HUMAN DNA POLYMERASE ε JUSSI Expression, phosphorylation and -protein interactions TUUSA

Biocenter Oulu, University of Oulu Department of Biochemistry, University of Oulu

OULU 2001 JUSSI TUUSA

HUMAN DNA POLYMERASE ε Expression, phosphorylation and protein-protein interactions

Academic Dissertation to be presented with the assent of the Faculty of Science, University of Oulu, for public discussion in Kajaaninsali (Auditorium L6), Linnanmaa, on December 17th, 2001, at 2 p.m.

OULUN YLIOPISTO, OULU 2001 Copyright © 2001 University of Oulu, 2001

Manuscript received 15 November 2001 Manuscript accepted 27 November 2001

Communicated by Professor Kalle Saksela Docent Hans Spelbrink

ISBN 951-42-6581-5 (URL: http://herkules.oulu.fi/isbn9514265815/)

ALSO AVAILABLE IN PRINTED FORMAT ISBN 951-42-6580-7 ISSN 0355-3221 (URL: http://herkules.oulu.fi/issn03553221/)

OULU UNIVERSITY PRESS OULU 2001 Tuusa, Jussi, Human DNA polymerase ε Expression, phosphorylation and protein- protein interactions Biocenter Oulu, University of Oulu, P.O.Box 5000, FIN-90014 University of Oulu, Finland, Department of Biochemistry, University of Oulu, P.O.Box 3000, FIN-90014 University of Oulu, Finland 2001 Oulu, Finland (Manuscript received 15 November 2001)

Abstract DNA replication is a process in which a cell duplicates its genome before cell division, and must proceed accurately and in organized manner to guarantee maintenance of the integrity of the genetic information. DNA polymerases are enzymes that catalyse the synthesis of the new DNA strand by utilizing the parental strand as a template. In addition to chromosomal replication, DNA synthesis and therefore DNA polymerases are also needed in other processes like DNA repair and DNA recombination. The DNA polymerase is an essential DNA polymerase in eukaryotes and is required for chromosomal DNA replication. It has also been implicated in DNA repair, recombination, and in transcriptional and cell cycle control. The regulation of the human enzyme was explored by analysing its expression, phosphorylation and protein-protein interactions. Expression of both the A and B subunits of the human DNA polymerase ε was strongly growth- regulated. After serum-stimulation of quiescent fibroblasts, the steady-state mRNA levels were up- regulated at least 5-fold. In actively cycling cells, however, the steady-state mRNA and protein levels fluctuated less than 2-fold, being highest in G1/S phase. The promoter of the B subunit was analysed in detail. The 75 bp core promoter was essentially dependent on the Sp1 transcription factor. Furthermore, mitogenic control of the promoter required an intact E2F binding element, and binding of E2F2, E2F4 and p107 was demonstrated in vitro. A down-regulation element, located immediately downstream from the core promoter, bound E2F1, NF-1 and pRb transcription factors. A model of the promoter function is presented. Topoisomerase IIβ binding protein 1 (TopBP1) was found to be associated with human DNA polymerase ε. TopBP1 contains eight BRCT domains and is homologous to Saccharomyces cerevisiae Dpb11, Schizosaccharomyces pombe Cut5, Drosophila melanogaster Mus101 and the human Breast Cancer susceptibility protein 1 (BRCA1). TopBP1 is a phosphoprotein, whose expression is induced at the G1/S border and is required for chromosomal DNA replication. It co- localizes in S phase with BRCA1 into discrete foci, which do not represent sites of ongoing DNA replication. However, if DNA is damaged or replication is blocked in S phase cells, TopBP1 and BRCA1 re-localize into proliferating cell nuclear antigen (PCNA) containing foci that represent stalled replication forks. Finally, phosphorylation of DNA polymerase ε was described and at least three immunologically distinct and differentially phosphorylated forms were shown to exist. Phosphorylation is on serine and threonine residues and shows a cell cycle dependent fluctuation, but is not affected by DNA damage or by inhibition of DNA replication. BRCA1 co-immunoprecipitates with a hypophosphorylated form of DNA polymerase ε. In contrast, TopBP1 was shown to be associated with a hyperphosphorylated form.

Keywords: , DNA replication, DNA-directed DNA polymerase, phospho- rylation, protein-protein interactions Acknowledgements

This research was carried out at the Biocenter Oulu and the Department of Biochemistry at the University of Oulu. I wish to thank my supervisor, professor Juhani Syväoja for providing the excellent opportunity for learning science. I would also like to thank professors Karl Tryggvason, Kalervo Hiltunen and Raili Myllylä for professional leadership of the Department of Biochemistry, excellent research facilities and the creation of a supportive atmosphere. I am grateful to Professor Kalle Saksela and Docent Hans Spelbrink for their valuable criticism of the manuscript of this thesis, and to Dr. Deborah Kaska and Dr. Lloyd Ruddock for rapid and accurate revision of the language. I am indebted to all former and present members of the JS-team. Especially, my warmest thanks belong to my lab mates Anna Rytkönen and Helmut Pospiech with whom I have shared so many “brilliant-breakthrough-ideas” as well as the development of the “Pospiech-Rytkönen-Tuusa modified Burgers-Campbell-Falaschi double-fork-replisome model”. I wish to express my thanks to all my co-authors. In particular, I would like to thank Docent Kari Poikonen for help in flow-cytometry and Professor Sven Skog for guidance in centrifugal elutriation. I would also like to thank all my colleagues and staff at the Department of Biochemistry for continuous help and interesting discussions. Special thanks belong to Jaakko Keskitalo, Maila Konttila and Eeva-Liisa Stefanius. I am deeply grateful for my wife Soile, my parents, my sister and brother and my family-in-law, Oulun OPKO, LUONA-piiri and all my friends for belonging to my life and more, for love and encouragement during these years, when my time was prioritised for cell cultures instead of them. Finally, in the end of the day, let my praise and glory be on Our Creator, Lord Jesus, by whom all things were created, things in heaven and on earth. With the words of Psalm writer: “I praise you because I am fearfully and wonderfully made; your works are wonderful, I know that full well.”

Oulu, November 2001 Jussi Tuusa

Abbreviations aa amino acid ARS autonomously replicating sequence ATM ataxia telangectesia mutated ATR ATM and Rad3 related bp BRCA1 breast cancer susceptibility gene/protein 1 BRCT BRCA1 C-terminus Cdc cell division control Cdk cyclin-dependent kinase DNA deoxyribonucleic acid dsDNA double-stranded DNA EMSA electrophoretic mobility shift assay kDa kilodalton Mcm minichromosome maintenance protein nt nucleotide ORC origin recognition complex PAGE polyacrylamide gel electrophoresis PBS phosphate buffered saline PCNA proliferating cell nuclear antigen PIP PCNA interacting protein pol DNA polymerase Rad radiation sensitive RC replication complex RFC RNA ribonucleic acid RPA replication protein A ssDNA single-stranded DNA SV40 simian virus 40 T-ag large T-antigen TLC thin-layer chromatography TLE thin-layer electrophoresis TopBP1 topoisomerase IIβ binding protein 1 UV ultraviolet WRN Werner syndrome protein List of original articles

In addition to some unpublished material, this thesis is based on the following articles, which are referred to in the text by their Roman numerals:

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

II 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.

III 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.

IV Tuusa J, Uitto L, Jokela M, Pospiech H & Syväoja JE (2001) Immunologically distinct forms of human DNA polymerase ε are differentially phosphorylated: Implications for the interaction with BRCA1 and TopBP1. Manuscript submitted.

Contents

Acknowledgements Abbreviations List of original articles Contents 1 Introduction...... 13 2 Review of literature...... 14 2.1 Concept of DNA replication...... 14 2.1.1 Structure of DNA and principles of polymerisation...... 14 2.1.2 DNA replication in prokaryotes ...... 16 2.1.3 DNA replication in eukaryotes...... 17 2.1.3.1 Simian virus 40 replication in vitro...... 18 2.1.3.2 Chromosomal DNA replication in yeast ...... 20 2.1.3.3 Regulation of DNA replication ...... 20 2.1.3.4 Replicative DNA polymerases ...... 23 2.1.3.5 S phase checkpoints ...... 23 2.1.4 DNA replication in Archaea ...... 27 2.2 Eukaryotic DNA polymerases...... 27 2.2.1 Classification, structure and function of DNA polymerases ...... 27 2.2.2 DNA polymerase alpha- ...... 30 2.2.3 DNA polymerase delta ...... 32 2.2.4 DNA polymerase epsilon ...... 34 2.3 associated with DNA replication and S phase checkpoint ...... 38 2.3.1 Mcm2, Mcm3, Mcm4, Mcm5, Mcm6 and Mcm7...... 38 2.3.2 Dpb11 and homologous proteins...... 39 2.3.3 Cdc45 ...... 39 2.3.4 Replication protein A ...... 40 2.3.5 Proliferating cell nuclear antigen ...... 40 2.3.6 Replication factor C ...... 42 2.3.7 Rad1-Rad9-Hus1-Rad17...... 43 2.3.8 BRCA1...... 43 2.3.9 E2F and pocket protein families...... 44 3 Aims of the present work ...... 46 4 Materials and methods ...... 47 4.1 Ribonuclease protection assay (I, II, III)...... 47 4.2 Immunoprecipitation and Western-analysis (I, III, IV) ...... 47 4.3 Serum stimulation of quiescent cells (I, II, III) ...... 48 4.4 Cell cycle specific synchronization and fractionation (I, II, III, IV)...... 48 4.5 Promoter activity analysis (II)...... 49 4.6 Electrophoretic mobility shift assay (EMSA) (II)...... 49 4.7 Deoxyribonuclease I footprinting analysis (II) ...... 49 4.8 DNA replication assay in isolated nuclei (III)...... 50 4.9 Immunostaining (III)...... 50 4.10 Ecdysone-inducible expression system (III, IV) ...... 50 4.11 Site directed mutagenesis...... 50 4.12 Metabolic labelling with 32P (III, IV)...... 51 4.13 Phosphoamino acid analysis and phosphopeptide mapping (III, IV)...... 51 4.14 Cross-linking with formaldehyde (IV)...... 52 5 Results...... 53 5.1 The expression of the human DNA polymerase ε is typical for replicative DNA polymerases (I, II)...... 53 5.2 The gene for human DNA polymerase ε B subunit is regulated by E2F and Sp1 transcription factors (II) ...... 54 5.3 DNA polymerase ε interacts with BRCA1 and TopBP1 (III, IV) ...... 54 5.4 TopBP1 is a phosphoprotein with eight BRCT-domains and is expressed in a proliferation dependent manner (III) ...... 55 5.5 TopBP1 is required for chromosomal DNA replication (III) ...... 56 5.6 TopBP1 and BRCA1 co-localize in S phase and are re-localized after replication block or DNA damage into the PCNA containing foci (III)...... 56 5.7 TopBP1 interacts physically with checkpoint protein Rad9 (III) ...... 57 5.8 DNA polymerase ε has at least three immunologically separable forms with distinct phosphorylation and different interacting proteins (IV) ...... 57 6 Discussion...... 60 6.1 The proliferation dependent expression of pol ε is similar to other replicative DNA polymerases with some unique features (I, II, IV)...... 60 6.2 TopBP1 is a pol ε associated protein with functions in DNA replication and damage response (III)...... 62 6.3 Pol ε has at least three differently phosphorylated forms with specific interactions with BRCA1 and TopBP1...... 63 7 Conclusions...... 65 8 References...... 66

1 Introduction

In eukaryotic cells four essential nuclear DNA polymerases (pols) are involved in chromosomal DNA replication, pol α, pol δ, pol ε and pol σ. Pol α has a well established role in the initiation of chromosomal DNA replication as well as in priming of the Okazaki fragments of the lagging strand. Pol σ, which was only recently discovered and is still poorly characterized, is needed in the coupling of DNA replication with the cohesion of sister chromatids. Both pol δ and pol ε are required during replicative DNA synthesis for the elongation of DNA strands as well as for the maturation of Okazaki fragments. However, their exact functions are not fully understood. In addition to DNA polymerases, a number of other proteins are required as structural or regulatory factors to guarantee accurate copying of the genome in a manner that is temporally and spatially coordinated with other cell cycle events. Furthermore, checkpoint mechanisms exist that prevent or slow down the DNA replication during DNA damage, prevent premature mitosis and allow time for repair of damage and for recovery of stalled replication forks. The functions of proteins can be regulated via changes in expression, post- translational modifications and interactions with other proteins. In this study the expression patterns, phosphorylation and protein-protein interactions of human pol ε were examined. Pol ε was found to be expressed similarly to other replicative pols with a strong response to mitogenic stimulation when quiescent cells were induced to proliferate, but with modest fluctuation during the mitotic cell cycle. Secondly, Topoisomerase ΙΙβ binding protein 1 (TopBP1), a homologue of Saccharomyces cerevisiae Dpb11 was characterized as a pol ε associated protein and was shown to function in DNA replication and the DNA damage response. Thirdly, we demonstrated that the Breast Cancer Susceptibility protein 1 (BRCA1) interacts physically with pol ε. Fourthly, pol ε was shown to be phosphorylated in vivo and at least three immunologically distinct and differentially phosphorylated forms were identified. Finally, a correlation was observed between differential phosphorylation and interaction with TopBP1 and BRCA1. 2 Review of literature

2.1 Concept of DNA replication

2.1.1 Structure of DNA and principles of polymerisation

In the history of mankind, the nature of genetic information remained unknown until our time. In 1944, Avery and co-workers showed for the first time that deoxyribonucleic acid, hereafter DNA, was a genetic substance (Avery et al. 1944). By the end of the 20th century, the blueprint of life, that is, the genome of more than 20 prokaryotic and 2 eukaryotic species were decoded and the world was anxiously waiting the announcement that sequencing of the was finished. As the complete determination of the exact order of the four bases of DNA, or guanine (G), cytosine (C), adenine (A) and thymine (T) in genomes has been a landmark in history of science, so was the discovery of the structure of DNA. In 1953 James D. Watson and Francis H. C. Crick published a model for the three-dimensional structure of DNA based on the X-ray data from work done by Rosalind Franklin (Watson & Crick 1953, Wilkins et al. 1953, Franklin & Gosling 1953). DNA is a double helix consisting of two anti-parallel strands. The strands are formed of alternating deoxyribose sugars and phosphates linked by phosphodiester bonds, and of purine (A, G) or pyrimidine (C, T) bases that are linked to the sugar moieties by N-glycosidic bonds. The two strands are bound together via stable hydrogen bonds between the bases: two hydrogen bonds between A and T and three bonds between G and C. Importantly, a complementary sequence of bases in anti-parallel strands is a prerequisite for a stable conformation that is further stabilized by the stacking of hydrophobic bases. Hardly any discovery of macromolecular structure has so directly revealed the function as did the double helix of DNA. The nature of genetic information itself had to wait for the cracking of the genetic code in the ensuing decades, but the other important character for the carrier of heredity, the ability to copy itself, that is, to replicate, was apparent: One strand of DNA could be copied from the complementary information of 15 the other strand. An obvious model, semi conservative DNA replication, was experimentally confirmed in 1957 by Meselson & Stahl (Meselson & Stahl 1958). Two daughter molecules both inherit one strand from the parental DNA, while the other strand is synthesized de novo. Another landmark in DNA replication research was the purification of DNA polymerase by Arthur Kornberg and co-workers in 1955 (Kornberg et al. 1955). Besides being invaluable reagents in molecular biology such as in DNA amplification and sequencing, DNA polymerases have been central objects for understanding DNA replication, repair and recombination. Indisputably, many fundamental and universal features of DNA replication reflect the properties of DNA polymerases. The two strands of the double helix must be separated, because DNA polymerase utilizes ssDNA as a template. DNA polymerases cannot start DNA synthesis de novo, but need a primer with an accessible hydroxyl group. The synthesis of a new DNA strand elongates in a 5' to 3' direction, when pol adds activated monomeric substrates, deoxynucleoside 5'- triphosphates, one by one, to the primer end. The way in which DNA polymerase is bound to the template-primer affects on the affinity for the incoming deoxynucleotide such that only a deoxynucleotide complementary to the opposite template base has high probability to be a target for a nucleophilic attack by the hydroxyl group of the primer terminus (Kornberg & Baker 1992, pp.108-109, 113-140). Therefore, the structure of DNA polymerase establishes a kinetic control mechanism that increases the likelihood for incorporation of the correct deoxynucleotide. This intrinsic property of DNA polymerase, together with the often associated 3'->5' exonuclease activity, provides the fidelity of replication that is essential for maintenance of genetic information. A direct consequence of the polarity of polymerisation is semi discontinuous replication. One strand, also called the leading strand, is synthesized continuously, whereas the other, lagging strand, is synthesized discontinuously in short patches, also called Okazaki fragments, that are later ligated together (Kurosawa et al. 1975). The schematic outline of the semi conservative, semi discontinuous DNA replication is presented in Figure 1. The replicative DNA polymerases are usually part of large multienzyme complexes that may contain primase activity, nucleases, accessory factors that provide processivity, help in loading on the primer terminus, stabilize ssDNA or couple the DNA synthesis into the context of cell cycle regulation. Importantly, these replication complexes do not start DNA synthesis randomly, but are assembled onto the specific origins of DNA replication by initiation factors. The initiation of cellular DNA replication is universally regulated by the interplay of the DNA containing origin sequence ("replicator") and origin recognition factor(s) ("initiator"). The initiator catalyses the opening of the DNA duplex at the origin and recruits the assembly of the DNA replication complex. DNA replication may proceed unidirectionally, but more often proceeds bi-directionally, when two replication forks are created and move in opposite directions.

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Fig. 1. Schematic outline of cellular DNA replication. DNA is duplicated in a semi- conservative, semi discontinuous process, where new strands are synthesized by one-by-one addition of nucleotide pre-cursors.

2.1.2 DNA replication in prokaryotes

Prokaryotes have usually a single circular . The size of the genome, for example 4,7 Mb in Escherichia coli, makes the direct analysis and manipulation of chromosomal DNA replication difficult. Therefore, it has been helpful to use viruses that are able to infect and replicate in bacterial cells. These viruses, or bacteriophages, have relatively small genomes from 5.4 kb of φ(X)174 phage to 166 kb of T4 phage. φ(X)174 provides an example of viruses that largely utilize the host's replication machinery and need only a few viral encoded proteins for DNA replication, whereas T4 represents a system relying exclusively on gene products of the bacteriophage's own genome. This has enabled the examination of DNA replication in vitro and, together with the versatile genetics of E. coli, has provided us with a relatively good understanding of DNA replication in this bacterium as well as in prokaryotes in general. In E. coli, DNA replication begins from a single origin, called oriC, and bi-directional replication continues until the two replication forks moving in opposite directions meet (Bird et al. 1972). The 245 bp minimal origin in oriC contains four 9-mers acting as a recognition site for the DnaA initiator protein, three AT-rich 13-mers that facilitate the melting of the DNA duplex and association of the DnaB helicase and 14 GATC sequences that are targets for Dam methylase (Oka et al. 1984, Kornberg & Baker 1992, 17 pp.521-533). DnaA initiation factor, a 52 kDa protein, accumulates in a co-operative manner at oriC during the growth phase of the bacterial cell. In addition to recognition of the origin for DNA replication, DnaA promotes the opening of double helical DNA and loads DnaB helicase with the help of the DnaC protein onto the origin (Fuller et al. 1984, Bramhill D & Kornberg A 1988). A hexamer of DnaB is a 5’->3’ helicase that opens dsDNA ahead of a replication fork. After DnaB has created ssDNA, DnaG primase readily synthesizes an RNA primer, which becomes elongated by DNA polymerase III holoenzyme (pol III). The re-initiation of DNA replication during the ongoing cell cycle is prevented by the hemimethylated state of newly synthesized DNA that also is sequestered by the cell membrane in a SeqA dependent manner. SeqA is a protein that binds tightly to the hemimethylated DNA (Crooke 1995). Pol III holoenzyme is a 900 kDa protein complex containing at least 20 polypeptides. It consist of a dimer of a catalytic core complex with DNA polymerase and 3’->5’ exonuclease activities, two β- clamps that function as processivity factors, a γ-complex called a clamp loader and a τ- complex with an auxiliary function (Kim et al. 1996).

2.1.3 DNA replication in eukaryotes

Eukaryotic genomes are much larger and more complex than genomes in prokaryotes. Instead of single circular molecules, they are organized and segregated as linear . The chromosomes with species-specific number and ploidy are compartmentalized inside the nucleus and associated with the nuclear matrix. In eukaryotes nuclear DNA exists in a form of nucleoprotein complex, called chromatin. Chromatin is formed when histone proteins are assembled on replicated DNA to form a nucleosome structure. This process includes the activities of several assembly factors, and mature chromatin contains a variety of structural proteins in addition to histones (Krude et al. 1999). Mitochondrial DNA exists in several hundred of copies in a circular form. Its replication has been reviewed (Shadel & Clayton 1997) and is not discussed here. The study of chromosomal DNA replication is very difficult because of the complexity of eukaryotic genomes. Recently, a few cell free systems have been introduced and chromosomal DNA replication can be examined in vitro in Xenopus or Drosophila egg extracts or in human permeable nuclei (Blow & Laskey 1986, Crevel & Cotterill 1991, Krude et al. 1997). However, most of our current understanding is based either on the use of animal viruses or on yeast genetics. The viral replication systems can be divided in two categories according to the need for host cell replication proteins. The papovaviruses, like Simian virus 40, polyomaviruses and papillomaviruses, rely heavily on the replication machinery of the host cell, whereas adenoviruses and herpesviruses provide examples of systems where all or most of the replication proteins are encoded by viral genomes. While the former give direct information about the cellular machinery, the latter systems are useful in understanding the minimal requirements for DNA replication and the roles and functions of the viral replication proteins and their cellular orthologues. 18

2.1.3.1 Simian virus 40 replication in vitro

The Simian virus 40 (SV40) cell-free replication system was used for the first time in 1984 by Li and Kelly (Li & Kelly 1984). SV40 belongs to the papovaviruses and its 5243 bp minichromosome type genome is replicated in a bi-directional semi discontinuous manner in infected cells (Kornberg & Baker 1992). Reconstitution of SV40 replication in vitro with purified replication factors has shown that one viral and nine cellular proteins are needed for complete replication (Waga & Stillman 1994a,b). The only viral protein required, the large T-antigen (T-ag), is a multifunctional 708 aa phosphoprotein with ssDNA and sequence specific dsDNA binding, ATPase and helicase activities (Fanning & Knippers 1992). First, T-ag recognizes and binds to the origin of replication as a double hexamer. This binding is negatively and positively regulated by phosphorylation in vivo (Fanning 1994), and indeed, SV40 DNA replication in vivo is tightly restricted to the S phase. Secondly, T-ag catalyses the melting of the DNA duplex. Thirdly, it recruits the replication protein A (RPA) and DNA polymerase alpha-primase (pol α) onto the origin. RPA is a eukaryotic single strand binding protein that protects and stabilizes ssDNA and promotes DNA unwinding. Fourthly, T-ag acts as a replicative helicase that unwinds DNA in front of the replication fork. Pol α primes the leading strand as well as every Okazaki fragment of the lagging strand by synthesizing a short RNA-DNA primer that consists of a ca. 10 nt long RNA with a 20-30 nt long DNA extension (Hay et al. 1984, Nethanel et al. 1988, Bullock et al. 1991, Murakami et al. 1992, Eki et al. 1992). The polymerase switch occurs both on the leading and lagging strands. RNA-DNA primers synthesized by pol α are elongated to long nascent products on the leading strand template and to ca. 300 bp Okazaki- fragments on lagging strand template by pol δ (Tsurimoto & Stillman 1991). The loading of pol δ requires the presence of a primer recognition complex that consists of proliferating cell nuclear antigen (PCNA), replication factor C (RFC) and ATP (Tsurimoto & Stillman 1991). PCNA is a processivity factor for pols δ and ε and forms “a sliding clamp” around the DNA. PCNA cannot assemble on covalently closed circular DNA itself, but needs a “clamp loader” – RFC (Yoder & Burgers 1991, Burgers & Yoder 1993). The primer recognition complex competes with pol α on binding to the primer end and replaces pol α in an ATP dependent manner (Eki et al. 1992, Tsurimoto & Stillman 1991). This reaction is mediated by RPA that associates with RFC and releases pol α (Yuzhakov et al. 1999). The maturation of Okazaki fragments into continuous DNA is catalysed in a series of reactions, where RNaseH first cleaves the RNA moiety endonucleolytically except for the last 3’ ribonucleotide that is removed by the exonuclease activity of Fen-1, also known as maturation factor 1 (MF1). The gap is filled by pol δ and finally the fragments are covalently linked by ligase I (Waga & Stillman 1994a,b). Pol ε is dispensable in SV40 DNA replication, but is able to partially replace pol δ (Lee et al. 1991, Eki et al. 1992). Finally, topoisomerases I and II are needed for removing the positive supercoiling in front of the replication fork and the negative supercoiling behind the fork as well as for the decatenation of daughter chromosomes. The basic concept of SV40 DNA replication is outlined in Figure 2.

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Fig. 2. SV40 DNA replication in vitro. (A) A model for the structure of the SV40 replication fork. (B) A model of the polymerase switch on lagging strand synthesis. Adapted from Burgers 1998, Waga & Stillman 1994.

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2.1.3.2 Chromosomal DNA replication in yeast

The yeast DNA replication assay in vitro was established only quite recently (Pasero et al. 1997) and therefore most of our knowledge about yeast DNA replication comes from genetic studies with budding yeast Saccharomyces cerevisiae and fission yeast Schizosaccharomyces pombe as well as from biochemical characterization of the individual replication proteins. Since the early mutation screens in these two organisms (Hartwell 1971, Hartwell et al. 1974, Nurse et al. 1976) a significant amount of information has accumulated about the roles and relationships of the replication-linked . In S. cerevisiae, DNA replication initiates from defined origin sequences, also called as autonomously replicating sequences (ARS) because of their ability to allow episomal replication of plasmid DNA. ARSs consist of an A element and a variable number of B elements (Newlon 1996). The A element is essential for origin function and it contains an ARS core consensus sequence (ACS) 5’-[A/T]TTTA[T/C][A/G]TTT[A/T]- 3’, an essential determinant for the binding of the origin recognition complex (ORC). The ORC consist of six subunits, recognizes the origin sequence in an ATP dependent manner and is necessary for the initiation of DNA replication (Newlon 1996, Marahrens & Stillman 1992, Bell & Stillman 1992). The B elements flanking the A element are less conserved in sequence, but functionally equivalent elements can be found from distinct origins (Maharens & Stillman 1992, Theis & Newlon 1994, Rao et al. 1994, Rashid et al. 1994). In ARS1, the best characterized ARS in S. cerevisiae, any two of the three B elements are sufficient for a functional origin (Maharens & Stillman 1992). The B1 element is next to the A element and partially protected by ORC binding (Bell & Stillman 1992). The B3 contains a binding site for a transcription factor Abf1, whose binding enhances origin utilization (Maharens & Stillman 1992). The function of the B2 element is not fully understood, but the loss of the ARS activity together with the increase in helical stability in B2 deletion mutants implies a role as a DNA unwinding element (DUE) (Natale et al. 1993). In the fission yeast S. pombe, the origins of DNA replication are much larger and more difficult to define. They contain AT-rich sequence blocks that are important for ARS activity, but lack any well-conserved consensus sequence (Dubey et al. 1996). In higher eukaryotes, ORC proteins are conserved, but initiation is largely determined in a sequence independent manner and DNA replication may start from alternative sites on the relative large initiation zone (reviewed by DePamphilis 1999).

2.1.3.3 Regulation of DNA replication

The regulation of eukaryotic DNA replication (reviewed by Kelly & Brown 2000, Nasmyth 1996) is primarily regulated at initiation and secondarily by S phase checkpoints that may also affect elongation and re-initiation. The DNA replication initiates from hundreds or thousands of temporally and spatially divergent origins per cell cycle and must be regulated so that the whole genome will replicate completely, but only once per cell cycle. The overall control of the cell cycle is based on the periodic fluctuation in the activities of the cyclin-dependent kinase (Cdk) that are regulated by 21 transient expression and degradation of the regulatory cyclin subunit, by a reversible phosphorylation of the catalytic subunit as well as by specific Cdk-inhibitor proteins (reviewed by Morgan 1997). With regard to DNA replication, the main targets of Cdk- activity are the protein complexes deposited on the origins of replication. In S. cerevisiae ORC binds to the origin throughout the cell cycle. However, the DNaseI footprint of the origin i.e. the region of origin DNA protected by associated proteins, changes during the cell cycle. The footprint represents ORC binding in the beginning of the mitosis, increases to its largest size in late mitosis and during the G1 phase and shrinks again in S phase (Diffley et al. 1994). The large footprint appearing in late mitosis and present during the G1 phase is due to the pre-replication complex (pre-RC) that assembles within the ORC. The Cdk-activity has a dual function on the pre-RC. On one hand G1/S-specific Cdks trigger the initiation of DNA replication by catalysing the conversion of the pre- RCs to active replication forks and ORC-containing post-replicative complexes (post- RC). On the other hand S/G2 phase specific Cdks maintain the replication forks and inhibit the re-initiation from post-RCs (Nasmyth 1996, Krude et al. 1997, Schwob & Nasmyth 1993, Strausfeld et al. 1996, Zou & Stillman 1998, Nougarede et al. 2000, Fotedar et al. 1996, Sever-Chroneos 2001, Hayles et al. 1994, Moreno & Nurse 1994, Dahmann et al. 1995, Tanaka et al. 1997, Hua et al. 1997, Nguyen et al. 2001). The sequence of the pre-RC assembly was first analysed in detail in Xenopus systems. Blow and Laskey observed that permeabilization of the nuclear membrane allowed re- replication (Blow & Laskey 1988). This indicates that a cytoplasmic factor, that is able to enter the nucleus during mitosis, “licenses” chromatin for replication. Later studies have shown that the licensing activity requires at least three components in addition to ORC: Cdc6, Cdt1 and Mcm proteins (Rowles et al. 1996, Cocker et al. 1996, Coleman et al. 1996, Madine et al. 1995, Chong et al. 1995, Thömmes et al. 1997, Maiorano et al. 2000, Nishitani et al. 2000, Tada et al. 2000). Cdc6 is an ATPase and shows homology to Orc proteins (Lee & Bell 2000). Mcm comes from “minichromosome maintenance” referring to the observation that corresponding genes are required for replication of episomal DNA in yeast cells. Mcms 2-7 are homologous to each other and can form several kind of heteromeric complexes, whereas another initiator protein, Mcm10 is unrelated (reviewed by Tye 1999). In animal cells, Cdc6 and Cdt1 enter the nucleus in mitosis and are required for the loading of Mcm proteins onto the origin and for the formation of the pre- RC. In G1/S phase, Cdc6 is phosphorylated by cyclin A/Cdk2 and exported from the nucleus, and Cdt1 is inhibited by the binding of the inhibitor protein, geminin (Petersen et al. 1999, Wohlschlegel et al. 2000). No new pre-RCs can form and replicated chromatin stays in a non-licensed stage until mitosis, when geminin is degraded and a new Cdc6 is synthesized and imported into the nucleus (McGarry & Kirschner 1998). In S. cerevisiae the nuclear membrane remains intact during mitosis and the import of licensing activity into nuclei requires active transport. Instead of transporting Cdc6 out of the nucleus, it is degraded after Mcm loading and new pre-RCs can form only when new Cdc6 is expressed during mitosis (Drury et al. 1997, Elsassar et al. 1999). Importantly, both in yeast and animal cells degradation of mitotic cyclins is a prerequisite for pre-RC assembly. The Mcm proteins assemble as a heterohexameric complex in the pre-RC (Prokhorova et al. 2000). They are needed for the loading of RPA, Cdc45, pol α and pol ε, as well as 22 for the accompanying origin unwinding (Tanaka & Nasmyth 1998, Walter & Newport 1998, Mimura & Takisawa 1998, Zou & Stillman 2000). The Mcm4-Mcm6-Mcm7 sub-

Fig. 3. A model for the assembly and activation of the eukaryotic pre-replication complex based on genetic data from yeast and immunodepletion studies with Xenopus (Nougarede et al. 2000, Coleman et al. 1996, Takisawa et al. 2000 and references therein.) complex harbours an intrinsic helicase activity and translocates with replication forks (Ishimi 1997, Aparacio et al. 1997). Despite the modest processivity of the Mcm4- Mcm6-Mcm7 helicase in vitro, it is a good candidate for a so far unidentified replicative helicase in eukaryotic cells (Labib et al. 2000). Several pre-RC proteins are substrates for Cdks including Orc1, Orc2 and Orc6, Cdc6, Mcm2 and Mcm4, RPA and pol α. This phosphorylation may either activate or inhibit initiation (Wolf et al. 1996, Finley et al. 1999, Nguyen et al. 2001, Elsassar et al. 1996, Fujita et al. 1998). Both the licensing by chromatin-bound Mcm proteins and the Cdk activity are required for the assembly of 23

RPA and Cdc45, whose binding to the chromatin is also mutually dependent (Zou & Stillman 2000). Finally, these factors load pols α and ε on the pre-RC in a process that at least in yeast is dependent on another protein, Dpb11 (Aparicio et al. 1999, Masumoto et al. 2000). In addition to Cdks, the activity of another protein kinase Cdc7-Dbf4 (or DDK) is essential for the loading of Cdc45, RPA and pols and for the final conversion of pre- RCs to replication forks (Zou & Stillman 2000). The Mcm2, Mcm3, Mcm4, Mcm6 and Mcm7, Cdc45 and pol α are substrates for Cdc7 (Lei et al. 1997, Weinreich & Stillman 1999, Nougarede et al. 2000). Interestingly Cdc7 itself is a substrate for several cyclin- Cdks and a corresponding phosphorylation is necessary for Cdc7 activity (Masai et al. 2000). The assembly of Cdc7-Dbf4 takes place after licensing and it interacts with ORC and Mcm proteins and the B2 element of ARS1 in yeast (Dowell et al. 1994, Hardy 1996, Lei et al. 1997). The observation that a mutation in Mcm5 can suppress a defective cdc7 mutant that is actually the last genetic marker in yeast before the onset of DNA replication, suggests that the kinase activity of Cdc7 causes conformational changes in the pre-RC leading to the formation of replication forks (Hereford & Hartwell 1974, Lei et al. 1997, Hardy et al. 1997). The sequence of pre-RC assembly and activation is presented in Figure 3.

2.1.3.4 Replicative DNA polymerases

Pols α and δ are required for the replication of SV40 DNA in vitro whereas pol ε is dispensable. However, genetic studies in yeast show that all three pols (α, δ, ε; reviewed in Hübscher et al. 2000) are essential for chromosomal DNA replication (Johnson et al. 1985, Sitney et al. 1989, Morrison et al.) Pols α and ε are loaded onto the early origins of replication at G1/S, onto the late origins at mid-S and associate with replication forks after initiation (Aparicio et al. 1999). Surprisingly, the polymerase domain of pol ε is dispensable in yeast but the C-terminal part of the catalytic subunit is essential for viability. This implies that some redundancy between pols exists and pol δ or another pol can replace pol ε in replicative DNA synthesis. Importantly, deletion mutants without the catalytic domain grow very slowly and the mutant yeast with full-length pol ε, but devoid of polymerase activity is non-viable (Kesti et al. 1999, Feng & D’Urso, 2001, Dua et al. 1999). The experiments with neutralizing antibodies and/or immunodepletion have shown that pols α and ε are also involved in chromosomal DNA replication in animal cells (Miller et al. 1985, Kaczmarek 1986, Pospiech et al. 1999, Waga et al. 2001). Furthermore, all pols α, δ and ε can be cross-linked to nascent DNA (Zlotkin et al. 1996).

2.1.3.5 S phase checkpoints

A checkpoint is a biochemical pathway that makes a cellular process dependent on another biochemically unrelated process: The cell cycle checkpoints ensure that the 24 previous stage of the cell cycle is fully completed before the entry into the next (Elledge 1996). The checkpoints that guarantee that the cell does not go to mitosis prematurely before the completion of DNA replication (S/M checkpoint, Murray 1992) and the checkpoints that slow down DNA synthesis to allow the repair of damaged DNA (intra-S checkpoint, Paulovich & Hartwell 1995) are collectively called S phase checkpoints. The S phase checkpoint may become activated due to a DNA synthesis block or because of DNA damage. Although the sensors of these two insults are separate, a common two- kinase signal transduction pathway is utilized (Figure 4). S. cerevisiae Mec1, S. pombe Rad3 and mammalian “Ataxia telangectasia mutated” (ATM) and “ATM and Rad3 related” (ATR) are serine/threonine protein kinases that belong to the family of phosphatidylinositol-3-kinases (reviewed in Durocher & Jackson 2001, Shiloh 2001). MEC1 and ATR are essential genes whereas Rad3 and ATM are dispensable for viability (Kato & Ogawa 1994, Brown & Baltimore 2000, Bentley et al. 1996, Barlow et al. 1996). Mammalian cells with mutated ATM are hypersensitive to ionising radiation, but have intact checkpoint function when exposed to UV or hydroxyurea (Siliciano et al. 1997, Brown et al. 1999, Tibbets et al. 2000). Hydroxyurea causes a replication block by inhibiting ribonucleotide reductase. Obviously, the checkpoint response after UV damage or replication block is channelled via ATR (Tibbets 2000). Two conserved protein kinases Chk1 and Chk2/Rad53/Cds1 are essential for many checkpoint functions. They act downstream from and are substrates of the Mec1/Rad3/ATM/ATR kinases. Chk1 and Chk2 have partially overlapping substrate spectra (Furnari et al. 1999) and are able to complement each other (Lindsay et al. 1998). They are needed for DNA damage or replication block induced transcription and for the S/M checkpoint in yeast and mammals (reviewed in Zhou & Elledge 2000). Furthermore, Rad53 in S. cerevisiae inhibits initiation from late origins via pol α and RPA as part of the intra-S checkpoint function (Marini et al. 1997, Longhese et al. 1996, Pelliciolli et al. 1999). The elegant work of Dimitrova & Gilbert suggests a similar mechanism in mammalian cells, which not only prevents the firing from late origins, but also maintains the integrity of the stalled replication complex (Dimitrova & Gilbert 2000). At least three different types of sensor systems seem to be involved in the detection of DNA damage and replication block. First, S. cerevisiae with mutations in replication protein genes, like POL2 (pol ε), SGS1, RFC5, DPB11 and SLD2, fail to restrain mitosis when cells are in S phase. In addition to their function at the S/M checkpoint, these genes participate in the intra-S checkpoint (Navas et al. 1995, Frei & Gasser 2000, Sugimoto et al. 1997, Araki et al. 1995, Wang & Elledge 1999). Secondly, in the G1 and in G2 phases, the checkpoint function is dependent on the Rad9 protein that also has a minor role in S phase (Navas et al. 1996). Both the replication machinery and Rad9 dependent systems use the same Mec1-Rad53 signal transduction pathway. Thirdly, S. pombe Rad26 associates with and becomes phosphorylated by Rad3, independent of the pathways described. Furthermore, Rad26 is needed for full Cds1 activation (Edwards et al. 1999). Importantly, the specific sensors and exact mechanism by which DNA damage or replication block is recognized have not been identified so far. Mec1, Rad3, ATM and ATR may have an intrinsic sensor function that is modulated by factors that are specific for certain types of damage or cell cycle stage. A PCNA-like clamp structure (Rad17- Ddc1-Mec3 in S. cerevisiae; Rad1-Rad9-Hus1 in S. pombe and in mammals) is loaded onto chromatin in a clamp-loader (Rad24 in S. cerevisiae; Rad17 in S. pombe and in 25 mammals) assisted process immediately after DNA damage. However, it is not likely to be the ultimate damage sensor, but rather a signal modifier and/or a processivity factor in translesion synthesis (Venclovas & Thelen 2000, Rauen et al. 2000). All four proteins, Rad1, Rad9, Hus1 and Rad17 are phosphorylated after DNA damage or replication block. Rad9 and Rad17 are substrates for ATM/ATR and in addition to these, Hus1 is a substrate for Rad3 in S. pombe (Chen et al. 2001, Bao et al. 2001, Kostrub et al. 1998). Significantly, the checkpoint function of pol ε is parallel to that of Rad24 in S. cerevisiae, and in S. pombe, the checkpoint function of Rad26 is independent of Rad1-Rad9-Hus1- Rad17 and of the replication machinery.

Fig. 4. The cell cycle checkpoint signal transduction pathways in eukaryotes. S. cerevisiae checkpoint proteins are indicated first and S. pombe or animal counterparts are shown in parentheses. Adapted from Zhou & Elledge 2000.

In animal cells BRCA1 is involved in the damage response during S phase. It has a role in transcriptional regulation, transcription-coupled repair of oxidative damage and homologous recombination mediated double-strand break repair (Chapman & Verma 1996, Scully et al. 1997, Anderson et al. 1998, Yarden & Brody 1999, Pao et al. 2000, Gowen et al. 1998, Moynahan et al. 1999, Scully et al. 1999, reviewed in Scully & Livingstone). BRCA1 co-localizes with Rad51 or with Rad50-Mre11-Nbs1 on sites of DNA damage or stalled replication forks (Scully et al. 1997, Zhong et al. 1999). These proteins are known to function in homologous recombination and/or in non-homologous end joining. After ionising radiation, the formation of BRCA1 foci is preceded by the accumulation of phosphorylated H2AX and p53 binding protein 1 (53BP1) (Paull et al. 26

2000, Rappold et al. 2001). Interestingly, budding yeast Rad9 is somewhat homologous to both p53BP1 and Brca1. In yeast, Mec1 phosphorylates Rad9 that thereafter becomes associated with Rad53 (Vialard et al. 1998). In mammalian cells, H2AX and 53BP1

Table 1. Comparison of cellular/viral replication proteins in the three kingdoms of life

Replication protein Eubacteria1 Eukaryotes SV402 HSV13 Archaea Origin recognition DnaA* ORC* T-ag* UL9 Cdc6?5* & unwinding4 Licensing/helicase DnaC* Cdc6/Cdc18 T-ag UL87 Cdc6 loading Licensing, helicase6 DnaB Mcm2-3-4-5-6-7* T-ag UL57 Mcm Pol α- Primase DnaG Pol α-primase UL527 primase primase ssDNA binding & SSB RPA RPA UL29 RPA stabilization β-subunit of Processivity clamp8 PCNA PCNA UL42 PCNA Pol III γ-complex of Clamp loader9 RFC* RFC - RFC Pol III*

PolC, PolA PolB PolB PolB, DNA polymerase UL30 (Pol III, Pol I) (Pol α, Pol δ, Pol ε) (Pol α, Pol δ) PolD

Maturation nuclease RnhA Fen1 Fen1 Fen1 Ligase Ligase Ligase I Ligase I LigaseI

1Nomenclature and data based on Escherichia coli. 2SV40 (Simian virus 40) uses the replication machinery of the host with a single viral encoded protein, T-ag. 3HSV1 (Herpes simplex virus 1) represents viruses that encode most of the proteins needed in replication. 4DnaA accumulates onto the origin as a homomultimer, ORC as a heterohexamer, T-ag as a double homohexamer, and UL9 as a dimer. 5Cdc6/Cdc18 is ATP-dependent loader of Mcm proteins. It shows to Orc proteins. In Archaea it possibly acts also as a origin recognition protein. 6DnaB and T-ag are essentially replicative helicases with mono and double homohexameric structure. Mcms are heteromultimers in eukaryotes, but homomultimers at least in some Archaea. They are required for loading of pols and a role as a replicative helicase has been proposed. 7HSV1 UL8, UL5 and UL52 protein form a heterotrimeric complex with helicase and primase activities. 8β-subunit dimer and PCNA trimer, although non-homologous in sequence, form similar ring-structure around DNA and function as a processivity factor for pol. UL42 is unrelated, and does not form ring-structure, but has intrinsic affinity on dsDNA. 10γ-complex is a heteropentamer of (γ, δ, δ’, χ, ψ). RFC is a heteropentamer of one large and four small subunits. In eukaryotes small subunits are distinct, but in Archaea all four are identical. * These proteins belong to the AAA+-ATPase superfamily. 27 become phosphorylated after double strand breaks by a Mec1 homologue (ATM, ATR and/or DNA-PK) (Paull et al. 2000, Rappold et al. 2001), whereas BRCA1 is phosphorylated by ATM after γ-irradiation and by ATR and Chk2 after several kinds of DNA damage or replication block (Cortez et al. 1999, Lee et al. 2000, Tibbets et al. 2000). The role of Rad9, 53BP1 and BRCA1 in damage recognition remains to be explored. Finally, p53, the tumour suppressor most often mutated in human cancers, functions downstream from the sensors and signal transducers mentioned above and plays a decisive role in the DNA damage response, whether the cell cycle is stopped for DNA repair or on a pathway toward apoptosis or senescence (reviewed in Appella & Anderson 2001, Somasundaram & El-Deiry 2001).

2.1.4 DNA replication in Archaea

The molecular dynamics in the third domain of life, Archaea, is still very poorly known. The cellular architecture and the genomic organization usually with a single circular chromosome is similar to that of prokaryotes whereas the proteins involved in DNA replication, transcription and protein synthesis show similarities to their eukaryotic counterparts. Currently, it is unclear if all Archaea utilize a single origin for chromosomal DNA replication as has been shown to be the case for Pyrococcus abyssi (Myllykallio et al. 2000). Theoretical studies based on strand specific bias on nucleotide, oligomer or codon frequencies suggest that some archaeal species like Methanococcus jannaschii utilize multiple origins (Mrazek & Karlin 1998). Archaeal homologues exist for the eukaryotic B-type DNA polymerase catalytic and B subunits, RPA, PCNA, RFC, Cdc6 and Mcm (Bernander 2000, MacNeill 2001, Mäkiniemi et al. 1999). It is possible that archaeal DNA replication is a model for eukaryotic replication with a minimal set of replication factors. A comparison of replication proteins with analogous function in the three domains of life is presented in Table 1.

2.2 Eukaryotic DNA polymerases

2.2.1 Classification, structure and function of DNA polymerases

So far, 15 DNA polymerases (pols) have been characterized in eukaryotes (Table 2). The catalytic subunits of pols can be divided into five families on the basis of sequence homologies to E. coli DNA polymerases (Ito & Braithwaite 1991, Aravind & Koonin, 1999, Friedberg et al. 2000, Ohmori et al. 2001). In addition, eukaryotes harbour template independent terminal transferases and RNA dependent telomerase, which are not discussed here. Pols γ and θ having roles in mitochondrial DNA replication and 28 interstrand cross-link repair, respectively, belong to family A, because of the similarity to E. coli pol I (Shadel & Clayton 1997, Sharief et al. 1999). The replicative pols α, δ and ε are related to pol II in E. coli and form the family B together with pol ζ that has a role in translesion synthesis (Nelson et al. 1996). All B family pols have similar polymerase

Table 2. Eukaryotic DNA polymerases

Pol Family Subunit composition Function(s)2 (kDa)1 Pol α B 166 (167) Replication, repair (DSBR), telomere maintenance, 66 (79) S phase checkpoint control 59 (62) 50 (48) Priming in replication Pol δ B 124 (125) Replication, repair (MMR, BER, NER, DSBR), 51 (55) recombination, telomere maintenance, S phase 51 (40) checkpoint control 12 Pol ε B 262 (256) Replication, repair (BER, NER, DSBR), 60 (79) recombination, transcriptional silencing, 17 (23) S phase checkpoint control 12 (22) Pol ζ B 353 (173) Translesion synthesis 24 (29) Pol γ A 140 (144) Mitochondrial DNA replication 55 Pol θ A 198 Interstrand cross-link repair? Pol β X 38 Repair (BER) Pol λ X 63 (68) Meiotic repair Pol µ X 55 Somatic hypermutation Pol σ1 X 57 (66) Establishment of sister chromatid cohesion (Trf4) Pol σ2 X 60 (73) Establishment of sister chromatid cohesion (Trf5) Pol η Y 78 (72) Translesion synthesis Pol ι Y 80 Translesion synthesis, mutagenesis? Pol κ Y 99 Translesion synthesis Rev1 Y 138 (112) Translesion synthesis

1Calculated from cDNAs for the human polypeptides. The mass of the S .cerevisiae orthologue is shown in parentheses. 2BER = base excision repair, DSBR = double strand break repair, MMR = mismatch repair, NER = nucleotide excision repair 29 domains with six conserved sequence motifs (Ito & Braitwaite 1991) and some of them share homologous N-terminal, exonuclease and zinc-finger domains (Figure 5). Furthermore the non-catalytic B subunits of B-family pols are related to each other and to the B subunits of archaeal D-family pols (Mäkiniemi et al. 1999). No homologues for E. coli pol III exist in eukaryotes. E. coli pol IV (also called DinB) and pol V (also called UmuC) are members of a recently recognized superfamily Y (Ohmori et al. 2001). Pols η, ι, κ and Rev1 also belong to this family and have denoted roles in translesion synthesis (Masutani et al. 1999, Johnson et al. 2000, Ohashi et al. 2000, Haracska et al. 2001). Originally, DNA polymerases that did not have a recognized counterpart in E. coli were classified as family X. That is now considered to be a subfamily in a larger superfamily (Aravind & Koonin 1999). Of these, a few have been characterized as DNA polymerases. Pol β is a major base excision repair pol in animals (reviewed in Wilson 1998), pol λ has an obvious role in meiosis-associated repair (Garcia-Diaz et al. 2000) and pol µ is involved in somatic hypermutation in lymph nodes (Dominguez et al. 2000). In addition, two redundant homologues, TRF4 and TRF5 in S. cerevisiae encode the fourth essential nuclear DNA polymerase, pol σ, that links DNA replication to the establishment of sister chromatid cohesion (Wang Z et al. 2000, Carson & Christman 2001). A relative of Trf4 and Trf5 in S. pombe, Cid1, has a S/M checkpoint function (Wang S et al. 2000). Finally, a distinct DNA polymerase family D exists in euryarchaeota.

Fig. 5. The comparison of conserved motifs between B family pols. Adapted from Hübscher et al. 2000.

A limited number of crystal structures for DNA polymerase catalytic subunits are known, and the three-dimensional structure of only one eukaryotic DNA polymerase, pol β, has been resolved so far (Sawaya et al. 1994). No structure of a replicative pol is known from eukaryotes, but structures of B family pols are known from bacteriophage RB69 and from four archaeal species: Desulfurococcus strain Tok, Thermococcus 30 gorgonarius, Pyrococcus kodakaraensis KOD1 and Thermococcus sp. 9 degrees N-7 (Wang et al. 1997, Zhao et al. 1999, Hopfner et al. 1999, Hashimoto et al. 2001, Rodriguez et al. 2000). Ternary complex structures with primer-template DNA and deoxynucleotides are known for pol β and RB69 (Pelletier et al. 1994, Franklin et al. 2001). All pols contain three distinct domains, “thumb”, “palm” and “fingers” with analogous functions. The catalytic centre is located on the “palm” domain with two conserved aspartate residues. The “thumb” and “fingers” domains, although analogous, are not homologous between the pol families. Despite the divergent evolutionary origins, structural and enzymological studies show that all pols obviously utilize a universal ‘two metal cation’ reaction mechanism. The binding of the correct deoxynucleotide induces a conformational change in the “fingers” domain that is essential for catalysis. The selection of the correct incoming deoxynucleotide is based on steric interactions and Watson-Crick base pair formation between the deoxynucleotide and the template DNA. The two aspartates coordinate the two magnesium ions. One of the metal ions activates the hydroxyl group of the primer by lowering its pKa enabling nucleophilic attack on the α-phosphate. This generates a penta-coordinated transition state that is stabilized by both metal ions. The other metal ion also participates in stabilizing the negative charge of the leaving β-phosphate oxygen. The pyrophosphate cleavage is followed by a conformational change and translocation of the DNA on the polymerase (Brautigam & Steitz 1998).

2.2.2 DNA polymerase alpha-primase

Mammalian DNA polymerase alpha-primase (pol α) consist of four subunits (A, B, C, D) with calculated molecular weights of 166 kDa, 66 kDa, 59 kDa and 50 kDa. The apparent molecular weights of the two largest subunits are 180 kDa and 90 kDa, due to post- translational modifications. The A subunit contains DNA polymerase activity without exonuclease proofreading. The B subunit is not known to have any catalytic function, but has a role in regulation and in tethering pol α to the replication machinery. The two smallest subunits form a primase that is required in the initiation of all chromosomal DNA synthesis de novo (Wang 1996). The expression of pol α (A subunit) is up-regulated when quiescent cells are mitogenically activated to re-enter the cell cycle. This is reflected at the mRNA and protein levels as well as in enzyme activity of pol α. However, pol α is constitutively expressed during the cell cycle in actively cycling cells with a minor increase at the G1/S border (Wahl et al. 1988). The other subunits are also expressed in a similar manner (Miyazava et al. 1993). The mitogenic activation of both the A and B subunit promoters is mediated by E2F transcription factors (Pearson et al. 1991, Nishikawa et al. 2000). In the budding yeast S. cerevisiae, the genes for all subunits of pol α are co-ordinately induced at the G1/S border by the MBF transcription factor (MluI cell cycle box binding factor) (Johnson et al. 1985, Foiani et al. 1989, Foiani et al. 1994). MBF and SBF (Swi4/6 Cell cycle box binding factor) are heterodimeric transcription factors consisting of a Swi6 regulatory subunit complexed with either Mbp1 or Swi4 DNA binding proteins, 31 respectively. They have binding sites in the promoters of several DNA replication related genes, such as the genes for DNA polymerases, and activate transcription of these genes in the late G1 phase only, being themselves regulated by the Cln3/Cdk1 kinase (Ho et al. 1999, reviewed in Koch & Nasmyth 1994). Both the yeast and the mammalian pol α is phosphorylated in a cell cycle dependent manner (Wong et al. 1986, Nasheuer et al. 1991, Foiani et al. 1995). The human pol α A subunit is phosphorylated throughout the cell cycle, but becomes hyperphosphorylated in the G2/M phase. The B subunit becomes phosphorylated in S phase and hyperphosphorylated in G2/M. Both subunits are phosphorylated by cyclin dependent kinases in vitro (Nasheuer et al. 1991). The polymerase activity is not affected by phosphorylation, but the primase activity is moderately stimulated in vitro (Voitenleitner at al. 1997). When the effects of several Cyclin-Cdks were studied in an SV40 initiation assay, Cyclin A/Cdk2 dependent phosphorylation of recombinant pol α inhibited initiation of DNA replication, whereas phosphorylation by Cyclin E/Cdk2 stimulated it. The two-dimensional tryptic phosphopeptide maps from the B subunit that had been phosphorylated by Cyclin E/Cdk2 in vitro, resembled those of the endogenous B subunit in G1/S synchronized cells. Correspondingly, the phosphopeptide map from the pol α B subunit from G2 cells was similar to that from the Cyclin A/Cdk2 phosphorylated recombinant B subunit (Voitenleitner et al. 1999). Recently, another study showed that newly synthesized pol α A and B subunits are hypophosphorylated, interact with Cyclin E/Cdk2 and protein phosphatase PP2A, and co-localize with Mcm2 to origins of DNA replication, but not on foci with high DNA synthesis. In contrast, pol α with strongly phosphorylated A and B subunits interacts with cyclin A and co-localizes in sites of ongoing DNA replication, but not with Mcm2 (Dehde et al. 2001). In addition to the essential role in DNA replication (discussed above in 2.1.3), several other functions have been implicated for pol α. These include cell cycle checkpoint control in S. pombe (D’Urso et al. 1995, Tan & Wang 2000) and in Xenopus (Michael et al. 2000), DNA double strand break repair by homologous recombination in S. cerevisiae (Holmes & Haber 1999) and by non-homologous end-joining in mammalian cells (Pospiech et al. 2001) and telomere maintenance in S. cerevisiae (Diede & Gottschling 1999). Pol α has been reported to interact physically with the single strand binding protein RPA (Dornreiter et al. 1992, Braun et al. 1997, Maga et al. 2001), initiation proteins Mcm3 and Cdc45 (Thömmes et al. 1992, Mimura & Takisawa 1998), the pol δ C subunit (Huang et al. 1999c), poly(ADP-ribose) polymerase (Dantzer et al. 1998), tumour suppressors p53 (Kuhn et al. 1999) and pRb (Takemura et al. 2001), mutually exclusively to yeast cohesion protein Ctf4 or chromatin structure proteins Cdc68/Spt16 and Pob3 (Miles & Formosa 1992, Wittmeyer & Formosa 1997), yeast telomeric binding protein Cdc13 (Qi et al. 2000), growth suppressor p12(DOC-1) (Matsuo et al. 2000) and several viral origin recognition proteins/helicases like SV40 large T-antigen, human papillomavirus E1 and herpes simplex virus UL9 (Dornreiter et al. 1990, Conger et al. 1999, Lee et al. 1995). 32

2.2.3 DNA polymerase delta

DNA polymerase delta (pol δ) has been purified from several species and shows some degeneracy in its subunit composition. S. cerevisiae pol δ contains three subunits (Gerik et al. 1998), whereas fission yeast S. pombe pol δ has four or five subunits (Zou et al. 1997). The mammalian enzyme consists of at least four subunits with apparent molecular weights of 125 kDa, 68 kDa, 50 kDa and 12 kDa (Liu et al. 2000). The message level and enzyme activity of pol δ are up-regulated when quiescent cells are induced to proliferate (Yang et al. 1992). During the cell cycle, both the mRNA and protein levels of pol δ fluctuate moderately with a two to three fold increase at the G1/S phase (Zeng et al. 1994a). An E2F consensus binding-like element is found in the promoter of human pol δ gene, but there are no reports of a role for E2F in the regulation of the pol δ gene. However, transcription factors Sp1 and Sp3 have well characterized roles in the stimulation of pol δ (Zhao & Chang 1997). The expression of pol δ is down-regulated by DNA damage and this repression is mediated by p53 (Li & Lee 2001). Pol δ is a phosphoprotein and is hyperphosphorylated in S phase (Zeng et al. 1994). When co-expressed with different Cyclin-Cdks in insect cells, pol δ interacts with and becomes phosphorylated by Cyclin D3/Cdk4 and Cyclin E/Cdk2. This result suggests that late G1 and G1/S specific Cdks potentiate pol δ for S phase. However, phosphorylation by Cdks does not affect polymerase activity in vitro (Wu et al. 1998).

Table 3. Comparison of the regulation of mammalian pol α and polδ

Pol subunit Expression Phosphorylation

Pol α subunit A G0/G1: 8-fold up in steady-state mRNA G1/S: substrate for Cyclin E/Cdk2, 11-fold up in de novo protein Cdc7/Dbf4

G1/S: < 2-fold up in steady-state mRNA S/G2: substrate for Cyclin A/Cdk2 no change in immunoreactive protein on T174, S209, T219

Pol α subunit B G0/G1: 10-fold up in steady-state mRNA G1/S: substrate for Cyclin E/Cdk2

G1/S: no significant change S/G2: substrate for Cyclin A/Cdk2

Pol δ subunit A G0/G1: 10-fold up in steady-state mRNA G1/S: substrate for Cyclin D3/Cdk4,

G1/S: 3-fold up in steady-state mRNA Cyclin E/Cdk2 2-fold up in immunoreactive protein

Pol δ subunit B G0/G1: no significant change? ?

Data from Wahl et al. 1988, Miyazawa et al. 1993, Dehde et al. 2000, Voitenleitner et al. 1999, Weinreich & Stillman 1999, Schub et al. 2001, Yang et al. 1992, Zeng et al. 1994, Wu et al. 1998, Iyer et al. 1999.

The characteristic feature of pol δ is its tight coupling to the proliferating cell nuclear antigen (PCNA), a processivity factor. When the polymerase activity of pol δ is assayed on a ssDNA primer template, PCNA increases the processivity of pol δ up to 100-fold by forming a “sliding clamp” - a ring structure that holds the polymerase on DNA (Tan et al. 1986, Prelich et al. 1987, Krishna et al. 1994). On the covalently closed DNA, the 33 formation of a PCNA clamp is dependent on the replication factor C (RFC) (Yoder & Burgers 1991). Both the N-terminus of the catalytic subunit and the C subunit of pol δ contact PCNA (Zhang et al. 1999, Reynolds et al. 2000). The latter interaction is mediated via a so-called ‘PIP’-box motif (PIP = PCNA interacting protein) that binds into a hydrophobic pocket beside the interdomain connector loop of PCNA. The B subunit is also essential for the functional interaction with PCNA at least in the absence of the C subunit (Zhou et al. 1997). In addition, Pol δ physically interacts with RFC (Pan et al. 1993), RPA (Yuzhakov et al. 1999), Werner syndrome protein (WRN) (Szekely et al. 2000), S. cerevisiae Srs2 helicase (Huang et al. 2000) and a hypophosphorylated form of the tumour suppressor pRb in the G1 phase (Krucher et al. 2000). Werner syndrome is an inherited human disorder with clinical manifestations of premature aging. The defective gene product, WRN, belongs to the family of RegQ class helicases and also has 3’- >5’exonuclease activity (Kamath-Loeb et al. 2000, reviewed in Enomoto 2001). The interaction of WRN with pol δ can recruit the latter into the nucleolus (Szekely et al. 2000) and WRN is also a pol δ specific accessory factor in resolving hairpin and tetraplex structures in repetitive DNA (Kamath-Loeb et al. 2000). In S. cerevisiae pol δ interacts with Srs2 helicase that is synthetically lethal with Sgs1 helicase – a WRN homologue (Lee et al. 1999). Although WRN inevitably is involved both in initiation and elongation of DNA replication, it is not essential for viability and not likely a major replicative helicase (Chen et al. 2001). Instead, it seems to have a central role in resolving and recruiting pol δ to aberrant DNA structures and stalled replication forks (Constantinou et al. 2000, Kamath-Loeb et al. 2000). Pol δ is a major replicative polymerase and its function in SV40 DNA replication has been described earlier. It has a crucial function in cellular DNA replication as well, but its exact role is not known. The replicative roles of pol δ and pol ε are discussed in the next chapter. Pol δ is also involved in several non-replicative functions. It has a minor role in base excision repair in yeast (Wang et al. 1993, Blank et al. 1994) and a gap-filling function in mammalian long-patch base excision repair, probably redundantly with pol ε (Stucki et al. 1998, Fortini et al. 1998). Both pol δ and pol ε are required for DNA synthesis in nucleotide excision repair (Zeng et al. 1994b, Budd & Campbell 1995, Araujo et al. 2000, Wu et al. 2001). The role of pol δ in nucleotide excision repair in S. cerevisiae is still unclear. The nucleotide excision repair in vitro has not been reconstituted with yeast proteins. Although Budd and Campbell (1995) found that temperature sensitive pol δ-pol ε double mutants were deficient in repair of UV-induced damage, this does not distinguish between nucleotide excision repair and RAD6 dependent post-replicative repair. Pol δ has been implicated in the latter (Huang et al. 2000) and a recent report indicates a role for pol δ together with pol ζ and Rev1 in the bypass of abasic sites as well (Haracska et al. 2001). However, a further study showed reduced DNA repair synthesis in cell extracts prepared either from pol δ or pol ε deficient cells. The addition of purified pol restored the original level of synthesis and this assay was specific for nucleotide excision repair (Wu et al. 2001). In human cells, pol δ is the major, if not the only, polymerase involved in mismatch repair (Longley et al. 1997). Finally, a function for pol δ in recombination, double strand break repair, telomere maintenance and cell cycle checkpoint control has been reported (Giot et al. 1997, Paquet & Haber 1997, Holmes & Haber 1999, Diede & Gottschling 1999, Francesconi et al. 1993). 34

2.2.4 DNA polymerase epsilon

DNA polymerase epsilon (pol ε) was first purified from yeast S. cerevisiae as DNA polymerase II in 1970 (Winterberger & Winterberger 1970) and from a mammalian source, rabbit bone marrow, in 1976, although at that time it was called pol δ (Byrnes et al. 1976, Lu & Byrnes 1992). Later pol ε was purified as a large, PCNA independent form of pol δ from calf thymus and from Hela cells (Crute et al. 1986, Syväoja & Linn 1989). It was not until the 1990’s that pol ε was shown to be a DNA polymerase distinct from pol α and pol δ (Syväoja et al. 1990), and a homologue of S. cerevisiae pol II (Kesti et al. 1993), nowadays also called pol ε (Burgers et al. 1990). So far pol ε has been purified from human, rabbit, bovine, silkworm Bombyx mori, mosquito and S. cerevisiae and the catalytic subunit (A subunit) of pol ε has been cloned from human, mouse, D. melanogaster, S. pombe and S. cerevisiae. In addition at least partial cDNAs are known from several species. The amino acid sequence of pol ε shows a high conservation with 90 % identity between human and mouse and about 40 % identity between distant eukaryotes like mammals vs. yeast. The highest conservation is located in the N-terminal half where there are three conserved motifs in the N-terminal domain, five (or six) in the exonuclease domain, seven in the polymerase domain and three more C-terminal motifs. Notably, even thought the exonuclease and polymerase motifs are found in other B class pols, a few unique features exist for pol ε. First, motif II has been split in two by a 65 aa residue insert. Secondly, there are differences in the sequence of three motifs (I-III) that form the catalytic centre of the pols. The well-conserved tyrosine in motif I is replaced by glutamate in pol ε. Similarly, two conserved serines in motif II are replaced by alanine (in metazoan pol ε) and asparagine. Also, critical threonine and arginine residues in motif III are not conserved in pol ε. Many of these residues are important for nucleotide binding and/or template-primer stabilization, but the significance of these replacements is not known. Finally, two potential zinc-finger motifs are present in the extreme C-terminus of pol ε with similarity to corresponding motifs in pol α, pol δ and pol ζ (Huang et al. 1999a). Most of the C-terminal half of the protein, that is the region between the polymerase domain and the C-terminal zinc-finger domain, is unique for pol ε and least conserved between species. This region shows very limited homology to the N-terminus predicting a partial duplication of an ancestral gene. The budding yeast pol ε consists of five polypeptides with molecular weights of 256, 79, 34, 30 and 29 kDa and a stoichiometry of 1:1:4:1:4 as determined from SDS-PAGE. The 256 kDa polypeptide may undergo a proteolytic cleavage and the major product, a 145 kDa polypeptide, without associated subunits, retains the catalytic properties of polymerase and exonuclease activities (Hamatake et al. 1990). The pol ε subunits are encoded by POL2 (256 kDa A subunit), DPB2 (79 kDa B subunit), DPB3 (34 kDa C subunit and its 30 kDa proteolytic product) and DPB4 (29 kDa D subunit). POL2 and DPB2 are essential for DNA replication and viability (Morrison et al. 1990, Araki et al. 1991a,b, Ohya et al. 2000). The polymerase and exonuclease domains in the N-terminal half of the A subunit are dispensable as is the checkpoint function of the C-terminus (Kesti et al. 1999). The non-redundant property of the C-terminal part is unknown, but it may be important in the protein-protein interactions that are essential for assembly or maintenance of replication complex. The C-terminal half of the A subunit is sufficient for 35 the interaction with the B subunit and this interaction is weakened by introducing a deletion into a zinc-finger domain in the A subunit (Dua et al. 1998). Later, it was shown that the B subunit could dimerize and mediate the formation of dimeric polymerase i.e. tetrameric complex with two dimers of A and B subunits. Furthermore, the C and D subunits form a tight complex that interacts with modest affinity with both A and B subunits (Dua et al. 2000). The mammalian pol ε has been purified as a dimeric enzyme of the 215 kDa catalytic (A) subunit and 55 kDa (B) subunit with unknown function (Kesti et al. 1991). A 140 kDa form of pol ε from calf thymus and a 170 kDa form from human placenta have also been described (Kesti et al. 1991, Weiser et al. 1991, Lee et al. 1987). These preparations are often devoid of a B subunit, and biochemical and immunological studies strongly imply that these smaller forms are produced from the 215 kDa polypeptide by proteolytic processing (Kesti et al. 1991, Uitto et al. 1995). Specifically, during an early stage of apoptosis, caspase 3 is responsible for this cleavage (Liu & Linn 2000). Recently, the C and D subunits for human pol ε were reported. They contain histone fold motifs and are homologous to the C and D subunits of yeast pol ε (Li et al. 2000). Interestingly, subunit C has been characterized as CHRAC17, a component of the human chromatin remodeling complex ISWI. Furthermore, a second subunit of ISWI, CHRAC15, is similar to subunit D of pol ε. Currently, the functions of the C and D subunits are unknown, but roles in replication associated chromatin remodeling and/or transcriptional (de) silencing are possible. The differences between pol ε and other class B pols in the primary structure of catalytically important motifs suggest that pol ε might have characteristic catalytic properties and sensitivity to inhibitors. Pol ε has a high inherent processivity and fidelity, the latter partially because of the associated 3’-5’ exonuclease activity. Pol ε utilizes poly(dA)-oligo(dT) as a preferential template in low salt and high (15 mM) Mg2+ concentrations (Syväoja & Linn 1989). An increase in monovalent salt concentration up to 100 – 200 mM (KCl, potassium glutamate) completely inhibits pol ε on this template- primer as well as on single primed M13 ssDNA (Syväoja & Linn 1989, Lee et al. 1991, Kesti et al. 1991). However, on gapped dsDNA pol ε is tolerant up to 140 mM KCl (Syväoja & Linn 1989, Kesti et al. 1991). The high Mg2+ optimum on poly(dA)- oligo(dT) makes pol ε different from pol α. However, on single primed M13 ssDNA both pols have their highest activity at 1 mM MgCl2 (Chui & Linn 1995). All pols α, δ and ε are unable to use RNA as a template, but all utilize RNA efficiently as primer (Syväoja & Linn 1989, Chui & Linn 1995). Pol ε does not need PCNA as an auxiliary factor for processive DNA synthesis on homonucleotide templates in vitro and that is a major difference from pol δ. Neither does PCNA increase the activity of human pol ε in vitro, but this maybe a property of the purified human enzyme without other replication factors present. A truncated form from calf thymus both interacts with and is stimulated by PCNA (Maga & Hübscher 1995) and PCNA together with RPA and RFC is able to overcome the inhibition of high salt concentration on poly(dA)-oligo(dT) or on single primed M13 ssDNA (Lee et al. 1991, Chui & Linn 1995). Pol ε is sensitive to both aphidicolin and araCTP as are pols α and δ, but not to dideoxynucleotides like ddTTP (Syväoja & Linn 1989, Mirzayans et al. 1993). In contrast to pol α, pol ε is not very sensitive to butylphenyl-dGTP or butylphenyl-dATP. Pol ε is also not sensitive to carbonyldiphosphonate or 9-(2-phosphonomethoxyethyl)-2,6-diaminopurine diphosphate 36

(PMEDAPpp) like pol δ, but is sensitive to DMSO and (S)-9-(3-hydroxy-2- phosphonomethoxypropyl)adenine diphosphate (HPMPApp) (Syväoja et al. 1990, Kramata et al. 1996). In addition, specific inhibition of pol ε by acidic phospholipids has been reported (Shoji-Kawaguchi et al. 1995). Pol ε is known to interact with several proteins involved in DNA replication, repair, recombination and cell cycle control. The biochemical interaction with pol ε has been shown for RPA (Lee et al. 1991, Shivji et al. 1995), PCNA (Lee et al. 1991, Maga & Hübscher 1995, Shivji et al. 1995), RFC (Lee et al. 1991, Podust et al. 1992, Pan et al. 1993, Shivji et al. 1995), Fen1 nuclease (Siegal et al. 1992, Turchi & Bambara 1993), ligase I (Mossi et al. 1998), poly (ADP-ribose) polymerase (Eki 1994), helicase E (Turchi et al. 1992) and stimulatory factor I (Smiley et al. 1992). A few proteins like PCNA (Coll et al. 1997), p53 regulator MDM2 (Vlatkovic et al. 2000), S. cerevisiae Dpb11 (Masumoto et al. 2000) and a complex of Cdc45, RPA and Mcm2 (Zou & Stillman) have been reported to have a direct physical interaction with pol ε. Furthermore, genetic interactions of pol ε with a potential DNA polymerase Cid1 and RFC- and the PCNA- associated protein Cdc24 in S. pombe (Wang et al. 2000, Tanaka et al. 1999) as well as with Sld2 and Sgs1 helicase in S. cerevisiae (Kamimura et al. 1998, Wang & Elledge 1999, Frei & Gasser 2000) are known. Finally pol ε has been purified from mammalian cells as a component in a replicative synthesome (Coll et al. 1996), in a recombinative complex (Jessberger et al. 1993) and in a RNA polymerase II complex (Maldonado et al. 1996), and has also been shown to be associated with pol α, Cdc45, Mcm4, Mcm6 and Mcm7 in yeast replication forks (Aparicio et al 1997, Aparicio et al. 1999). The specific role of pol ε in DNA replication has been under debate since its characterization. There is a consensus about the function of pol α in the initiation and in RNA-DNA primer synthesis for each Okazaki fragment. Based on the observations with SV40 DNA replication, pol δ was proposed to be a principal pol in elongation of both leading and lagging strands - also in cellular DNA replication (Waga & Stillman 1994). This model suggests a dimeric pol δ analogous to E. coli pol III. However, in addition to pol δ, pol ε was also found to be essential for DNA replication in yeast (Morrison et al. 1990, Araki et al. 1992). As mentioned earlier, pol ε associates with replication forks in yeast, can be cross-linked to newly synthesized replicating chromosomal DNA in mammalian cells, and its inactivation by neutralizing antibodies or by immunodepletion causes a cessation of DNA replication. So, what is pol ε doing there? Many observations support a model in which pol δ and pol ε replicate different strands. Burgers proposed a model, in which pol δ synthesizes the leading strand while pol ε is a lagging strand polymerase. This model was based on observations that human pol ε co-purifies with pol α and it synthesizes shorter DNA products in an SV40 system than pol δ (Lee et al. 1991) and that S. cerevisiae pol ε holoenzyme has a > 50 % slower elongation rate compared to pol δ (Burgers 1991). Sugino and co-workers proposed exactly the opposite based on the inherent processivity of pol ε (Morrison et al. 1990). In any case, several studies suggest that these two pols function on different strands. This is concluded from biased mutation frequencies in exonuclease defective pol δ and pol ε mutants, where a reporter gene is located in a defined orientation with regard to the origin of DNA replication (Morrison & Sugino 1994, Shcherbakova & Pavlov 1996, Karthikeyan et al. 2000). Despite that, some evidence still supports the original model of pol δ as the replicator of both strands, and suggests a role for pol ε in Okazaki fragment maturation or in another replication-linked 37 process with a requirement of DNA synthesis (Bambara 1997, Burgers 1998). These arguments include the existence of a dimeric form of pol δ (Burgers & Gerik 1998), a finding that pol ε synthesizes much less DNA than does pol δ (Zlotkin et al. 1996), the viability of yeast cells without the polymerase domain of pol ε (Kesti et al. 1999, Feng & D’Urso 2001) and the fact that pol ε is functionally coupled to the Fen-1 nuclease (Siegal et al. 1992, Turchi & Bambara 1993). However these results must be viewed with caution. First, a double replisome model with two coupled replication forks can explain the presence of a dimeric pol δ and pol ε with distinct strand preference: two lagging/leading strand pols are synthesizing lagging/leading strands of different forks as a dimer (Dua et al. 1999, Falaschi 2000). Secondly, the modest amount of DNA synthesis measured by Zlotkin et al. (1996) maybe characteristic for a certain form of pol ε (this study). Thirdly, the results by Kesti et al. (1999) and Feng & D’Urso (2001) suggest an essential role for the C-terminus of pol ε in the assembly of the replication complex. Altogether, the exact function of pol ε remains to be elucidated and the models described can be over-simplifications. The different structures of pol δ and pol ε and interactions with other proteins indicate a possibility that these pols, although partially redundant, are specialized for different kinds of templates like euchromatin vs. heterochromatin, transcribed vs. non-transcribed templates, regions near vs. far from origins or synthesis started from origin vs. started by recombination. Interestingly, pol ε has been shown to have an important role in the initiation of DNA replication (Masumoto et al. 2000, Feng & D’Urso 2001) and be involved in replication-linked recombination (Zou & Rothstein 1997). In addition to the role in DNA replication, pol ε has been implicated in base excision repair and in nucleotide excision repair (Wang et al. 1993, Stucki et al. 1998, Fortini et al. 1998, Budd & Campbell 1995, Araujo et al. 2000, Wu et al. 2001, see discussion in 2.2.3), in homologous recombination dependent double-strand break repair (Holmes & Haber 1999, in S/M and intra-S checkpoint control (Navas et al. 1995, Frei & Gasser 2000) and in transcriptional silencing (Ehrenhofer-Murray et al. 1999). Pol ε was originally purified as a factor needed for the repair synthesis of UV-damaged DNA (Nishida et al. 1988). Although a study with a specific antibody against pol δ has demonstrated the important role of pol δ, the experiments with highly purified pols show that both pol δ and pol ε are able to fulfill the synthesis step in reconstituted nucleotide excision repair in vitro (Araujo et al. 2000). Pol ε has a well-demonstrated S phase checkpoint function in S. cerevisiae. It acts upstream of Mec1 and Rad53 in the S/M checkpoint and this function is independent and parallel to that of S. cerevisiae Rad9 (Navas et al. 1995, 1996). Pol ε is also involved in the intra-S checkpoint, where it is epistatic to Sgs1 helicase, but parallel to Rad24 (Frei & Gasser 2000). These checkpoint functions require the presence of the extreme C-terminus of the pol ε A subunit. Although this region is moderately conserved, no S phase checkpoint function has been denoted for pol ε in other eukaryotes and it has been shown that S. pombe pol ε mutants are defective in the onset of DNA replication, but still have an intact S/M checkpoint (D’Urso & Nurse 1997). 38

2.3 Proteins associated with DNA replication and S phase checkpoint

2.3.1 Mcm2, Mcm3, Mcm4, Mcm5, Mcm6 and Mcm7

Minichromosome maintenance proteins Mcm2, Mcm3, Mcm4, Mcm5, Mcm6 and Mcm7 are homologous proteins that can form several kinds of hetero-oligomeric complexes. As discussed earlier, they have relatively well characterized roles in regulating the initiation of DNA replication, but they are also required in elongation of replication forks and because of their intrinsic 3’-> 5’ helicase activity, it has been proposed that they have a role as the replicative helicase in eukaryotic cells (Tye 1999, Labib et al. 2000, Labib & Diffley 2001). A recent study shows that a double hexameric complex, each consisting of two pairs of Mcm4-Mcm6-Mcm7 dimers, is indeed a processive helicase with a forked DNA substrate (Lee & Hurwitz 2001). The essential involvement both in initiation and elongation of DNA replication indicates they have a function analogous to the SV40 large T-antigen. The final demonstration of the Mcm-complex as a eukaryotic replicative helicase still requires reconstitution of cellular DNA replication from a defined origin in vitro. The genes for mammalian Mcms are expressed both in a growth and cell cycle dependent manner and this is mediated by transcriptional induction by E2F3 in the late G1 phase (Leone et al. 1997, Ishida et al. 2001). However, the amounts of Mcms at the protein level do not fluctuate during the cell cycle (Mendez & Stillman 2000). Indeed, it seems that Mcms are very abundant in all eukaryotic cells and the regulation of their function is dependent on sub-cellular targeting and on post-translational modifications (Tye 1999). In S. cerevisiae, the bulk of the Mcms is located in the nucleus in G1 and exported out during S phase in a process dependent on B-type Cyclin/Cdk activity (Hennessy et al. 1990, Dalton & Whitbread 1995, Labib et al. 1999, Nguyen et al. 2000). In mammalian cells, Mcms are mainly nuclear throughout the cell cycle, but they shuttle between chromatin and nucleoplasm, become chromatin-bound in late mitosis and are released during DNA replication (Mendez & Stillman 2000). Mcm2, Mcm3 and Mcm4 show cell cycle specific phosphorylation that correlates with chromatin binding. They are hypophosphorylated and chromatin-bound in G1, but become phosphorylated by Cdk activity at G1/S and later in the cell cycle (Young & Tye 1997, Fujita et al. 1998). Phosphorylation of Mcm proteins is also essential for pre-RC assembly and activation. This process is dependent on Cdk activity and Cdc7-Dbf4 kinase is also required. Mcm2 is phosphorylated by Cdc7 in vivo (Lei et al. 1997, Jiang et al. 1999, Masai et al. 2000) and, Mcm2, Mcm3, Mcm4 and Mcm6 are good substrates for Cdc7 in vitro (Sato et al. 1997, Weinreich & Stillman 1999).

39

2.3.2 Dpb11 and homologous proteins

DPB11 (DNA polymerase B possible subunit) was isolated as a multi-copy suppressor of S. cerevisiae pol ε A and B subunit mutants pol2-11 and dpb2-1 (Araki et al. 1995). A temperature sensitive mutant, dpb11-1, has both DNA replication and S/M checkpoint defects and is synthetically lethal with temperature sensitive mutants of each of the pol ε subunits (Araki et al. 1995, Kamimura et al. 1998, Ohya et al. 2000). Dpb11 has been shown to physically interact with pol ε and is required for the loading of pol α and pol ε onto the origins of DNA replication (Masumoto et al. 2000). Dpb11 is homologous to S. pombe Cut5, an essential protein for DNA replication and also involved in the DNA damage checkpoint (Saka & Yanagida 1993, Saka et al. 1997). In addition, Dpb11 is homologous to D. melanogaster Mus101, linked to DNA replication, repair, and chromosome condensation (Yamamoto et al. 2000). DPB11 interacts not only with pol ε, but also genetically with CDC45, RAD53, SLD2 and SLD3 (Sld = synthetically lethal with dpb11-1), all of which are essential genes encoding replication/checkpoint proteins (Kamimura et al. 1998, 2001). Dpb11 is expressed constitutively during the cell cycle (Kamimura et al. 1998). In a screen for topoisomerase IIβ interacting proteins a human protein with eight BRCT- domains was discovered. This protein was designated as topoisomerase IIβ binding protein 1 (TopBP1) and found to interact with topoisomerase IIβ in vitro (Yamane et al. 1997). Importantly, TopBP1 was found to show similarity to Cut5. Further studies have revealed that TopBP1 binds DNA strand breaks and DNA ends and that TopBP1 expression is down-regulated after UV irradiation (Yamane & Tsuruo 1999, Potter et al. 2000).

2.3.3 Cdc45

The human Cdc45 is a 66 kDa protein. It is essential for DNA replication as demonstrated in S. cerevisiae and in S. pombe (Hopwood & Dalton 1996, Zou et al. 1997, Miyake & Yamashita 1998). It associates with Mcm proteins in the pre-RC and is required for loading of pol α and pol ε onto the origin of DNA replication (Mimura & Takisawa 1998, Aparicio et al. 1999, Zou & Stillman 2000). Furthermore Cdc45 has an essential function in the elongation phase of DNA replication, although its specific role in this phase is not yet known. Cdc45 interacts physically with the Mcm proteins, RPA, pol α and pol ε (Kukimoto et al. 1999, Zou & Stillman 1998, 2000). The steady-state mRNA level for the human Cdc45 shows a dramatic up-regulation at the G1/S phase, but at the protein level it is constitutively expressed throughout the cell cycle (Saha et al. 1998). Cdc45 is regulated by phosphorylation at least in S. cerevisiae. It is phosphorylated by the Cdc7 kinase (Nougarede et al. 2000), and its assembly on the pre-RC is dependent on Cdk activity, although it is not known if it is a direct substrate (Zou & Stillman 1998, 2000). 40

2.3.4 Replication protein A

The replication protein A (RPA) is a heterotrimeric ssDNA binding protein implicated in DNA replication, recombination and repair (Longhese et al. 1994, reviewed in Wold 1997). The human RPA was purified as an essential factor for SV40 replication in vitro and consists of 70, 32 and 14 kDa subunits (p70, p32, p14; Wold & Kelly 1988). It binds to ssDNA with high affinity (>109 M-1) and low co-operativity. This interaction is mediated via the central domain of p70 and protects 30 nt of DNA (Kim et al. 1992, Kim & Wold 1995, Gomes & Wold 1996, Walther et al. 1999). Another mode of ssDNA binding also exists, where RPA covers 8-10 nt and this binding is co-operative (Blackwell & Borowiec 1994). RPA interacts with several proteins involved in DNA replication, repair and cell cycle control like SV40 T-antigen, pol α, p53 and the nucleotide excision repair proteins XPA, XPF and XPG (Wold 1997). Gene expression of the RPA subunits has been studied in S. cerevisiae, the trypanosamatid Crithidia fasciculate and in D. melanogaster, where RPA transcripts show accumulation in the G1/S phase (Brill & Stillman 1991, Brown & Ray 1997, Perdigao et al. 1999). Furthermore the gene for p32 has been shown to be under E2F-1 regulation (Kalma et al. 2001). However, the amount of RPA protein remains nearly constant throughout the cell cycle. By contrast, p32 is phosphorylated in a cell cycle dependent manner, being phosphorylated in the S and G2/M phases, but not in G1 (Din et al. 1990). The efficient phosphorylation requires ssDNA, when only DNA-bound RPA becomes phosphorylated (Fotedar & Roberts 1992, Blackwell et al. 1996). The cell cycle specific phosphorylation is catalysed by a DNA dependent protein kinase (DNA-PK), but RPA is also a substrate for other kinases. ATM is able to phosphorylate RPA after DNA damage (Brush et al. 1994, Liu & Weaver 1993, Carty et al. 1994, Gately et al. 1998). p32 is also a substrate for Cdks in vitro, but mutations of Cdk target sites do not affect RPA’s function in replication or in repair nor do they prevent hyperphosphorylation (Henricksen & Wold 1994). The phosphorylated residues are located in the N-terminus of p32 and the deletion of the first 33 amino acids does not affect RPA’s capacity to bind ssDNA or support replication (Henricksen et al. 1996). However, the phosphorylation of p32 does have an effect on the stability of the RPA trimer as well as its binding to chromatin (Treuner et al. 1999). DNA damage or inhibition of replication also induces the phosphorylation of the 70 kDa subunit of RPA in S. cerevisiae. This phosphorylation is Mec1-dependent, and neither yeast nor human RPA p70 is a substrate for DNA-PK in vitro (Brush & Kelly 2000). Interestingly, the association of RPA on pre-RC is positively regulated by Cdks and Cdc7-Dbf4 and negatively by Rad53 in S. cerevisiae (Tanaka & Nasmyth 1998). Whether RPA is a substrate for the latter two kinases is not known.

2.3.5 Proliferating cell nuclear antigen

The proliferating cell nuclear antigen (PCNA) is a processivity factor for pol δ and pol ε (Prelich et al. 1987, Maga & Hübscher 1995). Beyond this role it has several other functions in DNA replication, repair and recombination, such as recruitment of other 41 proteins in these processes (reviewed in Kelman & Hurwitz 1998). PCNA is a homotrimer of 29 kDa subunits and it is conserved between eukaryotic and archaeal domains. It forms a ring-like ‘clamp’ structure around DNA (Krishna et al. 1994). This structure is very similar to other ‘sliding clamps’ like the E. coli DNA pol III β subunit or T4 phage gp45 that are functionally analogous, but evolutionarily unrelated (Kong et al. 1992, Moarefi et al. 2000). Moreover there are other potential clamp-forming PCNA paralogs in eukaryotes like Rad1-Rad9-Hus1 (Thelen et al. 1999). PCNA is generally used as a proliferation marker. It is expressed in very low amounts in quiescent cells, but the expression is up-regulated when cells are induced to proliferate (Almendral et al. 1987). In cycling cells, PCNA expression fluctuates modestly (2-3-fold) being highest in S phase. However, chromatin binding is largely specific for S phase (Morris & Mathews 1989) and this correlates with the formation of nuclear PCNA foci that represent the sites of ongoing DNA replication (Bravo & Macdonald-Bravo 1987). The phosphorylation of this chromatin bound form of PCNA has also been reported (Prosperi et al. 1993), but has not been repeated by others. This question should be re- examined, especially because PCNA associates with several Cyclin-Cdks, some of which migrate similarly to PCNA in SDS-PAGE (Xiong et al. 1992, Zhang et al. 1993). PCNA is not phosphorylated by an associated kinase in vitro (Xion et al. 1992), but can recruit several proteins like RFC and ligase I for phosphorylation by Cdks (Koundrioukoff et al. 2000). PCNA interacts with many proteins involved in DNA replication and in other processes (Zvi & Kelman 1998). A number of proteins including pol δ and pol ε, Cdk inhibitor p21, excision nuclease XPG, the large subunit of RFC, DNA (cytosine-5) methyl transferase (MCMT), ligase I and Fen-1 have a conserved Q-x-x-[I/L/M]-x-x-[F/H/L]- [F/Y] motif also called the “PCNA-interacting-protein”-box or PIP-box (Jonsson et al. 1998, Warbrick 2000, Gomes & Burgers 2000). The crystal structure of the C-terminus of p21 bound on PCNA shows that the PIP-box motif binds deep in the hydrophobic pocket in PCNA that is surrounded by interdomain collector loop and the C-terminus of a PCNA monomer (Gulbis et al. 1996). This pocket is by no means the only interaction surface with other proteins as is evident from two illustrative examples: Fen-1 nuclease has a PIP-box mediated stable interaction with PCNA, but also a more transient interaction that regulates its activity. This has been shown from studies in which the S. cerevisiae PCNA mutant pcna-79 with a mutated interdomain collector loop fails to form a stable complex with Fen-1, but is still able to stimulate Fen-1 nuclease activity. Exactly the opposite is true for the pcna-90 mutant, where mutations localize on the C-terminus of PCNA (Gomes & Burgers 2000). The second example concerns pol ε, which seems to have complex interactions with PCNA. The effect of PCNA on the polymerase activity of the human pol ε in vitro is apparent only in the presence of RFC and RPA (Lee et al. 1991). However, when Maga and co-workers used calf thymus pol ε and linear templates, they found that PCNA stimulates pol ε both in primer binding and in DNA synthesis. The mutation that was introduced on the back side of PCNA significantly reduced the primer binding only, whereas the PCNA mutants with an altered hydrophobic pocket showed defects in both activities. Thus, it is possible that pol ε may interact with both sides of the PCNA ring (Maga et al. 1999). These results also suggest that pol ε may interact with PCNA via a PIP-box motif (human pol ε aa 1180-1187, S. cerevisiae pol ε aa 1193-1200), 42 which is obviously better exposed in the truncated pol ε from calf thymus than in the full- length form from human cells. The PIP-box mediated interaction with PCNA has been linked to the sub-nuclear targeting to replication foci with ligase I and MCMT (Cardoso et al. 1997, Rossi et al. 1999, Chuang et al. 1997). Ligase I is phosphorylated in vivo at Ser66, which is located near the PIP-box. This serine residue is transiently dephosphorylated in the G1 phase that allows the binding to PCNA (Rossi et al. 1999). Phosphorylation in S and G2 is essential for ligase activity (Prigent et al. 1992). This is a beautiful example of how protein-protein interactions, sub-cellular localization and enzyme activity can be regulated by (de) phosphorylation.

2.3.6 Replication factor C

Replication factor C (RFC) is a DNA dependent ATPase, a “clamp loader” that is required for PCNA loading in SV40 DNA replication in vitro. It consist of five subunits, 140, 40, 38, 37 and 36 kDa, as determined for the human protein in SDS-PAGE. Each of these subunits is essential for viability in yeast (Cullmann et al. 1995). The large subunit (p140 encoded by RFC1 in S. cerevisiae) binds to DNA, the 37 kDa subunit binds specifically to primer ends, whereas the smallest 36 kDa subunit has ATPase activity. In addition to association with p140, the four small subunits are also found in other complexes. S. cerevisiae checkpoint protein Rad24 as well as a complex of three cohesion factors Ctf18, Ctf8 and Dcc1 can associate with p40, p38, p37 and p36. It has been proposed that these alternative complexes can function as “clamp loaders” in DNA damage checkpoints and in the establishment of sister chromatin cohesion, respectively (Rauen et al. 2000, Mayer et al. 2001). During the lagging strand synthesis, RFC loads PCNA onto DNA in an ATP dependent manner in a reaction where it recognizes and binds primer end, displaces pol α, assembles PCNA on DNA and finally allows pol δ to bind PCNA (Yuzhakov et al. 1999). Besides its function in DNA replication, RFC has also been implicated in DNA repair (McAlear et al. 1996) and cell cycle checkpoint control (Sugimoto et al. 1996, 1997, Shimomura 1998, Noskov et al. 1998, Kim & Brill 2001, Krause et al. 2001). The checkpoint functions are linked to three small subunits (p40, p38 and p37 encoded by S. cerevisiae genes RCF4, RCF5 and RCF2 respectively) and at the moment it is unclear if these phenotypes are specific for RFC1, Rad24 or the Ctf18-Ctf8-Dcc1 complexes. The expression of RFC seems similar to that of many other replication proteins. The steady-state mRNA for p140 is highest in actively proliferating tissues and is up- regulated at the G1/S phase (Luckow et al. 1994, Ishida et al. 2001). However, when the expression of the p38 and p37 subunits was studied in yeast, they showed little variation during the cell cycle in contrast to many other replication proteins in S. cerevisiae (Noskov et al. 1994, Li & Burgers 1994). The PCNA binding domain of p140 is phosphorylated by calmodulin kinase II in vitro. This phosphorylation takes place only with soluble RFC that is not bound by PCNA or DNA, and thereafter prevents any association with PCNA and inhibits PCNA dependent DNA synthesis (Maga et al. 1997). 43

The same domain undergoes phosphorylation by PCNA associated Cdk2 in vitro (Koundrioukoff et al. 2000).

2.3.7 Rad1-Rad9-Hus1-Rad17

S. pombe Rad1, Rad9, Hus1 and Rad17 as well as their S. cerevisiae homologues Rad17, Ddc1, Mec3 and Rad24 are implicated in the DNA damage checkpoint mainly in the G1 and in G2 phases (Huberman, Weinert et al. 1994, Pellicioli et al. 1999). Rad1, Rad9 and Hus1 are believed to form a PCNA like ring structure and reconstitution with recombinant proteins shows that this complex is a heterotrimer where the N-terminus of Rad9 interacts with Rad1, the N-terminus of Rad1 with Hus1 and the N-terminus of Hus1 with the C-terminus of Rad9 (Burtelow et al. 2001). Both Rad1 and Rad9 have intrinsic 3’-5’ exonuclease activity (Parker et al. 1998, Bessho & Sancar 2000). Rad17 interacts with Rad1, Rad9 and Hus1 in a manner that is similar to the interaction between RFC and PCNA (Rauen et al. 2000) and this association is dependent on phosphorylation of Rad17 by ATM or ATR (Bao et al. 2001). Rad1, Rad9 and Hus1 are also phosphorylated after damage (Burtelow et al. 2000, St Onge et al. 1999, Kostrub et al. 1998). The expression of human Rad1 and Rad17 are elevated in cancer cell lines and in the testis (Parker et al. 1998a,b).

2.3.8 BRCA1

BRCA1 (BReast CAncer susceptibility gene 1, reviewed in Wang Q et al. 2000, Zheng et al. 2000, Scully & Livingston 2000) is a tumour suppressor that is the gene most often mutated in hereditary breast and ovarian cancer. Despite being a tumour suppressor, BRCA1 is essential for proliferation, and brca1 knockout mice die between embryonic days 6.5 and 13.5 (Gowen et al. 1996, Hakem et al. 1996, Liu et al. 1997). BRCA1 contains a ‘RING’ domain in the N-terminus and two copies of the BRCT domains (BRCA1 C-Terminus like domain), which mediate interactions to numerous proteins. BRCA1 is not conserved with lower eukaryotes, although S. cerevisiae Rad9 and S. pombe Crb2 share some limited sequence homology and partially analogous functions. BRCA1 is involved in the regulation of transcription and chromatin remodelling. It has an intrinsic transactivation function in the C-terminus and also interacts with the RNA polymerase II holoenzyme via RNA helicase A, the histone deacetylase complex, the CBP/p300 histone acetylase complex, transcription repressor CtIP, polyadenylation factor Cst50 via the BARD1 protein and tumour suppressors pRb and p53 (Anderson et al. 1998, Yarden & Brody 1999, Pao et al. 2000, Li et al. 1999, Kleiman & Manley 1999, Ouchi et al. 1998). It can induce the expression of the Cdk inhibitor p21 and the repair protein Gadd45 (Somasundaram et al. 1997, Harkin et al. 1999). BRCA1 is further implicated in homologous recombination dependent double-strand break repair, in transcription-coupled repair, chromosome segregation, cell cycle control and apoptosis. 44

BRCA1 interacts with BRCA2, Rad51 and the Rad50-Mre11-Nbs1 complex (Zhong et al. 1999). BRCA1 and Rad50-Mre11-Nbs1 are part of a larger complex of damage response associated factors including ATM kinase, the mismatch repair proteins MSH2, MSH6, MLH1 and BLM helicase (Wang Y et al. 2000). BRCA1 is expressed at a very low level in quiescent cells. When these cells are induced to proliferate, BRCA1 expression is up-regulated and in cycling cells reaches its highest level at the G1/S phase. Similar changes occur in cell cycle specific phosphorylation levels (Vaughn et al. 1996, Ruffner & Verma 1997). Parallel to the increased expression and phosphorylation, a proportion of BRCA1 localizes in S phase in nuclear foci that are distinct from the sites of ongoing DNA replication. Strikingly, both the block of DNA replication and DNA damage induce a hyperphosphorylation of BRCA1 that occurs simultaneously with sub-nuclear re-localization of BRCA1 into PCNA containing foci (Scully et al. 1997). Recently, it has been shown that after treatment with ionising radiation this re-distribution of BRCA1 is preceded by phosphorylation of the histone variant H2AX and p53 binding protein 1 (53BP1) and their accumulation into the same foci (Paull et al. 2000, Rappold et al. 2001). These ionising radiation induced foci (IRIF) contain also Rad50-Mre11-Nbs1 or Rad51 but never both (Zhong et al. 1999). Specifically, BRCA1 has been shown to be a substrate for Cdk2, ATM, ATR and Chk2 both in vitro and in vivo (Ruffner et al. 1999, Tibbetts et al. 2000, Gatei et al. 2001, Lee et al. 2000).

2.3.9 E2F and pocket protein families

E2F transcription factors (reviewed in Ohtani 1999, Black & Azizkhna-Clifford 1999) have an essential role in the induction of S phase in animal cells. They are heterodimeric proteins consisting of an E2F subunit (E2F1-6) and a DP subunit (DP1-3). E2F subunits share a homologous structure with an N-terminal DNA-binding domain, a dimerization domain in the central region of the polypeptide and a transactivation domain in the C- terminus that also contains a docking site for retinoblastoma family proteins. The retinoblastoma protein, pRb, is a tumour suppressor and a member of the so called “pocket protein” family consisting of pRb, p107 and p130 (reviewed in Mulligan & Jacks 1998). E2F6 is an exception to this family architecture as it lacks the transactivation domain. E2F transcription factors bind their recognition sequence 5’-TTT(G/C)(G/C) CGC-3’ by major groove contacts between bases in the GC-rich core and both E2F and DP subunits, while the TTT-extension is bound only by the E2F subunit (Zheng et al. 1999). Binding induces a DNA bend and this is a typical feature for all members in E2F family (Cress & Nevins 1996). In addition to a consensus binding sequence, several variant sequences are known to bind E2F and the sequence specificity is dependent on the subunit composition (Tao et al. 1997). E2F binding sites are found in promoters of several replication linked genes like dihydrofolate reductase, thymidine kinase, thymidylate synthatase and DNA polymerase α and in promoters for cell cycle regulators like cyclins, Cdc25, Orcs, Mcms, Cdc6 and Cdc45 (Ohtani 1999, Ishida et al. 2001). The function of E2Fs can be both stimulatory and repressive. The hypophosphorylated pocket 45 proteins bind E2Fs in G0 and G1. The phosphorylation of pocket proteins by Cyclin D/Cdk4 (or 6) and Cyclin E/Cdk2 is followed by the dissociation of E2F. Free E2F can activate transcription via several mechanisms. It can directly bind the basal transcription factors TFIIH and TBP, as well as recruit the CBP/p300 histone acetylase (Fry et al. 1999). Furthermore, E2F1 has been shown to physically interact with Sp1 transcription factor and synergistically activate promoters for dihydrofolate reductase and thymidine kinase (Lin et al. 1996, Karlseder et al. 1996). E2F associates with and becomes phosphorylated by Cyclin A/Cdk2 in S phase, which leads to its inactivation (Dynlacht et al. 1994). The pocket proteins not only sequester E2F, but also form functional repressor complexes and are often associated with the histone deacetylase 1 (HDAC1) (Ferreira et al. 1998). E2F4 and E2F5 bind preferentially p107 and p130 and are bound to promoters usually during G0 and G1, whereas E2F1, E2F2 and E2F3 bind preferentially pRb and are promoter bound most often during late G1 (Takahashi et al. 2000, Wells et al. 2000). However, the most abundant E2F, E2F4 is expressed throughout the cell cycle and is also bound to some promoters in S phase (Wells et al. 2000). In addition to their role as transcriptional regulators, E2Fs and pocket proteins seem to have a more direct role in the initiation of DNA replication, as they are found to interact with the ORC and localize into the early replication foci (Bosco et al. 2000, Kennedy et al. 2000). E2F also interacts with Mre11-Rad50-Nbs1 protein complex and co-localizes with it into the origins of DNA replication (Maser et al. 2001). The function of E2F in the initiation of DNA replication is not known but may relate to chromatin remodelling. 3 Aims of the present work

The DNA polymerases are central components in DNA replication, recombination, repair and cell cycle control. The role of pol ε has been enigmatic and elucidation of its function is of importance. To address this question, the expression and phosphorylation of pol ε were studied. Parallel to this, a search for proteins that interact with pol ε was conducted.

The specific aims of the present work were as follows,

1) To examine, how the expression of human pol ε is dependent on cell proliferation and the cell cycle.

2) To analyse the promoter of the gene for the human POLE2.

3) The search for proteins interacting with pol ε.

4) To characterize the phosphorylation of the catalytic subunit of human pol ε. 4 Materials and methods

4.1 Ribonuclease protection assay (I, II, III)

Total RNA was isolated either by a guanidium thiocyanate phenol-CHCl3-method (Chomczynski & Sacchi) (I) or by using the Trizol®LS reagent (GibcoBRL) (II &III). Total RNA (5 - 10 µg) was hybridised with a 32P-labelled RNA probe transcribed in vitro and purified in urea-PAGE. Digestion with RNase T1 and RNase A was performed using an RPA II™ ribonuclease protection assay kit (Ambion). RNA transcription in vitro was carried out as described (Tuusa et al. 1995, Huang et al. 2001, Mäkiniemi et al. 2001).

4.2 Immunoprecipitation and Western-analysis (I, III, IV)

Antibodies K19 and K20 (against human Pol ε, aa 473 - 657, GenBank accession no. 3192938), K27 (against human Pol ε, aa 1 – 203, GenBank accession no. 3192938), anti- TopBP1.1 (human TopBP1, aa 792 – 1003, GenBank accession no. D87448) and anti- TopBP1.2 (human TopBP1, aa 1023 – 1167, GenBank accession no. D87448) were raised in rabbits and purified by protein A-Sepharose CL4B (Amersham Pharmacia Biotech). Monoclonal antibodies 93H3B, 93G1A and 93E24A against Pol ε were described earlier (Uitto et al. 1995). Anti-PCNA (PC10), anti-tubulin, anti-actin and anti-FLAG antibodies were from Boehringer Mannheim, Clontech, Roche and Sigma, respectively. Cultivated cells (106 – 107) were washed with PBS and suspended in lysis buffer [100 mM Tris-Cl, 100 mM NaCl, 10 % glycerol, 0.1 % Nonidet P-40 and Complete™ protease inhibitors (Roche)]. Cells were broken with 2 x 15 s sonication (7.5 µm amplitude, Soniprep 150, SANYO) on ice and the extract was clarified by centrifugation. For pre- clearing, 4 µl of pre-immune rabbit serum was added per 1 ml of cell extract and, after incubation for 1 h, pre-immune IgG binding proteins were collected with protein A- Sepharose. Immunoprecipitation with 10 µg of specific antibody was performed in ice for more than 1h and immunocomplexes were collected with protein A-Sepharose / protein 48

G-Agarose (Sigma) and washed twice with lysis buffer, once with washing buffer II (50 mM Tris-Cl pH 7.5, 500 mM NaCl, 0.5 % Nonidet P-40) and once with washing buffer I (50 mM Tris-Cl pH 7.5, 150 mM NaCl, 0.5 % Nonidet P-40). Immunoprecipitation of denatured proteins is described in chapter 4.12 and the immunoprecipitation of cross- linked proteins in chapter 4.14. For Western analysis, proteins were separated in SDS-PAGE and blotted onto an Immobilon™ P PVDF membrane (Millipore). Non-fat milk (5%) and Tween-20 (0.05%) were used for blocking of non-specific binding. After incubation with specific antibody, the blots were washed with TBST (50 mM Tris-Cl pH 7.5,150 mM NaCl, 0.05% Tween- 20), further incubated with anti-mouse IgG alkaline phosphatase conjugate (Schleicher&Schulell 1:10000) or with anti-rabbit IgG alkaline phosphatase conjugate (Biorad 1:5000) and finally with the colour developing reagents 5-bromo-4-chloro-3- indolyl phosphate and nitro blue tetrazolium. Alternatively, the membrane was blocked with SuperBlock® blocking buffer (Pierce) and anti-mouse IgG HRP-conjugate (Pierce 1:10000) was used as the secondary antibody. The chemiluminescent signal was acquired by incubation in a mixture of Luminol and Stable peroxidase Solution (Pierce) and detected by Hyperfilm ECL (Amersham-Pharmacia Biotech).

4.3 Serum stimulation of quiescent cells (I, II, III)

IMR-90 fibroblasts were brought to a quiescent state (G0) by serum depletion (0.25 %) for 96 h. The cells were induced to proliferate by the addition of serum (10%). MCF-7 cells were serum starved 24 h, then the serum concentration was raised to 15 %. DNA- synthesis was monitored by measuring [3H]-thymidine incorporation from parallel cultures as described (Pearson et al. 1991).

4.4 Cell cycle specific synchronization and fractionation (I, II, III, IV)

Cells were synchronized at the G1/S border with a double thymidine block (exposure to 2 mM thymidine for 18 h, release into the thymidine-free medium for 10 h, a second exposure to 2 mM thymidine for 18 h) or at the G2/M border with a nocodazole block (100 nM nocodazole for 15 h) or fractionated into cell cycle specific fractions using counterflow centrifugal elutriation (Beckman JE-5.0 elutriator rotor, constant rotor speed, increasing flow rate). The cell cycle stage was monitored by measuring [3H]-thymidine incorporation or by flow cytometry (Becton Dickinson).

49

4.5 Promoter activity analysis (II)

The 1.1 kb genomic fragment containing the 5’-flanking region and exon 1 of POLE2 was subcloned into the pGL3-Basic luciferase reporter vector and subsequently used as a template for a series of deletion and site-directed mutation constructs. The deletions and point mutations were introduced by PCR. The promoter construct (2 µg) was transfected together with the pSV-β-galactosidase vector (0.5 µg; internal control) into Hela CCL2 cells using the FuGENE™ transfection reagent. The luciferase activity was normalized with β-galactosidase activity.

4.6 Electrophoretic mobility shift assay (EMSA) (II)

Two 32P labelled restriction fragments representing nt +101 - +175 and nt +101 - +254 were used as probes. Nuclear extracts were prepared from asynchronous and synchronized Hela S3 cells and from asynchronous, serum starved and serum stimulated IMR-90 fibroblasts as described (Lee et al. 1994). Nuclear extract (3 µg) in 10 mM Tris- Cl pH 7.5, 50 mM NaCl, 5 mM MgCl2, 0.5 mM DTT, 4 % glycerol was pre-incubated with 0.5 µg poly(dI-dC)-poly(dI-dC) at room temperature for 5 min. Radiolabelled probe was added, the mixtures incubated 30 min and the reactions stopped by the addition of loading buffer. The competition assays were performed by pre-incubation of extracts with double stranded unlabelled oligonucleotides for 15 min. The antibodies used for the supershift assays were E2F1 (C-20), E2F2 (C-20), E2F3 (N-20), E2F4 (C-20), E2F5 (E- 19), NF-1 (N-20), p107 (SD9), p130 (C-20), pRb (C-15) and Sp1 (1C6). Antibody (0.5- 2.0 µg) was added to the reaction after the probe and incubated at 4 °C for 2 h. DNA- protein complexes were analysed in a 4 % polyacrylamide gel in 0.25 x TBE buffer.

4.7 Deoxyribonuclease I footprinting analysis (II)

DNaseI footprinting was done using a SureTrack Footprinting Kit (Amersham Pharmacia Biotech). The genomic clone containing nucleotides from +101 to +254 was digested either with XhoI/NcoI or MluI/HindIII. Phosphatase treated and gel purified fragments were 5’-end-labelled with γ-[32P]-ATP and T4 polynucleotide kinase. Single-end labelled probes were prepared by digesting with HindIII or XhoI, followed by gel-purification. 20000 cpm of probe was incubated with nuclear extract prepared from asynchronous HeLa S3 cells. BSA was used as a negative control. A molar excess of cold probe was used for competition. The amount of DNaseI was optimised and 1 to 4 units was used per reaction. The G+A sequence ladder was prepared by the Maxam-Gilbert method (Bencini et al. 1984). The DNA fragments were analysed on a 7 M urea-8 % acryl amide- bisacrylamide gel.

50

4.8 DNA replication assay in isolated nuclei (III)

Permeable nuclei were prepared in lysolecithin containing buffer using a Dounce homogenizor as described (Stoeber et al. 1998). The DNA replication assay in vitro was done with triplicate samples as described (Pospiech et al. 1999).

4.9 Immunostaining (III)

For fixation, MCF-7 cells or IMR-90 fibroblasts were incubated 10 min in ice cold methanol or alternatively in 3 % paraformaldehyde followed by permeabilization either in 0.2 % TritonX-100 in PBS at room temperature for 10 min or in ice cold methanol for 2 min. The incubation with primary and secondary antibody was carried out in PBS with 0.2 % gelatin (Sigma) at 37˚C for 30 min. DNA was stained with bis-benzimide (Hoechst 33258, Sigma) at room temperature for 5 min. For detection of DNA synthesis de novo, cells were pulse-labelled (3 min) with 0.1 mM BrdU at 37˚C, fixed with 3 % paraformaldehyde for 10 min, denatured with 4 M HCl and immunostained with FITC- conjugated anti-BrdU antibody. Sample slides were mounted with IMMU-MOUNT (Shandon) and examined with a fluorescence microscope (Leitz or Olympus).

4.10 Ecdysone-inducible expression system (III, IV)

TopBP1, Rad9 and fragments of the catalytic subunit of pol ε were over-expressed using an Ecdysone-inducible expression system (Invitrogen). The cDNA for TopBP1 was subcloned into the pIND vector. The cDNA for Rad9 and cDNAs encoding aa 269-503 (EXO) and aa 2124-2286 (C-TERM) of pol ε were subcloned into the modified pIND vector with an SV40 T-antigen like nuclear localization signal and FLAG-epitope 5’-in- frame. The linearized constructs were transfected into EcR-293 cells (Invitrogen) with the FuGENE™ 6 transfection reagent (Roche). Cells were cultivated under selection for two weeks and G418 resistant colonies were recovered, expanded and tested for inducible expression by Western-analysis. Wild type and mutant Pol ε fragments were also used for transient transfection. Expression was induced with 2 µM ponasterone A for 24 h.

4.11 Site directed mutagenesis

A Transformer™ Site-Directed Mutagesis Kit (Clontech) was used to produce the pol ε EXO fragment Thr323Ala mutant. EXO fragment encoding aa 269-503 of pol ε in a modified pIND vector was used as a template. 5’-CCTTCGGTCCTCCCAT CGTT GTCAG-3’ primer was used in selection to destroy a unique PvuI restriction site in the 51 vector. The second oligonucleotide 5’-GAAGATTTTGAGTTCGCCCCCAAGCCAG-3’ carried the Thr323Ala mutation encoding an A -> G transition. Electroporation was used for the first transformation of mutant DNA into electrocompetent BMH71-18MutS cells. The mutant clone was verified by sequencing.

4.12 Metabolic labelling with 32P (III, IV)

EcR-293 cells (1-2 x 106) or Hela S3 cells (1 x 107) were washed once with phosphate- 32 free medium and labelled with 1 - 4 mCi of Pi in 2 ml of phosphate-free medium supplemented with fetal calf serum (10 %, dialyzed against 0.9 % NaCl) for 4 h. The cells were washed twice with PBS and lysed in 200 µl of SDS-lysis buffer (50 mM Tris-Cl pH 7.5, 2 % SDS, 70 mM β-mercaptoethanol) and incubated in a boiling water bath for 10 min. The genomic DNA was sheared by passing the lysate ten times through a 22G needle. For immunoprecipitation, the lysate was diluted 20-fold with washing buffer I (50 mM Tris-Cl pH 7.5, 150 mM NaCl, 0.5 % Nonidet P-40). Pre-immune rabbit serum (10 µl containing about 100 µg IgG) was added and after incubation in ice for more than 1 h, the samples were further incubated for 1 hr with immobilised protein A, either Staphylococcus aureus whole cells (Sigma) or protein A-Sepharose. The insoluble material together with pre-immune IgG binding proteins were removed by centrifugation. Pol ε was immunoprecipitated either with 10 µg of 93H3B, 93G1A or K19 antibody, TopBP1 with 10 µg of anti-TopBP1.2 antibody and pol ε fragments with 5 µg of anti- FLAG M2 antibody. Immunocomplexes were collected by protein A-Sepharose/ protein G-agarose. The pellets were washed three times with washing buffer II (50 mM Tris-Cl pH 7.5, 500 mM NaCl, 0.5 % Nonidet P-40), once with washing buffer I and suspended in SDS-PAGE sample buffer. Alternatively, cells were washed twice with PBS, suspended in 200 µl of hypotonic lysis buffer (50 mM Tris-Cl pH 7.5, 0.5 % Nonidet P-40 + protease inhibitors) and allowed to swell 5 min. Cells and genomic DNA were broken by passing the solution through a 27G needle 15 times. The NaCl concentration was adjusted to 100 mM and 300 µl lysis buffer [100 mM Tris-Cl, 100 mM NaCl, 10 % glycerol, 0.1 % Nonidet P-40 and Complete™ protease inhibitors (Roche)] was added. Pol ε was immunoprecipitated with 10 µg of K27 antibody as described in 4.2.

4.13 Phosphoamino acid analysis and phosphopeptide mapping (III, IV)

Metabolically 32P-labelled and immunoprecipitated proteins were purified in 6.25 %/12.5 % SDS-PAGE and blotted onto a PVDF-membrane. The radioactivity was visualized 52 either by a Phosphorimager (Molecular Dynamics) or by autoradiography (Kodak X- Omat or Kodak Biomax MS). Phosphoamino acid analysis and phosphopeptide mapping were performed essentially as described (Van der Geer et al. 1994). For phosphoamino acid analysis, metabolically labelled protein was cut from the membrane, washed several times with water and hydrolyzed for 60 min at 110°C in 6 M HCl. The acid was evaporated and sample was dissolved in 10 µl of TLE pH 1.9 buffer (2.2% formic acid, 7.8 % acetic acid) and mixed with 1 µg cold phosphoserine, phosphothreonine and phosphotyrosine standards. Phosphoamino acids were separated on cellulose-TLC plates (Merck) in two dimensions (1st dimension: 1.5 kV, 20 min, TLE pH 1.9 buffer; 2nd dimension: 1.3 kV, 16 min, TLE pH 3.5 buffer: 5 % acetic acid, 0.5 % pyridine). The location of the phosphoamino acids was mapped by ninhydrin staining and the presence of 32P-amino acids by autoradiography. For phosphopeptide mapping the piece of membrane with pol ε was cut from the membrane, and then reduced with 5 mM DTT and alkylated with 100 mM iodoacetamide to prevent spot doubling. Thereafter the membrane was blocked with polyvinylpyrrolidone in 100 mM acetic acid for 30 min at 37°C and digested with 2 x 10 µg trypsin (Promega, seq. grade) at 30°C overnight. After evaporation and re- solubilisation in 10 µl of TLE pH 1.9 buffer, the phosphopeptides were separated on a cellulose TLC plate by electrophoresis (1.0 kV, 45 min, TLE pH 1.9 buffer) for the first dimension and by ascending thin-layer chromatography in n-butanol:pyridine:acetic acid:water (75:50:15:60) for the second dimension. 32P-phosphopeptides were visualized by autoradiography.

4.14 Cross-linking with formaldehyde (IV)

EcR-293 cells cultivated on 75 cm2 plates were treated with 1 % formaldehyde in serum- free medium at 37˚C for 5 min. The cells were washed with PBS, incubated in PBS containing 100 mM glycine for 5 min and washed once more with PBS. The cells were scraped out, suspended in 900 µl of lysis buffer (10 mM Tris-Cl, 1 mM EDTA, 3 mM MgCl2 pH 8) and broken by sonication (3 x 10 s, 10 µm amplitude, Soniprep 150, SANYO). DNA was digested by incubating with 4 U DNaseI (Promega) at room temperature for 15 min. For denaturation of the proteins, 50 µl 20 % SDS, 40 µl 1 M Tris-Cl pH 7.5, 20 µl 0.5 M EDTA and 2.5 µl β-mercaptoethanol was added and incubated a further 60 min at room temperature. The lysate was clarified by centrifugation, diluted 10-fold with washing buffer I (50 mM Tris-Cl pH 7.5, 150 mM NaCl, 0.5 % Nonidet P-40), pre-cleared with pre-immune serum and protein A-Sepharose and used for immunoprecipitation with the antibodies indicated. Immunocomplexes were harvested and washed as described in 4.12 and suspended in SDS-PAGE sample buffer. The cross-links were reversed by incubation at +65˚C for 4 h and the samples were analysed in SDS-PAGE. 5 Results

5.1 The expression of the human DNA polymerase ε is typical for replicative DNA polymerases (I, II)

For analysing gene expression during the G0/G1 transition, logarithmically growing IMR- 90 fibroblasts were cultivated to sub-confluent density and deprived of serum for 96 h. The quiescent cells were induced to re-enter the cell cycle by the addition of serum. Total RNA was isolated at 4 h intervals and analysed by an RNase protection assay. Similarly total RNA was prepared from Hela S3 cells synchronized by double thymidine or nocodazole block or from cells fractionated by counter flow centrifugal elutriation. The relative levels of steady-state mRNA for the A and B subunit were similar, showing about a 5-fold up-regulation preceding DNA synthesis and reaching their highest levels after 18 – 22 h of serum stimulation. In contrast to the G0/G1 transition, both message levels were nearly constant during a mitotic cell cycle with less than a two-fold peak at G1/S border. These profiles were similar to that of pol α, but not to that of pol β. These pols were used as representatives of replicative and non-replicative polymerases. The amount of the pol ε catalytic subunit was analysed from mouse tissue samples and from cultivated human fibroblasts by immunostaining. The immunoreactive protein with a molecular weight > 200 kDa was present in all mouse tissues examined, but was most abundant in thymus extract. The immunoreactive pol ε was not detectable from quiescent fibroblasts but clearly observed from serum-stimulated cells after immunoprecipitation (data not shown). The amount of immunoreactive pol ε was nearly constant during the cell cycle with a minor fluctuation resembling that of the steady-state mRNA.

54

5.2 The gene for human DNA polymerase ε B subunit is regulated by E2F and Sp1 transcription factors (II)

The 5’-UTR and 5’-upstream flanking region of POLE2 encoding the B subunit of pol ε were cloned and the promoter region was characterized by standard promoter analysis methods. The minimal core promoter was mapped between nucleotides +101 to +175, and is located downstream, surprisingly, from the transcription initiation site(s). The promoter activity assay revealed the presence of down-regulation elements between nt – 360 to +101 and +175 to +254, the latter being immediately downstream from the core promoter. The identity and function of the transcription factors binding to the core promoter and downstream region were examined by promoter activity, electrophoretic mobility shift (EMSA) and DNaseI footprinting assays. An Sp1 binding element in the core promoter was essential for promoter activity and an Sp1 transcription factor was found to be bound on the core promoter in vitro both in extracts from quiescent cells as well as from proliferating cells throughout the cell cycle. The core promoter E2F/NF-1 consensus binding site was essential for full promoter activity and for serum response and it appeared that the mitogenic activation was due to the alleviation of G0-specific repression. The core promoter was bound by p107, E2F2 and E2F4 but not by NF-1 in vitro. When a longer probe consisting of the core promoter and a downstream region were used in EMSA, the binding of Sp1, p107, pRb, E2F1 and NF-1, but not of E2F2 or E2F4 was detected. The quality of the observed DNA-protein complexes were unchanged during the cell cycle and G0/G1 transition, but the quantities of these complexes were reduced when extracts from quiescent cells was used. Notably, the complex supershifted by the anti-p107 antibody was still as abundant as in extracts from cycling cells. DNaseI footprinting analysis showed full protection of the Sp1 element in the core promoter and the E2F elements in the downstream region as well as the protection of the antisense strand of the E2F element in the core promoter.

5.3 DNA polymerase ε interacts with BRCA1 and TopBP1 (III, IV)

In the search for pol ε -interacting proteins, co-immunoprecipitation was used. BRCA1 and TopBP1 were considered as putative pol ε associated proteins. For analysis of these proposed interactions, polyclonal antibodies against pol ε (K27 against pol ε aa 1- 203) and TopBP1 (anti-TopBP1.2 against aa 1023-1167) were raised in rabbits, purified by protein A-Sepharose and tested for immunoprecipitation and for immunostaining. Anti-pol ε and anti-TopBP1 antibodies efficiently immunoprecipitated pol ε and TopBP1 respectively. The rabbit polyclonal antibody Ab-2 against BRCA1 was able to co- immunoprecipitate pol ε from pre-cleared Hela cell extract. Similarly BRCA1 was present in the immunoprecipitate of anti-pol ε antibody K27. Neither of these proteins co- precipitated with matrix alone or with non-related control antibody. 55

Pol ε was also detected in low amounts in a anti-TopBP1 immunoprecipitate from a Hela cell extract. This interaction was confirmed by immunoprecipitating TopBP1 from a cell extract prepared from a stable cell line inducibly over-expressing TopBP1. The anti- TopBP1 antibody neither precipitated purified pol ε devoid of TopBP1, nor co- immunoprecipitated pol ε from denatured cell lysate, excluding the possibility of cross- reaction. To ensure that the interaction was not mediated via DNA, ethidium bromide was included in the immunoprecipitation reaction. This did not abolish the co- immunoprecipitation of pol ε. However, when pol ε was immunoprecipitated by the anti- pol ε antibody K27, co-immunoprecipitation of TopBP1 could not be demonstrated.

5.4 TopBP1 is a phosphoprotein with eight BRCT-domains and is expressed in a proliferation dependent manner (III)

Based on the homology to budding yeast Dpb11 and fission yeast Cut5, a human cDNA KIAA0259 was identified and found to encode a protein with eight BRCT domains. This protein was earlier characterized as Topoisomerase IIβ binding protein or TopBP1 (Yamane & Tsuruo 1997). Closer analysis showed that the pairs of BRCT domains 1-2 and 4-5 were conserved between human TopBP1 and S. cerevisiae Dpb11 (35 % and 27 % similar, 14 % and 11 % identical, respectively) as well as between TopBP1 and S. pombe Cut5 (44 % and 41 % similar, 26 % and 23 % identical, respectively) both having only these four BRCT domains. Obvious orthologs were found in searches of databases for the nematode Caenorhabditis elegans and the fly Drosophila melanogaster with six and seven BRCT domains, respectively. Western analysis from Hela, EcR-293 and IMR-90 cell extracts showed the presence of an immunoreactive polypeptide with a molecular weight of 180 kDa. The steady-state TopBP1 level increased 3-fold when quiescent IMR-90 cells were induced to proliferate. The steady-state mRNA level was measured by an RNase protection assay parallel to quantification of the protein. The message for TopBP1 showed a 1.8-fold increase during the G0/G1 transition with a profile similar to that of the TopBP1 protein. The steady-state mRNA level was also measured from cells that were synchronized by double thymidine or nocodazole block or from cells fractionated by counterflow centrifugal elutriation, and found to show a corresponding increase at the G1/S border. Metabolic labelling was used to determine if TopBP1 is phosphorylated in vivo. EcR- 293 cells were cultivated in multi-well dishes and about 1 x 106 cells were pulse-labelled 32 with 2 mCi of Pi in 2 ml of phosphate-free medium for 4 h. The washed cells were lysed in SDS-lysis buffer and boiled for 10 min for complete denaturation. TopBP1 was immunoprecipitated from the diluted lysate and analysed by SDS-PAGE and blotting. Radioactivity was detected by autoradiography. TopBP1 was found to be phosphorylated and subsequent phosphoamino acid analysis showed phosphorylation of serine and to a lesser extent of threonine residues. Two-dimensional phosphopeptide mapping revealed the presence of at least 10 distinct tryptic phosphopeptides. The phosphopeptide map -2 -1 remained unchanged after exposure to 20 Jm , UV254 nm, 2 mM hydroxyurea or 50 µgml methyl methane sulphonate (MMS). Because EcR-293 cells are resistant for zeozin, a 56 chemical catalysing single and double strand breaks, metabolic labelling was repeated with Hela S3 cells. An identical phosphorylation pattern was observed that was not affected by treatment with 200 µgml-1 zeozin (data not shown).

5.5 TopBP1 is required for chromosomal DNA replication (III)

A system to study mammalian DNA replication in vitro using permeable isolated nuclei and cytoplasmic extract has been developed (Krude et al. 1997) and was used to show the requirement of pol ε for chromosomal DNA replication (Pospiech et al. 1999). This assay was used to examine the role of TopBP1 in DNA replication. The polyclonal antibody anti-TopBP1.1 raised in rabbit against the TopBP1 aa 792-1003 inhibited DNA replication by 44%. This was comparable to the inhibition by anti-pol α antibody SJK- 132-20, whereas pre-immune antisera did not show any inhibition. Pospiech et al. 1999 showed that the specificity of the anti-pol ε antibody could be demonstrated by antigen blocking. When recombinant TopBP1 aa 792-1003 peptide was included in the reaction in stoichiometric excess, DNA replication was inhibited even more (83%) demonstrating independent inhibition by the antigen.

5.6 TopBP1 and BRCA1 co-localize in S phase and are re-localized after replication block or DNA damage into the PCNA containing foci (III)

The sub-cellular localization of TopBP1 was studied by immunofluorescence microscopy. In MCF-7 cells, TopBP1 was found to be mainly nuclear and concentrated in nuclear foci in interphase. Double immunostaining with the anti-PCNA antibody or with the anti- BrdU antibody after BrdU pulse-labelling showed that TopBP1 foci are distinct from sites of ongoing DNA replication. Next, possible co-localization with BRCA1 was studied and indeed, an extensive co-localization was evident. The number of TopBP1 foci was higher than that of BRCA1, but all BRCA1 positive foci were also TopBP1 positive. Because BRCA1 re-localizes to PCNA foci after a replication block or DNA damage, the behaviour of TopBP1 was studied. S phase MCF-7 cells were exposed to hydroxyurea -2 -1 -1 to block replication or treated with 10 Jm UV254 nm, 200 µgml zeozin or 50 µgml MMS. The blocked replication caused nearly complete co-localization of TopBP1 with PCNA together with BRCA1. 17 % of TopBP1 foci co-localized with PCNA after UV and a much smaller, although still statistically significant, amount after MMS treatment. Double strand breaks did not induce the co-localization between TopBP1 and PCNA. Interestingly, although TopBP1 and BRCA1 co-localized in S phase after zeozin treatment, in the G1 phase, zeozin induced TopBP1 foci that were devoid of BRCA1. 57

5.7 TopBP1 interacts physically with checkpoint protein Rad9 (III)

A two-hybrid screen in yeast was utilized in the search for proteins interacting with TopBP1. The BRCT domains 4 and 5 (TopBP1 aa 534-763) were fused to the LexA DNA-binding domain and used as bait in screening a human peripheral lymphocyte cDNA library. One of the interacting clones was found to encode 148 amino acids of the C-terminal part of the Rad9 protein. This interaction was specific for BRCT domains 4 and 5 and no interaction was found between Rad9 and BRCT domains 3 or 6 of TopBP1. Furthermore, the interaction was confirmed by demonstrating the interaction between full-length Rad9-LexA and full-length TopBP1-VP16 fusions. To find out if TopBP1 and Rad9 are complexed in vivo, co-immunoprecipitation was utilized. Because of the lack of anti-Rad9 antibody, a stable EcR-293 cell line inducibly expressing Rad9 with the FLAG-epitope was established. Rad9 was found to co- immunoprecipitate with TopBP1 and vice versa. anti-FLAG antibody was used both for immunoprecipitation and detection of Rad9.

5.8 DNA polymerase ε has at least three immunologically separable forms with distinct phosphorylation and different interacting proteins (IV)

The monoclonal antibodies 93H3B, 93G1A and 93E24A against pol ε were tested for immunoprecipitation. None of these were able to immunoprecipitate native pol ε, but 93G1A, and to a much lesser extent 93H3B and 93E24A, immunoprecipitated pol ε from denatured cell lysate. The specificity of the 93G1A antibody in immunoprecipitation was confirmed by blocking the immunoprecipitation with antigen and by showing that no pol ε precipitated with matrix alone or with a non-related antibody. The metabolic labelling 32 of Hela S3 as well as EcR-293 cells with Pi followed by an immunoprecipitation with 93G1A showed that pol ε is a phosphoprotein. The nature of the phosphorylation was characterized by phosphoamino acid analysis and tryptic phosphopeptide mapping. Pol ε was found to be phosphorylated on serine and threonine residues. Two dimensional analysis of tryptic phosphopeptides on a thin-layer chromatographic plate revealed at least nine phosphopeptides giving an estimate of the number of phosphorylation sites. The phosphorylation of pol ε was studied by metabolic labelling and phosphopeptide mapping during the cell cycle after serum, hydroxyurea, UV, MMS, bleomycin/zeozin and H2O2 -treatments. The de novo phosphorylation in nocodazole synchronized Hela S3 cells showed a moderate fluctuation with a 4-fold increase at the G1/S-border. Immunoblotting with a anti-phosphothreonine antibody showed a similar variation, whereas the amount of 93G1A immunoprecipitated pol ε remained nearly constant throughout the cell cycle. However, tryptic phosphopeptide mapping showed that this fluctuation in phosphorylation was not specific for any particular phosphopeptide. Indeed, the same phosphopeptides with the same relative intensities were present in each time point. DNA damage had no effect on the total phosphorylation of pol ε nor did it 58 change the phosphopeptide map pattern. Finally, inhibitors of MAP-kinases, protein kinase A or protein kinase C did not have any effect on the phosphorylation of pol ε (data not shown). When new antibodies against pol ε became available, they were tested for immunoprecipitation. The polyclonal antibody K27 raised in rabbit against the N- terminal aa 1-203 of pol ε was found to immunoprecipitate native, but not denatured pol ε. In contrast, two other polyclonal antibodies, K19 and K20, raised in rabbit against aa 473-657 of pol ε immunoprecipitated denatured but not native pol ε. The form of pol ε immunoprecipitated by K27 was poorly or not at all phosphorylated. Instead, pol ε immunoprecipitated by K19 or K20 was phosphorylated. The tryptic phosphopeptide map from pol ε immunoprecipitated by the K19 antibody was identical to that of K20 immunoprecipitated pol ε, but different from the phosphopeptide map of pol ε immunoprecipitated by 93G1A with only two apparently common phosphopeptides. As was the case with 93G1A specific phosphorylation, the K19 immunoprecipitable form of pol ε also showed no change in the phosphopeptide map after treatment with UV, MMS or zeozin. For an unknown reason, no de novo phosphorylation of K19 immunoprecipitable pol ε was observed after nocodazole synchronization, although immunoprecipitation of this form of pol ε worked well. Whether cell cycle specific variations in the phosphorylation of the K19 specific form of pol ε exist remains to be studied. These results show that there are at least three distinct forms of pol ε with different levels of phosphorylation. As described earlier, the polyclonal anti-pol ε antibody K27 co-immunoprecipitated BRCA1, but not TopBP1. TopBP1-pol ε interaction was studied in more detail by introducing cross-links between the associated proteins by treating the cells briefly with formaldehyde. To prevent chromatin-mediated carry-over, DNA was digested with DNaseI. The proteins were denatured by SDS and β-mercaptoethanol, which was necessary for efficient immunoprecipitation with the anti-pol ε antibodies 93H3B, 93G1A and K19. The immunoprecipitates were washed and heat-treated to reverse the cross- links. Immunoblotting with the anti-TopBP1 antibody showed TopBP1 in the 93G1A immunoprecipitate. Together with the observation that the anti-pol ε antibody K27 is specific for the hypophosphorylated form of pol ε and is not able to co- immunoprecipitate TopBP1, this result indicates that TopBP1 associates with the hyperphosphorylated form of pol ε. For determination of the phosphorylation sites of pol ε, cDNA fragments spanning the coding region of pol ε catalytic subunit cDNA were subcloned into a modified pIND vector and used for transient or stable transfection into EcR-293 cells. Only the constructs encoding aa 269-503 (EXO) and aa 2124-2286 (C-TERM) were successfully expressed after ponasterone A induction. Stable cell lines inducibly expressing these fragments were 32 established and used for metabolic labelling with Pi. Autoradiographic analysis showed phosphorylation of the EXO fragment, but not of the C-terminal fragment. This experiment was repeated by transient transfection of the wild type EXO construct and of the EXO mutant, where a putative Cdk phosphorylation site Thr 323 is changed to alanine. Both the wild type and mutant fragments were expressed, but only the wild type fragment was phosphorylated (Figure 6). Tryptic digestion of the labelled EXO fragment produced, however, a phosphopeptide that was distinct from all phosphopeptides from 93G1A or K19 immunoprecipitated endogenous pol ε (data not shown). 59

Finally, the effects of dephosphorylation on the polymerase activity of pol ε in vitro were examined. Purified pol ε was pre-incubated with soluble λ-protein phosphatase or with matrix-linked alkaline phosphatase, and polymerase activity was measured with a poly(dA)-oligo(dT) template-primer. A reduction in polymerase activity up to 27% with λ-protein phosphatase treated pol ε was observed compared to non-treated control. A higher reduction was evident with alkaline phosphatase treated pol ε, but in this assay, substrate hydrolysis cannot be completely excluded.

Fig. 6. A fragment of pol ε becomes phosphorylated at Thr323 in vivo. EXO-fragment (aa 269-503 of pol ε) was ectopically expressed as a FLAG-fusion in EcR-293 cells. The expression was induced with 2 µM ponasterone A for 24 h followed by metabolic labelling with 1 mCi of 32 Pi for 4 h. The fusion protein was immunoprecipitated by anti-FLAG antibody M2 and analysed in 12.5 % SDS-PAGE. The autoradiographic analysis of blotted sample shows phosphorylation on wild type, but not on the Thr323Ala mutant. 6 Discussion

6.1 The proliferation dependent expression of pol ε is similar to other replicative DNA polymerases with some unique features (I, II, IV)

Cell cycle dependent transcriptional regulation is different in the yeast S. cerevisiae and in higher eukaryotes. Most of yeast genes, whose gene products are needed during S phase such as genes for DNA polymerases, show periodic expression with the highest level of steady-state mRNA in the G1/S phase. This variation is mainly due to fluctuations in transcription that are regulated by two transcription factors, MBF and SBF (Merrill et al. 1992, Koch & Nasmyth 1994). In mammalian cells an analogous transcriptional regulation pathway exists orchestrated by E2F transcription factors (Ohtani 1999, Black & Azizkhan-Clifford 1999, Lavia & Jansen-Dürr 1999). However, the expression of several genes encoding DNA polymerases, replication factors and enzymes involved in nucleotide metabolism fluctuates only modestly (less than two-fold) in cycling cells and the yeast-like transcriptional burst in late G1 is restricted to the first cell cycle after mitogenic stimulation of quiescent cells (Wahl et al. 1988, Sherley & Kelly 1988, Morris & Matthews 1989, Zeng et al. 1994). Similarly, expression at the protein level is constant or shows only minor variation. The expression of pol ε is similar to that of other replicative polymerases and replication factors. The steady-state messages for pol ε A and B subunits were 5-8 fold up-regulated when quiescent IMR-90 fibroblasts were stimulated by serum. In cycling cells, the fluctuation was less dramatic showing approximately a 2-fold increase from early G1 to the G1/S border. At the protein level, pol ε showed even smaller changes being almost constantly expressed throughout cell cycle. This is very much in line with the expression of pol α and pol δ (Wahl et al. 1988, Miyazawa et al. 1993, Yang et al. 1992, Zeng et al. 1994a). In this work the transcriptional regulation of POLE2, gene encoding the B subunit of pol ε, was studied in detail. The B subunits of the replicative polymerases in eukaryotes and in Archaea belong to the same superfamily (Mäkiniemi et al. 1999). This suggests that they may share a common function, for example in DNA replication. However, no enzyme activity within B subunits has been reported and their exact role has remained 61 enigmatic. As the only B subunit promoter studied so far, mouse POLA2, encoding the B subunit of pol α, is known to be regulated by E2F transcription factor(s). The POLA2 promoter region contains binding sites for several transcription factors like E2F, Sp1, USF, ATF, Ap2 and p53. The USF binding elements or ‘E-boxes’ in POLA2 promoter bind a USF transcription factor that was essential for constitutive promoter activity.

Fig. 7. The structures of pol α, pol δ and pol ε promoters. Where transcription factor binding has been verified experimentally, the corresponding elements are shown in bold. Data is from Pearson et al., 1991, Chang et al., 1995, Hayhurst et al., 1995, Zhao & Chang, 1997, Huang et al. 1999b, Nishikawa et al. 2000, and Perez et al. 2000.

62

A mutated Sp1 binding element, however, did not disrupt promoter function. The mitogenic stimulation of POLA2 promoter is solely dependent on E2F binding elements on both sides of the transcription initiation site. The over-expression of E2F1 increased POLA2 promoter activity 15-fold and furthermore the downstream E2F element binds the E2F4 transcription factor in vitro (Nishikawa et al. 2000). The analysis of POLE2 promoter revealed a proliferation-induced activation that was dependent on E2F similar to POLA2. However, regulation of the basal level of expression is different and most importantly dependent on a Sp1 element within the 75 bp minimal core promoter. This region also binds E2F2, E2F4 and p107 in vitro. The sequence downstream from the minimal core promoter can repress promoter activity in vitro and furthermore it binds E2F1, NF-1 and pRb. A comparison of the promoters of the replicative polymerase A and B subunits is presented in Figure 7. We have proposed a model of two E2F-pocket protein complexes, one binding to the core promoter and the other to a downstream region, which regulate the promoter, probably in synergy with Sp1 (Huang et al. 2001).

6.2 TopBP1 is a pol ε associated protein with functions in DNA replication and damage response (III)

Our search for pol ε -associated proteins led to the cloning of cDNA and analysis of TopBP1, a human protein, reported to interact with topoisomerase IIβ (Yamane & Tsuruo 1997). TopBP1 is homologous to S. cerevisiae Dpb11 and S. pombe Cut5 within the two pairs of BRCT domains (1&2, 4&5), but has four additional BRCT domains: one pair similar to BRCA1 (7&8) and two further BRCT domains (3,6). Dpb11 is required for DNA replication and for the S phase checkpoint and is essential for loading pol α and pol ε at origins of DNA replication (Araki et al. 1995, Masumoto et al. 2000). Similarly, Cut5 is essential at early stages of DNA replication and involved in the DNA damage checkpoint function (Saka & Yanagida 1993, Saka et al. 1997). Although our attempts to complement temperature sensitive dpb11 and cut5 mutants with TopBP1 failed, we found some evidence that suggests an analogous function in DNA replication and in the damage response. In S. cerevisiae, Dpb11 interacts physically with pol ε throughout the cell cycle, most abundantly in S phase (Masumoto et al. 2000). We were able to co-immunoprecipitate pol ε with TopBP1. The amount of co-precipitating pol ε did not vary during the cell cycle, nor was it changed after DNA damage. Interestingly, it seems that interaction with TopBP1 is restricted to the hyperphosphorylated form of pol ε (see below). Secondly, TopBP1 was expressed in a growth and cell cycle dependent manner with the highest level of expression in G1/S that suggests a role in DNA replication. Thirdly, TopBP1 was found to be required for chromosomal DNA replication in isolated nuclei in vitro. This function was linked to BRCT domain 6 because both the antibody raised against that region as well as the antigen itself inhibited replication 44% and 83%, respectively, in our assay. Obviously, an essential protein-protein interaction is disturbed. Whether this is intermolecular or intramolecular remains to be clarified. 63

In addition to DNA replication, a function in DNA damage response was demonstrated for TopBP1. The sub-cellular localization in S phase is similar to that of BRCA1 with extensive co-localization. While these foci do not represent sites of ongoing DNA replication, both TopBP1 and BRCA1 were re-localized to PCNA containing foci after DNA damage or replication block. These foci represent stalled replication forks and indicate a function for TopBP1 in damage recognition and signalling and/or in DNA repair and re-initiation of replication. We could not demonstrate a physical interaction between TopBP1 and BRCA1, nor did we observe damage induced phosphorylation of TopBP1 or interaction with the damage responsive kinases Chk1 or Chk2. We favour a model, where TopBP1 acts at an early stage of the DNA damage response and allows maintenance or re-establishment of the integrity of the replication machinery in stalled replication forks.

6.3 Pol ε has at least three differently phosphorylated forms with specific interactions with BRCA1 and TopBP1

Pol α and pol δ show cell cycle specific phosphorylation, and with pol α this affects initiation and primase activity (Voitenleitner et al. 1999, Dehde et al. 2001, Zeng et al. 1994). In this study we have shown that pol ε is phosphorylated in vivo. It appeared very difficult to determine individual phosphorylation sites or to link phosphorylation to any function. A key discovery in the analysis of phosphorylation was made when different anti-pol ε antibodies were found to immunoprecipitate the differentially phosphorylated forms of pol ε. The polyclonal antibody K27 raised against the N-terminus of pol ε efficiently immunoprecipitated native, but not denatured, pol ε, and this form of pol ε was hypophosphorylated. The second antibody against the N-terminus of Pol ε, monoclonal 93H3B, modestly immunoprecipitated denatured pol ε. This form was hypophosphor- ylated as well. However, the monoclonal antibody against the exonuclease domain (93G1A) and polyclonal antibodies raised against the polymerase domain (K19, K20) immunoprecipitated the more phosphorylated forms of pol ε from denatured cell lysates. The 93G1A specific form was hyperphosphorylated and K19 (K20) specific form moderately phosphorylated. Furthermore, the tryptic phosphopeptide maps from the 93G1A and K19 specific forms of pol ε were distinct. This difference may originate from an almost completely different set of phosphorylation sites or it may, at least partially, arise from differences in trypsin digestion. The latter would require methylation or acetylation of lysine/arginine residues in one or both forms of pol ε. The existence of these kinds of modifications in pol ε is not known and remains to be explored. Pol ε has been implicated in several cellular processes and purified as a component of complexes associated with these processes. First the human pol ε has been shown to carry out a significant part of chromosomal DNA replication (Zlotkin et al. 1996, Pospiech et al. 1999) and has been purified with a replicative complex or synthesome from breast cancer cells (Coll et al. 1996). Secondly, S. cerevisiae and S. pombe pol ε are necessary for initiation of DNA replication and the budding yeast enzyme has been immunoprecipitated with an initiation complex (activated pre-RC) (Zou & Stillman 64

2000). Thirdly, budding yeast pol ε has been implicated in recombination (Zou & Rothstein 1997, Holmes & Haber 1999) and mammalian pol ε characterized as a component of a recombinative complex (Jessberger et al. 1993). Fourthly, yeast pol ε has been implicated in transcriptional silencing (Ehrenhofer-Murray et al. 1999) and mammalian pol ε has been purified with the RNA polymerase II holoenzyme (Maldonado et al. 1996). Finally, a soluble form of pol ε was originally purified as an essential factor needed in nucleotide excision repair (Nishida et al. 1988). What are the determinants that target pol ε to interact with different proteins in different DNA transactions? Does phosphorylation or other post-translational modifications play a role in these selective targetings? Interestingly, this seems to be the case with pol α, where the hypophosphorylated form is associated with initiation of DNA replication and the hyperphosphorylated form is involved in elongation (Dehde et al. 2001). We found that BRCA1 can be co-immunoprecipitated with the polyclonal K27 antibody that is specific for the hypophosphorylated pol ε. On the other hand TopBP1 is not co-precipitated with this antibody. However, TopBP1 became cross-linked to the hyperphosphorylated form of pol ε that can be immunoprecipitated with the monoclonal antibody 93G1A. It is intriguing to speculate that the hypophosphorylated form of pol ε might have a function in the same processes as BRCA1, such as in recombination, whereas the hyperphosphorylated pol ε would be involved in initiation/re-initiation of DNA replication or in other TopBP1 specific process. Importantly, these processes may be coupled and indeed, TopBP1 and BRCA1 do co-localize in S phase, even without a physical interaction. 7 Conclusions

In this study the expression of pol ε was found to be dependent on the proliferative status of the cell, but much less related to the stage of the cell cycle. This was similar to other replicative DNA polymerases. The transcriptional regulation of POLE2 that encodes the B subunit of pol ε was analysed in detail. The promoter of the gene is regulated by a Sp1 transcription factor and the mitogenic activation was mediated by E2F. We conclude that the results from POLE2 suggest a common E2F mediated activation of the promoters of replicative DNA polymerase B subunits, although the regulation of basal level promoter activity is not conserved. TopBP1 was characterized as a pol ε -associated protein with homology to S. cerevisiae Dpb11 and S. pombe Cut5. TopBP1 was required for chromosomal DNA replication in isolated nuclei in vitro and this property was linked to a region in TopBP1 containing BRCT domain 6. We proposed that this part of the protein interacts with an essential replication factor and that this interaction is required for replicative DNA synthesis. After DNA damage or replication block, TopBP1 was re-localized with BRCA1 in PCNA containing foci, obviously on the sites of stalled replication forks. Our results predict a function in an early stage of the DNA damage response, but whether it is in damage recognition, signal transduction or in maintenance or re-establishment of stalled replication forks is not yet clear. The modular structure of TopBP1 and the demonstrated interactions with pol ε and with Rad9 may suggest a role for TopBP1 as an “integrator” of the damage response. Finally, we identified at least three immunologically distinct forms of pol ε with differential phosphorylation and showed that the hypophosphorylated form interacted with BRCA1, whereas the hyperphosphorylated form was specific for TopBP1. This suggests, but does not prove, different functions for the differentially phosphorylated forms of pol ε. These results pave the way for further studies to determine whether the differential phosphorylation and unique protein-protein interactions target pol ε for specific functions.

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