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Tomi Hillukkala

ROLES OF DNA POLYMERASE EPSILON AND TOPBP1 INDNA REPLICATION AND DAMAGE RESPONSE

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A C T A U N I V E R S I T A T I S O U L U E N S I SA S c i e n t i a e R e r u m N a t u r a l i u m 4 7 8

TOMI HILLUKKALA

ROLES OF DNA POLYMERASE EPSILON AND TOPBP1 INDNA REPLICATION ANDDAMAGE RESPONSE

Academic dissertation to be presented, with the assent ofthe Faculty of Science of the University of Oulu, for publicdefence in Raahensali (Auditorium L10), Linnanmaa,on December 15th, 2006, at 12 noon

OULUN YLIOPISTO, OULU 2006

Copyright © 2006Acta Univ. Oul. A 478, 2006

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OULU UNIVERSITY PRESSOULU 2006

Hillukkala, Tomi, Roles of DNA polymerase epsilon and TopBP1 in DNA replicationand damage responseFaculty of Science, Department of Biochemistry, University of Oulu, P.O.Box 3000, FI-90014University of Oulu, Finland, Biocenter Oulu, University of Oulu, P.O. Box 5000, FI-90014University of Oulu, Finland Acta Univ. Oul. A 478, 2006Oulu, Finland

AbstractDuring DNA replication cells accurately copy their DNA to transfer the genetic information todaughter cells. DNA polymerases synthesise the new DNA strand using the old strand as a template.Other functions of DNA polymerases are recombination linked and DNA iamage repair linked DNAsynthesis, regulation of replication complex formation and regulation of transcription – a process inwhich the genetic information is transformed into an RNA sequence needed to guide proteinsynthesis.

In this study, the TopBP1 protein was shown to associate with DNA polymerase epsilon. TopBP1contains eight BRCT domains mediating interactions between phosphorylated proteins and is ahuman homolog of bakers yeast Dpb11 and fission yeast Cut5. These yeast proteins act on DNAreplication and cell cycle arrest after DNA damage. TopBP1 was found to be phosphorylated andexpressed in elevated amounts during S phase suggesting an involvement in DNA replication. Thiswas directly demonstrated by DNA synthesis inhibition by a competing TopBP1 fragment and by anantibody targeted to block TopBP1.

Ultraviolet irradiation damages DNA and decreases the amount of TopBP1 in the nucleus. Thetranscription factor Miz-1 was found to associate with TopBP1 and was released from this interactionafter UV damage. Free Miz-1 activated the expression of the cell cycle arresting proteins p15 and p21cooperatively with other transcription factors and allowed extra time for DNA damage repair.

TopBP1 was also found to interact with the breast cancer susceptibility protein 1 and both proteinslocalised together to arrested DNA synthesis apparatuses. The interaction of TopBP1 with thedamage recognition and processing protein Rad9 is still further evidence of a link between TopBP1and DNA damage.

DNA polymerase epsilon forms a complex with Cdc45, a protein involved in DNA replicationinitiation and elongation. This complex does not interact with Cdc45 complexed with DNApolymerase delta, suggesting that these complexes synthesise DNA independently of each other. Ourresults are in agreement with the view that polymerase epsilon synthesises the first strand of DNAand polymerase delta the other.

Finally,DNA polymerase epsilon binds to the RNA synthesising form of RNA polymerase II andnascent transcripts. The physiological meaning of this interaction needs to be determined.

Keywords: cell cycle, DNA polymerase, DNA repair, DNA replication, gene expression,nucleotide excision repair, phosphorylation, protein-protein interactions, RNA polymeraseII, transcription, ultraviolet rays

Acknowledgements

This work has been performed at the Department of Biochemistry and Biocenter Oulu at the University of Oulu during years 1998-2006.

I would like to express thanks to my supervisors Professor Juhani Syväoja and Helmut Pospiech for their never ending flow of research suggestions and the high level of freedom to complete these projects myself. Helmut Pospiech has also provided solid land to build research plans by making educated suggestions for key experiments. Thank you for that. I am grateful for Professor Kalervo Hiltunen who has organised excellent research facilities at the department of biochemistry. For example, when the immunofluorescence microscope was changed to a better one, my needs were taken into account in a nice way. I wish to thank Varpu Marjomäki and Minna Nyström for your valuable comments and criticism over the manuscript. I was positively surprised when I got your comments several weeks in advance. The Akin’s law “Schedules only move in one direction” was thereby proven wrong.

All JS-group members deserve my greatest thanks. I would especially like to thank Minna Mäkiniemi for guiding me into the research community. I am also thankful for Kaarina Reini and Anna Rytkönen for your peer support during all these years. Without you I would never have finished this project. I thank Maarit Jokela, Siri Lehtonen, and Jussi Tuusa for your positive attitude towards life. You created the good spirit which was characteristic to our work at the late nineties. I would also like to thank Markku Vaara and Miiko Sokka for your valuable collaboration in recent years. Professor Heinz-Peter Nasheuer is known for good work with Cdc45 and DNA replication. Collaboration with you was a vital part of my work. Thank you so much!

There are several people giving invaluable help for the researchers at the department of biochemistry. Thanks for all of you and especially Maila Konttila; you are like a mother for the department while keeping things organised. I also thank Jaakko Keskitalo, Kyösti Keränen, and Jari Karjalainen for helping me with several work and hobby related technical problems. Ari-Pekka Kvist deserves my special thanks for making data system to work so fluently that it rarely disturbed my work.

I wish to thank all the members of the volley ball team Pateniemen Vesa. Hitting the ball has given me a lot of strength to keep up the research work. Ursa, the astronomical

society of Finland, and Arktos, the astronomical society of Oulu, have taught me the beauty of the night sky. Thank you for supporting this great hobby.

I thank my mother Marja-Leena and father Seppo for taking care of me during the first twenty years of my life and keeping close touch since then. You have always been interested in my affairs and given the support I need. My sisters, brothers, and their families deserve the same thanks. My connection to Marjo and Mikko, Teija and Jouni, and Heikki and Pasi is a close one and I know I can always trust you. In addition to the previous I am thankful to my brother Mika for coffee and lunch break companionship and valuable comments on subjects whatsoever. Arja and Pauli, you have brought up a truly lovable daughter. You have always wished me welcome at your home, and I feel you have taken me as your son. Päivi and Rami, you are like brothers to me. You have always given me the help I need with some fresh, open minded criticism, which I really appreciate.

My warmest thanks belong to my wife Mervi, and our children Lauri and Martta. You are the most important thing in my life.

This work was supported by the Academy of Finland and Cancer Foundation.

Oulu, October the 26th 2006 Tomi Hillukkala

Abbreviations

ATM ataxia telangiectasia mutated ATP adenosine triphosphate ATR ATM and Rad3 related BER base excision repair BrdU bromodeoxyuridine bp base pair BRCA1 breast cancer susceptibility gene/protein 1 BRCT BRCA1 C-terminus BSA bovine serum albumin Cdc cell division control Cdk cyclin-dependent kinase CIP calf intestinal phosphatase cDNA complementary DNA CHRAC chromatin-remodelling complex CS Cockayne Syndrome CSA Cockayne syndrome A protein CSB Cockayne syndrome B protein CTD carboxyterminal domain C-terminus carboxy terminus DNA deoxyribonucleic acid DSB DNA double strand break dsDNA double-stranded DNA DTT dithiothreitol EDTA ethylenediaminetetraacetic acid Fen-1 flap endonuclease 1 GST glutathione S-transferase HEPES N-[2-hydroxyethyl]-piperazine-N’-[2-ethanesulphonic acid] IEM immuno-electron microscopy IPTG isopropyl-β-D-thiogalactoside kDa kilodalton(s) MCM minichromosome maintenance protein

MMR mismatch repair mRNA messenger RNA NER nucleotide excision repair NHEJ non-homologous end joining nt nucleotide N-terminus amino terminus ORC origin recognition complex PCNA proliferating cell nuclear antigen PCR polymerase chain reaction Pol DNA-dependent DNA polymerase Pre-RC pre-replication complex RFC replication factor C RNA ribonucleic acid RNA pol RNA polymerase RPA replication protein A TCR transcription coupled repair TFIIH transcription factor IIH TopBP1 topoisomerase IIβ binding protein 1 UV ultraviolet XP Xeroderma pigmentosum

List of original articles

This thesis is based on the following articles, which are referred to in the text by their Roman numerals:

I Mäkiniemi M, Hillukkala T, Tuusa J, Reini K, Vaara M, Huang D, Pospiech H, Majuri I, Westerling T, Mäkela TP & Syväoja JE (2001) BRCT domain-containing protein TopBP1 functions in DNA replication and damage response. J Biol Chem 276: 30399-30406.

II Herold S, Wanzel M, Beuger V, Frohme C, Beul D, Hillukkala T, Syväoja J, Saluz HP, Haenel F & Eilers M (2002) Negative regulation of the mammalian UV response by Myc through association with Miz-1. Mol Cell 10: 509-521.

III Rytkönen AK*, Hillukkala T*, Vaara M, Sokka M, Jokela M, Sormunen R, Nasheuer H-P, Nethanel T, Kaufmann G, Pospiech H & Syväoja JE (2006) DNA polymerase ε associates with the elongating form of RNA polymerase II and nascent transcripts. FEBS J, in press.

IV Hillukkala T, Vaara M, Koistinen H, Sokka M, Nasheuer H-P, Syväoja JE & Pospiech H (2006) Human Cdc45 forms distinct complexes with DNA polymerases δ and ε during nuclear DNA replication. Manuscript

* These authors contributed equally to this work.

Contents

Abstract Acknowledgements Abbreviations List of original articles Contents 1 Introduction ................................................................................................................... 13 2 Review of the literature ................................................................................................. 14

2.1 DNA replication mechanism and fidelity ...............................................................14 2.2 Eukaryotic DNA polymerases ................................................................................14 2.3 Introduction to DNA replication.............................................................................15

2.3.1 DNA polymerase α ..........................................................................................16 2.3.2 DNA polymerase δ and PCNA ........................................................................17 2.3.3 DNA polymerase ε and GINS..........................................................................17

2.4 Assembly of proteins to origins of DNA replication – Pre-replication complex formation.................................................................................................19

2.5 Initiation of DNA replication..................................................................................19 2.6 Replication fork progression...................................................................................21

2.6.1 Leading and lagging strand replication............................................................22 2.7 Maturation of Okazaki Fragments ..........................................................................24 2.8 Regulation of DNA replication...............................................................................25 2.9 Transcription and DNA replication.........................................................................27

2.9.1 Regulation of transcription ..............................................................................27 2.9.2 RNA polymerase II..........................................................................................28

2.10 DNA repair pathways ...........................................................................................29 2.10.1 Nucleotide excision repair .............................................................................30

3 Aims of the present work............................................................................................... 33 4 Materials and methods................................................................................................... 34

4.1 cDNA cloning (I) ....................................................................................................34 4.2 Gene chromosomal mapping (I) .............................................................................34 4.3 Production of antibodies (I, II) ...............................................................................34 4.4 Cell lines and synchronisation (I, III, IV)...............................................................35

4.5 Induction of DNA damage (I, III)...........................................................................35 4.6 Preparation of cell extracts (I, III) ..........................................................................35 4.7 T98G cell fractionation (IV)...................................................................................36 4.8 Immunoprecipitation (I, III, IV) .............................................................................36 4.9 SDS-PAGE and Western-analysis (I, III)................................................................36 4.10 Indirect immunofluorescence (I, III, IV) ..............................................................37 4.11 Immuno-electron microscopy (III, IV) .................................................................38

5 Results ........................................................................................................................... 39 5.1 cDNA cloning and chromosomal localisation of human TOPBP1 (I) ....................39 5.2 TopBP1 is a phosphoprotein expressed in a proliferation dependent

manner (I) ..............................................................................................................40 5.3 TopBP1 interacts with DNA polymerase ε and hRad9 (I) ......................................40 5.4 TopBP1 is required for chromosomal DNA replication (I).....................................41 5.5 TopBP1 and Brca1 colocalise in S phase and become relocalised into

PCNA containing foci after replication block (I)...................................................41 5.6 TopBP1 interacts with Miz-1 (II)............................................................................41 5.7 TopBP1 inhibits activation of p15INK4B promoter by Miz-1 (II) .............................42 5.8 After UV damage TopBP1 mRNA levels are downregulated and

interaction of TopBP1 with Miz-1 decreases followed by upregulation of p15INK4B (II) ...........................................................................................................42

5.9 The ATM/ATR kinase inhibitor caffeine blocks downregulation and redistribution of TopBP1 (II) .................................................................................43

5.10 TopBP1 binds to the p21Cip1 promoter together with Miz-1 (II) ...........................43 5.11 DNA polymerase ε interacts with the hyperphosphorylated RNA pol II

and recently synthesised RNA (III) .......................................................................43 5.12 DNA polymerase ε and hyperphosphorylated RNA polymerase II

colocalise (III) .......................................................................................................45 5.13 DNA polymerase ε interaction with RNA pol II may not be linked to

nucleotide excision repair or DNA replication (III)...............................................46 5.14 Cdc45, pol δ, pol ε, and PCNA behave differently in cell fractionation

(IV) ........................................................................................................................47 5.15 Cdc45 colocalises with DNA polymerases δ and ε (IV).......................................47

6 Discussion ..................................................................................................................... 49 6.1 TopBP1 interacts with DNA polymerase ε and is needed for replication (I) ..........49 6.2 Cdc45 forms separate complexes with DNA polymerases δ and ε (IV) .................50 6.3 DNA polymerase ε associates with transcribing RNA polymerase II and

nascent transcripts (III)..........................................................................................51 6.4 TopBP1 functions in the DNA damage response (I, II) ..........................................52

7 Conclusions ................................................................................................................... 56 References Original articles

1 Introduction

Genetic information stored as a nucleotide sequence needs to be maintained and transferred to the next cell generation without alterations. DNA polymerases α, δ, and ε are enzymes duplicating the chromosomal DNA. Pol α starts the DNA synthesis and pols δ and ε continue the work producing most of DNA. DNA polymerases also produce DNA patches to stabilise DNA, and to repair alterations caused by cellular metabolism and exogenous stress. To verify genome integrity cells have established several cell cycle and DNA damage checkpoints, which enable responses to environmental changes. This results in the formation and disruption of protein complexes, protein expression level alterations and different post-translational modifications of proteins. DNA polymerases can also deliberately change the genetic information in mitotic recombination or in somatic recombination of heavy chain variable regions of antibodies.

In this study TopBP1 was characterised as a pol ε associating phosphoprotein. TopBP1 levels peaked prior to S phase, which is typical for proteins involved with replication. ToBP1 inhibition downregulated DNA synthesis efficiently, manifesting its function in DNA replication. TopBP1 was found to interact with the DNA damage recognition and processing protein hRad9. TopBP1 also localised together with the breast cancer susceptibility protein 1 (Brca1) to stalled replication forks and, after DNA damage, to other nuclear structures, linking the TopBP1 and Brca1 damage responses. Finally, TopBP1 was found to associate with transcription factor Miz-1. In response to UV irradiation TopBP1 levels were downregulated and Miz-1 was released resulting in upregulation of cell cycle inhibitors and cell cycle arrest.

Both pols δ and ε formed complexes with Cdc45 although these polymerases did not interact with each other. Obviously Cdc45 does not mediate the interaction between leading and lagging strand DNA polymerases, a difference between prokaryotic and eukaryotic replication systems. Independent lagging strand replication may allow the synthesis of multiple Okazaki fragments at the same time shortening the time needed to duplicate the eukaryotic genome. Pol ε also associated with transcriptionally active RNA polymerase II and nascent transcripts. Interaction was specific for pol ε as RNA pol II was not detected in complex with either pol α or δ. Pol ε interaction with RNA pol II may involve coordination of replication or transcription initiation, facilitation of DNA repair or coordination of bypassing transcription apparatuses and replication forks.

2 Review of the literature

2.1 DNA replication mechanism and fidelity

DNA-dependent DNA polymerases (pols) produce nascent DNA preserving the genetic information stored in the template strand (Steitz et al. 1994). Pols catalyse the hydrolysis of deoxyribonucleotidetriphosphates (dNTPs) into deoxyribonucleotidemonophosphates (dNMPs) and pyrophosphate groups (PPs). Polymerases add the dNMP to a free 3’-hydroxylgroup of primer DNA and use the other strand of the double-stranded DNA as a template resulting in the lengthening of the primer-DNA by one nucleotide and a new free 3’-end. Processive pols repeat the addition of dNMP without dissociation from the template DNA while distributive ones add a single nucleotide at a time. Pyrophosphate is a side product that becomes hydrolysed rapidly to monophosphates providing energy for the reaction and making it irreversible.

The added nucleotides form Watson-Crick base pairs (adenine (A) – thymine (T) and guanine (G) – cytosine (C)) with the template strand to achieve the correct position in DNA. Pols have several fidelity mechanisms in the selection of dNTP and phosphodiester bond formation to prevent misincorporation resulting in the error rate of 10-3 to 10-5 per added nucleotide (reviewed by Echols & Goodman 1991). Some pols have a 3’ to 5’ exonuclease activity to scan for errors and to digest misincorporated nucleotides from the primer end. This proofreading activity increases the overall fidelity 100-fold. Mismatch repair is the third step in gaining the required 10-9 to 10-10 fidelity for large eukaryotic genomes.

2.2 Eukaryotic DNA polymerases

Genome sequencing projects during recent years have vastly increased the number of DNA polymerases known. There are 16 known DNA dependent DNA polymerases in eukaryotes (Marini et al. 2003, reviewed by Hübscher et al. 2002 and Rattray et al. 2003). In addition, there is a telomerase, an RNA directed DNA polymerase needed for telomere maintenance, and a terminal deoxynucleotidyltransferase, a template

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independent DNA polymerase acting in somatic hypermutation of the immunoglobulin genes. Many recently identified pols are specialised in certain types of DNA damage justifying the presence of a high number of different polymerases. Polymerases may also act as a backup for each other making single enzymes dispensable for certain processes.

Eukaryotic pols are named using Greek letters in the order of discovery (Burgers et al. 2001). Rev1 is the only exception probably because it only inserts cytidines in abasic sites on DNA. The eukaryotic pols listed in Table 1 can be divided into two functional groups; replicative or DNA repair DNA polymerases (reviewed by Hübscher et al. 2002). The groups are not exclusive as pol α, δ, and ε perform chromosomal DNA replication and participate in several repair pathways, for example. Replicative pols are very accurate whereas DNA repair polymerases specialised in damage bypass can easily make mistakes. These translesion synthesis polymerases provide damage tolerance for cells because DNA replication can proceed through the damaged area in DNA. A third group of polymerases can be called miscellaneous DNA polymerases including those required to produce immunoglobulins by somatic hypermutation.

Table 1. Eukaryotic DNA polymerases; Nomenclature and main functions

Proposed main function Greek name Other names for the polymerase DNA replication α (alpha) Base excision repair β (beta) Mitochondrial DNA replication γ (gamma) DNA replication δ (delta) DNA replication ε (epsilon) Translesion synthesis (extension of mispaired area) ζ (zeta) Translesion synthesis (relatively high accuracy) η (eta) XPV Interstrand crosslink repair θ (theta) Translesion synthesis (very low fidelity) ι (iota) Translesion synthesis (low fidelity, moderate processivity) κ (kappa) Base excision repair λ (lambda) Non-homologous end joining in hypermutation of immunoglobulin genes

μ (mu)

Interstrand crosslink repair ν (nu) Sister chromatid cohesion σ (sigma) TRF4 Role in rRNA production (high accuracy in DNA synthesis) φ (phi) Translesion synthesis (incorporates C at an abasic site) Rev1 Partially adopted from Burgers et al. (2001), Hübscher et al. (2002), Marini et al. (2003), and Prakash et al. (2005).

2.3 Introduction to DNA replication

Xenopus laevis egg extracts have been beneficial in the study of replication initiation factors (Pasero & Gasser 1998). Another source of information has been virus based replication systems, especially simian virus 40 (Li & Kelly 1984, Waga & Stillman

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1994). Viruses utilise the cellular replication apparatus to copy their own genome and therefore allow the analysis of cellular elongation phase components. The SV40 system uses a viral initiator and a helicase protein revealing the use of RPA in the coating of unwound DNA, pol α, which functions as a primase, polymerase switching with the help of RFC and PCNA and pol δ for replicating both leading and lagging strand DNA. Fen1, Dna2 and RnaseH were found to be components of Okazaki fragment maturation. The function of these proteins will be discussed below. The third important source of knowledge has been genetic studies in yeast, which has allowed the identification of replication or replication regulation related proteins (reviewed by Sugino 1995).

All replicative polymerases belong to the same family of DNA polymerases and share a common fold (Asturias et al. 2006, reviewed by Hübscher et al. 2002). The template strand is positioned in an active site cleft, where it makes a sharp turn at the active site. The catalytic residues are buried inside the domain where they bind to two metal ions and coordinate formation of the reaction intermediate. The polymerase may have another active site for exonuclease activity and, in that case, the pol must have a shuffling mechanism to transfer the primer end between the polymerase and exonuclease sites (Franklin et al. 2001, Shamoo & Steitz 1999).

2.3.1 DNA polymerase α

Pol α consists of four subunits containing a primase activity that is unique among DNA polymerases (reviewed by Garg & Burgers 2005a and Arezi & Kuchta 2000). Human 180 kDa Pol1 is the largest subunit containing the DNA polymerase activity but no exonuclease activity despite the presence of an exonuclease domain. The second largest 70 kDA-subunit binds tightly to and regulates the catalytic subunit (Nasheuer et al. 1991). The 55 kDa Pri2, primase accessory subunit, regulates the RNA polymerase activity of the 49 kDa Pri1 (Santocanale et al. 1993, Zerbe & Kuchta 2002). Pri2 is required for primase stability, initiation effectiveness, primer length determination and possibly for the transfer of the RNA primer terminus to the catalytic subunit.

RNA polymerase activity enables pol α to start DNA synthesis with an 8-12 nucleotide long primer RNA and a 20 nucleotide long DNA stretch (Eliasson & Reichard 1978, Nethanel & Kaufmann 1990). Pol α synthesises a primer to each origin on the leading strand and to each Okazaki fragment on the lagging strand. It is likely that pol δ or possibly p53 proofreads all DNA synthesised by pol α because of the lack of an active exonuclease domain in pol α (Melle & Nasheuer 2002).

Pol α activity is tightly regulated as it is the initiator of DNA synthesis. Mcm10 and Cdc45 bind to pol α targeting it to replication initiation sites (Mimura et al. 2000, Zou & Stillman 2000). During the S and G2 phases, Cdk2/CyclinA phosphorylates the second largest subunit regulating the polymerase activity of pol α. A hypophosphorylated form of pol α can be recruited to DNA while hyperphosphorylation inhibits the loading. (Nasheuer et al. 1991, Desdouets et al. 1998).

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2.3.2 DNA polymerase δ and PCNA

Human pol δ consists of a 125 kDa catalytic A-subunit containing DNA polymerase and proofreading exonuclease activities, a 50 kDa regulatory B-subunit, a 68 kDa C-subunit binding to the regulatory subunit, and a 12 kDa D-subunit (Liu et al. 2000, Podust et al. 2002). Saccharomyces cerevisiae pol δ consists of only three subunits (A, B and C), although the fourth D-subunit is needed for full activity of the pol δ complex in human and Schizosaccharomyces pombe (Zuo et al. 1997, Liu et al. 2000, Podust et al. 2002). The Pol δ complex is monotetrameric containing one molecule of each subunit (Johansson et al. 2001, Bermudez et al. 2002). The C-subunit is dispensable in S. cerevisiae, although the cells show poor growth, increased sensitivity to replication inhibitors and DNA damaging agents and synthetic lethality with many DNA metabolism related genes (Gerik et al. 1998, Huang et al. 2002, Tong et al. 2004). The C-subunit is capable of recruiting PCNA to the pol δ complex and is absolutely required for growth in S. pombe (Bermudez et al. 2002). The B-subunit does not have enzymatic activities but belongs to a conserved superfamily of polymerase B-subunits and is essential for stabilising the holoenzyme (Mäkiniemi et al. 1999).

The DNA replication processivity factor PCNA was first discovered as an auxiliary protein for pol δ enhancing DNA replication activity and existing in the nucleus in elevated amounts during S phase (Tan et al. 1986, Prelich et al. 1987, Zhang et al. 1995). Homotrimeric PCNA consists of 29 kDa-subunits and, with the help of the loading factor RFC, forms a ring-shaped sliding clamp around DNA (Krishna et al. 1994, Gulbis et al. 1996). PCNA binds to over 30 proteins in the nucleus forming an interaction network linked to DNA replication and repair, cell cycle control, and chromatin remodelling (Dua et al. 2002, Garg et al. 2005, Gulbis et al. 1996, Shibahara & Stillman 1999, reviewed by Maga & Hübscher 2003). PCNA is modified by ubiquitination to promote interaction with translesion DNA polymerases η and ζ enabling these damage tolerant DNA polymerases to perform lesion bypass DNA synthesis (Garg et al. 2005, Garg & Burgers 2005b). Pol δ and PCNA bind together through several interaction sites some of which are in use only when PCNA is bound to DNA (Tan et al. 1986, Prelich et al. 1987). Pol δ A and B-subunits form a stable complex with DNA bound PCNA underlining the importance of DNA association in revealing the interaction sites (McConnell et al. 1996, Mozzherin et al. 1999). The less conserved C-subunit also has a consensus PCNA binding sequence in the C-terminus (Liu et al. 2000, Johansson et al. 2004).

2.3.3 DNA polymerase ε and GINS

Pol ε was first isolated from calf thymus and later on as a complex from S. cerevisiae (Syväoja & Linn 1989, Morrison et al. 1990). Yeast pol ε has been ectopically expressed in a baculovirus system and in yeast as a heterotetramer of Pol2 (256 kDa A-subunit), Dpb2 (78 kDa B-subunit), Dpb3 (23 kDa C-subunit), and Dpb4 (22 kDa D-subunit) (Dua et al. 2002, Chilkova et al. 2003). Pol2 contains DNA polymerase and proofreading exonuclease activities, but the entire N-terminus containing both activities is dispensable (Kesti et al. 1999). Deletion of the N-terminus causes severe growth defects. The C-

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terminus of Pol2, however, contains a zinc finger region commonly mediating interactions between proteins, and this zinc finger is essential for growth and for S phase checkpoint function (Navas et al. 1995, Araki et al. 1995, Dua et al. 1999, Feng & D'Urso 2001). It is plausible that the C-terminus of Pol2 is an essential component of the replication initiation complex as pol ε is loaded onto origins before primer synthesis. A chromatin immunoprecipitation assay showed that both wild type pol ε and a catalytic domain deficient POL2-16 mutant bind to origins in early S phase (Hiraga et al. 2005). Wild type pol ε remained bound to DNA but the mutant was rapidly detached, suggesting that both proteins were able to carry out the essential function during initiation and this process did not require DNA synthesis activity. Replicase function during the elongation phase was performed by another polymerase enabling the viable phenotype. Interestingly the essential Pol2 C-terminus and the Dpb3-Dpb4 complex both have a double-stranded DNA binding domain that is uncommon among DNA polymerases (Tsubota et al. 2003). This suggests that the essential function of Pol ε is carried out while binding to DNA. Pol ε also has a non-replication related interaction with Trf4/pol σ acting in sister chromatid cohesion, and pol2 mutants are impaired in this process (Edwards et al. 2003). TRF4 is an essential gene, thus it is possible that the essential function of the Pol2 C-terminus is related to this interaction with Trf4. DPB2 is an essential gene while DPB3 and DPB4 are dispensable in S. cerevisiae (Araki et al. 1991a, Araki et al. 1991b, Ohya et al. 2000). In S. pombe Dpb2 and Dpb3 are essential proteins and only Dpb4 is non-essential working with Dpb3 in the initiation of DNA replication, in S phase progression and in cell separation (Feng et al. 2003, Spiga & D’Urso 2004). Both C and D-subunits contain histone fold motifs and form a stable dimer (Li et al. 2000). The D-subunit is a component of the CHRAC chromatin accessibility complex forming a link between DNA replication and chromatin remodelling (Corona et al. 2000, Poot et al. 2000, Iida & Araki 2004, Hartlepp et al. 2005).

Pol ε interacts with PCNA but also is a highly processive DNA polymerase without this interaction (Lee et al. 1991, Dua et al. 2002). The catalytic subunit of Pol ε has a globular structure containing a cleft for double-stranded DNA and forms an extended structure with three other subunits (Asturias et al. 2006). The latter part has a strong affinity to double stranded DNA enabling processive DNA synthesis by pol ε without the help of PCNA. PCNA increases the processivity of pol ε in high ionic strength solution in vitro, but this interaction may be needed for DNA repair rather than DNA replication (Lee et al. 1991, Dua et al. 2002). Deletion of the PCNA binding motif in Pol ε has almost no effect on growth but causes strong damage sensitivity. Instead of PCNA, GINS could be the processivity factor for pol ε during replication (Gambus et al. 2006, Kanemaki & Labib 2006). GINS is a ring-shaped complex of Sld5, Psf1, Psf2 and Psf3 conserved in all eukaryotes (Takayama et al. 2003). It is an essential component for both DNA replication initiation and elongation.

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2.4 Assembly of proteins to origins of DNA replication – Pre-replication complex formation

DNA replication begins from several sites in the chromosomes called origins (reviewed by Blow & Dutta 2005 and Kearsey & Cotterill 2003). S. cerevisiae origins are well characterised while origins in higher eukaryotes are less clearly defined (Wyrick et al. 2001). The origin function is analogous in all eukaryotes as manifested by precise replication initiation sites in mammalian origins (Abdurashidova et al. 2000, Kamath & Leffak 2001). Furthermore many origins are located in promoter areas of highly transcribed genes suggesting that transcription factors have a role in the initiation process (Murakami & Ito 1999, Todorovic et al. 1999).

The origin recognition complex (ORC), consisting of six subunits Orc1 - Orc6, binds to these sequences in an ATP dependent manner (Bell & Stillman 1992, Speck et al. 2005). S. cerevisiae and S. pombe ORC complexes are bound to origins in all cell cycle stages, whereas in Xenopus the whole complex, and in human, Orc1 binds to DNA in early G1 phase followed by Cdc6 and Cdt1 (Aparacio et al. 1997, Tanaka et al. 1997, Ogawa et al. 1999, Romanowski et al. 1996, Kreitz et al. 2001, Wohlschlegel et al. 2000). Minichromosome maintenance proteins 2-7 (MCMs) are loaded after that forming a complete pre-replicative complex (Kubota et al. 1997, reviewed by Blow & Dutta 2005). MCM loading factors Orc, Cdc6 and Cdt1 can be removed from DNA after Mcm loading without affecting DNA replication initiation (Rowles et al. 1999). The Mcm2-7 complex shows a DNA dependent ATPase activity, and a sub complex containing a double heterohexameric complex of Mcm4, Mcm6 and Mcm7 shows a weak but processive DNA helicase activity with forked DNA structures (Ishimi 1997, Schwacha & Bell 2000, Lee & Hurwitz 2001). The Mcm4,6,7 complex is capable of unwinding 600 base pairs of duplex DNA and is the best candidate to perform DNA helicase function in eukaryotic DNA replication. Mcms 2, 3, and 5 may have a regulatory role because all of them are essential for replication. Mcm2-7 are also needed for other functions like the S phase checkpoint and possibly only a small fraction of all DNA bound Mcms act as a helicase. Two Mcm complexes are needed per Orc for full DNA replication activity, but 10 to 40 complexes are loaded (Mahbubani et al. 1997, Edwards et al. 2002). Extra Mcm proteins may pump DNA to immobile replication factories assisting in the unwinding process or they may remove nucleosomes from DNA ahead of the replication apparatus (Laskey & Madine 2003).

2.5 Initiation of DNA replication

The pre-replicative complex is modified by cyclin dependent kinases (Cdks) and by Cdc7/Dbf4 activity to form an initiation complex at the G1/S transition (reviewed by Bell & Dutta 2002 and Blow & Dutta 2005). Cdc7 is a highly conserved serine/threonine kinase conserved from yeast to mammals. Its activity is dependent on either the regulatory subunit Dbf4 or an alternative regulatory subunit Drf1 in vertebrates, both peaking in the G1/S transition (Jares & Blow 2000, Takahashi & Walter 2005). Dbf4

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targets Cdc7 to chromatin and enables phosphorylation of Mcm proteins (Varrin et al. 2005). Cdks are serine/threonine kinases whose activity is dependent on an unstable positive regulatory subunit cyclin (reviewed by Sherr & Roberts 2004). Cdk activity is modulated by phosphorylation and Cdk inhibitors. Cyclins also contain an ubiquitinylation consensus sequence that targets them to degradation in a cell cycle dependent manner.

Phosphorylation of the pre-replication complex enables geminin to harvest Cdt1 away from the complex allowing Mcm10 loading to origins (Saxena et al. 2004, Sawyer et al. 2004). Mcm10 interacts directly with Cdc45 facilitating its loading and is needed for efficient initiation. Furthermore Mcm10 has a conserved chaperone-like domain required to stabilise the catalytic subunit of pol α (Ricke & Bielinsky 2004, 2006). Mcm10 is needed for loading pol α onto origins and for maintenance of the pol α chromatin association during replication fork progression in S. pombe. Differentiated phosphorylation of initiation complex proteins and loss of Cdt1 ensures that further loading of MCMs is prevented and there will be no more replication initiation sites formed before mitosis.

The initiation complex contains at least the presumed Mcm-helicase with regulatory subunits, RPA, Cdc45 and Sld proteins, Dpb11/Cut5, GINS, pol α and pol ε (Gambus et al. 2006, reviewed by Garg & Burgers 2005a). Furthermore topoisomerase I activity is required to release torsional stress in front of the replication fork during the unwinding process. Assembly of these proteins is presented in Figure 1.

RPA consists of three subunits all of which are important for the major cellular function of RPA, single-stranded DNA binding (Weisshart et al. 2004). The origin associations of RPA and Cdc45 are mutually dependent and can only happen after the activation of the pre-replication complex (Zou & Stillman 2000).

The loading of pol ε onto origins requires interdependent binding of Dpb11/Sld2, Cdc45/Sld3 and GINS to initiation sites (Masumoto et al. 2002, Kanemaki & Labib 2006, Takayama et al. 2003, Gambus et al. 2006, Pacek et al. 2006). DPB11 was originally isolated as a component, which when overexpressed was able to relieve mutations in two pol ε subunits in S. cerevisiae (Araki et al. 1995). It was found to contain four BRCT domains commonly mediating protein-protein interactions and later on it was recognised as a loading component for pols α and ε (Masumoto et al. 2000). All Sld proteins were found from yeast in a screen looking for mutations lethal with a dpb11-1 mutant (Kamimura et al. 1998). Cdc45 is a key player in the pre-replication to initiation complex transition as it is able to bind directly to Mcms, RPA, pol ε, and in many species pol α (Pacek et al. 2006, reviewed by Bell & Dutta 2002). It forms a complex with Sld3 in yeast and both proteins are needed for initiation (Kamimura et al. 2001, Nakajima et al. 2002). Cdc45 binding to Mcms and the loading of RPA to origins is prevented without Sld3.

The binding of pol ε enables Cdc45 and Mcm10 to recruit pol α together with RPA to origins and only after that can pol α perform its priming function (Masumoto et al. 2000, Mimura et al. 2000, Zou & Stillman 2000, Uchiyama et al. 2001, Ricke & Bielinsky 2004).

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Fig. 1. Interactions between replication initiation proteins. A double line represents DNA and solid arrows indicate conserved interactions between eukaryotic proteins. A dashed line between Cdc45 and pol α indicates a weak interaction not present in all eukaryotes. The figure was in part adopted from Nasheuer et al. 2006.

2.6 Replication fork progression

Sld3, one of the Cdc45 loading components, becomes dispensable in S. cerevisiae after replication fork firing (Kanemaki & Labib 2006). GINS moves along with the replication fork and maintains the association of Cdc45 with the MCM complex. It seems that Sld3 performs its essential function while loading Cdc45 onto origins and is not needed after that, while GINS is needed for both initiation and elongation. Cdc45, GINS and MCM2-7 form the components of the eukaryotic helicase complex (Masuda et al. 2003, Pacek et al. 2006). Biotin-streptavidin modified nucleotides were used to induce artificial replication pause sites in plasmids, and a chromatin immunoprecipitation approach was utilised to identify proteins bound to these areas in Xenopus egg extracts. In the presence of aphidicolin, a toxin uncoupling the helicase and polymerase activities of the replication fork, Cdc45, GINS, and MCM2-7 were present at the pause sites. Without aphidicolin pols α, δ, and ε were also precipitated together with MCM10.

Pol α is not able to start RNA primer synthesis by itself but requires the Mcm complex, Cdc45 and Mcm10 to facilitate pol α binding to RPA coated single stranded

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DNA (Masumoto et al. 2000, Mimura et al. 2000, Zou & Stillman 2000, Uchiyama et al. 2001, Ricke & Bielinsky 2004). Mcm10 also stimulates the RNA primer-end switch from primase to catalytic subunit as shown by the increased polymerase activity of pol α in the presence of Mcm10 (Fien et al. 2004). After DNA synthesis of approximately 20 nucleotides, the RFC-PCNA complex binds to DNA facilitating pol α dissociation from, and pol δ loading onto DNA (Tsurimoto & Stillman 1991a, 1991b, Maga et al. 2000, Mossi et al. 2000, Gomes & Burgers 2001). This reaction requires ATP as an energy source. RFC and PCNA probably exist as a complex already in the nucleoplasm because RFC binds to DNA just transiently and only soluble RFC is able to bind to PCNA (Gomes et al. 2001). The PCNA – pol δ complex still recruits FEN1 from the nucleoplasm and elongates DNA rapidly (Burgers 1991, Ayyagari et al. 2003, Garg et al. 2004).

Flap endonuclease FEN1 has a 5’ to 3’ exonuclease activity and an endonuclease activity needed for primer RNA degradation in Okazaki fragment maturation (reviewed by Liu et al. 2004). This essential enzyme is able to cut a DNA flap, a 5’-displaced ssDNA from duplex DNA, by loading from the 5’ end of the flap and tracking the ssDNA to the dsDNA junction (Harrington et al. 1994, Jonsson et al. 1998, Tom et al. 2000, Sakurai et al. 2005). To perform this activity Fen1 is able to interact with both single- and double-stranded DNA and the front surface of the PCNA ring.

2.6.1 Leading and lagging strand replication

Leading strand replication is not as well understood as lagging strand synthesis. It is likely that the helicase and leading strand DNA polymerase are immobilised on the nuclear matrix and act as a group of several replication forks called a replication factory (Radichev et al. 2005, reviewed by Anachkova et al. 2005). This organisation of replication allows minimal torsional stress on DNA and forms one means for replication regulation. The model systems used to study DNA replication have lacked the nuclear matrix effect on replication weakening the applicability of the results to in vivo conditions. A model for protein assembly in a replication fork is presented in figure 2.

Inactivating point mutations in the polymerase domains of yeast pols δ and ε cause lethality showing that these replication domains position themselves in essential places in the replication fork (Kesti et al. 1999, Dua et al. 1999, Pavlov et al. 2001). Both pols participate in chromosomal DNA replication although the pol ε polymerase domain can be replaced by another DNA polymerase under certain conditions (Zlotkin et al. 1996, Pospiech et al. 1999). Pols α, δ and ε are loaded onto the same origins in early S phase suggesting that these polymerases cooperate in the same replication fork and therefore pols δ and ε probably occupy opposite DNA strands (Hiraga et al. 2005). Another model suggests a specialised role for pol ε in heterochromatin replication (Fuss et al. 2002).

Several lines of evidence suggest that pol ε is the leading strand DNA polymerase and pol δ functions in lagging strand synthesis. Pol ε travels with replication forks fired from early replicating origins suggesting a position on the leading strand (Dua et al. 2002, Aparacio et al. 1997, 1999). Pol δ was found to be more active in late S phase cells consistent with a model of delayed lagging strand replication (Rytkönen et al. 2006).

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Furthermore inhibition of pol ε activity decreased replication more severely in early S phase.

Fig. 2. Replication protein assembly on the leading and the lagging strands. In this model GINS links the leading strand polymerase pol ε tightly to the helicase while pols α and δ act on the lagging strand more freely. The figure was adopted from Nasheuer et al. 2006.

Analysis of pols δ and ε exonuclease activities also places the polymerases on opposite strands. Exonuclease deficient pol2exo- and pol3exo- yeast mutants were exposed to 6-N-hydroxylaminopurine (HAP) induced mutagenesis (Shcherbakova & Pavlov 1996). HAP is a nucleotide analogue capable of base-pairing with thymidine and cytosine causing either GC-AT or AT-GC transitions depending on whether HAP exists as the incoming or template nucleotide. There was a significant difference in the missense mutation rate in the ura3 gene analysed depending on the position of the ura3 relative to the adjacent replication initiation site in a pol2-exo- strain. Relative to the pol2-exo-, the position of the ura3 gene showing a high mutagenesis rate was on the opposite side of the origin in the pol3-exo- mutant suggesting that the exonuclease activities of pol δ and ε scan opposite strands of the DNA replication fork. The same results were obtained when the tRNA gene was analysed in pol2-exo- and pol3-exo- mutants (Karthikeyan et al. 2000). Anyhow, the same strand specific bias also seems to exist in Escherichia coli where a single polymerase replicates all DNA (Fijalkowska et al. 1998). In Xenopus DNA replication decreases if cell extracts are depleted of either pol δ or pol ε suggesting that both polymerases act on replication (Fukui et al. 2004). Accumulation of single-stranded DNA gaps in the pol δ mutant indicates a role for pol δ in lagging strand Okazaki fragment synthesis.

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Pol α and δ are required for telomere replication in which telomerase forms the template DNA, and DNA replication consists only of lagging strand DNA synthesis (Diede & Gottschling 1999). This leaves the leading strand replicase role for pol ε, reinforced by the finding that Pol ε does not idle at a nick, a special mode of action needed for Okazaki fragment maturation (Garg et al. 2004). Idling DNA synthesis is supported by pol δ. Furthermore most rad27 mutants of S. cerevisiae encoding defective FEN1 are synthetically lethal with pol δ mutants and the viable double mutants show accumulation of small duplications resulting from defects in Okazaki fragment maturation (Jin et al. 2001, 2005). There is also genetic data showing that pol δ but not pol ε may proofread DNA synthesised by pol α (Pavlov et al. 2006). Finally pol δ interacts with pol α suggesting that these polymerases act in close proximity, a requirement for lagging strand DNA synthesis (Johansson et al. 2004).

2.7 Maturation of Okazaki Fragments

Lagging strand DNA synthesis consists of RNA primer synthesis by pol α primase activity, elongation of the primer by pol α DNA polymerase activity, switch of the primer terminus to pol δ, and further elongation followed by Okazaki fragment maturation (reviewed by Garg & Burgers 2005a). PCNA is the candidate protein for Okazaki fragment processing coordination and may enable the recycling of processing components (Sporbert et al. 2005). If leading and lagging strands are synthesised simultaneously there are only three seconds for each Okazaki fragment to be primed, elongated and matured assuming that Okazaki fragments are 100 – 200 nucleotides long, and forks advance 50 nucleotides per second (Raghuraman et al. 2001). The time consumed for RNA primer synthesis has not been measured in eukaryotes, but in E. coli, the primer synthesis initiation alone takes approximately 45 seconds (Johnson et al. 2000). Eukaryotic pols α and δ replicate 50 nt/s and Okazaki fragment maturation lasts an average of 16 seconds in vitro (Raghuraman et al. 2001, Ayyagari et al. 2003). This is much slower than the assumed in vivo rate. If the leading and the lagging strand DNA synthesis are not tightly linked together, Okazaki fragments can be synthesised in a distributive way, several at the same time, allowing enough time for each Okazaki fragment to be synthesised. However, the mean single-stranded DNA region in the yeast lagging strand is just 220 nucleotides revealed by electron microscopy (Sogo et al. 2002). Furthermore DNA adjacent to a single stranded region DNA was wrapped around nucleosomes indicating completed DNA synthesis and ligation. Therefore the number of unfinished Okazaki fragments behind a replication fork seems limited.

Maturation of Okazaki fragments is critical because unfinished fragments will result in double strand breaks during the next S phase. The Pol δ – FEN1 complex is able to maintain a ligatable nick (Jin et al. 2003, Garg et al. 2004). Pol δ alone is capable of 2 to 3 nucleotides long strand displacement DNA synthesis. This is a reversible process in which pol δ rapidly degrades the newly synthesised DNA using its 3’to 5’ exonuclease activity resulting in zero net DNA synthesis. Therefore it is called idling DNA synthesis. In the presence of FEN1, idling is inhibited. Residual idling synthesis produces DNA in one nucleotide fragments maintaining a ligatable nick for DNA ligase I. DNA ligase I is

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soluble and not present in Okazaki fragment maturation complexes (Ayyagari et al. 2003). Still the ligation is a very efficient process because the PCNA - pol δ - FEN1 complex advances only a few nucleotides past the RNA-DNA junction before nick translation is terminated by a ligation.

DNA ligase I encircles the DNA during the binding process, partially unwinds the nicked dsDNA area, and stabilises the structure while positioning itself on a nick and joining the two DNA ends together (Pascal et al. 2004). Cells lacking DNA ligase I activity are viable because DNA ligase I activity can be complemented by other ligases in chromosomal DNA replication (Prigent et al. 1994).

Dna2 is needed for Okazaki fragment maturation (Budd et al. 2000, Lee et al. 2000, Kao et al. 2004). It is able to cut single-stranded DNA with secondary structure which inhibits the activity of FEN1. These stretches result from problems in Okazaki fragment maturation and are so frequent that DNA2 is an essential gene (Budd & Campbell 1997, Bae et al. 2001, Ayyagari et al. 2003, Kao et al. 2004). The cooperative role of Dna2 in Okazaki fragment maturation is illustrated in the lethal pol3-exo- rad27 double mutant, which is rescued by overexpression of DNA2 (Jin et al. 2003). On the other hand the temperature sensitivity of the dna2-1 mutant is suppressed by overexpressing the FEN1 encoding gene RAD27 (Budd & Campbell 1997).

RNaseH is probably an auxiliary enzyme in Okazaki fragment maturation (Arudchandran et al. 2000). It is able to degrade RNA annealed to complementary DNA, but the loss of RNaseH activity has only a minor effect on nuclear DNA replication (Filippov et al. 2001, Cerritelli et al. 2003). RNaseH activity is essential in multicellular organisms because the activity is required for mitochondrial DNA replication. It seems that mitochondrial replication is significantly different from nuclear DNA replication, probably containing long RNA-DNA hybrids as intermediates, and thus making RNaseH essential (Gaidamakov et al. 2005).

2.8 Regulation of DNA replication

To transfer the genetic information from mother to daughter cells without any alterations DNA must be replicated once and only once during each cell cycle. To achieve this the ORC is destabilised in S phase by weakened DNA binding and ubiquitinylation of Orc1 establishing a rereplication prevention mechanism at the level of origin recognition in Xenopus and human cells (Romanowski et al. 1996, Kreitz et al. 2001, Li & DePamphilis 2002). On the other hand, yeast and hamster Orc1 binds to origins in all cell cycle stages (Bell & Stillman 1992, McNairn et al. 2005). The binding in hamster cells is a highly dynamic process illustrated by a fluorescence loss in a photo bleaching experiment making the achieved results questionable in other vertebrates. Rather, phosphorylation of Orc proteins by increased Cdk activity at the onset of S phase is the means for inhibiting the rereplication capability of ORC (Vas et al. 2001). Orc2 inactivation in S. pombe is an example of this control mechanism.

After ORC loading, Cdc6 and Cdt1 are sequestered at origins and both of them reinforce rereplication inhibition. At the onset of S phase Cdc6 is phosphorylated, followed by degradation in yeast, or export from the nucleus and degradation in higher

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eukaryotes (Drury et al. 2000, Petersen et al. 2000, Delmolino et al. 2001). Cdc6 physically interacts with the ORC modulating its DNA binding activity and Cdc6 also binds to MCMs recruiting them to origins cooperatively with Cdt1 (Speck et al. 2005). Thus removal of Cdc6 at the onset of S phase prevents further MCM loading and origin activation. The other MCM loader Cdt1 is degraded in S. pombe, Xenopus and human during S and G2 phases and the remaining small amount of Cdt1 is inactivated by an inhibitory protein, geminin, in higher eukaryotes (Gopalakrishnan et al. 2001, Li & Blow 2005, McGarry et al. 1998, Wohlschlegel et al. 2000). Geminin prevents MCM binding to Cdt1 by steric hindrance (Lee et al. 2004). Subsequently geminin is degraded during M phase to allow formation of preRCs in the next G1 phase (McGarry et al. 1998, Tada et al. 2001). In S. cerevisiae, the amount of nuclear Cdt1 is reduced by nuclear export (Tanaka et al. 2003). The importance of the low level of Cdt1 in the nucleus was shown in a Xenopus replication system where addition of Cdt1 in G2 phase nuclei induced DNA replication (Maiorano et al. 2005).

The activity of cyclin dependent kinases increases when cells enter S phase and this prevents further preRC formation inhibiting rereplication (Coverley et al. 1998, Ballabeni et al. 2004). An increase in Cdk activity results in the phosphorylation of several components of the pre-RC like Cdc6, Orc2 and also MCMs (Nguyen et al. 2001). On the other hand, Cdc6 phosphorylation by Cdks protects it from degradation and is required for preRC formation when quiescent cells enter the cell cycle (Mailand & Diffley 2005). PreRC formation is prevented in quiescent cells because of the low level of Cdc6 and phosphorylation regulated stabilisation of Cdc6 determines a start point for preRC formation.

Many replication initiation proteins like Sld2 require phosphorylation for their activity (Masumoto et al. 2002). The phosphorylated form of Sld2 is able to form a complex with Dpb11 and promote DNA replication initiation. Sld2 is a target protein for S phase cyclin dependent kinases and therefore represents one step in Cdk-activity regulated progression of the cell cycle. Cdk inactivation during mitosis allows the formation of new preRCs during the next G1 phase (Noton & Diffley 2000).

Mcm8 is a new member of the replication regulation related proteins being present in most but not all eukaryotes (Gozuacik et al. 2003, Maiorano et al. 2005, Blanton et al. 2005). Human Mcm8 participates in regulation of preRC formation as its depletion by the RNA interference technique causes reduced loading of Cdc6 and the Mcm complex and therefore delayed entry into S phase (Volkening & Hoffmann 2005). In Xenopus MCM8 depletion slows down DNA synthesis indicating a role in replication fork progression (Maiorano et al. 2005). Xenopus MCM8 has helicase and ATPase activities in vitro and it colocalises with replication forks in vivo suggesting a role as a helicase. It has other functions as well, because Drosophila offspring missing that gene show high levels of chromosome disjunction resulting from compromised meiosis (Blanton et al. 2005). Otherwise DNA replication is normal indicating that there is species to species variation in the function of Mcm8.

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2.9 Transcription and DNA replication

Eukaryotes have three RNA polymerases (reviewed by Paule & White 2000). RNA pol I synthesises ribosomal RNA and RNA pol III produces several small nuclear RNAs, all transfer RNAs needed for protein synthesis, and 5S ribosomal RNA. RNA pols I and III produce over 80 % of all RNA in cells while RNA pol II transcribes all protein encoding genes.

The gene for the human pol ε B-subunit can be viewed as a model of how transcription proceeds in eukaryotes (Huang et al. 2001). Genes have a coding region which contains the template for pre-mRNA production and after RNA processing the constructed messenger-RNA is translated to produce a protein. In addition to the coding sequence with interrupting intron areas, genes have a promoter region regulating the expression. Like most other DNA replication genes, the promoter for the pol ε B-subunit is GC-rich and contains several binding sites for transcription factors Sp1 and E2F. These transcription factors are able to recruit histone modifying enzymes to DNA and finally RNA pol II with the basal transcription factors. Many genes involved in housekeeping functions have a TATA-sequence in the promoter region, which acts as a loading site for RNA pol II, but many replication linked genes, like the pol ε B-subunit, do not have this sequence.

Transcriptionally active areas in the genome are likely to be replicated in early S phase and the RNA pol II complex, more precisely CTD, activates replication and probably mediates coordination between replication and transcription (MacAlpine et al. 2004, Gauthier et al. 2002). Transcription factors are able to regulate replication (reviewed by Kohzaki & Murakami 2005). The regulation may be indirect via modulation of chromatin structure and the availability of DNA binding sites or direct via recruiting protein complexes to DNA.

One of the best characterised origins is ARS1 in S. cerevisiae (Kohzaki et al. 1999). It contains an area called a B-element, which is a binding site for the transcription factor Abf1. Various transcription factors binding to this site repress or activate the DNA replication initiation activity of ARS1. The human c-MYC gene has a 2.4 kb long promoter region with several replication initiation sites (Ghosh et al. 2004). Deletion of transcription factor binding sites from this promoter abolished replication initiation. The addition of a GAL4-CREB transcription factor binding site restored the capacity to initiate replication indicating that transcription factors modulate replication.

2.9.1 Regulation of transcription

Nucleosomes pack DNA and act as general transcription repressors by restricting the accessibility of transcription factors to DNA. Active promoters usually contain modified histones and partially unfolded DNA to allow easier access for transcription factors (Boeger et al. 2004). Modified nucleosomes bind to DNA in a highly dynamic way by completely leaving the active promoter and then reassembling onto it. This results in a partially open DNA.

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Transcription factors c-Myc and Miz-1 can be viewed as examples how transcription factors act on promoter regulation. The transcription factor Myc/Max binds to DNA sequences termed E boxes in the promoter region and recruits the coactivator TRRAP complex with histone acetylation ability to activate the promoter (McMahon et al. 1998, reviewed by Eisenman 2001). Interactions with Max and TRRAP are critical for Myc-induced immortalisation of cells indicating that the transcription activation function is required (Park et al. 2001).

A hallmark of Myc-induced transformation is the inability to respond to anti-mitogenic signals (Warner et al. 1999). Myc induces both expression of several cell cycle promoting genes and downregulation of cyclin inhibitors (Warner et al. 1999, Claassen & Hann 2000). Keratinocytes do not respond to TGF-β induced growth arrest because of Myc dependent repression of the cyclin inhibitors p15INK4B and p21cip1. In primary cells Miz-1 binds to a core promoter of the p15INK4B gene with a p300 coactivator, but Myc represses p15 expression by competing with Miz-1 binding (Staller et al. 2001). The Miz-1/p300 activation complex also contains Smad3 and Smad4 transcription factors that bind to their own DNA sites on the p15 promoter acting cooperatively with Miz-1 in promoter activation (Seoane et al. 2001).

Many DNA replication and repair proteins act as positive or negative regulators of transcription (Yankulov et al. 1999, Mo & Dynan 2002, Krum et al. 2003). These include Mcm2, breast cancer associated protein Brca1 and an essential DNA double-strand break repair protein Ku70/Ku80 complex. The interaction deficient dominant negative Ku80 mutant showed decreased transcription levels and suppressed cell growth linking transcription and double-strand break repair together (Mo & Dynan 2002). Mcm2 is able to bind the C-terminal region of RNA pol II and antibodies against Mcm2 inhibited transcriptional activity of RNA pol II indicating a functional interaction (Yankulov et al. 1999).

2.9.2 RNA polymerase II

The RNA pol II core consists of a 12-subunit polymerase capable of RNA synthesis and proofreading of the transcript, five basal transcription factors TFIIB, -D, -E, -F, and –H mediating promoter recognition and promoter unwinding, and a 20-subunit Mediator complex which regulates the activity of RNA pol II (Kelleher et al. 1990, Kim et al. 1994, reviewed by Boeger et al. 2005). The RNA pol II structure has been resolved (Westover et al. 1994). RNA pol II contains an active cleft in which the DNA template is unwound having a 9 base pairs long RNA-DNA hybrid as a transcript construction intermediate. The eukaryotic RNA pol II structure is completely different from viral and bacterial RNA pols indicating an independent origin of these enzymes. The mediator complex forms an envelope around the RNA pol II/basal transcription factor complex forming many points of contact, which facilitate the regulatory function of Mediator (Asturias et al. 1999).

The C-terminal domain of RNA pol II (CTD) mediates the interactions of RNA pol II with other proteins (reviewed by Palancade & Bensaude 2003). CTD contains a species specific amount of a heptapeptide with a consensus YSPTSPS. Most RNA pol II

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molecules contain an unphosphorylated form of this heptapeptide and only the unphosphorylated form is capable of binding to a promoter region. Shortly after transcription initiation, serine 5 of the heptapeptide becomes phosphorylated in a transcription factor II H-dependent manner (Cho et al. 2001, Kim et al. 2002). During transcription elongation serine 5 is dephosphorylated and serine 2 becomes phosphorylated. After transcription termination CTD is dephosphorylated and a new round of transcription is enabled. Phosphorylated CTD also harvests pre-mRNA processing machinery to unfinished pre-mRNA and pre-mRNA modification can be initiated during ongoing transcription (Carty & Greenleaf 2002)

RNA pol II interacts with several DNA repair factors including pol ε, Ku, Rad51, RPA, RFC, Brca1, and the Brca1 associated ring domain protein 1, linking DNA repair and transcription together (Maldonado et al. 1996, Neish et al. 1998, Krum et al. 2003, Chiba & Parvin 2002, Yankulov et al. 1999, Holland et al. 2002, Mo & Dynan 2002, reviewed by Tornaletti 2005). RNA pol II is known to act as a DNA damage sensor in the transcription-coupled DNA repair pathway. RNA pol II pauses at a transcription blocking DNA lesion and starts a signalling cascade resulting in lesion repair. Most of these lesions are repaired via the nucleotide excision repair pathway. The need for coordination of DNA replication and transcription arises from progression of both replication forks and the trancription apparatuses on the same DNA template. Finally it must be noted that the composition of the RNA pol II holoenzyme depends on the purification method applied and many factors interacting with the RNA pol II holoenzyme can not be viewed as subunits of the holoenzyme (Neish et al. 1998, Holland & Yankulov 2003).

2.10 DNA repair pathways

The size of the human genome is 3 x 109 base pairs and because of this, DNA is an unstable molecule. The bulk of DNA damage results from spontaneous deamination of cytocine and oxidative damage caused by reactive oxygen species linked to cellular respiration (reviewed by Sung & Demple 2006, Wilson & Thompson 1997, Seeberg et al. 1995). It is estimated that 10 000 purines become hydrolysed in a cell each day. In addition to that, alkylating agents produce hundreds of 3-alkyladenines per day and the lipid metabolism side product epoxyaldehyde also damages DNA constantly. Cells have damage avoidance mechanisms, for example detoxification of reactive oxygen species, but there are still enough modified bases to make all components of the base excision repair (BER) pathway essential. There are actually no diseases linked to BER, propably because intrinsic base modifying agents and spontaneous deamination make all mutations in BER genes lethal. Most BER substrates arise from intrinsic sources while environmental sources contain γ-irradiation induced free radicals and harmful agents in food, for example alkylating nitrosoamines arising from bacteria processing nitrates and nitrites in the intestines.

BER is initiated by a DNA glycosylase catalysed deamination and completed via alternative routes requiring either DNA polymerase β activity or PCNA with either pol δ or pol ε (reviewed by Sung & Demple 2006). Cells have many DNA glycosylases recognising specific types of DNA damage, of which UNG2 is the major uracil-DNA

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glycosylase (Otterlei et al. 1999). BER is linked to replication as shown by upregulation of UNG2 during S phase and colocalisation to sites of DNA synthesis. A small fraction of DNA damage arises from the replication process itself but most of it is repaired via the mismatch repair pathway following replication (reviewed by Jun et al. 2006). The third excision pathway is nuclotide excision repair.

DNA double strand breaks (DSBs) and interstrand cross-links are caused by γ-irradiation or chemical stress (reviewed by Pastwa & Blasiak 2003 and Ishino et al. 2006). They are repaired via homologous recombination repair (HRR) or non-homologous end joining (NHEJ). HRR needs sister chromatin as a repair template, therefore acting in S and G2 phase cells, whereas NHEJ repairs DNA in all cell cycle stages. The NHEJ pathway can yield a direct ligation of two DNA ends or it can cause modifications to DNA, usually short insertions, while HRR can preserve the genetic information completely. NHEJ is the main DSB repair pathway in humans whereas HRR is more frequently utilised in yeast (Ferguson et al. 2000, Moore & Haber 1996).

Recombination is also needed for other processes than repair. During replication HRR helps the stalled replication forks to get past the blocking lesion (reviewed by Ishino et al. 2006). Homologous recombination is also used to relieve linkage between genes located in the same chromosome in a process called meiotic recombination (reviewed by Shinohara & Ogawa 1995). NHEJ is used for somatic hypermutation required for immunoglobulin and immunoglobulin receptor production (reviewed by Pastwa & Blasiak 2003).

Repair pathways share some components with each other and the same damage can be repaired via several pathways linking the repair systems together (Klungland et al.1999). For example the XPG nuclease working in nucleotide excision repair can also recognise modified bases to assist in the BER pathway.

2.10.1 Nucleotide excision repair

Nucleotide excision repair (NER) removes lesions that distort DNA structure and Watson-Crick base pairing (Balajee et al. 1999, reviewed by Batty & Wood 2000). Damage recognition does not require any specific damage, which allows repair of a wide variety of lesions. Ultraviolet light from the sun causes (6-4) photoproducts and cyclobutane thymine dimers on DNA, which are the main substrates for NER. Other substrates are covalently DNA bound polycyclic aromatic compounds and DNA interstrand crosslinks. Base excision repair pathway substrates, for example oxygen free radical and alkylating agent caused base modifications, can also be repaired by NER to some extent (Satoh et al. 1993). Nucleosomes protect DNA from radiation but some areas in DNA are naturally distorted, like promoter regions after TATA binding protein caused kinking, and are easily damaged by UV light (Gale et al. 1987 Aboussekhra & Thoma 1999).

The eukaryotic NER pathway is well known because several human cell lines are available having defects in NER (reviewed by Lehmann 2003). Patients having Xeroderma pigmentosum (XP) are sensitive to sunlight, suffer from accelerated ageing of skin and eye, and are predisposed to cancer, especially skin cancer. Defects in the neural

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system affect 18 % of patients. Before NER genes were cloned, patients were divided into seven groups (XPA-XPG) based on cell fusion caused defect complementation. After identifying the mutation behind each complementation group the genes were named based on this linkage. XPV (Xeroderma pigmentosum variant) cells differ from other XP cells having a fully functional NER pathway, but are defective in the replication of UV irradiated DNA (Masutani et al. 1999). XPV cells encode a DNA polymerase η capable of effective and accurate bypass of thymidine dimers. The sensitivity to UV light arises from defective replication caused by accumulation of double strand breaks and genome rearrangements (Limoli et al. 2000).

Approximately 40 genes are directly involved in NER and the repair pathway has been reconstituted in vitro from human and yeast (Wood et al. 2005, Araujo et al. 2000, Guzder et al. 1995). Damage recognition is the first step in the repair process. Actively transcribed genes are repaired faster than heterochromatin because RNA polymerase II is able to detect damage and start a signaling cascade (Donahue et al. 1994, Tornaletti et al. 1999). RNA pol II stalls on the DNA damage and withdraws by digesting a short piece of the transcript while remaining DNA bound. Transcription elongation factor SII helps in this process. Based on damage recognition mechanisms, NER can be divided to two pathways, namely transcription coupled repair (TCR) and global genome repair (GGR). In GGR the main damage sensor is the human XPC-HR23B or the yeast ortholog, the Rad4/Rad23 complex (Sugasawa et al. 1998, Guzder et al. 1998). The damage detector is able to slide along DNA consuming ATP and probably dissociating from DNA when sliding is blocked by a nucleosome, followed by association at another site (Guzder et al. 1999 Svetlova et al. 1999). This partial scanning of DNA may be enough to locate all NER substrates because distortion of DNA structure facilitates the sliding of nucleosomes away from the lesion resulting in a damage site between nucleosomes (Whitehouse et al. 1999). XPE consists of damage-specific DNA binding proteins 1 and 2 and participates in damage detection as a helper protein, recognising structures to which XPC has a low affinity (Nichols et al. 2000, Tang et al. 2000). Its role as an accessory protein results in a mild phenotype of XPE compared to other XP patients (reviewed by Lehmann 2003).

After damage detection TCR and GGR use the same mechanism to process the DNA damage. Damage detectors XPC, XPE and RNA pol II facilitate binding of XPA to the DNA damage site (Sugasawa et al. 1998). XPA acts as a coordinator in the NER pathway (Li et al. 1995, Robins et al. 1991). It forms a complex with RPA and the complex has a higher affinity to damaged DNA compared to XPA alone. With this damaged-DNA binding capability XPA can verify the damage detection done earlier in the pathway. Furthermore XPA interacts with the general transcription factor TFIIH recruiting it to DNA (Park et al. 1995). TFIIH binds also to XPC/HR23B and is needed to open DNA around the damage (Drapkin 1994, Evans et al. 1997). TFIIH contains XPB with 3’→5’ and XPD with 5’→3’ helicase activities to open a 24-32 nucleotides long stretch on DNA. The TFIIH complex also determines the strand to be removed. TFIIH involvement in NER links repair and cell cycle regulation together because three members of TFIIH, namely Cdk7, Cyclin H, and Mat1, form the mammalian cyclin-dependent kinase activating kinase (Araujo et al. 2000). It phosphorylates several Cdks and also phosphorylates the RNA pol II CTD when transcription proceeds from the initiation to the elongation phase (Rossi et al. 2001, reviewed by Kaldis 1999).

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XPA/RPA also interacts with the endonucleases XPG and XPF/ERCC1 catalysing damage removal (de Laat et al. 1998). XPG cuts DNA 2-9 nucleotides away from the damage on the 3’-side (O’Donovan et al. 1994, de Laat et al. 1998). XPF/ERCC1 performs 5’-incision 24-32 nucleotides away from 3’-site cut (Evans et al. 1997, de Laat et al. 1998). RPA controls the length of the DNA to be removed by binding ssDNA revealed by TFIIH (Lao et al. 2000).

NER requires RFC and PCNA for DNA synthesis indicating pol δ and/or pol ε involvement in the pathway (Araujo et al. 2000). DNA ligase I is probably the only enzyme capable of nick sealing in NER because the DNA ligase I deficient S. cerevisiae mutant is unable to perform NER (Wu et al. 1999).

Patients having Cockayne's syndrome (CS) suffer from severe physical and mental retardation and sensitivity to sunlight (reviewed by Lehmann 2003). They are not prone to skin cancer like XP patients, however, indicating a different defect in NER. Cells from patients form two complementation groups and the hallmark of these CSA and CSB cells is the inability to recover RNA synthesis after UV treatment (van Oosterwijk et al. 1996). In CS cells RNA pol II remains in a hypophosphorylated non-transcribing form for long periods of time after a UV pulse (Rockx et al. 2000). Both CSA and CSB interact with RNA pol II and assist RNA pol II to pass a secondary structure containing template or to dissociate from DNA facilitating NER (Groisman et al. 2003, van Gool et al. 1997, Tantin et al. 1997). CSB is also linked to RNA pol I and ribosomal RNA synthesis (Bradsher et al. 2002). Although TCR is compromised in CS cells it seems that symptoms of CS patients are rather a consequence of defects in transcription (reviewed by Lehmann 2003). Some XPA and XPG patients lack the NER pathway completely but suffer only from skin problems indicating a different mechanism causes the symptoms of CS patients. On the other hand some defects in XPB, XPD and XPG genes can cause either XP or combined CS/XP disease.

A third inherited disorder linked to NER is trichothiodystrophy (TTD), causing physical and mental retardation like CS and brittle hair (reviewed by Lehmann 2003). There are three complementation groups, TTDA with an unknown gene, TTD/XPB and the most common TTD/XPD. Involvement of the XPB and XPD genes suggest that also in TTD transcription is compromised. In fact all mutations causing XPD disease affect the endonuclease activity or the regulation of XPD, whereas in TTD/XPD cells, the basal transcription activity of TFIIH is affected (Winkler et al. 2000, Dubaele et al. 2003).

3 Aims of the present work

DNA replication initiation is a well characterised process while the elongation phase studies have been problematic. In addition, transcription and many individual DNA damage processing pathways are defined to the level in which in vitro reconstitution is possible. In contrast, the linkage between transcription, damage response and replication is poorly understood. The general goal of this study was to find links between transcription, damage response and replication by identification of factors interacting with pol ε followed by functional analysis of the interactions. The specific aims of the present work were:

1. Identification, cloning, and structural characterisation of a potential human homolog of yeast Dpb11.

2. Functional analysis of the human Dpb11 homolog TopBP1 including identification of interaction partners.

3. To study whether DNA polymerase ε interacts with RNA polymerase II (as has been previously proposed) and transcripts.

4. To study the involvement of Cdc45 in DNA replication by analysing its interaction with DNA polymerase δ and DNA polymerase ε during the cell cycle.

4 Materials and methods

4.1 cDNA cloning (I)

An expressed sequence tag database was searched to find EST clones containing parts of the human cDNA sequence KIAA0259 (GenBank accession D87448). Clones AA013344, AA195149, and R54257 (Genome Systems, Inc. USA), together with nucleotides 308–1560 and 1628–2620 amplified from human T-cell (HUT-78) and thymus cDNA libraries (CLONTECH) were used to construct full length TopBP1 cDNA.

4.2 Gene chromosomal mapping (I)

A 2062-base pair cDNA probe was used for chromosomal mapping of TopBP1 by fluorescence in situ hybridization (FISH). The analysis was done by SeeDNA Biotech Inc (Canada).

4.3 Production of antibodies (I, II)

Antigens were prepared using a GST gene fusion system (Amersham Pharmacia Biotech) according to Pospiech et al. (1999) and rabbits were immunised using standard protocols. Rabbit antiserum α-TopBP1.1 was raised against peptide containing amino acids R792-A1003 and α-TopBP1.2 against D1023-P1167. The antibodies were purified by protein A-Sepharose CL-4B (Amersham Pharmacia Biotech) and the specificity of the antibodies was tested by Western analysis of human HeLa S3 cell extracts and by antigen block assays. The immunofluorescence stainings were validated by showing colocalisation between our polyclonal antibodies and mouse α-TopBP1 T10620 (Becton Dickinson).

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4.4 Cell lines and synchronisation (I, III, IV)

HeLa S3 cervix adenocarcinoma cells (ATCC CCL2.2) were cultured in suspension in Joklik’s modification of minimal essential medium (Flow Laboratories) supplemented with sodium bicarbonate and 5 % calf serum. HeLa CCL-2 cervix adenocarcinoma monolayer cells and MCF-7 epithelial adenocarcinoma cells (ATCC HTB-22) were grown in Dulbecco’s modified Eagle’s medium (Gibco Life Technologies) containing 10 % foetal bovine serum and antibiotics. IMR-90 normal lung fibroblasts (ATCC CCL-186) and T98G brain glioblastoma cells (ATCC CRL 1690) were cultured in Eagle’s minimum essential medium (Sigma) supplemented with 10 % foetal bovine serum, glutamax, non essential amino acids, sodium pyruvate and antibiotics. Hela S3 cells were grown at 37 °C in a normal atmosphere and all other cells at 37 °C in a 5 % carbon dioxide atmosphere.

Cells were arrested in the G0 phase using serum deprivation (0.25 - 0.5 % serum). T98G cells were deprived for six days, IMR-90 cells for four days, and MCF-7 cells for one day (Rytkönen et al. 2006, Tuusa et al. 1995, Scully et al. 1997a). Entry into the cell cycle was induced by adding balanced, complete growth medium. Propidium iodine staining of DNA and flow cytometry assays were used to determine the quality of the synchrony (Vindelöv et al. 1983). Synchrony of IMR-90 and MCF-7 cells during S phase were analysed by incorporation of [3H]-thymidine in parallel cell cultures and by punctate PCNA staining.

4.5 Induction of DNA damage (I, III)

DNA replication was inhibited by 1 mM hydroxyurea (HU). To induce double-strand breaks DNA was damaged using 100 μg/ml zeocin, alkylating DNA damage was caused with 50 μg/ml methyl methanesulphonate (MMS), and 10 J/m2 ultraviolet light induced DNA damage was produced using UV hand lamp (UVP model UVGL-58). Samples were collected one hour after induction of the damage.

To irradiate only parts of T98G cell nuclei, cells were grown to logarithmic phase on glass slides, washed briefly with PBS and exposed to 60 J/m2 UV light through a 8 μm micropore filter (Millipore). Cells were collected after one hour of recovery.

4.6 Preparation of cell extracts (I, III)

HeLa CCL-2 cells were washed with PBS and collected with a cell scraper and 1 ml of lysis buffer [80 mM NaCl, 10 % glycerol, 0.1 % Nonidet P-40, Complete protease inhibitors (Roche), 10 mM Na3VO4, 10 mM NaF, and 100 mM Tris-HCl, pH 7.5]. T98G cells were collected in lysis buffer [100 mM KCl, 5 % glycerol, 0.1 % NP-40, Complete protease inhibitor (Roche), 1 mM EDTA, 10 mM NaF, 2 mM Na3VO4, and 50 mM Tris-HCl pH 7.5]. Cell suspensions were sonicated twice for 15 seconds, equilibrated on ice for 15 minutes and centrifuged for 10 min at 20 000 g and snap frozen.

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HeLa S3 cell nuclei and cytoplasmic extracts were prepared as described by Stoeber et al. (1998).

4.7 T98G cell fractionation (IV)

To prepare cytosolic, DNA bound, and nuclear matrix bound T98G extracts, cultures were cooled to +4 °C, followed by washing with cold TBS twice (150 mM NaCl, 20 mM Tris-Cl, pH 7.5) and twice with hypotonic KM buffer (10 mM NaCl, 1 mM MgCl2, 2 mM DTT and 10 mM MOPS-NaOH, pH 7.0). Cells were lysed with 1 ml KM buffer containing 0.5 % Nonidet P-40 and Complete EDTA-free protease inhibitors (Roche) for half an hour. The resulting extract containing detergent-soluble proteins was collected and snap frozen. Cell remnants were washed twice with KAc buffer (5 mM K-acetate, 0.5 mM MgCl2, 2 mM DTT and 30 mM Hepes-KOH pH 7.4), followed by incubation in 1 ml of DNase I solution (150 mM NaCl, 1.5mM CaCl2, 6mM MgCl2, Complete EDTA-free protease inhibitors, 50 μg/ml RNase A, 50 U/ml DNase I and 40mM Tris-Cl pH 8.0) at 22 °C for 30 minutes. The extract released by DNA digestion contained DNA bound proteins. The remaining cell matrix was washed twice with DNase buffer (DNase I solution without DNase I and RNase A). Matrix bound proteins were solubilised in lysis buffer (100 mM NaCl, 0.5 mM MgCl2, 0.5 mM DTT, 5 mM KCl, 0.5 % SDS and 20 mM Hepes-KOH, pH 7.7), collected with a cell scraper and snap frozen. A total cell extract was prepared by washing a parallel 100 mm plate twice with cold TBS followed by addition of lysis buffer and extract collection with a cell scraper.

4.8 Immunoprecipitation (I, III, IV)

For each immunoprecipitation (IP), 200-300 μg cell extract and 2-4 μg antibody were used. Proteins binding unspecifically to antibodies or protein G were removed with either 2 μl of preimmune rabbit serum or with 10 μg of non-specific mouse IgG (Pierce) in combination with GammaBind protein G Sepharose (Amersham Biosciences). Specific IP was done over night. Pellets were washed twice with wash buffer (150 mM NaCl, 10 % glycerol, 0.1 % NP-40, and 50 mM Tris-Cl pH 7.5), twice with high salt buffer (250 mM NaCl, 10 % glycerol, 0.1 % NP-40, and 50 mM Tris-Cl pH 7.5), and eluted with SDS-PAGE loading buffer.

4.9 SDS-PAGE and Western-analysis (I, III)

Proteins were separated by 6 % SDS-PAGE and transferred to a PVDF membrane (Millipore). Detection was based on chemiluminescence or alkaline phosphatase based color formation and results were quantified by densitometric scans of several Western blots.

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4.10 Indirect immunofluorescence (I, III, IV)

Cells were grown on glass slides, cleaned from growth media with three Eisen PBS washes (150 mM NaCl, 1.86 mM NaH2PO4, 12.64 mM Na2HPO4, pH 7.4), and water was removed with brief -20 °C methanol wash. Permeabilisation was done with -20 °C methanol, fixation with fresh 3 % paraformaldehyde (PFA) and quenching with 50 mM NH4Cl in PBS, each treatment for 10 minutes. Unspecific binding of antibodies was blocked using 0.2 % cold water fish skin gelatine (Sigma) for at least one hour. Antibodies were diluted at 5 μg/ml in the blocking solution. Labelling was performed sequentially with primary and secondary antibodies for 30 min at 37 °C. Bisbenzimide (Hoechst 33258, Sigma) was used for DNA labelling and Immu-Mount (Shandon) for mounting.

To visualise recently synthesised DNA,cells were grown with 0.1 mM BrdU (Sigma) for 5 minutes. The staining protocol was the same as above except for the primary antibody treatment which was performed in digestion buffer (150 mM NaCl, 1.5mM CaCl2, 6 mM MgCl2, 0.2% gelatin, 2 U/ml Dnase I and 40mM Tris-Cl pH 8.0) for 30 minutes at 37 °C to reveal incorporated BrdU. Other methods to reveal BrdU were 2 M HCl treatment for 15 minutes at 37 °C or 4 M HCl for 10 minutes at 22 °C.

Other permeabilisation and fixation methods used were fixation with 3 % PFA followed by permeabilisation using 0.2 % triton X-100, fixation with 3 % PFA followed by permeabilisation using cold methanol, permeabilisation with 0.2 % Triton X-100 followed by fixation with 3 % PFA, permeabilisation and hypotonic extraction with KM buffer containing NP-40 (10 mM NaCl, 1 mM MgCl2, 2 mM DTT, 0.5 % Nonidet P-40, Complete EDTA-free protease inhibitors (Roche), and 10 mM MOPS-NaOH, pH 7.0) for 30 minutes at 4 °C followed by digestion of DNA with 100 U/ml HindIII and PstI for 30 minutes at 37 °C and fixation with 3 % PFA.

To visualise DNA damage caused by UV irradiation through a 8 μm Millipore filter, cells were permeabilised with 0.2 % Triton X-100, fixed with 3 % PFA, and treated with 2 M HCl for 7 minutes at 22 °C to denature DNA.

The amount of unspecific signal was checked by omitting one primary antibody from the double staining. The unspecific signal was negligible in all experiments. Imaging was done either with an Olympus BX-61 microscope with a 60x or 100 x objectives and CCD camera or with a Leitz microscope with a 50x or 60x objective and Kodak Ectachrome 400 film. Adobe Photoshop was used for image processing.

Secondary antibodies were Alexa Fluor 488 goat α-mouse IgG, Alexa Fluor 594 goat α-mouse IgG, Alexa Fluor 488 goat α-rabbit IgG, Alexa Fluor 546 goat α-rabbit IgG, Alexa Fluor 488 goat α-rat IgG (Molecular Probes), fluorescein isothiocyanate-conjugated goat α-BrdU, fluorescein isothiocyanate-conjugated goat α-mouse IgG, and tetramethylrhodamine isomer R-conjugated swine α-rabbit IgG (DAKO).

To determine level of colocalisation, foci from several nuclei in immunofluorsescence images were calculated manually and classified as red, green or yellow. Statistical analysis was done using χ2 and likelihood ratio tests.

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4.11 Immuno-electron microscopy (III, IV)

T98G cells were fixed with 4 % paraformaldehyde for 30 minutes (4 % PFA, 2.5 % sucrose, and 0.1 M phosphate buffer, pH 7.5) Cells were collected using a cell scraper and centrifuged (2000 g for 3 min). The pellet was suspended in 2 % NuSieve agarose (FMC BioProducts), and moderate size block pieces infiltrated with 2.3 M sucrose in PBS.

Cryosections were cut from frozen samples with a Leica Ultracut UCT microtome (Leica Microsystems). To block unspecific signals, sections were treated with 5 % BSA with 0.1 % gelatine in PBS. All antibodies and gold conjugates were diluted in 0.1 % BSA-C (Aurion) in PBS and all washing steps were done with the same solution. All primary antibodies were diluted to 5 μg/μl. The first primary antibody was bound to sections for 60 minutes followed by incubation with secondary antibody and gold particles, both for 30 minutes [1.9 μg/ml rabbit α-mouse IgG (Zymed) or rabbit α-rat IgG (Jackson) and protein A-gold complex, size 5 nm (Slot & Geuze 1985)]. Free binding sites on protein A were blocked with 1 % glutaraldehyde. The second primary antibody was incubated for 60 minutes followed by secondary antibody and protein A-gold complex (size 10 nm) as described above. Amount of unspecific signal was controlled by performing the labelling procedure in the absence of the second primary antibody.

Methylcellulose embedded samples were examined in a Philips CM100 transmission electron microscope (FEI company). Images were recorded by CCD camera equipped with TCL-EM-Menu version 3 from Tietz Video and Image Processing Systems GmbH (Gaunting). Gold particles were manually calculated and classified as centre if three or more particles were in close proximity. Mixed centres contained at least one 5 nm and one 10 nm gold particle.

5 Results

5.1 cDNA cloning and chromosomal localisation of human TOPBP1 (I)

We used the Psi-Blast program to look for the human homolog of S. cerevisiae Dpb11 protein, which is known to interact with pol ε (Araki et al. 1995, Masumoto et al. 2000). A human cDNA, KIAA0259, was identified and predicted to encode a 1522 amino acid protein with eight BRCT domains (Nagase et al. 1996). The same protein was found in a two-hybrid screen looking for DNA topoisomerase IIβ -binding proteins and named TopBP1 (Yamane et al. 1997). Further sequence analysis showed that the BRCT domain pairs 1-2 and 4-5 together with the surrounding sequence are most similar between TopBP1 and Dpb11 (35% similar, 14% identical and 27% similar, 11% identical, respectively) and also between TopBP1 and the S. pombe homolog Cut5 (44% similar, 26% identical and 41% similar, 23% identical, respectively). Dpb11 and Cut5 do not have additional BRCT domains but the Drosophila homolog XTopBP1Mus101 has 7 BRCT domains showing overall 38% similarity and 28% identity with TopBP1 (Yamamoto et al. 2000). The fourth obvious homolog is a predicted F37D6.1 protein in Caenorhabditis elegans having six BRCT domains in three pairs. The C-terminus of Brca1 is needed for tumour suppressor activity and it is 35% similar and 25% identical to the C-terminus of TopBP1 containing the BRCT domain pair 7-8.

A full length TopBP1 cDNA was constructed from EST clones and sequences from a human thymus cDNA library. Based on the sequence homology between TopBP1 and Brca1 the possible link between the TOPBP1 locus and cancer was investigated. The gene is located in chromosome 3 in locus q21-q23 based on the FISH method done with 2 kb probe. This locus was not linked to any chromosomal abnormalities in human cancers according to the OMIM data base.

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5.2 TopBP1 is a phosphoprotein expressed in a proliferation dependent manner (I)

Polyclonal rabbit antibodies were raised against GST-TopBP1 peptides. Both α-TopBP1.1 against amino acids R792-A1003 and α-TopBP1.2 against D1023-P1167 recognise a 180 kDa protein from HeLa cell extracts. This is slightly larger than the calculated molecular mass of 171 kDa and this increase in size is likely due to the phosphorylation of serine and to a lesser extent threonine residues of TopBP1 shown by in vivo phosphate labelling and phosphoamino acid analysis.

In serum starved IMR-90 fibroblasts, TopBP1 protein levels started to increase 14 hours after serum stimulation concomitantly with cells entering into S phase. S phase entry was monitored by the thymidine incorporation method. An RNase protection assay showed an increase in the TOPBP1 transcript level at the S phase entry and a peak 22 hours after the stimulation at the end of S phase. This type of expression profile is common for proteins acting in DNA replication. A similar S phase peak was detected in HeLa S3 cells first synchronized to M phase using mimosine and then allowed to proceed in the cell cycle indicating that high expression of TopBP1 during S phase is not specific for the first round of the cell cycle.

5.3 TopBP1 interacts with DNA polymerase ε and hRad9 (I)

The presence of several BRCT domains suggests that TopBP1 may have several protein-protein interactions. A two-hybrid construct containing BRCT domains 4-5 interacted with hRad9. The Human Rad9 was neither able to interact with BRCT domains 3 or 6 of TopBP1 nor with pol ε, Cdc45, lamin, or Chk1 indicating that the interaction between TopBP1 BRCT domains 4-5 and hRad9 is specific. Full length TopBP1 and hRad9 also interacted in a two-hybrid system. Interaction was confirmed by immunoprecipitation of TopBP1 with FLAG-tagged hRad9 from Ecr-293 cells.

α-TopBP1.2 immunoprecipitations from HeLa S3 cells or from Ecr-293 cells overexpressing TopBP1 contained a hyperphosphorylated form of pol ε in addition to TopBP1. The phosphorylation status of Pol ε was studied by mobility shift loss on SDS-PAGE after phosphatase treatment. The interaction was not cell cycle dependent because all HeLa S3 extracts prepared from cell populations in different cell cycle phases by centrifugal elutriation showed the interaction. The interaction was not affected by DNA damage either as coimmunoprecipitation was detected after hydroxyurea induced replication arrest, alkylating DNA damage by methyl methanesulphonate, double-strand break DNA damage caused by zeocin, and ultraviolet light pulse. The interaction was not mediated through DNA because the addition of ethidium bromide to denature DNA had no effect on the coimmunoprecipitation (Lai & Herr 1992). Under denaturing conditions TopBP1 was precipitated without pol ε indicating that coimmunoprecipitation was not due to crossreaction. Pols α and δ were not present in the immunoprecipitations.

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5.4 TopBP1 is required for chromosomal DNA replication (I)

Because of the interaction between TopBP1 and pol ε we investigated whether TopBP1 is required for chromosomal DNA replication in human cells. α-TopBP1.1 inhibited replicative DNA synthesis 44 % in permeabilised HeLa cell nuclei measured by incorporation of dCMP into chromatin. An even higher level of inhibition, 83 %, was achieved when the TopBP1 R792-A1003 peptide was competing with TopBP1 in the assay. This peptide was used to produce α-TopBP1.1 antisera and contains BRCT domain 6 with some flanking sequence. α-TopBP1.2 or its target peptide D1023-P1167 did not affect replication efficiency in the isolated nuclei, indicating that the sixth BRCT domain of TopBP1 is required for replication.

5.5 TopBP1 and Brca1 colocalise in S phase and become relocalised into PCNA containing foci after replication block (I)

Indirect immunofluorescence staining techniques were used to further investigate the role of TopBP1 in DNA replication. TopBP1 localised into several nuclear foci in MCF-7 cells but did not colocalise with ongoing DNA synthesis visualised either with BrdU or PCNA staining. Brca1 also showed a staining pattern similar to that of TopBP1 (Scully et al. 1997b). Indeed, all Brca1 foci were found to contain TopBP1. Nuclei also had TopBP1 foci without Brca1 because of the greater number of TopBP1 foci. Nuclear localisation of TopBP1 was also detected in IMR-90 cells.

A hydroxyurea induced replication block caused TopBP1 and Brca1 to relocalise extensively into stalled replication forks visualised by PCNA staining. Also the number of foci increased. This kind of action has been reported for Brca1 earlier (Scully et al. 1997b). A 10 J/m2 UV pulse caused 19 % of TopBP1 foci to colocalise with PCNA, while 50 µg/ml methyl methanesulphonate and 200 µg/ml zeocin did not induce colocalisation with PCNA. Interestingly the amount of TopBP1 foci in G1 phase nuclei also increased significantly after zeocin treatment. In this respect TopBP1 differs from Brca1, which is present at low levels in the G1 phase and no foci are observed. All DNA damaging agents induced an increase in the number and intensity of TopBP1 foci and extensive colocalisation with Brca1, suggesting that these proteins act together in the DNA damage response.

5.6 TopBP1 interacts with Miz-1 (II)

POZ domains in transcription factors mediate the binding of cofactors and enable targeting of these effector proteins to DNA (Bardwell & Treisman 1994). The Miz-1 N-terminus has a POZ domain and overexpression of a Miz-1 lacking this domain (Miz-1 ΔPOZ) in Rat1 fibroblasts allowed cells to grow, while overexpression of full length Miz-1 completely inhibited cell growth. Miz-1 inhibits cell proliferation by activating transcription of the cell cycle inhibitor p15INK4B gene (Staller et al. 2001), but Miz-1

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ΔPOZ failed to activate p15INK4B according to a luciferase reporter assay. Full length Miz-1 activated the luciferase reporter and the expression levels of truncated and full length Miz-1 were the same. Therefore, the POZ domain is required for the transcriptional activity of Miz-1.

To identify effector proteins binding to the Miz-1 POZ domain, a two-hybrid screen was performed. One of the recovered clones encoded amino acids 861-1407 of TopBP1. TopBP1 did not interact with several control proteins including POZ domains of Bcl-6 and ZID-1 indicating the interaction between TopBP1 and Miz-1 is specific. TopBP1 BRCT domains 7-8 mediated the interaction with Miz-1 as shown by deletion analysis.

GST tagged TopBP1 bound to in vitro translated Miz-1 in a GST-pull-down assay while GST alone did not, indicating a direct interaction between TopBP1 and Miz-1. The binding of GST-TopBP1 to Miz-1 ΔPOZ was reduced four-fold indicating that the POZ domain is required for efficient interaction in vitro.

When both Miz-1 and TopBP1 were overexpressed in HeLa cells coimmunoprecipitation was detected indicating an interaction in vivo. Endogenous Miz-1 was also coimmunoprecipitated with endogenous TopBP1 and vice versa from HeLa and human keratinocyte cell extracts.

5.7 TopBP1 inhibits activation of p15INK4B promoter by Miz-1 (II)

Human keratinocytes were transiently transfected with a p15INK4B promoter-luciferase reporter and CMV-based expression plasmids to investigate the functional consequences of TopBP1-Miz-1 complex formation. TopBP1 expression alone had no effect on reporter activity, but TopBP1 was able to inhibit the transactivation activity of Miz-1. TopBP1 expression had no effect on reporter activation by transcription factors Smad3, Smad4 or TGFβ, indicating that TopBP1 is not a general transcription inhibitor. Expression of TopBP1 lacking BRCT domains 7-8 did not significantly inhibit transactivation by Miz-1 suggesting that TopBP1 interaction with Miz-1 masks the POZ domain and specifically inhibits transactivation by Miz-1.

5.8 After UV damage TopBP1 mRNA levels are downregulated and interaction of TopBP1 with Miz-1 decreases followed

by upregulation of p15INK4B (II)

In human keratinocytes TopBP1 mRNA was downregulated in response to UV irradiation as reported earlier (Pötter et al. 2000). The remaining TopBP1 protein relocalised from a quite diffuse nuclear localisation to discrete foci while the distribution of Miz-1 was not affected, suggesting that these proteins dissociate after UV irradiation. Indeed, densitometric quantification of Western blots showed that the amount of TopBP1 in Miz-1 immunoprecipitates decreased nine-fold after UV irradiation. .

The transcription level of the Miz-1 target gene, p15INK4B, was increased following the kinetics of TopBP1 downregulation which supports the idea that Miz-1 is released from

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TopBP1 in response to UV irradiation. To demonstrate the involvement of Miz-1 in the repression of p15INK4B, keratinocytes were transfected with a CMV-based expression plasmid containing either a wild type p15INK4B promoter-luciferase reporter or a mutated p15INK4B promoter lacking the Miz-1 binding site. The basal activity and transactivation by Smad3 and Smad4 of the reporter was the same in both plasmids but the mutant plasmid showed no reporter upregulation following UV irradiation. These results suggest that Miz-1 mediates the upregulation of the cell cycle inhibitor p15INK4B in human keratinocytes.

5.9 The ATM/ATR kinase inhibitor caffeine blocks downregulation and redistribution of TopBP1 (II)

ATM and ATR kinases are general regulators of the cellular UV response. Inhibition of these kinases with caffeine blocked downregulation of TopBP1 mRNA and relocalisation of the TopBP1 protein. Upregulation of the p15INK4B promoter was also prevented. These results suggest that TopBP1 is regulated via phosphorylation by ATM or ATR kinases.

5.10 TopBP1 binds to the p21Cip1 promoter together with Miz-1 (II)

Gel shift experiments showed that three complexes bound to the p21Cip1 promoter start site. Antibody masking of the DNA binding domain of Miz-1 removed all three complexes from DNA indicating that Miz-1 directly binds to the p21Cip1 promoter. Similar results were obtained earlier for the p15INK4B promoter (Staller et al. 2001). Control antibodies had no effect on Miz-1 binding to the promoter sequence while α-TopBP1.2 supershifted two of the complexes indicating a presence of TopBP1 in a fraction of the Miz-1 DNA binding complexes. After a UV pulse the supershift was diminished indicating the release of TopBP1 from the complex. Small interfering RNA against Miz-1 suppressed the p21Cip1 transcription showing directly the involvement of Miz-1 in the upregulation of p21Cip1 after UV damage. These results indicate that Miz-1 is able to bind the p21Cip1 promoter together with TopBP1, but Miz-1 is transcriptionally active only after TopBP1 removal accompanied by exposure of the POZ domain.

5.11 DNA polymerase ε interacts with the hyperphosphorylated RNA pol II and recently synthesised RNA (III)

When RNA pol II was immunoprecipitated from HeLa cell extracts, pol ε but not pols δ or α coimmunoprecipitated. Pol ε immunoprecipitations done with two polyclonal antibodies recognising different areas of pol ε both contained RNA pol II, whereas pol δ or pol α immunoprecipitations did not. Pol α, δ or ε immunoprecipitations also did not

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contain other replicative DNA polymerases. IPs were repeated in the presence of ethidium bromide to exclude DNA mediated interactions (Lai & Herr 1992).

Pol ε has been reported to be present in RNA pol II holoenzyme preparations purified by conventional chromatography (Maldonado et al. 1996). Therefore, we investigated whether RNA pol II was present in standard purification intermediates of pol ε (Syväoja & Linn 1989). RNA pol II copurified with combined pol ε peak fractions through diethylaminoethyl-sephacel, phosphocellulose, and hydroxyapatite chromatographies (Figure 3A). Furthermore, hyperphosphorylated RNA pol II purification profile from the hydroxyapatite column matched the pol ε profile suggesting that pol ε interacts specifically with the phosphorylated, transcriptionally active, form of RNA pol II (Figure 3B). The hypophosphorylated form of RNA pol II was present in all samples with no increase corresponding to the pol ε peak fractions.

Fig. 3. RNA Pol II and pol ε partially copurify A. Pol ε was purified as described in Syväoja & Linn 1989. Western analysis was performed with pol ε (G1A, H3B and E24A) and RNA pol II (N20) antibodies for whole cell extract (WCE) and for the combined peak fractions from the diethyl aminoethyl sephacel column (DEAE), for the combined peak fractions from the phosphocellulose column (PC), and for the peak fraction from the hydroxyapatite column (HA). B Elution profiles for pol ε and hyperphosphorylated form of RNA pol II from the hydroxyapatite column. DNA polymerase activity was measured as described in Syväoja & Linn 1989. Fractions containing DNA polymerase activity were analysed by Western blot with hyperhosphorylated RNA pol II antibody (H5) and with pol ε antibodies (G1A, H3B and E24A).

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The RNA pol II present in pol ε immunoprecipitates was the phosphorylated form. When immunoprecipitated RNA pol II was treated with calf intestinal phosphatase, the slowly migrating form was lost. During the transcription cycle the hypophosphorylated form of RNA pol II binds to promoters and becomes phosphorylated mainly from the serine 5 of the CTD during transcription initiation followed by serine 2 phosphorylation during the elongation phase. The N20 antibody recognising all forms of RNA pol II and phosphospecific antibodies H14 recognising the serine 2 phosphorylated form, 8W16G recognising the serine 2 non-phosphorylated form but insensitive to serine 5 phosphorylation, and H5 recognising the serine 5 phosphorylated form of RNA pol II all coimmunoprecipitated pol ε and vice versa. Therefore, serine 5 phosphorylation alone is sufficient to mediate the interaction while serine 2 phosphorylation alone may also be sufficient.

To further investigate whether RNA pol II phosphorylation is required for the interaction with pol ε, chemicals known to inhibit transcription were employed. CDK inhibitors DRB and roscovitine inhibit the phosphorylation of CTD thereby preventing transition from the initiation to the elongation complex, while α-amanitin inhibits both initiation and elongation by blocking the translocation of DNA and RNA through the catalytic centre of RNA pol II (Bushnell et al. 2002, Gong et al. 2004, reviewed by Kim et al. 2001 and by Schang 2004). RNA pol II already proceeding through the elongation phase was not affected. Both DRB and roscovitine strongly decreased the amount of hyperphosphorylated RNA pol II present in cell extracts or immunoprecipitates resulting in a strongly diminished interaction. In contrast, α-amanitin induced stalled replication did not affect RNA pol II phosphorylation and the interaction with pol ε remained.

Pols ε and α were crosslinked to recently synthesised RNA detected by the presence of labelled UTP in immunoprecipitates, while pol δ immunoprecipitates did not contain the label. The presence of RNA in the pol α precipitate was expected because pol α is responsible for RNA primer synthesis during replication. Pol α crosslinking to RNA was abolished when DNA replication was inhibited by aphidicolin, or the crosslinker BrdUTP was omitted. Pol ε crosslinking to RNA was not dependent on BrdUTP or affected by DNA replication inhibition, but inhibition of transcription with α-amanitin abolished interaction. Therefore the crosslinking of RNA to pol α is linked to DNA replication while crosslinking to pol ε is related to transcription.

5.12 DNA polymerase ε and hyperphosphorylated RNA polymerase II colocalise (III)

Hyperphosphorylated RNA pol II stained with the H5 antibody showed quite even nuclear localisation in human glioblastoma T98G cells with few foci, which became more visible after hypotonic extraction of soluble proteins and partial digestion of DNA. Most RNA pol II foci colocalised with the α-pol ε G1A signal while only a few pol ε foci colocalised with RNA pol II due to the higher number of pol ε foci.

To investigate the colocalisation more closely, T98G cells were synchronised and samples for immuno-electron microscopy were prepared from G0 to lateS/G2 populations. Nuclei were imaged, and three or more gold particles in close proximity of each other

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were classified as a centr. A centre containing at least one label for both pol ε and RNA pol II was classified as a mixed centre and a site of colocalisation. The specificity of pol ε staining was tested by performing double labelling with antibodies G1A and H3B recognising different areas of pol ε. Fiftytwo percent of the pol ε signal and 45 % of RNA pol II labelled with N20 antibody recognising the total amount of RNA pol II were present in centres. This means that both proteins appeared rather extensively in centres in immuno-electron microscopy stainings.

Thirtyseven percent of the RNA pol II signal colocalised with pol ε and colocalisation was even stronger (52 %) when RNA pol II labelling was performed with the elongating RNA pol II specific H5 antibody. H5 staining was slightly different from N20 staining because 66 % of all signal was located in centres. It seems that transcriptionally active H5 form of RNA pol II forms most of RNA pol II foci and colocalises with pol ε more often than the transcriptionally inactive form. Unfortunately, there is no antibody recognising the transcriptionally inactive form of RNA pol II. From the remaining centres 23 % contained only pol ε and 25 % only RNA pol II. Therefore, the colocalisation between RNA pol II and pol ε is quite extensive, but not complete.

5.13 DNA polymerase ε interaction with RNA pol II may not be linked to nucleotide excision repair or DNA replication (III)

Pol ε and TFIIH are both implicated in nucleotide excision repair and belong to the RNA pol II holoenzyme (Araujo et al. 2000). Also RNA pol II itself acts in the transcription coupled repair branch of NER as a damage sensor (Svejstrup 2001). To study the involvement of RNA pol II and pol ε in nucleotide excision repair T98G nuclei were partly irradiated through a filter containing 8 μm holes (Moné et al. 2001). The H3 antibody recognising cyclobutane thymidine dimers was used to visualise damaged areas in nuclei. Nucleotide excision repair factor XPB localised to damaged areas as shown earlier (Volker et al. 2001). Punctate pol ε staining remained after UV irradiation and there was a small but reproducible relocalisation into damaged areas of DNA. RNA pol II staining was not affected by UV irradiation in most cells while a few cells showed significant reduction of RNA pol II signal at the damaged areas. These results do not support the view that pol ε interaction with RNA pol II is needed for nucleotide excision repair.

For further studies HeLa cells were UV irradiated for 10 J/m2 but nucleotide excision repair proteins XPA, XPB or XPG were not present in RNA pol II or pol ε immunoprecipitates. Neither was the level of coimmunoprecipitation between RNA pol II and pol ε affected.

To study the effect of cell cycle phase on RNA pol II interaction with pol ε, T98G cells were synchronised using serum starvation and samples at G0, G1, G1/S, mid S and G2/M phases of the cell cycle were collected. Immunoprecipitation was equally efficient in all samples and immuno-electron microscopy revealed pol ε colocalising with RNA pol II in all cell cycle phases. Furthermore, the replication inhibitor aphidicolin was not able to abolish the interaction. These data do not support the view that the interaction of RNA pol II with pol ε is linked to DNA replication, altough not rule out this possibility.

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5.14 Cdc45, pol δ, pol ε, and PCNA behave differently in cell fractionation (IV)

Soluble, DNA bound, nuclear matrix bound, and whole cell T98G cell extracts were prepared and tested by Western blotting using control antibodies. β-tubulin was only in the soluble fraction indicating that other fractions were free of soluble proteins (Rytkönen et al. 2006). MCM2 was partially released by DNase I treatment as expected, locating in both DNA and matrix bound fractions (Todorov et al. 1995). Lamin was present exclusively in the matrix fraction indicating that other fractions were not contaminated by nuclear skeleton.

To follow the DNA binding dynamics of replication proteins, T98G cells were synchronised by serum starvation and after the release of the block, samples at different time points were collected. Synchronisation of cells was verified by flow cytometry. PCNA and pol δ were mainly soluble in the beginning of the cell cycle but a fraction of each protein became DNA bound after entry into S phase. The DNA bound fraction became larger towards the end of S phase. DNA bound Cdc45 levels also increased towards the end of S phase although a fraction of Cdc45 was chromatin bound in all cell cycle stages. As expected from its role in replication initiation, Cdc45 was present in the DNA bound fraction before PCNA and pol δ. Pol ε behaved differently compared to other replication proteins. Pol ε was mainly matrix bound in all cell cycle stages. The small DNA bound fraction of pol ε existed in normal and slowly migrating forms, the slowly migrating form being specific for the DNA bound fraction. DNA bound pol ε levels also increased towards the end of S phase.

5.15 Cdc45 colocalises with DNA polymerases δ and ε (IV)

Asynchronous T98G cells were pulse labelled with BrdU to reveal sites of DNA synthesis in indirect immunofluorescence. Pols δ and ε localised almost exclusively into nuclear foci in all interphase cells but these foci did not colocalise with DNA synthesis sites during S phase. Cdc45 formed diffuse nuclear and nucleolar staining.

To characterise the localisation of Cdc45, pol δ, and pol ε more precisely immuno-electron microscopy was utilised. Most of the Cdc45 signal was dispersed in nuclei in all interphase cells although the amount of Cdc45 signal was two times higher in S phase compared to G0 and G1 phases. Roughly 30 % of Cdc45 particles localised into centres but there was no difference between G0/G1 and S phase samples, suggesting that these centres are not dependent on DNA synthesis.

As in our previous immuno-electron study, no colocalisation between pols δ and ε was detected (Rytkönen et al. 2006). Pol ε localised mainly into centres while pol δ formed extended fibrous like structures and diffuse staining. In contrast, Cdc45-pol δ and Cdc45-pol ε colocalisations were frequently detected. 68 images from nuclei in G1, early S, and late S phases of the cell cycle were analysed. Most of the pol ε signal was in centres (53 %) while the rest was dispersed. The amount of pol ε particles colocalising with Cdc45 was 27 %, and 35 % of the Cdc45 particles colocalised with pol ε. There were slightly

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less than 20 % of centres containing only Cdc45 and slightly more than 20 % of centres containing only pol ε, while 59 % of the centres formed contained both Cdc45 and pol ε in all samples. The results are shown in table 2.

Table 2. Results from immuno-electron microscopy double stainings.

Staining Cdc451 Pol δ1 Cdc452 Pol ε2

Amount of particles 1803 468 2920 2186 Particles in centres3 521 (29%) 204 (44%) 905 (31%) 1157 (53%) Particles colocalising 409 (23%) 252 (54%) 778 (27%) 769 (35%) Number of particles per centre 3.5 2.0 2.5 3.0 Nuclei / images counted 31 / 61 82 / 182 1Particles representing Cdc45 and pol δ were calculated from the same images 2Particles representing Cdc45 and pol ε were calculated from the same images 3Three or more colocalising gold particles were counted as a centre

Amount of pol δ labelling in pol δ – Cdc45 centres was rather low; on average there was only 2.0 pol δ particles per center compared to 3.5 Cdc45 particles. Almost all pol δ centres also contained Cdc45, only 1 % of all centres contained exclusively pol δ. These data underline the observation that pol δ localises into fibrous like structures rather than centres. On average 64 % were mixed Cdc45 – pol δ centres and the rest were pure Cdc45 centres. Fifty four % of all pol δ particles colocalised with Cdc45, and 23 % of all Cdc45 particles colocalised with pol δ. Although the amount of centres increased towards the end of S phase, the relative amount of signal in centres and the level of colocalisation between pol δ – Cdc45 and pol ε – Cdc45 were quite constant in all samples analysed. Taken together, both pols δ and ε colocalised rather extensively with Cdc45 while there was no apparent colocalisation between pols δ and ε. Pol ε was frequently located into centres and pol δ into fibrous like structures.

Interactions between Cdc45, pol δ and ε were also investigated using immunoprecipitation. IPs were done from both the G1/S border and mid/lateS phase synchronised HeLa cells and crosslinked with paraformaldehyde prior to extract preparation. Under these crosslinking conditions Cdc45 coimmunoprecipitated both with pol ε and pol δ but the polymerases did not coimmunoprecipitate each other. Under native conditions, the proteins did not coimmunoprecipitate. In conclusion, there was no interaction between the pols δ and ε, but there were weak interactions between pol δ – Cdc45 and pol ε – Cdc45, detectable under crosslinking conditions.

6 Discussion

6.1 TopBP1 interacts with DNA polymerase ε and is needed for replication (I)

S. cerevisiae Dpb11 and S. pombe Cut5 interact with pol ε and are required for DNA replication and for the S phase checkpoint (Araki et al. 1995, Masumoto et al. 2000, Saka et al. 1997). We searched for human pol ε interacting proteins based on sequence homology with these yeast proteins and cloned TopBP1. TopBP1 was named earlier as topoisomerase IIβ binding protein (Yamane et al. 1997). TopBP1 has three pairs of BRCT domains and two single ones. BRCT pairs 1-2 and 4-5 are conserved from yeast to human. The other four BRCT domains of ToBP1 are not present in S. cerevisiae or in S. pombe. It seems that TopBP1 and TopBP1 homologs in higher eukaryotes have acquired additional functions mediated by these additional sequences (Kumagai et al. 2006). The third BRCT pair, domains 7 – 8, is homologous to the Brca1 C-terminus, which is important for the tumour suppressor activity of Brca1 (Friedman et al. 1994).

We were not able to complement temperature sensitive alleles of Dpb11 and Cut5 with TopBP1, but there is other evidence that TopBP1 is the functional homolog of Dpb11 and Cut5. TopBP1 has a role in the DNA damage response similar to Dpb11 and Cut5 (discussed below). The TopBP1 expression level is highest in S phase, which is common for proteins involved in DNA replication. In addition, Pol ε coimmunoprecipitates with TopBP1 and the interaction is not affected by DNA damage, which indicates that, like Dpb11, TopBP1 plays a role in replication. TopBP1 interactions are presented in figure 4.

The Pol ε present in the TopBP1 immunoprecipitate was in the hyperphosphorylated form (Tuusa 2001). This form may be linked to replication elongation because it was solely DNA bound and the amount increased towards the end of S phase. The faster migrating major form peaked at the beginning of S phase, suggesting this form acts in the initiation complex. Therefore, TopBP1 interaction with pol ε may be replication elongation related. This does not rule out a transient TopBP1 interaction with pol ε during the DNA replication initiation process. In yeast, Dpb11 is required for loading of pols α and ε to the origins of replication (Araki et al. 1995, Masumoto et al. 2000). Like Dpb11, TopBP1 is linked to initiation because an antibody recognising the BRCT domain 6 of

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TopBP1 inhibits replication to the same extent (44 %) as neutralizing antibodies against pols α and ε (Kaczmarek et al. 1986, Pospiech et al. 1999). Addition of the TopBP1.2 peptide containing the sixth BRCT domain to the replication reaction caused the highest level of inhibition we have measured in this system, 83 %. The high level of inhibition suggests that the initiation phase of DNA replication is affected. Apparently the peptide competes with endogenous TopBP1 in binding to a critical DNA replication component. Interestingly, we were able to precipitate p53 using GST-BRCT6 as bait (unpublished).

In Xenopus XTopBP1 participates in both replication initiation and elongation. XTopBP1 is required for Cdc45 loading onto replication origins (Van Hatten et al. 2002, Hashimoto & Takisawa 2003) and also participates in the elongation phase, although it is not an essential component there (Kim et al. 2005). Without DNA damage TopBP1 was bound to DNA only during S phase indicating involvement in DNA replication, but degradation of TopBP1 using the small interfering RNA technique allowed replication to continue. Removal of TopBP1 caused activation of ATM and Chk2 kinases, phosphorylation of histone H2AX, and aberrant cell division suggesting accumulation of double-strand breaks. Our results suggest a similar role for human TopBP1 both in replication initiation and elongation.

6.2 Cdc45 forms separate complexes with DNA polymerases δ and ε (IV)

Cdc45 is required for DNA replication. Cdc45 is essential for the loading of pols α and ε onto replication initiation sites and it is a component of the elongation apparatus (Mimura et al. 2000, Pacek et al. 2006). Cdc45 interactions with pols α and ε has been reported earlier (Aparacio et al. 1999, Uchiyama et al. 2001), whereas interaction with pol δ is new. Cdc45 coimmunoprecipitates with both pol δ and ε showing that these polymerases interact physically with Cdc45. Interaction was detected only after the addition of crosslinker, suggesting that the interaction is weak or even indirect. Cdc45 interactions with both pol δ and ε were equally strong in the early and mid/late S phase cells indicating that the interaction is present throughout S phase. Pols δ and ε did not coimmunoprecipitate each other even in the presence of a crosslinker suggesting that these polymerases do not exist in close proximity to each other and rather form completely separate complexes.

Immuno-electron microscopy results are supporting this view. Cdc45 colocalises with both polymerases but pols δ and ε do not colocalise with each other, as has been shown in our previous study (Rytkönen et al. 2006). The different structures formed by pol ε compared to pol δ indicate that the two pols are not localised to the same fine structures. Pol ε forms mainly centres and pol δ extended fibrous like structures. Obviously Cdc45 does not mediate interaction between these proteins but rather forms a separate complex with each polymerase.

Cell fractionation studies revealed that most of Cdc45, PCNA, and pol δ were in the soluble fraction while the DNA bound fraction accumulated towards the end of S phase. DNA bound Cdc45 was already present in G0/G1 phases before the DNA binding of PCNA and pol δ. This may show the temporal order of function of these proteins.

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Contrary to other replication proteins investigated, most of pol ε was bound to nuclear matrix. The minor DNA bound fraction of pol ε increased toward the end of S phase. Interestingly, DNA bound pol ε existed in two forms; the larger one was present only in the DNA bound fraction. Possibly one form is responsible for replication initiation and the other for the elongation process, and modification of DNA bound pol ε may be important for its regulation. There seems to be species to species variations in the organisation of DNA replication. In Xenopus pol ε is soluble and loading to chromatin is Cdc45 dependent (Mimura et al. 2000). The immobile, matrix bound human pol ε may coordinate DNA replication by recruiting DNA and soluble replication apparatus proteins into matrix bound replication factories.

Taken together, human Cdc45 has a role in the function of pols δ and ε during S phase. Furthermore, Cdc45 exists in separate complexes. The first one contains pol δ and the other pol ε. A plausible model is that pols δ and ε both participate in DNA replication but do not interact with each other. This is in contrast to bacteria in which the leading and lagging strand DNA polymerases form a physical dimer.

6.3 DNA polymerase ε associates with transcribing RNA polymerase II and nascent transcripts (III)

Pol ε coimmunoprecipitates the phosphorylated, actively transcribing form of RNA pol II. The phosphorylated form was detected by a mobility shift in electrophoresis which was lost after phosphatase treatment, and with antibodies recognising the phosphorylated form of RNA pol II. The RNA pol II precipitate contained pol ε, but not other replicative polymerases, showing that the immunoprecipitation is specific for pol ε. Maldonado et al. (1996) detected pol ε in purified RNA pol II preparation along with several other DNA replication and recombination factors. This holoenzyme contained mainly the hypophosphorylated form of RNA pol II and, after the addition of general transcription factors, the complex was able to start transcription in vitro. This clearly differs from our results because the interaction depended on the phosphorylation of the CTD. When the elution profile of pol ε from the last purification column was compared to the elution profile of hyperphosphorylated RNA pol II from the same column they were almost identical, while the hypophosphorylated RNA pol II was present in equal amounts in all fractions analysed. Secondly, the addition of chemicals reducing the phosphorylation of CTD and preventing the transition from the initiation to elongation phase of transcription inhibited RNA pol II interaction with pol ε significantly. Thirdly, TFIIH factors present on the promoter bound hypophosphorylated form of RNA pol II were not present in our immunoprecipitates. The interaction is not dependent on ongoing RNA synthesis though, because the interaction remained in the presence of aphidicolin arrested transcription. Although transcription is not needed for the interaction, pol ε is in close proximity to nascent transcripts because pol ε can be crosslinked to recently transcribed RNA.

Pol ε interaction with RNA pol II was also detected by colocalisation in indirect immunofluorescence. RNA pol II foci were more prominent after removal of soluble proteins and partial DNA digestion, and only after this treatment RNA pol II signal partially colocalised with pol ε. In immuno-electron microscopy samples, colocalisation

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between pol ε and RNA pol II was readily detected, and the RNA pol II antibody recognising the phosphorylated form of RNA pol II showed a higher amount of colocalisation with pol ε compared to antibody recognising all forms of RNA pol II. The localisation of RNA pol II in this study resembled the localisation reported previously (Iborra et al. 1996, Cmarko et al. 1999).

RNA interacts via CTD with several proteins regulating the transcription cycle. It is tempting to speculate that the interaction is sensitive to the phosphorylation status of CTD because pol ε binds to that domain. Nevertheless, it was not possible to precipitate pol ε with a hypo- or hyperphosphorylated recombinant CTD construct suggesting that the interaction requires other areas of RNA pol II or may be indirect.

Several possibilities for the functional significance of this interaction were considered. Both pol ε and RNA pol II are implicated in nucleotide excision repair, pol ε as a polymerase and RNA pol II as a damage sensor (Araujo et al. 2000, Donahue et al. 1994). An interaction between RNA pol II and pol ε would facilitate repair, but we were neither able to detect alteration in coimmunoprecipitation efficiency after UV irradiation nor a cooperative response to UV damage after irradiation of limited areas of T98G cell nuclei. Furthermore, we were not able to detect repair proteins XPA, XPB or XPG in our RNA pol II or pol ε immunoprecipitates. These findings do not support, although they do not rule out, the hypothesis that the interaction of RNA pol II with pol ε is linked to DNA repair.

The interaction may mediate the coordination of DNA replication and transcription, as has been reported in bacteria (Liu et al. 1993, Liu & Alberts 1995). However, the interaction was neither limited to S phase nor affected by aphidicolin induced replication arrest, suggesting that the interaction is not dependent on DNA replication. Furthermore, the extensive colocalisation detected by immuno-electron microscopy suggests a more general role for the interaction than coordination of colliding transcription apparatuses and replication forks.

One possible explanation for the interaction is the coordination of DNA replication initiation. Many promoter areas of actively transcribed genes act as replication initiation sites (Gauthier et al. 2002, Kohzaki & Murakami 2005). Sites of DNA replication were also found to contain RNA processing components (Philimonenko et al. 2006). However, the pre-initiation complex rather than RNA pol II recruit pol ε to these sites. Interestingly, RNA pol II also interacts with MCM proteins and antibodies against Mcm2 inhibited transcription (Yankulov et al. 1999). Furthermore, mutations in the yeast CTD interacted genetically with Mcm5 and affected the maintenance of plasmids and DNA replication (Gauthier et al. 2002).

6.4 TopBP1 functions in the DNA damage response (I, II)

TopBP1 localisation during the S and G2 phases of the cell cycle was similar to Brca1 (Scully et al. 1997b). We were not able to coimmunoprecipitate these proteins though, which suggests that TopBP1 and Brca1 do not interact directly. However, the extensive colocalisation during S and G2 suggests that Brca1 and TopBP1 are present in the same macroscale complexes. After a hydroxyurea block both proteins relocalised to numerous

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stalled replication forks visualised by PCNA staining. This links TopBP1 to recognition of DNA damage, the DNA damage induced signalling cascade, the transcriptional response to DNA damage, processing of stalled replication forks, reinitiation of stalled forks, or a combination of these processes. DNA damage signalling involves phosphorylation of mediator and effector proteins. TopBP1 is a phosphoprotein but its phosphorylation is not altered after DNA damage, rather supporting a function in damage recognition than a role in DNA damage signalling. In agreement with this TopBP1 is able to bind to DNA breaks favouring the function in damage recognition (Yamane & Tsuruo 1999). Furthermore, we have shown that TopBP1 interacts with the DNA damage recognition and repair protein hRad9, but interaction with the damage responsive kinases Chk1 or Chk2 was not detected.

Recent findings show TopBP1 is a positive regulator of ATR activity (Kumagai et al. 2006). Ataxia-telengiectasia mutated (ATM) and ataxia-telengiectasia and Rad3-related (ATR) kinases are the main DNA damage regulators in eukaryotic cells (reviewed by Traven & Heierhorst 2005 and Sancar et al. 2004). They share the work, ATM responding mainly to DNA double strand breaks and ATR to stalled replication forks and a wider spectrum of DNA damage. ATR exists as a complex with the ATR-interacting protein (ATRIP) (Falck et al. 2005, reviewed by Sancar et al. 2004). The region between TopBP1 BRCT domains six and seven interacts with ATR in an ATRIP dependent manner and stimulates ATR kinase activity (Kumagai et al. 2006). This region was both sufficient and necessary for ATR interaction because this peptide alone stimulated ATR activity and mutation in this area abolished the ability of TopBP1 to stimulate ATR activity. TopBP1 facilitated phosphorylation of several ATR substrates like Chk1, Mcm2, Rad17, Rad1, and Hus1 forming an important step in the initiation of ATR-dependent signalling (Yan et al. 2006, Kumagai et al. 2006, Lupardus & Cimprich 2006). TopBP1 is the only known ATR activator. TopBP1 interactions are summarised in figure 4.

Interestingly, the Rad9-Hus1-Rad1 clamp and clamp loader Rad17 are required for ATR-dependent phosphorylation of downstream Chk1 (reviewed by Sancar et al. 2004). Therefore, this damaged DNA binding complex must also act early in the signalling cascade and may explain our finding that hRad9 and TopBP1 interact. In S. pombe Rad9 was phosphorylated in response to DNA damage and phosphorylated Rad9 interacted with the TopBP1 homolog Cut5 (Furuya et al. 2004). The Rad9-TopBP1 interaction also was required for activation of the Chk1 damage checkpoint.

The DNA damage checkpoint function of Cut5 is mediated through BRCT domain three, which interacts with the DNA damage checkpoint protein Crb3 (Taricani & Wang 2006). By mutating this domain the DNA damage checkpoint was destroyed while retaining normal DNA replication, replication checkpoint and cell cycle progression. This means that Cut5 mediates several DNA replication and repair related functions in the cell and these functions can be mapped into separable domains in the protein. Because TopBP1 contains eight BRCT domains it likely has additional interaction partners compared to Cut5 and forms a more complex regulation network. One such interaction partner could be p53, which binds to the sixth BRCT domain of TopBP1 (unpublished).

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Fig. 4. TopBP1 interactions. Diamonds represent proteins involved mainly in DNA repair, rectangles indicate involvement in DNA replication and circles in trancription.

ATM phosphorylates Brca1 after double-strand break damage and ATR after UV irradiation or treatment with DNA replication inhibitors linking Brca1 to both major DNA damage signalling pathways (Cortez et al. 1999, Tibbetts et al. 2000). Brca1 is localised to sites specialized for the processing of replicating or replicated DNA (Scully et al. 1997b). TopBP1 was shown to coimmunoprecipitate with the overexpressed Brca1/Bard1 complex after DNA damage (Greenberg et al. 2006). The result is not in conflict with our immunoprecipitation experiments, which were performed without DNA damaging agents, and support the view of extensive cooperation of these proteins after DNA damage or stalled DNA replication. TopBP1 also has a Brca1 independent function because it forms nuclear foci in response to double-strand breaks in the G1 phase when Brca1 levels are low.

TopBP1 was linked to transcription regulation after UV damage. TopBP1 interacted with the transcription factor Miz-1 shown by the two-hybrid method, coimmunoprecipitation of native proteins, and GST-pull-down with in vitro transcribed proteins. In response to UV irradiation of human keratinocytes, TopBP1 mRNA and protein levels were downregulated and Miz-1 was released from the inhibitory interaction with TopBP1. After the release Miz-1 was able to induce p15Ink4b expression. Both free and TopBP1 bound Miz-1 were able to bind to the p21Cip1 promoter but only free Miz-1 cooperatively with p53 was able to increase p21 expression to arrest the cell cycle. A similar two-input regulation model has been reported for the p15Ink4b promoter in response to TGF-β signalling (Seoane et al. 2001). TGF-β downregulates c-myc

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expression and releases Miz-1 from this inhibitory interaction (Staller et al. 2001). Miz-1 still recruits the transcriptional activator p300 to the p15 promoter and, together with TGF-β activated Smad transcription factors, cooperatively activates the promoter (Feng et al. 2002, Seoane et al. 2001).

TopBP1 also interacts with the proto-oncogene c-Abl and regulates its expression level and acts as a coactivator for the papillomavirus E2 protein in transcription regulation (Zeng et al. 2005, Boner et al. 2002). Furthermore, TopBP1 binds to and regulates transcription factor E2F1 in DNA damage dependent way (Liu et al. 2003). It seems that TopBP1 modulates expression of several cell cycle, replication, and repair related genes.

7 Conclusions

In this study, we cloned TopBP1, which is a human homolog of the DNA replication and S phase checkpoint proteins Dpb11 and Cut5 from S. cerevisiae and S pombe, respectively. TopBP1 localised into stalled replication forks together with Brca1 and these proteins may cooperate in the DNA damage response. TopBP1 also has a Brca1-independent DNA damage response function because, after double-strand break damage in the G1 phase, TopBP1 formed foci not containing Brca1. Because the TopBP1 phosphorylation status did not change after the DNA damage, our results suggest a role for TopBP1 early in the damage response rather than in signal transduction or in damage processing. Actually, TopBP1 was later reported to positively regulate ATR forming a very early step in damage signalling. This is in agreement with our finding that TopBP1 interacts with the DNA damage recognition and processing protein hRad9.

After UV irradiation TopBP1 levels were downregulated and this released the transcription factor Miz-1 from an inhibitory interaction with TopBP1. Free Miz-1 was able to upregulate the transcription of cell cycle progression inhibitors p15 and p21, linking TopBP1 into cell cycle arrest induction after UV damage.

TopBP1 was required for chromosomal DNA replication because DNA replication in isolated cell nuclei was severely inhibited both by a peptide containing the sixth BRCT domain of TopBP1 and by an antibody binding to this domain. The high level of inhibition suggests a role for TopBP1 in the initiation of DNA replication. Secondly, TopBP1 associated with the form of DNA polymerase ε possibly acting in the replication elongation phase.

Pol ε forms a complex with Cdc45 during DNA replication and this complex acts independently of the Cdc45 complex with pol δ. Based on cell fractionations, coimmunoprecipitations, and cell stainings we argue that pols δ and ε act independently of each other in S phase. Our results are in agreement with the hypothesis that pol ε is the leading strand relicase and pol δ acts on the lagging strand.

Interestingly, pol ε interacts with the transcriptionally active form of RNA pol II and nascent transcripts suggesting an involvement in transcription. The actual meaning of this interaction is unclear. It may be needed to target pol ε to replication initiation sites. Both pol ε and RNA pol II are involved in UV damage repair, which is an alternative explanation for the interaction.

References

Abdurashidova G, Deganuto M, Klima R, Riva S, Biamonti G, Giacca M, Falaschi A (2000) Start sites of bi-directional DNA synthesis at the human lamin B2 origin. Science 287: 2023-2026.

Aboussekhra A & Thoma F (1999) TATA-binding protein promotes the selective formation of UV-induced (6-4)-photoproducts and modulates DNA repair in the TATA box. EMBO J 18: 433-443.

Anachkova B, Djeliova V & Russev G (2005) Nuclear matrix support of DNA replication. J Cell Biochem 96: 951-961.

Aparacio OM, Weinstein DM & Bell SP (1997) Components and dynamics of DNA replication complexes in S. cerevisiae: redistribution of MCM proteins and Cdc45p during S phase. Cell 91: 59-69.

Aparacio OM, Stout AM & Bell SP (1999) Differential assembly of Cdc45p and DNA polymerases at early and late origins of DNA replication. Proc Natl Acad Sci USA 96: 9130-9135.

Araki H, Hamatake RK, Johnston LH & Sugino A (1991a) DPB2, the gene encoding DNA polymerase II subunit B, is required for chromosome replication in Saccharomyces cerevisiae. Proc Natl Acad Sci USA 88: 4601-4605.

Araki H, Hamatake RK, Morrison A, Johnson AL, Johnston LH & Sugino A (1991b) Cloning DPB3, the gene encoding the third subunit of DNA polymerase II of Saccharomyces cerevisiae. Nucleic Acids Res 19: 4867-4872.

Araki H, Leem S-H, Phongdara A & Sugino A (1995) Dpb11, which interacts with DNA polymerase II(ε) in Saccharomyces cerevisiae, has a dual role in S-phase progression and at a cell cycle checkpoint. Proc Natl Acad Sci USA 92: 11791-11795.

Araujo SJ, Tirode F, Coin F, Pospiech H, Syväoja JE, Stucki M, Hübscher U, Egly JM & Wood RD (2000) Nucleotide excision repair of DNA with recombinant human proteins: definition of the minimal set of factors, active forms of TFIIH, and modulation by CAK. Genes Dev 14: 349-359.

Arezi B & Kuchta RD (2000) Eukaryotic DNA primase. Trends Biochem Sci 25: 572-576. Arudchandran A, Cerritelli AS, Narimatsu S, Itaya M, Shin DY, Shimada Y & Crouch RJ (2000)

The absence of ribonuclease H1 or H2 alters the sensitivity of Saccharomyces cerevisiae to hydroxyurea, caffeine and ethyl methanesulphonate: implications for roles of RNases H in DNA replication and repair. Genes Cells 5: 789-802.

Asturias FJ, Cheung IK, Sabouri N, Chilkova O, Wepplo D & Johansson E (2006) Structure of Saccharomyces cerevisiae DNA polymerase ε by cryo-electron microscopy. Nat Struct Mol Biol 13: 35-43.

58

Asturias FJ, Jiang YW, Myers LC, Gustafsson CM & Kornberg RD (1999) Conserved structures of mediator and RNA polymerase II holoenzyme. Science 283: 985-987.

Ayyagari R, Gomes XV, Gordenin DA & Burgers PMJ (2003) Okazaki fragment maturation in yeast. I. Distribution of functions between FEN1 AND DNA2. J Biol Chem 278: 1618-1625.

Bae SH, Bae KH, Kim JA & Seo YS (2001) RPA governs endonuclease switching during processing of Okazaki fragments in eukaryotes. Nature 412: 456-461.

Balajee AS, Dianova I & Bohr VA (1999) Oxidative damage-induced PCNA complex formation is efficient in xeroderma pigmentosum group A but reduced in Cockayne syndrome group B cells. Nucleic Acids Res 27: 4476-4482.

Ballabeni A, Melixetian M, Zamponi R, Masiero L, Marinoni F & Helin K (2004) Human Geminin promotes pre-RC formation and DNA replication by stabilizing CDT1 in mitosis. EMBO J 23: 3122–3132.

Bardwell VJ & Treisman R (1994) The POZ domain: a conserved protein-protein interaction motif. Genes Dev 8: 1664-1677.

Batty DP & Wood RD (2000) Damage recognition in nucleotide excision repair of DNA. Gene 241: 193-204.

Bell SP & Dutta A (2002) DNA replication in eukaryotic cells. Annu Rev Biochem 71: 333-374. Bell SP & Stillman B (1992) ATP-dependent recognition of eukaryotic origins of DNA replication

by a multiprotein complex. Nature 357: 128-134. Bermudez VP, MacNeill SA, Tappin I & Hurwitz J (2002) The influence of the Cdc27 subunit on

the properties of the Schizosaccharomyces pombe DNA polymerase δ. J Biol Chem 277: 36853-36862.

Blanton HL, Radford SJ, McMahan S, Kearney HM, Ibrahim JG & Sekelsky J (2005) REC, Drosophila MCM8, drives formation of meiotic crossovers. PloS Genet 1: e40.

Blow JJ & Dutta A (2005) Preventing re-replication of chromosomal DNA. Nat Rev Mol Cell Biol 6: 476-486.

Boeger H, Bushnell DA, Davis R, Griesenbeck J, Lorch Y, Strattan JS, Westover KD & Kornberg RD (2005) Structural basis of eukaryotic gene transcription. FEBS Lett 579: 899-903.

Boeger H, Griesenbeck J, Strattan JS & Kornberg RD (2004) Removal of promoter nucleosomes by disassembly rather than sliding in vivo. Mol Cell 14: 667-673.

Boner W, Taylor ER, Tsirimonaki E, Yamane K, Campo MS & Morgan IM (2002) A Functional interaction between the human papillomavirus 16 transcription/replication factor E2 and the DNA damage response protein TopBP1. J Biol Chem 2002 277: 22297-22303.

Bradsher J, Auriol J, Proietti de Santis L, Iben S, Vonesch JL, Grummt I & Egly JM (2002) CSB is a component of RNA pol I transcription. Mol Cell: 819-829.

Budd ME & Campbell JL (1997) replicative helicase, Dna2 helicase, interacts with yeast FEN-1 nuclease in carrying out its essential function. Mol Cell Biol: 2136-2142.

Budd ME, Choe W & Campbell JL (2000) The nuclease activity of the yeast DNA2 protein, which is related to the RecB-like nucleases, is essential in vivo. J Biol Chem 275: 16518-16529.

Burgers PM (1991) Saccharomyces cerevisiae replication factor C. II. Formation and activity of complexes with the proliferating cell nuclear antigen and with DNA polymerases δ and ε. J Biol Chem 266: 22698-22706.

Burgers PMJ, Koonin EV, Bruford E, Blanco L, Burtis KC, Christman MF, Copeland WC, Friedberg EC, Hanaoka F, Hinkle DC, Lawrence CW, Nakanishi M, Ohmori H, Prakash L, Prakash S, Reynaud C-A, Sugino A, Todo T, Wang Z, Weill J-C & Woodgate R (2001) Eukaryotic DNA polymerases: proposal for a revised nomenclature. J Biol Chem 276: 43487-43490.

Bushnell DA, Cramer P & Kornberg RD (2002) Structural basis of transcription: α-Amanitin-RNA polymerase II cocrystal at 2.8 Å resolution. Proc Natl Acad Sci USA 99: 1218-1222.

59

Carty SM & Greenleaf AL (2002) Hyperphosphorylated C-terminal repeat domain-associating proteins in the nuclear proteome link transcription to DNA/chromatin modification and RNA processing. Mol Cell Proteomics 1: 598–610.

Cerritelli SM, Frolova EG, Feng C, Grinberg A, Love PE & Crouch RJ (2003) Failure to produce mitochondrial DNA results in embryonic lethality in Rnaseh1 null mice. Mol Cell 11: 807-815.

Chiba N, Parvin JD (2002) The BRCA1 and BARD1 association with the RNA polymerase II holoenzyme. Cancer Res 62: 4222-4228.

Chilkova O, Jonsson BH & Johansson E (2003) The quaternary structure of DNA polymerase ε from Saccharomyces cerevisiae. J Biol Chem 278: 14082-14086.

Cho EJ, Kobor MS, Kim M, Greenblatt J & Buratowski S (2001) Opposing effects of Ctk1 kinase and Fcp1 phosphatase at Ser 2 of the RNA polymerase II C-terminal domain. Genes Dev 15: 3319-3329.

Claassen GF & Hann SR (2000) A role for transcriptional repression of p21CIP1 by c-Myc in overcoming transforming growth factor beta-induced cell-cycle arrest. Proc Natl Acad Sci USA 97: 9498–9503.

Cmarko D, Verschure PJ, Martin TE, Dahmus ME, Krause S, Fu XD, van Driel R & Fakan S (1999) Ultrastructural analysis of transcription and splicing in the cell nucleus after bromo-UTP microinjection. Mol Biol Cell 10: 211-223.

Corona DF, Eberharter A, Budde A, Deuring R, Ferrari S, Varga-Weisz P, Wilm M, Tamkun J & Becker PB. (2000) Two histone fold proteins, CHRAC-14 and CHRAC-16, are developmentally regulated subunits of chromatin accessibility complex (CHRAC) EMBO J 19: 3049-3059.

Cortez D, Wang Y, Qin J & Elledge SJ (1999) Requirement of ATM-dependent phosphorylation of brca1 in the DNA damage response to double-strand breaks. Science 286: 1162-1166.

Coverley D, Wilkinson HR, Madine MA, Mills AD & Laskey RA (1998) Protein kinase inhibition in G2 causes mammalian Mcm proteins to reassociate with chromatin and restores ability to replicate. Exp Cell Res 238: 63–69.

de Laat WL, Appeldoorn E, Sugasawa K, Weterings E, Jaspers NG & Hoeijmakers JH (1998) DNA-binding polarity of human replication protein A positions nucleases in nucleotide excision repair. Genes Dev 12: 2598–2609.

Delmolino LM, Saha P & Dutta A (2001) Multiple mechanisms regulate subcellular localization of human CDC6. J Biol Chem 276: 26947–26954.

Desdouets C, Santocanale C, Drury LS, Perkins G, Foiani M, Plevani P & Diffley JF (1998) Evidence for a Cdc6p-independent mitotic resetting event involving DNA polymerase α. EMBO J 1998 17: 4139-4146.

Diede SJ & Gottschling DE (1999) Telomerase-mediated telomere addition in vivo requires DNA primase and DNA polymerases α and δ. Cell 99: 723-733.

Donahue BA, Yin S, Taylor JS, Reines D & Hanawalt PC (1994) Transcript cleavage by RNA polymerase II arrested by a cyclobutane pyrimidine dimer in the DNA template. Proc Natl Acad Sci USA 91: 8502-8506.

Drapkin R, Reardon JT, Ansari A, Huang JC, Zawel L, Ahn K, Sancar A & Reinberg D (1994) Dual role of TFIIH in DNA excision repair and in transcription by RNA polymerase II. Nature 368: 769-772.

Drury LS, Perkins G & Diffley JFX (2000) Cdc28p regulates distinct modes of Cdc6p proteolysis during the budding yeast cell cycle. Curr Biol 10: 231−240.

Dua R, Levy DL & Campbell JL (1999) Analysis of the essential functions of the C-terminal protein/protein interaction domain of Saccharomyces cerevisiae pol ε and its unexpected ability to support growth in the absence of the DNA polymerase domain. J Biol Chem 274: 22283-22288.

60

Dua R, Levy DL, Li CM, Snow PM & Campbell JL (2002) In vivo reconstitution of Saccharomyces cerevisiae DNA polymerase ε in insect cells and characterization. J Biol Chem 277: 7889-7896.

Dubaele S, Proietti De Santis L, Bienstock RJ, Keriel A, Stefanini M, Van Houten B & Egly JM (2003) Basal transcription defect discriminates between xeroderma pigmentosum and trichothiodystrophy in XPD patients. Mol Cell 11: 1635-1646.

Echols H & Goodman MF (1991) Fidelity mechanisms in DNA replication. Annu Rev Biochem 60: 477-511.

Edwards S, Li CM, Levy DL, Brown J, Snow PM & Campbell JL (2003) Saccharomyces cerevisiae DNA polymerase ε and polymerase σ interact physically and functionally, suggesting a role for polymerase ε in sister chromatid cohesion. Mol Cell Biol 23: 2733-2748.

Edwards MC, Tutter AV, Cvetic C, Gilbert CH, Prokhorova TA & Walter JC (2002) MCM2-7 complexes bind chromatin in a distributed pattern surrounding the origin recognition complex in Xenopus egg extracts. J Biol Chem 277: 33049-33057.

Eisenman RN (2001) Deconstructing Myc. Genes Dev 15: 2023-2030. Eliasson R & Reichard P (1978) Replication of polyoma DNA in isolated nuclei. Synthesis and

distribution of initiator RNA. J Biol Chem 253: 7469-7475. Evans E, Moggs JG, Hwang JR, Egly J-M & Wood RD (1997) Mechanism of open complex and

dual incision formation by human nucleotide excision repair factors. EMBO J 16: 6559–6573. Falck J, Coates J & Jackson SP (2005) Conserved modes of recruitment of ATM, ATR and DNA-

PKcs to sites of DNA damage. Nature 434: 605-611. Feng W, Rodriguez-Menocal L, Tolun G & D´Urso G (2003) Schizosaccharomyces pombe Dpb2

binds to origin DNA early in S phase and is required for chromosomal DNA replication. Mol Biol Cell 14: 3427-3436.

Feng, W & D'Urso G (2001) Schizosaccharomyces pombe cells lacking the aminoterminal catalytic domains of DNA polymerase ε are viable but require the DNA damage checkpoint control. Mol Cell Biol 21: 4495-4504.

Feng XH, Liang YY, Liang M, Zhai W & Lin X (2002) Direct interaction of c-Myc with Smad2 and Smad3 to inhibit TGF-β-mediated induction of the CDK inhibitor p15(Ink4B). Mol Cell 9: 133–143.

Ferguson DO, Sekiguchi JM, Chang S, Frank KM, Gao Y, DePinho RA & Alt FW (2000) The non-homologous end-joining pathway of DNA repair is required for genomic stability and the suppression of translocations. Proc Natl Acad Sci USA 97: 6630-66330.

Fien K, Cho YS, Lee JK, Raychaudhuri S, Tappin I & Hurwitz J (2004) Primer utilization by DNA polymerase α-primase is influenced by its interaction with Mcm10p. J Biol Chem 279: 16144-16153.

Fijalkowska IJ, Jonczyk P, Tkaczyk MM, Bialoskorska M & Schaaper RM (1998) Unequal fidelity of leading strand and lagging strand DNA replication on the Escherichia coli chromosome. Proc Natl Acad Sci USA 95: 10020-10025.

Filippov VM, Filippov M & Gill SS (2001) Drosophila RNase H1 is essential for development but not for proliferation. Mol Genet Genomics 265: 771-777.

Franklin MC, Wang J & Steitz TA (2001) Structure of the replicating complex of a pol α family DNA polymerase. Cell 105: 657-667.

Friedman LS, Ostermeyer EA, Szabo CI, Dowd P, Lynch ED, Rowell SE & King MC (1994) Confirmation of BRCA1 by analysis of germline mutations linked to breast and ovarian cancer in ten families. Nat Genet 8: 399-404.

Fukui T, Yamauchi K, Muroya T, Akiyama M, Maki H, Sugino A & Waga S (2004) Distinct roles of DNA polymerases δ and ε at the replication fork in Xenopus egg extracts. Genes Cells 9: 179-191.

61

Furuya K, Poitelea M, Guo L, Caspari T & Carr AM (2004) Chk1 activation requires Rad9 S/TQ-site phosphorylation to promote association with C-terminal BRCT domains of Rad4TOPBP1. Genes Dev 18: 1154-1164.

Fuss J & Linn S (2002) Human DNA polymerase ε colocalizes with proliferating cell nuclear antigen and DNA replication late, but not early, in S phase. J Biol Chem 277: 8658-8666.

Gaidamakov SA, Gorshkova II, Schuck P, Steinbach PJ, Yamada H, Crouch RJ & Cerritelli SM (2005) Eukaryotic RNases H1 act processively by interactions through the duplex RNA-binding domain. Nucleic Acids Res 33: 2166-2175.

Gale JM, Nissen KA & Smerdon MJ (1987) UV-induced formation of pyrimidine dimers in nucleosome core DNA is strongly modulated with a period of 10.3 bases. Proc Natl Acad Sci USA 84: 6644-6648.

Gambus A, Jones RC, Sanchez-Diaz A, Kanemaki M, van Deursen F, Edmondson RD & Labib K (2006) GINS maintains association of Cdc45 with MCM in replisome progression complexes at eukaryotic DNA replication forks. Nat Cell Biol 8: 358-366.

Gauthier L, Dziak R, Kramer DJ, Leishman D, Song X, Ho J, Radovic M, Bentley D & Yankulov K (2002) The role of the carboxyterminal domain of RNA polymerase II in regulating origins of DNA replication in Saccharomyces cerevisiae. Genetics 162: 1117-1129.

Garg P & Burgers PMJ (2005a) DNA Polymerases that Propagate the Eukaryotic DNA replication Fork. Crit Rev Biochem Mol Biol 40: 115-128.

Garg P & Burgers PMJ (2005b) Ubiquitinated proliferating cell nuclear antigen activates translesion DNA polymerases eta and REV1. Proc Natl Acad Sci USA 102: 18361-18366.

Garg P, Stith CM, Majka J & Burgers PMJ (2005) Proliferating cell nuclear antigen promotes translesion synthesis by DNA polymerase zeta. J Biol Chem 280: 23446-23450.

Garg P, Stith CM, Sabouri N, Johansson E & Burgers PMJ (2004) Idling by DNA polymerase δ maintains a ligatable nick during lagging-strand DNA replication. Genes Dev 18: 2764-2773.

Gerik KJ, Li X, Pautz A & Burgers PM J (1998) Characterization of the two small subunits of Saccharomyces cerevisiae DNA polymerase δ. J Biol Chem. 273: 19747-19755.

Ghosh M, Liu G, Randall G, Bevington J & Leffak M (2004) Transcription factor binding and induced transcription alter chromosomal c-myc replicator activity. Mol Cell Biol 24: 10193-207.

Gomes XV, Burgers PM (2001) ATP utilization by yeast replication factor C. I. ATP-mediated interaction with DNA and with proliferating cell nuclear antigen. J Biol Chem 276: 34768-34775.

Gomes XV, Schmidt SL & Burgers PM (2001) ATP utilization by yeast replication factor C. II. Multiple stepwise ATP binding events are required to load proliferating cell nuclear antigen onto primed DNA. J Biol Chem 276: 34776-34783.

Gong XQ, Nedialkov YA & Burton ZF (2004) α-Amanitin blocks translocation by human RNA polymerase II. J Biol Chem 279: 27422-27427.

Gopalakrishnan V, Simancek P, Houchens C, Snaith HA, Frattini MG, Sazer S & Kelly TJ (2001) Redundant control of rereplication in fission yeast. Proc Natl Acad Sci USA 98: 13114-13119.

Gozuacik D, Chami M, Lagorce D, Faivre J, Murakami Y, Poch O, Biermann E, Knippers R, Brechot C & Paterlini-Brechot P (2003) Identification and functional characterization of a new member of the human Mcm protein family: hMcm8. Nucleic Acids Res 31: 570-579.

Greenberg RA, Sobhian B, Pathania S, Cantor SB, Nakatani Y & Livingston DM (2006) Multifactorial contributions to an acute DNA damage response by BRCA1/BARD1-containing complexes. Genes Dev 20: 34-46.

Groisman R, Polanowska J, Kuraoka I, Sawada J, Saijo M, Drapkin R, Kisselev AF, Tanaka K & Nakatani Y (2003) The ubiquitin ligase activity in the DDB2 and CSA complexes is differentially regulated by the COP9 signalosome in response to DNA damage. Cell 113: 357-367.

62

Gulbis JM, Kelman Z, Hurwitz J, O'Donnell M & Kuriyan J (1996) Structure of the C-terminal region of p21(WAF1/CIP1) complexed with human PCNA. Cell 87: 297-306.

Guzder SN, Habraken Y, Sung P, Prakash L & Prakash S (1995) Reconstitution of yeast nucleotide excision repair with purified Rad proteins, replication protein A, and transcription factor TFIIH. J Biol Chem 270: 12973-12976.

Guzder SN, Sung P, Prakash L & Prakash S (1998) Affinity of yeast nucleotide excision repair factor 2, consisting of the Rad4 and Rad23 proteins, for ultraviolet damaged DNA. J Biol Chem 273: 31541-31546.

Guzder SN, Sung P, Prakash L & Prakash S (1999) Synergistic interaction between yeast nucleotide excision repair factors NEF2 and NEF4 in the binding of ultraviolet-damaged DNA. J Biol Chem 274: 24257-24262.

Harrington JJ & Lieber R (1994) The characterization of a mammalian DNA structure-specific endonuclease. Embo J 13: 1235-1246.

Hartlepp KF, Fernandez-Tornero C, Eberharter A, Grune T, Muller CW & Becker PB (2005) The histone fold subunits of Drosophila CHRAC facilitate nucleosome sliding through dynamic DNA interactions. Mol Cell Biol 25: 9886-9896.

Hashimoto Y & Takisawa H (2003) Xenopus Cut5 is essential for a CDK-dependent process in the initiation of DNA replication. EMBO J 22: 2526-2535.

Hiraga, S, Hagihara-Hayashi A, Ohya T & Sugino A (2005) DNA polymerases α, δ, and ε localize and function together at replication forks in Saccharomyces cerevisiae. Genes Cells 10: 297-309.

Holland L, Gauthier L, Bell-Rogers P & Yankulov K (2002) Distinct parts of minichromosome maintenance protein 2 associate with histone H3/H4 and RNA polymerase II holoenzyme. Eur J Biochem 269: 5192-51202.

Holland L & Yankulov K (2003) Two forms of RNA polymerase II holoenzyme display different abundance during the cell cycle. Biochem Biophys Res Commun 302: 484-488.

Huang D, Jokela M, Tuusa J, Skog S, Poikonen K, Syväoja JE (2001) E2F mediates induction of the Sp1-controlled promoter of the human DNA polymerase ε B-subunit gene POLE2. Nucleic Acids Res 29: 2810-2821.

Huang ME, Rio AG, Galibert MD & Galibert F (2002) Pol32, a subunit of Saccharomyces cerevisiae DNA polymerase δ, suppresses genomic deletions and is involved in the mutagenic bypass pathway. Genetics 160: 1409-1422.

Hübscher U, Maga G & Spadari S (2002) Eukaryotic DNA polymerases. Annu Rev Biochem 71: 133-163.

Iborra FJ, Pombo A, Jackson DA & Cook PR (1996) Active RNA polymerases are localized within discrete transcription "factories" in human nuclei. J. Cell Sci 109: 1427-1436.

Iida T & Araki H (2004) Noncompetitive counteractions of DNA polymerase ε and ISW2/yCHRAC for epigenetic inheritance of telomere position effect in Saccharomyces cerevisiae. Mol Cell Biol 24: 217-227.

Ishimi Y (1997) A DNA helicase activity is associated with an MCM4, -6, and -7 protein complex. J Biol Chem 272: 24508-24513.

Ishino Y, Nishino T & Morikawa K (2006) Mechanisms of maintaining genetic stability by homologous recombination. Chem Rev 106: 324-339.

Jares P & Blow JJ (2000) Xenopus cdc7 function is dependent on licensing but not on XORC, XCdc6, or CDK activity and is required for XCdc45 loading. Genes Dev 14: 1528-1540.

Jin YH, Ayyagari R, Resnick MA, Gordenin DA & Burgers PM (2003) Okazaki fragment maturation in yeast. II. Cooperation between the polymerase and 3'-5'-exonuclease activities of Pol δ in the creation of a ligatable nick. J Biol Chem 1626-1633.

63

Jin YH, Garg P, Stith CM, Al-Refai H, Sterling JF, Murray LJ, Kunkel TA, Resnick MA, Burgers PMJ & Gordenin DA (2005) The multiple biological roles of the 3'-->5' exonuclease of Saccharomyces cerevisiae DNA polymerase δ require switching between the polymerase and exonuclease domains. Mol Cell Biol 25: 461-471.

Jin YH, Obert R, Burgers PM, Kunkel TA, Resnick MA &Gordenin DA (2001) The 3'-->5' exonuclease of DNA polymerase δ can substitute for the 5' flap endonuclease Rad27/Fen1 in processing Okazaki fragments and preventing genome instability. Proc Natl Acad Sci USA 98: 5122-5127.

Johansson E, Garg P & Burgers PMJ (2004) The Pol32 subunit of DNA polymerase δ contains separable domains for processive replication and proliferating cell nuclear antigen (PCNA) binding. J Biol Chem 279: 1907-1915.

Johansson E, Majka J & Burgers PMJ (2001) Structure of DNA polymerase δ from Saccharomyces cerevisiae. J Biol Chem 276: 43824-43828.

Johnson SK, Bhattacharyya S & Griep MA (2000) DnaB helicase stimulates primer synthesis activity on short oligonucleotide templates. Biochemistry 39: 736-744.

Jonsson ZO, Hindges R & Hübscher U (1998) Regulation of DNA replication and repair proteins through interaction with the front side of proliferating cell nuclear antigen. EMBO J 17: 2412-2425.

Jun SH, Kim TG & Ban C (2006) DNA mismatch repair system. Classical and fresh roles. FEBS J 273: 1609-1619.

Kaczmarek L, Miller MR, Hammond RA & Mercer WE (1986) A microinjected monoclonal antibody against human DNA polymerase-α inhibits DNA replication in human, hamster, and mouse cell lines. J Biol Chem 261: 10802-10807.

Kaldis P (1999) The cdk-activating kinase (CAK): from yeast to mammals. Cell Mol Life Sci 55: 284-296.

Kamath S & Leffak M (2001) Multiple sites of replication initiation in the human β-globin locus. Nucleic Acids Res 29: 809-817.

Kamimura Y, Masumoto H, Sugino A & Araki H (1998) Sld2, which interacts with Dpb11 in Saccharomyces cerevisiae, is required for chromosomal DNA replication. Mol Cell Biol 18: 6102-6109.

Kamimura Y, Tak Y-S, Sugino A & Araki H (2001) Sld3, which interacts with Cdc45 (Sld4), functions for chromosomal DNA replication in Saccharomyces cerevisiae. Embo J 20: 2097-2107.

Kanemaki M & Labib K (2006) Distinct roles for Sld3 and GINS during establishment and progression of eukaryotic DNA replication forks. EMBO J 25: 1753-1763.

Kao H-I, Veeraraghavan J, Polaczek P, Campbell JL & Bambara RA (2004) On the roles of Saccharomyces cerevisiae Dna2p and Flap endonuclease 1 in Okazaki fragment processing. J Biol Chem 279: 15014-15024.

Karthikeyan R, Vonarx EJ, Straffon AF, Simon M, Faye G & Kunz BA (2000) Evidence from mutational specificity studies that yeast DNA polymerases δ and ε replicate different DNA strands at an intracellular replication fork. J Mol Biol 299: 405-419.

Kearsey SE & Cotterill S (2003) Enigmatic variations: divergent modes of regulating eukaryotic DNA replication. Mol Cell 12: 1067-1075.

Kelleher RJ 3rd, Flanagan PM & Kornberg RD (1990) A novel mediator between activator proteins and the RNA polymerase II transcription apparatus. Cell 1990 61: 1209-1215.

Kesti T, Flick K, Keränen S, Syväoja JE & Wittenberg C (1999) DNA polymerase ε catalytic domains are dispensable for DNA replication, DNA repair, and cell viability. Mol Cell 3: 679-685.

Kim DK, Yamaguchi Y, Wada T & Handa H (2001) The regulation of elongation by eukaryotic RNA polymerase II: a recent view. Mol Cells 11: 267-274.

64

Kim JE, McAvoy SA, Smith DI & Chen J (2005) Human TopBP1 ensures genome integrity during normal S phase. Mol Cell Biol 25: 10907-10915.

Kim Y-J, Bjorklund S, Li Y, Sayre MH & Kornberg RD (1994) A multiprotein mediator of transcriptional activation and its interaction with the C-terminal repeat domain of RNA polymerase II. Cell 77: 599-608.

Kim YK, Bourgeois CF, Isel C, Churcher MJ & Karn J (2002) Phosphorylation of the RNA polymerase II carboxyl-terminal domain by CDK9 is directly responsible for human immunodeficiency virus type 1 Tat-activated transcriptional elongation. Mol Cell Biol 22: 4622–4637.

Klungland A, Hoss M, Gunz D, Constantinou A, Clarkson SG, Doetsch PW, Bolton PH, Wood RD & Lindahl T (1999) Base excision repair of oxidative DNA damage activated by XPG protein. Mol Cell 3: 33-42.

Kohzaki H, Ito Y & Murakami Y (1999) Context-dependent modulation of replication activity of Saccharomyces cerevisiae autonomously replicating sequences by transcription factors. Mol Cell Biol 19: 7428-7435.

Kohzaki H & Murakami Y (2005) Transcription factors and DNA replication origin selection. Bioessays 27: 1107-1116.

Kreitz S, Ritzi M, Baack M & Knippers R (2001) The human origin recognition complex protein 1 dissociates from chromatin during S phase in HeLa cells. J Biol Chem 276: 6337-6342.

Krishna TS, Kong XP, Gary S, Burgers PM & Kuriyan J (1994) Crystal structure of the eukaryotic DNA polymerase processivity factor PCNA. Cell 79: 1233-1243.

Krum SA, Miranda GA, Lin C & Lane TF (2003) BRCA1 associates with processive RNA polymerase II. J Biol Chem 278: 52012-52020.

Kubota Y, Mimura S, Nishimoto S, Masuda T, Nojima H & Takisawa H (1997) Licensing of DNA replication by a multi-protein complex of MCM/P1 proteins in Xenopus eggs. EMBO J 16: 3320-3331.

Kumagai A, Lee J, Yoo HY & Dunphy WG (2006) TopBP1 activates the ATR-ATRIP complex. Cell 124: 943-955.

Lai JS & Herr W (1992) Ethidium bromide provides a simple tool for identifying genuine DNA-independent protein associations. Proc Natl Acad Sci USA 89: 6958-6962.

Lao Y, Gomes XV, Ren Y, Taylor JS & Wold MS (2000) Replication protein A interactions with DNA. III. Molecular basis of recognition of damaged DNA. Biochemistry 39: 850-859.

Laskey RA & Madine MA (2003) A rotary pumping model for helicase function of MCM proteins at a distance from replication forks. EMBO Rep 4: 26-30.

Lee C Hong B, Choi JM, Kim Y, Watanabe S, Ishimi Y, Enomoto T, Tada S & Cho Y (2004) Structural basis for inhibition of the replication licensing factor Cdt1 by geminin. Nature 430: 913-917.

Lee JK & Hurwitz J (2001) Processive DNA helicase activity of the minichromosome maintenance proteins 4, 6, and 7 complex requires forked DNA structures. Proc Natl Acad Sci USA 98: 54-59.

Lee K-H, Kim DW, Bae S-H, Kim J-A, Ryu G-H, Kwon Y-N, Kim K-A, Koo H-S & Seo Y-S (2000) The endonuclease activity of the yeast Dna2 enzyme is essential in vivo. Nucleic Acids Res 28: 2873-2881.

Lee S-H, Pan Z-Q, Kwong AD, Burgers PMJ & Hurwitz J (1991) Synthesis of DNA by DNA polymerase ε in vitro. J Biol Chem 266: 22707-22717.

Lehmann AR (2003) DNA repair-deficient diseases, xeroderma pigmentosum, Cockayne syndrome and trichothiodystrophy. Biochimie 85: 1101-1111.

Li A & Blow JJ (2005) Cdt1 downregulation by proteolysis and geminin inhibition prevents DNA re-replication in Xenopus. EMBO J 24: 395−404.

65

Li C-J & DePamphilis ML (2002) Mammalian Orc1 protein is selectively released from chromatin and ubiquitinated during the S-to-M transition in the cell division cycle. Mol Cell Biol 22: 105–116.

Li JJ & Kelly TJ (1984) Simian virus 40 DNA replication in vitro. Proc Natl Acad Sci USA 81: 6973-6977.

Li L, Lu X, Peterson CA & Legerski RJ (1995) An interaction between the DNA repair factor XPA and Replication protein A appears essential for nucleotide excision repair. Mol Cell Biol 15: 5396-5402.

Li Y, Pursell ZF & Linn S (2000) Identification and cloning of two histone fold motif-containing subunits of Hela DNA polymerase ε. J Biol Chem 275: 23247-23252.

Limoli CL, Giedzinski E, Morgan WF & Cleaver JE (2000) Polymerase η deficiency in the xeroderma pigmentosum variant uncovers an overlap between the S phase checkpoint and double-strand break repair. Proc Natl Acad Sci USA 97: 7939-7946.

Liu B & Alberts BM (1995) Head-on collision between a DNA replication apparatus and RNA polymerase transcription complex. Science 267: 1131-1137.

Liu B, Wong ML, Tinker RL, Geiduschek EP & Alberts BM (1993) The DNA replication fork can pass RNA polymerase without displacing the nascent transcript. Nature 366: 33-39.

Liu K, Lin FT, Ruppert JM & Lin WC (2003) Regulation of E2F1 by BRCT domain-containing protein TopBP1. Mol Cell Biol: 3287-3304.

Liu L, Mo, J, Rodrigez-Belmonte EM & Lee MYWT (2000) Identification of a fourth subunit of mammalian DNA polymerase δ. J Biol Chem 275: 18739-18744.

Liu, Y., H. I. Kao and R. A. Bambara (2004) Flap endonuclease 1: a central component of DNA metabolism. Annu Rev Biochem 73: 589-615.

Lupardus PJ & Cimprich KA (2006) Phosphorylation of Xenopus Rad1 and Hus1 defines a readout for ATR activation that is independent of Claspin and the Rad9 carboxy terminus. Mol Biol Cell 17: 1559-1569.

MacAlpine DM, Rodriguez HK & Bell SP (2004) Coordination of replication and transcription along a Drosophila chromosome. Genes Dev 18: 3094-3105.

Maga G & Hübscher U (2003) Proliferating cell nuclear antigen (PCNA): a dancer with many partners. J Cell Sci 116: 3051-3060.

Maga G, Stucki M, Spadari S & Hübscher U (2000) DNA polymerase switching: I. Replication factor C displaces DNA polymerase α prior to PCNA loading. J Mol Biol 295: 791-801.

Mahbubani HM, Chong JP, Chevalier S, Thömmes P & Blow JJ (1997) Cell cycle regulation of the replication licensing system: involvement of a Cdk-dependent inhibitor. J Cell Biol 136: 125–135.

Mailand N & Diffley JF (2005) CDKs promote DNA replication origin licensing in human cells by protecting Cdc6 from APC/C-dependent proteolysis. Cell 122: 915-926.

Maiorano D, Cuvier O, Danis E & Mechali M (2005) MCM8 is an MCM2-7-related protein that functions as a DNA helicase during replication elongation and not initiation. Cell 120: 315-328.

Maiorano D, Krasinska L, Lutzmann M & Mechali M (2005) Recombinant Cdt1 induces rereplication of G2 nuclei in Xenopus egg extracts. Curr Biol 15: 146-153.

Maldonado E, Shiekhattar R, Sheldon M, Cho H, Drapkin R, Rickert P, Lees E, Anderson CW, Linn S & Reinberg D (1996) A human RNA polymerase II complex associated with SRB and DNA-repair proteins. Nature 381: 86-89.

Mäkiniemi M, Pospiech, H, Kilpeläinen S, Jokela M, Vihinen M & Syväoja JE (1999) A novel family of DNA-polymerase-associated B subunits. Trends Biochem Sci 24: 14-16.

Marini F, Kim N, Schuffert A & Wood RD (2003) POLN, a nuclear PolA family DNA polymerase homologous to the DNA cross-link sensitivity protein Mus308. J Biol Chem 278: 32014-32019.

66

Masuda T, Mimura S & Takisawa H (2003) CDK- and Cdc45-dependent priming of the MCM complex on chromatin during S-phase in Xenopus egg extracts: possible activation of MCM helicase by association with Cdc45. Genes Cells 8: 145-161.

Masumoto H, Muramatsu S, Kamimura Y & Araki H (2002) S-Cdk-dependent phosphorylation of Sld2 essential for chromosomal DNA replication in budding yeast. Nature 415: 651-655.

Masumoto H, Sugino A & Araki H (2000) Dpb11 controls the association between DNA polymerases α and ε and the autonomously replicating sequence region of budding yeast. Mol Cell Biol 20: 2809-2817.

Masutani C, Kusumoto R, Yamada A, Dohmae N, Yokoi M, Yuasa M, Araki M, Iwai S, Takio K & Hanaoka F (1999) The XPV (xeroderma pigmentosum variant) gene encodes human DNA polymerase η. Nature 399: 700-704.

McConnell M, Miller H, Mozzherin DJ, Quamina A, Tan C-K, Downey KM & Fisher PA (1996) The Mammalian DNA Polymerase -Proliferating Cell Nuclear Antigen-Template-Primer Complex: Molecular Characterization by Direct Binding. Biochemistry 35: 8268 -8274.

McGarry TJ & Kirschner MW (1998) Geminin, an inhibitor of DNA replication, is degraded during mitosis. Cell 93: 1043-1053.

McMahon SB, Van Buskirk HA, Dugan KA, Copeland TD & Cole MD (1998) The novel ATM-related protein TRRAP is an essential cofactor for the c-Myc and E2F oncoproteins Cell 94: 363-374.

McNairn AJ, Okuno Y, Misteli T & Gilbert DM (2005) Chinese hamster ORC subunits dynamically associate with chromatin throughout the cell-cycle. Exp Cell Res 308: 345-536.

Melle C & Nasheuer H-P (2002) Physical and functional interactions of the tumor suppressor protein p53 and DNA polymerase α-primase. Nucleic Acids Res 30: 1493-1499.

Mimura S, Masuda T, Matsui T & Takisawa H (2000) Central role for Cdc45 in establishing an initiation complex of DNA replication in Xenopus egg extract. Genes Cells 5: 439-452.

Mo X & Dynan WS (2002) Subcellular localization of Ku protein: functional association with RNA polymerase II elongation sites. Mol Cell Biol 22: 8088-8099.

Mone MJ, Volker M, Nikaido O, Mullenders LH, van Zeeland AA, Verschure PJ, Manders EM & van Driel R (2001) Local UV-induced DNA damage in cell nuclei results in local transcription inhibition. EMBO Rep 2: 1013-1017.

Moore JK & Haber JE (1996) Cell cycle and genetic requirements of two pathways of nonhomologous end-joining repair of double-strand breaks in Saccharomyces cerevisiae. Mol Cell Biol 16: 2164-2173.

Morrison A, Araki H, Clark AB, Hamatake RK & Sugino A (1990) A third essential DNA polymerase in S. cerevisiae. Cell 62: 1143-1151.

Mossi R, Keller RC, Ferrari E & Hübscher U (2000) DNA polymerase switching: II. Replication factor C abrogates primer synthesis by DNA polymerase α at a critical length. J Mol Biol 295: 803-814.

Mozzherin DJ, Tan CK, Downey KM & Fisher PA (1999) Architecture of the active DNA polymerase δ.proliferating cell nuclear antigen.template-primer complex. J Biol Chem 274: 19862-19867.

Murakami Y & Ito Y (1999) Transcription factors in DNA replication. Front Biosci 4: d824-d833. Nagase T, Seki N, Ishikawa K, Ohira M, Kawarabayasi Y, Ohara O, Tanaka A, Kotani H,

Miyajima N & Nomura N (1996) Prediction of the Coding Sequences of Unidentified Human Genes. VI. The Coding Sequences of 80 New Genes (KIAA0201-KIAA0280) Deduced by Analysis of cDNA Clones from Cell Line KG-1 and Brain. DNA Res 3: 321-329.

Nakajima R & Masukata H (2002) SpSld3 is required for loading and maintenance of SpCdc45 on chromatin in DNA replication in fission yeast. Mol Biol Cell 13: 1462-1472.

Nasheuer H-P, Moore A, Wahl AF & Wang TSF (1991) Cell cycle-dependent phosphorylation of human DNA polymerase α. J Biol Chem 266: 7893-7903.

67

Nasheuer HP, Pospiech H & Syväoja JE (2006) Progress towards the anatomy of the eukaryotic DNA replication fork. In: Lankenau D-H (ed) Genome Integrity: Facets and Perspectives. Springer-Verlag, Berlin Heidelberg: 27-68. DOI: 10.1007/7050_016.

Navas TA, Zhou Z & Elledge SJ (1995) DNA polymerase ε links the DNA replication machinery to the S phase checkpoint. Cell 80: 29-39.

Neish AS, Anderson SF, Schlegel BP, Wei W & Parvin JD (1998) Factors associated with the mammalian RNA polymerase II holoenzyme. Nucleic Acids Res 26: 847-853.

Nethanel T & Kaufmann G (1990) Two DNA polymerases may be required for synthesis of the lagging strand DNA of simian virus 40. J Virol 64: 5912-5918.

Nguyen VQ, Co C & Li JJ (2001) Cyclin-dependent kinases prevent DNA re-replication through multiple mechanisms. Nature 411: 1068–1073.

Nichols AF, Itoh T, Graham JA, Liu W, Yamaizumi M & Linn S (2000) Human damage-specific DNA-binding protein p48. Characterization of XPE mutations and regulation following UV irradiation. J Biol Chem 275: 21422-21428.

Noton E & Diffley JF (2000) CDK inactivation is the only essential function of the APC/C and the mitotic exit network proteins for origin resetting during mitosis. Mol Cell 5: 85-95.

O'Donovan A, Davies AA, Moggs JG, West SC & Wood RD (1994) XPG endonuclease makes the 3' incision in human DNA nucleotide excision repair. Nature 371: 432-435.

Ogawa Y, Takahashi T & Masukata H (1999) Association of fission yeast Orp1 and Mcm6 proteins with chromosomal replication origins. Mol Cell Biol 19: 7228-7236.

Ohya T, Maki S, Kawasaki Y & Sugino A (2000) Structure and function of the fourth subunit (Dpb4p) of DNA polymerase ε in Saccharomyces cerevisiae. Nucleic Acids Res 28: 3846-3852.

Otterlei M, Warbrick E, Nagelhus TA, Haug T, Slupphaug G, Akbari M, Aas PA, Steinsbekk K, Bakke O & Krokan HE (1999) Post-replicative base excision repair in replication foci. Embo J 18: 3834-3844.

Pacek M, Tutter AV, Kubota Y, Takisawa H & Walter JC (2006) Localization of MCM2-7, Cdc45, and GINS to the site of DNA unwinding during eukaryotic DNA replication. Mol Cell 21: 581-587.

Palancade B & Bensaude O (2003) Investigating RNA polymerase II carboxyl-terminal domain (CTD) phosphorylation. Eur J Biochem 270: 3859-3870.

Park C-H, Mu D, Reardon JT & Sancar A (1995) The general transcription-repair factor TFIIH is recruited to the excision repair complex by the XPA protein independent of the TFIIE transcription factor. J Biol Chem 270: 4896–4902.

Park J, Kunjibettu S, McMahon SB & Cole MD (2001) The ATM-related domain of TRRAP is required for histone acetyltransferase recruitment and Myc-dependent oncogenesis. Genes Dev 15: 1619–1624.

Pascal JM, O'Brien PJ, Tomkinson AE, Ellenberger T (2004) Human DNA ligase I completely encircles and partially unwinds nicked DNA. Nature 432: 473-478.

Pasero P & Gasser SM (1998) New systems for replicating DNA in vitro. Curr Opin Cell Biol 10: 304-310.

Pastwa E & Blasiak J (2003) Non-homologous DNA end joining. Acta Biochim Pol 50: 891-908. Paule MR & White RJ (2000) Survey and summary: transcription by RNA polymerases I and III.

Nucleic Acids Res 28: 1283-1298. Pavlov YI, Frahm C, Nick McElhinny SA, Niimi A, Suzuki M, Kunkel TA (2006) Evidence that

errors made by DNA polymerase α are corrected by DNA polymerase δ. Curr Biol 16: 202-207. Pavlov YI, Shcherbakova PV & Kunkel TA (2001) In vivo consequences of putative active site

mutations in yeast DNA polymerases α, ε, δ, and ζ. Genetics 159: 47-64. Petersen BO, Wagener C, Marinoni F, Kramer ER, Melixetian M, Denchi EL, Gieffers C,

Matteucci C, Peters J-M & Helin K (2000) Cell cycle- and cell growth-regulated proteolysis of mammalian CDC6 is dependent on APC-CDH1. Genes Dev 14: 2330-2343.

68

Philimonenko AA, Hodny Z, Jackson DA & Hozak P (2006) The microarchitecture of DNA replication domains. Histochem Cell Biol 125: 103-117.

Podust VN, Chang LS, Ott R, Dianov GL & Fanning E (2002) Reconstitution of human DNA polymerase δ using recombinant baculoviruses: the p12 subunit potentiates DNA polymerizing activity of the four-subunit enzyme. J Biol Chem 277: 3894-3901.

Poot RA, Dellaire G, Hulsmann BB, Grimaldi MA, Corona DF, Becker PB, Bickmore WA & Varga-Weisz PD (2000) HuCHRAC, a human ISWI chromatin remodelling complex contains hACF1 and two novel histone-fold proteins. EMBO J 19: 3377-3387.

Pospiech H, Kursula I, Abdel-Aziz W, Malkas L, Uitto L, Kastelli M, Vihinen-Ranta M, Eskelinen S & Syväoja JE (1999) A neutralizing antibody against human DNA polymerase ε inhibits cellular but not SV40 DNA replication. Nucleic Acids Res 27: 3799-3804.

Pötter T, Göhde W, Wedemeyer N & Köhnlein W (2000) Keratinocytes exposed to ultraviolet radiation reveal three down-regulated genes with potential function in differentiation and cell cycle control. Radiat Res 154: 51-15.

Prakash S, Johnson RE & Prakash L (2005) Eukaryotic translesion synthesis DNA polymerases: specificity of structure and function. Annu Rev Biochem 74: 317-353.

Prelich G, Tan C-K, Kostura M, Mathews MB, So AG, Downey KM & Stillman B (1987) Functional identity of proliferating cell nuclear antigen and a DNA polymerase-δ auxiliary protein. Nature 326: 517-520.

Prigent C, Satoh MS, Daly G, Barnes DE & Lindahl T (1994) Aberrant DNA repair and DNA replication due to an inherited enzymatic defect in human DNA ligase I. Mol Cell Biol 14: 310-317.

Radichev I, Parashkevova A & Anachkova B (2005) Initiation of DNA replication at a nuclear matrix-attached chromatin fraction. J Cell Physiol 203: 71-77.

Raghuraman MK, Winzeler EA, Collingwood D, Hunt S, Wodicka L, Conway A, Lockhart DJ, Davis RW, Brewer BJ & Fangman WL (2001) Replication dynamics of the yeast genome. Science 294: 115-121.

Rattray AJ & Strathern JN (2003) Error-prone DNA polymerases: when making a mistake is the only way to get ahead. Annu Rev Genet 37: 31-66.

Ricke RM & Bielinsky AK (2004) Mcm10 regulates the stability and chromatin association of DNA polymerase α. Mol Cell 16: 173-185.

Ricke RM & Bielinsky AK (2006) A conserved Hsp10-like domain in Mcm10 is required to stabilize the catalytic subunit of DNA polymerase-α in budding yeast. J Biol Chem 281: 18414-1825.

Robins P, Jones CJ, Biggerstaff M, Lindahl T & Wood RD (1991) Complementation of DNA repair in xeroderma pigmentosum group A cell extracts by a protein with affinity for damaged DNA. EMBO J 10: 3913-3921.

Rockx DA, Mason R, van Hoffen A, Barton MC, Citterio E, Bregman DB, van Zeeland AA, Vrieling H & Mullenders LH (2000) UV-induced inhibition of transcription involves repression of transcription initiation and phosphorylation of RNA polymerase II. Proc Natl Acad Sci USA 97: 10503-10508.

Romanowski P, Madine MA, Rowles A, Blow JJ & Laskey RA (1996) The Xenopus origin recognition complex is essential for DNA replication and MCM binding to chromatin. Curr Biol 6: 1416-1425.

Rossi DJ, Londesborough A, Korsisaari N, Pihlak A, Lehtonen E, Henkemeyer M & Mäkelä TP (2001) Inability to enter S phase and defective RNA polymerase II CTD phosphorylation in mice lacking Mat1. EMBO J 20: 2844-2856.

Rowles A, Tada S & Blow JJ (1999) Changes in association of the Xenopus origin recognition complex with chromatin on licensing of replication origins. J Cell Sci 112: 2011-2018.

69

Rytkönen AK, Vaara M, Nethanel T, Kaufmann G, Sormunen R, Laara E, Nasheuer H-P, Rahmeh A, Lee MY, Syväoja JE & Pospiech H (2006) Distinctive activities of DNA polymerases during human DNA replication. FEBS J 273: 2984-3001.

Saka Y, Esashi F, Matsusaka T, Mochida, S, Yanagida M (1997) Damage and replication checkpoint control in fission yeast is ensured by interactions of Crb2, a protein with BRCT motif, with Cut5 and Chk1. Genes Dev 11: 3387-3400.

Sakurai S, Kitano K, Yamaguchi H, Hamada K, Okada K, Fukuda K, Uchida M, Ohtsuka E, Morioka H & Hakoshima T (2005) Structural basis for recruitment of human flap endonuclease 1 to PCNA. EMBO J 24: 683-693.

Sancar A, Lindsey-Boltz LA, Unsal-Kacmaz K & Linn S (2004) Molecular mechanisms of mammalian DNA repair and the DNA damage checkpoints. Annu Rev Biochem 73: 39-85.

Santocanale C, Foiani M, Lucchini G & Plevani P (1993) The isolated 48,000-dalton subunit of yeast DNA primase is sufficient for RNA primer synthesis. J Biol Chem 268: 1343-1348.

Satoh MS, Jones CJ, Wood RD & Lindahl T (1993) DNA excision-repair defect of xeroderma pigmentosum prevents removal of a class of oxygen free radical-induced base lesions. Proc Natl Acad Sci USA 90: 6335–6339.

Saxena S, Yuan P, Dhar SK, Senga T, Takeda D, Robinson H, Kornbluth S, Swaminathan K & Dutta A (2004) A dimerized coiled-coil domain and an adjoining part of geminin interact with two sites on Cdt1 for replication inhibition. Mol Cell 23: 245-258.

Sawyer SL, Cheng IH, Chai W & Tye BK (2004) Mcm10 and Cdc45 cooperate in origin activation in Saccharomyces cerevisiae. J Mol Biol 340: 195-202.

Schang LM (2004) Effects of pharmacological cyclin-dependent kinase inhibitors on viral transcription and replication. Biochim Biophys Acta 1697: 197-209.

Schwacha A & Bell SP (2001) Interactions between two catalytically distinct MCM subgroups are essential for coordinated ATP hydrolysis and DNA replication. Mol Cell 8: 1093-1104.

Scully R, Chen J, Ochs RL, Keegan K, Hoekstra M, Feunteun J & Livingston DM (1997b) Dynamic changes of BRCA1 subnuclear location and phosphorylation state are initiated by DNA damage. Cell 90: 425-435.

Scully R, Chen J, Plug A, Xiao Y, Weaver, D, Feunteun J, Ashley T & Livingston DM (1997a) Association of BRCA1 with Rad51 in mitotic and meiotic cells. Cell 88: 265-275.

Seeberg E, Eide L & Bjørås M (1995) The base excision repair pathway. Trends Biochem Sci 20: 391-397.

Seoane J, Pouponnot C, Staller P, Schader M, Eilers M & Massague´ J (2001) TGFbeta influences Myc, Miz-1 and Smad to control the CDK inhibitor p15INK4b. Nat Cell Biol 3: 400–408.

Shamoo Y & Steitz TA (1999) Building a replisome from interacting pieces: sliding clamp complexed to a peptide from DNA polymerase and a polymerase editing complex. Cell 99: 155-166.

Shcherbakova PV & Pavlov YI (1996) 3´→5´ exonucleases of DNA polymerases ε and δ correct base analog induced DNA replication errors on opposite DNA strands in Saccharomyces cerevisiae. Genetics 142: 717-726.

Sherr CJ & Roberts JM (2004) Living with or without cyclins and cyclin-dependent kinases. Genes Dev 18: 2699-2711.

Shibahara K & Stillman B (1999) Replication-dependent marking of DNA by PCNA facilitates CAF-1-coupled inheritance of chromatin. Cell 96: 575-585.

Shinohara A & Ogawa T (1995) Homologous recombination and the roles of double strand breaks. Trends Biochem Sci 20: 387-390.

Slot JW & Geuze HJ (1985) A new method of preparing gold probes for multiple-labeling cytochemistry. Eur J Cell Biol 38: 87-93.

Sogo JM, Lopes M & Foiani M (2002) Fork reversal and ssDNA accumulation at stalled replication forks owing to checkpoint defects. Science 297: 599-602.

70

Speck C, Chen Z, Li H & Stillman B (2005) ATPase-dependent cooperative binding of ORC and Cdc6 to origin DNA. Nat Struct Mol Biol 12: 965-971.

Spiga MG & D'Urso G (2004) Identification and cloning of two putative subunits of DNA polymerase ε in fission yeast. Nucleic Acids Res 32: 4945-4953.

Sporbert A, Domaing P, Leonhardt H & Cardoso MC (2005) PCNA acts as a stationary loading platform for transiently interacting Okazaki fragment maturation proteins. Nucleic Acids Res 33: 3521-3528.

Staller P, Peukert K, Kiermaier A, Seoane J, Lukas J, Karsunky H, Moroy T, Bartek J, Massague´ J, Hanel F & Eilers M (2001) Repression of p15INK4b expression by Myc through association with Miz-1. Nat Cell Biol 3: 392–399.

Steitz TA, Smerdon SJ, Jager J & Joyce CM (1994) A unified polymerase mechanism for nonhomologous DNA and RNA polymerases. Science 266: 2022-2025.

Stoeber K, Mills AD, Kubota Y, Krude T, Romanowski P, Marheineke K, Laskey RA & Williams GH (1998) Cdc6 protein causes premature entry into S phase in a mammalian cell-free system. EMBO J 17: 7219-7229.

Sugasawa K, Ng JM, Masutani C, Iwai S, van der Spek PJ, Eker AP, Hanaoka F, Bootsma D & Hoeijmakers JH (1998) Xeroderma pigmentosum group C protein complex is the initiator of global genome nucleotide excision repair. Mol Cell 2: 223-232.

Sugino A (1995) Yeast DNA polymerases and their role at the replication fork. Trends Biochem Sci 20: 319-323.

Sung JS & Demple B (2006) of base excision repair subpathways in correcting oxidized abasic sites in DNA. FEBS J 273: 1620-1629.

Svejstrup JQ (2004) The RNA polymerase II transcription cycle: cycling through chromatin. Biochim Biophys Acta 1677: 64-73.

Svetlova MP, Solovjeva LV, Pleskach NA & Tomilin NV (1999) Focal sites of DNA repair synthesis in human chromosomes. Biochem Biophys Res Commun 257: 378-383.

Syväoja JE & Linn S (1989) Characterization of a large form of DNA polymerase δ from HeLa cells that is insensitive to proliferating cell nuclear antigen. J Biol Chem 264: 2489-2497.

Tada S, Li A, Maiorano D, Mechali M & Blow JJ (2001) Repression of origin assembly in metaphase depends on inhibition of RLF-B/Cdt1 by geminin. Nat Cell Biol 3: 107-113.

Takahashi TS & Walter JC (2005) Cdc7-Drf1 is a developmentally regulated protein kinase required for the initiation of vertebrate DNA replication. Genes Dev 19: 2295-2300.

Takayama Y, Kamimura Y, Okawa M, Muramatsu S, Sugino A & Araki H (2003) GINS, a novel multiprotein complex required for chromosomal DNA replication in budding yeast. Genes Dev 17: 1153-1165.

Tan C-K, Castillo C, So AG & Downey KM (1986) An auxiliary protein for DNA polymerase-δ from fetal calf thymus. J Biol Chem 261: 12310-12316.

Tanaka S & Diffley JF (2002) Interdependent nuclear accumulation of budding yeast Cdt1 and Mcm2-7 during G1 phase. Nat Cell Biol 4: 198-207.

Tanaka T, Knapp D & Nasmyth K (1997) Loading of an Mcm protein onto DNA replication origins is regulated by Cdc6p and CDKs. Cell 90: 649-660.

Tang JY, Hwang BJ, Ford JM, Hanawalt PC & Chu G (2000) Xeroderma pigmentosum p48 gene enhances global genomic repair and suppresses UV-induced mutagenesis. Mol Cell 5: 737-744.

Tantin D, Kansal A & Carey M (1997) Recruitment of the putative transcription-repair coupling factor CSB/ERCC6 to RNA polymerase II elongation complexes. Mol Cell Biol 17: 6803-6814.

Taricani L & Wang TS (2006) Rad4TopBP1, a Scaffold Protein, Plays Separate Roles in DNA Damage and Replication Checkpoints and DNA Replication. Mol Biol Cell 17: 3456-3468.

Tibbetts RS, Cortez D, Brumbaugh KM, Scully R, Livingston D, Elledge SJ & Abraham RT (2000) Functional interactions between BRCA1 and the checkpoint kinase ATR during genotoxic stress. Genes Dev 14: 2989-3002.

71

Thoma F (1999) Light and dark in chromatin repair: repair of UV-induced DNA lesions by photolyase and nucleotide excision repair. EMBO J 18: 6585-6598.

Todorov IT, Attaran A & Kearsey SE (1995) BM28, a human member of the MCM2-3-5 family, is displaced from chromatin during DNA replication. J Cell Biol 129: 1433-1445.

Todorovic V, Falaschi A & Giacca M (1999) Replication origins of mammalian chromosomes: the happy few. Front Biosci 4: d859-d868.

Tom S, Henricksen LA & Bambara RA (2000) Mechanism whereby proliferating cell nuclear antigen stimulates flap endonuclease 1. J Biol Chem 275: 10498-10505.

Tong AH, Lesage G, Bader GD, Ding H, Xu H, Xin X, Young J, Berriz GF, Brost RL, Chang M, Chen Y, Cheng X, Chua G, Friesen H, Goldberg DS, Haynes J, Humphries C, He G, Hussein S, Ke L, Krogan N, Li Z, Levinson JN, Lu H, Menard P, Munyana C, Parsons AB, Ryan O, Tonikian R, Roberts T, Sdicu AM, Shapiro J, Sheikh B, Suter B, Wong SL, Zhang LV, Zhu H, Burd CG, Munro S, Sander C, Rine J, Greenblatt J, Peter M, Bretscher A, Bell G, Roth FP, Brown GW, Andrews B, Bussey H & Boone C (2004) Global mapping of the yeast genetic interaction network. Science 303: 808-813.

Tornaletti S (2005) Transcription arrest at DNA damage sites. Mutat Res 577: 131-145. Tornaletti S, Reines D & Hanawalt PC (1999) Structural characterization of RNA polymerase II

complexes arrested by a cyclobutane pyrimidine dimer in the transcribed strand of template DNA. J Biol Chem 274: 24124-24130.

Traven A & Heierhorst J (2005) SQ/TQ cluster domains: concentrated ATM/ATR kinase phosphorylation site regions in DNA-damage-response proteins. Bioessays 27: 397-407.

Tsubota T, Maki S, Kubota H, Sugino A & Maki H (2005) Double-stranded DNA binding properties of Saccharomyces cerevisiae DNA polymerase ε and of the Dpb3p-Dpb4p subassembly. Genes Cells 8: 873-888.

Tsurimoto T & Stillman B (1991a) Replication factors required for SV40 DNA replication in vitro. I DNA structure-specific recognition of a primer-template junction by eukaryotic DNA polymerases and their accessory proteins. J Biol Chem 266: 1950-1960.

Tsurimoto T & Stillman B (1991b) Replication factors required for SV40 DNA replication in vitro. II. Switching of DNA polymerase α and δ during initiation of leading and lagging strand synthesis. J Biol Chem 266: 1961-1968.

Tuusa J (2001) Human DNA polymerase ε. Expression, phosphorylation and protein-protein interactions. Acta Univ Oul D661.

Tuusa J, Uitto L & Syväoja JE (1995) Human DNA polymerase epsilon is expressed during cell proliferation in a manner characteristic of replicative DNA polymerases. Nucleic Acids Res 23: 2178-2183.

Uchiyama M, Griffiths D, Arai K & Masai H (2001) Essential role of Sna41/Cdc45 in loading of DNA polymerase α onto minichromosome maintenance proteins in fission yeast. J Biol Chem 276: 26189-26196.

van Gool AJ, Citterio E, Rademakers S, van Os R, Vermeulen W, Constantinou A, Egly JM, Bootsma D & Hoeijmakers JH (1997) The Cockayne syndrome B protein, involved in transcription-coupled DNA repair, resides in an RNA polymerase II-containing complex. EMBO J 16: 5955–5965.

van Hatten RA, Tutter AV, Holway AH, Khederian AM, Walter JC & Michael WM (2002) The Xenopus Xmus101 protein is required for the recruitment of Cdc45 to origins of DNA replication. J Cell Biol 159: 541-547.

van Oosterwijk MF, Versteeg A, Filon R, van Zeeland AA & Mullenders LHF (1996) The sensitivity of Cockayne’s syndrome cells to DNA-damaging agents is not due to defective transcription-coupled repair of active genes. Mol Cell Biol 16: 4436-4444.

Vas A, Moc W & Leatherwood J (2001) Control of DNA rereplication via Cdc2 phosphorylation sites in the origin recognition complex. Mol Cell Biol 21: 5767–5777.

72

Vindelöv LL, Christensen IJ & Nissen NI (1983) Standardization of high-resolution flow cytometric DNA analysis by the simultaneous use of chicken and trout red blood cells as internal reference standards. Cytometry 3: 323-327.

Volkening M & Hoffmann I (2005) Involvement of human MCM8 in prereplication complex assembly by recruiting hcdc6 to chromatin. Mol Cell Biol 25: 1560-1568.

Volker M, Mone MJ, Karmakar P, van Hoffen A, Schul W, Vermeulen W, Hoeijmakers JH, van Driel R, van Zeeland AA & Mullenders LH (2001) Sequential assembly of the nucleotide excision repair factors in vivo. Mol. Cell 8: 213-224.

Waga S & Stillman B (1994) Anatomy of a DNA replication fork revealed by reconstitution of SV40 DNA replication in vitro. Nature 369: 207-212.

Wang Y, Cortez D, Yazdi P, Neff N, Elledge SJ & Qin J (2000) Genes Dev 14: 927-939. Wang H & Elledge SJ (1999) DRC1, DNA replication and checkpoint protein 1, functions with

DPB11 to control DNA replication and the S-phase checkpoint in Saccharomyces cerevisiae. Proc Natl Acad Sci USA 96: 3824-3829.

Warner BJ, Blain SW, Seoane J & Massagué J (1999) Myc downregulation by transforming growth factor beta required for activation of the p15(Ink4b) G(1) arrest pathway. Mol Cell Biol 19: 5913–5922.

Weisshart K, Pestryakov P, Smith RW, Hartmann H, Kremmer E, Lavrik O & Nasheuer H-P (2004) Coordinated regulation of replication protein A activities by its subunits p14 and p32. J Biol Chem 279:35368-35376.

Westover KD, Bushnell DA, Kornberg RD (2004) Structural basis of transcription: nucleotide selection by rotation in the RNA polymerase II active center. Cell 119: 481-489.

Whitehouse I, Flaus A, Cairns BR, White MF, Workman JL & Owen-Hughes T (1999) Nucleosome mobilization catalysed by the yeast SWI/SNF complex. Nature 400: 784-787.

Wilson DM 3rd, & Thompson LH (1997) Life without DNA repair. Proc Natl Acad Sci USA 94: 12754-12757.

Winkler GS, Araujo SJ, Fiedler U, Vermeulen W, Coin F, Egly JM, Hoeijmakers JH, Wood RD, Timmers HT & Weeda G (2000) TFIIH with inactive XPD helicase functions in transcription initiation but is defective in DNA repair. J Biol Chem 275: 4258-4266.

Wohlschlegel JA, Dwyer BT, Dhar SK, Cvetic C, Walter JC & Dutta A (2000) Inhibition of eukaryotic DNA replication by geminin binding to Cdt1. Science 290: 2309-2312.

Wood RD, Mitchell M, Lindahl T (2005) Human DNA repair genes. Mutat Res 577: 275-283. Wu X, Braithwaite E & Wang Z (1999) DNA ligation during excision repair in yeast cell-free

extracts is specifically catalyzed by the CDC9 gene product. Biochemistry 38: 2628-2635. Wyrick JJ, Aparicio JG, Chen T, Barnett JD, Jennings EG, Young RA, Bell SP & Aparicio OM

(2001) Genome-wide distribution of ORC and MCM proteins in S. cerevisiae: high-resolution mapping of replication origins. Science 294: 2357-2360.

Yamane K, Kawabata M & Tsuruo T (1997) A DNA-topoisomerase-II-binding protein with eight repeating regions similar to DNA-repair enzymes and to a cell-cycle regulator. Eur J Biochem 250: 794-799.

Yamane K & Tsuruo T (1999) Conserved BRCT regions of TopBP1 and of the tumor suppressor BRCA1 bind strand breaks and termini of DNA. Oncogene 18: 5194-5203.

Yamamoto RR, Axton JM, Yamamoto YY, Saunders RDC, Glover DM & Henderson DS (2000) The Drosophila mus101 Gene, Which Links DNA Repair, Replication and Condensation of Heterochromatin in Mitosis, Encodes a Protein With Seven BRCA1 C-Terminus Domains Genetics 156: 711-721.

Yan S, Lindsay HD & Michael WM (2006) Direct requirement for Xmus101 in ATR-mediated phosphorylation of Claspin bound Chk1 during checkpoint signaling. J Cell Biol 173: 181-186.

73

Yankulov K, Todorov I, Romanowski P, Ligatalosi D, Cilli K, McCracken S, Laskey R & Bentley DL (1999) MCM proteins are associated with RNA polymerase II holoenzyme. Mol Cell Biol 19: 6154-6163.

Zeng L, Hu Y & Li B (2005) Identification of TopBP1 as a c-Abl-interacting protein and a repressor for c-Abl expression. J Biol Chem 280: 29374-29380.

Zerbe LK & Kuchta RD (2002) The p58 subunit of human DNA primase is important for primer initiation, elongation, and counting. Biochemistry 41: 4891-4900.

Zhang S-J, Zeng X-R, Zhang P, Toomey NL, Chuang R-Y, Chang L-S & Lee MYWT (1995) A conserved region in the amino terminus of DNA polymerase δ is involved in proliferating nuclear antigen binding. J Biol Chem 270: 7988-7992.

Zlotkin T, Kaufmann G, Jiang Y, Lee MYWT, Uitto L, Syväoja J, Dornreiter I, Fanning E & Nethanel T (1996) DNA polymerase ε may be dispensable for SV40- but not cellular-DNA replication. EMBO J 15: 2298-2305.

Zou L & Stillman B (2000) Assembly of a complex containing Cdc45p, replication protein A, and Mcm2p at replication origins controlled by S-phase cyclin-dependent kinases and Cdc7p-Dbf4p kinase. Mol Cell Biol 20: 3086-3096.

Zuo S, Gibbs E, Kelman Z, Wang TS-F, O’Donnell M, MacNeill SA & Hurwitz J (1997) DNA polymerase δ isolated from Schizoschaccharomyces pombe contains five subunits. Proc Natl Acad Sci USA 94: 11244-11249.

Original articles

I Mäkiniemi M, Hillukkala T, Tuusa J, Reini K, Vaara M, Huang D, Pospiech H, Majuri I, Westerling T, Mäkela TP & Syväoja JE (2001) BRCT domain-containing protein TopBP1 functions in DNA replication and damage response. J Biol Chem 276: 30399-30406.

II Herold S, Wanzel M, Beuger V, Frohme C, Beul D, Hillukkala T, Syväoja J, Saluz HP, Haenel F & Eilers M (2002) Negative regulation of the mammalian UV response by Myc through association with Miz-1. Mol Cell 10: 509-521.

III Rytkönen AK*, Hillukkala T*, Vaara M, Sokka M, Jokela M, Sormunen R, Nasheuer H-P, Nethanel T, Kaufmann G, Pospiech H & Syväoja JE (2006) DNA polymerase ε associates with the elongating form of RNA polymerase II and nascent transcripts. FEBS J, in press.

IV Hillukkala T, Vaara M, Koistinen H, Sokka M, Nasheuer H-P, Syväoja JE & Pospiech H (2006) Human Cdc45 forms distinct complexes with DNA polymerases δ and ε during nuclear DNA replication. Manuscript

* These authors contributed equally to this work.

Permission to reprint the original articles has been obtained from the following copyright owners:

I ASBMB Journals

II Elsevier

III Blackwell Publishing

Original publications are not included in the electronic version of the dissertation.


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