M., Kawasaki, Y., Moon, K. Y., Hurwitz, J., Yatagai, F. and Hanaoka, F. (2000). The human homolog of Saccharomyces cerevisiae Mcm10 interacts with replication factors and

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dissociates from nuclease-resistant nuclear structures in G(2) phase. Nucleic Acids Res., 28, 4769-4777. Koberna, K., Stanek, D., Malinsky, J., Eltsov, M., Pliss, A., Ctrnacta, V., Cermanova, S. and Raska, I. (1999). Nuclear organization studied with the help of a hypotonic shift: its use permits hydrophilic molecules to enter into living cells. Chromosoma, 108, 325-335. Kuriyama, I., Mizuno, T., Fukudome, K., Kuramochi, K., Tsubaki, K., Usui, T., Imamoto, N., Sakaguchi, K., Sugawara, F., Yoshida, H. and Mizushina, Y. (2008). Effect of dehydroaltenusin-C12 derivative, a selective DNA polymerase alpha inhibitor, on DNA replication in cultured cells. Molecules (Basel, Switzerland), 13, 2948-2961. Miyazawa, H., Tandai, M., Hanaoka, F., Yamada, M., Hori, T., Shimizu, K. and Sekiguchi, M. (1986). Identification of a DNA segment containing the human DNA polymerase α gene. Biochem. Biophys. Res. Commun., 139, 637-643. Wang, T.S. (1991). Eukaryotic DNA polymerases. Annu. Rev. Biochem., 60, 513-552. Wang, J., Sattar, A. K., Wang, C. C., Karam, J. D., Konigsberg, W. H. and Steitz, T. A. (1997). Crystal structure of a pol alpha family replication DNA polymerase from bacteriophage RB69. Cell, 89, 1087-1099. Laskey, R.A. and Madine, M.A. (2003). A rotary pumping model for helicase function of MCM proteins at a distance from replication forks. EMBO Rep., 4, 26-30. McInerney, P. and O'Donnell, M. (2004). Functional uncoupling of twin polymerases: mechanism of polymerase dissociation from a lagging-strand block. J. Biol. Chem., 279, 21543-21551. Pages, V. and Fuchs, R.P. (2003). Uncoupling of leading- and laggingstrand DNA replication during lesion bypass in vivo. Science, 300, 13001303. Byun, T.S., Pacek, M., Yee, M.C., Walter, J.C. and Cimprich, K.A. (2005). Functional uncoupling of MCM helicase and DNA polymerase activities activates the ATR-dependent checkpoint. Genes Dev., 19, 1040-1052. Gorisch, S.M., Sporbert, A., Stear, J.H., Grunewald, I., Nowak, D., Warbrick, E., Leonhardt, H. and Cardoso, M.C. (2008). Uncoupling the replication machinery: replication fork progression in the absence of processive DNA synthesis. Cell cycle, 7, 1983-1990. Ishimi, Y., Sugiyama, T., Nakaya, R., Kanamori, M., Kohno, T., Enomoto, T. and Chino, M. (2009). Effect of heliquinomycin on the activity of human minichromosome maintenance 4/6/7 helicase. FEBS J., 276, 3382-3391.

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[43] Hara, C., Tateyama, K., Akamatsu, N., Imabayashi, H., Karaki, K., Nomura, N., Okano, H. and Miyawaki, A. (2006). A practical device for pinpoint delivery of molecules into multiple neurons in culture. Brain Cell Biol., 35, 229-237. [44] Nurse, P., Masui, Y. and Hartwell, L. (1998). Understanding the cell cycle. Nat. Med., 4, 1103-1106.

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Chapter V

Gel-Based Methods Using DNABinding Zinc(II) Complexes for the Detection of DNA Mutations

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Eiji Kinoshita∗, Emiko Kinoshita-Kikuta and Tohru Koike Department of Functional Molecular Science, Graduate School of Biomedical Sciences, Hiroshima University, Hiroshima, Japan

Abstract Detection of single-nucleotide polymorphisms (SNPs) or mutations in the human genome can contribute to the establishment of genetic linkages, the diagnosis of inherited diseases, and the selection of rational remedies. Several procedures have been developed to facilitate high-throughput detection of SNPs and mutations, but these usually require expensive apparatus and the services of skillful analysts, making it difficult for most ∗

Corresponding Author: the Department of Functional Molecular Science, Graduate School of Biomedical Sciences, Hiroshima University, Kasumi 1-2-3, Minami-ku, Hiroshima 734-8553, Japan; Tel: +81 82 257 5281; Fax: +81 82 257 5336; E-mail: [email protected].

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Eiji Kinoshita, Emiko Kinoshita-Kikuta and Tohru Koike clinical researchers and physicians to obtain useful data on SNPs and mutations. The establishment of a reliable and cost-effective detection method that uses conventional laboratory equipment is therefore desirable, particularly in medical applications. Here, we introduce two methods based on polyacrylamide gel electrophoresis (PAGE) of DNA-binding zinc(II) complexes that are suitable for detecting mutations. The first method, known as Zn2+–cyclen/PAGE, is based on a difference in the mobility in PAGE of mutant DNA compared with that of nonmutated DNA of the same chain length. This difference in mobility results from binding of the Zn2+–cyclen complex (cyclen = 1,4,7,10-tetraazacyclododecane) to thymine bases, which results in a decrease in the total charge and a local conformational change in the target DNA. The Zn2+–cyclen/PAGE method, when combined with polymerase chain reaction (PCR)-based heteroduplexing, permits visualization of heteroduplex bands on PAGE gels and allows screening for SNPs and mutations. The second method is known as Zn2+–Phos-tag/PAGE. This is based on a difference in the mobility of a phosphorylated DNA fragment compared with that of its nonphosphorylated analogue containing an identical numbers of base pairs when they are subjected to PAGE on a phosphate-affinity gel containing an immobilized polyacrylamide-bound dizinc(II) complex phosphate-binding tag molecule, Zn2+–Phos-tag {Phos-tag = 1,3-bis[bis(pyridin-2-ylmethyl)amino]propan-2-olate}. The Zn2+–Phostag/PAGE method, when combined with an allele-specific PCR method using a 1:1 mixture of 5'-phosphate-labeled and nonlabeled allele-specific primers, permits the separation of the less mobile 5'-phosphate-labeled PCR product from its more-mobile nonlabeled counterpart, and permits the determination of a genotype as heterozygotic or homozygotic. Here, we review some applications of gel-based methods based on zinc(II) complexes, and we compare their resolving power in separating mutant DNA with that of conventional gel-based electrophoresis techniques.

Introduction Since the completion of the Human Genome Project, the analysis of singlenucleotide polymorphisms (SNPs) has attracted considerable interest in relation to post-genome studies [1]. SNPs are defined as variations in DNA sequences that occur in at least 1% of the population. Most SNPs have no effect on biological or physiological functions, but some SNPs (so-called mutations) have been reported to be intimately related to diseases or to influence cellular responses to drugs. Although more than 99% of the sequence of human DNA is common to all populations, some SNPs/mutations can have a major impact on how humans respond to diseases, to environmental insults such as bacteria, viruses, toxicants,

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or chemicals, or to drugs or other therapies. In this sense, information on SNPs/mutations has considerable value in biomedical research on pharmaceutical products and in medical diagnostics [2]. Many methods are currently available for the detection of SNPs/mutations, and the choice of a particular method will depend on the scale of the assay [3–9]. We have developed two kinds of small-scale genetic-assay methods for the detection of SNPs that are based on polyacrylamide gel electrophoresis (PAGE): Zn2+–cyclen/PAGE [10] and Zn2+–Phos-tag/PAGE [11]. The Zn2+–cyclen/PAGE methodology is based on the properties of Zn2+–cyclen complex (cyclen = 1,4,7,10-tetraazacyclododecane). This complex binds selectively and reversibly to the imide group-containing nucleotide base thymine (Thy) in double-stranded DNA. This, in turn, induces dissociation of hydrogen bonds between Thy and adenine (Ade) bases, causing a local change in the conformation of the DNA. We have applied this Thy-recognizing property of Zn2+–cyclen in a method for separation of various DNA fragments by gel-based electrophoresis; this method is known as Zn2+–cyclen/PAGE. If this procedure is combined with a polymerase chain reaction (PCR)-based heteroduplexing method, it can be used to detect SNPs/mutations (genotyping and genomapping). The combined technique permits the visualization of heteroduplex bands that arise on a PAGE gel as a result of annealing during PCR of complementary strands from mutant and wild-type alleles. Furthermore, the technique does not require the use of radioactive labels or expensive fluorophore-labeled oligonucleotide probes. The appearance of slow less-mobile bands on the gel indicates the presence of a heteroduplex that, in turn, suggests the existence of a mutation or polymorphism. Furthermore, Zn2+– cyclen/PAGE can be used to separate a homoduplex of a mutant allele from a homoduplex of the corresponding homologous wild-type allele. In such cases, up to four distinct migration bands may be detected for each sample of DNA. The Zn2+–Phos-tag/PAGE method, on the other hand, relies on a difference in electrophoretic mobility between 5'-terminal phosphorylated and nonphosphorylated DNA fragments of equal length on the phosphate-affinityPAGE gel. The phosphate-affinity site consists of an immobilized phosphatebinding tag molecule, the polyacrylamide-bound dizinc(II) complex Zn2+–Phostag {Phos-tag = 1,3-bis[bis(pyridin-2-ylmethyl)amino]propan-2-olate}. When the Zn2+–Phos-tag/PAGE procedure is combined with an allele-specific PCR method using a 1:1 mixture of a 5'-phosphate-labeled allele-specific primer and the corresponding nonlabeled primers, it can be used to separate the less-mobile 5'phosphate-labeled PCR product from its more-mobile nonlabeled counterpart. The difference in the rates of migration of the allele-specific PCR products allows the

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identification of a genotype as heterozygotic or homozygotic, without the need for radioactive or fluorescent probes. In this review article, we introduce these two gel-based methods based on DNA-binding zinc(II) complexes and describe their use in the detection of mutations in DNA.

Zinc(II)–Cyclen

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In 1993, Shionoya et al. [12] reported that the macrocyclic tetraamine zinc(II) complex Zn2+–cyclen binds selectively and reversibly to the imide groupcontaining nucleoside deoxythymidine (dT) in aqueous solutions with a dissociation constant Kd = [free dT][free Zn2+–cyclen]/[dT–Zn2+–cyclen] = 0.3 mM at pH 8.0 (Figure 1). In the resulting 1:1 complex of dT and Zn2+–cyclen, a coordination bond is formed between the deprotonated imido N– anion of the pyrimidine moiety and the Zn2+ ion (Zn2+–N– distance = 2.053 Å), and the total charge on the dT molecule increases from 0 to +1. Zn2+–cyclen derivatives also bind selectively to dT-rich regions of double-stranded DNA; this results in a change the conformation of the DNA, for example, to a bulbous structure. This activity was identified by means of nuclease footprinting experiments and gel mobility-shift assays [13–17].

Figure 1. Equilibrium for binding of Zn2+–cyclen to deoxythymidine (dT). Zn2+–cyclen binds selectively and reversibly to the imide group-containing nucleoside dT in an aqueous solution with a dissociation constant Kd = [free dT][free Zn2+–cyclen]/[dT––Zn2+–cyclen] = 0.3 mM at pH 8.0. The resulting 1:1 complex of dT––Zn2+–cyclen has a total charge of +1.

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Figure 2. Molecular mechanism for the duplex-selective characteristic of bis(quinolin-4ylmethyl) derivative of Zn2+–cyclen. The primer and template DNA are shown in red and black, respectively. (A) Binding of the Zn2+–cyclen derivative to the Thy base (T) in a homoduplex, where the accurate priming occurs, causes only a local conformation change, resulting in the formation of a small bubble, and does not inhibit a subsequent PCR. (B) In a heteroduplex, where mispriming occurs, a bulbous structure is formed at the site of the mismatch. Binding to the T base then produces a greater change in conformation with the formation of a larger bubble that causes in a greater depression in the Tm and, as a result, the Zn2+–cyclen derivative suppresses nonspecific PCR amplification.

The dissociation of Ade–Thy hydrogen bonds is promoted by Zn2+–cyclen, and this is manifested by a lowering of the melting temperature (Tm) of DNA with increasing concentration of Zn2+–cyclen. We have utilized these Thy-recognizing and Tm-depressing properties of Zn2+–cyclen in a specificity-enhanced PCR procedure that uses micromolar concentrations of a 1,7-bis(quinolin-4-ylmethyl) derivative of Zn2+–cyclen (Figure 2) [18].

Zinc(II)–Cyclen Polyacrylamide Gel Electrophoresis We have extended the range of applications of Zn2+–cyclen to include PAGE separation of various DNA fragments, a technique that we refer to as Zn2+– cyclen/PAGE [10]. A combination of a PCR-based heteroduplex method and

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Zn2+–cyclen/PAGE permits more-accurate detection of single mutations introduced artificially, even for less readily detectable substitutions, such as guanine-to-cytosine or cytosine-to-guanine (Gua/Cyt-to-Cyt/Gua) single mutations. The combined procedure, which requires no radioisotopes or fluorescent probes, is capable of detecting heteroduplex bands that arise from the annealing of partially noncomplementary strands, one from the mutant DNA and one from wild-type DNA (heterozygosity), during PCR. The heteroduplex bands on the PAGE gel are developed by staining with ethidium bromide. The approach is based on three principles. First, a single-base mismatch can produce a local conformational change in the double-stranded DNA (bubble formation), leading to differential migration of heteroduplex and homoduplex bands. Secondly, the addition to the gel of Zn2+–cyclen, which selectively binds with Thy bases and disrupts the double strands, can intensify local conformational changes (formation of larger bubbles), resulting in an increased difference in the mobility of the two duplexes on the gel.

Figure 3. The principle of a combined procedure involving PCR-based heteroduplexing and Zn2+–cyclen/PAGE for the detection of SNPs/mutations.

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Figure 4. Genetic analysis by using Zn2+–cyclen/PAGE followed by direct DNA sequencing. (A) PCR products (244 bp) amplified from 19 healthy individuals were applied to a 15% (w/v) polyacrylamide gel containing 5.0 mM Zn2+–cyclen. The allele genotype of CYP3A5 (*1/*3 heterozygote, *1/*1 homozygote, and *3/*3 homozygote) and subject number for each individual are shown above the corresponding lanes. (B) Typical direct sequencing data around the SNP site from Ade (*1) to Gua (*3) at nucleotide position 6986, which is associated with enzymatic activity for drug metabolism [50], from the anti-sense strand (T/C) for five individuals (subjects numbers 2, 8, 13, 14, and 15). (C) Typical direct sequencing data around another SNP site from Gua to Ade at nucleotide position 6929, which was identified as a novel SNP in CYP3A5 gene by means of Zn2+– cyclen/PAGE [20] from the anti-sense strand (C/T) for subject number 8. This figure is reproduced from ref. 20 with the permission of the publisher: © 2009 Elsevier Inc.

Finally, binding of Zn2+–cyclen to Thy bases decreases the total anionic charge on the target DNA, thereby providing facilitating electrophoretic detection. The appearance of slow or differentially migrating bands on the gel indicates the presence of a heteroduplex, implying the presence of an SNP/mutation. The principles of method for detecting SNPs/mutations by using Zn2+–cyclen/PAGE are shown schematically in Figure 3. Up to four distinct migration bands per DNA sample can be detected in this method.

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Zn2+-cyclen/PAGE has been used to screen for mutations in the SCN5A gene that are related to inherited arrhythmia syndrome [19], and for genomapping of SNPs in the CYP3A5 gene, which is associated with enzymatic activity for drug metabolism (Figure 4) [20]. Zn2+–cyclen/PAGE can only detect mutations accurately if the length of the DNA fragments (i.e., PCR amplicons) for analysis is about 200 base pairs (bp) or less.

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Detection of Mutations in Uanine/CytosineRich DNA Sequences We initially showed that Zn2+–cyclen/PAGE can be used to detect Gua/Cytto-Cyt/Gua single mutations through binding of Zn2+–cyclen to Thy bases near the mutation sites [10]. The question remained, however, whether Gua/Cyt-toCyt/Gua mutations that are not situated near Thy/Ade bases could be detected by means of the Thy-dependent method. In other words, the limits of detection of Gua/Cyt-rich DNA regions were uncertain. Gua/Cyt-lined DNA sequences, (Gua/Cyt)n (20 ≥ n ≥ 10), which can influence the phenotype by altering the encoded proteins, frequently appear in genetic coding regions of the human genome. To determine the limits of detection of the Thy-dependent Zn2+– cyclen/PAGE technique, we examined its ability to detect Gua/Cyt-to-Cyt/Gua single substitutions in some artificial Gua/Cyt-lined sequences [21]. In the samples tested, Gua/Cyt-to-Cyt/Gua substitutions that were 1 to 10 bases away from the nearest Thy/Ade moiety could be successfully detected by determining the appropriate DNA length (176 to 241 bp). In other words, single mutations in Gua/Cyt-lined sequences up to 20 bp long can be detected through binding of Zn2+-cyclen moieties to Thy bases near the sites of mutations. In addition, we examined the DNA binding properties of the Zn2+–cyclen moiety by sequencing analyses of DNA bands eluted from the Zn2+–cyclen/PAGE gel. We found that the slow migration of heteroduplexes containing Cyt–Cyt base pairings in the presence of Zn2+–cyclen contributes to accurate detection when analyzing the Gua/Cyt-to-Cyt/Gua single substitutions in Gua/Cyt-lined sequences. The disruption of an Ade–Thy base pairing as a result of binding with Zn2+–cyclen can affect even a mismatched site that is ten bases away, causing a local conformational change in the target DNA that can be detected on the Zn2+– cyclen/PAGE gel.

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Detection of Mutations in denine/ThymineRich DNA Sequences Our original Zn2+–cyclen/PAGE method was limited to analyses of Ade/Thyrich DNA sequences of fewer than 100 bp that contained more than 65% of Ade/Thy bases. Because the Zn2+-cyclen in the gel binds selectively to Thy bases in double-stranded DNA and thereby dissociates the double strand, it does not permit visualization of short Ade/Thy-rich PCR amplicons as double-stranded DNA bands. To overcome this problem, we developed an improved Zn2+cyclen/PAGE procedure that uses a PCR primer with a Gua/Cyt-lined sequence (ten bases) attached at its 5'-end to increase the Tm of the resultant PCR product [22]. By using this improved procedure, we successfully detected an artificial single-nucleotide substitution in a 51-bp DNA region containing 73% of Ade/Thy bases. Furthermore, we extended the improved method to permit screening for mutations in human genomic material extracted from formalin-fixed paraffinembedded (FFPE) tissue, from which it is difficult to obtain long PCR products. As the first practical example of the use of this extended procedure, we demonstrated the reliable screening of an Ade-to-Thy mutation at nucleotide position 19861 in the breast cancer BRCA1 gene (GenBank accession No. AY273801) in short Ade/Thy-rich PCR products (96 bp) amplified from genomic material extracted from FFPE sections of breast-cancer tissue. Direct DNA sequencing did not detect any mutations in these PCR amplicons, suggesting that the content of mutated DNA may be much lower than that of wild-type DNA in cancer-tissue specimens. In this case, our improved gel-based Zn2+–cyclen/PAGE separation procedure was more sensitive and more accurate than direct sequencing analysis.

Zinc(II)–Phos-Tag In 2004, we reported that a dinuclear metal complex of 1,3-bis[bis(pyridin-2ylmethyl)amino]propan-2-olate acts as a novel phosphate-capture molecule in aqueous solutions [23]. The dinuclear zinc(II) complex forms a stable 1:1 complex with a phosphate monoester dianion. The X-ray crystal structure of the 1:1 dinuclear zinc(II) complex with a p-nitrophenyl phosphate dianion showed that each phosphate oxygen anion binds to a zinc(II) ion at the fifth coordination site, and that the two zinc(II) ions in the complex are separated by a distance of 3.6 Å. Thus, the dinuclear zinc(II) complex, which has a vacancy on the two

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zinc(II) ions, is accessible to a phosphate monoester dianion that acts as a bridging ligand (Figure 5). In aqueous solutions, the dinuclear zinc(II) complex binds strongly to the phenyl phosphate dianion (Kd = 25 nM) at neutral pH values. The anion selectivity indexes against SO42–, CH3COO–, Cl–, and the diphenyl phosphate monoanion [(PhO)2P(=O)O–] at 25 °C are 5.2 × 103, 1.6 × 104, 8.0 × 105, and >2 × 106, respectively. In addition, the formation of a 1:1 adduct consisting of one molecule of the dizinc(II) complex and an inorganic phosphate dianion (HOPO32–) has been clearly detected by means of matrix-assisted laser desorption/ionization-time of flight mass spectrometry (MALDI-TOF MS) from the characteristic mass shift and change in total charge of the phosphate (from –2 to +1) as a result of binding of the dizinc(II) complex with HOPO32– [24]. This finding led to a simple, rapid, and sensitive procedure for the analysis of phosphorylated compounds, such as phosphopeptides or phospholipids, by MALDI-TOF MS using the dizinc(II) complex as an MS probe [24–26]. We therefore named 1,3-bis[bis(pyridin-2-ylmethyl)amino]propan-2-olate “Phos-tag”, as an abbreviation of “phosphate-binding tag molecule”. Since this discover, we have developed our Phos-tag technology to permit the analysis of protein phosphorylation (Phos-tag consortium, http://www.phos-tag.com/english/ index.html).

Figure 5. Structure of acrylamide-pendant Phos-tag ligand and a scheme for the reversible capture of a phosphomonoester dianion (R-OPO32–) by polyacrylamide-bound Zn2+–Phostag in PAGE.

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Phos-Tag Technology The Phos-tag technology utilizes a Phos-tag molecule that binds selectively and reversibly to a phosphomonoester dianion under physiological pH conditions. The Phos-tag technology has been applied in a range of procedures, including MALDI-TOF MS analysis of phosphorylated compounds [24–28], electrospray ionization MS analysis for online mass tagging of phosphopeptides [29], immobilized zinc(II) affinity chromatography for the separation of phosphopeptides and phosphoproteins [30–32], surface plasmon resonance analysis for detection of reversible peptide phosphorylation [33–35], Western blotting analysis of phosphoproteins on a protein-blotted membrane [36,37], and fluorescence analysis of protein phosphorylation and dephosphorylation [38,39]. The Phos-tag technology is capable of replacing conventional methods that use radioisotopes or antibodies, and it provides innovative methods for the separation, purification, enrichment, visualization, and detection of phosphopeptides and phosphoproteins. We have also established a technique for phosphate-affinity gel electrophoresis in which the tag molecule provides an effective means of monitoring the phosphorylation status of proteins [36,40–45]. On the basis of this new technique for affinity electrophoresis, we have developed several convenient applications that involve analyses of the states of phosphorylation of proteins. These include in vitro kinase activity profiling for the analysis of the phosphoprotein species derived from various kinase reactions, in vivo kinase activity profiling for the analysis of extracellular signal-dependent protein phosphorylation, in vitro kinase inhibition profiling for quantitative analysis of kinase-specific inhibitors, and a two-dimensional mobility-shifting procedure using phosphate-affinity gel electrophoresis for the detailed analysis of phosphoprotein species.

Zinc(II)–Phos-Tag Polyacrylamide Gel Electrophoresis We have also adapted the phosphate-affinity gel-based separation method for SNP genotyping [11]. Phosphate-labeled (phosphorylated) and nonlabeled (nonphosphorylated) PCR amplicons (i.e., allele-specific DNA with identical numbers of base pairs) can be separated by phosphate-affinity gel electrophoresis using a Phos-tag complex with two zinc(II) ions (Zn2+–Phos-tag/PAGE). We used acrylamide-pendant Zn2+–Phos-tag as a novel additive in a normal PAGE gel (see

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Figure 5) with Laemmli’s buffer system [46] without sodium dodecyl sulfate. This method has several advantages. First, the procedure is almost identical to that of conventional PCR and PAGE. Secondly, no radioactive label is required. Thirdly, it permits the separation of a 5'-OPO32–-labeled DNA fragment with a length of ~1500 bp as a band that is less mobile than its nonlabeled counterpart. Fourthly, it enables the determination of whether a genotype is heterozygotic or homozygotic through the difference in the rates of migration of the allele-specific PCR products. Finally, Zn2+–Phos-tag/PAGE can be used in gene diagnosis for both dominant and recessive characteristics. The procedure for SNP genotyping by using 5'-OPO32–-labeled and nonlabeled (5'-OH) PCR products is shown schematically in Figure 6. The DNA bands can be visualized by staining with ethidium bromide.

Figure 6. The principle of the combined allele-specific PCR and Zn2+–Phos-tag/PAGE procedure.

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SNP Genotyping

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As a first practical demonstration of the adapted Zn2+–Phos-tag/PAGE procedure, we performed SNP genotyping of a silent mutation in the SCN5A gene of healthy individuals.

Figure 7. Zn2+–Phos-tag/PAGE of allele-specific PCR products (220 bp) amplified from three SCN5A genotypes of seven healthy individuals. Each panel shows the results of genotyping for each individual. The allele genotype (Gua/Ade heterozygote, Gua/Gua homozygote, or Ade/Ade homozygote) of each individual is represented on the left-hand side of the corresponding panel. The amounts of genomic template DNA used (500, 250, 125, 63, or 32 pg) are shown above the corresponding lanes. The Zn2+–Phos-tag/PAGE gels (20 µM polyacrylamide-bound Zn2+–Phos-tag and 18% (w/v) polyacrylamide) were stained with ethidium bromide. This figure is reproduced from ref. 11 with the permission of the publisher: © 2006 Elsevier Inc.

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This mutation, which causes no alteration in the amino acid sequence, involves a change in a single Gua or Ade nucleotide at the 87 position and it has been catalogued as GenBank Accession No. M77235 [19]. A 220-bp DNA fragment containing this SNP position was amplified by PCR using a 1:1 mixture of 5'-OPO32–-labeled Gua-allele-specific primer and a nonlabeled Ade-allelespecific primer, together with the reverse nonlabeled primer (see Figure 6) and genomic templates from each of seven healthy individuals. Two of these individuals were Gua/Ade heterozygotic, three were Gua/Gua homozygotic, and the other two were Ade/Ade homozygotic. The resultant PCR amplicons were analyzed by using Zn2+–Phos-tag/PAGE. The 5'-OPO32–-labeled Gua-allele product and the nonlabeled Ade-allele product should have appeared as a slowly migrating band and a rapidly migrating band, respectively. For genotyping, it is important to achieve significant differences in the quantities of allele-specific PCR amplicons from the simultaneous amplification of all the individual DNA samples. However, when a protocol involving a fixed thermal cycle (i.e., a fixed annealing temperature and a fixed number of amplification cycles) is used, it is difficult to identify a difference in the kinetics of allele-specific amplification of individual genomic templates if these differ in quality (for example, when they are present in different concentrations and/or they have different levels of purification). To avoid this problem, we performed five individual PCR reactions using 500, 250, 125, 63, or 32 pg of the genomic template for each individual genotyping. Figure 7 shows that the allele-specific PCR amplicons were clearly observed as a less-mobile band corresponding to the 5'-OPO32–-labeled Gua-allele product and more-mobile band corresponding to the nonlabeled Ade-allele product. In the case of the Gua/Ade heterozygote, signals of both alleles were observed almost equally in all lanes, whereas for the Gua/Gua or Ade/Ade homozygote samples, small amounts of nonspecific products were observed together with the specific product. There was, however, a significant difference in the quantities of the two amplified products. Genomic samples weighing 125–500 pg were found to be sufficient to permit the identification of the allele genotype when they were used as templates for allele-specific amplification.

Detection of Epigenetic Mutations Involving Methylation of DNA DNA methylation is a well-characterized epigenetic DNA modification that plays a role in the regulation of gene expression during cell and cancer

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development [47]. Many methods have been developed for analyzing the state of methylation of genomic DNA. Currently, almost all such methods employ the bisulfite-mediated deamination of denatured DNA, which effectively converts cytosine into uracil but leaves 5-methylcytocine intact [48]. Levels of uracil or 5methylcytocine in the resultant DNA sample can then determined by methylationspecific PCR (MSP) followed by DNA sequencing or other techniques. We developed a procedure for the analysis of DNA methylation by using a bisulfitemediated cytosine-to-uracil conversion of a target DNA, followed by MSP and phosphate-affinity Zn2+–Phos-tag/PAGE [49]. MSP was performed by using a 1:1 mixture of 5'-phosphorylated methylation-specific and 5'-OH nonmethylationspecific primers and a reverse nonlabeled primer. The combination of the MSP technique and Zn2+–Phos-tag/PAGE allowed shifts in methylation-specific products to be detected as less-mobile bands. We believe that the Zn2+–Phostag/PAGE method could be widely used in the medical field as a simple and effective method for identifying methylation of DNA.

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Conclusion We have introduced two gel-based methods that utilize DNA-binding zinc(II) complexes in the detection of mutations in DNA. Both methods are based on electrophoresis and use zinc(II) complexes to provide simple and accurate procedures for detecting SNPs/mutations without the use of radioisotopes, expensive fluorescent primers, or expensive special equipment. Because the procedures require only standard laboratory PCR apparatus and mini-slab PAGE systems, they could serve as convenient tools that would allow clinical researchers and physicians to obtain genetic information from small numbers of patients. Most other gel-based methods for small-scale assays, such as single-strand conformation polymorphism, denaturing gradient gel electrophoresis, or conformation-sensitive gel electrophoresis, involve complicated processes require skilled analysts and special apparatus. The Zn2+–cyclen/PAGE method has a high-resolution power and can distinguish not only between the homoduplexes and heteroduplexes, but also between pairs of homoduplexes (see Figure 4, CYP3A5*1 and CYP3A5*3) with the same numbers of base pairs (~200 bp). A single DNA band on the Zn2+– cyclen/PAGE gel after heteroduplexing indicates an absence of any mutation: the detection specificity would not be less than that (generally 70 to 90%) of other gel-based approaches described above [19,20]. The procedure with the Zn2+–

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cyclen is also very useful for the discovery of an unknown SNP/mutation. An assay using the Zn2+–Phos-tag/PAGE method has been shown to be 100% effective in detecting SNPs/mutations when allele-specific PCR amplicons of fewer than 1500 bp, optionally labeled with 5'-OPO32–, are used. The procedure with the Zn2+–Phos-tag is thus very useful for the identification of a key SNP/mutation which is targeted. Furthermore, the combination of Zn2+– cyclen/PAGE and Zn2+–Phos-tag/PAGE would permit more accurate and reliable genotyping and genomapping [20]. Regarding the Zn2+–cyclen dinitrate (commercially available from NARD Institute, Ltd., Amagasaki, Japan), which has a potency not only as a mobility shift-enhancer for PAGE but also as a specificity-enhancer for PCR, the cost of using Zn2+–cyclen for electrophoresis of 19 DNA fragments as samples on a minislab gel (1 mm × 9 cm × 9 cm) containing 5.0 mM of Zn2+–cyclen is less than 1 US dollar per run. In the case of the Phos-tag molecule, which is also a potent mobility shift-enhancer of phosphorylated proteins, one hundred mini-slab PAGE gels, each containing 20 μM of Zn2+–Phos-tag, can be prepared from 10 mg of the commercially available acrylamide-pendant Phos-tag ligand (Wako Pure Chemical Industries, Ltd., Osaka, Japan), so that the cost of the tag molecule for each gel is about 6 US dollars. It is therefore worth considering these procedures for highly sensitive and cost-effective medical screening and genotyping of various samples, including FFPE tissue samples, for a range of disease-causing mutations.

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[45] Kinoshita, E. and Kinoshita-Kikuta, E. (2011) Improved Phos-tag SDSPAGE under neutral pH conditions for advanced protein phosphorylation profiling. Proteomics, in press. [46] Laemmli, U. K. (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature (London, U. K.), 227, 680–685. [47] Shen, L., Kondo, Y., Guo, Y., Zhang, J., Zhang, L., Ahmed, S., Shu, J., Chen, X., Waterland, R. A. and Issa, J-P. J. (2007) Genome-wide profiling of DNA methylation reveals a class of normally methylated CpG island promoters. PLoS Genet., 26, 2023–2036. [48] Frommer, M., McDonald, L. E., Millar, D. S., Collis, C. M., Watt, F., Grigg, G. W., Molloy, P. L. and Paul, C. L. (1992) A genomic sequencing protocol that yields a positive display of 5-methylcytosine residues in individual DNA strands. Proc. Natl. Acad. Sci. U. S. A., 89, 1827–1831. [49] Kinoshita-Kikuta, E., Kinoshita, E. and Koike, T. (2008) A mobility shift detection method for DNA methylation analysis using phosphate-affinity polyacrylamide gel electrophoresis. Anal. Biochem., 378, 102–104. [50] Kuehl, P., Zhang, J., Lin, Y., Lamba, J., Assem, M., Schuetz, J., Watkins, P. B., Daly, A., Wrighton, S. A., Hall, S. D., Maurel, P., Relling, M., Brimer, C., Yasuda, K., Venkataramanan, R., Strom, S., Thummel, K., Boguski, M. S. and Schuetz, E. (2001) Sequence diversity in CYP3A4 promoters and characterization of the genetic basis of polymorphic CYP3A5 expression. Nat. Genet., 27, 383–391.

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In: DNA Replication and Mutation Editor: Raymond P. Leitner

ISBN: 978-1-61324-490-6 © 2012 Nova Science Publishers, Inc.

Chapter VI

DNA Mutations and Genetic Coding Jean-Luc Jestin∗

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Unité de Virologie Structurale, Département de Virologie, Institut Pasteur, France

Abstract There is a need to understand how biological organization at one level translates into biological organization at the next level. Biology encompasses numerous levels from nanometers to meters, from atoms and macromolecules to organisms and populations of individuals. Here we shall consider the DNA mutations at the level of single nucleotides and their impact at the higher level of codons, sets of synonymous codons and their symmetries linked to tRNA-aminoacylation and degeneracy in the genetic code. Our review will be restricted to DNA mutations, which are either single-base deletions known to be highly deleterious within open reading frames or base substitutions, which are transitions (substitutions of a pyrimidine by another pyrimidine or substitutions of a purine by another purine) or transversions. Concepts such as coevolution, minimization of the effects of errors or metrics which found so far little use in the natural sciences, can be

∗

Corresponding Author: Address: 25 rue du Dr. Roux 75724 Paris 15, France. Tel: +33 1 4438 9496.

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Jean-Luc Jestin successfully applied to understand the connections between biological properties at these molecular and macromolecular levels.

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Introduction The genetic code establishes the link between triplets of nucleotides or codons, and amino acids and thereby the link between genes and their corresponding proteins at the macromolecular level (Sonneborn 1965, Crick 1966, Nirenberg 2004). The genetic code is highly conserved among all living organisms and is therefore said to be quasi-universal (Ninio 1986, Jukes and Osawa 1990) (Table 1). The genetic code is also degenerate, as several codons can code for an amino acid or a chain termination signal. Several reports indicate that the genetic code properties could also help identifying conditions in which the genetic code emerged a few billion years ago (Di Giulio 2005, 2007, Gutfraind and Kempf 2008, Jestin and Kempf 2009). Understanding why the genetic code is the way it is, is also of high interest as it may provide new insights in the fields of biological chemistry and biochemistry. As a code, the genetic code could be expected to have general properties of codes as defined in coding theory (Conway). As coding theory and the genetic code remained independent fields, it has been useful to consider the basics of coding theory, that is number theory (Jestin 2003, Khrennikov and Nilsson 2009, Dragovich and Dragovich 2010). This provided also new insights on the understanding of why symmetries by base substitutions of degeneracy in the genetic code are the way they are (Jestin 2003, 2010). Interestingly, links between information theory and biological information have been established (Tlusty 2010). Table 1. Variations between the quasi-universal or standard genetic code and mitochondrial genetic codes from the organisms cited Genetic code Standard Vertebrate mitochondria Nematodes, molluscs and insects mitochondria

AGR Arg stop Ser

Source: (derived from from Jukes and Osawa, 1990).

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ATA Ile Met Met

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Importantly, these concepts have to be consistent with the theory of evolution and molecular evolution in particular as well as with the plethora of experimental data available in molecular biology and directed molecular evolution (Jestin and Kaminski 2004, Joyce 2004, Seelig and Szostak 2007). Genetic code and biological macromolecules have necessarily been coevolving, suggesting that DNA mutations may have had a central role in genetic coding. It is the aim of this review to detail essential links between these mutations and the genetic code. While base substitutions are the most frequent DNA mutations, frameshift mutations are the most deleterious ones.

Single-Base Deletions in DNA and Protein Chain Termination Coding

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Frameshift mutations are highly deleterious within open reading frames, as they yield truncated proteins which are generally non functional (Figure 1).

Figure 1. The BYAR sequences in which most of the single-base deletion occur yield truncated proteins within open-reading frames (ORF) or full-length proteins with a peptide added at their C-terminus within stop codons. These fusion proteins are likely to be functional while truncated proteins are unlikely to be functional.

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Frameshift mutations can be additions or deletions of one or more nucleotides within nucleic acids. The most frequent frameshift mutations are single-base deletions. These deletions occur in two types of sequence contexts, reiterated runs such as homopolymeric sequences or non-reiterated runs. In vitro polymerization assays with DNA polymerases from families A and B have shown that most single base deletions in non-reiterated runs occur opposite the template purine R within TR sequences. This was observed for 59% to 95% of the single-base deletions catalyzed by E. coli DNA polymerase I or its domains mutated or not, for 72% of the single-base deletions catalyzed by the exonuclease deficient T7 DNA polymerase, and for 46% to 56% of the single-base deletions catalyzed by three vertebrate DNA polymerases alpha (Kunkel 1985, De Boer and Ripley 1988, Bebenek et al. 1990). The consensus sequence for single-base deletions in non-reiterated runs was refined to 5’-YTRV-3’ template sequences (Jestin and Kempf 1997). Interestingly, mapping of these consensus sequences on the genetic code codon table indicates that the two stop codons TAA and TAG, as well as the two reverse-complementary sequences TTA and CTA are the codons in which most single-base deletions occur. Assigning the most deletion prone sequences to chain termination codons, i.e. stop codons rather than to codons encoding amino acids maximizes fitness by increasing the proportion of proteins which are functional (Jestin and Kempf 1997). This is because a single base deletion within an open reading frame yields a truncated protein, while a single base deletion at a chain termination codon yields a full-length protein fused to a peptide at its C-terminus (Figure 1). The observation that stop codons are single base deletion prone sequences (Jestin and Kempf 1997) supports the theory stating that the genetic code was selected so as to minimize the deleterious effects of mutations (Sonneborn 1965, Goldberg and Wittes 1966).

Nucleotide Substitutions and Symmetries of Degeneracy in the Genetic Code It has long been known that transitions, i.e. substitutions exchanging purines or exchanging pyrimidines (A into G, G into A, C into T, T into C) on the third base of codons are almost always neutral (Goldberg and Wittes 1966). This minimizes the deleterious effects of the most frequent substitutions catalyzed by DNA polymerases, the transitions.

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Figure 2. A representation of the vertebrate mitochondrial genetic code highlighting unique symmetries of degeneracy by base substitutions. Vertical arrows correspond to a symmetry which exchanges the degeneracy groups when applied to all three codon bases. Horizontal arrows correspond to a symmetry which leaves invariant the degeneracy group when applied to the first base of codons. (GT/AC) indicates the transversion which exchanges G and T as well as A and C. (GC/AT) indicates the transversion which exchanges G and C as well as A and T. Note that the transitions (AG/CT) applied to the third base of codons leave also invariant the degeneracy group.

The same observation can also be interpreted at the translational level by considering codon-anticodon recognition (Crick 1966, Ninio 1971). Transitions at third codon bases are always neutral in the vertebrate mitochondrial genetic code (Figure 2). This is consistent with the model according to which the genetic code was selected to reduce the impact of mutations (Sonneborn 1965, Goldberg and Wittes 1966, Jestin and Kempf 1997). Rumer dissected the 64 codons according to degeneracy into two groups of 32 codons, depending on whether the third codon base has to be defined so as to define unambiguously the amino acid or the stop signal (Rumer 1966, Shcherbak 1989). Interestingly, Rumer found a symmetry which exchanges both groups: this symmetry is the transversion substituting G and T as well as A and C, applied to all three codon bases (vertical arrow, Figure 2). A further symmetry that leaves unchanged both groups was identified (Jestin 2006): this symmetry is the transversion substituting G and C as well as A and T, applied to the first base of codons (horizontal arrow, Figure 2). It was then shown that there is no further symmetry exchanging or leaving invariant these degeneracy groups, except for the combination of the previously described ones (Jestin and Soulé 2007). The genetic code is then represented by two groups, one group of 8 amino acids with a four-fold degeneracy and one group of 16 amino acids or stop signal with a two-fold degeneracy. Note that amino acids encoded by six codons are considered as being encoded by four plus two codons. It should also be mentionned that in the standard genetic code, some of the amino acids are

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encoded by one or by three codons, while in the vertebrate mitochondrial genetic code, the degeneracy is always two-, four- or six-fold: these minor differences in genetic coding have been widely discussed (Ninio 1986, Jukes and Osawa 1990). Let us consider the matrix M of the numbers of substitutions from any codon to any other codon. Two codon to codon substitutions can be said similar if they have a mutation in common, that is if the two numbers of substitutions have a factor in common (Figure 3).

Figure 3. Matrix of the numbers of substitutions from any codon to any other codon. The two substitutions highlighted have a common A to T mutation on the first and third codon positions. The two numbers of this matrix that are highlighted have accordingly a factor in common, the probability for A to mutate to T, pAT. pA, pC and pG are respectively the probabilities for A, C and G not to mutate. Alpha and delta are proportionality factors.

Figure 4. A 3-adic tree. Note that 53 = 1 x 33 + 2 x 32 + 2 x 31 + 2 x 30 as evidenced by the tree. The arrows indicate the 3-adic distances d3 between two numbers. For example, d3 (80-53) = d3 (27) = d3 (33) = 1 / 33; d3 (74-5) = d3 (69) = d3 (23 x 31) = 1 / 31. By analogy with a phylogenetic tree, the power of 3 in the 3-adic distance indicates the level in the hierarchy at which the two numbers have a common ancestor; 80 and 53 have a common ancestor at generation 3, while 5 and 74 have a common ancestor at generation 1.

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Metrics in which close numbers are numbers which have a factor in common, are commonly used in number theory. This is the case of p-adic metrics (Serre 1973, Conway and Sloane 1988). 3-adic integers and 3-adic distances are given as an example in Figure 4. The number of non-synonymous substitutions S for any set of synonymous codons is the sum for all synonymous codons of the number of non-synonymous substitutions for each synonymous codon. It is then calculated from the matrix M according to the following equation:

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S = x M (1-x) where x is the vector, the n-tuple (for n codons) characterizing the set of synonymous and non-synonymous codons (x1, x2,… xi…, xn) with xi = 1 for the synonymous codons and xi = 0 for the non-synonymous codons (Jestin 2010). Each set of synonymous codons is then characterized by a class of equivalent quadratic forms over the field of rational p-adic numbers, Qp (Jestin 2010). There are 8 classes of quadratic forms for p ≠ 2 with the determinant belonging to the quotient group that can be considered as a 2-dimensional vector space over the field F2, and 16 classes of quadratic forms for p = 2 with the determinant belonging to the quotient group that can be considered as a 3dimensional vector space over the field F2 (Serre 1973). Consistently, there are 8 sets of synonymous codons, for which 2 bases have to be defined so as to define unambiguously the amino acid encoded (the four-fold degeneracy group), and 16 sets of synonymous codons, for which 3 bases have to be defined so as to define unambiguously the amino acid or the stop signal (the two-fold degeneracy group) (Figure 2).

Conclusion The notion of coevolution of the genetic code and biological macromolecules has been extremely fruitful in providing optimization models accounting for some properties of the genetic code. Answers have been for example provided for the following questions. Why are transitions at the last base of codons generally neutral? Why are stop codons what they are? Why are stop signals encoded as stop codons within the genetic code? Identifying and understanding the symmetries of objects has always been a central step for the progress of knowledge. Interestingly, discretization of the

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codon to codon substitutions matrix by making use of the field of p-adic rational numbers instead of the field of real numbers, highlights symmetries of the sets of synonymous codons, that would remain unexplained otherwise. Apart from the need to understand why the genetic code is the way it is, there remains some hope that the understanding of organization at the levels of nucleotides, of codons and of amino acids will be useful for a better understanding at the next higher levels of biological macromolecules.

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References Bebenek, K.; Joyce, C.; Fitzgerald, M. and Kunkel, T. (1990) The fidelity of DNA synthesis catalyzed by derivatives of E. coli DNA polymerase I. J. Biol. Chem., 265, 13878-13887. De Boer, J. and Ripley, L. (1988) An in vitro assay for frameshift mutations: hotspots for deletions of one base pair by Klenow fragment polymerase share a consensus DNA sequence. Genetics, 136, 709-719. Conway, J. H. and Sloane, N. J. A. (1988) Sphere packings, lattices and groups. New York, Springer. Crick, F.H.C. (1966) Codon-anticodon pairing: the wobble hypothesis. J. Mol. Biol. 1966, 19, 548-555. Di Giulio, M. (2005) The ocean abysses witnessed the origin of the genetic code. Gene. 346, 7-12. Di Giulio, M. (2007) The evolution of the genetic code took place in an anaerobic environment. J. Theor. Biol. 245, 169-174. Dragovich, B. and Dragovich, A. (2010) p-adic modelling of the genome and the genetic code. Computer Journal, 53, 432-442. Goldberg, A. and Wittes, R. (1966) Genetic code: aspects of organization. Science, 153, 420-424. Gutfraind, A. and Kempf, A. (2008) Error-reducing structure of the genetic code indicates origin in non-thermophile organisms. Orig. Life Evol. Biosph., 38, 75-85. Jestin, J. L. and Kempf, A. (1997) Chain termination codons and polymeraseinduced frameshift mutations. FEBS Letters, 419, 153-156. Jestin, J. L. (2003) Degeneracy in the genetic code and its symmetries by base substitutions. March, 11. Available from: http://arxiv.org/abs/physics/ 0303045.

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Jestin, J. L. and Kaminski, P. A. (2004) Directed enzyme evolution and selections for catalysis based on product formation. J. Biotech., 113, 85-103. Jestin, J. L. (2006) Degeneracy in the genetic code and its symmetries by base substitutions. Comptes Rendus Biol., 329, 168-171. Jestin, J. L. and Soulé, C. (2007) Symmetries by base substitutions in the genetic code predict 2’ or 3’ aminoacylation of tRNAs. J. Theor. Biol., 247, 391-394. Jestin, J. L. and Kempf, A. (2009) Optimization models and the structure of the genetic code. J. Mol. Evol., 69, 452-457. Jestin, J. L. (2010) A rationale for the symmetries by base substitutions of degeneracy in the genetic code. Biosystems, 99, 1-5. Joyce, G. F. (2004) Directed evolution of nucleic acid enzymes. Ann. Rev. Biochem., 73, 791-836. Jukes, T. H. and Osawa, S. (1990) The genetic code in mitochondria and chloroplasts. Experientia, 46, 1117-1126. Khrennikov A. Y. and Nilsson, M. (2009) A number theoretical observation about the degeneracy of the genetic code. Proc. Steklov Institute Math., 2009, 265, 140-142. Kunkel, T. (1985) The mutational specificity of DNA polymerases alpha and gamma during in vitro DNA synthesis. J. Biol. Chem., 260, 12866-12874. Ninio, J. (1971) Codon-anticodon recognition: the missing triplet hypothesis. J. Mol. Biol. 56, 63-74. Ninio, J. (1986) Divergence in the genetic code. Bioch. Systematics and Ecology, 14, 455-457. Nirenberg, M. (2004) Historical review: deciphering the genetic code. Trends Biochem. Sci., 29, 46-54. Rumer, Y. B. (1966) About the codon’s systematization in the genetic code. Proc. Acad. Sci. USSR, 167, 1393-1394. Seelig, B. and Szostak, J.W. (2007) Selection and evolution of enzymes from a partially randomized non-catalytic scaffold. Nature, 448, 828-831. Serre, J. P. (1973) A course in arithmetic. New York, Springer. Shcherbak, V. I. (1989) Rumer’s rule and transformation in the context of the cooperative symmetry of the genetic code. J. Theor. Biol., 139, 271-276. Tlusty, T. (2010) A colourful origin for the genetic code: information theory, statistical mechanics and the emergence of molecular codes. Phys. Life Rev., 7, 362-376. Sonneborn, T. M. (1965) Degeneracy in the genetic code: extent, nature and genetic implications. In V. Bryson and H. J. Vogel (Eds.), Evolving genes and proteins (pp. 377-397). New York, Academic Press.

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Chapter VII

Nuclear DNA Replication in Trypanosomatid Protozoa M. S. da Silva1, R. C. V. da Silveira1, A. M. Perez1, J. P. Monteiro1,2, S. G. Calderano3, J. P. da Cunha3, M. C. Elias3 and M. I. N. Cano1 Copyright © 2012. Nova Science Publishers, Incorporated. All rights reserved.

1

Departamento de Genética, Instituto de Biociências, UNESP, Botucatu, São Paulo, Brazil 2 EMBRAPA-Caprinos e Ovinos, Sobral, Ceará, Brazil 3 Laboratório de Toxinologia Aplicada, Instituto Butantan, São Paulo, Brazil

Abstract The parasites belonging to the family Trypanosomatidae (order Kinetoplastida) are among the most primitive eukaryotes. Some trypanosomatids are the etiologic agents of neglected human pathologies such as South American and African trypanosomiasis and leishmaniasis. As a consequence of their ancient phylogenetic position, nuclear DNA replication in trypanosomatid protozoa shows conserved and non-conserved features. DNA replication in trypanosomatids initiates nearly simultaneously in the nucleus and in the genetic material of the single mitochondrion (or kinetoplast), suggesting that DNA synthesis is coordinately regulated in both organelles. In eukaryotes, nuclear DNA replication is preceded by assembly of the pre-replication complex, which is coordinated by the Origen

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M. S. da Silva, R. C. V. da Silveira, A. M. Perez et al. Recognition Complex (ORC). However, in trypanosomatids, the prereplication complex differs from other eukaryotes and is similar to Archaea. All of these parasites contain only one protein that recognizes the replication origins and is found in the nucleus throughout the cell cycle, which suggests that it is not involved in the control of replication initiation. In the S phase, DNA replication starts at these origins and, in trypanosomes, occurs mainly at the nuclear periphery. In Leishmania spp., from the beginning up to mid S phase, replication sites are spread throughout the nuclear space to form subnuclear foci of active DNA replication. From mid-to-late S phase, replication is restricted to sites at the nuclear periphery. Few nuclear DNA polymerases have been described in trypanosomatid protozoa, although putative members of all polymerase families are found in their genomes. Structural and functional analyses indicate that most of these polymerases are highly conserved, with some of them being involved in polymerization and the repair of DNA damage. Although there are no descriptions of DNA polymerase δ in these protozoa, one of this protein´s partners, proliferating cell nuclear antigen (PCNA), is found in the nucleus throughout the cell cycle. Trypanosomatid PCNA forms distinct subnuclear foci in the S phase, whereas its distribution is more diffuse in the G2/M phase and in post-mitotic phase cells. This finding suggests that there may be phase-specific regulation of PCNA in the cell cycle. DNA replication in trypanosomatid telomeres is terminated by the action of telomerase. The biochemical properties of the trypanosomatid enzyme are conserved and resemble those described in other eukaryotes. Leishmania telomeres replicate late in S phase and at the beginning of G2 phase the chromosomes cluster at the nuclear periphery. Telomerase co-localizes with telomeres from the late S to G2 phases. These observations point to the existence of replication factories in trypanosomatids, the importance of which will be reviewed and discussed in this chapter.

Introduction 1. General Aspects of Nuclear DNA Replication in Higher Eukaryotes In most eukaryotes, DNA replication is precisely coordinated and tightly controlled in time and space to ensure that the entire genome is duplicated with high fidelity, exactly once per cell cycle, without errors. Thus, DNA replication presents a significant challenge to the cell given the size and complexity of eukaryotic genomes. The initiation and completion of DNA replication define, respectively, the beginning and end of the S phase and require the coordinated

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action of numerous factors that function in all phases of the cell cycle (Takeda and Dutta, 2005). DNA replication starts at a small number of sites within the nucleus and proceeds rapidly in early S phase. Replication is primarily confined to the heterochromatic regions in mid-to-late S phase and to the nuclear periphery at the end of S phase. DNA replication is regulated by key cell-cycle regulators, including cyclin-dependent kinases (CDKs), the anaphase promoting complex/cyclosome (APC/C), and pre-replicative complexes (pre-RCs), which include the Origin Recognition Complex (ORC) and Cell Division Cycle (CDC) (Diffley, 2004). Assembly of the replication machinery starts in late mitosis to early G1 phase at a large number of replication origins distributed on multiple chromosomes. Consequently, pre-RCs cannot assemble at origins during the S, G2 and M phases, but the number of replicating units clearly increases as the S phase progresses, becoming more coordinated in space (Fox et al., 1995; Diffley, 2004). The assembly of essential pre-RCs at origins occurs when CDK activity is low and APC/C activity is high, resulting in the elimination of cyclins and, in metazoans, geminin. This mechanism ensures that no origin can fire more than once in a cell cycle, thereby preventing re-initiation, which is crucial for maintaining genome integrity and avoiding the activation of checkpoint responses (Vaziri et al., 2003; Diffley, 2004). Despite distinct differences in origin sequence and structure, preRCs direct the order of assembly of replication factors, which are highly conserved across species. In higher eukaryotes, including mammals, the pre-RCs consist of the hexameric ORC (Orc1–Orc6), CDC6 and Cdt1 (Chromatin Licensing and DNA replication factor 1). During the transition to S phase, preRCs recruit the heptameric MCM2-7 (Mini Chromosome Maintenance) complex, which possesses helicase activity, to promote the unwinding and separation of the two strands of double-stranded DNA. In a cascade of events, MCM10 and CDC45 are loaded in the replication fork, the Y-shaped region of a replicating DNA molecule that results from separation of the DNA strands and where the synthesis of new strands occurs. The association of CDC45 with DNA polymerase and MCMs suggests that this complex may coordinate the function of these components in the replication fork, and its association with RPA (Replication Protein A) could help to tether the complex at the replication fork (Bell and Dutta, 2002). In summary, CDC45 is required for the assembly of many components of the DNA synthetic machinery at the replication fork, including RPA, PCNA (Proliferative Cell Nuclear Antigen), and DNA polymerases (pol) α and ε. In most eukaryotes, the assembly of DNA polymerases is highly ordered to ensure that all of the necessary polymerases are present at the origin prior to the synthesis

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of the first RNA primer by DNA pol α/primase. Although all three polymerases (pol α, pol δ and pol ε) share a conserved catalytic core, they perform specialized functions in elongation. DNA pol α is the only polymerase that can initiate synthesis de novo on single-stranded DNA, whereas DNA pol δ and DNA pol ε require a primed template for activity (Takeda and Dutta, 2005). RPA is required for the loading of DNA pol α but in some organisms RPA is not required for the association of DNA pol ε with the chromatin (Zou and Stillman, 2000; Mimura et al., 2000). Once DNA pol α has been recruited to origins and its primase activity has produced short RNA primers for synthesis of the leading and lagging strands, a polymerase switch replaces DNA pol α by DNA pol δ and/or pol ε. The latter two enzymes are very active 5’-3’ polymerization enzymes that have also 3’-5’ proofreading exonuclease activity. DNA pol δ and/or pol ε associate with PCNA, which in turn is loaded onto DNA by RFC (Replication Factor C), also known as the clamp loader (Elison and Stillman, 2001). DNA pol ε does not significantly elongate primed templates in the absence of DNA pol δ (Fukui et al., 2004), suggesting that DNA pol δ is the main polymerase for the synthesis of both strands. After mid S phase, the sites of DNA replication condense and late in the S phase, replication subsides in highly condensed heterochromatin. At the end of the S phase, replication is restricted to sites at the nuclear periphery, which probably represents the replication of telomeres that are attached to the nuclear membrane (Fox et al., 1995). Telomere replication by a telomere-specific reverse transcriptase (telomerase) solves the “end-replication problem”, or the inability of conventional DNA polymerases to replicate linear molecules fully (Greider and Blackburn, 1985). The sequence loss that is predicted to occur as a result of the deletion of the RNA primer of the most distal Okazaki fragment in the lagging strand is considerably less than has been observed in primary human cells (losses of ~100–200 bases of TTAGGG repeats per cell division). Thus, telomere loss appears to be caused by a combination of the end-replication problem and the exonucleolytic processing that must occur to create the single-stranded G-rich overhang on the telomeres of both leading- and lagging-strands (Harley et al., 1990; Counter et al., 1992; Sfeir et al., 2005; Chai et al., 2006; reviewed in Baird, 2008). These distinctive patterns demonstrate a programmed control of replication sites in the spatial domain in the nuclei of differentiated cells. The time at which a eukaryotic origin initiates replication within the S phase is characteristic of each origin, with early replicating origins being associated mostly with transcriptionally active regions and late replicating origins generally being

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associated with transcriptionally repressed genes. This arrangement indicates that the surrounding chromatin and not the origin itself is the major determinant of replication timing (Diller and Raghuraman, 1994; Friedman et al., 1996). Although proteins and many features involved in DNA replication have been conserved in eukaryotes, and nuclear DNA replication was established at an early stage of eukaryotic evolution, the process is still poorly understood in trypanosomatids. Recent findings have revealed surprising variations in how replication is controlled in these protozoa, implying that a conserved mechanism has not necessarily resulted in the conservation of regulatory pathways. Another interesting point is the fact that in different organisms some replication factors, such as ORC and MCM, work outside the replication machinery (Foss et al., 1993; Zhang et al., 1998; Prasanth et al., 2002; Chesnokov et al., 2003), which suggests species-specific differences in their regulation. These differences may yield clues to understanding the problem of origin timing and spacing, and provide insights into how DNA replication is influenced by factors such as gene density, epigenetic changes and the genetic background of the cell (Takeda and Dutta, 2005). Overall, variations in the control of DNA replication among lower and higher eukaryotes represent different evolutionary paths for the same purpose since the replication mechanism is apparently very well conserved (Kearsey and Cotterill, 2003).

2. Peculiarities of Kinetoplastida Protozoa: The Family Trypanosomatidae Protozoan parasites belonging to the order Kinetoplastida exhibit many peculiar and particular biological features that mirror their very ancient evolutionary origin. Among these parasite are members of the family Trypanosomatidae (dubbed Tri-tryps) that include Trypanosoma cruzi, Trypanosoma brucei and various species of Leishmania, the etiologic agents of medical and veterinary diseases that are common in tropical and subtropical areas of the world (Johnston et al., 1999; Gull, 2001). Most of these diseases are considered neglected because they affect the poor and because of the lack of effective treatment and prophylaxis. Consequently, in recent years, considerable effort has been devoted to the development of new therapeutic alternatives (Yamey and Toreele, 2002; Alvar et al., 2006). Many pathogenic trypanosomatids are digenetic parasites with complex life cycles that include different developmental forms in vertebrate and invertebrate

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hosts. This variation is central to successful parasite adaptation and the movement of these parasites between vector and host. Their life cycle is characterized mainly by changes in cell shape, cell cycle, metabolism, surface coat and gene expression which, in this case, also has its peculiarities (Johnston et al., 1999; Gull, 2001; Barrett et al., 2003; Simpson et al., 2004). In the sections below, we describe the peculiarities of these organisms and stress the evolutionary differences between them and their eukaryotic hosts. Trypanosoma cruzi: The etiological agent of American trypanosomiasis (Chaga’s disease), T. cruzi, alternates its life cycle between flagellate epimastigotes that live in the gut of triatomine bugs, non-proliferative trypomastigotes that can infect the mammalian host, and the dividing, intracellular and infective amastigote forms. Trypomastigotes are taken from the mammalian host by triatomine bugs while feeding on blood. In the insect vector, they transform into epimastigotes, thus completing the life cycle (Pereira, 1990; Barrett et al., 2003) (Fig. 1A). The high genetic diversity of T. cruzi compared to other trypanosomatids has led to this species being classified into six major genotypes (TcI to TcVI) or ‘Discrete Typing Units’ (DTUs). This classification groups parasite subspecies by biological, biochemical, genetic and eco-epidemiologic similarities since increasing evidence indicates that genetic exchange between parasites has contributed to the present population structure (Zingales et al., 2009; Sturm and Campbell, 2010). The T. cruzi nuclear genome (~112 Mb) has been sequenced and contains about 41 pairs of chromosomes with approximately 23,000 genes, as well as repetitive sequences (large gene families of surface proteins, retrotransposons and subtelomeric sequences) that account for ~50% of the genome (El-Sayed et al., 2005a; Weatherly et al., 2009). As in other trypanosomatids, most of the genes are organized in polycistronic arrangements at internal chromosomal positions, whereas some, including those encoding virulent surface antigens such as certain trans-sialidase family members (i.e. gp85), occur at subtelomeres (Cano, 2001). This organization suggests that T. cruzi chromosomal ends could have been the site for the generation of new variants of important adhesion molecules involved in the invasion of mammalian cells (Kim et al., 2005). Trypanosoma brucei: Most T. brucei subspecies have a more complex developmental cycle than observed in other trypanosomatid. This cycle alternates between proliferative and non-proliferative forms. The procyclic trypomastigote forms, which live in the insect (tse-tse fly) midgut, divide and differentiate into epimastigotes that in turn change into non-dividing metacyclic trypomastigotes in the salivary gland of the insect. Metacyclic trypomastigotes can infect the mammalian host and differentiate into multiplying, proliferative bloodstream

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trypomastigote forms that do not invade cells. They can transform into slender trypomastigotes and multiply in various body fluids. Some slender trypomastigotes can transform into non-dividing stumpy trypomastiogtes, which are ingested by the tse-tse fly when sucking blood from an infected mammalian host (Barrett et al., 2003) (Fig. 1B). Two T. brucei subspecies that are morphologically indistinguishable cause distinct diseases in humans: T. b. gambiense causes West African sleeping sickness and T. b. rhodesiense causes East African sleeping sickness. A third member of the complex, T. b. brucei, does not normally infect humans (Garcia et al., 2006). The nuclear genome of T. brucei, which has also been sequenced and is available online, is divided in three classes of chromosomes: 11 pairs of large chromosomes (1-6 Mb), 3-5 intermediate chromosomes (200-500 kb) and ~100 mini chromosomes (50-100 kb), present in multiple copies per haploid genome. The large chromosomes contain most of the genes, while the small chromosomes tend to carry genes involved in host-cell evasion by antigenic variation, including the VSG (Variant Surface Glycoprotein) genes (Berriman et al., 2005). The VSG genes are always located at an expression site that contains a number of Expression Site-Associated Genes (ESAGs) found in subtelomeres of the large and intermediate chromosomes. Current evidence indicates that only a single expression site is active at a given time, with the majority of VSG genes remaining silent; this selective activation results in antigenic variation. VSG switching replaces the single VSG isoform of one coat with another one and is responsible for the evasion of host immunity by African trypanosomes. The mechanism behind this gene switch is the major focus in trypanosome research (Pays, 2005; Horn and Barry, 2005). Leishmania spp.: Parasites belonging to the genus Leishmania are the etiological agents of leishmaniasis, which has a large spectrum of clinical forms (cutaneous, diffuse cutaneous, mucocutaneous and visceral). The forms of leishmaniasis are determined by parasite, host factors and immunoinflammatory responses, and may range from subclinical to disseminated infection, with varying presentations and outcome (Murray et al., 2005; Besteiro et al., 2006). Compared to trypanosomes, Leishmania spp. have a very simple developmental cycle. The extracellular flagellates and the proliferative promastigotes, which differentiate into the non-proliferative, infective metacyclic form, live in the sandfly midgut. Once metacyclics are inoculated by the insect vector into mammalian skin, the parasites are phagocytized and develop within the cells (inside the parasitophorus vacuole), where they differentiate into dividing, intracellular and non-flagellate amastigote forms capable of infecting new cells and a non-infected sandfly during blood feeding (Alexander and Russel, 1992) (Fig. 1C). The genomes of three

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Leishmania species (L. major, L. braziliensis, L. infantum) are highly conserved with regard to gene order and sequence; the genome of L. mexicana is currently being assembled. Old World Leishmania (L. donovani and L. major groups) have 36 chromosomal pairs, whereas New World species (L. mexicana and L. viania groups) have 34 or 35, with chromosomes 8+29 and 20+36 in the L. mexicana group and 20+34 in the L. braziliensis group being fused. The entire nuclear genome size is ~32.8 Mb, with more than 8,000 transcribed genes arranged in polycistronic clusters. Only 200 genes show a differential distribution between the three species; these genes may be related to host-pathogen interactions and parasite survival in macrophages. Unlike the trypanosomes, in Leishmania spp. the subtelomeric region is not the site for the expression of species-specific genes, and there are few multigene families near telomeres (Cano, 2001; Conte and Cano, 2005; Ivens et al., 2005, Horn and Barry, 2005).

(A)

Figure 1. (Continued).

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(B)

(C) Figure 1. Life cycle of Tri-tryps in invertebrate and vertebrate hosts. A) T. cruzi. B) T. brucei. C) Leishmania spp. Adapted from the animations available at http://apps.who .int/tdr /svc/diseases.

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The advent of complex ultrastructural and biochemical studies, the availability of genome sequences for the main Tri-tryps and the increasing number of post-genomic studies have revealed inter-specific similarities and helped to decipher the peculiarities of each species (Gull, 2001; Teixeira and Rocha, 2003; El-Sayed et al., 2005b). Some of the trypanosomatid peculiarities include: (i) complex, energy-consuming mitochondrial RNA editing, (ii) a unique mitochondrial DNA architecture known as the kinetoplast, (iii) mRNA maturation by trans-splicing, (iv) the arrangement of genes into giant polycistronic clusters, (v) unprecedented modifications of nucleotides, (vi) the compartmentalization of glycolysis, (vii) evasion of the host immune response by using a variable surface coat, and (viii) the ability to escape destruction by migrating out of phagocytic vacuoles (reviewed in Simpson et al., 2006). Trypanosoma brucei and probably L. braziliensis, but not T. cruzi or other Leishmania spp., also have RNA-mediate interference (RNAi) machinery (Ngo et al., 1998; Ivens et al., 2005). Another intriguing feature shared by Tri-tryps is variation in the DNA replication machinery and its control, as discussed below. 2.1. The Cell Cycle in Trypanosomatids The eukaryotic cell cycle is a progression of linked events that culminate in the segregation of identical genetic material into progeny. In trypanosomatids, the cell cycle also allows the correct duplication and segregation of single-copy and specialized organelles. Trypanosomatids contain a single Golgi complex (He et al., 2004) as well as a large mitochondrion that hosts a single kinetoplast formed by minicircle and maxicircle DNA molecules. A single flagellum connected through cytoskeletal filaments to the kinetoplast at its basal body, which serves as a microtubule organizing center, emerges from an invagination of the plasma membrane known as the flagellar pocket (Robinson and Gull, 1991; for good reviews see de Souza, 1999, 2002; Gull, 2003). The duplication and segregation of these single-copy organelles requires precise temporal and spatial duplication and segregation of the structures. These events trigger a very clear morphological pattern that changes according to the cell cycle. Detailed morphological descriptions of the cell cycle for T. brucei (Vaughan and Gull, 2008), T. cruzi (Elias et al., 2007) and L. mexicana (Wheeler et al., 2011) show that the morphological patterns found in each stage of the cell cycle are specific for each trypanosomatid. In addition, DNA replication in the nucleus and in the single mitochondrion (or kinetoplast) initiates nearly simultaneously, suggesting that the phases of DNA synthesis in the nucleus and mitochondrion are coordinately regulated (Pasion et al., 1994). In T. brucei procyclic forms, the presence of a duplicated basal body and of a small new flagellum indicates that the cell has

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started a new cell cycle and is initiating its DNA synthesis (S phase of the cell cycle). At the end of DNA replication, the cell reaches the G2 phase and kinetoplast segregation occurs. Hammarton et al. (2003) showed that there are also fundamental differences in cell cycle control among the life cycle forms of T. brucei and that key cell cycle checkpoints present in higher eukaryotes are absent from trypanosomes. In contrast to T. brucei, T. cruzi epimastigote cells reach the end of S phase with just one nucleus, one kinetoplast and one flagellum. During G2, the new flagellum emerges from the flagellar pocket and then the kinetoplast divides. After mitosis, the new flagellum reaches the size of the old one and cytokinesis is concluded (Elias et al., 2007; reviewed in Elias et al., 2009). The cell cycle of L. mexicana differs from other trypanosomatids, especially from L. amazonensis, T. cruzi and T. brucei. In this species, the new flagellum emerges just after the S phase, as in T. cruzi, but the kinetoplast divides after nuclear division (Wheeler et al., 2011; da Silva, Monteiro and Cano, personal communication). Morphological alterations during the cell cycle are not equal among trypanosomatids. This suggests that the mechanisms that determine precise division and segregation of the single-copy organelles may also be different. Indeed, in T. brucei, the growth of the new flagellum alongside the old flagellum plays a major role in morphogenesis. The new flagellum grows attached to the cell body, which is guided by the migration of a transmembrane mobile junction or flagella connector along the old flagellum. When growth or attachment of the new flagellum is perturbed, a range of morphogenetic defects occurs, including changes in cell shape, cell size and organelle segregation (reviewed in Vaughan, 2010). The new flagellum growth must be intimately linked with subpellicular microtubule dynamics to assemble the cytoskeleton of the new flagellum daughter cell. However, although the flagellum, basal body, golgi, nucleus and kinetoplast are closely associated with the single flagellum, the mechanism by which this assembly occurs remains unknown (reviewed in Ralston and Hill, 2008; Vaughan, 2010). In contrast to T. brucei, in T. cruzi, L. mexicana and L. amazonensis, the new flagellum does not grow attached to the old one or to the cell body (Elias et al., 2007; Wheeler et al., 2011; da Silva, Monteiro and Cano personal communication), indicating that different mechanisms were acquired during the evolution of trypanosomatids in order to guarantee the precise division of singlecopy organelles.

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Nuclear Dna Replication in Trypanosomatids

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1. Assembly of the Pre-Replication Complex at the Replication Origins Prior to DNA duplication, the start points of replication must be established. In 1963, Jacob and collaborators proposed the replicon model in which the replicator (the DNA element) must interact with the initiator (proteins) to facilitate DNA replication. The replicator element, now known as the origin of replication, is the DNA region where the pre-RC (initiator) binds during transition from the M phase to G1 phase of the cell cycle, thereby allowing the initiation of DNA replication during the S phase. This model has been extensively confirmed, although the pre-replication machineries differ considerably among prokaryotes and eukaryotes (reviewed by Sclafani and Holzen, 2007). To illustrate this, a brief description about the composition and function of the pre-RCs of metazoans, the Archaea and trypanosomatids is given below (Fig. 2). Metazoans: In metazoans, the pre-RC is formed during transition from the M phase to G1 phase, when CDK activity is low since high activity of this kinase prevents the pre-RC formation (Nguyen et al., 2001). The first component to reach the origin of replication is the ORC that consists of six subunits named according to their sizes in yeast: Orc1 (120 kDa), Orc2 (72 kDa), Orc3 (62 kDa), Orc4 (56 kDa), Orc5 (53 kDa) and Orc6 (50 kDa) (Bell and Stillman, 1992). Of these subunits, Orc1, Orc4 and Orc5 have the family domain AAA+ (ATPase activity involved in diverse cellular activities) that binds and hydrolyzes ATP. Orc subunits 1-5 also have a DNA binding domain (winged helix or WH). The ATPase activity and WH domain allow the ORC to select and bind to the origin of replication, respectively. Once ORC is bound to DNA, other two proteins, Cdc6 and Cdt1, must be recruited to load the MCM complex onto the replication origin to complete the pre-RC. Cdc6 also has the AAA+ ATPase domain that is important for its recruitment to the origin along with Cdt1 (da-Silva and Duncker, 2007). The main role of Cdt1 is to load the MCM complex to the origin. Cdt1 has three different regions: an N-terminal region that binds DNA, an intermediate region of interaction with Gemini (Cdt1 inhibitor) and a C-terminal region that binds the MCM complex (Lee et al., 2004). Through the C-terminal region Cdt1

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Figure 2. Pre-replication P c complexes ﴾pre-RCs﴿ from meetazoans, trypannosomatids and the Archaea. In I metazoans, thhe pre-RC is composed of the ORC O complex, Cdc6, Cdt and the MCM heteero-hexamer complex. In trypaanosomatids, thee pre-RC is com mposed of Orc1/Cdc6 6 and the MCM M hetero-hexameer complex. In Archaea, A the pre-RC consists of o Orc/Cdc6 and the MCM homo-hexamer h complex.

interacts directly with the MCM coomplex and, together t with Cdc6, carriess this p compoosed of ORC C, Cdc6, Cdt11 and complex to the originn. Thus, the pre-RC MCM, is completed. The MCM complex is a heterro-hexamer foormed by six different subbunits o Mcm7). Theese subunits innteract to form m a ring-shapedd structure thaat can (Mcm2 to encircle double- strannded DNA (ddsDNA). Thee MCM com mplex has hellicase hat melts the dsDNA d to proovide access during d replicattion in the S phase. p activity th All of thee six subunits have the sam me AAA+ ATP Pase domain that t is essentiaal for helicase activity a (da-Silva and Dunckker, 2007). The pre-RC p is form med in the M//G1 phase andd remains inacctive and bouund to DNA unttil the beginninng of the S phhase. Only at the G1/S trannsition is the MCM M complex activated by Cdk2 C and Cdc7 and other prroteins that stiimulate its hellicase o melt dsDNA A. DNA repliication starts after a the loading of RPA, DNA D activity to pol α/prim mase, RFC, PC CNA and pol δ (Truong andd Wu, 2011). Archaea: The Arcchaea are one of the three forms of lifee (the others being b Eukarya and Eubacterria). These orrganisms havee no nucleus or organelless and i the Archaea are resemble bacteria in shape. Howevver, the cellulaar functions in milar to eukarryotes than too prokaryotes.. In contrast to metazoanss, the more sim Archaea have no OR RC or Cdc6 and a Cdt1 prooteins, but haave an Orc1/Cdc6

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complex that recognizes and binds to replication origins and recruits the homohexamer MCM complex to the origin (de Koning et al., 2010). Trypanosomatids: Like the Archaea, trypanosomatids have no ORC, Cdc6 and Cdt1, but just a single protein (Orc1/Cdc6) that is similar to Orc1 and Cdc6. In T. brucei and T. cruzi, Orc1/Cdc6 is a component of the pre-RC and its absence results in parasites without a nucleus and a decrease in the number of cells in the G2 and M phases, indicating no DNA replication (Godoy et al., 2009). In L. major, Orc1/Cdc6 is found in the nucleus throughout the entire cell cycle (Kumar et al., 2008), in accordance with its involvement in DNA replication. Recently, one further ORC-like factor was identified in T. brucei (Dr. Richard McCulloch, personal communication). As in metazoans, trypanosomatids have the six subunits of the heterohexameric MCM complex. The pre-RC in trypanosomatids therefore probably consists of Orc1/Cdc6 and the MCM heterohexameric complex. As there is no Cdt1 in trypanosomatids, the mechanism by which the MCM complex is recruited to the origin remains unknown. We speculate that the MCM complex is recruited by Orc1/Cdc6, as in the Archaea, or by Mcm9, which has been proposed to be a “colicenser” of the MCM complex (Lutzmann and Michali, 2008) and is present in trypanosomatid genomes. Another possibility is that an unknown protein could recruit MCM to the replication origin, in a manner analogous to Cdt1. Further studies should confirm this hypothesis. 2. The Action of Toposiomerases at Replication Forks At the time of their replication and behind the replication fork, the two newly replicated DNA duplexes need be decatenated and separated from the parental DNA strands. In eukaryotes, an enzyme known as DNA topoisomerase type II is required to decatenate the chromosomal rings and promote chromosomal separation. DNA topoisomerases catalyze changes in the topological state of duplex DNA by introducing or removing supercoiling, knots or catenations in DNA molecules (Roca, 1998; Zuma et al., 2011). These enzymes are highly conserved and are active during replication, transcription, recombination and DNA repair (Wang, 1996). The first DNA topoisomerases discovered were Escherichia coli DNA topoisomerase I and E. coli DNA gyrase; animal and bacterial viruses also encode their own toposiomerases and this enzyme activity has been described in many eukaryotic cells (Roca, 1998). Because DNA topoisomerases play key roles in different cellular processes, it is important to define the physiological functions they are involved in and understand the molecular basis of their action.

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The topoisomerases can be divided into two types: type I (A and B) and type II. Both have been characterized in several trypanosomatids, as discussed below. DNA topoisomerase I is an abundant nuclear enzyme mostly associated with the nucleolus and chromatin regions undergoing transcription. In contrast, DNA topoisomerase II has a wider distribution in the chromosome and an important role in DNA replication and chromosome condensation (Muller, 1985). The following discussion will therefore concentrate on nuclear topoisomerase type II. The type II family of topoisomerases, which includes the eukaryotic enzyme, E. coli DNA gyrase, E. coli topoisomerase IV and archaic topoisomerase VI, is essential and highly conserved in all living organisms. In contrast to type I, type II DNA topoisomerases function as homodimeric molecules that are ATP-dependent (Roca, 1998). Eukaryotic topoisomerase II is one of the most abundant scaffold and nuclear matrix proteins (Earnshaw et al., 1985). This enzyme promotes the relaxation of positive and negative supercoils by a trans-esterification reaction involving nucleophilic attack at an active site tyrosine on a DNA phosphodiester bond; this action results in the formation of a covalent DNA 3' phosphotyrosyl linkage (Liu et al., 1980). During the reaction, each monomer remains bound at the 5' end of both DNA strands and the bonds between the enzyme and DNA conserve the energy of the phosphodiester bonds to facilitate rejoining after the "strand-passing" process or catalytic activity (reviewed in Douc-Rassy et al., 1988). Consistent with these properties, these enzymes do not require the exposure of single-strand regions to cause DNA relaxation (Champoux, 2001). In trypanosomatids, topoisomerase II activity is essential for nuclear and kinetoplast DNA replication and segregation. Topoisomerase II purified from T. cruzi and L. donovani has ATP-dependent and ATP-independent decatenating activities, although the ATP-independency activity may be artefactual (reviewed in Das et al., 2004). Leishmania donovani topoisomerase II (LdTOP2) occurs in the nucleus and kinetoplast, and T. cruzi topoisomerase II is also found in the nucleus (Fragoso et al., 1998; Das et al., 2004). Apart of the conserved C-terminal homodimerization domain, identified by modeling using yeast topoisomerase II data, LdTOP2 shows all of the features of the homodimeric model, including formation of the cavity that can accommodate the DNA double helix, the conservation of amino acid residues for Mg2+ binding and the presence of catalytic tyrosine (Sengupta et al., 2003; Das et al., 2006a). Most trypanosomatid topoisomerases II share high sequence identity and function almost exclusively as mitochondrial ezymes with no nuclear function. Recently, two nuclear type II topoisomerases (TbTOP2α and TbTOP2β) were described in T. brucei brucei. Phylogenetically, they share a common node with other nuclear topoisomerases from higher eukaryotes. TbTOP2α encodes an ATP-dependent topoisomerase that

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appears as a single band of ‫׽‬170 kDa on immunoblots and is located in the nucleus; RNA interference leads to pleomorphic nuclear abnormalities and early growth arrest, with no effect on kinetoplast DNA (kDNA). The role of TbTOP2β is unclear. Although transcribed in trypanosomes, TbTOP2β is not detected by βspecific antiserum and RNAi silencing results in no obvious phenotype (Kulikowicz and Shapiro, 2005). The potential of DNA topoisomerases as drug targets has expanded research interests in this field into pharmacology and clinical medicine (Roca, 1998). The identification of DNA topoisomerases as a promising drug target and recent studies showing that Kinetoplastida topoisomerases are markedly distinct from their human counterparts supports the belief that topoisomerase-based antiparasitic drugs have a role in future therapies (Das et al., 2006b). For example, selective targeting of leishmanial topoisomerase II and human topoisomerase II was shown to be achieved by some mitonafide analogs (Slunt et al., 1996, Das et al., 2006b). Other compounds, such as luteolin and quercetin, were found to specifically inhibit the kDNA topoisomerase II in L. donovani and T. brucei (TbTOP2mt), but not the nuclear topoisomerase II (Das et al., 2004).

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3. DNA Polymerases: The DNA Replication Catalysis DNA polymerases are essential enzymes for DNA replication in all living cells (Kornberg and Baker, 1992; Johnson and O'Donnell, 2005). All cellular organisms possess several DNA polymerases that operate during DNA chain elongation, recombination, replication and in diverse repair processes (Goodman and Tippin, 2000; Pavlov et al., 2006). Regardless of the range of processes DNA polymerases are involved in, they display several common features. The basic mechanism of nucleotide addition involves a pair of metal ions coordinated by carboxylated amino acid residues that are widely conserved among DNA and RNA polymerases (Oliveros et al., 1997; Steitz, 1998). The introduction of DNA cloning and sequencing techniques allowed the identification and isolation of many genes and putative sequences coding for DNA polymerases from phylogenetically diverse organisms. Based on the sequence similarities and homology, DNA polymerases can be further subdivided into seven families: A, B, C, D, X, Y, and RT. Family A is represented by replicative and repair polymerases that include the phage T7 DNA pol, mitochondrial DNA pol γ, E. coli DNA pol I, Thermus aquaticus pol I, and Bacillus stearothermophilus pol I. Family B includes the major eukaryotic DNA polymerases (α, δ and ε) and also DNA pol ζ. A hallmark of this family is a

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Figure 3. A putative evolutionary scenario for the origin of eukaryotic B family DNA polymerase. This scheme is based on the framework of the symbiotic scenario for the origin of eukaryotes in which the symbiosis of an archaeon with an α-proto-mitochondrion gave rise to the mitochondrion and triggered eukaryogenesis. Adapted from Tahirov et al. (2009).

remarkable accuracy during replication and, with the exception of DNA pol α and ζ, they also possess 3’-5’ proofreading activity. Family C is represented by the E. coli and other bacterial chromosomal replicative enzymes, such as DNA pol III from the α and ε subunits. Recent research has classified the C family polymerase as a subgroup of the X family. The polymerases in family D are not well characterized and include enzymes found in the Archaea. Family X includes the eukaryotic pol β, as well as other eukaryotic polymerases such as pol σ, pol λ, pol μ, and terminal deoxynucleotidyl transferase (TdT). The Y family members differ from other enzymes by their low fidelity on undamaged templates and their ability to replicate through damaged DNA, hence they are also known as translesion synthesis (TLS) polymerases. Finally, the reverse transcriptase family (RT family) includes examples from retroviruses and eukaryotic RNA-dependent DNA

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polymerases. The eukaryotic polymerases are usually restricted to telomerases (Burgers et al., 1998). According to Tahirov et al. (2009), the evolution of eukaryotic DNA polymerases seems to have involved previously unnoticed complex events. Their hypothesis is based on the fact that the archaeal ancestor of eukaryotes encoded three DNA polymerase, i.e., two B family polymerases and a D family of polymerases, all of which contributed to evolution of the eukaryotic replication machinery. These authors also suggested that the important Zn finger domain, present in most eukaryotic polymerases, might have been acquired from archaeal pol D by polymerases of family B. This gave rise to pol ε prior to or in the course of eukaryogenesis; this enzyme was subsequently captured by the ancestor of other eukaryotic B family polymerases (Fig. 3 summarizes the hypothesis proposed by Tahirov et al., 2009). 3.1. Trypanosomatid DNA Polymerases DNA synthesis of the two strands occurs by two different mechanisms, with one strand being synthesized continuously while the other is synthesized discontinuously. Several enzymes and protein factors are involved in these processes and DNA pol δ and pol ε play a major role in DNA synthesis. Although distinct DNA polymerases have been described in many eukaryote species (Burgers et al., 2001; Hübscher et al., 2002), the number of DNA polymerases in the Trypanosomatidae and their relationship with homologous enzymes are not well established. Early reports on the isolation and purification of trypanosomatid DNA polymerases indicated the existence of at least three DNA polymerase activities in extracts from T. cruzi (Rojas et al., 1992; Venegas et al., 2000). A few reports have also characterized the genes encoding for DNA pol α and pol β from T. brucei (Leegwater et al., 1991), T. cruzi (Venegas et al., 2000, 2009), Crithidia fasciculata (Torri and Englund, 1995) and L. donovani (Luton and Johnson, 1997); there are no descriptions of other DNA polymerases from these protozoa. In 2005, the genomes of the Tri-tryps were published and greatly improved our understanding of the genetic and evolutionary characteristics of these parasites (Ivens et al., 2005; El-Sayed et al., 2005a, b). The results also confirmed previous findings based on analysis of the T. brucei genome which showed that the polymerase β gene and its parolog sequence (polymerase β-PAK) occurred in tandem, although the function of this protein is not well characterized (Ivens et al., 2005). A search for eukaryotic DNA polymerase homologues in the Tri-tryps genome database yielded the information compiled in Tables 1 and 2. Here, trypanosomatid DNA polymerases are classified by their subcellular location,

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putative or known function and by the presence or absence of conserved domains. The following paragraphs summarize the major findings of the few reports on trypanosomatid DNA polymerases. Trypanosoma spp.: Dube et al. (1979) first identified a major DNA polymerase activity in T. brucei using chromatographic techniques. Subsequently, Chang et al. (1980) reported the presence of DNA pol α and pol β activities in T. brucei extracts subjected to sucrose density gradient analysis. In 1992, two DNA polymerases were partially purified from T. cruzi extracts using sequential chromatographic steps on DEAE-cellulose, phosphocellulose and DNA agarose (Rojas et al., 1992). Venegas et al. (2000) subsequently partially purified and biochemically characterized three DNA polymerase activities from T. cruzi. Curiously, one of these activities was classified as a potential new DNA polymerase. More recently, Lopes et al. (2008) proposed that DNA pol β and DNA pol β-PAK activities in T. cruzi are involved in the maintenance of kDNA. A year later, Rajão et al. (2009) described DNA pol κ activity as also being mitochondrial. A DNA pol η from T. cruzi (TcPolη) was cloned and characterized by de Moura et al. (2009). To date, this is the only report of a polymerase able to bypass cyclobutane pyrimidine dimers (CPD) in an error-free manner. It is also able to bypass other DNA lesions (8-oxoguanine, cisplatin, Apurinic/Apirimidinic sites) and is mainly involved in homologous recombination. TcPolη is the first polymerase found to be localized in the nucleus of T. cruzi if DNA polymerase β, β-PAK, and κ are considered to be mitochondrial (kinetoplast) polymerases (Lopes et al., 2008; Rajão et al., 2009). Leishmania spp.: Chang et al. (1980) reported one of the first studies on the presence of DNA pol β activity in extracts of amastigote and promastigote forms of L. mexicana. DNA pol A and B (β-like) from L. mexicana were also purified and characterized and found to be related to DNA synthesis associated with kDNA (Nolan and Rivera, 1991, 1992; Nolan, et al., 1992). Luton and Johnson (1997) cloned and performed a sequence analysis of the gene encoding a DNA pol α of L. donovani. Their results suggested that this enzyme was considerably different from the human homologue. Several years later, Taladriz et al. (2001) described a nuclear DNA pol β activity in L. infantum. This nuclear localization was similar to that of mammalian DNA pol β, but contrasted with the mitochondrial localization of DNA pol β in other trypanosomatids (Torri and Englund, 1992, 1995; Lopes et al., 2008). Table 1 summarizes the few trypanosomatid polymerases described to date, their putative functions and subcellular localization. As discussed above, the vast majority of these enzymes are mitochondrial polymerases.

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M. S. da Silva, R. C. V. da Silveira, A. M. Perez et al. Table 1. DNA polymerases from Trypanosomatidae protozoa

DNA polymerases (pol)

Putative function

Organism(GenBank Accession number)

Subcellular localization

Polymerization catalysis Leishmania donovani Mitochondrial Luton and Johnson, 1997 Unknown Crithidia fasciculata Holmes et al., Pol α unidentified 1984 Polymerization catalysis Trypanosoma brucei Mitochondrial Chang et al., Pol α (Tb927.10.15370) 1980 DNA binding, Leishmania infantum Pol β nucleotide binding, (AAF00495) Nuclear Taladriz et al., 2001 dRPase and Polymerization catalysis (BER pathway) 5'-dRP lyase activity Crithidia fasciculata Saxowsky et Pol β (BER pathway) (AAA68599) Mitochondrial al., 2002 Polymerization of DNA Trypanosoma cruzi Strand displacement (Tc00.104705350395 Pol β (long-patch) 5.20) Mitochondrial Lopes et al., Cleavage of the 5'-dRP 2008 (BER pathway) Polymerization of DNA Trypanosoma brucei Strand displacement (Tb927.5.2780) Pol β (long-patch) Cleavage Mitochondrial Saxowsky et al., 2003 of the 5'-dRP (BER) Trypanosoma cruzi Ivens et al., Pol β-PAK Unknown (Tc00.104705350395 Mitochondrial 2005; El-Sayed 3.59) et al., 2005 Trypanosoma brucei Ivens et al., Pol β-PAK Unknown (Tb927.5.2790) Mitochondrial 2005; El-Sayed et al., 2005a Leishmania major Ivens et al., Pol β-PAK Unknown (LmjF08.0900) Mitochondrial 2005; El-Sayed et al., 2005a Error-free bypass of cis- Trypanosoma cruzi Moura et al., Pol η syn cyclobutane (Tc00.104705351191 Nuclear 2009 pyrimidine dimers 1.120) (CPDs) Bypass of N2-adducted Trypanosoma cruzi Pol κ dG lesions (Tc00.104705350375 Mitochondrial Rajão et al., Extension of 5.10) 2009 mismatched primer termini Pol α

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Reference

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Table 2. Classification of the Tri-tryp nuclear DNA polymerases according to the Tri-tryp GeneDB DNA polymeras es Pol α

Pol α Pol β Pol δ

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Pol δ Pol δ

Pol δ

Putative domains

Organism (GenBank Acc. number)

catalytic subunit -exonuclease 3' to 5' T. brucei (Tb927.8.4880) -ribonuclease H-like T. cruzi -DNA- and dNTP-binding activities (Tc00.1047053508837.180) catalytic subunit L. major (LmjF16.1540) -ribonuclease H-like L. infantum (LinJ16_V31640) -exonuclease 3' to 5' -DNA-binding and dNTP binding L. brasiliensis activities (LbrM16_V2.1600) catalytic subunit L. infantum (AAF00495) catalytic subunit -ribonuclease H-like T. brucei (Tb927.2.1800) -exonuclease 3' to 5' T. cruzi (Tc00.1047053510259.6) -DNA-binding and dNTP binding activities subunit 2 T. brucei (Tb927.3.1130) -DNA polymerase subunit B T. cruzi (Tc00.1047053509455.70) catalytic subunit L. major (LmjF33.1690) -ribonuclease H-like L. infantum (LinJ33_V3.1790) -exonuclease 3' to 5' L. brasiliensis -DNA-binding and dNTP binding (LbrM33_V2.1960) activities subunit 2 L. major (LmjF25.1410) L. infantum (LinJ25_V3.1450) -DNA polymerase subunit B L. brasiliensis (LbrM25_V2.2060)

Although most trypanosomatid nuclear DNA polymerase are poorly described and/or characterized, gene sequences encoding for some of them, such as pol α, pol β and pol δ, have been identified in the public Tri-TrypDB database. Using this information for in silico analysis we identified functional domains, such as the catalytic domains, in each polymerase. Curiously, the percentage of identity/similarity between the trypanosomatid sequences and their mammalian counterparts was very low. In contrast, there was a high degree of conservation among the trypanosomatid’s polymerases used in this study (see Table 2 for details).

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4. Telomere Replication: The End-Replication Problem Is Solved Telomeres are DNA/protein complexes located at the extremities of linear chromosomes and are responsible for protecting chromosomes against degradation, recombination and fusion. Telomeres help to regulate telomeric DNA length and stability and are also essential for the completion of DNA replication during cell division. Telomeres are commonly associated with a number of other important processes such as the control of cell division, aging, transcription regulation, genome integrity and the maintenance of nuclear architecture (Dmitriev et al., 2003; Chan and Blackburn, 2004). Telomeric DNA is characterized by the presence of tandem repeats ending with a single strand 3´ G-rich protrusion known as the 3´ G-overhang, first demonstrated in Tetrahymena and Oxytrichia, and later in higher eukaryotes (Klobutcher et al., 1981). The repeats are highly conserved across a wide range of eukaryotic organisms and the 3´ G-overhang is the substrate for telomere elongation by telomerase and subsequent second strand synthesis by DNA pol α. The 3´ G-overhang regulates telomerase access to the telomeres and also protects telomeric DNA against the DNA repair machinery by adopting special structural conformations that prevent enzymes from binding and modifying the single strand. To date, two conformations have been described in a variety of organisms: t-loops, which also occur in T. brucei, and single strand G-quadruplexes (Gquartet or G4 DNA) (Griffith et al., 1999; Munoz-Jordan et al., 2001; Chan and Blackburn, 2004; Smogorzewska and de Lange, 2004; Teixeira and Gilson, 2005; Gilson and Geli, 2007). Single- and double-stranded telomeric regions contain binding sites for protein complexes responsible for telomeric maintenance, with increasing degrees of complexity in higher eukaryotes (Dmitriev et al., 2003; Lira et al., 2007b). These complexes are dynamic and vary according to the cell cycle phase, cell aging and external stimuli (Laroche et al., 2000; Stewart and Weinberg, 2002; Smogorzewska and de Lange, 2004). Telomeric proteins may interact directly with the DNA or with other proteins and their functions appear to be related to the telomeric region which they interact. Protein complexes that bind the 3´ Goverhang participate in telomerase recruiting and affect telomere elongation. On the other hand, protein complexes that bind the double-stranded region help to maintain the t-loop and regulate telomerase negatively (Dmitriev et al., 2003; Smogorzewska and de Lange, 2004; de Lange, 2005). Telomerase is a ribonucleoprotein first characterized in Tetrahymena thermophila (Greider and Blackburn, 1985) as an enzyme able to replicate the extreme ends of linear chromosomes and thus solve the “end-replication

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problem”. The requirement for a 3’ overhang in telomere elongation was demonstrated in Euplotes aediculatus around 11 years after the discovery of telomerase (Lingner and Cech, 1996). Without the presence of this enzyme, cells lose genetic material at their chromosomal ends in each replication cycle. In addition, cells would have to either stop dividing or continue losing genetic material until the deletion of essential genes caused the loss of homeostasis and consequent cell death (Harley et al., 1990, Counter et al., 1992). The essential components of human telomerase are the protein component, telomerase reverse transcriptase (TERT), or Est2 in yeasts, and the RNA component, telomerase RNA (TER), or Tlc1 in yeasts; both components are essential for proper telomere maintenance in vivo (Chiang et al., 2004). The limiting factor for telomerase activity is the presence of the RNA component while the assembly of telomerase components into a functional unit in human cell lines is dependent on the presence of other proteins, including the heat shock proteins hsp70 and hsp90 (Holt et al., 1999; Forsythe et al., 2001). Sequence analysis of the telomerase reverse transcriptase protein component from E. aediculatus, Saccharomyces cerevisiae, Schizosaccharomyces pombe and humans show an evolutionary relationship with non-LTR retrotransposons, but the definitive mechanism of telomere maintenance that first appeared in evolutionary history is still unclear (Eickbush, 1997; Nakamura et al., 1997; Nosek et al., 2006). The regulation of telomerase expression is very complex and depends on many unknown factors. So far, telomerase expression has been shown to be influenced by TER expression levels, expression of enzymatic components, subcellular localization, interactions with telomeric DNA, 3´ G-overhang size and interactions with telomeric and non-telomeric proteins (Collins, 2006; Lydall, 2009). In S. cerevisiae and vertebrates, the accumulation of functional telomerase depends on the association of TER with a series of ribonucleoproteins also involved in small nuclear RNA stabilization. These are the Sm heteroheptameric complex in yeasts and proteins that recognize the H/ACA motif in human TER (Mitchell et al., 1999; Seto et al., 1999; Collins, 2006). In Tetrahymena, on the other hand, functional telomerase acumulation is dependent on an association with the p65 protein, which specifically binds TER (Witkin and Collins, 2004). Many biological situations involve an increase in telomerase expression, including embryonic and pluripotent cells and most kinds of cancers (Blasco and Hahn, 2003; Phatak and Burger, 2007; Reichman et al., 1997; Savoysky et al., 1996; Sharma et al., 1995). In S. cerevisiae, increased telomerase activity has been detected during histone depletion, in certain phases of the cell cycle and

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during stress caused by osmotic and heat shock (Wyrick et al., 1999; Gasch et al., 2000; Taggart et al., 2002; Fisher et al., 2004; Longhese, 2008). Telomerase recruitment to the telomeres allows access of the enzyme to its substrate, the telomeric DNA, and is dependent on protein complexes that vary in several model organisms. In mammals, a six-protein complex known as shelterin consists of proteins that bind double-stranded (TRF1 and TRF2, TTAGG Repeat binding Factors 1 and 2) and single-stranded DNA (POT1, Protection of Telomeres 1) and proteins that bridge both complexes (RAP1, Repressor/Activator Protein 1; TPP1, TFRs and POT1 interacting factor; and TIN2, TRF1 Interacting Factor 2). Shelterin is responsible for telomere protection, t-loop maintenance and the regulation of telomerase access in a telomere lengthdependent manner. Protein complexes similar to shelterin also occur in yeasts and plants (Wellinger, 2009; Price et al., 2010), although in budding yeast, a minimal telomeric complex known as CST (CDC13/Stn1/Tin2) interacts with RPA and controls the DNA replication at telomeres by recruiting both telomerase and DNA pol α for C-strand synthesis (Gao et al., 2007). In addition, orthologues to DNA damage checkpoint kinases (Tel1 and Mec1) and components from the DNA repair machinery (i.e., KU70/80, Mre11 and RAD50) also partcipate in the recruitment of telomeric proteins for telomerase elongation (Longhese, 2008). On the other hand, protein complexes similar to the yeast minimal CST telomeric complex are also present in higher eukaryotes. Together, these findings indicate that the current concept on how telomerase gains access to short telomeres is evolving towards a balance between multiprotein complexes that share many similarities to the DNA damage response, including the use of numerous proteins shared with other DNA metabolism pathways (Collins, 2006; Gilson and Geli, 2007; Palm and de Lange, 2008; DeZwaan et al., 2009; Lydall, 2009; GiraudPanis et al., 2010). 4.1. Trypanosomatid Telomere Replication Trypanosomatid telomeres show the usual architecture found in other eukaryotes. They all possess the conserved 5’-TTAGGG-3’ repeat, although variations to this sequence occur in some Leishmania species (reviewed in Cano, 2001). In addition, telomeric and subtelomeric regions of trypanosomatids contain the modified base beta-D-glucosyl-hydroxymethyluracil (base J) and it has been postulated that this modified base participates in the regulation of gene expression in subtelomeric regions of T. cruzi and T. brucei (van Leeuwen et al., 1996, 1997; Ekanayake et al., 2007; Genest et al., 2007). Trypanosome subtelomeric regions show extensive architectural variations among species. The T. cruzi subtelomeric regions are variable and harbor a small number of hexameric repeats, a conserved 189 bp element, gp85/sialidase-like

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genes related to the parasite virulence and retrotransposons that may aid in transposition, duplication and recombinantion of nearby genes (Chiurillo et al., 1999; Freitas-Junior et al., 1999; Kim et al., 2005). The most extensively studied subtelomeric region in the Trypanosomatidae is from T. brucei and can be divided into four types that contain a variable number of copies of variant surface glycoprotein (VSG) genes known to be involved in parasite evasion of the mammalian immune system. The first two types are silent repositories of VSG copies in which the first harbors large arrays of VSG genes and pseudogenes (up to 1500 copies) flanked by 70 bp A+T rich repeat elements and ingi retrotransposons, and the second harbors intact single copies of VSGs and 70 bp elements proximal to the telomeres of around 100 minichromosomes. The third type of subtelomeric region, denominated bloodstream expression sites (BES), shows extensive architectural variations (Hertz-Fowler et al., 2008), and contains retrotransposons and retrotransposon hot spots, a 50 bp A+T rich repeat region, a strong RNA polymerase I promoter, a polycistronic region containing several expression-site-associated genes (ESAGs), a 70 bp repeat region and the telomere-proximal VSG gene. These sites can be transcriptionally active in the bloodstream form of the parasite and 15-20 of such structures may occur. However, only one of the copies is expressed at any given time and switching of active sites may occur. The fourth type of telomeric region is a metacyclic VSG (MVSG) that is active when the parasite is in the insect host. Around 25 of these structures may occur and they differ from the BES in the positioning of the promoter close to the VSG. The positioning of VSGs adjacent to the telomere facilitates antigen switching through increased rates of recombination involving duplicate gene conversion and reciprocal telomere translocation (Horn and Barry, 2005; Navarro et al., 2007). Subtelomeric regions have also been extensively investigated and tend to be relatively short in Leishmania. Although variations may occur in different species and even between chromosomes, they consist mainly of 100 bp repeats known as LCTAS (Leishmania conserved telomere-associated sequence) interspersed with CCCTAA repeats (Fu and Barker, 1998; Fu et al., 1998; Chiurillo et al., 2000; Conte and Cano, 2005). Leishmania subtelomeric regions may also contain a 781 bp non-repeated telomeric associated sequence (NRTAS), as well as developmentally transcribed noncoding RNA elements that also vary between species and individual chromosomes (Chiurillo and Ramirez, 2002; Dumas et al., 2006). Few telomeric proteins have been identified so far in trypanosomatids. T. brucei telomeres may contain functional homologues to the higher eukaryote proteins histone H3V, TRF, Tin2, Tpp1 and Rap1 (Lowell and Cross, 2004;

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Figure 4. Current view of the composition of L. amazonensis telomeric chromatin. A representation of the parasite´s chromosomal terminus showing the telomeric location of some of the proteins with known and hypothetical telomeric functions. The telomeric repeat (TTAGGG) and the position of the subtelomeric region (LCTAS, Leishmania conserved telomere associated sequence) in the chromosome are shown. Double-stranded telomeric proteins are LaTbp1, LaRbp38 and LaTRF (Lira et al., 2007a,c; da Silva et al., 2010). Proteins LaRbp38 and LaRpa-1 associate with the G-rich single-stranded telomeric DNA and with each other. New L. amazonensis telomeric components and their protein interactions (gray circle with an interrogation mark) are under characterization (Fernandez et al., 2004; Siqueira-Neto et al., 2007; Cano et al., personal communication). The telomerase complex is represented by LaTERT and LaTER (Cano et al., 1999; Giardini et al., 2006, 2011; Cano et al., personal communication). The diagram is not to scale.

Dreesen et al., 2005, 2007; Yang et al., 2009). T. cruzi, L. major and L. amazonensis have TRF orthologues and, together with L. tarentolae and T. brucei, have been shown to have telomerase activity (Cano et al., 1999; Munoz and Collins, 2004; da Silva et al., 2010; Giardini et al., 2011). The gene encoding the telomerase protein component (TERT) from four Leishmania species has been cloned and characterized. Apart from being one of the largest telomerases described so far, TERT shows species-specific modifications in some amino acid residues within the conserved telomerase motifs, and a specific motif T (Giardini et al., 2006), similar to trypanosome TERTs. The L. amazonesis enzyme was partially purified using complementary columns and further G-rich telomeric DNA affinity chromatography; the protein showed enzymatic features similar to those shared with the previously semi-purified T. cruzi enzyme (Cano et al., 1999; Munoz and Collins, 2004; Giardini et al., 2011). Currently, the best characterized trypanosomatid telomeric complex is that of L. amazonensis. This complex consists basically of the proteins LaRBP38 and LaRPA-1 that bind the 3´G-overhang, which are probably the functional homologues of yeast CDC13 and eukaryote POT1 since orthologues for both proteins are not found in the Tri-tryps genomes. LaTBP1, LaRBP38 and LaTRF are the proteins that bind the double-stranded telomeric region, and LaTRF shares structural and functional similarities to mammalian TRF1/TRF2 proteins

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(Fernandez et al., 2004; Lira et al., 2007a,b,c; Siqueira-Neto et al., 2007; da Silva et al., 2010). Figure 4 summarizes our knowledge of the composition of Leishmania telomeres.

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Assembly of Chromatin and DNA Replication: A Tight Event In each round of replication, the DNA, chromatin and its epigenetic status must be accurately transmitted to daughter cells. In other words, histones (including their post-translation modifications - PTM and variants) and DNA methylation must be faithfully replicated and also precisely localized on chromatin. As discussed above, Trypanosomatidae replication contains many peculiarities. Below, we summarize what is known about Trypanosomatidae chromatin, with particular emphasis on chromatin replication and the differences within this family compared with other eukaryotes. In contrast to other eukaryotes, trypanosomatid chromatin is organized in 10 nm fibers and chromosomes are not visible during mitosis (Vickerman and Preston, 1970; Solari, 1980). In T. cruzi, chromatin is more susceptible to nucleases when compared to other eukaryotes (Hecker et al., 1994). This finding is suggestive of a different chromatin structure that probably reflects the presence of divergent protein components in trypanosomatid chromatin (as discussed below). These differences in chromatin structure were also noted when different life cycle stages of T. cruzi were analyzed. For instance, in replicative forms, the densely packed chromatin is located preferentially at the nuclear periphery whereas in non-replicative forms the densely packed chromatin is dispersed over the nucleus (Elias et al., 2001). These patterns indicate that chromatin structure may contribute to differences in transcription and replication observed between the two life cycle stages. Chromatin consists of repeating units of nucleosomes. Four histones (H2A, H2B, H3 and H4) form the nucleosome core and a fifth histone (H1) interacts with DNA between each nucleosome to connect one nucleosome to another. These proteins are some of the most conserved proteins in eukaryote genomes, yet trypanosomatid histones differ greatly from eukaryote histones. Histone H1 is the most divergent histone as it lacks the N-terminus and globular domains of histone H1 from other eukaryotes. Curiously, functions unrelated to chromatin-binding have been attributed to Leishmania histone H1. The overexpression of histone H1

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genes not only delays progress of the Leishmania cell cycle and differentiation but also significantly reduces Leishmania infectivity in mice (Smirlis et al., 2006). In most eukaryotes, histone synthesis is dependent on DNA replication, mainly because of transcriptional regulation and the stabilization of histone transcript mRNA (Wu and Bonner, 1981; Stein and Stein, 1984). The levels of mammalian histone mRNAs increase 15 fold or more during the S phase. In trypanosomes, histone synthesis appears to be regulated in a different way. For example, in T. cruzi, the levels of histone mRNA increase only 2-4 fold in the Sphase, most probably because of mRNA stabilization since there is no increase in the rate of transcription in the S-, G1- and G2-phases (Recinos et al., 2001; Sabaj et al., 2001). Core histones are synthesized mainly during the S phase, as in other eukaryotes, but histone H1 is synthesized at low levels throughout the cell cycle (Sabaj et al., 1997). In Leishmania, the steady-state levels of core histones are independent of the cell cycle phase, although histone synthesis increases dramatically when cells enter the S phase. The regulation behind this phenomenon may be related to the fact that during the S phase histone mRNAs are co-localized with ribosome fractions whereas in other cell cycle phases they are unrelated to the ribosomal fraction (Soto et al., 2004). As with other eukaryotes, trypanasomatid histones are modified by methylation, phosphorylation and acetylation. Histone modification plays an essential role in many features of eukaryote cells, including replication, transcription and DNA repair. In general, histone acetylation is associated with an open chromatin and the activation of transcription (reviewed by Jenuwein and Allis, 2001). Distinct roles have been proposed for histone methylation, depending on the residue that is modified. For instance, tri-methylation of H4K4 correlates with transcriptional activity (Scheinder et al., 2004) whereas methylation at histone H3K9 is correlated with heterochromatin, transposons and silent genes (Gendrel et al., 2002). Currently, more than 30 residues are known to be post-translationally modified by acetylation, methylation and/or phosphorylation (Table 3), although the exact role of these modifications remains to be determined. Trypanosoma cruzi histones H3 and H2B are methylated, while H4 and H2A are mainly acetylated. The T. cruzi histone H4 modifications were identified by using mass spectrometry (da Cunha et al., 2006). In T. brucei, a combination of Edman degradation and mass spectrometry showed that H4 is largely modified whereas H2A and H2B contain few modifications (Janzen et al., 2006a; Mandava et al., 2007). Table 3 summarizes the currently recognized posttranslational modifications in trypanosomatid histones. Although there has been no systematic analysis of histone modifications in Leishmania, this parasite appears to contain modifications since commercial antibodies against acetylated

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Table 3. Histone post-translational modifications in trypanosomatids

T. cruzi

T. brucei

methylation

H2A A1

Histone types H2B H3 A1 K4, K32, K76

acetylation

A1, K4, K115, K4, K12, K16 K119, K120, K122, K125, K128 phosphorylation --Histone types H2A H2B methylation yes* yes* acetylation phosphorylation

H4 A1, K2, K17, K18, K53 K2, K4, K5, K10, K14

H1 --

--

--

--

H3 yes*

H4 A1, K18, R53 K4, K10, K14, K57 --

H1 --

S1, K23

yes*

yes*

yes*

--

--

--

--

S1 S12

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* observed by incorporation of acetylation and methylation precursors.

histone H3 are enriched at the 5' ends of polycistronic protein-coding gene clusters (Thomas et al., 2009). Some post-translational modifications are related to cell cycle phase and replication. During replication, the nucleosomes are displaced by movement of the replication fork so that it is still unclear how the epigenetic status is faithfully maintained, even in mammals. Two models have been proposed to explain this phenomenon. In the semi-conservative model, old H3-H4 and H2A-H2B dimers and their modifications are split between the daughter and parental DNA, thereby diluting the parental modification. In the random model, the dimers of H2A-H2B and tetramers of H3-H4 are randomly distributed in the daughter and parental DNA. In the latter model, the protein H1P (heterochromatin protein 1) may function as a “reader” for H3K9 methylation and stimulate the spreading of this modification through local chromatin (reviewed in Blomen and Boonstra, 2011). Histone H4 acetylation at K4 is a conserved event from yeasts to humans (Sobel et al., 1994, 1995) and, together with K12ac, is associated with histone deposition and cell cycle progression (Megee et al., 1990). In general, histone H4 is synthesized, acetylated in the cytosol at K4 and K12, and transported to the nucleus where it is incorporated into chromatin during the S phase and deacetylated afterwards. In trypanosomatids, the pathway involved is different. The K12 residue is absent in trypanosome histone H4 sequences, but H4K4 is

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present and acetylated in ~80% of histone H4 in T. cruzi and T. brucei (da Cunha et al., 2006; Janzen et al., 2006a). In contrast to all other eukaryotes, H4K4 is acetylated in the nucleus by a histone acetyltransferase (HAT3) (in agreement with this, no HAT is detected in the cytosol). Moreover, acetylation of H4K4 is apparently irreversible (Siegel et al., 2008). In T. cruzi, the level of H4K4 acetylation is lower in non-proliferative forms than in replicative forms, suggesting that H4K4ac may also be associated with the replicative status. In addition, in this species, there is also an increase in the acetylation of H4K10 and H4K14 after the S phase (Nardelli et al., 2009). An indirect way to understand the role of chromatin modifications in replication is to analyze the effect of depleting modified chromatin enzymes, such histone deacetylases, kinases and histone methyltransferases. These enzymes are found in trypanosome genomes but are poorly conserved. In T. brucei, histone deacetylases DAC1 and DAC3 are essential whereas DAC2 and DAC4 are not critical for viability. However, DAC4 delays progression of the G2/M cell phase (Ingram et al., 2002), suggesting that acetylation may be important for cell cycle progression. Two histone methyltransferases (DOT1A and DOT1B) were found in T. brucei. DOT1A dimethylates H3K76 while DOT1B trimethylates histone H3K76. Depletion of both enzymes (individually) causes defects in the cell cycle. RNAi for DOT1A results in a high proportion of cells with a haploid content suggesting that the mitotic checkpoint was perturbed (Janzen et al., 2006b). Whether these cell cycle defects are a direct effect of the methylation status of H3K76 requires more investigation. Another example of modifying enzymes that affect the cell cycle involves histone H3 phosphorylation. Trypanosoma brucei TLK1, a Tousled-like kinase homolog, interacts with chromatin assembly factor (ASF-1 and ASF2) and phosphorylates histone H3. RNAi for TLK1, Asf1A and Asf1b (single, double or triple depletions) increases the number of S phase cells, suggesting that these proteins may be required for S phase progression in this species (Li et al., 2007). Again, the effect of histone H3 phosphorylation on the S phase or mitosis has not been investigated. In addition to histone modifications, histone variants also have an important influence on chromatin structure and function. In general, the histone H2A variant, H2AX, is enriched at sites of DNA damage. H3.3, a histone H3 variant, is enriched at active transcriptional sites and its deposition is independent of replication. Interestingly, trypanosomatids lack this variant as well as the transcriptional regulation of coding genes. Variants have been reported for each histone in trypanosomes (Alsford and Horn, 2004; Siegel et al., 2009), but they have a different role than in eukaryotes. In T. brucei, the histone H2A variant, H2Az, interacts with an H2B variant and localizes at repetitive DNA (Lowell et

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al., 2005). Interestingly, CENH3, a histone H3 variant found in the centromers of eukaryotes was not found in trypanosomatids; instead, a histone H3 variant (H3v) was expressed and founded to be associated with telomeres and subtelomeric regions (Lowell and Cross, 2004). Histone H1 variants have also been found in Leishmania (Belli et al., 1999). There is increasing evidence linking histone variants and histone posttranslational modifications to putative transcription start and termination sites (Respuela et al., 2008; Siegel et al., 2009; Thomas et al., 2009). With regard to replication, it would be interesting if similar studies were done with the sequences of trypanosomatid replication origins in order to determine whether histone posttranslational modifications and histone variants are preferentially localized. Another epigenetic modification that is widely studied in eukaryotes is DNA methylation. Methylation involves cytosines that are located mainly in CpG islands and is an important mechanism of gene silencing. As with histone posttranslational modifications, the pattern of DNA methylation must also be accurately transmitted to the new DNA strand. After replication, only one strand of DNA contains the methylation pattern of the parental cell. DNMT-1 (DNA methyltransferase 1), together with PCNA and NP95 (a protein that binds to hemimethylated DNA), restores the methylation pattern (for a review see Blomen and Boonstra, 2011). Despite having a DNA methyltransferase that is more similar to prokaryotes than eukaryotes, T. brucei expresses DNMT1 in bloodstream and procyclic forms, both of which have 5-methylcytosine in their nuclear DNA (Militello et al., 2008). However, to date, neither a role for DNA methylation nor how this modification is propagated during the cell cycle has been proposed for trypanosomes. Nevertheless, 5-methylcytosine is present in T. cruzi nuclear DNA, indicating that this modification may be conserved in trypanosomes (Rojas and Galanti, 1990). In summary, trypanosomatids have divergent histones, enzymes that modify the chromatin status, and share few similarities with other eukaryotes. Despite considerable efforts to study the epigenetic status of trypanosomatids, more investigation is needed to understand the role of epigenetics in these protozoa.

Control of DNA Replication Significant advances in our understanding of cell cycle control in trypanosomatids have been made over the past few years, helped considerably by genomic studies and the use of RNAi in phenotype analysis. DNA replication in

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yeast and metazoans is a highly regulated process involving a number of licensing and replication factors and requires replicated DNA during every cell cycle and only once in each cell cycle. However, the mechanism involved in controlling nuclear DNA replication in trypanosomatids is only now beginning to be studied. DNA replication must be carefully coordinated with the events of the cell cycle to ensure the maintenance of a stable genome. Re-replication leads to the accumulation of single-stranded DNA and the formation of DNA double-strand breaks (DSBs) (Davidson et al., 2006; Liu et al., 2007) that activate checkpoint pathways (Cook, 2009). In yeast and metazoans, re-replication is avoided by assembling the pre-replication machinery in the G1 phase of the cell cycle, while the ability to license new replication origins is downregulated before entry into the S phase. Since ORC, Cdc6 and Cdt1 are required for loading MCM onto DNA but are not required for the continued MCM-DNA interaction (Donovan et al., 1997; Rowles et al., 1999; Edwards et al., 2002; Bowers et al., 2004), the downregulation of their expression/activity at the end of the G1 phase is an effective way of blocking DNA re-replication in the S and G2 phases after DNA replication has been initiated (Blow and Dutta, 2005). The precise mechanism involved in this control varies among organisms. In S. cerevisiae, Cdc6 is phosphorylated by CDK and degraded by proteolysis at the onset of the G1-S transition (Elsasser et al., 1999; Drury et al., 2000). Cdc6 transcription is also regulated during the cell cycle, reaching its maximum expression in late mitosis and early G1 (McInerny at al., 1997; Mendez and Stillman, 2003). As with Cdc6, Cdt1 levels may be controlled by CDK-dependent transcription and proteolysis (Nishitani et al., 2000). In addition, MCM are exported from the nucleus during the S phase, G2, and early mitosis, thus preventing the license of new origins in non-S-phase stages (Nguyen et al., 2000). Proteolytic degradation of Cdt1 is a conserved regulatory mechanism that extends to higher eukaryotes including Caenorhabditis elegans, Drosophila spp., Xenopus and mammals (Arias and Walter 2007; Cook, 2009). In mammalian cells, the Orc1 subunit is degraded in the S phase by a polyubiquitinylation reaction that is CDK-dependent (Mendez et al., 2002), while Cdc6 remains bound to chromatin during the cell cycle. Hence, the main way by which mammalian cells restrict replication to the S phase is through Cdt1 activity, which is regulated by geminin. Geminin binds and sequesters Cdt1 on chromatin during S and G2 thereby inhibiting Cdt1 association with Mcm2-7 and preventing pre-RC reassembly within one cell cycle (reviewed in Truong and Wu, 2011). As mentioned above, the trypanosomatid genome does not contain homologues to Cdt1. Consequently, the mechanism that normally uses Cdt1 to block re-replication may not be present in trypanosomatids. In addition, although

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Orc1/Cdc6 is upregulated in G1 in L. major (Kumar et al., 2008), this protein is expressed during the entire cell cycle of T. brucei and T. cruzi. Moreover, trypanosome Orc1/Cdc6 remains bound to chromatin during the cell cycle (Godoy et al., 2009), which strongly suggests that the restriction of replication to the S phase may be due to post-translational modifications in Orc1/Cdc6, changes in MCM expression and/or localization during the cell cycle, or the actions of other still unidentified proteins. In eukaryotes, a cell cycle checkpoint is activated when re-replication is induced in order to arrest the cell cycle or eliminate cells with over-replicated DNA; this elimination is generally by apoptosis or senescence. However, some cell cycle checkpoints are absent in particular life cycle stages of T. brucei (McKean, 2003), and some checkpoint regulators such as the spindle checkpoint protein BUB1, centromeric histone (CenH3), and Rho GTPases, are also apparently absent in trypanosomatids. In T. brucei, the inhibition of cytokinesis by a variety of mechanisms results in repeated DNA re-replication (McKean, 2003), which suggests that the timing of cell division plays an important role in ensuring that DNA is not re-replicated in a single cell cycle. Additionally, DNA re-replication occurs asynchronously in two-nuclei cells (Rothberg et al., 2006), suggesting that the nuclei are re-licensed for replication at different rates. The inhibition of DNA replication by silencing Orc1/Cdc6 expression in T. brucei cells results in cells with no nucleus (Godoy et al., 2009), demonstrating that these cells are also able to divide in the absence of two nuclei. Further studies are necessary to clarify the role of checkpoint mechanisms in the prevention of rereplication in trypanosomatids.

Nuclear Organization of DNA Replication There is substantial evidence that the events of DNA metabolism, such as RNA transcription, DNA replication and DNA repair, are compartmentalized within the nuclear space (Misteli, 2007). With regard to replication, these data support the notion that DNA passes through factories during its replication and that the assembly of macromolecular replication complexes on chromatin occurs via the recruitment of soluble units from a nucleoplasmic pool. Indeed, the formation of replication factories in yeast is a consequence of DNA replication itself (Kitamura et al., 2006). The nuclear organization of nuclear DNA replication in trypanosomatids is beginning to be unravelled. In T. cruzi and L. amazonensis, DNA replication occurs close to the nuclear periphery where chromosomes remain constrained during the S phase of the cell

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cycle (Elias et al., 2002, da Silva, Monteiro and Cano, personal communication). In T. cruzi, Orc1/Cdc6 and then PCNA are recruited in a sequential manner to a region close to the nuclear periphery as the cells go from the G1 to S phase (Calderano et al., 2011). This finding indicates that DNA replication foci in T. cruzi are acquired during self-organization as a consequence of DNA replication. In L. amazonensis, telomerase co-localizes with telomeres in the mid S phase and at the end of the S phase parasite telomeres are duplicated and found at the nuclear periphery (da Silva, Monteiro and Cano, personal communication). The molecular mechanisms that permit DNA replication close to the nuclear periphery in T. cruzi and Leishmania are unknown. One possibility is that the importation of nucleotides from the cytoplasm establishes this organization. Another hypothesis is that since these protozoa do not control transcription they should keep transcription and replication physically separate. At least in T. cruzi, transcription occurs throughout the nuclear space and has a main transcription center close to the nucleolus (Dossin and Schenkman, 2005). Recent work has shown that TcNUP-1, a component of the nuclear lamina, associates with chromosomal regions (Picchi et al., 2011). These associations could be involved in establishing the nuclear organization for DNA replication by anchoring T. cruzi chromosomes to the nuclear envelope. In L. donovani, distinct subnuclear foci (detected using PCNA as a marker) appear during the S phase (Kumar et al., 2009); the same appears to be true for L. amazonensis (da Silva, Monteiro and Cano, personal communication). These foci are the sites of active DNA replication, which confirms that DNA replication in trypanosomatids occurs in specific nuclear regions.

Conclusion In recent years, post-genomic studies have improved our knowledge of the biological aspects that make trypanosomatids such interesting organisms. Although considerable knowledge has been accumulated many aspects of nuclear DNA replication in these protozoa remain to be established. Current evidence indicates that the general aspects of DNA replication are well conserved in tripanosomatids, although Tri-tryps have simpler replication machinery when compared to higher eukaryotes. While lacking some conserved replication components, trypanosomatids have other components that share low sequence identity/similarity with their eukaryote counterparts and others that are probably exclusive to this group of parasites. In addition, some structural aspects that are common among eukaryotes are absent in trypanosomatids. For example, nuclear

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division in trypanosomatids does not involve the breakage of karyotheca and their cleavage furrow components have not been identified. Their chromosomes are too small to be seen by light microscopy, and chromosomal condensation and some phases of mitosis are not always observed during nuclear division. Consequently, the timing of nuclear replication in trypanosomatids is another aspect that deserves to be explored. For example, the L. mexicana nuclear S phase requires 2.9 h for completion (Wheeler et al., 2011) and is particularly long in comparison to T. brucei (1.51 h) (Woodward and Gull, 1990) and T. cruzi (2.4 h) (Elias et al., 2007). Curiously, the nuclear DNA content in L. mexicana (64 Mb) is similar to that of T. brucei (60 Mb) and is approximately half of that of T. cruzi (110 Mb), which raises many interesting questions. For example: Why does L. mexicana take twice as long to duplicate the same amount of nuclear DNA? Does this reflect differences in chromosome number and genome architecture? Is this phenomenon the result of lower processivity of the particular DNA replication machinery or is it caused by unknown epigenetic modifications? There is little information on the activity of Tri-tryps nuclear DNA polymerases. However, their genes are annotated in the genome databases and the putative encoded proteins show conserved structural and catalytic domains (see Table 2 of this chapter), which should make their characterization a lot easier than during the pre-genome era. Despite extensive telomere characterization in T. brucei and, to some extent, in L. amazonensis (Lira et al., 2007b), much remains to be done to understand telomere maintenance and replication in trypanosomatids. As for the components of the nuclear DNA replication machinery, differences in protein structure and telomeric complex composition may be useful in developing alternative therapies that target the pathogen with minimal side effects for the host. Trypanosomatid topoisomerase inhibitors have been extensively studied but no candidate able to target nuclear toposiomerase II, which is directly involved in DNA replication, has been identified (Das et al., 2004, 2006b). Further research should therefore be directed to elucidate the peculiarities of the Tri-tryps DNA replication machinery.

Acknowledgments The authors thank Dr. Stephen Hyslop for editing the English and critical reviewing the manuscript. This work was supported by grants from Fundação de Amparo a Pesquisa do Estado de São Paulo, Brazil (MINC, FAPESP 09/53336-0;

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MCE FAPESP 09/53693-8). MSS, AMP, SGC and JPM received fellowships from FAPESP. RCV received a fellowship from Conselho Nacional de Pesquisa (CNPq, Brazil).

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A access, 161, 172, 174 accounting, 143 acetic acid, 33 acetylation, 179, 180, 181, 201, 202, 205, 207 acid, 131, 136, 143 acrylate, 49 active site, 163, 175 adaptation, 152 adenine, 34, 51, 111 adhesion, 87, 153 adhesives, 48 aesthetic, 27 age, 51 allele, xiii, 110, 112, 117, 122, 123, 124, 125, 127, 128 ALS, 72 amino, xiii, 97, 110, 112, 120, 124, 136, 139, 140, 143, 163, 164, 176 amino acid, 97, 124, 136, 139, 140, 143, 163, 164, 176 amino acids, 136, 139, 140, 143 amsacrine, 50 ancestors, 208 anchoring, 186 anemia, ix, x, 2, 16, 17, 18, 19, 20, 21, 22, 23 animations, 156 annealing, 15, 111, 115, 125 ANOVA, 60, 63

antibiotic, 30, 101 antibody, 59, 96 anti-cancer, 81, 94 anticodon, 140, 144, 145 antigen, xv, 81, 93, 148, 175, 199, 204 aphasia, 54 apoptosis, x, xii, 26, 29, 30, 31, 35, 42, 43, 47, 52, 80, 93, 184 apraxia, 54 aqueous solutions, 112, 120 arithmetic, 145 arrest, ix, 2, 3, 21, 93, 100, 184 arrhythmia, 118 aspartate, 132 assessment, 49 astrocytes, 74 atmosphere, 30 atoms, xiv, 135 ATP, 10, 160, 163, 188, 189, 193, 194 atrophy, xi, 53, 55, 72, 75, 77 attachment, 158 autosomal recessive, 5 avoidance, 102 axon terminals, 57 B bacteria, 5, 111, 161, 198 bacteriophage, 106, 133 barriers, 7

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Index

base, xiii, xiv, 33, 42, 44, 51, 52, 110, 111, 114, 115, 118, 119, 122, 127, 128, 129, 135, 136, 137, 138, 139, 140, 143, 144, 145, 174, 193, 196, 209 base pair, xiii, 110, 118, 119, 122, 127, 144 biochemistry, 136 biomaterials, 27 biosynthesis, 208 Bis-GMA, x, 26, 27, 28, 29, 30, 31, 33, 34, 35, 37, 38, 39, 40, 41, 42, 43, 44, 45, 46, 47 bisphenol, x, 26, 27, 29, 37, 38, 39, 41, 42, 43, 44, 45 blood, 153, 154 bloodstream, x, 26, 27, 28, 45, 153, 175, 182 body fluid, 153 bonding, 29, 50 bonds, 163 bone marrow, 8 brain, 58, 60, 62, 64, 66, 71 Brazil, 147, 187 breast cancer, 12, 17, 20, 119 budding, 174, 192, 193

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C cabbage, 131 cancer, x, 3, 4, 8, 19, 20, 21, 26, 27, 45, 46, 50, 51, 89, 90, 93, 94, 95, 101, 102, 119, 126, 189 cancer cells, 50, 89, 90, 93, 94, 95, 101 carcinoma, xii, 80, 89 catalysis, 144, 168 catalytic activity, 163 C-C, 200 CCA, 90 CDC, 149 cell biology, 192 cell body, 158 cell culture, 30 cell cycle, ix, x, xiv, 1, 13, 20, 26, 29, 30, 31, 42, 44, 47, 49, 52, 90, 91, 93, 100, 104, 105, 107, 148, 149, 152, 157, 158, 159, 161, 172, 173, 178, 179, 180, 181, 182, 183, 184, 185, 189, 193, 197, 198, 199, 201, 202, 203, 205, 206, 207, 208, 210

cell death, 15, 56, 94, 172 cell division, 2, 103, 151, 171, 185, 209 cell line, xii, 20, 30, 52, 80, 89, 95, 101, 173 cell lines, 52, 89, 101, 173 cell size, 158 cellulose, 167 cerebral cortex, 66, 67 cervix, xii, 80, 89 Chagas disease, 194 challenges, 197 chemicals, x, 25, 27, 30, 97, 98, 99, 111 chemotherapy, 204 Chicago, 36, 59 chromatid, 6, 7, 16 chromatographic technique, 167 chromatography, 83, 121, 131, 177 chromosomal instability, 14 chromosome, 7, 16, 46, 72, 73, 75, 101, 162, 176, 187, 191, 192, 193, 204 classes, 142, 154, 208 classification, 62, 75, 153 cleavage, 13, 14, 186 clinical syndrome, 54 cloning, 104, 165, 196 clustering, 65, 66, 71 clusters, xi, 54, 57, 61, 65, 66, 67, 155, 157, 179 CO2, 30 coding, ix, 4, 118, 136, 137, 141, 165, 179, 182 codon, 139, 140, 141, 142, 143, 145 color, 96, 128 commercial, 29, 30, 179 communication, 176, 185 complexity, 149, 172 composites, 26, 27, 45, 50 composition, 159, 176, 177, 187 compounds, 28, 45, 47, 49, 86, 102, 120, 121, 130, 164 computer, 33 condensation, 162, 186 conflict, 48 conflict of interest, 48 connectivity, 73 consensus, 73, 75, 139, 144, 210

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Index consent, 58 conservation, 151, 163, 171 contingency, 66, 193 contour, 35 cooperation, 48 coordination, 14, 113, 120 corpus callosum, 58 correlation coefficient, 62 cortex, 55, 57, 59, 60, 61, 62, 63, 64, 65, 68, 69, 71 cortical pathway, 67, 70 corticobasal degeneration, 60, 71 cost, xiii, 110, 127, 130 crossing over, 17, 23 crystal structure, 120 CST, 174, 196, 205, 210 CTA, 139 culture, 31, 35, 107 cycles, 125 cyclins, 149 cytokinesis, 158, 184, 201, 204, 205 cytometry, 35, 37, 43, 90 cytoplasm, 56, 186 cytosine, 115, 126 cytoskeleton, 158 cytotoxicity, 45, 48, 49, 50, 52, 101 Czech Republic, 33 D damages, 27 database, 167, 171 defects, 8, 14, 158, 181 deficiency, 12 degenerate, 9, 136 degradation, 27, 51, 171, 179, 184 dementia, 54, 56, 58, 63, 66, 69, 71, 72, 73, 74, 75, 77 dental restorations, xi, 26, 45 dentin, 27, 45, 48 deoxyribose, 204, 206 dephosphorylation, 121, 132 deposition, 180, 182, 207 depression, 114 derivatives, 51, 104, 105, 113, 144

181

desorption, 120, 130, 131 destruction, 157, 194 detectable, 115 detection, ix, xiii, 46, 51, 100, 101, 110, 111, 112, 115, 116, 117, 118, 119, 121, 126, 127, 128, 130, 131, 133, 195 digestion, 131 dimethacrylate, x, 26, 27, 29, 37, 38, 39, 41, 42, 43, 44, 49 diploid, 101 discretization, 143 discrimination, 128 diseases, xiii, 77, 110, 111, 152, 153, 156, 210 disorder, 5, 19, 54, 55 displacement, 169 dissociation, 106, 111, 112, 113, 114 distribution, xi, xv, 54, 55, 57, 58, 61, 62, 65, 66, 67, 68, 69, 71, 73, 90, 148, 155, 162, 199 diversity, 133, 207 DNA breakage, 33 DNA damage, ix, x, xv, 1, 3, 8, 10, 12, 15, 17, 18, 19, 22, 23, 25, 26, 27, 31, 33, 34, 38, 40, 41, 44, 46, 47, 48, 49, 50, 51, 52, 148, 174, 182, 191, 200, 201, 202 DNA lesions, x, 2, 168 DNA ligase, 95 DNA polymerase, ix, xii, xv, 3, 80, 81, 84, 85, 86, 102, 103, 104, 105, 106, 130, 138, 139, 144, 145, 148, 150, 151, 164, 165, 166, 167, 168, 169, 170, 171, 187, 190, 192, 193, 195, 197, 199, 200, 201, 203, 204, 205, 208, 209 DNA repair, x, xi, 2, 3, 4, 6, 8, 9, 13, 15, 16, 17, 19, 20, 21, 22, 23, 26, 29, 34, 40, 41, 46, 47, 52, 162, 172, 174, 179, 185, 190, 203 DNA sequencing, 117, 119, 126 DNA strand breaks, 46 double helix, 163, 202 double strand breaks, ix, 1 Drosophila, 184, 190 drug discovery, 128 drug metabolism, 117, 118 drugs, xii, 27, 80, 102, 111, 164

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E egg, 98, 193, 195, 202 electric field, 33, 35 electrophoresis, x, xiii, 26, 32, 33, 34, 39, 46, 49, 50, 51, 93, 94, 110, 111, 122, 126, 127, 128, 129, 130, 132, 133 electroporation, 99 elongation, 19, 81, 98, 150, 164, 172, 174, 202 elucidation, 103 e-mail, 25 encoding, 5, 20, 139, 153, 167, 170, 171, 176 endonuclease, xi, 7, 10, 13, 15, 17, 18, 23, 26, 33, 42 endothelial cells, 45 energy, 132, 157, 163 energy transfer, 132 enlargement, 70 environment, 144 environmental change, 196 enzymatic activity, 117, 118 enzyme, xv, 13, 14, 34, 40, 42, 81, 83, 86, 87, 88, 97, 102, 144, 148, 162, 166, 170, 172, 174, 177 enzymes, xi, xii, 4, 5, 13, 15, 26, 33, 40, 46, 52, 80, 84, 85, 86, 104, 145, 150, 162, 163, 164, 166, 171, 172, 181, 183, 190, 200, 206 EPA, 48 epigenetic modification, 182, 187 epigenetics, 183 equilibrium, 33 equipment, xiii, 110, 126 ester, 27, 31 ethanol, 31 eukaryote, 166, 176, 177, 178, 179, 186, 199 eukaryotic, xii, 18, 80, 82, 98, 102, 103, 104, 149, 151, 152, 157, 162, 165, 166, 167, 172, 188, 199, 202, 208, 209 eukaryotic cell, 98, 102, 157, 162, 188, 209 eukaryotic DNA polymerase, xii, 80, 103, 104, 166, 167 evidence, 7, 29, 47, 72, 73, 153, 154, 182, 185, 186

evolution, 137, 144, 145, 151, 159, 166, 207, 209 excision, 34, 51, 52, 204 excitation, 31, 33 exonuclease, 138, 150, 170, 208 exploitation, 197 exposure, xii, 40, 41, 42, 45, 50, 80, 163 extraction, 95 extracts, 11, 91, 167, 168, 193, 195, 197, 202 F factories, xv, 98, 99, 101, 148, 185 families, xv, 56, 70, 82, 138, 148, 153, 155, 165, 199 family members, 153, 166 Fanconi anemia (FA), x, 2 fatty acids, 103 fertility, 5 fever, 203 fibers, 177 fibroblasts, 29, 30, 37, 38, 39, 40, 41, 43, 44, 45, 50, 105, 197 fidelity, 2, 102, 143, 149, 166 filament, 6, 7, 57, 72 fish, xii, 80, 86 fission, 21, 202, 203 fitness, 139 flagellum, 157, 158, 205, 209 flight, 120, 130, 131 fluorescence, 33, 99, 121, 128, 132 food, 28, 45 formaldehyde, 30, 96 formation, 5, 11, 23, 71, 93, 98, 114, 115, 120, 144, 159, 163, 183, 185 fragile site, 16 fragments, 56, 111, 112, 115, 118, 127, 128, 130 frameshift mutation, 137, 138, 144 frontal cortex, 59, 61, 64, 65 frontal lobe, 55, 58, 64 Frontotemporal lobar degeneration (FTLD), xi, 53, 54 fungus, xii, 80, 83, 101 fusion, 46, 138, 171

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G gel, ix, x, xiii, 26, 32, 34, 39, 46, 50, 51, 83, 89, 94, 110, 111, 112, 113, 115, 117, 118, 119, 122, 126, 127, 128, 129, 130, 132, 133 gene expression, 49, 104, 126, 129, 152, 174, 188, 203, 204, 208, 209, 210 gene silencing, 182 genes, 4, 5, 8, 13, 21, 46, 136, 146, 151, 153, 154, 155, 157, 165, 167, 172, 175, 178, 179, 182, 187, 192, 193, 199, 203, 205, 206, 210 genetic background, 151 genetic code, xiv, 135, 136, 137, 139, 140, 143, 144, 145 genetic defect, 55 genetic disease, 22, 27 genetic diversity, 153 genetic information, 126 genetic linkage, xii, 75, 109 genetics, 74, 76, 196 genome, 5, 7, 16, 17, 19, 20, 23, 111, 128, 144, 149, 153, 154, 155, 157, 167, 171, 183, 184, 187, 194, 196, 198, 205, 210 genomic instability, 2, 4, 6, 8, 22 genomic stability, x, 2, 4, 6 genomics, 194, 197 genotype, xiii, 110, 112, 117, 122, 124, 125 genotyping, 111, 122, 123, 124, 125, 127, 129 genus, 154 germ cells, 28 Germany, 30, 32, 90 gingival, x, 26, 29, 30, 37, 38, 39, 40, 41, 43, 44, 45, 50 glutamine, 30 glycol, 27, 49, 103 glycolysis, 157 grants, 187 growth, xii, 5, 14, 31, 80, 89, 93, 94, 95, 99, 158, 163, 201 growth arrest, 94, 164 GTPases, 184 guanine, 115 guidelines, 50, 60

183 H

haploid, 154, 181 harbors, 175 healing, 131 heat shock protein, 48, 173 heterochromatin, 150, 179, 180 heterogeneity, 55, 72, 76 heterozygote, 117, 124, 125 hippocampus, 56, 57, 59, 60, 63, 64, 66, 67 histology, 55 histone, 21, 173, 176, 178, 179, 180, 181, 182, 184, 191, 196, 198, 200, 201, 202, 205, 207, 208, 210 histone deacetylase, 181 histones, 177, 178, 179, 183, 188, 207 history, 173 HIV, 86 homeostasis, 172 homologous recombination, x, 2, 3, 6, 8, 11, 14, 19, 21, 23, 168, 201 homozygote, 117, 124, 125 host, 152, 153, 154, 157, 175, 187, 190, 196 hot spots, 21, 175 hotspots, 144 human, x, xii, xiv, 3, 12, 13, 19, 20, 22, 23, 26, 27, 29, 37, 38, 39, 40, 41, 43, 44, 45, 48, 49, 50, 51, 52, 71, 80, 81, 86, 89, 90, 93, 94, 95, 97, 101, 104, 105, 106, 109, 111, 118, 119, 128, 147, 151, 164, 170, 173, 188, 192, 195, 197, 201, 202, 204, 206, 208 human genome, xii, 81, 97, 109, 118 human immunodeficiency virus, 86 humidity, 30 hybrid, 210 hybridization, 128 hydrogen, 31, 34, 40, 111, 114 hydrogen bonds, 111, 114 hydrogen peroxide, 31, 34, 40 hydrolysis, 189 hydroxyethyl methacrylate, x, 26, 27, 29, 37, 38, 39, 41, 42, 43, 44, 52 hypothesis, 10, 12, 69, 70, 144, 145, 162, 166, 186

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I ideal, 2 identification, 15, 61, 112, 125, 127, 164, 165, 204 identity, 9, 163, 171, 186 IFN, 210 image analysis, 33 immobilization, 132 immune response, 157 immune system, 5, 175 immunity, 154 immunofluorescence, 96, 99 immunohistochemistry, 55, 59, 74 immunoprecipitation, 9 immunoreactivity, 56, 70 in vitro, 11, 14, 46, 50, 93, 95, 98, 101, 102, 103, 122, 132, 144, 145, 195 in vivo, 6, 14, 45, 50, 93, 94, 98, 101, 102, 106, 122, 132, 173, 190, 200, 207 incidence, 3 incubator, 30, 31, 40 Independence, 23 individuals, xiv, 12, 117, 123, 124, 125, 135 induction, 48, 87, 92 industry, 28, 48 infection, 154, 201 information processing, 192 inhibition, xii, 80, 83, 87, 88, 93, 94, 97, 98, 99, 100, 101, 102, 122, 184, 199 inhibitor, ix, xii, 80, 81, 82, 83, 87, 97, 99, 101, 103, 105, 106, 160, 211 initiation, xiv, 3, 18, 97, 98, 104, 148, 149, 159, 202 injuries, 45 insects, 137 integration, 21 integrity, 2, 20, 23, 149, 171 interface, 10 interference, 46, 157, 163, 188 interstrand crosslinks, ix, 1 intervention, 204 iodine, 31, 35, 36, 43 ionization, 120, 121, 130, 131 ions, 120, 122

Iowa, 76 ipsilateral, 74 irradiation, 12, 32 islands, 182 isolation, 165, 167 J Japan, 33, 79, 83, 102, 109, 127 Jordan, 172, 202 K kinase activity, 122 kinetics, 40, 125 knots, 162 KOH, 34 L laminar, xi, 54, 57, 61, 67, 68, 69 landscape, 210 latency, x, 26 lattices, 144 lead, ix, 1, 4, 15, 45, 94 leishmaniasis, xiv, 147, 154 lesions, ix, xi, 1, 2, 3, 54, 55, 59, 61, 62, 64, 65, 66, 67, 71, 73, 74, 169, 205 leucine, 93 leukemia, 51, 89 life cycle, 152, 158, 178, 184, 194 ligand, 120, 121, 127 light, 9, 13, 15, 32, 33, 35, 82, 101, 103, 186 linear molecules, 151 Lion, 131 localization, 16, 96, 168, 170, 173, 184, 193, 195, 196, 199 loci, 16 locus, 188 lung cancer, 89 lupus, 16 lymphocytes, 48, 49 lysine, 198 lysis, 32, 34

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Index

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M machinery, 106, 149, 150, 151, 157, 166, 172, 174, 183, 186, 187, 196 macromolecules, xiv, 135, 137, 143 macrophages, 155, 188 macular degeneration, 51 magnitude, 45, 68 majority, 56, 57, 67, 154, 171 mammalian cells, x, 2, 48, 49, 52, 98, 101, 153, 184, 200 mammals, 3, 9, 149, 174, 179, 184 management, 22 manipulation, 208 mapping, 104, 130, 139 mass, 83, 120, 121, 130, 131, 179 mass spectrometry, 120, 130, 131, 179 materials, xi, 26, 28, 45, 47, 48, 49 matrix, 120, 131, 141, 142, 143, 163 matter, 61 MCP, 211 measurements, 37, 43 medical, x, xiii, 25, 110, 111, 126, 127, 152 medicine, 27, 28, 164 melt, 161 melting, 30, 87, 114 melting temperature, 87, 114 Metabolic, 83 metabolism, 152, 174, 185 metabolites, 93 metal ions, 164 methacrylate composite, x, 26 methacrylates, ix, 26, 27, 28, 29, 37, 38, 39, 41, 42, 43, 44, 47 methodology, 111, 207 methylation, 126, 133, 177, 179, 180, 181, 182, 196, 205 Mg2+, 163 mice, 14, 94, 101, 178 microinjection, 99 micrometer, 61 microorganisms, 103 microscope, 32, 33 microscopy, 186

185

migration, 6, 10, 18, 32, 45, 112, 115, 118, 119, 122, 158 Ministry of Education, 102 mitochondria, 137, 145, 205 mitochondrial DNA, 157, 165, 200, 203, 206, 209 mitosis, 16, 149, 158, 177, 181, 184, 186 model system, 14, 73 modelling, 144 models, 143, 145, 179 modifications, 32, 40, 42, 47, 157, 176, 177, 179, 180, 181, 182, 184, 189, 191, 198, 201 molds, 34 molecular biology, 137 molecular weight, 35 molecules, xii, 80, 98, 99, 105, 107, 153, 157, 162, 163, 199 monoclonal antibody, 96 monomers, x, 26, 27, 28, 29, 43, 44, 45, 47, 48 Moon, 105 morphogenesis, 158, 197, 209 morphology, 192 motif, 5, 11, 97, 173, 177 motor neuron disease, 54, 57, 75, 77 mRNA, 157, 178, 208 mutagen, 49 mutagenesis, 4, 29, 48, 102 mutant, xiii, 11, 56, 101, 110, 111, 115 mutation, ix, x, xi, 4, 17, 25, 26, 44, 54, 55, 56, 57, 58, 59, 63, 65, 66, 67, 68, 69, 70, 72, 75, 76, 112, 117, 118, 119, 123, 124, 127, 130, 141 mutations, ix, x, xi, xii, xiv, 2, 4, 12, 14, 20, 26, 27, 28, 54, 55, 56, 64, 70, 73, 74, 75, 109, 111, 112, 115, 116, 118, 119, 126, 127, 128, 130, 135, 137, 138, 139, 140 mycelium, 83 myosin, 82, 103 N NaCl, 32, 34 nanometers, xiv, 135 natural compound, 83

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186

Index

natural science, xiv, 136 necrosis, 35 neocortex, 56, 64 neurodegeneration, 71 neurodegenerative diseases, 56, 74 neurodegenerative disorders, 57, 71 neurofibrillary tangles, 56, 58, 71 neuronal cytoplasmic inclusions (NCI), xi, 54, 55, 68 neurons, 69, 71, 74, 107 neutral, x, 26, 33, 34, 38, 39, 46, 120, 133, 139, 140, 143 next generation, 28 NMR, 83 normal distribution, 36 nuclear genome, 153, 154, 155 nuclear membrane, 150 nuclei, 96, 97, 151, 185, 199, 210 nucleic acid, 87, 138, 145 nucleolus, 162, 186 nucleosome, 178, 210 nucleotide sequencing, 128 nucleotides, xiv, 2, 6, 81, 129, 135, 136, 138, 143, 157, 186, 206 nucleus, xiv, 56, 95, 96, 97, 98, 99, 101, 148, 149, 158, 159, 161, 163, 168, 178, 181, 184, 185, 192, 194, 199 O OH, 123, 126 Okazaki fragment, 151 oncogenes, 46 optimization, 99, 143 oral cavity, x, 26, 27, 45 organ, 27 organelles, xiv, 148, 157, 158, 161 organism, 27, 44 organs, x, 26 Origen Recognition Complex (ORC), xiv, 148 overlap, 72 oxidative damage, 33, 40, 46, 192 oxidative stress, 48, 49 oxygen, 50, 120

P p53, 7, 21, 209 PAA, 30 pairing, 119, 144 parallel, xi, 17, 33, 54, 57, 59, 61, 65, 66, 67, 194 parasite, 152, 153, 154, 175, 176, 179, 185, 190, 191, 193, 195, 196, 197, 198, 199, 202, 207, 210 parasites, xiv, 147, 152, 153, 154, 161, 167, 186, 191, 195, 197, 198 pathogenesis, 71, 73 pathology, 28, 55, 60, 64, 67, 70, 71, 72, 73, 74, 75 pathways, ix, xi, 2, 8, 9, 11, 13, 17, 21, 52, 54, 57, 70, 151, 174, 183 PCR, xiii, 110, 111, 112, 114, 115, 116, 117, 118, 119, 122, 123, 124, 125, 126, 127 penetrance, 20 penicillin, 30 peptide, 121, 132, 138, 139 peptides, 132 periodicity, 57, 66 peripheral blood, 48 permission, 117, 124 permit, 119, 120, 125, 127, 185 personal communication, 158, 159, 161, 176, 185 pH, 32, 33, 34, 46, 112, 113, 120, 121, 131, 133 phage, 165 pharmaceutical, 102, 111 pharmacology, 164 phenocopy, 14 phenotype, 45, 72, 74, 75, 98, 118, 164, 183 phosphate, xiii, 30, 47, 59, 110, 112, 120, 122, 126, 129, 130, 131, 132, 133, 204, 206 phosphatidylcholine, 131 phosphatidylserine, 35 phospholipids, 120 phosphorylation, 11, 12, 120, 121, 132, 133, 179, 180, 181 phylogenetic tree, 142 physicians, xiii, 110, 126

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Index physics, 144 pia mater, xi, 54, 57, 59, 61, 62, 65, 67, 68, 69 plants, 174 plaque, 71 plasma membrane, 157 plasmid, 31, 32, 35, 37, 38, 46, 49, 98 platform, 7 PM, 190 point mutation, 28 Poland, 25, 30, 32 policy, 48 polyacrylamide, xiii, 110, 111, 112, 117, 121, 124, 129, 130, 132, 133 polyacrylamide gel electrophoresis (PAGE), xiii, 110, 111 polymer, x, 25, 27 polymerase, xiii, xv, 2, 18, 81, 86, 106, 110, 111, 128, 138, 144, 148, 150, 166, 167, 168, 171, 175, 192, 196, 200, 205, 206, 208, 209 polymerase chain reaction, xiii, 110, 111, 128 polymerization, x, xv, 25, 27, 81, 88, 138, 148, 150 polymers, x, 26, 27 polymorphism, 112, 126, 128, 129 polymorphisms, 128, 129 polynucleotide kinase, 86 population, 36, 43, 111, 153, 208 population structure, 153, 208 poverty, 188 preparation, 31, 47 prevention, 7, 185 priming, 104, 114 principles, 115, 117 probability, 45, 141 probe, xii, 80, 81, 82, 120, 128 progressive supranuclear palsy, 60 prokaryotes, 159, 161, 182 proliferating cell nuclear antigen (PCNA), xv, 93, 148 proliferation, xii, 28, 80, 89, 104, 105, 204, 211 promoter, 175 prophylaxis, 152 proportionality, 141

187

protection, 20, 29, 98, 174 protein components, 178 protein structure, 187 protein synthesis, 92, 93 proteinase, 30, 35 proteins, 4, 7, 8, 9, 10, 12, 13, 14, 15, 19, 22, 46, 91, 92, 95, 98, 105, 106, 118, 122, 127, 131, 132, 133, 136, 137, 138, 139, 146, 151, 153, 159, 160, 161, 163, 172, 173, 174, 176, 177, 178, 181, 184, 187, 192, 195, 196, 199, 200, 203, 207, 210 proteolysis, 183, 193, 202 pulp, x, 26, 27, 45, 48, 52 purification, 30, 121, 125, 167, 196, 203 purines, 34, 139 pyrimidine, xiv, 113, 136, 168, 169 Q quadratic curve, 62 quantification, 50, 131 quercetin, 164 R radiation, 51 radicals, 50 radius, 37 reactions, 122, 125, 130 reactive oxygen, 52 reactivity, 66 reading, xiv, 135, 137, 138, 139, 199 reagents, 132 real numbers, 143 recognition, 22, 129, 130, 140, 145, 188, 193, 195, 202, 206 recombination, x, 2, 3, 4, 5, 6, 8, 11, 14, 17, 19, 20, 21, 23, 162, 164, 168, 171, 175, 201, 204, 205 recommendations, 75 recruiting, 172, 174 redundancy, 15 regression, 10, 62 relaxation, 32, 37, 46, 163 relevance, 14

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188

Index

repair, ix, x, xv, 1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 20, 22, 23, 26, 40, 47, 51, 52, 148, 164, 165, 204 replication forks, ix, 1, 2, 3, 5, 7, 9, 10, 15, 18, 20, 21, 106 reptation model, 51 requirements, 28, 49 researchers, xiii, 82, 110, 126 residues, 133, 163, 164, 177, 179 resistance, 11 resolution, 6, 12, 127 response, xi, 3, 8, 10, 15, 21, 22, 23, 49, 53, 55, 132, 174, 196, 200 restoration, 27 restorative dentistry, ix, 28 restorative material, x, 25, 47 restorative materials, x, 25, 47 restriction enzyme, 49 reticulum, 56 retroviruses, 166 reverse transcriptase, 86, 151, 166, 173, 190, 196 ribosome, 179 rings, 162 risk, 4, 8, 76 risks, 48 RNA, 56, 86, 92, 93, 101, 104, 150, 151, 157, 163, 164, 166, 173, 175, 176, 185, 188, 190, 192 RNAi, 157, 164, 181, 183 room temperature, 35 root, 48 Royal Society, 74 S salivary gland, 153 salmon, 85, 86, 101 scavengers, 50 sclerosis, 56, 70 scope, 29 secretion, 56 segregation, 7, 16, 157, 158, 163, 204, 205 selectivity, 120 self-organization, 185

senescence, 14, 184 senile dementia, 73 sensitivity, 8, 11, 14, 50 sequencing, 117, 118, 120, 133, 165 serum, 30, 34, 87 serum albumin, 34, 87 services, xiii, 110 shape, 60, 152, 158, 161 shelter, 174, 196, 203 shock, 173 showing, 59, 164, 176 side effects, 187 signal peptide, 56, 76 signalling, 10, 19 signals, 125, 143 silica, 83 single-nucleotide polymorphism, xii, 109, 111, 130 single-nucleotide polymorphisms (SNPs), xii, 109, 111 siRNA, 101 sister chromatid exchange, 5, 6, 11 skin, 154 sleeping sickness, 154 SNP, 117, 122, 123, 125, 127, 129 sodium, 33, 122 software, 31, 32, 35, 36, 60, 62, 90 solid tumors, 94, 101 solution, 30, 31, 32, 113 South America, xiv, 147 species, xii, 52, 80, 82, 102, 122, 149, 151, 152, 153, 154, 157, 158, 167, 174, 175, 176, 181, 188, 199, 204 S-phase, ix, xii, 1, 3, 12, 19, 80, 90, 91, 93, 178, 184, 199, 210, 211 spindle, 60, 184, 199, 208 Spring, 102, 198 stability, ix, 2, 5, 13, 17, 23, 171 stabilization, ix, 2, 173, 178 standard deviation, 37 standard error, 43, 62, 63 state, 126, 162, 178 statistics, 62 stomach, 89 stress, 3, 16, 21, 152, 173

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Index stretching, 99 striatum, 64 structural protein, 133 structure, 6, 7, 9, 10, 13, 14, 17, 18, 19, 20, 22, 29, 46, 61, 82, 83, 84, 104, 106, 113, 114, 144, 145, 149, 161, 177, 181, 190, 193, 208 subsidy, 102 substitution, 119 substitutions, xiv, 115, 118, 119, 135, 136, 137, 139, 140, 141, 142, 143, 144, 145 substrate, xii, 6, 16, 80, 82, 86, 87, 88, 172, 174 substrates, 5, 13, 14 sucrose, 167 sulfate, 122 Sun, 49, 200 suppression, 5, 6, 93, 129 surface chemistry, 132 surveillance, 7 survival, 2, 52, 76, 155 susceptibility, 21 Switzerland, 106 symbiosis, 165 symmetry, 140, 145 syndrome, 4, 5, 8, 11, 16, 17, 18, 19, 21, 22, 23, 118 synthesis, x, xii, xiv, 2, 3, 17, 80, 81, 83, 92, 94, 97, 98, 103, 104, 106, 143, 145, 148, 150, 158, 166, 168, 172, 174, 178, 201, 205, 206, 210

temperature, 32, 101, 125 temporal lobe, xi, 53, 55, 57, 58, 63, 64, 69, 70 termination codon, 139, 144 testing, 28, 29, 50 therapeutics, 191 thymine, xiii, 103, 110, 111, 129, 130 tissue, 31, 57, 59, 61, 66, 70, 71, 119, 127 tooth, 27, 45 toxicity, 27 toxicology, 48, 49, 50 transactive response (TAR), xi, 53, 55 transcription, 162, 171, 178, 179, 182, 184, 185, 186, 200, 201, 205, 207, 209 transduction, 3 transfection, 99 transformation, 46, 145 translation, 177, 208 translesion synthesis (TLS), x, 2, 166 translocation, 175 transmission, 189 transport, 21 treatment, 31, 40, 44, 51, 91, 93, 96, 97, 98, 100, 152 Trypanosomatidae, xiv, 147, 152, 167, 168, 175, 177, 208 trypanosomiasis, xiv, 147, 152, 196 trypsin, 30, 31 tumor, 20, 45, 46, 94, 101, 105 tumor growth, 94, 101 tyrosine, 163

T tandem repeats, 171 tangles, 74 target, xiii, 45, 100, 110, 117, 119, 126, 164, 187, 193 tau, 56, 66, 71, 72, 74, 76 techniques, xiii, 95, 99, 110, 126, 132, 165 technology, 120, 121, 129, 131 telangiectasia, 3, 19 telomere, 151, 172, 173, 174, 175, 176, 187, 190, 192, 195, 196, 197, 202, 205, 206, 208, 210

189

U UK, 32, 35, 53, 72 ultrastructure, 190, 211 universe, 18, 196 urethane, 27 USA, 30, 31, 36, 57, 60, 62, 76, 102, 190, 200, 206, 207 USSR, 145 UV, 32, 33, 37, 38, 94 UV irradiation, 32, 37

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190

Index V

vacuole, 154 variations, 61, 111, 151, 152, 174, 175, 199 vector, 142, 152, 153, 154 velocity, 87 vertebrates, 173 viruses, 111, 162 visualization, xiii, 110, 111, 119, 122 vitamins, 52

Western blot, 91, 93, 121 white matter, 57, 61, 69 wild type, 12 Y yeast, 5, 9, 13, 15, 17, 20, 21, 35, 101, 159, 163, 174, 177, 183, 185, 192, 193, 195, 196, 199, 202, 203, 209, 210 yield, 137, 138, 151

W

zinc, ix, xiii, 110, 112, 120, 121, 122, 126, 129, 130, 131

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Washington, 57 wells, 35 West Africa, 154

Z

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