DNA Replication Across Taxa [1st Edition] 9780128051146, 9780128047354

Table of contents :
Content:
CopyrightPage iv
ContributorsPages ix-x
PrefacePage xiLaurie S. Kaguni, Marcos Túlio Oliveira, Fuyuhiko Tamanoi
Chapter One - Replication Initiation in BacteriaPages 1-30S. Chodavarapu, J.M. Kaguni
Chapter Two - The E. coli DNA Replication ForkPages 31-88J.S. Lewis, S. Jergic, N.E. Dixon
Chapter Three - The Replication System of Bacteriophage T7Pages 89-136A.W. Kulczyk, C.C. Richardson
Chapter Four - Protein-Primed Replication of Bacteriophage Φ29 DNAPages 137-167M. Salas, M. de Vega
Chapter Five - Archaeal DNA Replication Origins and Recruitment of the MCM Replicative HelicasePages 169-190R.Y. Samson, S.D. Bell
Chapter Six - The Eukaryotic Replication MachinePages 191-229D. Zhang, M. O'Donnell
Chapter Seven - The Many Roles of PCNA in Eukaryotic DNA ReplicationPages 231-254E.M. Boehm, M.S. Gildenberg, M.T. Washington
Chapter Eight - Animal Mitochondrial DNA ReplicationPages 255-292G.L. Ciesielski, M.T. Oliveira, L.S. Kaguni
Chapter Nine - Fidelity of Nucleotide Incorporation by the RNA-Dependent RNA Polymerase from PoliovirusPages 293-323C.E. Cameron, I.M. Moustafa, J.J. Arnold
Author IndexPages 325-351
Subject IndexPages 353-359

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Academic Press is an imprint of Elsevier 50 Hampshire Street, 5th Floor, Cambridge, MA 02139, USA 525 B Street, Suite 1800, San Diego, CA 92101-4495, USA The Boulevard, Langford Lane, Kidlington, Oxford OX5 1GB, UK 125 London Wall, London, EC2Y 5AS, UK First edition 2016 Copyright © 2016 Elsevier Inc. All rights reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, recording, or any information storage and retrieval system, without permission in writing from the publisher. Details on how to seek permission, further information about the Publisher’s permissions policies and our arrangements with organizations such as the Copyright Clearance Center and the Copyright Licensing Agency, can be found at our website: www.elsevier.com/permissions. This book and the individual contributions contained in it are protected under copyright by the Publisher (other than as may be noted herein). Notices Knowledge and best practice in this field are constantly changing. As new research and experience broaden our understanding, changes in research methods, professional practices, or medical treatment may become necessary. Practitioners and researchers must always rely on their own experience and knowledge in evaluating and using any information, methods, compounds, or experiments described herein. In using such information or methods they should be mindful of their own safety and the safety of others, including parties for whom they have a professional responsibility. To the fullest extent of the law, neither the Publisher nor the authors, contributors, or editors, assume any liability for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions, or ideas contained in the material herein. ISBN: 978-0-12-804735-4 ISSN: 1874-6047 For information on all Academic Press publications visit our website at https://www.elsevier.com/

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CONTRIBUTORS J.J. Arnold The Pennsylvania State University, University Park, PA, United States S.D. Bell Indiana University, Bloomington, IN, United States E.M. Boehm Carver College of Medicine, University of Iowa, Iowa City, IA, United States C.E. Cameron The Pennsylvania State University, University Park, PA, United States S. Chodavarapu Michigan State University, East Lansing, MI, United States G.L. Ciesielski Institute of Biosciences and Medical Technology, University of Tampere, Tampere, Finland; Michigan State University, East Lansing, MI, United States M. de Vega Instituto de Biologı´a Molecular “Eladio Vin˜uela” (CSIC), Centro de Biologı´a Molecular “Severo Ochoa” (CSIC-UAM), Universidad Auto´noma de Madrid, Cantoblanco, Madrid, Spain N.E. Dixon Centre for Medical & Molecular Bioscience, University of Wollongong, Wollongong, NSW, Australia M.S. Gildenberg Carver College of Medicine, University of Iowa, Iowa City, IA, United States S. Jergic Centre for Medical & Molecular Bioscience, University of Wollongong, Wollongong, NSW, Australia J.M. Kaguni Michigan State University, East Lansing, MI, United States L.S. Kaguni Michigan State University, East Lansing, MI, United States; Institute of Biosciences and Medical Technology, University of Tampere, Tampere, Finland A.W. Kulczyk Harvard Medical School, Boston, MA, United States J.S. Lewis Centre for Medical & Molecular Bioscience, University of Wollongong, Wollongong, NSW, Australia I.M. Moustafa The Pennsylvania State University, University Park, PA, United States ix

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Contributors

M. O’Donnell Howard Hughes Medical Institute, The Rockefeller University, New York, NY, United States M.T. Oliveira Faculdade de Ci^encias Agra´rias e Veterina´rias, Universidade Estadual Paulista “Ju´lio de Mesquita Filho”, Jaboticabal, SP, Brazil C.C. Richardson Harvard Medical School, Boston, MA, United States M. Salas Instituto de Biologı´a Molecular “Eladio Vin˜uela” (CSIC), Centro de Biologı´a Molecular “Severo Ochoa” (CSIC-UAM), Universidad Auto´noma de Madrid, Cantoblanco, Madrid, Spain R.Y. Samson Indiana University, Bloomington, IN, United States M.T. Washington Carver College of Medicine, University of Iowa, Iowa City, IA, United States D. Zhang The Rockefeller University, New York, NY, United States

PREFACE Nearly 100 years passed from the discovery of DNA by Friedrich Miescher in 1869 until the determination of its 3D structure by James Watson and Francis Crick in 1953. Then, just several years later, Arthur Kornberg and colleagues identified the mechanism of its synthesis and the first DNA polymerase. Since that time, the field of DNA replication has grown from the early biochemical investigations to the elucidation of the myriad modes of replication employed in biological systems. In this volume, contributions from nine leading scientists summarize recent progress in studies of systems across taxa. Early research on replication in bacteria revealed the strategies of the less complicated single-stranded DNA bacteriophages G4, M13, and ϕX174. Studies followed of double-stranded bacterial plasmids and bacteriophages, including the bacteriophages T7 and ϕ29, for which recent progress is presented in Chapters 3 and 4, respectively. The in vitro reconstitution of E. coli chromosomal replication in 1982 has led to the elucidation of its mechanism of initiation and elongation, discussed in Chapters 1 and 2, respectively. Although complete replication systems for more complex organisms have remained elusive for a lengthy period, studies of various eukaryotic viruses yielded much information revealing similar basic strategies with multifarious variations. Chapter 5 presents current knowledge in the archaeal system, and Chapters 6–8 describe nuclear and mitochondrial replication in eukaryotes. Similarities and differences in nucleotide polymerization among the polymerases that copy both DNA and RNA genomes are discussed in the final chapter (Chapter 9). We thank the authors for their unique and comprehensive contributions. We are also grateful to Mary Ann Zimmerman and Helene Kabes of Elsevier for their help in launching this project and to Kirsten Shankland and Hannah Colford for seeing it to completion. LAURIE S. KAGUNI MARCOS TU´LIO OLIVEIRA FUYUHIKO TAMANOI April 2016

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CHAPTER ONE

Replication Initiation in Bacteria S. Chodavarapu, J.M. Kaguni1 Michigan State University, East Lansing, MI, United States 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Bacterial Replication Origins 2.1 The Replication Origin of E. coli 2.2 Replication Origins of Other Bacteria 3. Mechanism of Initiation 3.1 Replication Initiation is a Stepwise Process that Begins with the Formation of a Complex of DnaA Assembled at oriC 3.2 DnaA: Domain I Interacts with Other Proteins that Affect Self-Oligomerization of DnaA at oriC 3.3 DnaB 3.4 The DnaB–DnaC Complex and Its Formation 3.5 Formation of the Prepriming Complex 3.6 Helicase Activation Requires the Dissociation of DnaC from DnaB 3.7 Events Leading to Assembly of the Replisome 4. Summary Acknowledgments References

2 2 2 9 11 11 11 14 15 16 19 20 20 21 21

Abstract The initiation of chromosomal DNA replication starts at a replication origin, which in bacteria is a discrete locus that contains DNA sequence motifs recognized by an initiator protein whose role is to assemble the replication fork machinery at this site. In bacteria with a single chromosome, DnaA is the initiator and is highly conserved in all bacteria. As an adenine nucleotide binding protein, DnaA bound to ATP is active in the assembly of a DnaA oligomer onto these sites. Other proteins modulate DnaA oligomerization via their interaction with the N-terminal region of DnaA. Following the DnaA-dependent unwinding of an AT-rich region within the replication origin, DnaA then mediates the binding of DnaB, the replicative DNA helicase, in a complex with DnaC to form an intermediate named the prepriming complex. In the formation of this intermediate, the helicase is loaded onto the unwound region within the replication origin. As DnaC bound to DnaB inhibits its activity as a DNA helicase, DnaC must dissociate to activate DnaB. Apparently, the interaction of DnaB with primase (DnaG) and primer formation leads to the release of DnaC from DnaB, which is coordinated with or followed by

The Enzymes, Volume 39 ISSN 1874-6047 http://dx.doi.org/10.1016/bs.enz.2016.03.001

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2016 Elsevier Inc. All rights reserved.

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translocation of DnaB to the junction of the replication fork. There, DnaB is able to coordinate its activity as a DNA helicase with the cellular replicase, DNA polymerase III holoenzyme, which uses the primers made by primase for leading strand DNA synthesis.

1. INTRODUCTION DNA replication is a central process in all organisms. This biochemical pathway is divided into three stages: initiation, elongation, and termination. Individual proteins act in concert with others at each stage, performing specific functions that lead to the duplication of DNA. The study of DNA replication has its foundation in studies of Escherichia coli as a model system. Of interest, studies of other bacterial species, such as Bacillus subtilis, Caulobacter crescentus, and Helicobacter pylori, have revealed that the molecular mechanisms used among bacteria are remarkably similar. Moreover, the program of molecular events in these organisms at the stage of replication initiation is surprisingly like that of eukaryotes and archaebacteria. Together, these studies of bacteria may lead to the development of novel antibiotics that act to inhibit DNA replication in bacterial pathogens. These shared features include recognition of a replication origin by a DNA binding protein, or in some cases a complex of proteins, unwinding of the parental duplex DNA, loading of the replicative DNA helicase onto each strand of unwound DNA, and activation of the helicase. Recruitment of the cellular replicase follows, which leads to the assembly of the enzymatic machinery that acts at a replication fork to duplicate the chromosome. This chapter summarizes replication initiation in E. coli, and highlights similarities and important differences in the biochemical mechanisms of initiation in other bacterial species.

2. BACTERIAL REPLICATION ORIGINS 2.1 The Replication Origin of E. coli DNA replication in bacteria starts at a unique locus called the replication origin, or oriC. Among bacteria, this site contains a variety of DNA sequence elements described in more detail below that are recognized by proteins that are either directly involved in replication initiation, or that regulate the frequency of this event to coordinate it with bacterial growth and cell division. In E. coli oriC, this 250 base pair (bp) sequence is located between the gidA

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and mioC genes at 84.3 min on the genetic map (reviewed in Ref. [1]). Originally, this site was identified by measuring the gene frequency in replicating cells [2,3], or in cells manipulated to initiate DNA replication synchronously. In the latter approach, a growing culture of a dnaC(Ts) mutant was transferred to an elevated temperature to block new rounds of DNA replication while permitting ongoing DNA replication to continue to completion [4]. A downshift to the permissive temperature leads to a new round of DNA replication and also an elevated ratio of oriC to distal sites, which were measured by radioactivity. Alternate methods rely on the ability of oriC to confer autonomous replication to derivatives of bacteriophage lambda under conditions in which the bacteriophage replication origin is inhibited [5,6], or to plasmids that confer antibiotic resistance [7–9]. For bacterial species that are not closely related evolutionarily to E. coli, the location of the respective replication origins is often found adjacent to the dnaA gene, but the length of oriC is variable, ranging from 100 to 1000 bp. E. coli oriC has been the subject of molecular analyses by a variety of methods. One approach relies on comparative DNA sequence analysis of E. coli oriC and the replication origins of evolutionarily related bacteria to identify conserved DNA regions that presumably represent sequences recognized by specific proteins. Another is DNA footprinting to demonstrate that a protein is able to bind to a particular DNA sequence. A third relies on mutational analysis of minichromosomes wherein oriC joined to a drug resistance gene provides for maintenance of this DNA in bacterial cells. These approaches have led to the identification of five copies of a DNA motif named the DnaA box that is recognized by DnaA. To indicate the positions of these DnaA boxes, they have been named R1 through R5. These DnaA boxes are essential for oriC function on the basis that individual mutations in each cause impaired activity [10–12]. Although R1 and R4 are identical in DNA sequence, DnaA binds with higher affinity to R4 presumably because of its differential interaction with nucleotides that flank these sites [13–15]. Inasmuch as DnaA binds specifically to adenine-containing nucleotides [16], both DnaA-ATP and DnaA-ADP comparably recognize these DnaA boxes [16–18]. In contrast, DnaA-ATP binds with lesser affinity to R2 followed by R3, and with even lower affinity to sites named I1, I2, I3, R5, τ1, τ2, C1, C2, and C3 [17,18]. Of interest, recent studies suggest that DnaA-ATP may actually interact with C2 and C3 instead of R3, which are on either side and overlap R3 [19]. Hence, DnaA may not recognize the R3 DnaA box. These observations with DnaA-ATP differ from those with DnaA complexed to ADP, which poorly recognizes the low-affinity sites.

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These differences in the adenine-bound forms of DnaA correlate with the activity of DnaA-ATP and not DnaA-ADP in replication initiation [16]. Considering that B-form DNA has about ten base pairs per turn of the DNA helix, other studies strongly suggest that the relative arrangement of DnaA molecules assembled at oriC is important. Phylogenetic comparison of the replication origins of Gram negative bacteria reveals conserved regions that are separated by DNA of fixed length whose sequences are not conserved, suggesting that spacing or phasing between the conserved regions is critical for oriC function [20]. Moreover, inverting the DnaA boxes R1, R2, or R4 [12], or changing the distance between R3 and R4 or R2 and R3 led to inactivation of oriC [21,22]. In contrast, insertion or deletion of 10–12 base pairs did not render oriC nonfunctional [21,23]. Together, these observations strongly suggest that the orientation of DnaA bound to a specific DnaA box relative to other DnaA-DnaA box complexes in oriC is essential. As will be described in more detail below, DnaA self-oligomerizes via sites in its N-terminal domain (domain I), and its ATP binding module (domain III). In view of observations that DnaA oligomerization is required for replication initiation [24–26], these findings support a model in which DnaA complexed to the individual DnaA boxes at oriC forms a specific nucleoprotein complex. An attractive idea is that the DnaA boxes serve as sites for nucleation of DnaA-ATP. These DnaA molecules bound in an ordered manner at these sites serve to recruit additional DnaA molecules to the nearby lower affinity sites, which presumably stabilizes a DnaA oligomer. On the basis of the orientation of the DnaA boxes, τ-, I-, and C-sites [17–19], Leonard and Grimwade proposed that oriC is a bipartite structure, and suggested that DnaA assembles on the left and right portions of oriC as separate opposing oligomers [1,19]. In support, alterations in the DNA sequence, orientation, or spacing of the low-affinity sites in the left or right portions of oriC disrupted DnaA binding, and caused impaired function of oriC in DNA replication [19]. The work summarized above relies on the analysis of plasmids that carry oriC. Of interest, studies of these DNA motifs when oriC is in its natural environment of the E. coli chromosome revealed important differences. An early study showed that insertion of a two kilobase pair DNA fragment between R3 and R4 inactivates oriC when borne by a plasmid, but not when this insertion mutation was placed into the E. coli chromosome [27]. This work also showed that deletion of R4 in an oriC-containing plasmid

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blocks origin function, but its removal from the chromosome did not interfere with viability. However, initiation in the R4 deletion mutant under culture conditions that lead to rapid growth and multiple initiations was no longer synchronous.a More recent studies showed that deletion of the region that removes R4, the DNA motifs C3, C2, C1, I3, and R3, like the R4 deletion mutation described above, did not impair viability [33]. Physiological characterization of this mutant revealed a decreased rate of growth, fewer origins per cell, and asynchronous initiations. These defects were exacerbated in rich growth media but were suppressed in poor media. These findings suggest that the segment at the right in oriC is uniquely required when the conditions call for multiple initiations within a single cell. Hence, the DNA elements within oriC needed for its function are not fixed but depend on physiological conditions. Bearing in mind the idea that R4 serves as a DnaA nucleation site that directs its cooperative binding to the C3, C2, C1, and I3 sites nearby, the formation of this DnaA oligomer may be needed under the conditions of multiple initiations. In addition to the DNA motifs recognized by DnaA, E. coli oriC has an unusual abundance of 11 GATC sequences that are targeted by DNA adenine methylase (reviewed in Ref. [28]). For simplicity, Fig. 1 omits these sites, which are also preferentially recognized by SeqA when they exist in the hemi-methylated state. Shortly after initiation, SeqA complexed to the resultant hemi-methylated DNA appears to occlude oriC from DnaA and other proteins needed for replisome assembly. After SeqA dissociates from oriC, DNA adenine methylase is able to convert the hemi-methylated sites in oriC to the fully methylated form to permit another cycle of initiation.

a

Although bacteria are considered to be haploid, individual cells in rapidly growing cultures initiate new rounds of DNA replication prior to cell division and before the previous cycle has been completed [28]. Depending on the culture conditions, such cells bear chromosomes that are partially diploid, partially tetraploid, or even partially octaploid. As measured by flow cytometry under conditions in which cell division and new initiations are blocked, but ongoing DNA replication can proceed to completion, cells normally carry 2n chromosomes where n
0 and is an integer [29–31]. Hence, cells growing quickly initiate DNA replication synchronously and at a particular time in the bacterial cell cycle. In contrast, asynchronous DNA replication leads to odd numbers of chromosomes as observed in mutants carrying dnaA(Ts) alleles defective in ATP binding, suggesting that ATP binding by DnaA is necessary for proper regulation of initiation [32]. Other mutations (fis, himA, seqA, dam, hupA, and hda) also cause asynchronous initiations because the respective mutations perturb the initiation process by direct or indirect effects.

Escherichia coli L M R

R1

τ1

gidA DUE

R5

τ2 I1 I2

R2

C3

R3 R4 C2 I3 C1 mioC

IHF

Fis

G2 W5

W4 W3W2 W1

DnaA box τ-site I-site C-site

13mer

Caulobacter crescentus G1

duf299 DnaA box (G, moderate affinity) (W, weak affinity) CtrA GANTC (CcrM) IHF

hemE AT-rich region

Bacillus subtilis dnaA

rpmH

dnaN

oriC1

16mer

oriC2

DUE 27mer

DnaA box 16mer AT-rich cluster

Helicobacter pylori punB

dnaA oriC1

comH oriC2

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DnaA box

Vibrio cholerae oriC1 gidA

mioC DUE

DnaA box

13mer

Vibrio cholerae oriC2 inc2

oriC2

11mer 39mer

12mer

29mer

rctA

rctB DnaA box

AT-rich region

11mer 12mer RctB binding sites 29mer 39mer DnaA box

Fig. 1 Bacterial replication origins carry DNA sequence motifs recognized by DnaA and other proteins, and an AT-rich region named the DNA unwinding element (DUE). Among bacteria however, the presence, relative arrangement, and position of these DNA motifs are not conserved. For E. coli, DnaA in a complex with either ATP or ADP recognizes a specific DnaA box with comparable affinity, but the relative affinity among the individual DnaA boxes varies. In comparison, DnaA-ATP and not DnaA-ADP specifically binds to I-, τ-, and C-sites. The affinities of DnaA-ATP to these sites are less than to the DnaA boxes. E. coli oriC also carries binding sites for Fis and IHF, whereas the replication origin of C. crescentus contains a binding site for IHF but not Fis. In contrast with these replication origins, which are contiguous DNA sequences, the replication origins of B. subtilis and H. pylori are bipartite in which oriC1 and oriC2 are separated by the dnaA gene. B. subtilis oriC1 has three AT-rich 16mer repeats upstream of dnaA [34,35]. A 27mer AT-rich cluster in the oriC2 region is unwound by DnaA [36]. V. cholerae carries two chromosomes. Initiation from oriC1 appears to be similar to E. coli oriC. Initiation from oriC2 requires RctB, which recognizes 11mer and 12mer iterons, and also a 39mer and a truncated form of the 39mer named the 29mer. The 39mer in the inc2 region and the 29mer bound by the monomeric form of RctB together with the DnaA box bound by V. cholerae DnaA regulate the frequency of initiation from oriC2 [37]. Adapted from Refs. [36–38].

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2.1.1 The DNA Unwinding Element of oriC and DnaA Replication initiation from oriC in vitro requires its presence in a supercoiled DNA. Assembled at oriC, DnaA complexed to ATP or ATPγS, an analogue that is poorly hydrolyzed by most ATPases, induces the unwinding of an AT-rich region located near the left border of oriC. This region, which is sometimes described as a DNA unwinding element or DUE [39], contains three 13mer repeats that when unwound at an optimal temperature of 38°C are sensitive to nucleases or chemicals that are specific for single-stranded DNA [40]. Presumably, this condition aids in the disruption of base pairs. An interesting conundrum that is not understood, DnaA binds with relatively high affinity to ATP (KD ¼ 0.03 μM), yet unwinding requires much higher concentrations of 5 mM ATP [16]. Despite this lack of understanding, ATPγS in place of ATP but not ADP supports unwinding. Hence, ATP hydrolysis is not required. Apparently, DnaA complexed to ATP or ATPγS is in a conformation that is active in unwinding whereas DnaA-ADP is not. As evidence that the conformation of DnaA complexed to ADP is different from DnaA-ATP, X-ray crystallographic analysis of domains III and IV of Aquifex aeolicus DnaA complexed to ADP revealed a different structure in comparison with DnaA bound to AMP-PCP, an ATP analogue (see later) [41,42]. The co-crystal containing ADP supports a model of a closed ring composed of six molecules of DnaA, each arranged with ADP at the interface separating adjacent protomers. The co-crystal containing AMP-PCP underpins a model of a right-handed helical filament. Each model has hydrophobic and positively charged amino acids in the interior channel. Evidence presented below supports a model that the unwound DNA interacts with accessible amino acids of the channel’s interior. To test the role in DNA binding of specific hydrophobic and positively charged amino acids that line the inner channel, mutant DnaA proteins bearing alanine substitutions for Val221 and Arg245 were shown to be impaired in unwinding of oriC contained in a linear duplex DNA [43]. Under comparable conditions, wild-type DnaA complexed to ATP was active but DnaA-ADP was not. An independent study showed that wild-type DnaA is active in strand displacement of a duplex oligonucleotide of 15 base pairs, but not of a 30 base pair duplex DNA, suggesting that DnaA exploits the relative thermodynamic instability of the DUE by binding to the transiently unwound DNA [44]. These biochemical observations correlate with the X-ray crystallographic structure of domains III and IV of A. aeolicus DnaA complexed to AMP-PCP and polydA12 [44], supporting the model that

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unwinding of oriC involves the binding of the unwound DNA to the inner cavity of DnaA assembled as a helical filament. As noted above, the DUE of E. coli oriC contains three 13mer motifs. Within each 13mer, a 6mer sequence (consensus: AGATCT) is present that is specifically recognized by DnaA-ATP when this DNA is single-stranded [45,46]. Preferential binding to the 6mers in the top strand of the DUE is thought to require the initial binding of DnaA to the DnaA box named R1 that is proximal to the DUE as well as to DNA to the right up to and including the DnaA box named R5 [43,47,48]. Presumably, the interaction between adjacent DnaA monomers stabilizes their binding to this region of duplex DNA. Single-molecule fluorescence assays showed that DnaA assembles one monomer at a time in the 30 -to-50 direction as a dynamic filament on single-stranded DNA [49]. Confirming results from the X-ray structure of the truncated form of A. aeolicus DnaA bound to polydA12 [44], each monomer binds to three nucleotides of ssDNA [49]. 2.1.2 Binding Sites for Fis and IHF Within E. coli oriC E. coli oriC also carries binding sites for Fis (factor for inversion stimulation), and IHF (integration host factor) [50,51]. Fis was originally discovered as a required factor for Hin and Gin site-specific DNA recombinases of Salmonella and bacteriophage Mu, respectively [52,53]. IHF was identified by its involvement in the integration of bacteriophage λ DNA during lysogeny [54]. Despite the different locations of their binding sites, a common feature of both proteins is their ability to bend DNA. Mutation of their respective binding sites in oriC disrupts origin function [55], which correlates with the asynchronous initiations observed in mutants that lack fis or ihfB [56,57]. The latter gene encodes one of the two subunits of IHF [58]. Other work showed that these DNA binding proteins act during the initiation stage of DNA replication. One study showed that IHF stimulates the DnaA-dependent unwinding of the 13mer region of oriC [59], which correlates with in vitro footprinting studies in which IHF apparently leads to the redistribution of DnaA to R2, R3, R5, and to low-affinity sites recognized by DnaAATP [60]. These findings complement in vivo footprinting results, which showed that both R3 and the IHF site became protected at the time of initiation in synchronized cells [61]. Other studies suggest that IHF and Fis have opposing effects on the binding of DnaA to oriC. Low levels of Fis apparently inhibit the unwinding of oriC by interfering with the binding of IHF and DnaA [62,63]. However, higher levels of DnaA augmented by the presence of IHF overcome this inhibition. These events correlate with the lack of

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occlusion of the Fis site at initiation in vivo whereas this site is protected throughout other periods of the cell cycle [61]. Evidently, the dynamic influence of Fis and IHF on the binding of DnaA to R3 suggests that these proteins modulate the frequency of initiation (reviewed in Ref. [1]).

2.2 Replication Origins of Other Bacteria The replication origins of bacteria closely related to E. coli are similar in both DNA sequence and the relative organization of the conserved DNA sequence motifs described above [20,64] (reviewed in Refs. [1,65]). These observations suggest that the mechanism of replication initiation is similar in evolutionarily related bacteria. As evidence, one of its two chromosomes named chromosome 1 of Vibrio choleraeb has a single replication origin (oriC1) that is flanked by gidA and mioC, and is similar in DNA sequence and the organization of essential DNA motifs compared with E. coli oriC. As oriC1 can functionally replace E. coli oriC [66], E. coli DnaA is capable of recognizing this DNA, and can direct the assembly of the replication fork machinery at this site to duplicate the chromosome. However, the V. cholerae origin is a less effective substitute for E. coli oriC as indicated by less frequent initiations. E. coli oriC contains sequence motifs that are bound with varying affinities by DnaA. Likewise, the replication origin of Caulobacter crescentus has DNA motifs named G1 and G2 that are bound by DnaA with moderate affinity, whereas DnaA has a weak affinity for sites named W1–5 [69]. Presumably, these differences in affinity reflect the need for the ordered assembly of DnaA molecules at the C. crescentus origin, leading to the expectation that the chromosomal origins of other bacteria also contain sites that vary in their affinity for DnaA. If so, assembly of a DnaA oligomer may be a common requirement at all bacterial origins despite their diverse DNA sequences, the variation in the relative positions and orientations of DNA motifs within including the number and spacing of DnaA boxes, their locations in the chromosome relative to flanking genes, and their different lengths. b

In bacteria with divided genomes, a protein not related to DnaA recognizes a specific replication origin of a chromosome. For example, the second chromosome of V. cholerae contains a single replication origin named oriC2 located between two genes, rctA and rctB (Fig. 1). oriC2 contains an array of tandem repeats named the 11mer and 12mer that are bound by monomers and dimers of RctB [37,38]. Another DNA motif named the 39mer is specifically bound by monomeric RctB. The methylation status of GATC sites that are recognized by DNA adenine methylase influences the binding of RctB to these sites wherein the dimer of RctB negatively regulates the frequency of initiation from oriC2 by binding to the 39mers [66]. DnaA is essential for initiation from oriC2, but apparently performs an auxiliary role [67,68].

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The C. crescentus origin also contains sites that are recognized by CtrA, which negatively regulates initiation to coordinate duplication of the chromosome with cellular development [69,70]. Biochemical studies reveal that CtrA, which is regulated by phosphorylation, inhibits the binding of DnaA to the C. crescentus origin [70,71]. Of interest, a protein analogous to E. coli IHF overlaps a binding site for CtrA. Occupancy of the IHF site is thought to displace CtrA, leading to initiation [72]. A DNA methyltransferase, CcrM, methylates the adenine base in the sequence GANTC [73]. Unlike the indirect role of DNA adenine methylase in controlling the frequency of initiation in E. coli [28], CcrM-dependent methylation is not involved in the control of replication initiation in C. crescentus. However, methylation of promoter regions by CcrM is required for the efficient transcription of many genes that are essential for cell cycle progression [74,75]. Compared with the replication origins summarized above, which are contiguous DNA sequences, the origins of some bacteria are bipartite. As represented by the replication origins of B. subtilis and H. pylori, a gene (dnaA for these but not all other organisms with bipartite origins) separates two essential regions named oriC1 and oriC2 (Fig. 1). These regions bear clusters of DnaA boxes, but oriC2 also contains the DUE. For the B. subtilis origin, deletion analysis of the intervening dnaA gene revealed that much of it could be removed without affecting the function of the remaining DNA as a replication origin when carried in a plasmid [76]. It has not been determined if the B. subtilis origin can tolerate the removal of all of dnaA, or if a minimal length of spacer DNA is required. The latter result suggests that pairing of the left and right portions is essential for replication initiation, which also requires two additional proteins named DnaD and DnaB [77,78]. Homologues of B. subtilis DnaB or DnaD are not present in E. coli. Note that E. coli DnaB is unrelated to B. subtilis DnaB. DnaD is recruited to the origin through an interaction with DnaA. Similar deletion analysis of the dnaA gene that separates the two parts of the H. pylori origin has not been performed. However, the binding of DnaA to oriC2 requires that it is supercoiled, which is essential for DnaA to unwind the DUE [79,80]. Only two of the three DnaA boxes display this requirement for supercoiling. By comparison, supercoiling does not affect the binding of E. coli DnaA to the DnaA boxes of E. coli oriC [14,81]. For the H. pylori origin, the region at the left (oriC1) affects the amount of unwound DNA produced. In its absence, about 20 base pairs is unwound compared with as much as 52 base pairs when oriC1 is present, suggesting that oriC1 and oriC2 interact, but are separated by a loop containing the dnaA gene [79].

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3. MECHANISM OF INITIATION 3.1 Replication Initiation is a Stepwise Process that Begins with the Formation of a Complex of DnaA Assembled at oriC DnaA protein initiates DNA replication by forming a specific DnaA-oriC complex. The types of high and low-affinity sites, the influence of the adenine nucleotide bound to DnaA on recognition of these sites, and evidence that describes the structure of this complex have been summarized above. Structure–function studies of DnaA indicate that it has four functional domains (Fig. 2). Domain IV is necessary and sufficient for binding to the DnaA box motif. Specific residues that confer DNA binding activity as well as specificity in DNA binding have been identified by molecular genetic methods [84,85]. Structures of this domain bound to the DnaA box complement the analyses described above [15,42], and also identify amino acid residues that contact nucleotides flanking the DnaA box sequence. As mentioned above, domains I and III contribute to DNA binding by promoting cooperative interactions between neighboring DnaA molecules assembled at oriC [24,25,86].

3.2 DnaA: Domain I Interacts with Other Proteins that Affect Self-Oligomerization of DnaA at oriC Following the assembly of DnaA-ATP at oriC, DnaA induces strand opening of the AT-rich region, creating an intermediate named the open complex [40]. Early studies showed that HU, a protein that binds to DNA nonspecifically and is also a major component of the bacterial nucleoid, or IHF stimulate the formation of this intermediate [54,59,87]. More recent studies have identified a protein named DiaA that, like HU and IHF, stimulates the DnaA-dependent unwinding of oriC [88]. The underlying biochemical mechanism for DiaA and HU appears to involve their ability to interact with domain I of DnaA to facilitate DnaA oligomerization or to stabilize the DnaA oligomer after it has formed [89,90]. The evidence that DiaA interacts with domain I is based on the observation that the substitution of phenylalanine with alanine at residue 46 of DnaA inhibits its interaction with DiaA [90], and on reports that DiaA, a structural homologue of H. pylori HobA [91], apparently interacts with the same surface in domain I of DnaA as H. pylori HobA [92]. In contrast with these observations and as summarized earlier, IHF is suggested to reorganize

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DnaA Domain I Domain II

Domain IIIa and IIIb

Interaction with DnaB

Sensor 1 Box VII (RLKSR)

Walker A

Interaction with DnaB, HU, Dps, DiaA, L2, and Hda DnaA oligomerization

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Domain IV 347

Sensor 2

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1

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DNA binding

AAA+ domain, DnaA oligomerization

Interaction with phospholipids

DnaB Interaction with primase

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80

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1 15

RecA domain, interaction with ATP and DnaC

DnaC AAA+ domain

Sensor 1 Box VII (RVMDRM) Sensor 2

Interaction with DnaB

245

Walker B

73

Walker A

1

Fig. 2 Domain organization of DnaA, DnaB, and DnaC protein. The numbers in the respective lines refer to the coordinates for E. coli DnaA, DnaB, and DnaC protein. DnaA: Domain I interacts with DnaB, HU, Dps, DiaA, Hda, and ribosomal protein L2, and is also required for DnaA oligomerization. Domain II may function as a flexible linker to join domain I and III. Domain III carries the amino acid sequence motifs shared among AAA+ family members that act in ATP binding and its hydrolysis. This domain also functions in DnaA oligomerization, and appears to carry a site denoted by a filled square that interacts with DnaB. Domain IIIa carries an abbreviated RecA-type fold. Domain IIIb contains a three-helix bundle. Domain IV binds to the DnaA box and presumably also to I-, τ-, and C-sites. A region that interacts with acidic phospholipids is in domain IV. The borders separating the domains have been determined by functional analysis of DnaA together with a homology model based on the X-ray crystal structure of domain III and IV of A. aeolicus DnaA. DnaB: Its N-terminal domain interacts with primase and its larger C-terminal domain functions in ATP binding and hydrolysis. On the basis of the X-ray crystallographic structures of Geobacillus kaustophilus and Geobacillus stearothermophilus DnaB [82,83], this C-terminal domain that also interacts with DnaC is similar in structure to RecA. The filled symbols represent the Walker A and B boxes and the arginine finger (arginine 442 of E. coli DnaB), and the DnaC-interacting domains. DnaC: The interacting domains of DnaC with DnaB and with ATP, including the AAA+ motifs and the conserved arginines in box VII are shown. Reviewed in Ref. [42].

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individual DnaA molecules bound to respective low- and high-affinity DnaA sites within oriC [60]. Assuming that IHF operates at oriC by a single biochemical pathway, it appears that this protein uses a different mechanism than DiaA and HU to stimulate unwinding by DnaA. Domain I of E. coli DnaA also interacts with Dps [93], ribosomal protein L2 [94], and Hda [95]. Dps is a stress-induced protein that protects the bacterial chromosome from DNA damage by sequestering and oxidizing Fe2+ [96,97] (reviewed in Refs. [98,99]). It also inhibits strand opening of oriC in vitro, which correlates with in vivo evidence that Dps reduces the frequency of initiation [93]. Together, these observations suggest that Dps may act as a checkpoint to inhibit initiation during oxidative stress so that the cell has an opportunity to repair its DNA. Dps may inhibit initiation by interfering with DnaA oligomerization at oriC. By interacting with the N-terminal region of DnaA, ribosomal protein L2 as well as a truncated form lacking 59 N-terminal residues inhibits initiation of oriC-containing plasmids in vitro by disrupting the ability of DnaA to self-oligomerize at oriC [94]. Like Dps, L2 obstructs the DnaAdependent unwinding of oriC. In free-living organisms, both replication initiation and ribosome biogenesis are coordinated with cell growth but the mechanism underlying this coordination is not understood (reviewed in Ref. [100]). L2 is encoded by rplB, which is in the S10 ribosomal operon that is autoregulated by ribosomal protein L4 at the transcriptional and translational level [101]. As a possible explanation, when ribosomefree L2 is in excess, such as during the transition from rapid to slow cell growth, it may interact with DnaA to inhibit initiation while L4 that is also in oversupply due to expression of the S10 operon inhibits expression of the operon. This model provides a feedback mechanism to coordinate the control of ribosome biogenesis with DNA replication. In a pathway in which Hda complexed to the β clamp stimulates the hydrolysis of ATP bound to DnaA to regulate its activity (reviewed in Ref. [102]), Hda interacts with the AAA+ domain of DnaA to stimulate ATP hydrolysis, and also with domain I [95,103]. Deletion analysis revealed that the N-terminal 56 residues of DnaA are involved in this interaction [95]. An N44A substitution in DnaA caused a small reduction in the physical interaction between DnaA and Hda in a pull-down assay. It is not known if this substitution affects DnaA oligomerization. In B. subtilis, SirA and other proteins have been identified that interact with DnaA. Studies have shown that SirA, which is under SpoOA-P

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regulation, interacts with domain I of B. subtilis DnaA to inhibit replication initiation in diploid cells committed to the formation of dormant spores [104–106]. Both B. subtilis SirA and H. pylori HobA bind to the same surface within domain I of their respective DnaAs [106]. However unlike HobA, which is essential for self-oligomerization of H. pylori DnaA [107], SirA inhibits DnaA in initiation probably by destabilizing the DnaA oligomer or by interfering with its assembly at oriC [92,108]. (In contrast, YabA, DnaD, and Soj of B. subtilis interact with domain III [109–115].) These and other observations suggest that proteins of respective bacteria bind to the same or other sites in domain I to affect either formation of the DnaA oligomer at oriC or its stability. A model has been proposed that domain I acts as a sensor by its interaction with other proteins in order to adjust the frequency of initiation with changes in cell physiology, or the cell’s environment [94].

3.3 DnaB DNA helicases are enzymes that coordinate nucleotide hydrolysis with their translocation and unwinding of duplex nucleic acids. Of the six superfamilies of these enzymes, DnaB is in Superfamily 4 that is categorized by the presence of a RecA-type domain (Fig. 2) [116,117]. Whereas other members of this superfamily are assemblies of both hexameric and heptameric subunits, DnaB is a hexamer of identical subunits; a DnaB heptamer has not been observed. Each of the six protomers of DnaB has an N-terminal domain joined by a linker to the C-terminal RecA domain, which functions to coordinate nucleoside triphosphate hydrolysis with both unwinding of duplex DNA and translocation of the enzyme in the 50 -to-30 direction relative to the single-stranded DNA to which the enzyme is bound [118–124]. Both early cryo-electron microscopy of E. coli DnaB [125,126], and X-ray crystallographic analysis of DnaB from thermophilic bacteria revealed that DnaB is a toroid [82,83,127]. These and other X-ray structures integrated into the framework of cryo-electron microscopic structures support a model that the form of DnaB complexed with DnaC is an open ring with a right-handed turn, which contrasts with the closed ring structure of DnaB in the absence of DnaC [128,129]. X-ray structures of A. aeolicus DnaB in the absence of DNA revealed constricted and dilated conformations of the portion of the DnaB ring formed by the N-terminal domains of neighboring DnaB protomers [128]. These observations strongly suggest dynamic movement of its

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domains, which have been corroborated in hydrogen-deuterium exchange experiments discussed below. With DnaB bound to an artificial replication fork, fluorescence energy transfer experiments indicated that the larger C-terminal domain is oriented nearest the fork junction [130]. The DNA strand bound by DnaB passes through its central cavity. A mechanism has been proposed in which this single-stranded DNA interacts with specific amino acids in the central cavity while the other DNA strand of the replication fork is excluded during translocation [116,131–133].

3.4 The DnaB–DnaC Complex and Its Formation Studies characterizing the DnaB–DnaC complex revealed that as many as six DnaC molecules bind to DnaB [134–136]. Although early studies suggested that ATP is needed for DnaC to form a complex with DnaB [137], later studies showed that ATP or ADP is not required [136,138,139]. These nucleotides do not substantially affect binding affinity. Several lines of evidence show that DnaC interacts via residues near its N-terminus with specific residues near the C-terminus of DnaB. Genetic and biochemical studies demonstrated that amino acid substitutions within residues 8–11 and 31–44 of DnaC abrogate its interaction with DnaB and the function of DnaC in replication initiation [140]. In hydrogendeuterium exchange experiments, which measures protein dynamics by the relative ability of amide hydrogens in a protein to exchange with deuterium in a buffer made with heavy water, peptides corresponding to these segments of DnaC were found to become occluded when DnaC is complexed to DnaB [141]. Together, these results indicate that these regions of DnaC interact directly with DnaB. Other evidence showed that mutant DnaBs bearing I297A, L304A, or E435A substitutions are defective in complementing a temperature-sensitive dnaB mutant at nonpermissive temperature [141]. To substantiate that the E435A substitution interferes with DnaB function, biochemical experiments were performed, which showed that this protein is inactive in DNA replication of an oriC-containing plasmid. In addition, this substitution blocks the interaction of DnaB with DnaC. These results are complemented by hydrogen-deuterium exchange experiments, which indicated that peptides bearing residues 295–304 and 431–435 display greatly reduced rates of exchange when DnaB is complexed to DnaC, but not when the proteins were analyzed separately [141]. Hence, these regions of DnaB interact with DnaC in formation of the DnaB–DnaC complex.

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3.5 Formation of the Prepriming Complex After formation of the open complex, the DnaB–DnaC complex binds to the unwound region of oriC to form an intermediate named the prepriming complex (Fig. 3). Quantitative analysis combined with permanganate footprinting experiments indicated that each separated strand is bound by a single DnaB–DnaC complex [142–144]. The one bound to the top strand is near the left border of oriC [142]. The complex on the bottom strand is near the DnaA box named R1. On the basis that DnaB and not DnaC interacts directly with DnaA in E. coli, it is thought that DnaA directs the loading of the DnaB–DnaC complex on each unwound strand [145,146]. Two regions of DnaA interact with DnaB. Deletion mutants lacking domain I or portions of it as well as E21A or F46A substitutions of DnaA are defective in interacting with DnaB, indicating a requirement for domain I [90,146]. A second interacting region in domain III is indicated by evidence that a monoclonal antibody that recognizes a conformational epitope within amino acids 111–148 of DnaA inhibits its interaction with DnaB [145]. Deletion analysis restricted this interacting region to residues 135–148 [147]. Formation of a DnaA oligomer is also required to load the DnaB–DnaC complex at oriC. As evidence, a mutant DnaA bearing a W6A substitution is unable to form a DnaA oligomer at oriC, and fails to load the DnaB–DnaC complex [25]. Thus, DnaA must be able to interact with DnaB and also form a DnaA oligomer to load the DnaB–DnaC complex at oriC. In contrast with the results summarized above, DnaC from A. aeolicus is able to interact physically with DnaA from this bacterium. These results suggest that DnaC assists in helicase loading in this organism, raising the possibility of a similar mechanism in E. coli. However, enzyme-linked immunosorbent assays to measure an interaction between DnaA and DnaC of E. coli were unsuccessful under conditions that were able to detect an interaction between DnaA and DnaB, or DnaB and DnaC [145]. Considering that the dnaC gene is proposed to have arisen by gene duplication of dnaA followed by the separate evolution of dnaA and dnaC so that the respective proteins perform separate functions in initiation, an alternate possibility is that the respective DnaCs have evolved to differ in their ability to interact with DnaA. As genetic evidence indicates that DnaC is required for replication initiation but not at the elongation stage during which DNA is duplicated [148–151], what is its role? Initial biochemical characterization of DnaC

17

Replication Initiation in Bacteria

E. coli oriC L M R

R1

τ1

gidA DUE

R5

τ2 I1 I2

R3 R4 C3 C2I3C1

R2

mioC

IHF

Fis

C-site I-site τ-site

DnaA box IHF Fis

13mer

1 2 3

1

Assembly of DnaA at oriC

τ1

τ2 R5 I1I2

R1

DnaA

4

C3

R2

C2 C2

4

Primer formation and helicase movement

C1 C1

R4 R I3 I3

R3 R

DnaB

R1

DnaC

DnaB–DnaC complex

2

Loading of DnaB–DnaC

DnaC

5

Extension of primers coordinated with helicase movement

R1

R1

Primase (DnaG)

3

6

Primase binds to DnaB

Leading strand synthesis by DNA polymerase III Primers for Okazaki fragments Okazaki fragment synthesis by DNA polymerase III

R1

Fig. 3 Assembly of DnaA, and the DnaB–DnaC complex at E. coli oriC, and activation of DnaB is a step-wise process. Step 1: DnaA recognizes specific DNA elements in E. coli oriC (see Fig. 1 and the text) to assemble a DnaA oligomer at this site. The numbers in the figure of DnaA represent its functional domains. Upon assembly of DnaA-ATP as a selfoligomer, it induces unwinding of a region containing the 13mers by binding to the top strand. At this step, domain IV of DnaA binds to the DnaA boxes and presumably to C-, I-, and τ-sites whereas domain I and III of a DnaA monomer interacts with another to assemble the DnaA oligomer. The interaction of HU with domain I of DnaA stabilizes the DnaA oligomer, but this interaction is not shown for simplicity. Step 2: Domain I of DnaA interacts with the N-terminal domain of DnaB in the DnaB6–DnaC3 complex to load this complex onto each DNA strand of the unwound region. Step 3: Primase interacts with the N-terminal domain of DnaB, which is required for primer synthesis. Step 4: As primase synthesizes a primer (shown as red (dark gray in the print version) squiggles) on the top and bottom strands, this event is coupled with the translocation of DnaB and the dissociation of DnaC from DnaB. Step 5: Primase completes primer synthesis and will then dissociate from DnaB after the primer is transferred to DNA polymerase III holoenzyme. DnaB moves on each single-stranded DNA toward the junction of each replication fork. Step 6: DNA polymerase III holoenzyme (not shown) extends these primers for the synthesis of each leading strand. During this elongation stage of DNA replication, DnaB at the junction of each replication fork will act as a DNA helicase to unwind the parental duplex DNA. The translocation of DnaB and its transient interaction with primase leads to the synthesis of subsequent primers on the top and bottom DNA strands. These primers are used by DNA polymerase III holoenzyme for the synthesis of Okazaki fragments.

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showed that its binding to DnaB attenuates its ATPase and helicase activities [152]. As additional support of the inhibitory effect of DnaC on DnaB, elevated ratios of DnaC to DnaB beyond the optimum for in vitro DNA replication of an oriC-containing plasmid is inhibitory [153]. Likewise, overproduction of DnaC in vivo causes inviability [140,153], and inhibits replication fork movement [154]. Hence, DnaC negatively regulates the helicase activity of DnaB, but DnaC evidently provides an essential function at an event prior to the binding of DnaB to DNA. Other studies demonstrate that DnaC binds adenine nucleotides, which modulate its function [137,138,155]. Specifically, ATP stimulates its binding to single-stranded DNA [138,139,144,156,157]. However, ATP or ADP is not needed for DnaC to interact with DnaB [138,139], nor does a nucleotide bound to DnaB dramatically affect the strength of this interaction [136]. Despite the work summarized above, ATP bound to DnaC is essential for DnaC function at oriC [136,138,139]. For example, an arginine substitution of the conserved lysine in the Walker A box caused undetectable ATP binding and inactivity in DNA replication of an oriC-containing plasmid [138]. In support, genetic evidence showed that missense mutations in each AAA+ motif interfere with DnaC function [140,158]. Together with the results of the preceding paragraph, these findings strongly suggest that ATP binding by DnaC is essential at a step after formation of the DnaB– DnaC complex. Considering that the structure of DnaB is a closed ring, how does ring opening occur so that the DnaB–DnaC complex can bind to the unwound region of oriC, and is ATP binding by DnaC required at the step of helicase loading? Insight into the role of ATP is suggested by the X-ray crystallographic structure of the ATP binding domain of A. aeolicus DnaC bound to ADP-BeF3, an ATP mimetic [139]. The results support a model of DnaC self-oligomerized as a right-handed helical filament. 3D reconstruction of electron microscopic images of negatively stained DnaB compared with the DnaB–DnaC complex and biochemical studies suggest that DnaC assembled as a helical filament onto DnaB opens the DnaB ring [129]. When the DnaB–DnaC complex binds to the unwound DNA within oriC, it is attractive to consider that the single-stranded DNA passes through the gap in the ring. In contrast, hydrogen-deuterium exchange studies of the DnaB–DnaC complex in comparison with these proteins alone support a different conclusion [141]. Specific predictions can be made for DnaB and the DnaB–DnaC complex. If DnaB is a closed ring, the interface between its protomers is expected to be occluded. In the open ring form

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of DnaB in the DnaB–DnaC complex, one interface relative to the remainder should exchange more readily. However, results from hydrogendeuterium exchange experiments indicated that the interface between protomers of the DnaB hexamer opens and closes spontaneously, and that DnaC traps DnaB in the open ring conformation [141].

3.6 Helicase Activation Requires the Dissociation of DnaC from DnaB A proposed mechanism for loading of the DnaB–DnaC complex at oriC involves occupancy of the interior channel of the helical DnaC filament by the unwound DNA of oriC [129]. An interaction between DnaA oligomerized at oriC and protomers of DnaC in the helical filament is suggested to mediate helicase loading [139]. DnaC must then dissociate from DnaB to reveal its helicase activity. Apparently, the interaction of the N-terminal domain of DnaB with primase and primer formation induces a conformational change in the C-terminal domain of DnaB where DnaC is bound, leading to the dissociation of DnaC from DnaB [144]. This dissociation appears to be coordinated with hydrolysis of ATP bound to DnaC on the basis of the following evidence. With wild-type DnaC bound to ATP but not ATPγS, DnaC is able to dissociate from DnaB, indicating that ATP hydrolysis is required for its release [144]. Like other members of the AAA+ superfamily of ATPases, DnaC contains a highly conserved arginine named the arginine finger in a conserved amino acid motif named box VII. This residue is proposed to coordinate ATP hydrolysis with a conformational change. In experiments that characterize a mutant protein bearing an alanine substitution for the arginine finger residue, it unlike wild-type DnaC failed to dissociate from DnaB under conditions that support the interaction of primase with DnaB and primer formation. Hence, this mutant is either unable to hydrolyze ATP or sense whether ATP has been hydrolyzed, so one or both activities are evidently required for DnaC to dissociate from DnaB. Whereas DnaC is essential in E. coli for the DnaB–DnaC complex to be placed at oriC, not all bacteria carry a dnaC gene. As an example, H. pylori does not encode dnaC. Like the replicative helicases found in some archaea, eukaryotic cells, and some viruses, the helicase of H. pylori is a dodecamer formed by the arrangement of two hexamers via their respective N-terminal domains [159,160]. Using an E. coli dnaC(Ts) mutant, a plasmid encoding H. pylori dnaB but not one bearing E. coli dnaB was able to complement the dnaC mutant at nonpermissive temperature, suggesting that

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this heterologous DnaB can bypass the requirement for DnaC in vivo [161,162]. Presumably, the interaction of E. coli DnaA with H. pylori DnaB mediates loading of the helicase at oriC. Recent experiments showed that the binding of H. pylori DnaB to single-stranded DNA requires ATP, but that ATP hydrolysis and ssDNA binding are not sufficient to separate the hexamers of the dodecamer [160]. Instead, its interaction with H. pylori primase causes the dodecamer to dissociate into individual hexamers as measured by multiangle light scattering and small angle X-ray scattering. Of interest, the interface joining the two hexamers in the X-ray crystallographic structure of H. pylori DnaB partially obscures the primase interaction site described in G. stearothermophilus DnaB [82]. Somehow, this interface becomes available for primase to interact, leading to dissociation of the dodecamer.

3.7 Events Leading to Assembly of the Replisome In the model described in Fig. 3, DnaA assembled at oriC loads a single DnaB–DnaC complex on each strand of the unwound region. The interaction of primase with DnaB and primer formation then leads to the dissociation of DnaC and activation of DnaB. To elaborate on the interaction of primase with DnaB, this process is transient in that primase remains associated with DnaB during primer synthesis but then dissociates after primer synthesis is complete [163–166]. The 30 -end of the primer, now available after primase has left, is extended by DNA polymerase III holoenzyme as it synthesizes the leading strand in concert with the translocation of DnaB as it unwinds the parental duplex DNA (reviewed in Refs. [167,168]). Positioned at the apex of each replication fork, DnaB moves in the 50 -to-30 direction relative to the DNA strand to which it is bound. DnaB is subsequently recognized by primase, leading to the synthesis of another primer. This primer and those synthesized later are used for the synthesis of Okazaki fragments.

4. SUMMARY Several lines of evidence indicate that DnaA assembles as a selfoligomer. In addition, the adenine nucleotide bound to DnaA, the interaction between and possibly among DnaA protomers of the oligomer, and DNA motifs to which these protomers are bound critically affect the assembly process. IHF bound to its cognate recognition site and Fis presumably at its binding site in oriC as well as other proteins that interact with DnaA via its

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N-terminal domain I also affect DnaA assembly at oriC, which is required for DnaA to unwind the AT-rich DUE element. DnaA then places the DnaB– DnaC complex onto each unwound strand within oriC and models have been proposed to explain the biochemical mechanism. Following helicase loading, activation of DnaB involves the dissociation of DnaC that appears to be induced by the interaction of primase with DnaB and primer formation. As primase and DnaC interact with the N-terminal and C-terminal domains of DnaB, respectively, a presumed conformational change in DnaB induced by primase binding apparently causes the dissociation of DnaC. Although structures of these proteins from thermophilic bacteria are known, the conformational change in DnaB has not been determined.

ACKNOWLEDGMENTS This work was supported by grant RO1 GM090063 from the National Institutes of Health, and by the USDA National Institute of Food and Agriculture, Hatch project number MICL02370, and Michigan AgBioResearch.

REFERENCES [1] A.C. Leonard, J.E. Grimwade, The orisome: structure and function, Front. Microbiol. 6 (2015) 545. [2] R.E. Bird, J. Louarn, J. Martuscelli, L. Caro, Origin and sequence of chromosome replication in Escherichia coli, J. Mol. Biol. 70 (1972) 549–566. [3] M. Masters, P. Broda, Evidence for the bidirectional replications of the Escherichia coli chromosome, Nat. New Biol. 232 (1971) 137–140. [4] R.C. Marsh, A. Worcel, A DNA fragment containing the origin of replication of the Escherichia coli chromosome, Proc. Natl. Acad. Sci. U.S.A 74 (1977) 2720–2724. [5] K. von Meyenburg, F.G. Hansen, L.D. Nielsin, E. Riise, Origin of replication, oriC, or the Escherichia coli chromosome on specialized transducing phages lambda asn, Mol. Gen. Genet. 160 (1978) 287–295. [6] K. von Meyenburg, F.G. Hansen, E. Riise, H.E. Bergmans, M. Meijer, W. Messer, Origin of replication, oriC, of the Escherichia coli K12 chromosome: genetic mapping and minichromosome replication, Cold Spring Harb. Symp. Quant. Biol. 43 (1979) 121–128. [7] S. Yasuda, Y. Hirota, Cloning and mapping of the replication origin of Escherichia coli, Proc. Natl. Acad. Sci. U.S.A 74 (1977) 5458–5462. [8] T. Miki, S. Hiraga, T. Nagata, T. Yura, Bacteriophage lambda carrying the Escherichia coli chromosomal region of the replication origin, Proc. Natl. Acad. Sci. U.S.A 75 (1978) 5099–5103. [9] W. Messer, M. Meijer, H.E. Bergmans, F.G. Hansen, K. von Meyenburg, E. Beck, H. Schaller, Origin of replication, oriC, of the Escherichia coli K12 chromosome: nucleotide sequence, Cold Spring Harb. Symp. Quant. Biol. 43 (1979) 139–145. [10] M. Matsui, A. Oka, M. Takanami, S. Yasuda, Y. Hirota, Sites of dnaA protein-binding in the replication origin of the Escherichia coli K-12 chromosome, J. Mol. Biol. 184 (1985) 529–533.

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[124] R. Galletto, M.J. Jezewska, W. Bujalowski, Unzipping mechanism of the doublestranded DNA unwinding by a hexameric helicase: the effect of the 3’ arm and the stability of the dsDNA on the unwinding activity of the Escherichia coli DnaB helicase, J. Mol. Biol. 343 (2004) 101–114. [125] L.E. Donate, O. Llorca, M. Barcena, S.E. Brown, N.E. Dixon, J.M. Carazo, pHcontrolled quaternary states of hexameric DnaB helicase, J. Mol. Biol. 303 (2000) 383–393. [126] S. Yang, X. Yu, M.S. VanLoock, M.J. Jezewska, W. Bujalowski, E.H. Egelman, Flexibility of the rings: structural asymmetry in the DnaB hexameric helicase, J. Mol. Biol. 321 (2002) 839–849. [127] B. Liu, W.K. Eliason, T.A. Steitz, Structure of a helicase-helicase loader complex reveals insights into the mechanism of bacterial primosome assembly, Nat. Commun. 4 (2013) 2495. [128] O. Itsathitphaisarn, R.A. Wing, W.K. Eliason, J. Wang, T.A. Steitz, The hexameric helicase DnaB adopts a nonplanar conformation during translocation, Cell 151 (2012) 267–277. [129] E. Arias-Palomo, V.L. O’Shea, I.V. Hood, J.M. Berger, The bacterial DnaC helicase loader is a DnaB ring breaker, Cell 153 (2013) 438–448. [130] M.J. Jezewska, S. Rajendran, W. Bujalowski, Complex of Escherichia coli primary replicative helicase DnaB protein with a replication fork: recognition and structure, Biochemistry 37 (1998) 3116–3136. [131] E.J. Enemark, L. Joshua-Tor, Mechanism of DNA translocation in a replicative hexameric helicase, Nature 442 (2006) 270–275. [132] N.D. Thomsen, J.M. Berger, Running in reverse: the structural basis for translocation polarity in hexameric helicases, Cell 139 (2009) 523–534. [133] A.Y. Lyubimov, M. Strycharska, J.M. Berger, The nuts and bolts of ring-translocase structure and mechanism, Curr. Opin. Struct. Biol. 21 (2011) 240–248. [134] S. Wickner, J. Hurwitz, Interaction of Escherichia coli dnaB and dnaC(D) gene products in vitro, Proc. Natl. Acad. Sci. U.S.A 72 (1975) 921–925. [135] J.A. Kobori, A. Kornberg, The Escherichia coli dnaC gene product. III. Properties of the dnaB-dnaC protein complex, J. Biol. Chem. 257 (1982) 13770–13775. [136] R. Galletto, M.J. Jezewska, W. Bujalowski, Interactions of the Escherichia coli DnaB helicase hexamer with the replication factor the DnaC protein. Effect of nucleotide cofactors and the ssDNA on protein-protein interactions and the topology of the complex, J. Mol. Biol. 329 (2003) 441–465. [137] E. Wahle, R.S. Lasken, A. Kornberg, The dnaB-dnaC replication protein complex of Escherichia coli. I. Formation and properties, J. Biol. Chem. 264 (1989) 2463–2468. [138] M.J. Davey, L. Fang, P. McInerney, R.E. Georgescu, M. O’Donnell, The DnaC helicase loader is a dual ATP/ADP switch protein, EMBO J. 21 (2002) 3148–3159. [139] M.L. Mott, J.P. Erzberger, M.M. Coons, J.M. Berger, Structural synergy and molecular crosstalk between bacterial helicase loaders and replication initiators, Cell 135 (2008) 623–634. [140] A.V. Ludlam, M.W. McNatt, K.M. Carr, J.M. Kaguni, Essential amino acids of Escherichia coli DnaC protein in an N-terminal domain interact with DnaB helicase, J. Biol. Chem. 276 (2001) 27345–27353. [141] S. Chodavarapu, A.D. Jones, M. Feig, J.M. Kaguni, DnaC traps DnaB as an open ring and remodels the domain that binds primase, Nucleic Acids Res. 44 (2015) 210–220. [142] L. Fang, M.J. Davey, M. O’Donnell, Replisome assembly at oriC, the replication origin of E. coli, reveals an explanation for initiation sites outside an origin, Mol. Cell 4 (1999) 541–553. [143] K.M. Carr, J.M. Kaguni, Stoichiometry of DnaA and DnaB protein in initiation at the Escherichia coli chromosomal origin, J. Biol. Chem. 276 (2001) 44919–44925.

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[144] M. Makowska-Grzyska, J.M. Kaguni, Primase directs the release of DnaC from DnaB, Mol. Cell 37 (2010) 90–101. [145] J. Marszalek, J.M. Kaguni, DnaA protein directs the binding of DnaB protein in initiation of DNA replication in Escherichia coli, J. Biol. Chem. 269 (1994) 4883–4890. [146] M.D. Sutton, K.M. Carr, M. Vicente, J.M. Kaguni, E. coli DnaA protein: the N-terminal domain and loading of DnaB helicase at the E. coli chromosomal origin, J. Biol. Chem. 273 (1998) 34255–34262. [147] H. Seitz, C. Weigel, W. Messer, The interaction domains of the DnaA and DnaB replication proteins of Escherichia coli, Mol. Microbiol. 37 (2000) 1270–1279. [148] P.L. Carl, Escherichia coli mutants with temperature-sensitive synthesis of DNA, Mol. Gen. Genet. 109 (1970) 107–122. [149] W.H. Schubach, J.D. Whitmer, C.I. Davern, Genetic control of DNA initiation in Escherichia coli, J. Mol. Biol. 74 (1973) 205–221. [150] J.A. Wechsler, Genetic and phenotypic characterization of dnaC mutations, J. Bacteriol. 121 (1975) 594–599. [151] H.L. Withers, R. Bernander, Characterization of dnaC2 and dnaC28 mutants by flow cytometry, J. Bacteriol. 180 (1998) 1624–1631. [152] E. Wahle, R.S. Lasken, A. Kornberg, The dnaB-dnaC replication protein complex of Escherichia coli. II. Role of the complex in mobilizing dnaB functions, J. Biol. Chem. 264 (1989) 2469–2475. [153] G.J. Allen, A. Kornberg, Fine balance in the regulation of DnaB helicase by DnaC protein in replication in Escherichia coli, J. Biol. Chem. 266 (1991) 22096–22101. [154] K. Skarstad, S. Wold, The speed of the Escherichia coli fork in vivo depends on the DnaB:DnaC ratio, Mol. Microbiol. 17 (1995) 825–831. [155] R. Galletto, S. Rajendran, W. Bujalowski, Interactions of nucleotide cofactors with the Escherichia coli replication factor DnaC protein, Biochemistry 39 (2000) 12959–12969. [156] S.B. Biswas, S. Flowers, E.E. Biswas-Fiss, Quantitative analysis of nucleotide modulation of DNA binding by the DnaC protein of Escherichia coli, Biochem. J. 379 (2004) 553–562. [157] B.A. Learn, S.J. Um, L. Huang, R. McMacken, Cryptic single-stranded-DNA binding activities of the phage lambda P and Escherichia coli DnaC replication initiation proteins facilitate the transfer of E. coli DnaB helicase onto DNA, Proc. Natl. Acad. Sci. U.S.A 94 (1997) 1154–1159. [158] K. Hupert-Kocurek, J.M. Sage, M. Makowska-Grzyska, J.M. Kaguni, Genetic method to analyze essential genes of Escherichia coli, Appl. Environ. Microbiol. 73 (2007) 7075–7082. [159] M. Stelter, I. Gutsche, U. Kapp, A. Bazin, G. Bajic, G. Goret, M. Jamin, J. Timmins, L. Terradot, Architecture of a dodecameric bacterial replicative helicase, Structure 20 (2012) 554–564. [160] A. Bazin, M.V. Cherrier, I. Gutsche, J. Timmins, L. Terradot, Structure and primasemediated activation of a bacterial dodecameric replicative helicase, Nucleic Acids Res. 43 (2015) 8564–8576. [161] R.K. Soni, P. Mehra, N.R. Choudhury, G. Mukhopadhyay, S.K. Dhar, Functional characterization of Helicobacter pylori DnaB helicase, Nucleic Acids Res. 31 (2003) 6828–6840. [162] R.K. Soni, P. Mehra, G. Mukhopadhyay, S.K. Dhar, Helicobacter pylori DnaB helicase can bypass Escherichia coli DnaC function in vivo, Biochem. J. 389 (2005) 541–548. [163] K. Tougu, K.J. Marians, The interaction between helicase and primase sets the replication fork clock, J. Biol. Chem. 271 (1996) 21398–21405. [164] K. Tougu, K.J. Marians, The extreme C terminus of primase is required for interaction with DnaB at the replication fork, J. Biol. Chem. 271 (1996) 21391–21397.

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[165] P. Chang, K.J. Marians, Identification of a region of Escherichia coli DnaB required for functional interaction with DnaG at the replication fork, J. Biol. Chem. 275 (2000) 26187–26195. [166] L.E. Bird, H. Pan, P. Soultanas, D.B. Wigley, Mapping protein-protein interactions within a stable complex of DNA primase and DnaB helicase from Bacillus stearothermophilus, Biochemistry 39 (2000) 171–182. [167] C.S. McHenry, Bacterial replicases and related polymerases, Curr. Opin. Chem. Biol. 15 (2011) 587–594. [168] N.Y. Yao, M. O’Donnell, SnapShot: the replisome, Cell 141 (2010). 1088, 1088 e1081.

CHAPTER TWO

The E. coli DNA Replication Fork J.S. Lewis, S. Jergic, N.E. Dixon1 Centre for Medical & Molecular Bioscience, University of Wollongong, Wollongong, NSW, Australia 1 Corresponding author: e-mail address: [email protected]

Contents 1. 2. 3. 4. 5.

Introduction Initiation at oriC, the Origin of Replication The Primosome, DnaB, and DnaG Role of SSB at the Replication Fork The Elongation Stage 5.1 Assembly and Action of the DNA Polymerase III Holoenzyme 5.2 Dynamics on the Lagging Strand 6. Concluding Comments Acknowledgments References

32 36 36 41 43 44 60 68 69 69

Abstract DNA replication in Escherichia coli initiates at oriC, the origin of replication and proceeds bidirectionally, resulting in two replication forks that travel in opposite directions from the origin. Here, we focus on events at the replication fork. The replication machinery (or replisome), first assembled on both forks at oriC, contains the DnaB helicase for strand separation, and the DNA polymerase III holoenzyme (Pol III HE) for DNA synthesis. DnaB interacts transiently with the DnaG primase for RNA priming on both strands. The Pol III HE is made up of three subassemblies: (i) the αεθ core polymerase complex that is present in two (or three) copies to simultaneously copy both DNA strands, (ii) the β2 sliding clamp that interacts with the core polymerase to ensure its processivity, and (iii) the seven-subunit clamp loader complex that loads β2 onto primer–template junctions and interacts with the α polymerase subunit of the core and the DnaB helicase to organize the two (or three) core polymerases. Here, we review the structures of the enzymatic components of replisomes, and the protein–protein and protein–DNA interactions that ensure they remain intact while undergoing substantial dynamic changes as they function to copy both the leading and lagging strands simultaneously during coordinated replication.

The Enzymes, Volume 39 ISSN 1874-6047 http://dx.doi.org/10.1016/bs.enz.2016.04.001

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2016 Elsevier Inc. All rights reserved.

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1. INTRODUCTION All the genetic information essential for the inheritance of traits that define the phenotypes of cells is packaged in the double helix of chromosomal DNA. A single DNA molecule from Escherichia coli contains just over 4.6 million base pairs (Mbp) [1] and is
1.5 mm in length, roughly 1000 times the length of an individual cell. Replication of chromosomal DNA occurs on a remarkable scale and is a fundamentally complex, yet carefully orchestrated process that requires the assembly and action of a molecular machine at regions of the DNA called replication forks. In 1963, John Cairns reported the first autoradiograph of a replicating [3H]-thymidine-labeled E. coli chromosome [2]. This was the first time that a bacterium was caught in the act of replicating its circular chromosomal DNA. In the replicating chromosomes, two replication forks were identified, representing the sites of active DNA synthesis. At each of them, parental DNA must first be unwound into two template strands that are copied simultaneously with a high degree of accuracy and with great efficiency. Furthermore, the antiparallel structure of DNA and the requirement that DNA polymerases make DNA by extending a preexisting primer from the 30 -OH end restrict the mechanisms that can be used at replication forks to copy DNA. At each fork, the two nascent DNA strands are necessarily synthesized by different processes, with the leading strand synthesized continuously while the lagging strand is synthesized discontinuously as short Okazaki fragments that are later joined [3,4]. Although the mechanisms on the two strands are different, their synthesis is believed to be coupled together, and to be carried out by the same replication apparatus, called the replisome. Replisomes are multiprotein molecular machines, evolved to coordinate all the necessary enzymatic activities for coupled DNA replication. The E. coli replisome is undoubtedly the best understood across all species, and is composed of over a dozen individual subunits, some of which are present in multiple copies (Fig. 1; Table 1). Once assembled, the E. coli replisome unwinds and duplicates DNA at remarkable speeds, approaching 1000 bp/s with an error rate of approximately one mutation for each 106 to 107 nucleotides incorporated (reviewed in Ref. [87]). These individual subunits interact and exchange in a hierarchy of strong and weak functional interactions (pairwise KD values range from low pM to high μM), the kinetic properties of which are tightly controlled by external conditions such as cellular protein concentrations and nucleotide

The E. coli Replication Fork

33

Pol III core

SSB

}

Leading strand

ψ

δ⬘

Replication fork

χ

δ τ

DnaB helicase

}

Lagging strand

Clamp loader complex (CLC)

DnaG primase

α Primer ε β2

θ

Fig. 1 Architecture of the E. coli replisome at the chromosomal replication fork derived from in vitro studies and direct observation in vivo. The DnaB helicase is located at the apex of the replication fork on the lagging strand where it uses the energy of ATP hydrolysis to unwind dsDNA. The lagging-strand template produced by helicase action is protected by SSB. The DNA polymerase III holoenzyme (Pol III HE) uses the single strands of DNA produced by DnaB as templates to synthesize new DNA on both the leading and lagging strands. The β2 sliding clamp confers high processivity on the DNA Pol III HE by tethering the Pol III αεθ cores onto the DNA. The clamp loader complex (CLC) uses ATP hydrolysis to assemble the β2 clamp onto RNA primer junctions on template DNA. Up to three Pol III cores are coupled through the τ subunits of the CLC through their extreme C-terminal domains, and the τ subunits also interact with DnaB, thus organizing and coupling the DNA Pol III HE to DnaB. Due to the opposite polarity of the two DNA strands, the lagging strand is synthesized in a series of short Okazaki fragments in the opposite direction to the leading strand. DnaG primases interact with DnaB to synthesize RNA primers to initiate DNA synthesis on the lagging strand, signaling the start of synthesis of an Okazaki fragment. When an Okazaki fragment is complete, Pol I replaces RNA primers with DNA and DNA ligase A joins the fragments into a contiguous lagging-strand DNA chain. Figure adapted from A. Robinson, A.M. van Oijen, Bacterial replication, transcription and translation: mechanistic insights from single-molecule biochemical studies, Nat. Rev. Microbiol. 11 (2013) 303–315 with permission.

availability. This hierarchy of functional interactions enables the replisome to transition through multiple conformational states to accomplish simultaneous, concerted copying of both DNA strands at each replication fork while its overall integrity is maintained [88].

Table 1 E. coli Replisome Components and Associated Functions Enzyme Subunit Molecules/Cell Gene Mass (kDa) Interaction Partners

Function

References

[5]

Pol III HE (αεθ)2–τ2γδδ0 ψχ–(β2)2 10a

790.7

Replicates E. coli chromosome

60b

165.8

DNA synthesis and [6,7] proofreading

Pol III core α

dnaE 129.9

τ, ε, β2, UmuD

DNA polymerase

[8–16]

ε

dnaQ 27.1

α, θ, β2

30 –50 exonuclease

[11–13,17–22]

θ

holE

ε

Stimulates [22–27] exonulease activity

Clamp loader

8.8

Loads β2 clamp, organizes Pol III HE

297.1

[28]

τ/γ

140

dnaX 71.1/47.5 δ, δ0 , α, τ/γ, ψ, DnaB ATPase, dimerizes Pol III cores

δ

930

holA 38.7

τ/γ, δ0 , β2

ATPase, opens β2 clamp

[30,35,37–39]

δ0

140

holB

τ/γ, δ

ATPase

[30,37,38]

χ

1200

holC 16.6

ψ, SSB

Bridges clamp loader to SSB

[40–43]

ψ

340

holD 15.2

χ, τ/γ

Stabilizes clamp loader

[38,40–42]

36.9

[14,29–36]

Sliding clamp

81.2

Tethers polymerase on DNA

β

300c

dnaN 40.6

Processivity factor α, δ, ε, Hda, MutS, MutL, ligase A, Pol I, Pol II, Pol IV, Pol V

DnaB

–

15–20

dnaB 52.4

DnaA, DnaC, DnaG, τ, Rep

50 –30 helicase, activates DnaG

[34,53–61]

DnaG

–

50–100

dnaG 65.6

DnaB, SSB

RNA primer synthesis

[43,62–67]

Pol I

–

400

polA 103.1

β2

Primer removal and [6,68,69] gap filling

Ligase A

–

300

lig

73.6

β2

Joins nascent DNA [6,69–72] fragments

SSB

–


1000–2000d ssb

18.8

DnaG, χ, TopB, UmuC, PriA, RecJ, RecO, RecQ, ExoI, ExoIX, GroEL

Protects ssDNA

a

May be up to 20 molecules/cell. Including 40 free Pol III cores molecules/cell. c As dimers of β. d As tetramers of SSB in wild-type E. coli cells during mid-log phase. b

[44–52]

[43,73–86]

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J.S. Lewis et al.

Our focus is on the E. coli replisome as a representative of those of other eubacteria. Recent work reviewed, for example in Ref. [89], indicates that all bacterial species use highly conserved replisomal components to carry out conserved functions [90,91], but a picture is emerging where the hierarchy of protein interactions varies such that interactions that are strong in some species may be weaker in others, with the necessity for a multiplicity of interactions to enable dynamics while preserving replisomal integrity being satisfied in different ways. Access to three-dimensional (3D) structural information for most of the individual proteins and domains of the E. coli replisome over the last 25 years, combined with the ability to faithfully reconstitute replisomal complexes on various DNA templates in vitro, has led to the discovery of many dynamic protein–protein and protein–DNA interactions at the replication fork. This chapter explores the assembly of replisomal subunits, their structures, and their dynamic behavior.

2. INITIATION AT oriC, THE ORIGIN OF REPLICATION Initiation of bacterial DNA replication occurs through highly coordinated assembly of nucleoprotein complexes at the unique origin of replication (oriC). DnaA is the specialist initiator protein that recognizes the origin and guides the formation of these nucleoprotein complexes [92–94]. To initiate DNA replication, this complex must be activated and undergo large conformational changes that result in the local unwinding of duplex DNA. DnaB, the ring-shaped replicative DNA helicase that unwinds double-stranded (ds) DNA, is opened and loaded onto the single-stranded (ss) DNA bubble (open complex) from a complex with the helicase loader, DnaC. This loading requires DnaB-bound DnaC and oriC-bound DnaA to form a prepriming complex that eventually results in dissociation of DnaC and loading of DnaB to encircle the ssDNA. The helicase must then migrate along the ssDNA, opening the region to expose enough ssDNA for DnaG primase to interact with it to produce an RNA primer for the replicative polymerase, the DNA polymerase III (Pol III) holoenzyme (HE), to assemble on each replication fork to complete replisome assembly. The mechanics of initiation of DNA replication at oriC are described in detail in chapter “Replication Initiation in Bacteria” by Chodavarapu and Kaguni [95].

3. THE PRIMOSOME, DnaB, AND DnaG DnaB is the replicative helicase and the first replisomal protein to assemble at the replication fork [96]. Following initial strand separation at

The E. coli Replication Fork

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oriC by DnaA, a DnaB molecule is loaded onto each of the single strands of the open complex. Once loaded, each DnaB initiates ATP hydrolysisdependent unwinding of the duplex parental strands by translocating in the 50 –30 direction on what will become the lagging-strand template strands of the two replication forks that will invade the duplex DNA on either side of the origin. The result of translocation of DnaB (at a rate up to
300 bp/s [97] in the absence of other replisomal proteins) is progressive separation of the parental dsDNA into two ssDNA templates [98]. In E. coli and all other eubacteria, DnaB is a toroid of six identical 52-kDa subunits oriented in the same direction. The formation of a functional E. coli homohexamer is Mg2+ dependent, unlike bacteriophage T4 helicase that requires binding of ATP or GTP for hexamerization [99]. DnaB has two domains. In E. coli, the N-terminal domain consists of a short, probably unstructured, region followed by a helical domain that although monomeric in isolation [57,58] forms a trimer of dimers in the hexamer to generate sites for interaction with up to three molecules of DnaG primase [59,65,100]. The C-terminal RecA-like motor domains of DnaB oligomerize as a ringshaped hexamer to generate potentially six catalytic sites for ssDNAdependent ATP hydrolysis [101,102]. Biochemical studies suggest that
20 nucleotides (nt) of ssDNA pass through the central channel of DnaB when it is loaded on a forked DNA molecule with a 50-nt ssDNA tail [103–105]. In this configuration, the C-terminal motor domains are at the apex of each replication fork, and the trailing N-terminal domains are positioned to interact with DnaG to synthesize RNA primers complementary to the strands on which they are translocating. The first primer synthesized on each strand at oriC becomes the leading-strand primer for the other replication fork, and subsequent primers initiate Okazaki fragment synthesis on the lagging strand. Despite there being no available high-resolution 3D structure of the functional DnaB hexamer from E. coli alone or in any protein complex, several structures of full-length versions or domains of DnaB and complexes from other bacterial species are available [100,106–109]. The recent cryoelectron microscopy (cryo-EM) structure of the E. coli DnaB6–(DnaC)6 complex determined at 25 A˚ resolution shows an open helical arrangement of DnaB–DnaC heterodimers, suggesting a mechanism for loading of the helicase at a nascent replication fork [110]. Early negative stain EM studies established the hexameric ring structure of E. coli DnaB6 but raised questions about the exact conformation of the ring. Both C3 and C6 rotational symmetries were observed regardless of the presence of nucleotide cofactors [101,111,112]. Reexamination of

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J.S. Lewis et al.

˚) DnaB alone with a bound ATP analog (ADPBeF3) by cryo-EM (at 25 A suggested C3 symmetry (in the N-terminal domains) produced shapes most closely matching the reference-free 2D class averages and known crystal structures of other DnaB homologs [110]. All of these structures show a wide open (dilated) central channel with the N-terminal domains appearing as a trimer of dimers (C3 symmetry) while the C-terminal RecA-like motor domains have pseudo-sixfold symmetry. Subsequently, the Berger group reported the crystal structure of Aquifex aeolicus DnaB, which showed a quite different arrangement of the N-terminal domains. Although their threefold symmetry is maintained, the central channel is severely constricted. This suggests that DnaB can adopt different arrangements of the N-terminal domains when it interacts with its different binding partners, and with different nucleotides, implying that transition between constricted and dilated structures may occur as DnaB translocates on ssDNA [113]. A much earlier structure of the N-terminal domain of DnaB (residues Lys24–Ser136) determined in solution by NMR [58] and the crystal structure of a similar domain construct [57] showed a dimer interface that assembles around Phe102, which was initially thought to contribute to dimerization of the N-terminal domain in the C3 symmetrical state of the DnaB hexamer [114]. However, the crystal structures of full-length DnaB homologs and the recent cryo-EM structures disprove this, and the existence of the C6 structure in the early micrographs appears now to be an artifact. In fact, Phe102 is now predicted to be a part of the interface with the DnaG primase (Fig. 2B, inset). The translocating helicase interacts through its N-terminal domains with the DnaG primase, the specialist RNA polymerase that synthesizes short RNA primers on the ssDNA regions produced by helicase action [115–117]. In E. coli, the DnaB–DnaG interaction is weak and transient, though in some other species like the Firmicute Geobacillus stearothermophilus, it is stable and the complex can be crystallized [100] (Fig. 2). Bacterial DnaG primases contain three domains that are linked together by regions predicted to be flexible: the N-terminal zinc-binding domain (ZBD), the central catalytic RNA polymerase domain (RPD), and the C-terminal helicase-binding domain (HBD or DnaGC). The ZBD contains a zincribbon motif, which coordinates a Zn2+ ion [118]. The ZBD is essential for primase activity and is thought to recognize trinucleotide priming sequences in the ssDNA template [119,120]. The bacterial RPD domains share high structural similarity with types IA and II topoisomerases and primase-like proteins (TOPRIM fold) [121,122], unlike the evolutionarily

The E. coli Replication Fork

39

A

B

Phe102

90 degrees

Fig. 2 Homology model of the E. coli DnaB6–(DnaGC)3 primosomal complex based on the corresponding structure from Geobacillus stearothermophilus [100]. (A) Top view of a cartoon representation of the DnaB6–(DnaGC)3 complex showing the three DnaG molecules (red (dark gray in the print version)) bound at the periphery of DnaB's N-terminal domains (blue (light gray in the print version)). (B) Side view, rotated 90 degrees around the y-axis of the view in (A). The inset in (B) shows a zoomed view of the predicted interface between DnaGC (red (dark gray in the print version)) and DnaB (light blue (light gray in the print version)), showing the position of Phe102 of DnaB (purple (gray in the print version) sticks). From a homology model prepared in collaboration with Thomas Huber, Australian National University. This and all later figures were prepared using PyMOL v1.8.0.5 (Schro€dinger).

and structurally distinct archaeo-eukaryotic primase RPD domains [123]. Within the RPD are two conserved catalytic motifs [64], one containing a conserved Glu presumed to act as a general base during ribonucleotide incorporation. The other contains two strictly conserved Asp (DxD) residues required for Mg2+-mediated NTP binding [124,125]. DnaG primase binds through the DnaGC domain to the N-terminal domain of the DnaB helicase to form the primosomal complex DnaB6–(DnaG)3 [59,65]. Although the ZBD and RPD are sufficient to catalyze template-directed oligonucleotide synthesis in phage T7, E. coli, and A. aeolicus systems [126–128], coordination of all three domains is required to allow effective initiation of de novo RNA primer synthesis. DnaGs preferentially utilize defined trinucleotide sequences in DNA. For E. coli, this is 50 -dCTG in the leading-strand and 50 -dC(A/T)G in the lagging-strand templates [119,129,130]. Trinucleotide specificity varies somewhat among species, eg, primases from Firmicutes preferentially initiate synthesis from 50 -dCTA [131]. Acting alone, DnaG is slow to synthesize a 12–16-nt RNA primer once every 1000 s in vitro [131,132], yet much higher rates are required to support in vivo Okazaki fragment synthesis. The presence of DnaB

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stimulates DnaG activity to in vivo rates and limits the length of RNA primers to 10–14 nucleotides with 12 nucleotides being the predominant RNA primer length [130]. The ZBD recognizes and interacts with the trinucleotide recognition sequences, bringing the templates in close proximity to the RPD for RNA primer synthesis. This RNA priming activity is specifically stimulated by direct interaction with the N-terminal domain of DnaB through DnaGC [130]. Although E. coli DnaG has been shown to be monomeric [118], it is thought to bind to template DNA as a dimer [133] and to function in trans with another DnaG protomer bound to the DnaB hexamer, ie, the ZBD of one protomer recognizes the priming site in the DNA template while the RPD of the other protomer synthesizes the primer [127]. There is no high-resolution structural information on a full-length primase from any eubacterial species, but structures of each of the individual domains of DnaG have been determined either by X-ray crystallography or NMR spectroscopy. These structures include the RPD and DnaGC domains from E. coli and other bacterial species, as well as the ZBD from G. stearothermophilus and RPD/ZBD from A. aeolicus [36,59,64,122,127,134,135]. Without a full-length DnaG primase structure, how each of the three domains in DnaG cooperates to enable efficient DnaB-dependent primer synthesis remains speculative. Following RNA primer synthesis, the next step in replisome assembly at the fork requires the loading of the replicative polymerase, the Pol III HE at the primer termini. The polymerase never needs to dissociate from the template as it continuously and processively extends the first primer laid down on the leading strand at oriC. The situation on the other strand is quite different: replication of the lagging strand needs to occur discontinuously, generating Okazaki fragments 1–2 kb in length that need to be processed by DNA polymerase I and joined by DNA ligase [6]. This means that lagging strand, unlike leading-strand synthesis, generates considerable amounts of ssDNA, which is coated by single-stranded DNA-binding protein (SSB). The lagging-strand ssDNA is produced by helicase action, and consumed by DNA synthesis by the polymerase, and in a fully coupled situation where the rates of the two enzymes should be near identical, the amount of ssDNA at each replication fork would be nearly constant at about half of the average Okazaki fragment length, or 0.5–1 kb (Fig. 1). Thus, the amount of SSB at each fork should also be relatively constant. It is appropriate at this stage to consider the properties of SSB and its still incompletely understood roles at the replication fork.

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4. ROLE OF SSB AT THE REPLICATION FORK The basic function of SSB is to bind preferentially to ssDNA with high affinity in a sequence-independent manner [74]. By doing this it is able to protect ssDNA from nucleolytic digestion and prevent intrastrand pairing such as hairpin formation, so as to preserve ssDNA in a conformation suitable for the action of DNA metabolic enzymes [84]. But SSB is also a pivotal interaction hub, interacting with a large number of proteins, directing and organizing them to sites of DNA replication, recombination, and repair (reviewed in Ref. [84]). Historically SSB has been viewed as only providing inert protection to regions of ssDNA. However, there is increasing evidence that SSB–ssDNA complexes are highly dynamic and have great functional significance. To illustrate, SSB is able to rapidly sample alternative binding topologies without dissociation [136,137], and to utilize a direct transfer mechanism between ssDNAs, hypothesized to enable recycling of SSB tetramers from old to new lagging-strand templates at the replication fork [138]. Earlier studies also demonstrated SSB’s role in RNA priming by DnaB–DnaG, which occurs randomly on naked ssDNA but is restricted, eg, to defined origins of replication on SSB-coated single-stranded phage DNAs [139]. In E. coli, SSB is the product of the ssb gene, which is essential for cell viability [73,140,141]. SSB forms a stable homotetramer of 177 amino acid subunits [73], separated into two distinct domains. The N-terminal domain (112 residues) is structured and forms a classic oligonucleotide (oligosaccharide)-binding (OB) fold responsible for ssDNA binding [78,142,143]. Several X-ray crystal structures of the OB domain of E. coli SSB have been solved (Fig. 3), both in isolation and bound to two 35-mer ssDNAs per tetramer [78,144,145]. The four ssDNA-binding domains in the tetramer enable it to bind tightly to ssDNA in a variety of modes with different binding properties depending on the monovalent or divalent cation concentrations, and input stoichiometry. The dominant binding modes observed in in vitro studies are referred to as SSB65, SSB56, and SSB35 [146,147]. The subscript number reflects the average number of nucleotides bound by each tetramer [13,148–152]. The SSB65 mode (Fig. 3B), favored by moderately high salt concentration (>2 mM Mg2+ or >200 mM Na+), utilizes all four ssDNA-binding sites and exhibits little cooperativity between neighboring ssDNA-bound tetramers [78,153–155]. The SSB35 binding mode, on the other hand is favored in low salt concentrations (120 nt/s.

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The second “active” model proposes that the helicase destabilizes dsDNA by using the energy of nucleotide hydrolysis to move forward and displace dsDNA at the fork. The above two models were tested in a single-molecule assay by measuring rates of gp4 translocation along ssDNA and unwinding of dsDNA [150]. The observed dependence of unwinding rates on DNA sequence agrees with the active model. The kinetic data demonstrate that the average number of base pairs unwound per molecule of dTTP hydrolyzed is 1–2 for a GC-rich and 3–4 for an AT-rich regions [143]. Single-molecule studies of DNA unwinding indicate that T4 helicase functions as a passive motor [151].

5.3 Single-Stranded DNA-Binding Protein T7 phage gene 2.5 encodes a single-strand DNA-binding protein gp2.5 (Fig. 1). Through interactions with gp5 [91,93,152,153] and gp4 [91,154,155] gp2.5 coordinates simultaneous synthesis of leading- and lagging-strands [156], described in the sections later. In addition, gp2.5 facilitates homologous DNA base pairing [149,157], a step involved in the formation of T7 concatemers. Gp2.5 also plays a role in recombination [149,158,159], and in the repair of double-stranded breaks in phage DNA [160]. The crystal structure of gp2.5 [65] presented in Fig. 9 reveals that the core of the protein adopts an oligosaccharide/oligonucleotidebinding fold (OB-fold), the structural feature common in proteins that

Fig. 9 Crystal structure of gene 2.5 single-strand DNA-binding protein. The structure of gp2.5 contains a five-stranded antiparallel β-barrel (OB-fold) shown in yellow (gray in the print version); the structural motif characteristic for ssDNA-binding proteins. The two alpha-helices (αA and αB) capping the OB-fold are shown in blue (dark gray in the print version). An acidic C-terminal tail that participates in dimer formation and interactions with other replication proteins and DNA protrudes away from the OB-fold and αB.

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function by binding to ssDNA [161,162]. Interestingly, despite structural and functional resemblance between T7 gp2.5 and other ssDNA-binding proteins, for instance E. coli SSB protein and T4 gene 32 protein, the above proteins lack any significant amino-acid sequence homology [163]. Another important feature of gp2.5 is an acidic C-terminal tail. The essential protein– DNA and protein–protein interactions of gp2.5 with gp4 and gp5 are mediated in part by the C-terminal tail. 5.3.1 Gp2.5 Binding to ssDNA In the absence of ssDNA, gp2.5 forms stable dimers in solution [164]. Removal of the C-terminal 21 residues alters the dimerization properties of gp2.5; the protein in solution is now monomeric and binds to ssDNA with a higher affinity [155]. Although the protein construct used for structure determination lacked the C-terminal 26 residues, it crystallized with two protomers in the crystallographic asymmetric unit in the trans orientation. It is believed that the protein arrangement in the crystal reflects the native gp2.5 dimer in solution [65]. The binding geometry of protomers, and specifically the orientation of the adjoining residues, suggests an extended conformation of the C-terminal tail. In such a case, the C-terminal tails of the two adjacent protomers could stabilize the dimer through a domain-swapping arrangement involving electrostatic interactions between acidic residues of the C-terminal tail and basic residues on the surface of the OB-fold. Consequently, the interaction of the C-terminal tail with the DNA binding site of gp2.5 could modulate its affinity for ssDNA. A similar mechanism was proposed for the acidic C-terminal tail of the T4 gene 32 single-stranded binding protein [165]. In the model, the binding of ssDNA displaces the tail from the DNA binding site on gp2.5. Conversely, the displacement of the C-terminal tail can increase the affinity for binding of gp2.5 to gp4 or gp5. The C-terminal tail is sensitive to a proteolytic digestion in gp2.5–DNA complex suggesting that it might be available for other interactions. The above model was confirmed by a crosslinking analysis [166]. The analysis revealed a covalent binding of the residues from the C-terminal tail and the residues located in the OB-fold of gp2.5 [166]. The presence of ssDNA in the reaction mixtures abolishes the above cross-linking pattern. Also, the cross-linked proteins lose their affinity to ssDNA. Alteration of residues in the OB-fold reduces the affinity of gp2.5 for ssDNA [167]. The above model is confirmed by NMR chemical shift mapping analysis. NMR revealed that both the C-terminal tail and ssDNA associate with the same part of gp2.5 surface [166]. Interestingly,

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E. coli SSB protein and T4 gp32 also have acidic C-terminal tails that modulate their interactions with other replication proteins and ssDNA [165,168–173]. 5.3.2 Interactions of gp2.5 with Other Replication Proteins As indicated earlier, the removal of the C-terminal tail from gp2.5 eliminates its interaction with gp5/trx [91,93,152] and gp4 [155]. The 15 out of 26 residues present in the C-terminal tail are aspartic or glutamic acids. Substitution of these acidic residues with basic or neutral amino acids eliminates the binding of gp2.5 to gp4 or gp5/trx [152]. Interestingly, the last C-terminal residue in the tail is phenylalanine. Removal of phenylalanine 232 or its replacement with a nonaromatic amino-acid eliminates an interaction of gp2.5 with gp5/trx [152]. The C-terminal tail of gp2.5 binds to the two basic loops [93] located within the TBD of gp5 to which trx also binds [75]. A screening for suppressors of gp2.5 lacking the C-terminal phenylalanine identified residues that map in the DNA-binding cleft of gp5/trx [174]. The altered gp2.5 lacking the C-terminal phenylalanine has a lower affinity for gp5/trx relative to the wild-type protein. The binding of the altered gp2.5 and gp5/trx is partially restored by the suppressor mutations in gp5 [175]. As described in previous sections, gp4 has a C-terminal tail analogous to the tail of gp2.5. The chimeric versions of gp4 and gp2.5, in which the C-terminal tails are swapped, bind to gp5/trx like their respective wild-type counterparts. Both chimeric proteins also support phage growth [120,152]. The overlapping interactions of gp2.5 with gp5/trx and gp4 modulate synthesis of the primer [154,176], and are important for coordination of leading- and lagging-strand synthesis [177].

6. LEADING-STRAND SYNTHESIS Efficient leading-strand synthesis requires numerous interactions between gp5/trx, gp4, and DNA. The gp4–gp5/trx complex mediates the simultaneous unwinding of duplex DNA and DNA synthesis at the rate of 114 bp/s with a processivity greater than 17 kb [68,69].

6.1 Gp4–gp5/trx Interactions in Leading-Strand Synthesis The interaction between a basic patch on gp5/trx and the C-terminal tail of gp4 is essential for the initiation of leading-strand synthesis. This solvent exposed patch consisting of four basic residues is located on the surface of

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Fig. 10 Interactions of gp5/trx and gp4 during synthesis of the leading-strand. (A) The gp4 assembles as a hexamer on the lagging-strand DNA. Gp5/trx is recruited to the replication fork by an initial interaction with the acidic C-terminal tail of gp4 through its basic loading patch (Fbp). (B) Gp5/trx can also initially bind to the C-terminal tail of gp4 via two basic loops located within the TBD (TBDbp). (C) The interaction of the loading patch of gp5 with the C-terminal tail of helicase facilitates the loading of gp5/trx to the primer-template and the interaction is maintained during leading-strand synthesis. Gp5/trx and gp4 also interact through the helicase domain of gp4 and a part of the surface of the polymerase domain of gp5/trx. (D) The C-terminal tails of the gp4 hexamer not only bind any dissociating gp5/trx but also accommodate additional gp5/trx through interactions with the exchange patch of gp5. Any of these gp5/trx can be reloaded onto the primer-template without involvement of the loading patch.

gp5 (Fig. 10) facing the duplex region of the DNA adjacent to the template strand as it exits the active site of gp5 [153]. Substitution of any of the basic residues from the patch with acidic residues eliminates the ability of altered gp5/trx to interact with gp4 to initiate strand-displacement synthesis [153]. Protein–protein binding studies show that the altered polymerase is defective in binding to gp4, and that the acidic C-terminal tail of gp4 is involved in this interaction. Similarly to gp2.5, the acidic tail of gp4 contains a phenylalanine residue at its C-terminus. The replacement of the terminal phenylalanine with a nonpolar residue abolishes the binding between gp4 and gp5/trx [120]. The results suggest that the interaction between the C-terminal tail of the helicase and the basic patch of DNA polymerase is important for loading of gp5/trx onto the replication fork to initiate

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strand-displacement synthesis. Once gp5/trx and gp4 have initiated leadingstrand synthesis this basic patch is no longer necessary for processive leadingstrand synthesis [153]. Most studies on leading-strand synthesis use a preformed replication fork with a 50 -ssDNA tail onto which gp4 can load. Gp5/trx and gp4 are unable to initiate synthesis at a nick in duplex DNA [175]. Gp2.5 enables gp5/trx to catalyze limited strand-displacement synthesis at a nick, creating a 50 -ssDNA tail to which gp4 binds. Gp5/trx and gp4 then mediate leading-strand DNA synthesis. Gp2.5 lacking the C-terminal phenylalanine does not support the loading of gp5/trx, thus implying that a gp5/trx–gp2.5 interaction is necessary in addition to binding of gp2.5 to DNA [175]. Suppressor mutations in gp5 that allow T7 mutants expressing gp2.5 lacking a C-terminal phenylalanine to grow also allow this altered gp2.5 to support initiation of DNA synthesis at a nick. Gp5/trx has a processivity of 0.8 kb on ssDNA, but a processivity higher than 17 kb during leading-strand synthesis when coupled to gp4 [67,69]. A deletion of the C-terminal tail of gp4 reduces the processivity to 5 kb [69,75]. A small-angle X-ray scattering (SAXS) model of the leading-strand complex reveals that the stable interaction of gp4 and gp5/trx that is essential for the high processivity of 5 kb observed during DNA synthesis in the absence of gp5/trx exchange is mediated through the helicase and the palm–exonuclease of gp5/trx [69]. This interaction can be observed by surface plasmon resonance (SPR), when gp5/trx is bound to a primer-template in the presence of the dideoxynucleoside 50 -triphosphate corresponding to the next incoming dNTP [69,93]. The complex mediating leading-strand synthesis is extremely stable with a half-life of more than 10 min [93]. The binding between gp4 and ssDNA is also stable with a dissociation rate of 0.002/s [178]. However, the complex of gp5/trx, in the absence of gp4, readily dissociates from a primer-template with a dissociation constant of 0.2/s [179]. The ZBD and RPD are not directly involved in binding to gp5/trx as indicated by nearly identical Rmax measured using SPR for wild-type gp4, and the truncation constructs lacking ZBD or the entire primase domain [69]. The primase domain of gp4 is involved in formation of the leading-strand complex by stabilizing the helicase. Removal of the ZBD from gp4 has a minimal influence on leading-strand synthesis as measured in the single-molecule assay. Removal of both the ZBD and the RPD results in a significant reduction of leading-strand synthesis indicative of the major role of the RPD in stabilization of the helicase, as discussed earlier in the

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section concerned with an oligomerization of gp4. The relative orientation of gp5/trx and gp4 in the SAXS model of the leading-strand complex allows for binding of the C-terminal tail of the helicase to a basic patch on gp5/trx. Thus we propose that this electrostatic interaction is maintained during leading-strand synthesis. Removal of the C-terminal tail decreases the binding of gp4 to gp5/trx in the leading-strand complex by 30% as evidenced by SPR measurements [69], abolishes initiation of strand-displacement synthesis, and reduces the rate of ongoing synthesis by 50% [153]. In the model, a primer-template bound to gp5/trx is located such that the extension of the 50 -end of the template would position it in the close proximity to the subunit interface β-hairpin consisting of phenylalanine 523 of gp4. Phenylalanine 523 interacts directly with the extruded DNA leading-strand during the process of DNA unwinding [138]. Gp5/trx–gp4 interaction in the leadingstrand complex helps to stabilize partially unwound dsDNA at the junction between ssDNA and dsDNA at the replication fork, thus helping to separate duplex DNA. Additionally, gp5/trx binding to the helicase can change the equilibrium between the forward and the backward Brownian sliding of gp4 along ssDNA [102] consistent with an active model for DNA unwinding described in a previous section. Gp5/trx not only destabilizes the duplex to facilitate forward Brownian push of the helicase but it also prevents backward slipping of the helicase by synthesizing the complementary strand [84,85]. Gp4 variants defective in ssDNA binding as well as unwinding can mediate efficient strand-displacement synthesis in the presence of gp5/trx, thus confirming the above hypothesis [180].

6.2 Exchange of DNA Polymerases Replisomes of E. coli, and T4 and T7 bacteriophages remain highly processive even when challenged by dilution [181–184]. At the same time, however, the T4 and T7 polymerases exchange rapidly when challenged with excess polymerase in solution [181,182]. In the phage T7 system, this exchange requires the presence of gp4, suggesting that the interactions between gp4 and gp5/trx mediate the process. This ability of DNA polymerase in solution to exchange with the replicating DNA polymerase is medicated through the interaction between the C-terminal tail of gp4 and the TBD of gp5. As described earlier, when trx binds to the TBD two basic loops are formed in the TBD. It is to these two basic loops that gp2.5 and gp4 bind via their acidic C-terminal tails. These basic loops allow for the exchange of polymerases [71,75].

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Abolishment of the charges in the loops or removal of the C-terminal tail of gp4 reduces the processivity of the replisome from 17 kb to approximately 5 kb as measured by single-molecule techniques [69,71]. Interestingly, in the absence of trx, the T7 replisome can mediate leading-strand synthesis, but at a rate and processivity 4.5 times lower, ie, 26 bp/s and 3.9 kb, respectively [74]. Thus, apart from functioning as a processivity factor, trx also plays an important role in the process of polymerase exchange. In addition, one lysine in trx located in close proximity to one of the two basic loops also contributes to the interaction [185]. The exchange of DNA polymerases during leading-strand synthesis has been visualized directly by correlating the arrival of fluorescently labeled polymerases at the replication fork with changes in DNA synthesis [71]. Performing these experiments at the level of individual replisomes allows for quantification of the kinetics of the polymerase exchange reaction. In the experiments, a mixture of an altered unlabeled polymerase and a fluorescently labeled wild-type gp5 was used. The altered polymerase described in the section concerned with discrimination of gp5/trx against ddNTPs contains a single amino-acid substitution in the active site, namely Tyr 526 is replaced with Phe. This mutation increases the ability of the altered enzyme to discriminate against ddNTPs by several 1000-fold in comparison to wild-type gp5. Interestingly, the replacement of Tyr at position 526 with phenylalanine in the nucleotide-binding pocket slows the average rate of DNA synthesis 2.5-fold (122  5 to 48  2 bp/s), [71]. This rate change can be readily detected at the single-molecule level. Strikingly, with DNA replication reactions containing a 1∶1 mixture of both polymerases, abrupt changes in the rate of synthesis carried out by individual replisomes can be observed. These changes correlate with the appearance and disappearance of the fluorescently labeled wild-type gp5/trx at the replication fork. For instance, the appearance of the wild-type polymerase is followed by an increase in the rate of synthesis. The exchange process is not immediate with a delay between the time when a free polymerase from solution associates with gp4 at the replisome. On average this delay is approximately 1 min. The observed delay between polymerase association and exchange has important implications for how the replisome might regulate access of various proteins at the replication fork. For instance, translesion DNA polymerases suggest that the replisome is highly dynamic, with components exchanging and entire replisomes collapsing and reassembling [186]. Studies involving the E. coli [183,184] and bacteriophage T4 [181] replisomes have revealed that polymerase exchange is mediated through interactions between the polymerase and the sliding clamp.

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In summary, we envision the following scenario for leading-strand synthesis and exchange of polymerases at a T7 replication fork. Gp5/trx joins the replication complex by binding to the C-terminal acidic tail of gp4 via its basic loading patch located at the front side of gp5 facing its direction of movement along the ssDNA template (Fig. 10A). Alternatively, gp5/trx can bind to the C-terminal tail through a basic exchange patch located in the TBD of gp5 (Fig. 10B). Subsequently, gp5/trx has to switch its binding to the loading patch in order to initiate DNA synthesis [153] (Fig. 1C). Elimination of the basic charges located within the exchange patch does not interrupt strand-displacement synthesis [75]. Once leading-strand synthesis is initiated a highly stable complex of gp5/trx and gp4 is formed, increasing the processivity from 0.8 kb to approximately 5 kb per binding event. Interactions of the C-terminal tail of gp4 with the exchange patch of gp5/trx further increase the processivity of the replisome from 5 to >17 kb by capturing gp5/trx that transiently dissociate from DNA [71,75] (Fig. 10D). On the lagging-stand DNA polymerase repeatedly dissociates from DNA and recycles to a new primer upon completing of each Okazaki fragment. Single-molecule analysis has shown that lagging-strand polymerases are also exchanged at the replication fork [72]. Albeit this exchange process is more frequent than on the leading-strand. The lagging-strand gp5/trx remains associated with the replisome for the synthesis of one or two Okazaki fragments.

7. LAGGING-STRAND SYNTHESIS The lagging-strand DNA polymerase requires an oligoribonucleotide, synthesized by DNA primase, to initiate the synthesis of an Okazaki fragment. In the replication system of bacteriophage T7, gp4 provides DNA primase and DNA helicase activity. The primase activity resides within the amino-terminal half of gp4 [104]. The primase domain contains the N-terminal ZBD connected to the RPD by a flexible linker (Fig. 6A). The current model for lagging-strand synthesis involves several events [187]. Both the ZBD and RPD are involved in synthesis of the primer. First, the trinucleotide sequence, 50 -GTC-30 in the lagging-strand is recognized by the primase, an event for which there is little information [176,188]. The cytosine in the template is “cryptic” in that it is essential for recognition but is not copied into the primer. Both the ZBD and the RPD appear to play a role in recognition of the cryptic cytosine [188]. At this trinucleotide recognition site, T7 primase catalyzes the synthesis of the diribonucleotide

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50 -pppAC-30 and further extends it to the functional tetraribonucleotides, 50 -pppACCC-30 , 50 -pppACCA-30 , and 50 -pppACAC-30 provided cognate sequences are available in the template. The interaction of the ZBD with the RPD can occur in cis or trans within the functional gp4 hexamer (Fig. 11) [189]. In the trans mode the ZBD on one subunit of the hexamer contacts the RPD on an adjacent subunit to catalyze the synthesis of the oligonucleotide. Results obtained by modifying the length of the linker connecting the RPD and the ZBD suggest a model in which the primase catalyzes the synthesis of pppAC in a cis mode and the extension occurs in a trans mode [190]. A similar swapping arrangement, in which the ZBD of one molecule packs against the adjacent RPD from another molecule was observed in structures of bacteriophage T7 primase [64] and DnaG from Aquifex aeolicus and E. coli [191]. The trans mode has been postulated to explain the halting of helicase movement and hence leading-strand replication, the phenomenon observed during synthesis of primers using single-molecule methods [67]. This pause allows for the relatively slow process of primer synthesis and, the subsequent events that lead to the initiation of a new Okazaki fragment [67]. Without such a pause leading-strand synthesis would outpace lagging-strand

Fig. 11 Primer synthesis by gp4. The two distinct modes of interaction between the zinc-binding domain (ZBD) and RNA polymerase domain (RPD) are involved in primer synthesis at the primase recognition sequence by the primase domain of gp4. A primer is synthesized at the 50 -GTC-30 basic recognition sequence through interactions between the ZBD and RPD within the same subunit (cis mode). Alternatively, the primers can be synthetized in a trans mode, in which the ZBD of one subunit contacts the RPD of an adjacent subunit within the ring of gp4 primase–helicase. The cis configuration is likely adopted during synthesis of diribonucleotides. The diribonucleotides are then extended to tetraribonucleotides in the trans configuration of the RPD and the ZBD.

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synthesis. Finally, the tetraribonucleotide is transferred to T7 DNA polymerase where it serves as a primer to initiate the synthesis of Okazaki fragments [176,188]. A protein–nucleic acid complex resembling a conformation of the replisome adopted during extension of an RNA primer to an Okazaki fragment by the lagging-strand DNA polymerase was reconstituted in vitro [188]. Formation of this complex is dependent on the presence of a six nucleotide RNA/DNA primer [176]. The complex is formed only when the 30 -end of the primer contains the chain-terminating ddC and the next incoming nucleoside 50 -triphosphate is present. The presence of ddC at the 30 -terminus of the oligonucleotide prevents gp5/trx from extension of the primer and consequently dissociation from the priming complex. The incoming nucleotide base with the next nucleotide in the template and locks gp5/trx on DNA since the 30 -ddC on the primer has no 30 -hydroxyl group to carry out a nucleophilic attack on the α-phosphate of the dNTP. The RPD is crucial in stabilization of the complex, as confirmed by fluorescence anisotropy measurements with an altered gp4 lacking the primase. The presence of the ZBD is not essential for complex formation, although the complex is less stable. The C-terminal tail of gp4 does not play a role in stabilization of the priming complex.

7.1 RNA Primer Handoff Once the tetranucleotide is synthesized it must be transferred into the DNAbinding crevice of gp5 such that the 30 -terminus is situated in the active site. Precisely how the tetranucleotide is transferred to the DNA-binding crevice is not known. Earlier studies suggested that the ZBD plays a major role in primer delivery [64,176]. However, gp4 lacking the ZBD is also capable of primer delivery, necessitating the involvement of other elements [192]. Once the primer has been delivered to the active site of polymerase, it is stabilized by interactions between gp5/trx and the ZBD and RPD of gp4 [177,188,193]. Oligonucleotides less than 21-nt in length are ineffective primers for gp5/trx clearly indicative of an interaction of gp4 with gp5/ trx to secure the primer in the DNA-binding cleft of gp5 [60]. Taking into consideration the biochemical data, we propose the following model (Fig. 12) for a conformation of the replisome adopted during extension of lagging-strand primers. In this model, the ZBD and RPD interact with each other to form a positively charged interface stabilizing a primer-template in the DNA-binding crevice of gp5/trx.

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Fig. 12 A model for the RNA primer handoff from gp4 to gp5/trx. Once synthetized, the RNA primer is transferred from gp4 to gp5/trx for an extension to an Okazaki fragment. Binding between the ZBD from one subunit and the RPD of the adjacent subunit leads to formation of a positively charged interface that stabilizes a primer-template (shown as a black ribbon) in the DNA-binding crevice of DNA polymerase. Amino acids important in primer synthesis and primer delivery from gp4 to gp5/trx are displayed as black sticks in the model. See text for detailed description.

This protein–protein interface encircles the DNA, preventing dissociation of the primer from the template. NMR chemical shift mapping and alanine scanning mutagenesis implicated several amino acids from the ZBD and RPD in primer delivery [176]. Mapping of these amino acids into the structural model reveals that they are clustered in the interface formed by the ZBD and RPD. For instance, residues: F35, Y37, E40, and W42 form a part of the interface that is located in close proximity to the primer strand, whereas residues S30 and V38 are positioned close to the template strand. The ZBD amino acid: D31 is involved in the recognition of the “cryptic” cytosine [194]. In the model, D31 is located in proximity to the 30 -terminus of the template strand, the position where the cryptic cytosine is expected to reside after primer delivery to gp5/trx. Three other amino acids, K122, K128, and K131 are also involved in DNA binding [195]. These residues form a shallow groove between the TOPRIM and the N-terminal subdomain of RPD. Tryptophan 69 located in the RPD is crucial in primer delivery [192]. In the model, W69 is located in the protein–protein interface between the RPD and gp5/trx, supporting an important role of this residue in formation of the lagging-strand complex. The four amino acids loop (residues: E401–K404) in the DNA-binding

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crevice of gp5/trx is also implicated in the process of primer delivery. Gp5/ trx lacking this loop is defective in the use of tetraribonucleotides as primers to initiate DNA synthesis [196]. In the structural model, the loop forms a part of the DNA-binding interface located directly across the ZBD close to the template strand (Fig. 12). The primase active site contains five invariant residues: E157, D161, D207, D209, and D237 coordinating two magnesium ions. These acidic residues are located in a cleft between the TOPRIM and the N-terminal subdomain of the RPD. A mechanism has been suggested in which the linker region between the ZPD and the RPD acts as a hinge to bring the two domains together for primer synthesis [64,176]. In our model, the RPD ˚ from the ZBD active site from one subunit is located approximately 20 A of the neighboring subunit (Fig. 12). We propose that prior to adopting this conformation, both the ZBD and RPD from the same primase domain come together in a cis conformation to facilitate synthesis of pppAC, consistent with biochemical data. A subsequent conformational change in the linker region allows for the delivery of pppAC to the RPD from the neighboring subunit, and the extension of pppAC to a functional tetraribonucleotide in trans. In this new conformation gp5, the ZBD and RPD binding in trans form a tunnel-like structure encircling a primertemplate. Since the catalytic sites of polymerase and primase are located in close proximity across this tunnel, the tetraribonucleotide can be extended without any additional conformational changes.

7.2 Formation and Release of the Lagging-Strand Replication Loop The reverse polarity and antiparallel nature of the two strands of duplex DNA pose a topological problem for their simultaneous synthesis. The “trombone” model of DNA replication postulates that the lagging-strand forms a loop such that the leading and lagging-strand replication proteins contact one another [197]. This replication loop contains a nascent Okazaki fragment and allows for coordination of leading and lagging-strand synthesis; the replisome now can move in one direction along the DNA while synthesizing both strands (Fig. 1). Replication loops were visualized for the first time by negative-stain electron microscopy (EM) [198] using bacteriophage T7 replication proteins assembled on a preformed M13 replication fork [66]. Subsequently, these loops were characterized using a synthetic minicircular DNA (minicircle) molecule [156,199] and confirmed by single-molecule techniques [67,76].

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The minicircle consists of a 70 bp circular dsDNA with a replication fork bearing a 50 -ssDNA tail of 40 nt [156]. The 30 -hydroxyl terminus on the duplex provides the site for assembly of gp5/trx, and the single-stranded tail provides a loading site for gp4 so that it can translocate along ssDNA in a 50 to 30 direction to assist DNA polymerase in synthesis of the leading-strand. A sequence on the lagging-strand contains a defined number of recognition sites (50 -TGGTC-30 ) at which the primase can catalyze the synthesis of tetraribonucleotide primers when provided with ATP and CTP. The minicircle offers several advantages for studies of DNA replication. Firstly, it provides a desirable concentration of replication forks necessary for proper stoichiometry between DNA and the replisomes. Such stoichiometry is difficult to achieve using larger naturally occurring DNA molecules. In addition, large DNA molecules may harbor nicks and single-stranded regions to which the replication proteins would bind and catalyze nonspecific reactions. Second, the minicircle permits for selective base composition. If one strand, eg, the leading-strand is mainly composed of cytosine residues, the complementary lagging-strand consequently lacks cytosine residues. Thus, it is possible to measure quantitatively the amount of leading- and lagging-strand synthesis by monitoring the incorporation of dGMP and dCMP, respectively. In the reaction consisting of a minicircle, gp5/trx, gp4, and gp2.5, leading- and lagging-strand synthesis proceeds at an equal rates [156,199]. Coordinated DNA synthesis yields a continuous leading-strand of >10 kb, and lagging-strand containing multiple Okazaki fragments with an average length of 1 kb. The size of Okazaki fragments is constant regardless of the number of primase recognition sites on the minicircular template. In the absence of a primase recognition site, no lagging-strand synthesis occurs. A distinguishing feature of the coordinated DNA synthesis is the presence of a lagging-strand replication loop containing ssDNA and dsDNA segments [199]. The double-stranded portion of the replication loop represents the nascent Okazaki fragment. The loop on the lagging-strand implies changes in the orientation of gp5/trx bringing it into the replisome. In this new orientation, both leadingand lagging-strand synthesis occur at a single point and proceed in the same direction. Precisely how the replication loop is formed and released is unknown. As gp4 unwinds dsDNA the resultant ssDNA passes through the central channel in the hexameric ring of DNA helicase [62,69,134]. The crystal structure of the hexameric gp4 reveals the presence of three loops protruding into the central cavity [62]. These loops contain a number of

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basic residues involved in DNA binding [140]. Hydrolysis of dTTP by gp4 is coupled with a sequential transfer of ssDNA by the DNA-binding loops from one subunit to the adjacent subunit within the central channel [62,140]. ssDNA exiting from the channel has the correct polarity for binding by the primase and the DNA-binding cleft of gp5/trx. Most likely the polymerization activity of gp5/trx combined with the movement of ssDNA through the central cavity of the helicase account for formation of the replication loop on the lagging-strand. Two models have been proposed for the release of the replication loop: (i) the collision model and (ii) the signaling model (Fig. 13). In the collision model, the encounter of gp5/trx with the 50 -terminus of previously synthesized Okazaki fragment triggers dissociation of gp5/trx and loop release [76,200–202]. In the E. coli and phage T4 replication systems, after the collision the lagging-strand polymerase disengages from the clamp protein leaving the clamp bound to the lagging-strand [168,203,204]. The observation that the length of the Okazaki fragment is related to primer synthesis led to an alternative-signaling model, in which RNA polymerization activity of the primase is the signal triggering release of the replication loop [76,202,205]. Several reports suggest that the signaling mechanism functions in the E. coli, phage T4 and T7 systems [205]. One of the consequences of the signaling model is the fact that a fraction of replication loops is released before completion of the Okazaki fragment. EM analysis and gel

Fig. 13 Models for release of the lagging-strand replication loop. Two models have been proposed for release of the replication loop on lagging-strand. In the signaling mechanism, the lagging-strand polymerase releases the nascent Okazaki fragment as a result of signaling by the primase upon synthesizing a dinucleotide at a primase recognition sequence (highlighted in a dotted circle). In the collision mechanism, an encounter between gp5/trx completing the synthesis of the Okazaki fragment with the 50 -end of the previously synthesized Okazaki fragment (highlighted by a dotted circle) triggers release of the replication loop.

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electrophoresis reveal the presence of gaps of ssDNA between Okazaki fragments synthesized by the T4 replisome [205,206]. In the phage T7 system, loop release was correlated with the first step of primer synthesis, the condensation of ATP and CTP to pppAC by the primase active site [76]. In addition, varying the concentration of ATP and CTP results in changes in the size of Okazaki fragments in E. coli, phage T4 and T7 systems [199,205,207]. The size of Okazaki fragments is dependent on the concentration of the clamp and the lamp loader, the two components of the lagging-strand complex in T4 replisome [205]. In bacteria, association of DnaG primase with DnaB helicase is thought to function as a primary regulator of Okazaki fragment length [208]. Despite these observations, the detailed mechanism of how primase activity leads to the release of the replication loop remains unclear. We envision a switch mechanism, in which primase dissociates from gp5/trx and binds to the primase recognition sequence that exits from the central cavity of the helicase. This simple switch mechanism is driven by relative differences in binding affinities of primase to gp5/trx and to the primase recognition sequence. Dissociation of the primase destabilizes gp5/trx on the replisome, subsequently leading to detachment of gp5/trx from DNA and release of the replication loop. An analogous mechanism can function in bacteria. One possibility is that DnaG binding to DnaB destabilizes the interaction between the accessory subunit tau (τ) of DNA polymerase III with DnaB. The average length of E. coli Okazaki fragments decreases from 1–1.5 to 0.5 kb in the absence of τ [209]. Single-molecule analysis of the sizes and lag times between release and formation of subsequent replication loops reveals that both collision and signaling mechanisms function in the T7 system [76]. The signaling mechanism releases the replication loop if the primase locates the primer recognition site before synthesis of the Okazaki fragment is completed. Alternatively, if synthesis of the Okazaki fragment is finished, but the primase did not engage a new recognition sequence, the collision mechanism triggers loop release. The presence of two triggers may represent a fail-safe mechanism ensuring the timely reset of the replisome after the synthesis of each Okazaki fragment (Fig. 13). Following loop release, gp5/trx remains bound to the replisome through the C-terminal tail of gp4. In this way, the same molecule of gp5/trx can be recycled for synthesis of subsequent Okazaki fragments [72]. It has been also postulated that the binding of gp2.5 to the replication results in condensation of the DNA with the increasing mass leading to resolution of the replication loop [156].

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8. COORDINATION OF LEADING- AND LAGGINGSTRAND SYNTHESIS Coordination of leading- and lagging-strand synthesis is critical for rapid and accurate replication of a chromosome by the replisome. However, the enzymatic events on the lagging- and leading-strand are not equivalent. The lagging-strand DNA polymerase repeatedly dissociates from DNA and recycles to a new primer upon completing of each Okazaki fragment. Furthermore, one of the slowest events on the lagging-strand is recognition of primase recognition sites and synthesis of oligoribonucleotides by the primase. In the 6–12 s required for these steps [67,76,210,211], approximately a thousand nucleotides of leading-strand DNA would accumulate [59,67,69]. Why does the synthesis of leading-strand not outpace the synthesis of the lagging-strand? Two models have been proposed to achieve coordination of DNA replication. In the first model, leading-strand synthesis transiently stops allowing the completion of the slow enzymatic events on the lagging-strand. Single-molecule studies with T7 replication complexes undergoing coordinated leading- and lagging-strand synthesis reveal the presence of the 12 s lag between the release of one replication loop and formation of the next loop [76]. Reducing the concentration of ATP and CTP required for primer synthesis by the primase increase the lag time to approximately 28 s. Simultaneously, the average length of synthesized Okazaki fragments increase from 1.4 to 3.6 kb. These observations suggest the role of the signaling mechanism in release of replication loops in the T7 system. Interestingly, addition of a preformed pppAC restores the average lag time and loop length to these observed with optimal concentrations of ribonucleotides [76]. This result demonstrates that condensation of ATP and CTP to pppAC by the ZBD and RPD triggers loop release, the event that likely takes place in the cis configuration (Fig. 11). A subsequent transfer of pppAC to the RPD in the adjacent subunit has been postulated to explain the halting of helicase movement observed during primer synthesis [76]. It the model, the trans interaction (Fig. 11) triggers a conformational change that halts the sequential hydrolysis of dTTP by gp4, and consequently temporarily stops the forward movement of the replisome [190]. In the second model, coordination is achieved by different synthesis rates of the leading- and lagging-strand. Studies of E. coli, phage T4, and T7 replication systems demonstrated that the lagging-strand is synthetized faster than the leading-strand [212,213]. Contrary to the results described by Lee et al. [67], single-molecule studies of the T7 replisome by Pandey

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et al. [212] indicate that lagging-strand gp5/trx synthesizes DNA approximately 38% faster than does the leading-strand gp5/trx. In addition, no pausing of the replisome movement was detected during primer synthesis [212]. Structural and single-molecule studies aiming to discriminate between the two models are currently underway. Gp2.5 stimulates activities of gp4 and gp5 during coordinated DNA synthesis [156]. Gp2.5 increases the frequency of initiation of laggingstrand synthesis >10-fold [59]. The acidic 26-residue C-terminal tail of gp2.5 interacts with the basic exchange patch of gp5/trx [75], the region where also the acidic C-terminal tail of DNA helicase binds (Fig. 10). A competition between these two overlapping interactions during coordinated synthesis can provide a switching mechanism to release gp5/trx from the lagging-strand and allow for new primer utilization. In an analogous manner, gp2.5 can control polymerase exchange and leading-strand synthesis by the interaction of its C-terminal with a basic loading patch on the surface of gp5. The overlapping interactions between gp4, gp5, and gp2.5 provide a structural framework for coordination of timing and kinetics in leading- and lagging-strand synthesis. Primase recognition sites are abundant in the phage T7 genome. Despite the fact that the majority of these sites are used for tetraribonucleotide synthesis, not all tetraribonucleotides are used as primers by lagging-strand gp5/ trx [210,212]. A similar phenomenon has been observed in replication systems from other organisms [205]. Why are not all the tetraribonucleotides extended? Our model based on structural and biochemical data shows that RNA and DNA polymerization activities at the replication fork are spatially decoupled with the exception of events that occur during primer transfer [177]. Consequently, multiple tetraribonucleotides can be synthesized during the time necessary for completion of an Okazaki fragment. Previous single-molecule studies indicated that tetraribonucleotides are synthetized concomitantly during ongoing DNA synthesis [212].

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[192] B. Zhu, S.J. Lee, C.C. Richardson, Direct role for the RNA polymerase domain of T7 primase in primer delivery, Proc. Natl. Acad. Sci. U.S.A. 107 (2010) 9099–9104. [193] J.R. Wallen, J. Majka, T. Ellenberger, Discrete interactions between bacteriophage T7 primase-helicase and DNA polymerase drive the formation of a priming complex containing two copies of DNA polymerase, Biochemistry 52 (2013) 4026–4036. [194] T. Kusakabe, A.V. Hine, S.G. Hyberts, C.C. Richardson, The Cys4 zinc finger of bacteriophage T7 primase in sequence-specific single-stranded DNA recognition, Proc. Natl. Acad. Sci. U.S.A. 96 (1999) 4295–4300. [195] S.J. Lee, C.C. Richardson, Essential lysine residues in the RNA polymerase domain of the gene 4 primase-helicase of bacteriophage T7, J. Biol. Chem. 276 (2001) 49419–49426. [196] K. Chowdhury, S. Tabor, C.C. Richardson, A unique loop in the DNA-binding crevice of bacteriophage T7 DNA polymerase influences primer utilization, Proc. Natl. Acad. Sci. U.S.A. 97 (2000) 12469–12474. [197] N.K. Sinha, C.F. Morris, B.M. Alberts, Efficient in vitro replication of doublestranded DNA templates by a purified T4 bacteriophage replication system, J. Biol. Chem. 255 (1980) 4290–4293. [198] K. Park, Z. Debyser, S. Tabor, C.C. Richardson, J.D. Griffith, Formation of a DNA loop at the replication fork generated by bacteriophage T7 replication proteins, J. Biol. Chem. 273 (1998) 5260–5270. [199] J. Lee, P.D. Chastain 2nd., J.D. Griffith, C.C. Richardson, Lagging strand synthesis in coordinated DNA synthesis by bacteriophage T7 replication proteins, J. Mol. Biol. 316 (2002) 19–34. [200] T.E. Carver Jr., D.J. Sexton, S.J. Benkovic, Dissociation of bacteriophage T4 DNA polymerase and its processivity clamp after completion of Okazaki fragment synthesis, Biochemistry 36 (1997) 14409–14417. [201] F.J. Lopez de Saro, R.E. Georgescu, M. O’Donnell, A peptide switch regulates DNA polymerase processivity, Proc. Natl. Acad. Sci. U.S.A. 100 (2003) 14689–14694. [202] X. Li, K.J. Marians, Two distinct triggers for cycling of the lagging strand polymerase at the replication fork, J. Biol. Chem. 275 (2000) 34757–34765. [203] P.T. Stukenberg, J. Turner, M. O’Donnell, An explanation for lagging strand replication: polymerase hopping among DNA sliding clamps, Cell 78 (1994) 877–887. [204] R.E. Georgescu, I. Kurth, N.Y. Yao, J. Stewart, O. Yurieva, M. O’Donnell, Mechanism of polymerase collision release from sliding clamps on the lagging strand, EMBO J. 28 (2009) 2981–2991. [205] J. Yang, S.W. Nelson, S.J. Benkovic, The control mechanism for lagging strand polymerase recycling during bacteriophage T4 DNA replication, Mol. Cell 21 (2006) 153–164. [206] N.G. Nossal, A.M. Makhov, P.D. Chastain 2nd., C.E. Jones, J.D. Griffith, Architecture of the bacteriophage T4 replication complex revealed with nanoscale biopointers, J. Biol. Chem. 282 (2007) 1098–1108. [207] C.A. Wu, E.L. Zechner, J.A. Reems, C.S. McHenry, K.J. Marians, Coordinated leading- and lagging-strand synthesis at the Escherichia coli DNA replication fork. V. Primase action regulates the cycle of Okazaki fragment synthesis, J. Biol. Chem. 267 (1992) 4074–4083. [208] K. Tougu, K.J. Marians, The interaction between helicase and primase sets the replication fork clock, J. Biol. Chem. 271 (1996) 21398–21405. [209] C.A. Wu, E.L. Zechner, A.J. Hughes Jr., M.A. Franden, C.S. McHenry, K.J. Marians, Coordinated leading- and lagging-strand synthesis at the Escherichia coli DNA replication fork. IV. Reconstitution of an asymmetric, dimeric DNA polymerase III holoenzyme, J. Biol. Chem. 267 (1992) 4064–4073.

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CHAPTER FOUR

Protein-Primed Replication of Bacteriophage Φ29 DNA M. Salas1, M. de Vega1 Instituto de Biologı´a Molecular “Eladio Vin˜uela” (CSIC), Centro de Biologı´a Molecular “Severo Ochoa” (CSIC-UAM), Universidad Auto´noma de Madrid, Cantoblanco, Madrid, Spain 1 Corresponding authors: e-mail address: [email protected]; [email protected]

Contents 1. Introduction 2. The Protein-Primed Replication Mechanism 2.1 Initiation of ϕ29 TP–DNA Replication 2.2 A Sliding-Back Mechanism for Protein-Primed DNA Replication 2.3 Transition from Protein-Primed to DNA-Primed Replication 2.4 DNA-Primed Elongation 2.5 TP-Primed DNA Amplification 3. In Vivo Compartmentalization of ϕ29 DNA Replication Acknowledgments References

138 138 140 153 155 156 158 158 161 161

Abstract The requirement of DNA polymerases for a 30 -hydroxyl (30 -OH) group to prime DNA synthesis raised the question about how the ends of linear chromosomes could be replicated. Among the strategies that have evolved to handle the end replication problem, a group of linear phages and eukaryotic and archaeal viruses, among others, make use of a protein (terminal protein, TP) that primes DNA synthesis from the end of their genomes. The replicative DNA polymerase recognizes the OH group of a specific residue in the TP to initiate replication that is guided by an internal 30 nucleotide of the template strand. By a sliding-back mechanism or variants of it the terminal nucleotide(s) is(are) recovered and the TP becomes covalently attached to the genome ends. Bacillus subtilis phage ϕ29 is the organism in which such a mechanism has been studied more extensively, having allowed to lay the foundations of the so-called protein-primed replication mechanism. Here we focus on the main biochemical and structural features of the two main proteins responsible for the protein-primed initiation step: the DNA polymerase and the TP. Thus, we will discuss the structural determinants of the DNA polymerase responsible for its ability to use sequentially a TP and a DNA as primers, as well as for its inherent capacity to couple high processive synthesis to strand displacement. On the other hand, we will review how TP primes initiation followed by a transition step for further DNA-primed replication by the same polymerase molecule. Finally, we will review how replication is compartmentalized in vivo. The Enzymes, Volume 39 ISSN 1874-6047 http://dx.doi.org/10.1016/bs.enz.2016.03.005

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1. INTRODUCTION DNA polymerases are unable to start de novo DNA synthesis. Their ability to insert nucleotides relies on the presence of a 30 OH group, usually provided by the 30 end of a DNA or an RNA molecule, that primes DNA synthesis. This fact implies a problem during the replication of the termini of linear chromosomes as once the most terminal primer is removed, the remaining short region of unreplicated single-stranded DNA (ssDNA) would eventually lead to a continuous shortening of the daughter DNA molecule after successive rounds of DNA replication that would cause cell death. To guarantee replication of the chromosome ends, organisms containing linear genomes have developed diverse replication strategies. Thus, bacteriophages T4, T7, and SPP1 form head–tail concatemers because of terminal redundancies. Others, as phage λ, contain terminal complementary sequences that allow them to circularize early after infection to be replicated by rolling circle (reviewed in Ref. [1]). In higher eukaryotes telomerase extends directly the 30 end, producing an overhanged ssDNA end [2] that finally can invade homologous double-stranded (ds) telomeric tracts, enlarging and protecting chromosome ends [3]. Other organisms, including bacteriophages, animal viruses like adenovirus and human hepatitis B virus, mitochondrial and cytoplasmic plasmids of yeast and filamentous fungi, linear chromosomes and plasmids of Streptomyces [4], several virus infecting Archaea [5–7], and eukaryotic transposable elements and virophages [8,9], possess replication origins at both ends of their linear chromosomes formed by inverted terminal repetitions (ITR) together with a terminal protein (TP) [10]. Therefore, this unusual location of the two replication origins allows simultaneous and continuous duplication of both DNA strands without requiring asymmetric complexes of DNA polymerase with accessory proteins to control the different mechanics of continuous and discontinuous synthesis [11]. In these cases, the OH group of a specific serine, threonine, or tyrosine of the TP primes initiation of DNA replication from the ends of the linear chromosome, circumventing the “end replication problem,” the TP becoming covalently linked to the 50 -termini of the genome (TP–DNA) [4,10,12].

2. THE PROTEIN-PRIMED REPLICATION MECHANISM ϕ29 is a lytic phage that infects the Gram-positive bacterium Bacillus subtilis [13]. It has a linear dsDNA genome 19.3-kbp long, with a 6 bp ITR

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(30 -TTTCAT) [14,15] and a 31 kDa TP covalently linked to each 50 end by a phosphoester bond between the OH group of Ser232 and 50 -dAMP (the initial nucleotide) [16]. The development of a soluble in vitro replication system with highly purified proteins and TP–DNA from bacteriophage ϕ29 has allowed to lay the foundations of the so-called protein-primed mechanism of DNA replication [12,17,18]. Fig. 1 shows a summary of the protein-primed mechanism of ϕ29 DNA replication [10]. Initiation of replication starts by the formation of a TP/ DNA polymerase heterodimer that recognizes the TP-containing DNA ends, the origins of replication. The formation of a nucleoprotein complex of the ϕ29-encoded double-stranded (ds)DNA binding protein (DBP) at the DNA ends facilitates opening of the latter and, in the presence of the initial nucleotide, dATP, stimulates the formation of the covalent linkage between dAMP and the OH group of the Ser232 residue of the TP, a reaction

Fig. 1 Schematic representation of bacteriophage ϕ29 TP–DNA replication. Primer and parental TP are shown in black and green (light gray in print version), respectively. ϕ29 DNAP, DBP, and SSB are colored in blue (gray in print version), red (dark gray in print version), and yellow (white in the print version), respectively. A scheme of the type I and type II replication intermediates is shown at the left. Reproduced from M. de Vega, M. Salas, Protein-primed replication of bacteriophage ϕ29 DNA, in: J.K. Tisma (Ed.), DNA Replication and Related Cellular Processes, InTech, Croatia, 2011, pp. 179–206. See text for details.

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catalyzed by the ϕ29 DNA polymerase (hereafter ϕ29 DNAP). After a transition step, the same polymerase molecule starts the elongation step (DNAprimed) of replication giving rise to the formation of the type I replication intermediates that consist in full-length dsDNA molecules with one or more ssDNA tails covered by the binding of the phage-encoded ssDNA binding protein (SSB). When the two opposite replication forks meet, type I molecules result in two type II molecules where a portion of the DNA, starting from one end, is double-stranded, and the portion spanning to the other end is single-stranded. Type II molecules are fully replicated by the ϕ29 DNAP with concomitant dissociation of the SSB protein to yield two ϕ29 TP– DNA molecules. The final dissociation of the DNA polymerase from each DNA molecule allows the formation of new heterodimers with free TP molecules to initiate a new round of replication.

2.1 Initiation of ϕ29 TP–DNA Replication The first step in ϕ29 TP–DNA replication is the formation of a nucleoprotein complex of the replication origins with the ϕ29 DBP. This is recognized by a heterodimer formed by two viral encoded proteins: the replicative DNA polymerase and a free molecule of TP. 2.1.1 ϕ29 DNA Polymerase 2.1.1.1 Coupling of Processive DNA Synthesis to Strand Displacement

ϕ29 DNAP is a 66 kDa monomeric enzyme, fully responsible for viral DNA replication [19]. Based on amino acid sequence similarities and its sensitivity to specific inhibitors, ϕ29 DNAP was categorized as a family B (eukaryotictype) DNA-dependent DNA polymerase [20]. As other DNA polymerase, it accomplishes sequential template-directed addition of dNMP units onto the 30 OH group of a growing DNA chain, with insertion discrimination values ranging from 104 to 106 and with an efficiency of mismatch elongation from 105- to 106-fold lower than that of a properly paired primerterminus [21]. In addition, ϕ29 DNAP catalyzes 30 –50 exonucleolysis, ie, the release of dNMP units from the 30 end of a DNA strand [22], showing a preferential degradation of a mismatched primer-terminus, agreeing with a role in proofreading of DNA insertion errors that enhances replication fidelity 102-fold [23,24], as it occurs in most DNA replicases. An extensive mutational analysis of individual residues contained in regions of high amino acid similarity among family B DNA polymerases allowed to identify the ϕ29 DNAP catalytic residues required for the synthetic and degradative activities and those responsible for the stabilization of

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the substrates at both active sites (reviewed in Refs. [25,26]; see also Table 1). Thus, sequence alignments and site-directed mutagenesis served to identify the catalytic and ssDNA ligand residues responsible for the 30 –50 exonuclease activity, located at the N-terminal domain (residues 1–189; exonuclease domain), and to propose the hypothesis, widely demonstrated later, of an evolutionary conserved 30 –50 exonuclease active site among distantly related DNA-dependent DNA polymerases [33]. Such an active site is formed by three conserved N-terminal amino acid motifs, ExoI, ExoII, and ExoIII, that contain four carboxylate groups that bind two metal ions, and one tyrosine residue that orients the attacking water molecule [33] (see also Table 1). In addition, these analyses led to identify a new motif (Kx2hxA) whose lysine residue plays an auxiliary role in catalysis, specifically in family B DNA polymerases [34]. Similarly, site-directed mutagenesis studies of ϕ29 DNAP pioneered the functional analyses of specific amino acids at motifs YxGG, Dx2SLYP, LExE, Kx3NSxYG, Tx2GR, YxDTDS, and KxY, placed at the C-terminal domain (residues 190–572; polymerization domain) and highly conserved among the eukaryotic DNA polymerases from family B (see Table 1). These investigations demonstrated the overlapping between polymerization and protein-primed initiation domains and served to identify the amino acids involved in metal binding and catalysis, as well as DNA, TP, and dNTP ligands [25,26,29,30] (see Table 1). ϕ29 DNAP has three distinctive functional features compared to most replicases. First, it initiates DNA replication at the origins located at both ends of the linear genome utilizing a TP as primer and catalyzing the addition of the initial dAMP onto the OH group of Ser232 of the phage TP (reviewed in Refs. [4,10,27]). Second, ϕ29 DNAP performs highly processive DNA synthesis, the highest described for a DNA polymerase (>70 kb; Ref. [11]) without the assistance of processivity factors, unlike most replicases that rely on accessory proteins to be stably bound to the DNA, as thioredoxin for T7 DNA polymerase [35,36], the β-subunit of Escherichia coli Pol III holoenzyme [37], or the eukaryotic clamp protein, PCNA [38,39]. Third, during replication through the ds genome regions, ϕ29 DNAP efficiently couples processive DNA polymerization to strand displacement without the assistance of a helicase-like protein [11]. Resolution of the ϕ29 DNAP structure gave the insights into these two unique properties of the enzyme, processivity and strand displacement capacity. These structural studies, performed in collaboration with Tom Seitz’s lab (Yale University), showed the ϕ29 DNAP structure formed by an N-terminal exonuclease domain, containing the 30 –50 exonuclease active

Table 1 Proposed Role for Individual Residues Belonging to Highly Conserved N-Terminal and C-Terminal Motifs of ϕ29 DNA Polymerase as Defined by Site-Directed Mutagenesis Regiona

Motif

Metal Binding and Catalysis

DNA Binding

TP Binding

dNTP Binding

Strand Displacement

Exonuclease domain

Exol

“DxE”

D12, E14

T15

Exoll

“Nx2–3F/YD”

D66

Y59, H61, N62, F65, F69

Pre-“(S/T)Lx2h” “(S/T)Lx2h” “kx2hxA” Exolll

“Yx3D” “YxGG”

Subdomain Motif

96

K143

D12, E14

110

112

R ,K ,K ,K S122, L123, F128, b 148 Y

F65, Y69, F69, H61

113

96

,

D66

114

R ,K , S122, F128 K143

Y165, D169 Y226, F230, R223

R223, G228, G229, F230

Metal Binding and Catalysis

DNA Binding

TP Binding

D249

S252

Y165, D169 dNTP Binding Strand Displacement

Polymerization domain

Palm

“Dx2SLYP”

c

V250, Y254

K305, Y315

TPR1

D332, K305, Y315 K371, K379

Fingers N387, G391, F393, L384

K383, Y390, K392

Fingers

“Kx3NSxYG”

Palm

“Tx2GR”

R438

Palm

“YxDTDS”

D456, D458

Y454

Palm

“LExE”

d 486

d

Palm

“KxY”

Detailed description of the role of these residues is compiled in Ref. [27]. From Ref. [28]. From Ref. [29]. d From Ref. [30]. e From Ref. [31]. f From Ref. [32]. b c

T434, R438

G481, dW483

E

Thumb a

T434, R438

K498, Y500, eK529 f

538

K

,K

555

,L

567

,T

e 573

,K

575

K529

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Fig. 2 Three-dimensional structures formed by ϕ29 DNAP with DNA and TP. (A) ϕ29 DNAP–DNA complex. (B) Structure of the ϕ29 DNAP/TP complex. The figure was rendered with UCSF Chimera software.

site, and a C-terminal polymerization domain (see Fig. 2) that, like in other DNA polymerases, is subdivided into the universally conserved palm (containing the catalytic residues as well as DNA ligands, Fig. 2A; in dark blue (black in the print version) letters in Fig. 3), fingers (mainly involved in binding the incoming dNTP, Fig. 2A; in pink (gray in the print version) letters in Fig. 3), and thumb (containing DNA ligands which confer stability to the primer-terminus, Fig. 2A; in green (gray in the print version) letters in Fig. 3) subdomains [40]. Although a priori this bimodular structure would be a common theme among proofreading DNA polymerases, the main structural novelty was the presence in the polymerization domain of ϕ29 DNAP of two additional subdomains, both corresponding to sequence insertions that we had previously identified as specifically conserved in the protein-primed subgroup of DNA polymerases, called TPR1 and TPR2 [41,42]. TPR1 lies at the edge of the palm, while TPR2 contains a β-hairpin and forms with the apex of the thumb subdomain an arch-like structure. Palm, thumb, TPR1, and TPR2 subdomains form a doughnutshaped structure that wraps the growing duplex DNA at the polymerization active site [43], functioning as an internal clamp that provides the enzyme with the maximal DNA binding stability required for its intrinsic processivity, analogously to the sliding clamps used by other replicative polymerases (see Fig. 2A). On the other hand, TPR2, palm, and fingers’

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Fig. 3 A schematic representation of ϕ29 DNAP-DNA interactions from one copy of the ternary complex. The base of each nucleotide is represented by a rectangle and a circle represents the phosphate and sugar of a nucleotide. The incoming dNTP is indicated in yellow (light grey in the print version). The residues are color coded according to the subdomain they belong to: dark blue (underlined black in the print version; palm); pink (grey in the print version; fingers); green (underlined grey in the print version; thumb); cyan (light grey in the print version; TPR2); orange (italics grey in the print version; TPR1) and red (black in the print version; exonuclease domain). Asterisks indicate the catalytic residues D249 and D458 from the Dx2SLYP and the YxCDTS motifs, respectively. Figure adapted from “A.J. Berman, S. Kamtekar, J.L. Goodman, J.M. Lázaro, M. de Vega, L. Blanco, M. Salas, T.A. Steitz: Structures of phi29 DNA polymerase complexed with substrate: the mechanism of translocation in B-family polymerases. EMBO J. (2007) 26, 3494-3505. Reproduced with permission from EMBO”.

subdomains, together with the exonuclease domain, encircle the downstream template strand [43], forming a narrow tunnel whose dimensions ˚ ) do not allow dsDNA binding. This fact forces the unwinding of (10 A the downstream dsDNA to allow threading of the template strand through this tunnel to reach the polymerization site, using the same topological mechanism as hexameric helicases to open dsDNA regions, and providing the structural basis for the strand displacement capacity of ϕ29 DNAP [40,44]. However, an intriguing question that remained to be answered was how ϕ29 DNA polymerase could couple the polymerization to the strand displacement, as the polymerase may behave as a “passive” unwinding motor, if translocation of the protein traps transient unwinding fluctuations of the fork, or as an “active” motor, if the polymerase actively destabilizes the

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duplex DNA at the junction. By using optical tweezers it was concluded that the DNA polymerase destabilizes the two nearest base pairs of the fork by maintaining a sharp bending of the template and the complementary strands at a closed fork junction [45]. Bending of the two DNA strands would generate mechanical stress at the junction promoting its active destabilization. The deficient strand displacement capacity displayed by mutants at the exonuclease active site residues [23,34,46] could be explained by a less stable tunnel that would not be able to keep the mechanical stress at a closed fork junction [45]. 2.1.1.2 On the Translocation Mechanism

Additionally, recent resolution of ϕ29 DNAP tertiary complex structures has allowed to dissect the subtle changes in the polymerization active site that take place upon dNTP binding, providing the structural basis for the mechanism of translocation. The incoming dNTP diffuses through a large pore enclosed by residues from the exonuclease domain and from the palm, fingers, and thumb subdomains to gain access to the polymerization active site [40,43]. The insertion site is initially occupied by the aromatic ring of the two conserved residues, Tyr390 (from the fingers subdomain, motif Kx3NSxYG) and Tyr254 (from the palm subdomain, motif Dx2SLYP; see Fig. 4). Binding of the nucleotide at the polymerization site triggers a 14° rotation of the fingers’ subdomain toward the polymerization active site, going from an open to a closed state and allowing electropositively charged residues from the fingers subdomain to bind the α- and β-phosphates of the dNTP. Closing of the fingers moves Tyr390 and Tyr254 out of the nucleotide insertion site into their position in the nascent base pair binding pocket, allowing the base moiety of the incoming nucleotide to form a Watson–Crick base pair with the templating nucleotide, whereas the deoxyribose ring stacks on the phenolic group of Tyr254. Once the catalysis of the phosphoester bond formation between the α-phosphate of the incoming dNTP and the OH group of the priming nucleotide occurs (pretranslocation state), the pyrophosphate produced leaves the DNA polymerase, breaking the electrostatic cross-link that kept the fingers subdomain in the closed state. Concomitant to the fingers opening, residues Tyr254 and Tyr390 move back into the nucleotide insertion site, and the nascent base pair translocates backwards one position (posttranslocation state; Ref. [43]). This translocation allows the 30 OH group of the newly added nucleotide to be in a competent position to prime the following nucleotide insertion event [43] (see Fig. 4).

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Fig. 4 Comparison of the binary (yellow; light gray in the print version) and ternary (green; gray in the print version) complex structures of ϕ29 DNAP. The mechanistically significant amino acid movements are indicated. Reproduced from A.J. Berman, S. Kamtekar, J.L. Goodman, J.M. Lázaro, M. de Vega, L. Blanco, M. Salas, T.A. Steitz, Structures of phi29 DNA polymerase complexed with substrate: the mechanism of translocation in B-family polymerases, EMBO J. 26 (2007) 3494–3505, with permission from EMBO.

Although the structures of the binary and ternary complexes of ϕ29 DNAP provided a structural basis for comprehending the mechanism of translocation, they could not, by themselves, address the thermodynamic and kinetic aspects of the translocation process [43]. In this sense, direct observation of translocation in individual ϕ29 DNAP complexes monitored with single-nucleotide resolution and using the hemolysin nanopore has allowed to conclude that ϕ29 DNAP translocation occurs discretely from the pretranslocation state to the posttranslocation state, driven by Brownian thermal motion [47]. Although nucleotide does not drive translocation, the fluctuation of the binary complexes between the pretranslocation and posttranslocation states is rectified to the posttranslocation state by the binding of complementary dNTP [48]. The movement from the open, posttranslocation state, to the closed pretranslocation state most probably reflects an equilibrium between the fingers-open and fingers-closed states to relieve the steric clash of the primer-terminus with residues Tyr254 and Tyr390 (see earlier), which occlude the nucleotide insertion site when the fingers are open [47].

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2.1.1.3 Coordination of the Polymerization and Proofreading Activities

Remarkably, during the processive DNA replication, a tight and fine-tuned coordination between the polymerization and exonucleolytic cycles should exist to permit a productive but at the same time faithful replication. The dynamics of this coordinated action, the kinetic intermediates involved, and their interconversion remained largely unsolved. 3D resolution of ϕ29 DNAP structure gave the clues about how primer-terminus switches between polymerization and exonuclease active sites during proofreading of polymerization errors. In other proofreading DNA polymerases their thumb subdomains are composed of two helicoidal microdomains linked by a flexible region [49–54] and the primer-terminus switches between both active sites by the rotation of the top microdomain that established the same contacts with the 30 -end of the template strand in both the polymerization and the editing modes. Conversely, the ϕ29 DNAP thumb subdomain has an unusual structure principally constituted by a long β-hairpin without microdomains [40]. Moreover, comparison of the apo enzyme with the binary complexes showed that the thumb subdomain does not rotate upon DNA binding [43]. In fact, prevention of potential thumb movements by cross-linking the tips of the TPR2 and thumb subdomains did not affect the partitioning of the primer-terminus between the polymerization and editing active sites [55]. The impeded motion of the TPR2 subdomain suggested that in ϕ29 DNAP the primer-terminus switching between both active sites is not guided by a rotation of the thumb tip. Moreover, the precluded motion of the TPR2, together with the narrow dimensions of the downstream template tunnel suggested that rotation of the DNA is not required to transfer the primer-terminus between the polymerization and editing active sites in ϕ29 DNAP, most likely as there is not any structural barrier in between to be solved. Then, how does the frayed terminus travel to the exonuclease active site? Recently, by using a single-molecule manipulation method, it has been possible to study the dynamics of the partitioning mechanism by applying varying tension to a processive single ϕ29 DNAPDNA complex [56]. The application of increasing tension has the same effect on the DNA that the insertion of a wrong nucleotide during DNA polymerization. Thus, it has been shown that the application of mechanical force to the template triggers the gradual intramolecular transfer of the primer between the active sites of the protein by decreasing the affinity of the polymerization active site for the template strand and further disrupts the proper dsDNA primer–template structure provoking a fraying of 4–5 bp of dsDNA, allowing primer-terminus to reach the exonuclease active site

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Fig. 5 Schematic representation of the proposed conformational transitions: pol1, initial protein–DNA conformation with DNA bound at the pol domain (blue triangle; black in the print version) and the DNA correctly aligned with the pol active site (yellow circle; light gray in the print version). pol2, polymerase loses contact with template strand, the duplex DNA is still correctly aligned with the pol active site, but translocation is affected (d2). Under these conditions, the protein could explore alternative nonfunctional conformations (pause state) where the DNA and the pol active site may not be correctly aligned. From the pause state, the primer is transferred to the exo domain (red square; dark gray in the print version) by the unwinding of four to five nucleotides of the dsDNA. Red arrows (dark gray in the print version) show the localization of the tension sensitive transitions. Reproduced from B. Ibarra, Y.R. Chemla, S. Plyasunov, S.B. Smith, J.M. Lázaro, M. Salas, C. Bustamante, Proofreading dynamics of a processive DNA polymerase, EMBO J. 28 (2009) 2794–2802, with permission from EMBO.

intramolecularly [56] (see Fig. 5), agreeing with previous in vitro results [57]. Therefore, considering the ϕ29 DNAP thumb subdomain as a nearly static structure, the frayed primer-terminus should switch by a passive diffusion mechanism. The energetically unfavorable gradual melting of 4–5 bp of dsDNA should be progressively offset by new and specific interactions established with DNA ligands of the thumb subdomain, as suggested [32]. Such interactions would also channel the primer-terminus in the appropriate orientation to contact with ssDNA ligands of the exonuclease domain responsible for the stabilization of the primer-terminus at the editing active site [32,40,55,58,59]. Recent development of a single-molecule approach using a nanoscale pore has allowed to quantify with single-nucleotide spatial precision and submillisecond temporal resolution, the rates of translocation, primer strand transfer between the polymerase and exonuclease sites, and dNTP binding in ϕ29 DNAP [48]. It has been shown that transfer of the primer strand from the polymerase to the exonuclease site takes place before translocation, the pretranslocation state being therefore the branch point between the DNA synthesis and editing pathways. Once the 30 terminal nucleotide is cleaved, the primer-terminus returns to the polymerase site and pairs with the

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template strand, the resulting complex being in the posttranslocation state and poised to bind the incoming dNTP and resume DNA synthesis [48]. 2.1.1.4 Biotechnological Applications of ϕ29 DNAP

The two distinctive features of ϕ29 DNAP, high processivity, and strand displacement capacity, together with a remarkably faithful replication, contributed by a high nucleotide insertion fidelity and an intrinsic proofreading activity, led to the development of isothermal multiple displacement amplification (MDA) currently exploited [60,61]. These amplification technologies based on ϕ29 DNAP have two main advantages with respect to classical PCR DNA amplification: first, no previous sequence information is required, due to the use of random hexamer primers, any DNA being susceptible to be amplified, and second, amplicons performed by the ϕ29 DNAP are much larger than those obtained by PCR. On the other hand, the ability displayed by ϕ29 DNAP to use circular multiply primed ssDNA as template has led to the development of the multiply primed rolling circle amplification, one of the most robust technologies to amplify circular templates of variable size [61]. This amplification technology is widely used for genome sequencing, efficient amplification and detection of known and unknown circular viral genomes [62], genotyping of single-nucleotide polymorphisms [63], whole genome analysis of noncultivable viruses [62], detection and identification of circular plasmids in zoonotic pathogens [64], and the description of new metagenomes [65]. Recently, we have achieved improvements of isothermal MDA by fusing DNA binding domains to the C-terminus of ϕ29 DNAP [66]. The results showed that the addition of Helix–hairpin–Helix domains increases DNA binding of the hybrid DNA polymerases without hindering their replication rate. In addition, the chimerical DNA polymerases displayed an improved DNA amplification efficiency on both circular plasmids and genomic DNA and are the only ϕ29 DNAP variants with enhanced amplification performance. These chimerical DNA polymerases will contribute to make ϕ29 DNA amplification technology one of the most powerful tools for genomics, consolidating MDA technology as the alternative to PCR for many applications. 2.1.2 ϕ29 Terminal Protein As already mentioned, the primer TP forms a heterodimer with the DNA polymerase for recognition and further initiation of TP-primed DNA replication. Hereafter, the TP molecule linked to the 50 DNA ends will be called parental TP and the one forming the complex with DNA polymerase,

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primer TP. Crystallographic resolution of the structure of ϕ29 DNAP/ primer TP heterodimer showed that the TP has an elongated three-domain structure (Fig. 2B) [67]. The N-terminal domain (residues 1–73) is structurally disordered, likely because it is not interacting with the polymerase [67,68], and is responsible for DNA binding and TP nucleoid localization [69]. Recent Far-UV circular dichroism spectroscopic analyses have shown a high content of α-helices in this domain [70]. The intermediate domain (residues 74–172) contains two long α-helices and a short β-turnβ structure and makes extensive contacts with the TPR1 subdomain of the polymerase. This interface has many charged residues and includes two salt bridges between arginine residues in the TP and glutamic acid residues in the TPR1 subdomain of the polymerase (Arg158:Glu291; Arg169: Glu322) responsible for conferring stability to the TP/DNA polymerase heterodimer as well as for guaranteeing a functional interaction between both proteins [71]. The intermediate domain is connected through a hinge region to the C-terminal priming domain (residues 173–266), a region highly electronegative that has a four-helix bundle topology. The priming Ser232, which provides the hydroxyl group for DNA synthesis, lies in a disordered loop (residues 227–233) at the end of the priming domain and should be positioned at the active site of the DNA polymerase to allow the initiation reaction to take place (see Fig. 6, left panel). Positioning of the priming loop at the polymerase active site is stabilized through a salt bridge between TP residue Asp233 and DNA polymerase residue Lys529 [31]. The priming domain structure shows interactions between many of their acidic residues and positively charged residues of the thumb subdomain of the polymerase (eg, between Glu191:Lys575 and Asp198: Lys557), with residue Arg96 of the exonuclease domain and with TPR2 subdomain residues [67,72]. Those contacts play a role in the maintenance of the equilibrium between TP-priming domain stabilization and its gradual exit from the polymerization active site of the DNA polymerase as the new DNA is synthesized [71]. The overall dimensions and negative charge of the TP-priming domain allow it to occupy during the initiation reaction the same upstream DNA “tunnel” in the polymerase as duplex DNA does during elongation [67,73] (see also Fig. 2B). This explains why DNA synthesis by the heterodimer cannot begin at internal sites of the phage genome, as the upstream 30 template would sterically clash with the TP [67]. Our previous studies showed that the DNA polymerases of ϕ29 and the related phages GA-1 and Nf display a great specificity for their corresponding primer TPs, as the heterologous systems did not give any

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Fig. 6 Modeling of the placement of the TP-priming loop at the ϕ29 DNAP active site. Left: placement of TP-priming residue Ser232 (in lemon green; gray in the print version) and penultimate template nucleotide at the ϕ29 DNAP active site (catalytic aspartates and Mg2+ ions are shown in red (dark gray in the print version) and gray, respectively). Right: flexible orientations of the TPR1 loop in the apoenzyme (colored in magenta; gray in the print version) and its stable and moved out structural conformation shown in the DNA polymerase/TP complex (colored in orange; gray in the print version). TP is colored in yellow (light gray in the print version). Green arrows (dark gray in the print version) indicate the suggested conformational changes of both, the DNA polymerase TPR1 loop and the TP-priming domain to allow the formation of a stable heterodimer. Reproduced from M. de Vega, M. Salas, Protein-primed replication of bacteriophage ϕ29 DNA, in: J.K. Tisma (Ed.), DNA Replication and Related Cellular Processes, InTech, Croatia, 2011, pp. 179–206 and reproduced with permission from P. Pérez-Arnaiz, E. Longás, L. Villar, J.M. Lázaro, M. Salas, M. de Vega, Involvement of phage ϕ29 DNA polymerase and terminal protein subdomains in conferring specificity during initiation of protein-primed DNA replication, Nucleic Acids Res. 35 (2007) 7061–7073.

detectable initiation product [74,75]. By means of chimerical proteins, constructed by swapping the priming domain of the related ϕ29 and GA-1 TPs, we showed that DNA polymerase can form catalytically active heterodimers exclusively with the chimerical TP containing the intermediate domain of the homologous TP, indicating that the interaction between the polymerase TPR1 subdomain and the TP-intermediate domain is the one main responsible for the specificity between both proteins [68]. In addition, the independent expression of the ϕ29 TP-priming domain and intermediate plus N-terminal domains showed that the former can only prime initiation in the presence of the latter that assists the TP-dAMP formation most probably by inducing a conformational change in the DNA

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polymerase [68]. The structure of the ϕ29 DNAP forming a complex with the TP is very similar to that of the apo enzyme, the main conformational changes being restricted to TPR1 residues 304–315 [67]. Such residues form loops with a high degree of flexibility in the apo enzyme. By the contrary, the ϕ29 DNAP/TP heterodimer structure shows that this loop moves out to allow the TP to access the polymerase active site. Altogether, these results led to propose a model for the DNA polymerase–TP interaction in which the TP-intermediate domain would recognize specifically and interact with the DNA polymerase TPR1 subdomain. Such interaction would promote the change of the TPR1 loop from a flexible to the stable moved out conformation that now would allow the proper (prone to catalysis) placement of the TP-priming domain into the DNA polymerase structure [68] (see Fig. 6, right panel). 2.1.3 Recognition of Replication Origins by the TP/ DNA Polymerase Heterodimer The ϕ29 TP/DNAP heterodimer recognizes the replication origins at the genome ends (see Fig. 1). Blunt-ended DNA fragments containing the left or right ϕ29 DNA ends, but not internal ϕ29 DNA fragments, were active as templates in in vitro initiation reactions [76–78]. However, the activity was 6- to 10-fold lower than that obtained with TP–DNA [77,78]. These results indicated, on the one hand, that specific DNA sequences located at the ϕ29 DNA ends are involved in origin recognition and, on the other hand, that the parental TP is a major signal in the template for such a recognition, strongly suggesting that the heterodimer is recruited to the origin through interactions with the parental TP. In agreement with this, detection of initiation activity by using heterologous systems in which DNA polymerase, primer TP, and TP–DNA were from the ϕ29- and Nf-related phages, showed that initiation was selectively enhanced when the DNA polymerase and the TP–DNA were from the same phage, implying a specific interaction between DNA polymerase and parental TP [74]. In line with this, a chimerical ϕ29 DNAP containing the GA-1 DNA polymerase TPR1 subdomain catalyzed the initiation reaction primed by GA-1 TP but only in the presence of ϕ29 TP–DNA, supporting the hypothesis that a major contribution to the parental TP recognition is performed by the DNA polymerase [68]. Similarly, mutations introduced at several TP-intermediate domain residues rendered TP mutants that could not support DNA replication when they acted as parental TP, indicating also a contribution of the primer TP in the specific recognition of the replication origins [71,79,80].

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2.2 A Sliding-Back Mechanism for Protein-Primed DNA Replication As already indicated, the DNA ends of ϕ29 and those of the ϕ29-related phages have a reiteration of three nucleotides (30 -TTT…50 ). Once the replication origins are specifically recognized by the TP/DNA polymerase heterodimer [68,74,81–83], the DNA polymerase catalyzes the formation of a phosphoester bond between the initial dAMP and the hydroxyl group of Ser232 of the TP (see Fig. 1), a reaction directed by the second T at the 30 end of the template strand [84] and performed by the same DNA polymerase catalytic residues responsible for canonical polymerization [25,26]. During the initiation reaction, the priming Ser232 of TP should be placed at the catalytic site of the DNA polymerase in line to attack nucleophilically the α-phosphate of the initial nucleotide (Fig. 6, left panel). Therefore, the 30 end of the template strand should enter deep into the catalytic site of the DNA polymerase to place the penultimate 30 dTMP of the template strand at the catalytic site, as it directs the first nucleotide insertion reaction. By using chimerical TPs, constructed by swapping the priming domain of the related ϕ29 and Nf proteins, we showed that this domain is the main structural determinant that dictates the internal 30 nucleotide used as template during initiation, the second and third in ϕ29 and Nf DNA, respectively [85]. Recently, we have shown that the aromatic residue Phe230 of the ϕ29 TP-priming loop is the one responsible for positioning the penultimate nucleotide at the polymerization site to direct insertion of the initial dAMP during the initiation reaction, most probably by interacting with the 30 terminal base, limiting the internalization of the template strand [86]. To perform TP–DNA full-length synthesis, the TP-dAMP initiation product translocates backward one position to recover the template information corresponding to the first 30 -T, the so-called sliding-back mechanism that requires a terminal repetition of 2 bp. This reiteration permits, prior to DNA elongation, the asymmetric translocation of the initiation product, TP-dAMP, to be paired with the first T residue [84] (see scheme in Fig. 7). Our studies have shown how the sliding-back mechanism, or variations of it, seems to be a common feature of protein-priming systems to restore full-length DNA. Thus, in the case of the ϕ29-related phage GA-1, initiation occurs mainly at the 30 second nucleotide of the template (30 TTT) [87]. The ϕ29-related phage Nf and the Streptococcus pneumoniae phage Cp-1 initiate at the 30 third nucleotide of their terminal repetition (30 -TTT) [85,88], whereas the E. coli phage PRD1 initiates at the fourth nucleotide

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Fig. 7 Sliding-back (jumping-back) model for the transition from initiation to elongation. The figure compares the three mechanisms described during initiation of TP–DNA replication: sliding-back (bacteriophage ϕ29), stepwise sliding-back (bacteriophage PRD1), and jumping-back (Adenovirus), see text for details. TP is represented as a pink (dark gray in the print version) oval and DNA polymerase as a gray square. The internal template nucleotide that directs the insertion of the initial nucleotide is shown in bold red (gray in the print version) letter. Yellow (light gray in the print version) box represents the catalytic active site of the DNA polymerase. Reproduced from M. de Vega, M. Salas, Protein-primed replication of bacteriophage ϕ29 DNA, in: J.K. Tisma (Ed.), DNA Replication and Related Cellular Processes, InTech, Croatia, 2011, pp. 179–206.

(30 -CCCC) [89], requiring two and three consecutive sliding-back steps, respectively, to recover the DNA end information (stepwise sliding-back). The adenovirus genome ends present a more complex reiteration (30 GTAGTA), the 30 fourth to six template positions directing the formation of the TP-CAT initiation product. Thus, recovery of the 30 ends is

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performed by a single jump, after which TP-CAT is paired with the terminal 30 -GTA (jumping-back) [90] (see scheme in Fig. 7). What is the rationale of the sliding-back mechanism? ϕ29 proteinprimed initiation is an unfaithful reaction with a nucleotide insertion discrimination factor about 102. Moreover, unlike the exonucleolytic elimination of a misinserted dNMP during the DNA-primed elongation stage, the proofreading activity of ϕ29 DNAP is unable to act on the TP-dAMP initiation complex, precluding the possibility that a wrong dNMP covalently linked to TP could be released [21]. The sliding-back mechanism and its variants have been envisaged to increase the fidelity during the initiation reaction, as several base pairing checking steps have to occur before definitive elongation of the initiation product takes place [84,90]. Thus, if a nucleotide is misincorporated during the initiation reaction, the erroneous TP-dNMP complex will not pair with the terminal 30 -T of the template after the sliding-back, hindering its further elongation. Moreover, if an incorrect TP-dNMP product were elongated, it would be corrected in the subsequent round of replication, because it could not serve as a template [21]. The presence of sequence repetitions at the ends of other TP-containing genomes supports the hypothesis that the sliding-back type of mechanism could be a common feature of protein-primed replication systems [84].

2.3 Transition from Protein-Primed to DNA-Primed Replication Functional analyses established that the ϕ29 DNAP/primer TP heterodimer does not dissociate immediately after initiation or after sliding-back [91]. The same DNA polymerase molecule incorporates five nucleotides to the primer TP while it is still complexed with the latter (initiation mode), undergoes some structural change during incorporation of nucleotides 6–9 (transition), and finally dissociates from the primer TP when nucleotide 10 is incorporated into the nascent DNA chain (elongation mode) [91]. These results probably reflect the polymerase requirement for a DNA primer of a minimum length to catalyze DNA elongation efficiently. We have shown that the strength of the ϕ29 DNAP/primer TP interaction is differently contributed by the TP-priming and intermediate domains [68], supporting the model proposed for the transition from the protein-primed initiation to the DNA-primed elongation modes [67]. Thus, the TP-intermediate domain would be in a fixed orientation on the polymerase through stable contacts with the TPR1 subdomain. The weak interaction observed with the DNA polymerase would facilitate

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Fig. 8 A model for the transition from initiation of replication to elongation (see main text for details). Reproduced from S. Kamtekar, A.J. Berman, J. Wang, J.M. Lázaro, M. de Vega, L. Blanco, M. Salas, T.A. Steitz, The phi29 DNA polymerase:protein-primer structure suggests a model for the initiation to elongation transition, EMBO J. 25 (2006) 1335–1343, with permission from EMBO.

the TP-priming domain to rotate following the helicoidal pathway as DNA is synthesized. The relative motion of the TP-priming domain with respect to the fixed TP-intermediate domain would be possible due to the flexibility of the hinge region that connects both domains. After incorporation of six to seven nucleotides the proximity of the priming Ser to the hinge region would impede a further priming domain rotation, promoting complex dissociation [67] (see Fig. 8).

2.4 DNA-Primed Elongation Once the initiation, sliding-back, and transition steps have been fulfilled and ϕ29 DNAP has separated from the primer TP, the DNA polymerase resumes TP–DNA replication; therefore, the same DNA polymerase molecule accounts for complete genome replication from a single binding event [11]. As mentioned before, the high stability of the ϕ29 DNAP/ DNA complex, by virtue of the "internal sliding-clamp-like" structure formed by thumb, palm, TPR1, and TPR2 subdomains, allows the polymerase to perform complete DNA replication without the assistance of

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processivity factors, unlike most replicative DNA polymerases. In addition, the singular TPR2 subdomain enables the ϕ29 DNAP to couple polymerization to the unwinding of the downstream dsDNA regions (strand displacement capacity) making unnecessary the intervention of a helicaselike protein [11] (see Fig. 9). As described earlier, since replication starts at both ϕ29 DNA ends and is coupled to strand displacement, this results in the generation of the so-called type I replication intermediates (see Fig. 1). The ssDNA stretches generated are bound by the viral SSB, essential for elongation of replication in vivo [92]. Binding of ϕ29 SSB to ϕ29 DNA replicative intermediates has been

Fig. 9 Modeling processivity and strand displacement in ϕ29 DNAP. The TPR2 insertion would contribute to a full encirclement of the DNA substrate, conferring a remarkable processivity, and also acts as a structural barrier, which would force the DNA strands of the parental DNA to diverge (melt). Because ϕ29 DNAP translocates after each polymerization cycle, the TPR2 subdomain would act as a wedge to couple polymerization to strand displacement. In the figure the subdomains palm, thumb, TPR1, and TPR2 are colored in pink (gray in the print version), green (gray in the print version), orange (gray in the print version), and cyan (gray in the print version), respectively. The 30 -50 exonuclease domain is colored in red (dark gray in the print version). Reproduced with permission from I. Rodríguez, J.M. Lázaro, L. Blanco, S. Kamtekar, A.J. Berman, J. Wang, T.A. Steitz, M. Salas, M. de Vega, A specific subdomain in f29 DNA polymerase confers both processivity and strand displacement capacity, Proc. Natl. Acad. Sci. U.S.A. 102 (2005) 6407–6412. Copyright (2005) National Academy of Sciences, U.S.A.

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demonstrated to occur in vitro [93]. The protein binds in a cooperative way [94] stimulating dNMP incorporation during ϕ29 DNA replication [95] and increasing the elongation rate, mainly when ϕ29 DNAP mutants impaired in strand displacement are used, probably due to the helix destabilizing activity of the ϕ29 SSB [96]. When the two converging DNA polymerases merge, a type I replication intermediate becomes physically separated into two type II replication intermediates [93,97]. Continuous elongation by the DNA polymerase completes replication of the parental strand.

2.5 TP-Primed DNA Amplification By using appropriate amounts of the four ϕ29 DNA replication proteins, primer TP, DNA polymerase, DBP, and SSB, we could amplify limited amounts of the ϕ29 TP–DNA molecule by three orders of magnitude after 1 h of incubation at 30°C [98]. Moreover, the quality of the amplified DNA was demonstrated by transfection experiments, in which infectivity of the synthetic (amplified) ϕ29 TP–DNA, measured as the ability to produce phage particles, was identical to that of the natural ϕ29 TP–DNA obtained from virions, leading us to establish some of the requisites for the development of isothermal DNA amplification strategies based on the ϕ29 DNA replication machinery to amplify very large (>70 kb) segments of exogenous DNA. In this sense, we have recently reported a method to amplify in vitro heterologous DNAs using the ϕ29 DNA replication proteins mentioned earlier. The DNA to be amplified was inserted between two ϕ29 DNA replication origins without parental TP that recruited the TP/DNA polymerase heterodimer [99]. In the presence of the DBP and SSB, the ϕ29 replication system amplified 30-fold the heterologous DNA, rendering DNA with TP covalently attached to the 50 ends. Therefore, this method opens the possibility to amplify in a defined way long linear DNAs, like viral genomes, chromosome fragments, or plasmids. Additionally, it would be potentially achievable to produce in vitro high amounts of DNAs bound to TP fused to diverse targeting peptide modules required for gene delivery. Finally, it would be possible to use the ϕ29 replication machinery to generate TP–DNAs with different biological activities [99].

3. IN VIVO COMPARTMENTALIZATION OF ϕ29 DNA REPLICATION It is well established that replication of phage genomes occurs at specific intracellular locations using large organizing structures that bring

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together replication factors to enhance the efficiency of the replication process. Some lines of evidence support that replication of phage DNA takes place in close association with the bacterial membrane [100–102]. Recent investigations have also given evidences concerning compartmentalization of phage ϕ29 DNA replication in B. subtilis cells. We have shown that ϕ29 TP binds in vitro to dsDNA and in vivo to the host nucleoid, thanks to its sequence-independent DNA binding capacity [69,103]. This ability is conferred to the TP by the high density of positively charged amino acids of the N-terminal domain [70]. Recent results have shown that nucleoid localization is not restricted to ϕ29 TP, as the TPs from phages as PRD1 and CP-1 that infect a variety of Gram-negative bacteria and the Gram-positive bacterium S. pneumoniae, respectively, also target the nucleoid through the N-terminal positively charged motifs [104]. This capacity enables the parental TP, and therefore the viral TP–DNA, to associate with the bacterial nucleoid early after injection of the ϕ29 genome (see scheme in Fig. 10) where the B. subtilis RNA polymerase is also located [69]. There, synthesis of the ϕ29 early proteins DNA polymerase, primer TP, SSB, and DBP, essential for in vivo ϕ29 DNA replication, takes place. Once synthesized, the TP/DNAP heterodimer binds the bacterial chromosome and recognizes the TP–DNA replication origins. At this stage, replication of TP–DNA will start from both terminal origins giving rise to the

Fig. 10 Model of nucleoid-associated early ϕ29 DNA replication organized by the TP. (A) Attachment of ϕ29 TP–DNA to the bacterial nucleoid surface (gray mass at bottom) through the N-terminal domain (red; dark gray in the print version) of the parental TPs (red (dark gray in the print version) and green (light gray in the print version)). (B) Recruitment of the ϕ29 DNAP/primer TP heterodimer to the replication origins of TP–DNA. (C) Processive elongation of the nascent DNA strands (red (dark gray in the print version) lines) coupled to strand displacement. (D and E) Once DNA replication is completed, two ϕ29 TP–DNA molecules are ready for another round of replication. For simplicity, other viral proteins involved in DNA replication are not drawn. Reproduced with permission from D. Muñoz-Espín, R. Daniel, Y. Kawai, R. Carballido-López, V. Castilla-Llorente, J. Errington, W.J. Meijer, M. Salas, The actin-like MreB cytoskeleton organizes viral DNA replication in bacteria, Proc. Natl. Acad. Sci. U.S.A. 106 (2009) 13347–13352.

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replicative intermediates type I and II, as it has been observed by electron microscopy [97,105,106]. By fluorescence microscopy, it has been shown that at middle infection times, the DNA polymerase, TP, and viral TP–DNA are reorganized adopting a peripheral helix-like distribution toward the poles of the cell [69,107]. Although the pathway followed by the ϕ29 replicative machinery from the nucleoid to bacterial peripherical regions remains to be determined, it has been proposed that it could travel associated to the replication of the bacterial chromosome [69], as newly synthesized bacterial DNA is translocated toward the cell poles via a helical structure [108]. The ϕ29 membrane protein p16.7 has a nonsequence-specific DNA binding capacity [109] that enables it to interact with the ϕ29 replication origins through recognition of the parental TP [110]. This protein also shows a helix-like pattern at the membrane of infected cells, most probably being involved in the compartmentalization of in vivo membrane-associated ϕ29 DNA replication through a direct contact with TP–DNA, organizing the viral replicating intermediates at numerous peripheral locations [107,111] (see Fig. 11). Different experimental approaches have shown that

Fig. 11 Model of membrane-associated late ϕ29 DNA replication organized by the MreB cytoskeleton. MreB, Mbl, and MreBH are shown to form a putative triple helical structure closely associated with the inner surface of the membrane. Each dimeric unit of protein p16.7 is represented by a yellow (gray in the print version) hexagon. The tridimeric p16.7 units form oligomers in a helix-like localization at the cell membrane. For simplicity, other viral proteins involved in DNA replication are not drawn. Reproduced with permission from D. Muñoz-Espín, R. Daniel, Y. Kawai, R. Carballido-López, V. Castilla-Llorente, J. Errington, W.J. Meijer, M. Salas, The actin-like MreB cytoskeleton organizes viral DNA replication in bacteria, Proc. Natl. Acad. Sci. U.S.A. 106 (2009) 13347–13352.

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protein p16.7 interacts directly with the B. subtilis actin-like cytoskeleton protein MreB [107]. This protein forms helix-like filamentous structures in vivo essential for the control of the bacterial rod-shaped morphology [112], suggesting that MreB would contribute to efficient ϕ29 DNA replication by recruiting protein p16.7 to the appropriate sites at the cell membrane allowing simultaneous replication of multiple templates at numerous peripheral locations. Further evidence is the finding that ϕ29 DNA replication is severely affected in ΔMreB cytoskeleton mutants [107]

ACKNOWLEDGMENTS This work has been supported by grants from the Spanish Ministry of Economy and Competitiveness (BFU2014-52656-P to M.S.) and (BFU2014-53791-P to M.V.) and by an Institutional grant from Fundacio´n Ramo´n Areces to the Centro de Biologı´a Molecular “Severo Ochoa.”

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CHAPTER FIVE

Archaeal DNA Replication Origins and Recruitment of the MCM Replicative Helicase R.Y. Samson, S.D. Bell1 Indiana University, Bloomington, IN, United States 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Origin Recognition in Eukaryotes 3. Recruitment of the Replicative Helicase 4. Archaeal Replication Origin Specification 5. Structure of Archaeal Orc1/Cdc6 Proteins 6. Role of ATP in MCM Loading 7. The Mechanism of MCM Recruitment by Orc1–1 8. Concluding Remarks Acknowledgments References

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Abstract DNA replication is fundamental to the propagation of all life on the planet. Remarkably, given the central importance for this process, two distinct core cellular DNA replication machineries have evolved. One is found in the bacterial domain of life and the other is present in Archaea and Eukarya. The archaeal machinery represents a simplified and presumably ancestral form of the eukaryotic DNA replication apparatus. As such, archaeal replication proteins have been studied extensively as models for their eukaryal counterparts. In addition, a number of archaea have been developed as model organisms. Accordingly, there has been a considerable increase in our knowledge of how archaeal chromosomes are replicated. It has become apparent that the majority of archaeal cells replicate their genomes from multiple origins per chromosome. Thus, at both organizational and mechanistic levels, archaeal DNA replication resembles that of eukarya. In this chapter, we will describe recent advances in our understanding of the basis of archaeal origin definition and how the archaeal initiator proteins recruit the replicative helicase to origins.

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1. INTRODUCTION It is estimated that approximately 20% of the Earth’s biomass is archaeal, yet we still know very little about the biology of these diverse organisms. Until Carl Woese’s landmark work published in 1977, archaea were merely regarded as quirky bacteria [1]. We now know that Archaea are distinct from Bacteria and belong to a separate domain of life. Based on currently available sequencing data, Archaea have been divided into five major phyla: Euryarchaea, Thaumarchaea, Aigarchaea, Crenarchaea, and Korarchaea. Due to close phylogenetic relatedness between the latter four groups, it has been suggested that they represent a coherent group, named the TACK superphylum [2]. However, a quiet revolution has been happening in the field of evolutionary biology, with the emergence of data supporting the notion that all life on Earth can be categorized into two taxonomic domains, not three as we have believed for almost four decades [3]. In this two domain tree of life, Eukarya are actually a sister group to the TACK superphylum within the domain Archaea. Improvements in sampling and sequencing procedures as well as new methods in phylogenetic reconstruction have provided evidence that an archaeon was the host that engulfed the bacterial protomitochondrial endosymbiont, a key event in the prokaryote-to-eukaryote transition. In agreement with the proposed archaeal origin of eukaryotic cells, components of many biological processes essential to extant life forms are shared between these two groups. Certainly, it appears that key factors involved in chromosome replication were established before the divergence of the archaeal and eukaryotic lineages [4]. There are no better examples of this ancient origin than the Orc1/Cdc6 candidate replication initiator proteins and MCM replicative helicase. With the exception of the euryarchaeon Methanopyrus kandleri, all archaeal genomes encode at least one orc1/cdc6 gene and all archaea have MCM. As seen in eukaryotes, there seems to be flexibility in the constitution of and the interactions between initiator proteins and DNA replication origins [5,6]. For example, Pyrococcus species have only one origin of replication that is governed by a single Orc protein, whereas marker frequency analysis of genomic DNA from Pyrobaculum calidifontis has suggested that this organism possesses four replication origins [7,8]. Additionally, as discussed later, one of the three chromosomal origins in Sulfolobus solfataricus is controlled by a

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complex of two Orc1/Cdc6 proteins, in contrast to a one-Orc-per-origin rule that seems to be adopted by other archaea [9–11]. In the Euryarchaea, there is a great deal of variation in the number of genes for Orc1/Cdc6 proteins. For example, while Pyroccocus has a single orc1/cdc6, some halophiles encode 15 or more Orc1/Cdc6 paralogs. In addition, Euryarchaea are generally highly polyploid with no evidence currently for gap-phases between DNA synthesis and cell division [12,13]. In contrast, all studied TACK archaea appear to oscillate between one and two chromosomes throughout a cell cycle that has defined G1 and G2 phases [14]. While the Thaumarchaeaon Nitrosopumilus maritimus has a single replication origin [15], a diverse range of Crenarchaea possess two or more replication origins per chromosome [8,9,14,16–18]. Since archaeal DNA replication has a fundamental relationship to the far more extensively studied eukaryotic machinery, we will briefly summarize recent findings on the mechanisms of origin specification and replicative helicase recruitment in eukaryotes before describing studies of the archaea.

2. ORIGIN RECOGNITION IN EUKARYOTES Several eukaryotic model systems have been exploited to study DNA replication including: Saccharomyces cerevisiae, Schizosaccharomyces pombe, Drosophila melanogaster, Xenopus laevis and Mus musculus, and Homo sapiens cell lines. However, arguably the most detailed molecular biological work on the initiation phase of replication has been done in yeast systems. In S. cerevisiae, the process of replication initiation begins when a complex of six proteins, called ORC (origin recognition complex), binds to specific sites on the chromosome. Orc1–Orc5 belong to a family of proteins called the AAA + (ATPases associated with a variety of cellular activities) family [19]. Sequence specificity of the S. cerevisiae ORC appears to be affected by a related AAA+ protein called Cdc6. Indeed, phylogenetic studies have revealed Orc1 and Cdc6 to be related to one another and presumably derived from a common ancestor. Origin-bound ORC recruits Cdc6 and this protein assembly directs the recruitment and loading of a replicative helicase complex. Loading of the heterohexameric MCM(2–7) replicative helicase requires an additional factor, Cdt1, that serves as a coloader. Conceptual, if not mechanistic, parallels can be drawn with the well studied, yet nonorthologous, Escherichia coli replication system in which the initiator DnaA recruits the replicative helicase, DnaB, via the actions of a dedicated helicase loader protein, DnaC [19]. Thus, both eukaryotes and bacteria exploit

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dedicated helicase loader proteins as intermediaries between origin-bound initiator proteins and incoming replicative helicases. While specific consensus sequences for origins have been identified in S. cerevisiae, this is not the case in other yeast species. Indeed, while there is a preference for initiation to occur at AT-rich sequences in some fission and budding yeast, origins in Schizosaccharomyces japonicus are GC rich. It is believed that chromatin environment is an important factor in determining replication origins in yeast and metazoans, and both poly(G) motifs and AT-rich regions have been shown to exclude nucleosomes. In metazoans, origins appear to be very plastic in nature and are probably not hardwired by specific sequences in the DNA. Epigenetic factors, such as histone acetylation, affect the distribution of origins on chromosomes and changes in epigenetic marks as an organism transitions through different developmental stages affect the landscape of active replication origins [20]. Thus, the molecular determinants for origins of replication in many eukaryotes are still poorly resolved and the specific requirements clearly vary between organisms. Remarkably, the composition of the complex that defines origins also appears to vary throughout the eukaryotic domain. With the exception of members of the microsporidia, parasites with reduced genomes, members of the supergroup Opisthokonta, including metazoans and fungi, possess the complete array of Orc proteins, Orc1–Orc6, Cdc6, and the helicase coloader Cdt1 (Fig. 1). However, examination of the initiator gene content of earlier-branching eukaryotes indicates that many of these organisms have an ORC of considerably reduced complexity. A notable example is seen in the deepest-branching eukaryotic lineage, the Excavata. This supergroup includes a number of significant human pathogens such as Trypanosomes. These organisms encode a single protein that is related to both Orc1 and Cdc6 [22,23]. Presumably gene duplication and diversification during the evolution of the eukaryotic domain of life led to the appearance of the separate Orc1 and Cdc6 proteins found in later branching eukaryotic lineages. It should be recognized that the apparent absence of genes in some lineages could arise from extreme sequence divergence of ORC components, rendering orthologs undetectable by conventional BLAST algorithms used to compile this table. Alternatively, gene loss in the streamlined genomes of parasitic organisms could account for the patterns observed. However, we favor the interpretation that the complexity of ORC has increased over evolutionary time, concomitant with the increasing size of eukaryotic genomes. Presumably increased subunit complexity confers increased regulatory potential to help ensure the fidelity of initiation in organisms that possess thousands of replication origins [24].

Fig. 1 Distribution of prereplicative complex proteins in the eukaryal and archaeal domains of life. The question marks in the Orc2 and Orc5 columns arise because Orc1/Cdc6 in Trypanosomes associated with additional AAA+-related proteins that may correspond to Orc2 and Orc5 in higher eukaryotes [21]. Figure adapted from C. Tiengwe et al., Identification of ORC1/CDC6-interacting factors in Trypanosoma brucei reveals critical features of the origin recognition complex architecture, PLoS One 7 (3) (2012) e32674.

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Furthermore, the closest prokaryotic ancestors of eukaryotes, the archaea, like the deepest-branching eukaryotes, possess the presumptive ancestral Orc1/Cdc6 protein. Given that Orc1 serves a role as an initiator protein within ORC and Cdc6 is viewed has a helicase loader protein, it has long been speculated that the archaeal Orc1/Cdc6 proteins may fulfill both origin specification and helicase loading functions [25].

3. RECRUITMENT OF THE REPLICATIVE HELICASE The compositional complexity of the origin specification and MCM helicase recruitment machinery vary across the eukaryotic domain. However, the enzyme responsible for melting double-stranded DNA during replication, the MCM helicase, appears to operate under tighter selective forces. All sequenced eukaryotic genomes possess six genes encoding the constituents of a conserved heterohexameric complex, MCM(2–7). The subunits assemble with defined organization of MCM2–6–4–7–3–5. All six subunits appear to be derived from a common ancestor and belong to the AAA+ superfamily of ATPases [26]. Based primarily on evidence compiled in yeast labs, we know that ORC and Cdc6 are bound to origins of replication during the G1 phase of the cell cycle. Through the concerted actions of Cdc6 and Cdt1, MCM(2–7) is recruited and loaded onto doublestranded DNA as two head-to-head hexameric rings [19]. The exact details of this loading process are currently unclear, but the recruitment of MCM (2–7) appears to be mediated by interactions between the C-terminus of MCM3 and Cdc6 and also between Cdt1 and Orc6 [27]. In addition to supplying a recruitment module, Cdt1 has also been shown to stabilize the MCM2,4,6 subunits within the MCM hexamer [27]. Initially, only a single hexamer of MCM is recruited to DNA, forming what has been called the OCCM (ORC, Cdc6, Cdt1, MCM) complex. ATP hydrolysis then occurs together with ejection of Cdc6 from the complex, this is followed rapidly by departure of Cdt1 [28]. A second molecule of Cdc6 then binds to ORC
MCM just prior to the recruitment of a second MCM
Cdt1 complex. This work confirms that MCM(2–7) is loaded in two distinct steps by a single ORC and it has been proposed that the mechanism for loading the second hexamer is distinct from the first and relies on N-terminal interactions between the MCM subunits [28]. One of the biggest mysteries in the loading of budding yeast MCM is how one asymmetrical ORC can load two MCM hexamers in opposite orientations on the same side of ORC using the same protein–protein interaction interfaces [21,27,29,30].

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4. ARCHAEAL REPLICATION ORIGIN SPECIFICATION The first archaeal origin of DNA replication characterized was from Pyrococcus abyssi. Radiolabeling studies and subsequent 2D neutral–neutral agarose gel analyses provided evidence for a single replication origin in the P. abyssi chromosome [7,31]. Chromatin immunoprecipitation (ChIP) studies revealed that the single Orc1/Cdc6 paralog encoded by P. abyssi bound to the origin. Notably, the orc1/cdc6 gene was encoded adjacent to the origin. This cis-relationship of initiator and origin is conserved in a broad range of archaea from both TACK and Euryarchaea groupings and is also common in bacteria, where DnaA is often encoded adjacent to oriC. However, the single origin system of Pyrococcus seems to be something of an oddity in the archaeal domain, as a range of archaea possesses multiple origins [8,9,13,16–18,32–35]. Genetic studies in multiorigin organisms such as Sulfolobus and the halophilic archaea have revealed that the activities of individual origins are nonessential [18,35,36]. However, in Sulfolobus at least one origin is required to ensure genome maintenance [18]. Similarly, in Haloarcula hispanica, either one of the two origins of the main chromosome can be deleted, however, it was not possible to delete both origins [35]. A recent study of another haloarchaeon, Haloferax mediterranei, revealed that its main chromosome is replicated from three origins. Remarkably, it was possible to delete all three origins. Cell viability was maintained by activation of a dormant replication origin, not normally used in wild-type cells in logarithmic growth [37]. An even more extreme example was seen in the related species Haloferax volcanii. In H. volcanii, it was possible to delete all main chromosome origins (three or four depending on whether the isolate in question has an integrated chromosomal element) and retain viability [36]. In contrast to H. mediterranei, there was no evidence for the activation of a single dormant origin in H. volcanii to restore growth, rather, viability was dependent on the product of the radA gene. RadA is the archaeal ortholog of bacterial RecA and eukaryal Rad51 recombinase and so it was proposed that replication of the “origin-less” chromosome was dependent on homologous recombination, as has been seen in studies of stable DNA replication in E. coli [38]. At the current time, probably the most highly studied replication origins are those of the Sulfolobus species. As alluded to earlier, the genomes of Sulfolobus are replicated from three origins, oriC1, oriC2, and oriC3 [9,17,32]. All three origins fire once in every cell cycle [39]. Sulfolobus

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species encode three Orc1/Cdc6 candidate initiator proteins [40]. For the reasons outlined later, Orc1–1 and Orc1–3 are believed to promote replication. In contrast, several studies have revealed that mRNA levels for Orc1–2 are specifically upregulated in response to a variety of cellular stresses [41–45] and thus Orc1–2 is believed to be a negative regulator of replication. In addition, a divergent homolog of Cdt1, termed WhiP, was identified in the Sulfolobus genus and other members of the Sulfolobales and Desulfurococcales [16]. Initial ChIP studies have suggested some species-specific variations with regard to origin binding by the Orc1/Cdc6 initiators. More specifically, in S. solfataricus Orc1–1 bound to both oriC1 and oriC2 and Orc1–3 bound to oriC2 [9]. The cooccupancy of oriC2 by Orc1–1 and Orc1–3, with direct protein–protein interactions between Orc1–1 and Orc1–3, has been confirmed biochemically and structurally [9–11]. ChIP studies of S. solfataricus oriC3 have not been reported. In contrast, Sulfolobus acidocaldarius origins oriC1 and oriC2 appeared to have simpler binary rules of specification, with Orc1–1 binding solely to oriC1 and Orc1–3 binding to oriC2. None of the Orc1/Cdc6 proteins showed any significant enrichment at S. acidocaldarius oriC3 [39]. More recently, studies in Sulfolobus islandicus have exploited the development of robust genetic tools for this model organism to dissect the rules governing specification of the three replication origins in vivo [18]. It was possible to individually delete the genes for all candidate initiator proteins and retain cell viability. In addition, it was possible to delete all possible pairs of orc1/cdc6 genes. Origin firing in the resultant strains was tested using 2D gel analyses and whole-genome marker frequency analyses. A simple set of rules emerged with Orc1–1 uniquely specifying oriC1 and Orc1–3 specifying oriC2. Orc1–2, in agreement with its proposed role as a negative regulator of replication, was not required for initiation at any of the three origins. The unique origin specification traits of the Orc1–1 and Orc1–3 proteins were further confirmed using ChIP-Seq and locus-specific ChIP qPCR analyses. It is intriguing that the available data support Orc1–3 being the sole initiator for oriC2 in both S. islandicus and S. acidocaldarius, whereas a more complicated situation is seen in the closely related S. solfataricus where oriC2 is bound by both Orc1–1 and Orc1–3. It seems likely that S. solfataricus oriC2 has acquired the Orc1–1-binding ability and it appears that alterations in both Orc1–1 protein sequence and oriC2 DNA sequence have contributed to the evolution of this enhanced complexity [18]. However, genetic studies have not been performed in S. solfataricus to test the functional significance of the Orc1–1/Orc1–3 heterodimer formation.

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Significantly, the genetic studies in S. islandicus revealed that neither individual nor paired deletions of Orc1/Cdc6 protein-coding genes impacted upon firing of oriC3. Indeed, a strain lacking both Orc1–1 and Orc1–3 maintained chromosome replication solely from oriC3, with a lengthening of the S-phase of the cell cycle. Furthermore, none of the Orc1/Cdc6 paralogs bound to oriC3 in vivo, as adjudged by ChIP. However, the gene for WhiP, the Cdt1 homolog, is immediately adjacent to oriC3. Biochemical DNA footprinting and ChIP-Seq studies revealed the WhiP bound specifically to sequence repeats within its own open-reading frame, a situation reminiscent of bacterial RepC plasmids. Accordingly, the whiP gene was disrupted by introduction of a stop codon immediately downstream of the ATG start codon, resulting in loss of expression of the WhiP protein. Consequently, replication initiation was lost at oriC3 but maintained at oriC1 and oriC2. Thus, although WhiP was initially identified as a distant sequence homolog of Cdt1, in Sulfolobus it is functioning as an initiator factor for a single, specific origin rather than serving the global coloader protein role that Cdt1 performs in eukaryotes [18,19]. Taken together, the genetic, ChIP, and biochemical studies revealed that the S. islandicus chromosome is a mosaic of three distinct replicons, each specified by an unique initiator protein that is encoded adjacent to its cognate origin. More recently, this phenomenon of replicon-mosaic chromosomes has been extended to the euryarchaeal phylum. Studies of H. hispanica revealed that the two replication origins on the main chromosome are individually controlled by distinct Orc1/Cdc6 proteins that are encoded adjacent to the origin [35]. Importantly, both Sulfolobus and Haloarcula revealed that the cis-relationship between origin and specifying initiator was not absolutely required, as the initiator could be supplied in trans. The combination of the nonessentiality of the S. islandicus orc1–1 gene and the ability to supply it in trans has allowed parallel in vivo and in vitro dissections of the structure and function of this conserved initiator protein.

5. STRUCTURE OF ARCHAEAL ORC1/CDC6 PROTEINS The hyperthermophilic archaea have been well exploited by structural biologists. The combination of the ease of purification of the inherently thermostable proteins, coupled with their reduced complexity compared to their eukaryotic orthologs, has facilitated the determination of the structures of a number of components of the archaeal replication machinery, including the Orc1/Cdc6 proteins [11,46–48]. Initial studies of homologs

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of the Sulfolobus Orc1–2 protein from the Crenarchaea Pyrobaculum aerophilum and Aeropyrum pernix confirmed the bioinformatic predictions that these proteins possess a AAA + ATPase domain followed by a C-terminal winged helix (wH) domain [46,47]. Intriguingly, both structures revealed that the recombinant proteins purified from E. coli possessed an ADP moiety stably bound in their ATPase active site. Studies with the A. pernix protein from which ADP had been stripped by sequential denaturation, dialysis, and refolding, revealed that it underwent a single turnover ATP hydrolysis event [47,49]. This behavior has subsequently been confirmed for other Orc1/ Cdc6 proteins. Thus, Orc1/Cdc6 exhibits a switch-like behavior between ATP and ADP-bound states, rather than catalyzing the iterative cycles of ATP binding, hydrolysis, and release found in many other AAA+containing proteins. The Orc1/Cdc6s belong to the so-called initiator clade of AAA+ proteins that also includes the bacterial initiator DnaA [50]. Members of the initiator clade have a characteristic embellishment to the core AAA+-fold, in the form of the initiator-specific motif (ISM)—an alpha-helical insertion in the base subdomain of the AAA + module. In DnaA, the ISM is a wedgeshaped pair of essentially antiparallel alpha-helices that facilitates helical polymerization of the DnaA monomers into a filament-like structure [51]. In contrast, the Orc1/Cdc6 proteins have a pair of parallel alpha-helices connected by a elongated linker, resulting in a block-like appendage. Modeling of the Orc1/Cdc6 proteins onto the structure of a DnaA filament results in a steric clash. Indeed, as discussed later, the Orc1/Cdc6 proteins appear to function as monomers at the replication origin. In 2007, the Berger and Wigley laboratories published landmark studies in which they determined the crystal structures of Orc1/Cdc6 proteins bound to replication origin DNA [11,48]. The Wigley lab described the structure of A. pernix Orc1–1 bound to the consensus ORB (origin recognition box) element from the organism’s oriC1 [48]. Berger and colleagues described the structure of a heterodimer of the S. solfataricus Orc1–1 and Orc1–3 proteins bound to their adjacent sites at oriC2 [11]. As described earlier, all three proteins contained ADP in their active site. One of the most striking features of the structures was that each Orc1/Cdc6 protein made bidentate contacts with DNA. Previous work had implicated the wH domain in origin recognition and this was confirmed in the structures. However, in addition, the AAA+ domain contacted DNA via the ISM motif. The binding sites for Orc1–1 at oriC1 and at S. solfataricus oriC2 are related in sequence. The sites at oriC1 conform to a consensus sequence, termed

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ORB, that can be recognized in a broad phylogenetic diversity of the archaea [9]. Indeed, Sulfolobus Orc1–1 can bind specifically to the ORB elements from origins in the chromosomes of Pyrococcus and Halobacterium [9]. ORB elements contain a dyad symmetric element that is flanked on one side only by a string of 3–5 guanine nucleotides. The Orc1–1 binding sites at oriC2 in S. solfataricus possess a shortened version of the ORB element (termed mini-ORB) that possesses just the dyad symmetric element and lacks the G-string [9]. Based on the presence of the dyad symmetric element within an ORB, it might have been anticipated that Orc1–1 would bind as a symmetric dimer. However, the Aeropyrum Orc1–1
ORB structure, and subsequent isothermal calorimetry experiments, revealed that a single monomer of Orc1–1 contacted the ORB element [48] (Fig. 2). The G-string distal element of the symmetric dyad is contacted by the extended wing of the wH and the G-string proximal element is recognized by the recognition helix of the wH domain. The G-string itself was contacted by the ISM in the Orc1–1 AAA + domain. Considerable deformation of DNA is seen in the structure with a global bend of 35°C. In addition, there is considerable local deformation to the helical geometry. Over the ORB element, DNA is untwisted by 60 degree and the minor groove is significantly broadened throughout. This protein-induced deformation accounts for why only a single monomer of Orc1–1 binds—the perturbation of the DNA structure inhibits recognition of the dyad by a second symmetrically apposed

ADP

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TCTCCACAGGAAACGGAGGGGT Dyad element

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Fig. 2 Structure of the Aeropyrum pernix Orc1–1 protein bound to ORB element DNA (PDB 2V1U). The ISM and wH DNA-binding units are highlighted in red (dark gray in the print version). ADP is in dark blue (black in the print version) and the catalytic magnesium ion shown as a black sphere. The dyad element of the ORB is shown in yellow (light gray in the print version) and the G-string in cyan (light gray in the print version).

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Orc1–1 monomer [48]. The interaction of the ISM with the G-string helps explain the observed polarity of binding. Significantly, the arrangement of ORB elements at a number of origins, although varying in number in a species-specific manner, shares some global architectural features. Most notably, at least two ORB elements are found in inverted repeat orientation flanking a highly AT-rich region. The inverted ORB elements are situated such that the wH domains of the DNA-bound Orc1–1 proteins face one another. Replication initiation point (RIP) mapping, a high-resolution technique that reveals the transition point between leading and lagging strand replication, in S. solfataricus and A. pernix revealed that leading strand synthesis begins within or just adjacent to the intervening AT-rich region, suggesting it may act as a duplex unwinding element (DUE) [9,52]. Similarly, RIP mapping in H. volcanii and P. abyssi revealed candidate DUEs and initiation sites in the proximity of inverted-orientation ORB elements [33,53] (Fig. 3). This organization is suggestive of a mechanism for MCM

Fig. 3 Organization of oriC1 from a range of archaeal species. Arrows indicate the position of the start of leading strand synthesis mapped by RIP assays. The candidate duplex unwinding elements are shown as gray boxes and ORB elements are yellow (light gray in the print version) and blue (dark gray in the print version) arrows. The polarity is indicated by color and direction of the arrow. The color convention is as in Fig. 2, with the dyad in yellow (white in the print version) and G-string in blue (gray in the print version). The full complement of ORB elements are shown for all organisms with the exception of Haloferax which has an additional five elements extending downstream (from the DUW distal face) of ORB3.

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loading whereby two single hexamers of MCM are individually loaded onto the AT-rich DUE region by Orc1–1 proteins bound to the flanking ORB elements.

6. ROLE OF ATP IN MCM LOADING The Orc1/Cdc6 proteins are AAA + proteins and have the unusual property of catalyzing a single turnover ATP hydrolysis event, resulting in ADP bound stably in the active site. This manifestation of end-product inhibition serves to confer a switch-like behavior on these proteins. Studies in S. acidocaldarius have demonstrated that expression of the orc1–1 and orc1–3 genes are tightly regulated with mRNA levels peaking prior to the initiation of S-phase [18]. Interestingly, the timing of firing of oriC1, specified by Orc1–1, and oriC2, specified by Orc1–3, are offset such that oriC2 fires later than oriC1 [39]. The expression profiles of the cognate initiators parallel this temporal separation of firing, with orc1–1 mRNA levels peaking prior to those of orc1–3 [18]. Nascent Orc1/Cdc6 proteins presumably bind ATP upon completion of protein folding. As in bacteria, transcription and translation are coupled in archaea [54]. Therefore, the cis-relationship of the initiator protein-coding gene with its origin of replication will ensure rapid association of the ATP-bound initiator with the replication origins. The origin-associated ATP-bound form of the protein will then have a defined period of time prior to ATP hydrolysis. Notably, the Orc1/Cdc6 proteins remain origin associated for the duration of the cell cycle, yet replication only initiates once per origin per cell cycle [39]. We speculate that the shortlived ATP-bound form of the protein confers a permissive window in the cell cycle by promoting MCM loading. Subsequent hydrolysis of ATP by the initiator protein would lead to its inactivation and thus prevent further rounds of replication from occurring. The origin incumbent, ADP-bound form of the protein could either be degraded at the end of the cell cycle or conceivably ADP in the active site could be exchanged for ATP. However, given the wave of de novo synthesis of Orc1/Cdc6s at the beginning of the subsequent cell cycle, we favor the former hypothesis [18]. Data to support this binary switch model of Orc1/Cdc6 function has been obtained in Sulfolobus, taking advantage of the nonessentiality of the orc1–1 gene [18]. A so-called Walker B mutant, E147A, was expressed in Δorc1–1 S. islandicus cells. In this form of the protein, the catalytic glutamate of the Walker B box has been changed to alanine, resulting in a protein that binds but fails to hydrolyze ATP. Strikingly, expression of the Walker

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B mutant protein restored firing at the oriC1 origin in an orc1–1 null background. Replication initiation at oriC1 appeared to be enhanced as the initiation arc detected in 2D gels was more pronounced than in wild-type cells. Furthermore, fluorescence microscopy and flow cytometry revealed the presence of a significant population of cells of enlarged size and with elevated DNA content, indicative of overreplication. Parallel biochemical studies examining MCM recruitment to immobilized DNA on paramagnetic beads revealed that preincubation of the origin-specific beads with the Walker B mutant, the constitutively ATP-bound form of the protein, resulted in recruitment of the MCM helicase in vitro. In contrast, incubation of beads with the wild-type ADP-bound form of Orc1–1 failed to recruit any MCM [18]. Thus, the combined in vivo and in vitro data support the model whereby Orc1–1 is active in its ATP-bound state and hydrolysis of ATP to ADP inactivates the protein with respect to its ability to recruit MCM. Whether ATP hydrolysis simply acts to switch the Orc1–1 protein off, of whether it actively helps release MCM remains unknown. However, it is clear that the Walker B mutant version of the protein still supports replication in vivo, so hydrolysis per se is clearly not essential for Orc1–1 function in the cell.

7. THE MECHANISM OF MCM RECRUITMENT BY ORC1–1 Electron microscopy and X-ray crystallography studies of MCM, mostly of Methanothermobacter thermautotrophicus MCM, have demonstrated that it forms double hexamers via interactions between the N-terminal domains of the protein [55,56]. Overall, a single MCM hexamer has a three-tier structure with one tier formed by the N-terminal domains, followed by an AAA + module, and finally wH domains at the C-terminus of the protein. Crystallographic studies have revealed the structures of the N-terminal and AAA+ domains, both in a monomeric single subunit and in a hexameric assembly [57–59]. The wH domains have not been visualized in the context of the full-length protein, presumably due to their flexibility. However, the structures of isolated wH domains have been solved for both M. thermautotrophicus and S. solfataricus MCM by NMR [60] (Fig. 4). The model proposed earlier for MCM recruitment by Orc1–1 protomers bound to inverted ORB elements would lead to a head-to-head double hexamer structure of MCM on the origin (Fig. 5). As double hexamer formation occurs via N-terminal–N-terminal tier interactions, the model

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Fig. 4 Structure of archaeal MCM. The structure shown is the Pyrococcus–Sulfolobus hybrid created by Enemark and colleagues (PDB 4R7Y, Ref. [58]). Protomers within the hexamer are alternately colored red (dark gray in the print version) and cyan (light gray in the print version). The structure lacks the C-terminal wH domain of each monomer. However, the structure of that domain has been solved by NMR and a single wH domain from S. solfataricus is shown to the same scale in the right-hand panel [60]. wH

AAA+

NTD

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Orc 1–1 AAA+ wH

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MCM hexamer

MCM hexamer

Fig. 5 Cartoon model for an origin-associated double hexamer of MCM, loaded as two independent hexamers by inversely oriented Orc1–1 proteins.

predicts that the C-terminal wH domains of MCM would be adjacent to the wH-face of DNA-bound Orc1–1. Initial in vitro MCM recruitment assays employed purified recombinant Orc1–1 protein but used cell extract as the source of MCM. It was therefore unclear whether Orc1–1 was, in addition to acting as the origin-specifying initiator protein, also playing a role as a helicase loader. Very recently, however, a fully recombinant MCM recruitment system using purified recombinant Orc1–1 and MCM has been described [61]. The system makes use of the constitutively ATP-bound Walker B mutant form of Orc1–1 to effect origin-dependent recruitment

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of MCM. It appears, therefore, that the archaeal Orc1–1 is a bifunctional initiator and helicase recruiter. This is in agreement with the proposal that the archaeal Orc1/Cdc6 proteins were the progenitors of both the eukaryal Orc1 component of the initiator ORC and the helicase coloader, Cdc6. However, this observation highlights a distinction from the bacterial and eukaryal replication machineries, where dedicated helicase loaders act as intermediaries between initiator and helicase and makes the archaeal system the simplest cellular replication system characterized, at least in terms of the number of different proteins required. Notably, studies with DNA substrates containing mutated origin sequences revealed that ORB2 and ORB3 were necessary and sufficient for recruitment of MCM to a high-salt stable complex on DNA—presumably reflective of loading of the double hexamer and thus supportive of the model shown in Fig. 5 [61]. Studies using mutated derivatives of MCM revealed that there was no requirement for MCM to bind or hydrolyze ATP for the loading process. However, deletion of the C-terminal wH domain of the MCM protein abrogated recruitment. Thus, it seems likely that the wH domain of MCM is serving as a protein–protein interaction site [61]. Notably, the C-terminal wH domain of eukaryotic MCM3 is required for recruitment of MCM(2–7) by the ORC
Cdc6 complex [27], suggesting an evolutionarily conserved mode of interaction. Examination of the structure of the wH domain of MCM reveals its closest structural homolog in the PDB database to be a wH domain from human RPA32 [60–62]. RPA32 is a component of the single-stranded DNA-binding protein RPA. The wH domain in RPA32 has no role in DNA binding, rather, it is a protein–protein interaction module that mediates RPA’s interaction with a number of distinct factors [62–64]. Strikingly, residues directly involved in protein–protein interactions in RPA32 are conserved in Sulfolobus MCM and mutation of these residues impacts upon the ability of MCM to be recruited to oriC1 by Orc1–1 [61]. The region of Orc1–1 responsible for interaction with MCM was similarly identified based on homology with the known interaction partners of RPA32. More specifically, alignment of the RPA32-binding peptide from human UNG2 (a uracil N-glycosylase) with Orc1/Cdc6 proteins identified a conserved region in the initiator proteins [62]. The motif, which maps to a surface-exposed α-helix on the lid domain of the Orc1/Cdc6 proteins’ AAA + domain, is conserved in Orc1–1 and Orc1–3 but, notably, is absent from Orc1–2 proteins. Thus, this candidate MCM recruitment motif

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(MRM) is a signature feature of replication-promoting Orc1/Cdc6 proteins and is broadly conserved across the archaeal domain of life. As can be seen in Fig. 6, the MRM is located on the wH-proximal face of the AAA+ domain and would face toward the candidate DUE when Orc1–1 is bound at the inverted ORB elements flanking the DUE. The functional relevance of the MRM was confirmed by mutation of the conserved residues. The resultant proteins bound origin DNA in vivo and in vitro, yet failed to recruit MCM in vitro and could not support initiation in vivo. A clue to the ATP-dependence of MCM recruitment comes from the location of the MRM in the AAA + domain. The motif is four residues removed from the R250 (in S. islandicus) Sensor 2 active site residue. This arginine is a conserved feature of many AAA+ proteins and acts to coordinate the γ-phosphate when ATP is present. Coordination of the Sensor 2 results in relative repositioning of the lid and base domains of the AAA+ module [50,65,66]. R250 thus identifies the nature of the nucleotide in the active site and modulates the structure of the AAA+ module accordingly. Intriguingly, like a Walker B mutant of Orc1–1, the Sensor 2 mutant form of the protein (R250A) binds but fails to hydrolyze ATP. However, in contrast to the active Walker B mutant, Sensor 2 mutants of Orc1–1 fail to recruit MCM. Similar behaviors of Walker B and Sensor 2 mutants have been observed in studies of budding yeast Cdc6 [67–70]. Importantly, a Sensor 2/Walker B double mutant Orc1–1 binds ATP and DNA normally but

Fig. 6 Position of the MCM recruitment motif (MRM) in A. pernix Orc1–1. The MRM is highlighted in purple (dark gray in the print version) and the adjacent Sensor 2 residue (R250 in Sulfolobus) is shown in blue (black in the print version). Residues A240 and R244 of the MRM are shown in purple (dark gray in the print version) stick form, and the atoms of ADP in this structure is in green (dark gray in the print version) sphere representation. DNA coloring is as in Fig. 2 with the dyad in yellow (light gray in the print version) and G-string in cyan (light gray in the print version).

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fails to recruit MCM in vitro or support origin firing in vivo. Taken together, these data support the proposal that the Sensor 2 residue, as its name implies, senses the presence of ATP in the active site and, as in other AAA + proteins, leads to a conformational modulation of the AAA+ domain. The consequence of this conformational switch would be to facilitate the interaction between the adjacent MRM and the wH domain of the incoming MCM. Upon hydrolysis of ATP to ADP, the AAA+ would rearrange into an inactive configuration. The current crystal structures of Orc1–1 are in the ADP-bound state and modeling of the MRM–MCM wH interaction with the ADP-bound Orc1–1 indicates a modest steric clash between the two proteins. A relatively subtle repositioning of the MRM could be sufficient to alleviate this clash and permit the interaction. Clearly, determination of the structure of the ATP-bound form of Orc1–1 is a major goal for future work. Finally, how does Orc1–1 mediate MCM loading? Numerous models have been proposed for this process, ranging from ring assembly to dynamic ring-opening models. MCM is a highly pleomorphic complex and a range of structures have been described, from ring-shaped hexamers, double hexamers and heptamers, through to filaments [55]. Notably, one electron microscopy study investigated the effect of treatment of M. thermautotrophicus MCM at its physiological growth temperature of 60°C. Strikingly, the consequence of the heat treatment was that greater than 50% of the particles observed were in an open-ring conformation [71]. Biochemical loading assays in which S. islandicus MCM was preheated at 75°C resulted in a significant increase in the yield of high-salt stable MCM complexes recruited to the oriC1 replication origin. Additionally, gel filtration chromatography was used to separate open- and closed-ring forms of the MCM hexamer. Use of these populations in recruitment assays revealed the open-ring form was far more efficiently recruited to origins in vitro than the closed-ring form [55]. These data indicate that the open-ring form of MCM is thermodynamically favored at the physiological growth temperature of archaea and that this open-ring form of MCM is preferentially recruited to and loaded on replication origins.

8. CONCLUDING REMARKS A considerable body of structural data is now available for components of the archaeal DNA replication machinery. However, our understanding of the nature of the dynamic transactions between replication factors on the

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DNA template remains less well resolved. With the development of in vitro systems and robust genetic tools for parallel in vivo analyses, it is anticipated that rapid progress will be made in understanding the molecular manipulations that underpin the transitions from origin-bound MCM to the ultimate assembly of the replication fork. Still less is known about regulation of these pivotal events in the archaeal cell cycle. Certainly, transcriptional regulation of the initiator proteins genes appears to be a component of the regulatory network; however, the identity of the transcriptional regulators and how they in turn are regulated remain completely unknown at this time. While the binary switch model that we have proposed applies to oriC1 and oriC2 in Sulfolobus, we do not know how oriC3, the WhiP-dependent and Orc1/ Cdc6-independent origin, is regulated and limited to firing once per cell cycle. One possibility is that WhiP could be regulated at the level of protein stability. It is also conceivable that, in addition to initiator-level regulation, regulation of initiation could also be effected post-MCM recruitment. This notion appeals, as a regulatory system that acts upon origin-bound MCM would target the first common component to replication initiation at all three replication origins and could supply an over-arching coordinate control system. It may be significant that archaea possesses a number of serine/ threonine kinases related to those found in eukaryotes [72]. Additionally, ubiquitin-related modifier proteins have been implicated in protein degradation in Sulfolobus [73] and other archaea [74]. It will be of considerable interest to determine if regulation of replication by posttranslational modification is a phenomenon shared by archaea and eukaryotes.

ACKNOWLEDGMENTS We are indebted to Richard McCulloch and Thorsten Allers for helpful discussions. We also thank the College of Arts and Sciences, Indiana University for funding.

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CHAPTER SIX

The Eukaryotic Replication Machine D. Zhang*, M. O'Donnell*,†,1 *The Rockefeller University, New York, NY, United States † Howard Hughes Medical Institute, The Rockefeller University, New York, NY, United States 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. The CMG Helicase 3. The Polymerase Alpha-Primase 4. The Leading and Lagging Strand DNA Polymerases Epsilon and Delta 5. The PCNA Clamp and RFC Clamp Loader 6. The Eukaryotic Replisome Structure and Function 7. Comparison of Bacterial and Eukaryotic Replisomes 8. Future Perspectives Acknowledgments References

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Abstract The cellular replicating machine, or “replisome,” is composed of numerous different proteins. The core replication proteins in all cell types include a helicase, primase, DNA polymerases, sliding clamp, clamp loader, and single-strand binding (SSB) protein. The core eukaryotic replisome proteins evolved independently from those of bacteria and thus have distinct architectures and mechanisms of action. The core replisome proteins of the eukaryote include: an 11-subunit CMG helicase, DNA polymerase alphaprimase, leading strand DNA polymerase epsilon, lagging strand DNA polymerase delta, PCNA clamp, RFC clamp loader, and the RPA SSB protein. There are numerous other proteins that travel with eukaryotic replication forks, some of which are known to be involved in checkpoint regulation or nucleosome handling, but most have unknown functions and no bacterial analogue. Recent studies have revealed many structural and functional insights into replisome action. Also, the first structure of a replisome from any cell type has been elucidated for a eukaryote, consisting of 20 distinct proteins, with quite unexpected results. This review summarizes the current state of knowledge of the eukaryotic core replisome proteins, their structure, individual functions, and how they are organized at the replication fork as a machine.

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2016 Elsevier Inc. All rights reserved.

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1. INTRODUCTION DNA replication is a once-in-a-lifetime event. It must go from start to finish without failure, unlike the other nucleic acid-based informational pathways of transcription and translation that can afford to sometimes fail and then start over. Furthermore, eukaryotic genomes are very large, over 4 billion base pairs in the human. Replication of the genome must not only finish the job, but must do so with exquisitely high fidelity, leaving no mistakes, or exceedingly few, to preserve the species. The replication process is initiated at origins by highly regulated proteins and cell cycle kinases [1–3]. Once started, replication of both strands of duplex DNA is carried out by many different proteins and enzymes that function in a highly choreographed fashion [4,5]. Many of these proteins bind one another relatively tightly, forming a replisome complex that acts somewhat like the moving gears of a sewing machine. Some proteins that are central to the DNA replication process act in a dynamic fashion, coming in and out of the replisome at a moving replication fork. This review will focus on the major set of proteins that function to propel the eukaryotic replication fork. The term “core replisome” will refer to those proteins that are necessary to propagate the replication fork, regardless of their affinity to the protein complex that moves with the fork. Numerous other proteins act after the fork has passed, to fix mismatches and to repair and ligate Okazaki fragments. These repair proteins will not be within the focus of this review. The eukaryotic replisome is much more complicated than bacterial and viral replisomes, as one might expect from the greater genome size and complexity of a eukaryotic cell. The eukaryotic replisome contains all the core replisome proteins of the bacterial replisome, plus many others. The core replisome proteins are the helicase, primase, DNA polymerases, sliding clamp, clamp loader, and single-strand binding (SSB) protein. The eukaryotic core components are multisubunit complexes, each with more subunits than their bacterial counterparts. There are many other factors that move with the eukaryotic replisome and have no analogous bacterial protein. These additional factors, outside the core machinery, appear to be involved in two major processes. One is regulation of the fork, as some of the components of the eukaryotic replisome are involved in checkpoint and repair pathways. Another is the need for the eukaryotic replisome to handle nucleosomes, which bacteria do not have. Nucleosomes are at the heart of animal development.

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This review will discuss each of the eukaryotic replisome core components and how they fit and function together. We conclude with a brief comparison of the eukaryotic replisome to the bacterial replisome. There are a few additional facts to mention in this section before starting off on these subjects. One is that eukaryotes have linear chromosomes and use numerous origins, unlike most bacteria and archaea that typically use one or a few origins within a circular genome [6]. Thus, eukaryotic origins are often spaced only tens of kilobases apart, and eukaryotic replication forks do not have to cover the megabase distances that bacterial forks must cover [7]. This may be why the rate of eukaryotic replication forks average about 10–50 nucleotides compared to the 10- to 20-fold faster rate of bacterial and phage replisomes [7–9]. The use of linear chromosomes requires that eukaryotes have telomere ends, which are replicated by the telomerase ribozyme. This review will not cover telomere biology, and the reader is referred to review on this subject [10–12]. Genome sequencing of cells from the three domains of life, bacteria, archaea, and eukaryotes, reveal that most of the core replisome components evolved twice, independently [13,14]. Thus, the bacterial core replisome enzymes do not share a common ancestor with the analogous components in eukaryotes and archaea [13,14], while the archaea and eukaryotic core replisome machinery share a common ancestor [15,16]. An exception to this are the clamps and clamp loaders, which are homologous in all three domains of life [14,17,18]. This is quite unlike the processes of transcription and translation, the two other major nucleic acid informational pathways, both of which have homologous core components and a universal genetic code in bacteria, archaea, and eukaryotes. Why the core replisome machinery evolved independently in bacteria and archaea/eukaryotes is unknown. One possibility is that the last universal common ancestor (LUCA) cell replicated its nucleic acid genome in a much more streamlined and simple fashion compared to modern day cells, such as seen in some phage and viruses. For example, many phage and viruses do not replicate both strands of double-strand (ds) DNA at the same time. If the two strands of dsDNA are not made simultaneously (ie, one strand is completed before the other strand is started), replication simplifies tremendously. For example, priming is only needed once or twice and can be substituted by a nick generated by an endonuclease for a covalently closed genome, a tRNA primer as in retroviruses, terminal protein priming or by RNA polymerase [6]. Instead of using a helicase to unwind DNA, the DNA polymerase may be capable of strand displacement (eg, phi29 bacteriophage). Or if RNA is the genome, it may be digested during replication as in retrovirus replication [6]. Interestingly, the

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helicase and primase are the most different among the core replisome factors of bacterial and eukaryotic replisomes. The ATPase motor of bacterial replicative helicases (eg, E. coli DnaB) is based on a RecA fold, while the six ATPase subunits of the eukaryotic Mcm helicase are based on the AAA+ fold [19,20]. The primase of bacteria is based on a Toprim fold, that has an evolutionary relationship to topoisomerases [21–23], while the primase of eukaryotes shares homology to the X-family of DNA polymerases [24]. There are six classes of DNA polymerases that are unrelated in sequence [25,26]. The eukaryotic replicative DNA polymerases are in the B-family, while the only domain of life that contains C-family DNA polymerases are bacteria, which use them for genome replication [27]. Only the sliding clamp processivity factor and the clamp loader of bacteria and eukaryotes share a common ancestor, and one can question why this may be so. The structure and function of the circular clamp and clamp loader were first discovered in the context of E. coli replication [28,29], but later studies have shown that the clamp and clamp loader are used by numerous different proteins in various DNA repair, checkpoint, and cell cycle regulatory pathways in all cell types [30,31]. Thus, the clamp/clamp loader may have evolved for a nonreplicative use in LUCA and then was recruited for replication later in cellular evolution after the split of bacteria from archaea/eukaryotes. The core components of the eukaryotic replisome are listed in Table 1 for Saccharomyces cerevisiae (budding yeast). Other important systems for replisome studies include: Schizosaccharomyces pombe, Drosophila melanogaster, Xenopus laevis, and human. The composition of the full eukaryotic replisome is still an active area of investigation, and new factors that move with the replisome continue to be found. Despite the many proteins that move with eukaryotic replisomes, biochemical reconstitution studies confirm that an active moving replisome that replicates both leading and lagging strands assembles from the components listed in Table 1 [32,33]. This does not lessen the importance of the many additional factors that move with replisomes, because the ability to regulate the replisome, to adhere the sister chromatids, and to handle nucleosomes are all central to genomic integrity. On that note, we begin an exploration of the individual components of the replisome.

2. THE CMG HELICASE The eukaryotic replicative helicase is an 11-subunit complex referred to as CMG [34,35]. CMG is an acronym of the three components required for helicase activity. The “C” refers to Cdc45, a protein that shares

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Table 1 Core Replication Proteins of the Saccharomyces cerevisiae Replisome Subunit (MW in kDa) Complex (MW in Da) Complex Function

Mcm2 98,779

Mcm2–7 605,627

CMG 786,038 Helicase

Mcm3 107,516 Mcm4 105,002 Mcm5 86,410 Mcm6 112,977 Mcm7 94,943 Sld5 33,947

GINS 105,163

Psf1 24,203 Psf2 25,061 Psf3 21,952 Cdc45 74,248 Pol1 166,808

Pol alpha 355,534

Primase

Pol epsilon 378,670

Leading polymerase

Pol delta 220,222

Lagging polymerase

RPA 114,099

Single-strand binding protein

Pol12 78,774 Pri1 47,690 Pri2 62,262 Pol2 255,669 Dpb2 78,229 Dpb3 22,665 Dpb4 21,997 Pol 3 124,618 Pol31 55,295 Pol32 40,309 Rfa1 79,347 Rfa2 29,936 Rfa3 13, 816

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homology to the RecJ family of nucleases but is not known to contain nuclease activity [36,37]. The “M” refers to the six Mcm motor subunits that form the ring shaped heterohexamer Mcm2–7 complex [38,39]. The “G” stands for the heterotetramer GINS complex, which is an acronym formed from the first letter of the Japanese spelling of the numbers 5,1,2,3 (go-ichi-ni-san) and their individual subunit names are Sld5, Psf1, Psf2, and Psf3 [40]. Together, the 11-subunit CMG assembly is an active helicase [34,35]. Archaea use a single Mcm subunit that hexamerizes to form an active Mcm helicase [41,42], although protein homologs of Cdc45 and GINS are present in archaea [36]. The Mcm2–7 of budding yeast displays some helicase activity [38], but human and Drosophila Mcm2–7 display no detectable helicase activity without the Cdc45, GINS accessory factors [34,43]. CMG is formed during activation of an origin, a fascinating multistep reaction (reviewed in Ref. [1–3,5]). Briefly, origin activation is separated into two distinct phases of the cell cycle, G1 and S. In G1, an origin is “licensed” in a process that requires the six-subunit origin recognition complex, along with Cdc6, Cdt1, and the Mcm2–7 complex. The licensing process results in two Mcm2–7 hexamers that encircle dsDNA and are positioned head-to-head (ie, the two hexamers bind via their N-terminal regions). In proceeding to S phase, several origin activation factors work together, along with two cell cycle kinases, to form the CMG complex in which Cdc45 and GINS become tightly associated with Mcm2–7, and each CMG complex encircles only one strand of DNA. The CMG helicases proceed to unwind DNA, forming the ssDNA needed for priming and replisome assembly. The detailed mechanism of the origin activation process is still an active area of research, and the complete reconstitution of this process from pure proteins has recently been achieved in the S. cerevisiae system [44]. It is interesting to note that a mixture of Mcm2–7, Cdc45, and GINS does not result in formation of CMG [34,35]. Thus, CMG requires some or all of the origin activation factors for its formation. Presumably some type of energetic barrier must be surpassed to put CMG together from its component parts. Helicases have been subdivided into superfamilies (SF), monomeric and dimeric helicases in SFI and SFII, hexameric AAA+ helicases of SF3 and SF6 which track 30 -50 along ssDNA, and RecA-based helicases of SF4 and SF5 that track 50 -30 along ssDNA [20]. The eukaryotic Mcm2–7 and archaeal Mcm are SF6 helicases, the eukaryotic SV40 (T-antigen (T-Ag)) and papilloma virus (E1 protein) are SF3 helicases, and the bacterial cellular and

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animal mitochondrial helicases, and phage T4 and T7 helicases are in the SF4 family. All the replicative helicases function as circular homohexamers, except CMG which contains the circular Mcm2–7 heterohexamer motor plus five accessory factors. A feature that is shared by the motor subunits of all replicative helicases is their construction from two major domains, the N- and C-terminal domains (NTDs and CTDs) (Fig. 1A). The CTDs contain the ATP sites, and thus are the motor domains. The ATP sites are located at subunit interfaces which may facilitate intersubunit communication driven by ATP hydrolysis. The NTDs of the different helicases are quite distinct but appear to form a scaffold on which the motor domains are attached. The NTDs of the Mcm subunits contain an OB fold (oligonucleotide/oligosaccharide-binding fold), which often bind ssDNA, and are demonstrated to do so in the NTD hexameric ring of an archaeal Mcm [45]. The ring-shaped helicases of all cell types are currently thought to act by the steric exclusion mechanism of DNA unwinding (Fig. 1B). An alternative mechanism is the side channel extrusion model. The distinguishing characteristic of these models is whether one or both strands of DNA enter the central pore [46–51]. In the steric exclusion mechanism only one DNA strand enters the helicase, while the other strand is excluded, and as the helicase tracks along the ssDNA it acts as a wedge to split the parental duplex. The side channel extrusion mechanism requires both strands to enter the central pore, and DNA is split inside the helicase, followed by one strand

Fig. 1 Replicative helicases are hexamers that have the appearance of two stacked rings. (A) All replicative helicases are composed of a hexameric motor. The individual subunits consist of two large globular domains, a N-terminal domain (NTD) and C-terminal domain (CTD). Thus, helicase hexamers have the appearance of two rings stacked on one another. (B) Steric exclusion mechanism of unwinding. In the steric exclusion mechanism, one strand is excluded, while the tracking strand threads through the central pore of the helicase. The eukaryotic helicases encircle the leading strand, as illustrated.

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being directed out a side channel between the CTD and NTD. Interestingly, the bacterial helicases and CMG have been shown capable of tracking over either ssDNA or dsDNA, and thus they could possibly function by either mechanism [43,48,49,52,53]. But the available experimental evidence is that all replicative helicases, from bacteria to eukaryotes, function by the steric exclusion mechanism [47,52–56]. It is not entirely clear whether nucleotide hydrolysis is required to actively melt the duplex DNA, or whether the duplex frays by thermal melting, and that helicase tracking along ssDNA prevents reannealing. These modes are referred to as “active” and “passive” unwinding, respectively. The rate of thermal fraying has been measured to be rapid and possibly sufficient to account for the rate of the fast bacterial helicases [57–59]. Current estimates of ATP per base pair (bp) unwound are one ATP every 1–2 bp, and given the 12 kcal/ATP compared to 3.6 kcal for 2 bp of dsDNA, the energetics are compatible with either case [60–62]. Considering the 10- to 20-fold slower rate of eukaryotic fork movement, active unwinding would not seem necessary, but displacement of nucleosomes may become a major energetic barrier. The main evidence for CMG acting by the steric exclusion mechanism derives from studies in the Xenopus system, using DNA substrates that contain a streptavidin–biotin block on either the leading or lagging strand [54]. In the steric exclusion mechanism, a bulky group attached to the excluded strand that lies on the outside of the central channel should not slow the helicase, but a bulky block on the leading strand should restrict DNA entry into the central channel and thus prevent unwinding. This is, in fact, the observed result for E. coli DnaB [48]. Xenopus egg extracts replicate exogenous DNA in a manner that depends on all the known components of the replisome, and therefore this system is believed to reflect the behavior of a complete replisome. In the Xenopus study, the “block” was formed by two adjacent biotinylated nucleotides, each bound to streptavidin. When presented with a leading strand block, the replisome is halted, as expected of a helicase that tracks along this strand. But the lagging strand block did not stop the replisome, indicating that the lagging strand stays outside the helicase and supporting a steric exclusion mechanism [54]. Unlike studies of E. coli DnaB, the lagging strand block resulted in replisome pausing in the Xenopus study. The nature of this pause is not understood at the current time but suggests that the lagging strand may have extensive interaction with the outside surface of CMG. Indeed, studies with archaeal Mcms, and crosslinking studies of Drosophila CMG with DNA, support the proposal that the lagging strand wraps around the outside of the Mcm ring [63,64].

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Unlike other hexameric helicases, the Mcm2–7 has an opening between two of its subunits, sometimes referred to as the Mcm2/5 gate. This is observed by EM 3D reconstruction studies of Mcm2–7 from two different organisms and shown in Fig. 2 for D. melanogaster Mcm2–7 [65,66]. A gap between the Mcm2/5 subunits was predicted from earlier biochemical studies and is required during the initiation process [39,67]. At an origin, Mcm2–7 must first be loaded onto dsDNA to form a double hexamer. Although EM studies of the double hexamer do not visualize a gap between subunits, biochemical studies demonstrate that the Mcm2/5 interface must

Fig. 2 EM structures of the Mcm2–7 helicase motors. (A) CryoEM atomic structure of Saccharomyces cerevisiae Mcm2–7 double hexamer. Left, surface view; each subunit is a different color (gray shades in the print version). Right, cut away illustration of the central channel of the double hexamer, with an inset that illustrates a spiral arrangement of loops that may bind dsDNA. (B) EM structure of Mcm2–7 from Drosophila melanogaster adopts two conformations; one (right) has a prominent gap between the AAA + CTD domains of Mcm2 and Mcm5. (C) EM structure of Drosophila melanogaster CMG with and without ADP-BeF3. Panel (A) adapted by permission from Macmillan Publishers Ltd. N. Li, Y. Zhai, Y. Zhang, W. Li, M. Yang, J. Lei, B.K. Tye, N. Gao, Structure of the eukaryotic MCM complex at 3.8 A, Nature 524 (2015) 186–191. Panels (B) and (C) are adapted by permission from Macmillan Publishers Ltd. A. Costa, I. Ilves, N. Tamberg, T. Petojevic, E. Nogales, M.R. Botchan, J.M. Berger, The structural basis for MCM2-7 helicase activation by GINS and Cdc45, Nat. Struct. Mol. Biol. 18 (2011) 471–477.

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be allowed to open during Mcm2–7 double hexamer assembly at an origin [68]. During formation and activation of CMG, an opening between Mcm subunits is needed a second time to exclude one of the DNA strands from the central channel for the steric exclusion mechanism of DNA unwinding. Although it is not known which subunit interface opens for the transition to encircling ssDNA, it could be the Mcm2/5 gate. In a tour de force, the >1.2 MDa yeast Mcm2–7 double hexamer structure has been solved at atomic resolution by cryoEM [69] (Fig. 2). The Mcm2–7 hexamers are closed in the double hexamer, but interestingly the two hexamers are rotated relative to one another such that if the Mcm2/5 subunits were to open, the two gaps would not be aligned, and the double hexamer would remain topologically linked to DNA. The exact mechanism by which dsDNA at the origin is melted is not yet known. The double hexamer structure shows the two hexamers are askew, indicating that the DNA between them would be kinked and distorted. It is proposed that ATP may fuel a twisting motion that could lead to DNA unwinding [69]. The atomic model of the Mcm2–7 double hexamer reveals a wide cen˚ in diameter, consistent with ability of Mcm2–7 to tral channel, 30–40 A encircle dsDNA. Interestingly, two constriction points were observed in each hexamer, one in the NTD and one in the CTD. The constriction point in the CTD motor domains corresponds to loops that form a right hand spiral and are predicted to track the grooves of DNA. These loops may correspond to the loops in E1 helicase that appear to move with nucleotide hydrolysis and are proposed to “escort” ssDNA through the hexamer during translocation, one nucleotide per ATP hydrolyzed [70]. The EM structure of Drosophila CMG, and more recently the yeast CMG, has been determined (Fig. 2) [65,71]. The accessory factors bind the outside surface of the Mcm2–7 ring and span the Mcm2/5 interface, effectively trapping DNA inside even if the Mcm2/5 interface were to open. Furthermore, the accessory factors form a second, smaller channel lined by the outside surfaces of Mcm2/5/3 and the GINS and Cdc45 subunits. It was originally suggested that this second channel may hold the lagging strand [65], and this could possibly explain the observed replisome pausing by lagging strand-specific blocks in the Xenopus system [54]. Association of several other proteins with CMG has been identified in pull-down experiments of nuclease-treated cell extracts using epitopes directed against the GINS complex [72]. The resulting “replisome progression complex,” or RPC, contained, in addition to CMG, the following proteins: Ctf4, Pol alpha, Mcm10, Topo I, the checkpoint response factors

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Mrc1, Tof1, Csm3, and the nucleosome mobility factor, FACT [72]. A criteria for RPC isolation were the ability to withstand two washes of the immunoprecipitation beads. Hence, one can anticipate other factors bind the replisome but associate too weakly to survive these conditions. We will revisit some of the factors of the RPC in sections to follow.

3. THE POLYMERASE ALPHA-PRIMASE DNA polymerases cannot initiate DNA synthesis de novo; they require a preexisting primed site (ie, a 30 ss/ds junction). Thus, initiation of replication on both the leading and the lagging strands requires a primase enzyme that synthesizes an RNA primer to initiate DNA synthesis. Since the leading strand is extended in the direction of fork movement, it may only need to be primed one time. But due to the antiparallel structure of DNA the lagging strand is extended in the opposite direction of fork unwinding, and this requires formation of multiple Okazaki fragments that each initiated by a primer synthesized by primase. Okazaki fragments are only 100–200 nucleotides in eukaryotes, and therefore priming events must be frequent [6]. Primases synthesize a short section of RNA of a dozen nucleotides or less. Why is RNA used for the primer, and not DNA? The answer to this question is not firm, but there are some reasonable possibilities. One is that rNTPs are in 10- to 100-fold higher concentration in cells compared to dNTPs [73]. The initial condensation of two nucleotides during de novo synthesis of a primer requires the protein to bind two nucleoside triphosphates at the same time. Thus, the higher concentration of rNTPs over dNTPs may have favored the use of RNA over DNA. Another possible reason is that the de novo synthesis of a primer is an inherently low-fidelity process [74–76]. Thus, use of RNA for primer formation could serve as a convenient “marker” for removal of the low-fidelity primer, and its ultimate replacement by DNA by a high-fidelity polymerase. Interestingly, archaeal primase can use dNTPs [77,78], but they bind rNTPs tighter [79] and Okazaki fragments are tipped with 50 RNA in the archaeal cell [80]. The eukaryotic primase is a four-subunit enzyme that contains both RNA primase and DNA polymerase activities in separate subunits, unlike the single subunit bacterial primase [81–83]. This enzyme is called DNA polymerase alpha-primase, referred to in this review as Pol alpha. Pol alpha was the first eukaryotic polymerase discovered. After finding that Pol alpha is both a DNA polymerase and contains an inherent primase activity [84], it

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was thought that this one enzyme complex could perform both leading and lagging strand synthesis. Indeed, for many years Pol alpha was thought to be the only eukaryotic replicative polymerase. The primase activity was found to be associated with the two smallest subunits, Pri1 (large) and Pri2 (small) [85]. Subsequent studies have shown that the catalytic site of primase is in the smallest subunit, Pri1 [21,77,86], but the larger of the two primase subunits is also required for activity, and its conserved C-terminal region contains an iron–sulfur cluster that is essential for the initiation of primer synthesis [87–89]. After synthesis of 7–9 rNMPs, the enzyme switches to the DNA polymerase site, located in the largest subunit, Pol1. This transfer occurs internally, without enzyme dissociation from DNA [90–92]. Interestingly, DNA synthesis terminates within about 20 nucleotides of extension [93,94], but Pol alpha will rebind and continue DNA synthesis, and by repeated cycles of extension can produce DNA chains of many kilobases. In the presence of replicative polymerases, Pol alpha acts as a primase by synthesis of a hybrid RNA/DNA primer of 20–30 nucleotides [94] that is then captured and extended by processive replicative DNA polymerases [95]. Structural studies of the polymerase subunit imply that it distinguishes A form RNA–DNA from B form DNA–DNA, and that the enzyme may not bind well to B form dsDNA, possibly explaining why the enzyme dissociates after a few tens of dNMPs are added to the RNA site [93]. Crystal structure analysis show the polymerase and primase active site regions contain three conserved acidic residues that bind two metal ions, common to DNA polymerase active sites [96–98]. While the Pol1 polymerase subunit is a B family polymerase, the primase shares homology to the X-family of DNA polymerases [99]. Low-resolution EM structures (eg, 25 A˚) of the entire Pol alpha shows a CTD that is connected to the B family polymerase structure by a flexible linker and that the CTD binds to the B subunit (Fig. 3) [97,100,101]. The leading and lagging strand DNA polymerases (Pol epsilon and Pol delta, respectively) also contain a B subunit of similar size that is essential to cell viability and contain an oligosaccharidebinding domain and phosphodiesterase-like region [99,102]. We know very little about the function of the B subunit in any of these DNA polymerases. The Pol1 CTD also binds the primase subunits, and thus the Pol alpha holo˚ and enzyme consists of two enzymatic components separated by >80 A connected by a flexible tether (Fig. 3). The functional consequences of this architecture and how it relates to the intramolecular hand-off of the RNA primer to the DNA polymerase are not certain.

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Fig. 3 Polymerase and primase are separated by a flexible tether in Pol alpha. (A) EM reconstruction of Pol alpha from budding yeast. (B) Cartoon illustration summarizing the EM structure. Reproduced from Fig. 1e of R. Nunez-Ramirez, S. Klinge, L. Sauguet, R. Melero, M.A. Recuero-Checa, M. Kilkenny, R.L. Perera, B. Garcia-Alvarez, R.J. Hall, E. Nogales, L. Pellegrini, O. Llorca, Flexible tethering of primase and DNA Pol alpha in the eukaryotic primosome, Nucleic Acids Res. 39 (2011), 8187–8199 by permission of Oxford University Press.

Pol alpha lacks 30 -50 proofreading exonuclease activity and this was puzzling during the years that Pol alpha was thought to be the replicative DNA polymerase. Upon discovery of the high-fidelity replicative DNA polymerases, Pol delta, and Pol epsilon (which have proofreaders), Pol alpha was recognized to function primarily as a primase that generates a hybrid RNA/DNA primer for the replicative polymerases [95,103]. Since DNA polymerase alpha has a lower fidelity than the replicative polymerases delta and epsilon, the DNA portion of the primer must either be removed or efficiently proofread by mismatch repair and ExoI [104,105]. Removal of the RNA is performed by either the Fen1 flap endonuclease or Dna2 nuclease, and replaced with DNA by Pol delta [106,107]. Why Pol alpha has a DNA polymerase is not understood. In fact, the archaeal primase consists only of the two small RNA priming subunits, homologous to those of Pol alpha, and does not contain the DNA polymerase or B subunits [98]. It is possible that the DNA polymerase activity of Pol alpha serves a role beyond priming. For example, telomeres are extended by telomerase that generates a ssDNA 50 tail. The 50 ssDNA of a telomere binds the shelterin complex and makes a DNA loop [108], but some of the telomeric ssDNA must be converted to dsDNA and perhaps Pol alpha serves this role. Pol alpha has long been known to bind a protein called Ctf4 [109]. Ctf4 is a component of the RPC and helps to stabilize the association of Pol alpha with the replisome [72,110]. EM studies indicate that full-length Ctf4 contains a central flexible hinge, complicating crystal structure analysis of the full-length protein [111]. However, the crystal structure of the C-half of

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Ctf4 has been determined (Fig. 4A and B), and it binds a peptide sequence within the N-terminal region of the polymerase subunit of Pol alpha [111]. Ctf4 is a homotrimer, with a beta-propeller forming the trimer interface, and C-terminal alpha helices that bind the Pol alpha peptide sequence (Fig. 4C). Ctf4 also binds a peptide of the Sld5 subunit of the GINS complex in the same position as the Pol alpha peptide [110,112]. Hence, the Ctf4 trimer bridges the leading strand CMG helicase to the lagging strand Pol alpha (Fig. 4D). The third protomer of Ctf4 is available to bind a third protein.

Fig. 4 Ctf4 is a homotrimer that binds peptide sequences within Pol alpha and GINS. (A) and (B) Crystal structure of the C-half of Ctf4. The trimer is formed by a beta-propeller motif (blue (gray in the print version)) that supports an alpha helical structure (yellow (light gray in the print version)). (A) Top view, (B) side view. (C) Close up of a Pol1 peptide (green (gray in the print version)) bound to the alpha helical region of Ctf4. (D) Proposed interactions of the Ctf4 trimer with Sld5 in the GINS component of CMG, and Pol alpha. The yellow (light gray in the print version) sphere suggests that an unidentified protein(s) may bind Ctf4. Adapted by permission from Macmillan Publishers Ltd. A.C. Simon, J.C. Zhou, R.L. Perera, F. van Deursen, C. Evrin, M.E. Ivanova, M.L. Kilkenny, L. Renault, S. Kjaer, D. Matak-Vinkovic, K. Labib, A. Costa, L. Pellegrini, A Ctf4 trimer couples the CMG helicase to DNA polymerase alpha in the eukaryotic replisome, Nature 510 (2014) 293–297.

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The finding that Ctf4 binds a consensus peptide is similar to studies of PCNA, which binds a short consensus peptide found in many proteins and acts as a platform for dynamic interactions with its many protein partners [30]. If Ctf4 acts in a similar fashion, it may act as a “hub” for protein trafficking within the replisome.

4. THE LEADING AND LAGGING STRAND DNA POLYMERASES EPSILON AND DELTA Eukaryotes use two different B-family DNA polymerases for leading and lagging strand synthesis, Pol epsilon and Pol delta, respectively [113–115]. Both polymerases contain a 30 -50 exonuclease proofreader, and a mutation in the exonuclease site of either enzyme is associated with human cancers [116]. Pol delta was first identified as the replicative polymerase from biochemical studies of simian virus 40 (SV40) replication [103]. This small virus only encodes one protein for its own replication, the SV40 large T-Ag. T-Ag is both an origin binding protein and a helicase. Hence, all the remaining enzymes for replication are derived from the host. The in vitro SV40 DNA replication system initially required cell extracts and pure T-Ag [117], but years of intensive study and fractionation eventually identified each of the required host proteins. The studies showed that Pol alpha synthesized the primed sites for both strands, and that bulk replication was performed by a second DNA polymerase, Pol delta, along with its required accessory factors, PCNA and replication factor C (RFC) [103]. PCNA and RFC were later shown to be the clamp and clamp loader, upon discovery of the function of the analogous accessory factors in E. coli, discussed in Section 5 [18,28,29]. Although SV40 DNA replication only requires Pols alpha and delta, genetic studies identified Pol epsilon as a third DNA polymerase that is essential for cellular replication [118]. Assignment of Pol epsilon to the leading strand and Pol delta to the lagging strand has followed from several experiments by the Kunkel lab and his collaborators [113–115,118,119]. Initial studies used a subtle mutation in the active site of each polymerase that generates an asymmetric mutation; AT is converted to GC only when the polymerase replicates one strand but not the other. The mutant Pol delta or Pol epsilon retains wild-type catalytic and proofreading activity, and the mutator phenotype is quite mild. Strains having either polymerase mutant were constructed, and a Ura marker gene was placed in one direction or the other near the strong, early firing origin ARS306 [114,115]. Mutations

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in the URA3 gene were sequenced from strains containing either the Pol delta or the Pol epsilon active site mutant. These studies strongly supported the use of Pol epsilon on the leading strand and Pol delta on the lagging strand. Further studies were performed with polymerase mutants that misincorporate ribonucleotides at relatively high frequency, enabling misincorporation events to be examined genome wide by deep sequencing technology, and the results supported Pol epsilon as the major leading strand polymerase and Pol delta on the lagging strand [119]. These studies were extended to S. pombe with similar conclusions [120]. Furthermore, DNA cross-linking studies demonstrated that Pol epsilon cross-linked to the leading strand and Pol delta cross-linked to the lagging strand [121]. Biochemical study of the polymerases showed that Pol delta functions with Fen1 nuclease to efficiently process RNA from Okazaki fragments and provides a ligatable nick, but Pol epsilon is not capable of this reaction, supporting a role of Pol delta on the lagging strand [122,123]. While these studies support the assignment of Pol epsilon on the leading strand, there is one study that arrived at a different conclusion in which Pol delta replicates both strands [124]. The different conclusions may possibly be explained by the need to mutate certain repair pathways to perform studies of this sort [124,125]. For example, use of the mutant polymerases converting AT to GC required the mismatch repair pathway to be knocked out, and use of rNTP misincorporation mutants of the DNA polymerases required the RNaseH repair pathway to be knocked out. Hence, the asymmetric distribution of mutations might possibly originate from differential pathways for repair of the leading or lagging strands. Given the wealth of data on Pol epsilon as the leading polymerase, this review will assume that Pol epsilon operates on the leading strand. There is no disagreement about use of Pol delta on the lagging strand. The reader should keep in mind that a conclusive assignment of Pol epsilon as the leading strand polymerase awaits further studies. The active polymerase region of Pol epsilon, encompassing the first half of the POL2 gene, can be deleted without killing the cell, although progression through S phase is severely compromised [126,127]. It was proposed that a back-up DNA polymerase, probably Pol delta, takes over leading strand synthesis under this circumstance [126,127]. This observation is reminiscent of the case in E. coli in which the dnaE gene encoding the replicative polymerase (alpha subunit of Pol III) can be deleted but slow growing mutant cells still survive through use of Pol I [128]. Interestingly, the essential part of Pol2 is the C-terminal half, having the sequence of an inactive B-family polymerase, and it is thought to serve an essential structural role.

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Pol epsilon is composed of four subunits, the large polymerase subunit, the B subunit called Dpb2 (DNA polymerase-binding protein 2), and two small histone-like subunits, Dpb3 and Dpb4 [129]. Of these, only Dpb2 is essential in budding yeast, although mutants in Dpb3 or Dpb4 disrupt the ability to maintain heterochromatin. All three accessory subunits bind the C-half of Pol2, and considering that Dpb2 is essential, this may possibly explain the essential role of the C-half of Pol2. Pol epsilon is stimulated in synthesis by the PCNA clamp, but there are no obvious PCNA-binding motifs in the Pol epsilon subunits. Thus, the points of contact between Pol epsilon and PCNA are not yet identified. Pol delta consists of three subunits in yeast: Pol3 (the polymerase), Pol31 (B subunit), and Pol32 [130]. Pol delta in higher organisms contains a small fourth subunit, p12 [131]. Pol delta requires the PCNA clamp for detectable activity, and the Pol32 subunit contains a PCNA-binding motif. However, the POL32 gene is not essential in yeast, and therefore other, less obvious points of contact must link Pol delta to PCNA. Pol delta, like Pols epsilon and alpha, is a B family DNA polymerase. Eukaryotes contain a fourth B family polymerase, Pol zeta, which acts as a translesion bypass polymerase capable of extending DNA over a template lesion [132]. Interestingly, Pol zeta contains the Pol31 and Pol32 subunits [133,134]. For information about translesion polymerases, the reader is referred to reviews on this fascinating topic [26,132]. Pols delta, epsilon, and alpha are B family polymerases, but they all share a feature that does not generalize to other B family polymerases. Specifically, these polymerases have a cysteine-rich region in their CTDs that contain two metal-binding sites, CysA and CysB [135]. These sites have been characterized in the Pol3 subunit of Pol delta, and the findings are likely to generalize. The CysB site binds an iron–sulfur cluster and is required for Pol3 to bind the B subunit [135]. The CysA site binds zinc, and this site is needed for Pol delta to function with PCNA [135]. The fact that iron– sulfur clusters have different oxidation states suggest that it may sense the oxidation–reduction state of the cell, but further studies are needed to explore the functions of these metal-binding sites. The crystal structure of the catalytic regions of DNA polymerases alpha, delta, and epsilon shows the expected structure of B family polymerases [97,136,137]. As with DNA polymerases of all six families (A, B, C, X, Y, RT), the B family polymerases are composed of three subdomains (fingers, palm, and thumb) that form the shape of a right hand, and the palm contains the active site, in which three conserved acidic residues bind two

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metal ions that catalyze DNA polymerization [25,26]. Pol epsilon contains a distinct feature not present in Pols delta and alpha, referred to as a “processivity domain” [136]. The processivity domain closes the gap between the fingers and thumb, thereby completely encircling the DNA substrate.

5. THE PCNA CLAMP AND RFC CLAMP LOADER The first discovery of a circular protein that surrounds DNA was the E. coli beta clamp [28,29]. Since that time numerous proteins have been shown to encircle DNA for function. The initial biochemical study identifying the ring shape demonstrated that beta binds DNA topologically and slides along dsDNA and can slide off the end of a linear DNA [138]. Proof that beta was a circular protein soon followed, with the crystal structure showing a homodimer that forms a ring with a hole of sufficient diameter to encircle dsDNA [28]. The fact that each beta protomer is constructed of three globular subdomains enabled the prediction that eukaryotic PCNA, which is 2/3 the size of beta, would be a homotrimer of subunits composed of two globular subdomains [28]. This prediction was proven correct with the crystal structure of PCNA (Fig. 5A) [139,140]. These clamps bind to DNA polymerases and hold them to DNA for high processivity. Many subsequent studies of PCNA showed that it binds numerous proteins involved in maintaining genomic integrity, including mismatch repair proteins, nucleosome assembly factors, nucleases, ligase, lesion bypass DNA polymerases, and cell cycle kinases. In fact, the E. coli beta clamp has also been shown to bind and function with numerous other proteins involved in maintaining genomic integrity. For more details, the reader is referred to a recent review on this topic [30] and to chapter “The many roles of PCNA in eukaryotic DNA replication” by Boehm et al. Sliding clamps require a five-subunit clamp loader that uses ATP to assemble the clamp at a primed site [18]. In eukaryotes, the clamp loader is referred to as RFC that was discovered as a polymerase accessory protein in the SV40 DNA replication system [103]. The subunits of bacterial and eukaryotic clamp loaders are homologous to one another, and each are in the AAA+ family of ATPases (ATPases associated with diverse cellular activities) [141,142]. The ATP-binding region of AAA+ proteins is an ancient fold found in different types of proteins, including proteases, vesicle fusion factors, helicases, origin-binding proteins, and many others [19]. AAA + proteins generally function as oligomers that remodel other proteins.

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Fig. 5 Structure and function of the PCNA clamp and RFC clamp loader. (A) Crystal structure of human PCNA (1AXC) [139]. The subunits of the homotrimer are shown in different colors (gray shades in the print version). (B) Structure of yeast RFC bound to PCNA (1SXJ). RFC is shown in the space filling representation. PCNA is in the ribbon representation. DNA (green (dark gray in the print version)/orange (gray in the print version)) is modeled into the structure. (C) Scheme of RFC clamp loader function in assembly of PCNA onto a primed site. Panel (B) adapted with permission of Nature Publishing Group from Fig. 6b of G.D. Bowman, M. O'Donnell, J. Kuriyan, Structural analysis of a eukaryotic sliding DNA clamp-clamp loader complex, Nature 429 (2004) 724–730.

The first AAA+ protein to be solved structurally was the delta prime subunit of the E. coli clamp loader [142]. Subsequently, the structures of clamp loaders from E. coli, yeast, and phage T4 were solved [143–145]. These clamp loaders from diverse organisms have a similar structure and mechanism of action. The ATP sites are located at the interface between subunits, requiring residues from both protomers for hydrolysis, and this strategic architecture organizes the structural changes in a multiprotein complex that are needed to open a clamp, position it on a primed site, and to close the clamp around DNA. The structure of yeast RFC bound to PCNA is shown in Fig. 5B. The five-clamp loading subunits are arranged in a right hand spiral with the AAA + domains held together by strong interactions between their CTDs that form a tight pentameric collar. The AAA+ domains contain a gap between two of the subunits; the gap enables DNA to pass into a central chamber

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formed by the AAA + domains of the five subunits. ATP is required for RFC to open the PCNA clamp, but PCNA is not open in the crystal structure despite the presence of ATP. This is due to use of ATP site mutations in RFC to prevent hydrolysis during crystal formation. However, by analogy to other clamp–clamp loader structures [145,146], the PCNA interface that opens is directly below the gap between RFC1 and RFC5. An outline of the mechanism by which clamp loaders function is illustrated in Fig. 5C and is an amalgam of structural and biochemical information on clamp loaders from E. coli, phage T4, yeast, and human. Three ATP sites are located at subunit interfaces, and residues from both subunits at each ATP site are essential to nucleotide hydrolysis. Upon PCNA ring opening, a primed site can pass through the gap in PCNA and between the AAA+ domains of RFC1 and RFC5 to fit into the inner chamber of RFC. The DNA-binding site of RFC (and the bacterial clamp loader) contains conserved DNAbinding residues located on loops in the five AAA+ domains [143,147]. The sequence independent, primed site structure-dependent specificity is facilitated by the pentameric collar that has no opening for DNA, and thus the DNA must sharply bend out the side of the clamp loader. The ss/ds DNA junction of a primed site contains the flexibility needed for this sharp bend, but dsDNA is too rigid. Binding of DNA induces the five subunits to adopt a pitch that closely follows the DNA duplex, and this organizes the ATP site residues into an active conformation [145,148]. ATP hydrolysis enables the clamp loader to eject, and the PCNA ring to close around the primed site for use by DNA polymerase (or other enzymes).

6. THE EUKARYOTIC REPLISOME STRUCTURE AND FUNCTION Recent biochemical studies have reconstituted an active eukaryotic replication fork from purified proteins (Fig. 6A) [32,33]. These protein factors also enabled reconstitution of replisome assemblies that have been studied by single-particle EM 3D reconstruction techniques, revealing a surprising architecture of the core eukaryotic replisome [71]. The factors required to reconstitute a functional eukaryotic replication fork include: Pol alpha, Pol delta, Pol epsilon, the RFC clamp loader, PCNA clamp, RPA SSB protein, and the 11-protein CMG helicase complex, for a total of 31 different subunits. Replication studies utilized a linear synthetic 3 kb forked DNA that lack dC on one strand and dG on the other, enabling specific monitoring of either the leading or lagging strand depending on the radionucleotide added to the assay. The CMG helicase functioned with Pol

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Fig. 6 Reconstitution of eukaryotic replication forks reveals the mechanism of assembly. (A) SDS polyacrylamide gels of proteins used in replication fork reconstitution studies. (B) Scheme of replication fork assembly having distinct DNA polymerases on the two strands. Left, Pol alpha interacts with CMG to prime both strands. Middle, Pol epsilon functions with CMG on the leading strand (top), while Pol delta does not (bottom). Right, Pol delta functions on the lagging strand, while Pol epsilon does not. Adapted with permission from Fig. 8 of R.E. Georgescu, G.D. Schauer, N.Y. Yao, L.D. Langston, O. Yurieva, D. Zhang, J. Finkelstein, M.E. O'Donnell, Reconstitution of a eukaryotic replisome reveals suppression mechanisms that define leading/lagging strand operation, Elife 4 (2015) e04988.

epsilon, RFC, PCNA, and RPA to synthesize the leading strand, while reactions using Pol delta in place of Pol epsilon were much less efficient, indicating that CMG specifically stabilized Pol epsilon on the leading strand [32]. This result is consistent with genetic studies identifying Pol epsilon as the leading strand polymerase. A subsequent study revealed the

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underlying basis by which Pol epsilon is stabilized on the leading strand by CMG. Specifically, CMG directly binds to Pol epsilon, forming a 15-subunit CMG–Pol epsilon (CMGE) complex that can be isolated on a glycerol gradient [149]. Pol alpha functions with CMG to synthesize primed sites and can even function distributively to extend the leading and lagging strands in the absence of Pol epsilon [33]. Interestingly, Pol alpha required CMG for priming activity in the presence of the RPA SSB protein, indicating a direct interaction of Pol alpha with CMG [33]. Indeed, previous studies found that the primase subunits of Pol alpha bind Mcm4 and that the Mcm4,6,7 complex stimulates priming activity [150]. When Pol epsilon is present with Pol alpha and CMG, Pol epsilon switches with Pol alpha after priming [33]. Unexpectedly, Pol epsilon was incapable of producing lagging stand products with CMG/Pol alpha and even inhibited Pol alpha in the extension of lagging strand primers [33]. This result was unanticipated, especially given that Pol epsilon enhanced leading strand synthesis in the very same reactions that it decreased lagging strand synthesis. These opposite results on the two strands suggest that in the presence of CMG, Pol epsilon displays an inherent bias for activity on the leading strand. Addition of Pol delta to reactions containing Pols alpha and epsilon resulted in efficient Okazaki fragment synthesis, with similar levels of both leading and lagging strand products [33]. In overview, the biochemical studies of the eukaryotic replication enzymes indicate that the asymmetric replisome architecture, with distinct polymerases on the leading and lagging strands, is inherent to the 31 core replisome proteins, and the many additional proteins known to travel with the replisome are not required for this aspect of fork architecture and function. Asymmetry is based in the recruitment of Pol epsilon to the leading strand by CMG, the inability of Pol epsilon to function on the lagging strand, the high activity of Pol delta on the lagging strand, and the low activity of Pol delta with CMG on the leading strand. These findings imply that the eukaryotic replication fork uses distinct polymerases for the leading and lagging strands because it is the only active solution for fork progression with CMG, as illustrated in Fig. 6B. As described earlier, a 15-protein leading strand CMGE complex can be reconstituted from CMG and Pol epsilon [149]. In addition, a mixture of Pol epsilon, CMG, Ctf4, and Pol alpha yielded a 20-protein particle (CMG–PolE–Ctf4–Pol alpha) that could be imaged in the EM [71]. To image the replisome and help identify each component, the replisome was built up in stages. First, the 3D structure of the CMGE was determined

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Fig. 7 EM 3D reconstruction of the CMGE leading strand replisome. Single-particle EM of the 15-subunit CMGE complex from Saccharomyces cerevisiae resulted in a 15 Å resolution structure with protein assignments as shown. Adapted with permission from Nature Publishing Group (NSMB). J. Sun, Y. Shi, R.E. Georgescu, Z. Yuan, B.T. Chait, H. Li, M.E. O'Donnell, The architecture of a eukaryotic replisome, Nat. Struct. Mol. Biol. 22 (2015) 976–982.

˚ resolution (Fig. 7). Then additional protein complexes were imaged; at 15 A the final particle contains a helicase, primase, a leading strand polymerase, and a lagging strand polymerase (Fig. 8A and B). Surprisingly, the two DNA polymerases are located on the two opposite sides of the helicase. Prior to this, DNA polymerases were thought to act only behind the helicase. DNA is demonstrated to enter CMG via the CTDs of the Mcm2–7 ring [151], and given this DNA orientation, the polymerase on the C-side of CMG must ride ahead of the replication fork. Application of protein crosslinking with mass spectrometry readout identified Pol epsilon as the polymerase that occupies the position at the C-side of CMG, and thus Pol epsilon rides on top of the helicase, adjacent to the unwinding point where the parental DNA enters into CMG (Fig. 8B). CMG functions by the steric exclusion model of helicase unwinding [54], and therefore the lagging strand is excluded to the outside of CMG, and the leading strand ssDNA threads though the Mcm2–7 central channel. To accommodate this unwinding mode, the leading strand must make a U-turn when it comes out the NTD side of Mcm2–7 in order to reach Pol epsilon at the C-side of CMG. This DNA path requires about 40 nucleotides of ssDNA, unless there is a shortcut for the leading ssDNA to reach Pol epsilon through the middle of CMG (eg, through the Mcm2/5 gate). Interestingly, there is evidence for substantial ssDNA on the leading strand in the Xenopus system in which use

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Fig. 8 Architecture of the eukaryotic replisome. (A) 2D class averages replisome subcomplexes reconstituted using proteins of the Saccharomyces cerevisiae replisome. (B) Protein arrangement within the eukaryotic replisome as inferred from the 3D structure of CMGE along with the 2D class averages of subcomplexes [71], and the experimentally determined direction of DNA threading through CMG [151]. Panel (A) adapted with permission from Fig. 2a and Fig. 5 of J. Sun, Y. Shi, R.E. Georgescu, Z. Yuan, B.T. Chait, H. Li, M.E. O'Donnell, The architecture of a eukaryotic replisome, Nat. Struct. Mol. Biol. 22 (2015) 976–982. Panel (B) adapted with permission from Nature Publishing Group (NSMB). J. Sun, Y. Shi, R.E. Georgescu, Z. Yuan, Chait, H. Li, M.E. O'Donnell, The architecture of a eukaryotic replisome, Nat. Struct. Mol. Biol. 22 (2015) 976–982.

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of DNA containing a site-specific intrastrand cross-link showed that the replisome stops on the leading strand 20–40 nucleotides before the crosslink [54]. It should be noted that it is still possible that the leading ssDNA could enter CMG through the N-terminal face, which would place Pol epsilon under CMG, and Pol alpha-primase would ride on top of CMG adjacent to the DNA split point where it could prime the leading strand. For example, the many proteins that assemble CMG on DNA at an origin may be different than the biochemical in vitro studies that identified the DNA direction through CMG. If upon further study the replisome architecture depicted in Fig. 8 stands, one may ask why cells would have evolved a replisome architecture with a polymerase riding ahead of the helicase? Pol epsilon has been documented to bind histone octamers [152,153]. Hence, one possible reason for Pol epsilon at the top of the fork is that this would enable Pol epsilon to encounter nucleosomes on the parental DNA. Furthermore, genetic studies show that the Dpb3/4 subunits of Pol epsilon are required for heterochromatin maintenance during replication, consistent with Pol epsilon interaction with nucleosomes [153–155]. The placement of Pol epsilon on top of CMG would bring the leading strand duplex product DNA above the helicase and close to the parental DNA, and this may facilitate nucleosome transfer from the parental duplex to the leading strand daughter duplex. Nucleosomes are heavily modified by posttranslational modifications, providing epigenetic information that underlies the differentiated state of the different cells of a multicellular organism. Hence, cells must divide asymmetrically during embryogenesis, enabling daughter cells to take distinct developmental paths for the different tissues of the body. The replisome is the only cellular machine that must deal with every nucleosome in the cell, and its asymmetric architecture could play a role in the production of two different daughter cells that have unique epigenetic nucleosome signatures. Specifically, if asymmetric distribution of nucleosomes occurs upon passage of the replication fork, even if this happens at only a few strategic sites in the genome, it could underlie asymmetric cell division during development of a multicellular animal.

7. COMPARISON OF BACTERIAL AND EUKARYOTIC REPLISOMES Most of the core replisome proteins of bacteria and eukaryotes are unrelated in both sequence and structure [13–15]. Thus, the organization

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Fig. 9 Comparison of bacterial and eukaryotic replisomes. (A) The bacterial replisome. (B) The eukaryotic replisome. See text for details.

and internal workings of bacterial replisomes may be quite different from eukaryotic replisomes. The current view of the organization of the bacterial replisome is illustrated in Fig. 9A. The bacterial replisome is organized by the clamp loader, which contains three copies of the tau subunit [156]. The tau subunit has a C-terminal extension that is not required for clamp loading and binds the DnaB helicase and the DNA Pol III [157]. Thus, the bacterial replisome has three identical copies of Pol III linked to one central clamp loader that also adheres to the hexameric helicase encircling the lagging strand. Three molecules of Pol III in the bacterial replisome were first observed in vitro [156], and then confirmed in vivo [158]. Use of three Pol IIIs at one replication fork was unexpected given there are only two strands of DNA to replicate. Both in vitro and in vivo single molecule studies indicate that two of the Pol IIIs function on the lagging strand [158–160]. Considering that the lagging strand has numerous primed sites, use of multiple polymerases may be advantageous. Bacterial primase interacts with DnaB helicase for priming activity, but primase does not form an integral part of the moving replisome; E. coli primase repeatedly binds and releases from DnaB during priming activity and fork movement [161]. As the bacterial fork progresses, the lagging strand ssDNA is coated with SSB protein that protects and organizes the ssDNA [6].

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As RNA primers are formed, the clamp loader loads new clamps onto RNA primers for function with Pol III. Single molecule studies show that the beta clamp can tether Pol III to DNA for many tens of kilobases [162], but bacterial Okazaki fragments are only 1–2 kb in length [6]. Therefore, some mechanism must limit the processivity of Pol III-beta and loosen the grip of the lagging Pol III from its beta clamp, freeing Pol III to recycle from a completed Okazaki fragment to a new primed site. There are at least two mechanisms that perform this polymerase recycling function. One is called “collision release” and occurs when Pol III-beta completes a section of DNA and bumps into a 50 terminus (ie, of the previous Okazaki fragment). When this occurs, Pol III loosens its grip on beta and ejects from DNA, leaving the clamp behind [138]. Collision release is also observed in the phage T4 system [163]. One study indicates that the rate of collision release may not be fast enough for a moving fork [164], but experiments with reconstituted E. coli replisomes demonstrate that collision release occurs about 50% of the time [165]. There is a second mechanism, referred to as “signal release” in which the lagging polymerase receives a signal to release from its clamp before an Okazaki fragment is complete, first observed in the T4 system [166]. Signal release has since been observed in E. coli and phage T7 systems [165,167]. The signal in phage replisomes may be either primase, the primer generated by primase, or the clamp assembled on a new primed site [166,167]. Study of the signal in the E. coli system rules out primase and indicates instead that the signal is the torsional strain incurred by a replisome having connected polymerases, each of which travel in spirals to make helical dsDNA products [165]. Independent (nonconnected) leading and lagging DNA polymerases would be able to freely rotate during synthesis of their respective strands. Torsional strain incurred by the rotation of connected leading/lagging strand polymerases generate the high amount of force required to separate the tight Pol III-beta connection, and experimental evidence supports the role of torsional strain in signal release in the E. coli system [165]. For further information on the bacterial replisome, see chapter “The E. coli DNA replication fork” by Lewis et al. The eukaryotic replisome has certain features in common with the bacterial replisome, but also many important differences. We will review some of these distinctions later, but it is important to keep in mind that replisome studies in the eukaryotic system are not as advanced as the study of bacterial systems, and thus there is still much to be learned about eukaryotic replisomes before a true comparison can be considered firm or complete. One of the obvious differences between eukaryotic and bacterial replisomes

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is that the RFC clamp loader is not reported to stably associate with any component of the eukaryotic replisome, nor is it identified as a component of the RPC. Thus, at the current time the clamp loader is not thought to be an integral part of the eukaryotic replisome. This contrasts with the bacterial replisome where the clamp loader is the central organizer of the replisome. However, both replisomes utilize sliding clamps that provide processivity to their respective polymerases. Another difference between bacterial and eukaryotic replisomes is the location of the helicase. The CMG helicase tracks along the leading strand, opposite the tracking direction of bacterial replicative helicases. However, like bacterial systems [168–171], the eukaryotic leading polymerase (Pol epsilon) interacts with the helicase for function [32,149]. Thus, a functional polymerase–helicase connection generalizes from bacteria to eukaryotes. Another similarity is that Pol alpha priming activity requires CMG during fork progression in the presence of the RPA SSB protein [33]. Hence, the property of a primase–helicase interaction required for priming also appears to generalize from bacteria to eukaryotes. The use of a Pol alpha complex that generates a RNA–DNA primer appears unique to eukaryotes, but the reader is reminded that the exact function of the DNA polymerase component of Pol alpha is probably yet to be discovered (discussed earlier). Distinctive to the eukaryotic replisome is the fact that Pol alpha-primase is an integral part of the replisome, while the primase of bacteria is not [72]. The EM 3D reconstruction of the core eukaryotic replisome places the leading polymerase above the helicase [71]. If the function of this architectural facet relates to nucleosomes, this feature may not generalize to bacteria. The use of a Ctf4 trimer “hub” in the eukaryotic replisome might be analogous to the trimeric tau CTD in bacteria. In both cases, the trimer mediates a polymerase-to-helicase interaction, but Ctf4 is not essential in budding yeast and Pol alpha still functions with CMG without Ctf4 in vitro [33], while the CTD of E. coli tau is essential both in vivo and for in vitro replisome activity [170,172]. It is too early to tell if Ctf4 should be compared to the CTD of bacterial tau. Another distinction of eukaryotic replisomes, compared to the E. coli replisome, is the use of two different DNA polymerases for the leading and lagging strands in eukaryotes. Interestingly, some bacteria have two essential C-family polymerases [173]. But there is conflicting evidence whether they both function at the fork or whether one may be specific to repair [174,175]. For example, the second C-family polymerase in the

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Gram-positive system is essential, but its synthetic rate with beta is 10 times slower than fork movement, and it has low fidelity because it lacks a proofreading exonuclease [174]. Further study is required to resolve this interesting issue. The eukaryotic lagging strand Pol delta is not known to travel with the replisome, and this is in sharp contrast to E. coli, phage T4, and phage T7 systems in which the lagging polymerase travels with the fork. When a lagging polymerase is part of the replisome and travels with the fork, it forms DNA loops, one for each Okazaki fragment [176]. These DNA loops have been observed in the EM and by single molecule experiments [176]. If eukaryotic Pol delta does not travel with the replisome, there would be no Okazaki fragment loops at eukaryotic forks. Further studies are required to address this issue. Eukaryotic replisomes are highly regulated by posttranslational modifications, unlike bacteria. Eukaryotic replisomes are also known to carry with them several factors involved in the DNA damage checkpoint response that controls fork movement or stability at times of DNA damage [72,177–179]. The CMG helicase is phosphorylated during DNA damage, and treatment with checkpoint kinases downregulates CMG helicase activity in vitro [180]. DNA damage also results in ubiquitination of the PCNA clamp, and this modification correlates with enhanced translesion synthesis [181,182]. Several other eukaryotic replisome proteins are known to be phosophorylated or ubiqutinated in a cell cycle-specific fashion, but the function of most of these modifications is unclear and will require further study.

8. FUTURE PERSPECTIVES We have learned numerous important insights about how replisomes function from studies of bacterial cells and their bacteriophages. However, the different evolutionary lineage of the eukaryotic core replication proteins, and the many additional factors they contain that have no known bacterial counterpart make future studies of the eukaryotic replisome very important. This is especially true given the central importance of DNA replication to genomic integrity, and thus to human health and disease. The increased complexity of eukaryotic replisomes relative to bacterial replisomes delayed the development of a fully reconstituted in vitro eukaryotic system needed to understand the internal workings of the replisome machinery. Only recently have studies accomplished the reconstitution of the eukaryotic replisome from pure recombinant proteins, and these systems will provide

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a basis for further mechanistic investigations on the structure and function of the eukaryotic replisome. Important questions include: Why does the CMG helicase have 11 subunits, while all other replicative helicases are homohexamers? What is the function of the numerous additional proteins that travel with the eukaryotic replication fork? How do various posttranslational modifications of replisome proteins alter their activity? How does the replisome deal with nucleosomes? Do nucleosome chaperones cooperate with the replisome during fork progression? How does the eukaryotic replisome deal with encounters with RNA polymerase and how does it cope with various types of DNA damage? The many new and recent findings about eukaryotic replisomes detailed in this review have provided a rich foundation upon which further advances can be made.

ACKNOWLEDGMENTS The authors are grateful for funding from the National Institutes of Health, US 1R01GM115809 and the Howard Hughes Medical Institute. We also appreciate the assistance of Dr. Nina Yao for help in preparing figures.

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CHAPTER SEVEN

The Many Roles of PCNA in Eukaryotic DNA Replication E.M. Boehm, M.S. Gildenberg, M.T. Washington1 Carver College of Medicine, University of Iowa, Iowa City, IA, United States 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. PCNA Structure and Function 3. The Role of PCNA in Normal DNA Replication 4. The Role of PCNA in Translesion Synthesis 5. The Role of PCNA in Error-Free Damage Bypass 6. The Role of PCNA in Break-Induced Replication 7. The Role of PCNA in Mismatch Repair 8. The Role of PCNA in Replication-Coupled Nucleosome Assembly 9. Putting It All Together Acknowledgments References

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Abstract Proliferating cell nuclear antigen (PCNA) plays critical roles in many aspects of DNA replication and replication-associated processes, including translesion synthesis, error-free damage bypass, break-induced replication, mismatch repair, and chromatin assembly. Since its discovery, our view of PCNA has evolved from a replication accessory factor to the hub protein in a large protein–protein interaction network that organizes and orchestrates many of the key events at the replication fork. We begin this review article with an overview of the structure and function of PCNA. We discuss the ways its many interacting partners bind and how these interactions are regulated by posttranslational modifications such as ubiquitylation and sumoylation. We then explore the many roles of PCNA in normal DNA replication and in replication-coupled DNA damage tolerance and repair processes. We conclude by considering how PCNA can interact physically with so many binding partners to carry out its numerous roles. We propose that there is a large, dynamic network of linked PCNA molecules at and around the replication fork. This network would serve to increase the local concentration of all the proteins necessary for DNA replication and replication-associated processes and to regulate their various activities.

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1. INTRODUCTION Since its discovery in the late 1970s, our view of proliferating cell nuclear antigen (PCNA) and its roles in DNA replication and genome maintenance has expanded considerably. PCNA was originally identified as the target of an autoimmune antibody derived from patients with systemic lupus erythematosus [1]. This protein was later shown to be one produced predominantly in proliferating and transformed cells [2–4]. By the middle of the 1980s, the involvement of PCNA in DNA replication was suggested based on its pattern of staining throughout the cell cycle [5]. Definitive evidence of a role for PCNA in DNA replication came a couple years later with the discovery that PCNA is required for the replication of simian virus 40 in vitro [6,7]. It was soon realized that PCNA was an auxiliary protein for DNA polymerase delta (pol δ) that increases its activity by making it more processive [8–10]. PCNA was subsequently shown to be an auxiliary factor for DNA polymerase epsilon (pol ε) [11–14]. By the early 1990s, the role of PCNA came to be viewed as being the processivity factor of eukaryotic replicative polymerases. An understanding of how PCNA confers high processivity to DNA polymerases was achieved when the X-ray crystal structure of PCNA was determined [15]. PCNA was shown to be a ring-shaped trimer similar to the structure of the Escherichia coli beta clamp determined a couple years earlier [16]. By the middle of the 1990s, it was known that PCNA is loaded onto double-stranded DNA by replication factor C (RFC) [17,18], where the PCNA functions as a sliding clamp that binds and anchors polymerases onto the DNA. As more and more PCNA-interacting partners were identified, it became clear that PCNA is not simply a processivity factor for replicative polymerases. It interacts with and regulates the activities of many proteins involved in Okazaki fragment maturation [19,20], mismatch repair [21], nucleotide excision repair [22], and translesion synthesis [23–26]. It also interacts with proteins involved in other processes such as cell cycle control [27–29], sister chromatid cohesion [30], epigenetic inheritance [31], and S-phase-specific proteolysis [32]. By the early 2000s, PCNA came to be viewed as an important hub protein that is critical for organizing and orchestrating events at the replication fork and other sites of DNA synthesis. Since the early 2000s, it has become clear that the regulation of several DNA metabolic processes is governed by posttranslational modifications

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of PCNA, most notably ubiquitylation and sumoylation [33,34]. Ubiquitylation of PCNA promotes translesion synthesis via the recruitment of translesion synthesis polymerases to stalled replication forks [35]. Sumoylation of PCNA inhibits recombination via the recruitment of antirecombinases to sites of DNA synthesis [36,37]. In this chapter, we will describe the many roles of PCNA in eukaryotic DNA replication and in replication-associated processes. We will begin by discussing the features of the structure and function of PCNA common to all of its roles. Then we will focus on its roles in normal DNA replication, translesion synthesis and error-free damage bypass, break-induced replication, mismatch repair, and replication-coupled nucleosome assembly.

2. PCNA STRUCTURE AND FUNCTION Sliding clamps are proteins that encircle double-stranded DNA and are found in all three domains of life. Although these proteins have different oligomeric states, they all possess a general pseudo-sixfold ring-shaped structure. Bacterial sliding clamps form homodimers, whereas archaeal and eukaryotic sliding clamps form homotrimers and heterotrimers. These sliding clamps function as platforms for recruiting and regulating various enzymes that function in DNA replication and repair, such as polymerases, nucleases, and ligases [38]. Although there is little sequence similarity among the sliding clamps across the domains of life, their striking structural similarity demonstrates the evolutionary importance of having such scaffolds for bringing proteins to sites of DNA synthesis. Eukaryotic PCNA is a homotrimer with each monomer composed of two similarly folded domains connected by an interdomain-connecting loop [15] (Fig. 1A). Domain 1 is comprised of residues 1–117, domain 2 is comprised of residues 135–258, and the interdomain-connecting loop is comprised of residues 118–134. The six structural domains form a ring with an outer layer of 6 β-sheets and an inner layer of 12 α-helices that line the central hole of the ring. The central hole is lined with positively charged residues that can form electrostatic interactions with the duplex DNA. The ˚ , which is wider than the diameter of B-form diameter of the hole is
35 A ˚ DNA (
20 A). Models derived both from X-ray diffraction data [42] and from molecular dynamics simulations [43] provide strong evidence that the DNA is significantly tilted at an angle as it passes through the center of the hole in order to contact the positively charged residues on the α-helices.

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A

B QTSMTDFY QGRLDDFF MQTLESFF QRSIMSFF QxxhxxFF

C

D

Fig. 1 PCNA structure. (A) The structure of yeast PCNA (PBD: 1PLQ) [15] is shown from a front view and a side view. Domain 1 is blue (dark gray in the print version), domain 2 is green (light gray in the print version), and the interdomain-connecting loop (IDCL) is red (gray in the print version). (B) The structure of yeast PCNA bound to the PIP motif of the Cdc9 DNA ligase (PDB: 2OD8) [39] shown from a front view. The PIP motif is blue (dark gray in the print version). Shown also are the sequences of several PIP motifs. (C) The structure of human PCNA bound to three full-length FEN1 proteins (PDB: 1UL1) [40] is shown from a front view. The three FEN1 molecules are blue (dark gray in the print version), green (light gray in the print version), and red (gray in the print version). (D) The structure of yeast SUMO-modified PCNA (PDB: 3PGE) [41] is shown from a front view. The SUMO moieties are blue (dark gray in the print version).

Single-molecule analysis showed that there are two distinct modes by which PCNA moves along double-stranded DNA [44]. As PCNA diffuses along the DNA, most of time the ring rotates following the helical pitch of the DNA. This rotation ensures that polymerases and other proteins bound to PCNA would be positioned consistently with respect to the DNA helix as

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the protein complex moves along it. A small fraction of the time, however, PCNA translocates along the DNA without following the helical pitch of the DNA. This mode of movement allows PCNA to more rapidly slide along the DNA. It also allows greater freedom of rotation, which under certain circumstances would allow optimal repositioning of PCNA-bound proteins with respect to the DNA. The PCNA ring has two faces, which we refer to as the front face and the back face (Fig. 1A). The front face points in the direction of DNA synthesis and contains the C-terminus of each monomer as well as the interdomainconnecting loop. The majority of proteins that interact with PCNA do so within a hydrophobic pocket on the front face of PCNA near the interdomain-connecting loop [45–47]. Binding on the front face allows these interacting proteins to access the primer terminus of the replicating DNA. The back face of PCNA, by contrast, points away from the direction of DNA synthesis and contains several extended loops and lysine-164, a site of ubiquitylation and sumoylation [33]. It has been suggested that these posttranslational modifications bind specific proteins and hold them in reserve on the back face of PCNA until they are needed [41,48,49]. The proteins that interact with the hydrophobic pocket on the front face of the PCNA ring generally contain one or more PCNA-interacting protein (PIP) motifs [50–52] (Fig. 1B). PIP motifs are sequences of eight amino acids with a conserved glutamine at position 1, a conserved aliphatic residue (leucine, isoleucine, or methionine) at position 4, and two adjacent, conserved aromatic residues (phenylalanine or tyrosine) at positions 7 and 8. The conserved aromatic side chains bind within the pocket comprised of isoleucine-128 in the interdomain-connecting loop and proline-234 and proline-253 in domain 2. A point that has not been widely appreciated is that the conformation of PCNA changes upon binding a PIP motif. Comparing X-ray crystal structures of PCNA in the presence and absence of a bound PIP motif shows that the backbone of isoleucine-128 moves by ˚ and the side chain of this residue moves by
5 A˚. This conformational
4 A change is necessary in order to accommodate the aromatic residue in position 8 in the PIP motif. Recently greater attention has been paid to contacts that PCNAinteracting proteins make with PCNA that occur outside of the canonical PIP motif. For example, the X-ray crystal structure of full-length flap endonuclease 1 (FEN1) bound to PCNA shows that this is a bipartite interaction [40]. FEN1 has a PIP motif on its C-terminal tail that binds to the front face of PCNA in the usual fashion. In addition, the core domain of

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FEN1 directly contacts domain 2 of PCNA (Fig. 1C). Three different conformations of the PCNA-bound FEN1 are observed in the crystal structure. In one of these states, the active site is oriented away from where the DNA would be positioned. It has been argued that this represents an inactive state of FEN1 and that a large rotation of the core domain of FEN1 about a flexible hinge region is necessary to achieve the active conformation. Thus, these additional contacts not only increase affinity of PCNA-interacting proteins for PCNA, but they also can play an important role in regulating these proteins. Posttranslational modifications of PCNA are critical events in the regulation of the DNA metabolic processes in which PCNA participates. These modifications change the binding specificity of PCNA and in some cases act to recruit specific PCNA-interacting proteins to replication forks. The best characterized of these PCNA posttranslational modifications are ubiquitylation and sumoylation of lysine-164 [33,34], and X-ray crystal structures of these modified forms of PCNA have been determined (Fig. 1D) [41,48,53]. Ubiquitylation of PCNA at this position promotes translesion synthesis by recruiting translesion synthesis polymerases, which themselves contain PIP motifs as well as ubiquitin-binding motifs in their C-terminal tails [35]. Sumoylation of PCNA at this position inhibits recombination by recruiting an antirecombinase [36,37], which contains a PIP-like motif that binds on the front face of PCNA as well as a SUMO-binding motif [53].

3. THE ROLE OF PCNA IN NORMAL DNA REPLICATION In eukaryotes, DNA replication is an extraordinarily complex, dynamic, multistage process that initiates at origins of replication [54–57]. Before an origin can fire, it must be licensed. Origin licensing occurs during late M phase and early G1 phase, when the pre-replication complex (preRC) forms at the origin. The pre-RC includes Cdt1, Cdc6, and two hexamers of the Mcm2–7 helicase. Origin firing occurs at the onset of S phase. At this point, the origin is melted by the Mcm2–7 helicases, and the resulting single-stranded DNA is coated by replication protein A (RPA). Two replication forks are then assembled at the origin. First, Cdc45 and go-ichi-ni-san (GINS) complexes are recruited, which together with the Mcm2–7 hexamers form two Cdc45/MCM2–7/GINS (CMG) complexes, the replicative helicase complexes. Next, other protein factors are recruited to the nascent replication forks, including RFC, PCNA, pol δ, and pol ε. Finally, DNA synthesis begins as the replication forks move away from the origin bidirectionally.

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It is critical that each origin fires only once per cell cycle, and PCNA plays an important role in limiting each origin to firing once. Cdt1, a component of the pre-RC, is degraded in S phase in a PCNA-dependent manner [32,58,59]. Cdt1 contains a specialized PIP motif called a PIP degron, which contains a threonine at position 5, an aspartate at position 6, and a basic amino acid located four residues following the PIP motif [59]. These specialized PIP motifs bind PCNA with greater affinity than do classical PIP motifs. In the case of Cdt1, this PIP degron is responsible for making the protein a substrate for the E3 ubiquitin ligase CLR4Cdt2. When Cdt1 is bound to a PCNA ring that has been loaded on DNA, CLR4Cdt2 facilitates the polyubiquitylation of Cdt1 leading to its degradation. This ensures that Cdt1 is not available to relicense origins once they have fired. There is a division of labor among the replicative DNA polymerases at the eukaryotic replication fork. The leading strand is synthesized in a continuous manner by pol ε, and the lagging strand is synthesized in a discontinuous manner by pol δ [60,61] (Fig. 2). Biochemical studies using purified, reconstituted systems have shown that PCNA interacts with and affects DNA synthesis by pol δ and pol ε differently [62]. For example, pol δ binds PCNA with high affinity, whereas pol ε binds PCNA with low affinity. Pol δ alone synthesizes DNA with low processivity, only incorporating up to six nucleotides before dissociating from the DNA template. In the presence of PCNA, the processivity of pol δ increased nearly 100-fold. By contrast, pol ε has a greater intrinsic processivity, incorporating approximately

Fig. 2 The replication fork. A representation of the replication fork is shown with the leading strand on the bottom and the lagging strand on the top. PCNA is gray, pol ε is red (dark gray in the print version), pol δ is orange (gray in the print version), RPA is purple (black in the print version), and the CMG complex is green (light gray in the print version).

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60 nucleotides before dissociating. The presence of PCNA increases the processivity of pol ε by about sixfold. Overall, on PCNA-primed and RPA-coated single-stranded DNA, pol δ and pol ε have nearly the same processivity, incorporating up to 600 nucleotides per DNA-binding event. The differential interactions between these polymerases and PCNA are partly responsible for selecting the appropriate polymerases for leading and lagging strand synthesis [63]. PCNA strongly favors extension by pol δ over extension by pol ε in competition experiments in vitro on RPA-coated single-stranded DNA, a situation analogous to lagging strand DNA synthesis. This preference for pol δ over pol ε is reversed, however, in the presence of the CMG helicase complex [63], a situation analogous to leading strand DNA synthesis. The pol δ–CMG complex synthesizes DNA about 5- to 10-fold slower than does the pol ε–CMG complex. In addition, the CMG complex selectively utilizes pol ε in competition experiments, and pol ε readily replaces pol δ from an actively extending pol δ–CMG complex. Thus PCNA, together with the CMG complex, maintains the division of labor between pol δ and pol ε at the replication fork. On the leading strand, only one PCNA ring needs to be loaded, and this occurs when the origin fires and the replication fork is assembled. On the lagging strand, by contrast, one PCNA ring needs to be loaded for each Okazaki fragment. An Okazaki fragment is initiated by DNA polymerase alpha (pol α), which has an associated primase that synthesizes a short RNA primer [64,65]. Pol α then extends this RNA primer by 10–20 nucleotides of DNA. Next, PCNA is loaded onto DNA by RFC [17,18,66]. RFC binds the primer–template junction synthesized by pol α and catalyzes the loading of PCNA in an ATP-dependent manner. RFC binds to the front of the PCNA ring and loads it with the front of the ring facing toward the 30 end of the primer strand [67]. This ensures that polymerases and other PCNA-interacting enzymes will have access to the primer terminus. Once PCNA is loaded onto the primer strand, pol δ is recruited to synthesize the remainder of the Okazaki fragment. PCNA also orchestrates the events on the lagging strand during the maturation of each Okazaki fragment. This process occurs when pol δ encounters the 50 end of the previous Okazaki fragment. Pol δ displaces the 50 end of the fragment containing the RNA primer and a segment of the DNA, creating a flap. FEN1 contains a PIP motif and is recruited to the maturing Okazaki fragment via its interaction with PCNA. FEN1 catalyzes cleavage of the flap structure to create a nicked duplex, an activity that is stimulated in the

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presence of PCNA [68]. DNA ligase I also contains a PIP motif and is recruited via its interaction with PCNA. DNA ligase I catalyzes the sealing of the nick, an activity that is also stimulated by PCNA [69]. Exactly how these sequential enzymatic activities are coordinated by PCNA remains an important unanswered question. In the yeast system, PCNA is sumoylated during S phase on lysine-164 by the complex of the E2 SUMO-conjugating enzyme Ubc9 and the E3 SUMO ligase Siz1 [33,34]. It has been estimated that the majority of DNA-bound PCNA is sumoylated during normal DNA replication [70]. Sumoylation of PCNA inhibits recombination by recruiting an antirecombinase Srs2 [36,37], which contains a PIP-like motif that binds on the front face of PCNA as well as SUMO-binding motif [53]. Srs2 acts by disrupting the formation of Rad51 nucleoprotein filaments [71,72], an active species in homologous recombination. PCNA is also sumoylated to a lesser extent on lysine-127, which is located in the interdomainconnecting loop, although the biological significance of this modification is unknown.

4. THE ROLE OF PCNA IN TRANSLESION SYNTHESIS DNA damage causes replication forks to stall, because classical DNA polymerases, such as pol δ and pol ε, are unable to efficiently incorporate deoxynucleotides opposite damaged DNA templates. Without a means of overcoming these replication blocks, replication forks collapse resulting in DNA strand breaks, chromosomal rearrangements, and cell death. Translesion synthesis is one of the means by which damaged DNA is bypassed during DNA replication. During translesion synthesis, one or more nonclassical DNA polymerases, such as DNA polymerase zeta (pol ζ), DNA polymerase eta (pol η), DNA polymerase iota (pol ι), DNA polymerase kappa (pol κ), and Rev1, are recruited to stalled replication forks [73–79]. The mechanisms of these nonclassical polymerases differ from those of their classical counterparts so that they are able to incorporate deoxynucleotides opposite damaged DNA with high efficiency. In addition, these nonclassical polymerases incorporate nucleotides with very low fidelity, so translesion synthesis is generally error-prone. When DNA is damaged, a single ubiquitin moiety is attached to lysine-164 on one or more of the PCNA subunits by the complex of the E2 ubiquitin-conjugating enzyme Rad6 and the E3 ubiquitin ligase Rad18 [33]. The resulting ubiquitin-modified PCNA (Ub-PCNA) plays

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important regulatory roles in translesion synthesis. It acts as a hub protein for recruiting the nonclassical polymerases to stalled replication forks. Many nonclassical polymerases, including pol η, pol κ, and pol ι, possess tandem ubiquitin-binding motifs and PIP motifs [35]. These motifs are important for allowing nonclassical polymerases to preferentially interact with Ub-PCNA over unmodified PCNA and to colocalize with Ub-PCNA in cells [35,80–84]. Structural and computational studies have shown that the ubiquitin moiety of Ub-PCNA is dynamic, yet predominantly occupies preferred orientations on the back and on the side of the PCNA ring [48,85,86]. This would allow Ub-PCNA to regulate the access of nonclassical polymerases to the primer terminus by altering the orientation of its attached ubiquitin. For example, nonclassical polymerases can be held in reserve without affecting the activity of enzymes bound on the front face of the PCNA ring when the ubiquitin moiety is on the back of the PCNA ring. Nonclassical polymerases can gain access to the primer terminus when the ubiquitin moves to the side of the PCNA ring. In addition to recruiting nonclassical polymerases to stalled replication forks, PCNA and Ub-PCNA both regulate the catalytic activity of nonclassical polymerases. The catalytic efficiencies of pol η, pol κ, and pol ι are increased in the presence of PCNA [23–26]. In the case of pol η and the nonclassical polymerase Rev1, the catalytic efficiency of nucleotide incorporation is increased more in the presence of Ub-PCNA than in the presence of unmodified PCNA [48,87,88]. By contrast, although unmodified PCNA does stimulate the activity of pol ζ, Ub-PCNA does not stimulate it to a greater extent [87]. Two separation-of-function mutations were identified in yeast PCNA that block translesion synthesis in cells [89,90] (Fig. 3). Both of these substitutions (G178S and E113G) are of residues in the β strands that constitute the subunit interface, and the X-ray crystal structures of both mutant proteins reveal perturbations that reduce the number of hydrogen bonds between these strands [91,92]. The mutant PCNA trimers are less stable than wild-type PCNA, and they are unable to stimulate the catalytic activity of nonclassical polymerases [88]. There are two possible explanations for the inability of these mutant proteins to stimulate these polymerases. First, decreased PCNA trimer stability may lead to breathing (ie, transient opening and closing of the ring) that disrupts the activity of nonclassical polymerases to a greater extent than it does other PCNA-interacting enzymes. Second, the PCNA subunit interface may serve as an additional point of contact for

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Fig. 3 Separation-of-function mutations in PCNA. The locations of the separation-offunction mutations in yeast PCNA are shown from a front view. Mutations affecting translesion synthesis are blue (dark gray in the print version), mutations affecting errorfree damage bypass are red (dark gray in the print version), mutations affecting breakinduced replication are green (light gray in the print version), mutations affecting mismatch repair are yellow (white in the print version), and mutations affecting replication-coupled nucleosome assembly are orange (gray in the print version).

nonclassical polymerases. Further studies are necessary to distinguish between these possible scenarios.

5. THE ROLE OF PCNA IN ERROR-FREE DAMAGE BYPASS In addition to translesion synthesis, which is mutagenic, another pathway for circumventing DNA lesions in the template strand during DNA replication is error-free damage bypass. The detailed mechanism of errorfree damage bypass has yet to be elucidated, but it is believed to involve a template-switching event whereby the replicative DNA polymerases moves to the newly synthesized sister strand and uses it as a template [93,94]. The model for how this template-switching event occurs is that a forkremodeling enzyme catalyzes regression of the stalled replication fork (Fig. 4). In yeast, this is carried out by the Rad5 helicase, and in mammals this is carried out by the Rad5 homologs HLTF and SHPRH. Fork regression leads to the formation of a “chicken foot” intermediate in which the primer terminus of the nascent strand is paired with the newly synthesized sister strand. Extension of the nascent strand to the 50 end of the sister strand and subsequent restoration of the replication fork result in bypass of the DNA damage and resumption of normal DNA replication. Although the detailed mechanism of error-free damage bypass is not yet known, it is clear that PCNA plays a central regulatory role in this process.

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3′

3′

Fig. 4 Error-free damage bypass. A schematic of a stalled replication fork is shown. The leading strand is blue (gray in the print version), the lagging strand is red (dark gray in the print version), and the location of the DNA damage is indicated by a red square (dark gray in the print version). The stalled replication fork is converted into the chicken foot intermediate, the chicken foot intermediate is extended, and the replication fork is then reestablished.

As was the case in translesion synthesis, the initiating event in error-free damage bypass is the attachment of a single ubiquitin moiety to lysine164 on one or more of the PCNA subunits by Rad6 and Rad18 to form Ub-PCNA [33]. Next, the complex of the E2 ubiquitin-conjugating enzymes Ubc13/Mms2 and the E3 ubiquitin ligase Rad5 are recruited to Ub-PCNA. (Rad5 functions in error-free damage bypass both as an E3 ubiquitin ligase and as a fork-remodeling helicase.) This results in the formation of lysine 63-linked polyubiquitin chains on PCNA [33]. The attachment of lysine 63-linked polyubiquitin chains to PCNA is required for error-free damage bypass, but it is unknown how it facilitates the template-switching process. Recent studies are providing some interesting clues. The presence of these polyubiquitin chains on PCNA decreases the formation of complexes between pol δ and PCNA, which prevents normal DNA replication from occurring [95]. It also decreases the ability of nonclassical pol η to bypass DNA lesions, which prevents translesion synthesis. This latter effect is not due to decreased binding of pol η, but rather may be due to pol η being trapped in a nonproductive complex by the polyubiquitin chains. Thus error-free DNA damage bypass may be facilitated by the polyubiquitylation of PCNA inhibiting the alternative pathways. Two separation-of-function mutations were identified in yeast PCNA that block error-free damage bypass [96] (Fig. 3). Cells producing the E104A/D105A or the D256A/E257A PCNA mutant proteins are more sensitive to ultraviolet radiation. Glutamate-104 and aspartate-105 are located on a loop immediately adjacent to the subunit interface, and aspartate-256 and glutamate-257 are located at the C-terminus of the protein. Cells producing the quadruple mutant protein, which has all four of these amino acid substitutions, are more sensitive to ultraviolet radiation than those expressing the other two mutant proteins. In these cells, error-free

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damage bypass is blocked, but error-prone translesion synthesis is unaffected. This latter point suggests that the ability of this mutant protein to be ubiquitylated is not compromised, but this has not been confirmed experimentally. The structural changes in PCNA caused by these substitutions are unknown, and the mechanistic basis for the mutant protein’s inability to support error-free damage bypass is not yet understood. Error-free damage bypass is an active area of research, and several interesting questions remain regarding the role of PCNA in this process. First, does the polyubiquitylation of PCNA signal for the recruitment of other protein factors necessary for fork remodeling? Recently, the genome maintenance factor Mgs1 has been suggested as a potential downstream effector of ubiquitylated PCNA that may play a role in this process [97]. Second, once the chicken foot intermediate is formed, is another PCNA ring loaded onto the middle branch of this structure? If so, this would nicely explain how pol δ is recruited to this new primer–template to carry out extension to the end of the newly synthesized sister strand.

6. THE ROLE OF PCNA IN BREAK-INDUCED REPLICATION If translesion synthesis and error-free damage bypass fail to allow the resumption of DNA replication, several other pathways may be used to restart the stalled replication fork. These pathways generally involve the use of the recombination machinery and are beyond the scope of this review; these pathways are described in detail elsewhere [98–100]. However, one such pathway, break-induced replication, does warrant attention. This is because PCNA is required for break-induced replication, and separationof-function mutations in PCNA have been identified that block breakinduced replication without significantly affecting normal replication and other recombination-dependent replication restart pathways [101]. Break-induced replication is used to repair one-sided double-strand breaks (Fig. 5) [100,102–105]. One-sided breaks are formed during DNA replication in several ways. They are formed when a replication fork encounters a single-stranded nick in the template strand. They are also formed when the chicken foot intermediate of a stalled replication fork, which is actually a Holliday junction, is cleaved by a junction-specific endonuclease. Break-induced replication proceeds by processing of the one-sided break in order to form a Rad51 nucleoprotein filament. Strand exchange occurs with the intact sister duplex resulting in the formation of a D-loop, a key recombination intermediate. DNA synthesis starts from the 30 end of

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Fig. 5 Break-induced replication. A schematic of a replication fork with a nick in the leading strand template is shown. The leading strand is blue (gray in the print version), and the lagging strand is red (dark gray in the print version). This gives rise to a one-sided break. A schematic of a chicken foot intermediate is shown. Resolution of this four-way junction by cutting at the indicated sites also gives rise to a one-sided break. The onesided break is converted into a D-loop, the D-loop is extended, and the replication fork is then reestablished.

the invading strand, and this will become the leading strand of a reestablished replication fork. The lagging strand is likely initiated subsequently, and replication proceeds until the end of the replicon or the end of the chromosome. Break-induced replication requires all of the proteins needed for normal DNA replication, except those specific for formation of the preRC complex [101]. Two separation-of-function mutations were identified in yeast PCNA that block break-induced replication in cells [101] (Fig. 3). Unlike other separation-of-function mutations in PCNA, both of these substitutions (F248A/F249A and R80A) are dominant. This means that only one mutant subunit in a PCNA trimer is sufficient to inhibit break-induced replication. The F248A/F249A substitution is in a β strand within the cleft formed by the two domains of the same monomer. It is near the binding site for PIP motifs on PCNA-interacting proteins, but is likely not close enough to directly contact these motifs. The R80A substitution is on an extended loop near the subunit interface. It is not known how these amino acid substitutions interfere with break-induced replication. These same amino acid substitutions, however, suppress the cold sensitivity of yeast lacking the nonessential Pol32 subunit of pol δ, a subunit that is also essential for breakinduced replication [101]. Moreover the decreases in break-induced

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replication caused by these amino acid substitutions in PCNA are epistatic with the decrease in break-induced replication caused by the absence of the Pol32 subunit of pol δ [101]. This suggests that these two mutant forms of PCNA interact with pol δ in an aberrant manner that interferes with the ability of its Pol32 subunit to perform its essential role in break-induced replication, whatever that may be.

7. THE ROLE OF PCNA IN MISMATCH REPAIR Replicative polymerases make errors when synthesizing DNA that can lead to base–base mismatches or short insertions and deletions. These errors are recognized and corrected by mismatch repair [106–112]. The first step of mismatch repair is recognition of the mismatches or insertions/deletions in the newly synthesized DNA. This involves either the MutSα heterodimer composed of Msh2 and Msh6 or the MutSβ heterodimer composed of Msh2 and Msh3. These mismatch recognition complexes have partially overlapping specificities with MutSα preferring base–base mismatches and small insertions/deletions and with MutSβ preferring larger insertions/ deletions. The next step of mismatch repair is excision of the mismatch and surrounding DNA from the newly synthesized strand. This requires the MutLα heterodimer composed of Mlh1 and Pms1 in yeast (PMS2 in humans) and the exonuclease ExoI. The final step of mismatch repair is synthesis of new DNA by pol δ to fill the gap. PCNA is required for all three steps of mismatch repair [21,113–116]. It plays an essential role in recognizing the mismatch by interacting with MutSα to form an active mismatch recognition complex [114]. Mutations in MutSα that disrupt its interaction with PCNA lead to a loss of mismatch repair in cells. PCNA plays an important role in excising the mismatch and surrounding DNA from the newly synthesized strand by binding MutLα and activating its latent endonuclease activity [115] and by binding ExoI and regulating its exonuclease activity [116]. Finally, PCNA plays a critical role in repair synthesis by interacting with pol δ. Two separation-of-function mutations in PCNA (C22Y and C81R) have been identified that inhibit mismatch repair in yeast [114] (Fig. 3). The crystal structures of these mutant proteins showed distinct structural alterations caused by these two substitutions [117]. Cysteine-22 is in a loop adjacent to one of the α-helices lining the central hole of the PCNA ring, and the C22Y mutation causes a significant shifting of several of these α-helices. Cysteine-81 is in an extended loop near the subunit interface,

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and the C81R mutation induces a change in the conformation of this loop leading to a slight destabilization of the PCNA trimer. Both of these mutant proteins bind MutSα, but the ternary complex formed by these mutant proteins, MutSα and DNA is abnormally large. Although the exact nature of these aberrant complexes is unknown, it is likely that this is responsible for the defect in mismatch repair. Recently, additional mutations in PCNA that block mismatch repair were identified. Three of these mutations were clustered around cysteine-22, and seven were near the subunit interface destabilizing the PCNA trimer [118]. These mutant PCNA proteins likely have the same structural perturbations as noted for the C81R and the C22Y mutant proteins, respectively. Arguably the most important unanswered question about mismatch repair in eukaryotes is how the newly synthesized strand and the template strand are distinguished so that only the newly synthesized strand is excised. The close coupling of the mismatch repair machinery and the replication machinery may allow strand discrimination to be achieved through several means. Nicks in the newly synthesized strand such as those occurring between Okazaki fragments on the lagging strand or those resulting from RNase H2-catalyzed removal of ribonucleotides are important for strand discrimination [119,120]. Because PCNA is loaded at the replication forks in a precise orientation with the front of the ring facing the direction of DNA synthesis, it may play an important role in strand discrimination. Evidence for this comes from the fact that not only does PCNA activate the latent exonuclease activity of MutLα, but it also makes it specific for nicking the newly synthesized strand [121].

8. THE ROLE OF PCNA IN REPLICATION-COUPLED NUCLEOSOME ASSEMBLY Immediately following DNA replication, nucleosomes must be assembled on the newly synthesized daughter duplexes behind the replication fork. In transcriptionally silent, heterochromatic regions of the genome including the centromeric and telomeric regions of chromosomes, this is carried out in part by chromatin association factor-1 (CAF-1). CAF-1 is a heterotrimer comprising Cac1, Cac2, and Cac3 subunits. It functions as a histone H3–H4 chaperone that catalyzes the deposition of nucleosomes onto newly synthesized DNA during S phase. The Cac1 subunit of CAF-1 contains a PIP motif that mediates its interaction with PCNA.

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Genetic studies in yeast showed that PCNA plays an essential role in CAF-1-mediated nucleosome assembly [31]. Three separation-of-function mutations in PCNA (D41A/D42A, R61A/D63A, and L126A/I128A) were identified that were defective in gene silencing at telomeric regions and mating-type loci. Aspartate-41 and aspartate-42 are close to the hydrophobic pocket on PCNA where PIP motifs bind, and leucine-126 and isoleucine-128 are on the interdomain connector loop near this binding pocket. Arginine-61 and glutamate-63, by contrast, are on a loop on the back face of the PCNA ring. Although it is not known what structural alterations in PCNA are caused by these substitutions, the mutant proteins are all defective in binding CAF-1 in vitro [31]. This suggests that the interaction with PCNA acts to target CAF-1 to newly synthesized DNA.

9. PUTTING IT ALL TOGETHER One of the most important, unanswered questions regarding PCNA’s role in DNA replication is how does it interact with and regulate the activity of so many binding partners. One widely discussed idea is that PCNA can function as a “tool belt” by binding several partners simultaneously. Because PCNA is a trimer, it can potentially bind up to three PIP motif-containing proteins at the same time. Although there is so far no direct evidence that eukaryotic PCNA forms such tool belts, archaeal PCNA has been shown to form tool belts. PCNA from the thermophile Sulfolobus solfataricus P2 is a heterotrimer that can simultaneously bind the DNA polymerase, the flap endonuclease, and the DNA ligase [122]. Given this, it is very likely that PCNA tool belts are also formed in eukaryotes. Even if tool belts form, however, there are still far too many proteins that need to interact with PCNA than can be accommodated by the PCNA rings loaded on the leading strand and lagging strand at the replication fork. A possible answer to this question emerges when one considers a number of seemingly disparate facts. First, it has been widely appreciated that replication occurs in eukaryotic cells in discrete regions of the nuclei known as replication foci or replication factories [123]. Immunofluorescent imaging of PCNA shows that in early S phase, there are roughly 1200 replication factories, each containing as few as two replication forks, in the nuclei of human cells [124]. These replication factories are 150 nm in diameter on average and contain multiple PCNA molecules. Second, many PCNA-interacting proteins have their PIP motifs within intrinsically disordered regions often near their N-termini or C-termini [125]. This allows these proteins to bind

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PCNA via flexible tethers without their folded domains remaining in the immediate vicinity of PCNA or being otherwise geometrically constrained. Third, some PCNA-interacting proteins contain multiple PIP motifs often within the same flexible tethers [126]. Multiple PIP motifs in the same protein would allow such a protein to simultaneously interact with two or more different PCNA rings, thereby linking them together. Taken together, these observations suggest that there is a large, dynamic network of linked PCNA molecules at and around the replication fork (Fig. 6). Some of the PCNA molecules will be loaded on the DNA, perhaps one per Okazaki fragment on the lagging strand. Others will not be loaded on the DNA, but linked to the DNA by a flexible meshwork of protein– protein interactions. Overall, a single replication factory with one or two replication forks at its core may contain hundreds of PCNA molecules and hundreds of PCNA-binding proteins surrounding the forks, all linked through a network of protein–protein interactions. Such a network would serve to increase the local concentration of all the proteins necessary for normal replication, translesion synthesis, error-free damage bypass, breakinduced replication, mismatch repair, and chromatin assembly and regulate their various activities during DNA replication.

Fig. 6 A replication factory. A representation of a single replication factory is shown containing two replication forks drawn to scale. Many PCNA molecules (gray) are shown on the DNA (black lines) as well as off the DNA. These PCNA molecules are linked together by many PCNA-interacting proteins (various colors; different gray shades in the print version) to form a large, flexible network surrounding the replication forks. The various colored (different gray shades in the print version) lines depict the intrinsically disordered regions of these PCNA-interacting proteins.

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ACKNOWLEDGMENTS We thank Christine Kondratick, Kyle Powers, Brittany Ripley, Lynne Dieckman, and Maria Spies for their valuable discussions. This work was supported by National Institute of Health Grants R01 GM108027, R01 GM081433, and T32 GM067795.

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CHAPTER EIGHT

Animal Mitochondrial DNA Replication G.L. Ciesielski*,†,1, M.T. Oliveira{,1, L.S. Kaguni*,†,2 *Institute of Biosciences and Medical Technology, University of Tampere, Tampere, Finland † Michigan State University, East Lansing, MI, United States { Faculdade de Ci^encias Agra´rias e Veterina´rias, Universidade Estadual Paulista “Ju´lio de Mesquita Filho”, Jaboticabal, SP, Brazil 2 Corresponding author: e-mail address: [email protected]

Contents 1. Overview 2. Structure–Function Relationships in mtDNA Replication Proteins 2.1 DNA Polymerase γ, the Mitochondrial Replicase 2.2 The Replicative mtDNA Helicase 2.3 mtSSB 2.4 Functional Interactions Among Replisome Proteins and Evolutionary Implications 3. Mechanisms of Mitochondrial DNA Replication In Vivo 3.1 The Rolling Circle Model in C. elegans 3.2 The Theta Model in D. melanogaster 3.3 The Theta Model in Sea Urchin 3.4 mtDNA Replication in Vertebrates 4. Perspectives Acknowledgments References

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Abstract Recent advances in the field of mitochondrial DNA (mtDNA) replication highlight the diversity of both the mechanisms utilized and the structural and functional organization of the proteins at mtDNA replication fork, despite the relative simplicity of the animal mtDNA genome. DNA polymerase γ, mtDNA helicase and mitochondrial singlestranded DNA-binding protein—the key replisome proteins, have evolved distinct structural features and biochemical properties. These appear to be correlated with mtDNA genomic features in different metazoan taxa and with their modes of DNA replication, although substantial integrative research is warranted to establish firmly these links. To date, several modes of mtDNA replication have been described for animals: rolling circle, theta, strand-displacement, and RITOLS/bootlace. Resolution of a 1

These authors contributed equally to this work.

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continuing controversy relevant to mtDNA replication in mammals/vertebrates will have a direct impact on the mechanistic interpretation of mtDNA-related human diseases. Here we review these subjects, integrating earlier and recent data to provide a perspective on the major challenges for future research.

1. OVERVIEW The discovery of mitochondrial DNA established the unique character of mitochondria as the only organelles in the animal cell with an “extrachromosomal” genome [1]. Its compact structure and organization engendered much interest in the study of its replication, expression, inheritance, and evolution, and with the identification of pathogenic and heritable mutations that result in human disease, the field of mitochondrial medicine emerged [2–4]. We focus our attention here on the proteins and mechanisms involved in animal mtDNA replication, keeping in mind the essential and dynamic nuclear–mitochondrial interactions that drive its evolution. Indeed, mtDNA per se evolves at a relatively high rate as compared to the nuclear genome, and despite its similar general structure and organization, it varies even among closely related animal species, such as higher primates [5,6]. Unveiling the mechanisms by which mtDNA sequence variation is introduced, inherited, and fixed, and their relationship to mtDNA replication and repair provide a major challenge for future research.

2. STRUCTURE–FUNCTION RELATIONSHIPS IN mtDNA REPLICATION PROTEINS Three nuclear-encoded proteins play key roles at the mtDNA replication fork (Fig. 1): DNA polymerase γ (Pol γ), the replicative mtDNA helicase Twinkle, and the mitochondrial single-stranded DNA-binding protein (mtSSB). Together, these proteins are sufficient for effective DNA synthesis in vitro, and their knockdown or ablation in vivo results in replication defects and mtDNA depletion (reviewed in Refs. [10–12]). The replicative mtDNA helicase catalyzes dsDNA unwinding at the fork that is driven by NTP hydrolysis, releasing single-stranded DNA that is stabilized and protected by mtSSB, and is used subsequently as template by Pol γ for DNA synthesis. In this section, we review the biochemical properties of the proteins and their implications in human health and disease, with a final subsection on features relevant to various mtDNA replication systems.

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Fig. 1 Proteins at the mitochondrial DNA replication fork. The crystal structure of the heterotrimeric human Pol γ bound to primer-template DNA (PDB: 4ZTZ; [7]), the model of the homotetrameric human mtSSB wrapped around ssDNA [8], and the model of the ring-shaped heptameric human mtDNA helicase [9] were used to create a representation of the nuclear-encoded proteins that function at the mtDNA replication fork. Pol γ-α, the proximal Pol γ-β, and the distal Pol γ-β are shown in pink (gray in the print version), and light and dark gray, respectively; mtDNA helicase and mtSSB are shown in green (gray in the print version) and cyan (light gray in the print version), respectively. The diagram is to scale, but not meant to depict specific protein–protein interactions. Structures and models were analyzed and the figure was produced using Pymol (www. pymol.org).

2.1 DNA Polymerase γ, the Mitochondrial Replicase Three catalytic activities have been ascribed to Pol γ: 50 –30 DNA polymerase, 30 –50 exonuclease, and 50 -dRP lyase (reviewed in Refs. [10,13]). All are contained in its catalytic subunit, Pol γ-α (or POLGA, encoded by the POLG gene), although the residues required for its lyase activity have not yet been identified. These activities are regulated by its accessory subunit Pol γ-β (or POLGB, encoded by the POLG2 gene) (Fig. 2), which itself has no catalytic activity. However, its effects on DNA synthesis are substantial: it enhances DNA and nucleotide binding, stimulating DNA synthesis by Pol γ-α, and increasing the processivity of the holoenzyme 100-fold. Pol γ is highly accurate in nucleotide polymerization, with an in vitro error rate of only 1 misincorporated nucleotide per half-million bases polymerized.

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Fig. 2 Structure of human Pol γ bound to primer-template DNA and nucleotide. (A) Representation (to scale) of domains and subdomains in the Pol γ-α and β polypeptide sequences. NTD, N-terminal domain; Exo, exonuclease domain; AID, accessoryinteracting determinant subdomain; IP, intrinsic processivity subdomain; Pol, DNA polymerase domain; T, the bipartite thumb subdomain. (B) Cartoon representation of the crystal structure of the heterotrimeric human Pol γ holoenzyme bound to primertemplate DNA and nucleotide (PDB: 4ZTS; [7]). The Pol γ-α domains are colored (different shades of gray in the print version) as shown in (A); the proximal and distal Pol γ-βs are shown in light and dark gray, respectively. (C) Enlargement of the pol active site, showing conserved residues for Mg2+ and incoming nucleotide-binding, and the identical positioning of dCTP (left panel, PDB: 4ZTZ) and the inhibitor ddCTP (right panel, PDB: 4ZTU) [7].

However, replicative bypass of abasic sites and sites of oxidative damage by Pol γ is error prone. The high base-substitution fidelity of Pol γ is also compromised by nucleotide imbalances, a situation that may be physiologically relevant to the normal fluctuation of metabolites known to occur in the

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mitochondrial matrix, as well as pathogenic defects in the nuclear genes involved in nucleotide synthesis and transport to mitochondria. The high base-substitution fidelity in human Pol γ derives both from the highnucleotide selectivity of its polymerase (pol) domain and from exonucleolytic proofreading catalyzed by its exonuclease (exo) domain. Unlike numerous nuclear animal DNA polymerases, Pol γ displays a remarkable ability to utilize diverse primer-template substrates, most likely because it is involved in all processes of DNA metabolism in mitochondria (reviewed in Ref. [10]). Pol γ is the most extensively studied protein of the mtDNA replisome, and its biomedical relevance is well documented: pathogenic mutations in the genes encoding both the catalytic and accessory subunits have been identified in association with numerous human diseases (reviewed in Ref. [13]); impaired proofreading has been shown to cause premature aging in mammalian models [14,15]; and the sensitivity of Pol γ to specific nucleoside analog reverse transcriptase inhibitors (NRTIs) used to treat HIV infection can result in mitochondrial toxicity (reviewed in Ref. [16,17]). Structure– function relationships in Pol γ have been forged by a combination of many years of biochemical study and recent advances in the determination of the 3D structure of the human apo-holoenzyme [18] and of the holoenzyme bound in a ternary complex with primer-template DNA together with either normal or derivatized nucleotides [7,19]. The heterotrimeric organization of the human replicase, comprising a single Pol γ-α and two Pol γ-β polypeptides, appears to be common to all vertebrate mtDNA polymerases [20]. Pol γ-α consists of three domains, arranged spatially to interact with the Pol γ-β dimer, and to facilitate the transition between its 50 –30 polymerase and 30 –50 exonuclease activities (Fig. 2). The pol domain carries the canonical “right-hand” fold common to DNA polymerases, formed by palm, fingers, and thumb subdomains. Notably, the thumb subdomain is bipartite, a feature that is apparently unique to Pol γ-α as compared to other Family A DNA polymerases, and makes extensive interactions with the proximal Pol γ-β protomer [18]. However, the major binding site for the accessory subunit is in the accessory-interacting determinant (AID) subdomain within the spacer domain (Fig. 2). The spacer domain also contains an intrinsic processivity (IP) subdomain not found in the homologous T7 gp5 DNA polymerase, and is implicated in the relatively high processivity of the catalytic subunit alone (reviewed in Ref. [10]). Interestingly, in the absence of DNA, Pol γ-α contacts the distal Pol γ-β protomer only by interaction of the Pol γ-α R232

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and Q540 residues with Pol γ-β E394 and R122, respectively [18]. The contact region expands greatly upon primer-template DNA binding due to conformational changes that rotate the dimeric Pol γ-β by 22 degrees toward the polymerase domain of the catalytic subunit [7,19]. The 3D structure of the DNA-bound Pol γ provides a mechanistic explanation for human disease-causing substitutions in both the catalytic and accessory subunits. For example, the deleterious effects of the Pol γ-β G451E mutation on subunit association and holoenzyme processivity [21,22] are apparent only in the ternary complex: upon DNA binding, the G451 residues in the proximal and distal Pol γ-βs interact with the hydrophobic L-helix and the R232 region of Pol γ-α, respectively [7]. Furthermore, the 3D structure enabled us to establish genotype–phenotype correlations that in turn have led to the development of a powerful pathogenicity prediction tool to evaluate the likely effects of chromosomal variants in the compound heterozygous form in which they are most often manifest in patients with POLG syndromes [23,24]. The 3D structures of Pol γ have also shed light on how the holoenzyme is affected by NRTIs and provide a strong basis for rational drug design. Fig. 2C shows that ddCTP, the antiviral agent known as zalcitabine, is stabilized in the pol active site by R943, K947, and Y951, and a bent conformation of the template imparted by Y955. This configuration is highly similar to that observed upon dCTP binding, providing a molecular basis for the susceptibility of Pol γ to this HIV reverse transcriptase inhibitor [7]. On the other hand, new classes of NRTIs, such as ()-FTC [()2,30 -dideoxy-5-fluoro-30 -thiacytidine, emtricitabine] causes much reduced mitochondrial toxicity because Pol γ discriminates against it more efficiently than does the viral reverse transcriptase [25]; the nucleotide-binding site in the pol domain of the mitochondrial replicase can distinguish the ribose and base modifications in ()-FTC as a result of the potential steric clashes between the rigid Y951 residue and the modified ribose. Moreover, the hydrophobic nature of the I948 residue precludes interactions with the 5-fluorine, forcing both the nucleoside and the α-phosphate of the inhibitor to be misaligned [19]. Less experimental focus has been placed on the proofreading exonuclease activity of Pol γ, although it is also implicated strongly in human disease and in aging. The exo domain of human Pol γ-α contains the highly conserved residue D274 (D257 in the mouse, D263 in Drosophila melanogaster). Its substitution with alanine results in mtDNA mutator mouse lines [14,15], and more recently a mutator fly line [26]. Whereas the mtDNA mutator mice have been analyzed extensively to show the substantial impact of a

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proofreading-deficient Pol γ in causing premature aging phenotypes and in shortening life span, the homozygous flies carrying the equivalent mutant enzyme surprisingly die during development in the late third larval instar stage, indicating potentially major differences in the requirements for the exonuclease activity of Pol γ in mtDNA maintenance in mammals and insects. Indeed, other major differences exist between the vertebrate and insect Pol γs, such as the heterodimeric structure of the latter [27,28].

2.2 The Replicative mtDNA Helicase Recently, we reviewed comprehensively the literature on the structure, catalytic activity, and evolution of the animal mtDNA helicase, including an evaluation of human pathogenic variants and a discussion of current animal models [9]. Among the important findings, two features of the replicative mtDNA helicase are notable: its remarkable resemblance to the bacteriophage T7 gp4, a bifunctional primase–helicase (see chapter “The Replication System of Bacteriophage T7” by Kulczyk and Richardson), and the numerous mutations in the human gene associated with mitochondrial disorders. Its primary sequence suggests a modular architecture as in T7 gp4 [29], comprising of a zinc-binding-like domain (ZBD), an RNA polymerase-like domain (RPD), a Linker region, and a C-terminal helicase domain (CTD) (Fig. 3). Molecular modeling and mapping of the diseaserelated residues reported to date in the human enzyme identified two major structural regions that we explored in detail. First, the abundance of pathogenic residues found in the Linker region and CTD argues that their effects on subunit interactions are a major cause of mtDNA replication defects leading to human disorders [9]. Oligomerization in replicative helicases is key to the formation of the nucleotide-binding pocket at the protomer–protomer interface that is required for proper positioning of the substrate for hydrolysis, which is coordinated with translocation of the helicase on DNA [31]. The putative role of these disease-related residues in maintaining the stability of the oligomeric mtDNA helicase is apparent only when two protomers are analyzed together, revealing that the Linker region of one protomer likely interacts with the CTD of the adjacent protomer. At present, several studies support such a hypothesis [32–35]. Second, pathogenic residues that map onto a human RPD model cluster on a positively charged surface area, which might represent a new DNAbinding region. If so, the RPD in mtDNA helicase binds DNA in a configuration not described for T7 gp4 or other prokaryotic primases [9]. Evidence

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Fig. 3 Schematic representation and structural model of the human mtDNA helicase. (A) Upper image, schematic representation of the conserved amino acid sequence motifs in human mtDNA helicase. MTS, mitochondrial target sequence; I–VI and R, conserved sequence motifs I–IV and RNAP basic motif of prokaryotic primases; 1–4 and 1a, conserved sequence motifs 1–4 and 1a of ring-shaped helicases [30]. The size and position of the conserved sequence motifs are represented to scale. Lower image, structural model of a protomer of the human mtDNA helicase highlighting its modular architecture organized in a zinc-binding-like domain (ZBD), RNA polymerase-like domain (RPD), Linker region, and a C-terminal helicase domain (CTD). *ZBD portion is represented as the polypeptide backbone of the bacteriophage T7 gp4 ZBD. (B) Model of the heptameric human mtDNA helicase, CTD view. (C) Electron microscopic image of the recombinant human mtDNA helicase at 100 mM NaCl in the presence of Mg2+ and ATPγS, showing its heptameric configuration. Reproduced with permission from L.S. Kaguni, M.T. Oliveira, Structure, function and evolution of the animal mitochondrial replicative DNA helicase, Crit. Rev. Biochem. Mol. Biol. 51 (1) (2016) 53–64 and from T.D. Ziebarth, R. Gonzalez-Soltero, M.M. Makowska-Grzyska, R. Núñez-Ramírez, J.M. Carazo, L.S. Kaguni, Dynamic effects of cofactors and DNA on the oligomeric state of human mitochondrial DNA helicase, J. Biol. Chem. 285 (2010) 14639–14647.

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suggesting the importance of this putative DNA-binding region includes the following: (1) expressing D. melanogaster mtDNA helicase bearing analogous mutations found in human patients causes severe mtDNA depletion in S2 cells [36], consistent with the fact that the recombinant N-terminal domain of the insect enzyme binds to both ss and dsDNA [37] and (2) recombinant human mtDNA helicases lacking part or all of the RPD exhibited lower ssDNA binding, ATPase and unwinding activities [38], in agreement with the observed decrease in mtDNA copy number in human cultured cells [39]. Interestingly, the RPD of the mtDNA helicase appears to have evolved these novel functions without losing either the polypeptide fold or some of the conserved amino acid residues of a prokaryotic primase, even though it does not synthesize RNA primers [9,38]. Whereas oligomerization clearly involves residues in the Linker region and CTD, both the oligomeric form and the conformation of individual protomers in the human mtDNA helicase appear to be dynamic [35,40,41]. A high proportion of structurally heterogeneous homohexamers with threefold symmetry was observed at high ionic strength with protein purified in the baculovirus expression system, which contrasts with a more homogeneous homoheptameric species with a clear sevenfold symmetry (Fig. 3C) observed at low ionic strength and in the presence of Mg2+ and ATPγS [40]. Interestingly, the opposite balance between forms is found for T7 gp4, which is predominantly homohexameric upon nucleotide triphosphate binding [42]. Because the heptamers of T7 gp4 are unable to bind DNA efficiently, it is proposed that the loss of a protomer during the switch from heptamers to hexamers promotes ring-opening for loading of the enzyme on ssDNA. By analogy, and considering evidence that the mtDNA helicase can load on a circular ssDNA template in the absence of a helicase loader [43], it seems plausible that ring-opening in the hexameric mtDNA helicase occurs by acquiring an extra subunit upon ATP binding, suggesting a role for the heptameric form in DNA loading, nucleotide hydrolysis, and dsDNA unwinding in mtDNA replication. Clearly, substantial research is warranted to test this hypothesis in vitro and in vivo. The most intriguing feature of the mtDNA helicase is perhaps the sequence and structural diversity of its ZBD among different species. Whereas the ZBD of T7 gp4 contains four cysteine residues that coordinate zinc that is essential for primase activity, it has been suggested that the absence of three of four cysteine residues in the ZBD of human (and other vertebrate) mtDNA helicases is the reason for their lack of primase activity [44]. Underscoring the structural differences among mtDNA helicases,

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we showed that a catalytic Mg2+-binding pocket in the RPD cannot be modeled in either the human or the D. melanogaster enzymes [9]. Although invertebrates do possess the conserved cysteine residues in the ZBD, physical characterization of the D. melanogaster ZBD showed that it binds an iron– sulfur cluster, coordinated by the homologous cysteine residues of T7 gp4 [37]. No evidence of zinc binding has been reported in human mtDNA helicase, and amino acid sequence analyses indicate that the ZBD in vertebrate mtDNA helicases has diverged significantly from their prokaryotic counterparts. Implying a potentially major shift in function, the evolutionary novelty of the iron–sulfur cluster makes the insect ZBD a candidate for sensing the redox state inside mitochondria, linking mtDNA replication to mitochondrial responses to excess reactive oxygen species. At present, it is not clear if the presence of an iron–sulfur cluster-binding domain is an ancestral condition in animal mtDNA helicases, or if it is a derived feature of the insect enzyme, but it is clearly absent in vertebrates, and could contribute to the diversification of mtDNA replication modes identified across animal taxa (see Section 3).

2.3 mtSSB Single-stranded DNA-binding proteins (reviewed in Ref. [45]) including mtSSB are essential in DNA metabolism, serving to bind and protect ssDNA during replication, repair, and recombination. At the mtDNA replication fork, mtSSB most likely coordinates interactions within the replisome, stimulating Pol γ and mtDNA helicase function. In vitro stimulation of the DNA polymerase and exonuclease activities of Pol γ by mtSSB has been documented in both the human and D. melanogaster systems [8,46–49]. Human mtSSB stimulates both the unwinding activity of the human mtDNA helicase up to eightfold [8,48,50], and the concerted actions of the human Pol γ and mtDNA helicase in strand-displacement DNA synthesis [51]. The role of mtSSB in mtDNA replication has also been documented genetically in cultured cells and in whole animals. Disruption of the D. melanogaster gene (lopo) resulted in developmental lethality, accompanied by severe mtDNA depletion and loss of cellular respiratory capacity [52]. Interestingly, Sugimoto et al. showed that viability of the nematode Caenorhabditis elegans is not affected by systemic mtSSB knockdown, but mtDNA levels and animal fertility are reduced drastically [53]. mtDNA depletion is consistently observed when mtSSB protein is knocked down by RNAi either in Drosophila Schneider cells [8,54,55] or in human HeLa cells [56].

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As a homologue of the well-studied SSB from Escherichia coli (EcSSB), sequence analysis indicates that the mtSSB of all metazoans is likely homotetrameric and binds to ssDNA similarly to the bacterial protein (Fig. 4). We retrieved 56 animal homologue sequences of the human and D. melanogaster proteins from diverse databanks, making structural predictions and drawing functional correlations, as recently reported for Pol γ [20] and the mtDNA helicase [9]. Because animal mtSSB sequences are highly similar, we also evaluated them using the Evolutionary Trace Server [63,64] to identify important amino acid residues. These are highlighted in yellow in Fig. 4A–C as those conserved only in metazoans. A comparison with the EcSSB residues that are known to be crucial for ssDNA binding and subunit interactions (red and light green arrows, respectively) reveals a remarkable overlap. This is consistent with the long-standing concept that bacterial and mitochondrial SSBs are structurally and functionally similar, especially in regard to ssDNA-binding activity [8,62,65]. The only residues identified by the Evolutionary Trace analysis that do not appear to be involved directly in DNA binding are D52 and D105 (in reference to the human sequence) (Fig. 4B and C). Interestingly, we have shown that deletion of the loop containing D52 (loop 2,3), and alanine substitutions in the loop containing D105 (loop 4,5) of human mtSSB do not affect substantially its ssDNAbinding affinities, but instead compromise its ability to stimulate either human Pol γ or mtDNA helicase [8] (see Section 2.4).

2.4 Functional Interactions Among Replisome Proteins and Evolutionary Implications The protein components of the minimal mtDNA replisome have distinct evolutionary origins, prompting the question of how they have coevolved to function coordinately to promote mtDNA replication. The mtDNA helicase and the catalytic subunit of Pol γ share ancestry with the gp4 primase–helicase and the gp5 DNA polymerase of bacteriophage T7, respectively [66]. mtSSB and Pol γ-β are eubacterial-like proteins with strong structural similarity to the homotetrameric E. coli SSB [67] and a class II aminoacyl-tRNA synthetase [68–70], respectively. However, unlike mtSSB, which most likely originated from the endosymbiotic α-proteobacterium that became the eukaryotic mitochondrion, sequence alignments indicate that Pol γ-β evolved as the accessory subunit of the mitochondrial replicase by lateral gene transfer involving a eubacterial species and early metazoans [20,70].

Fig. 4 Evolutionary Trace analysis of animal mtSSBs. (A) Multiple amino acid sequence alignment using selected animal mtSSBs retrieved from public databases (NCBI, Ensembl Metazoa and 959 Nematode Genomes). TBLASTN [57] and HMMR3 BLAST [58] searches were performed using the translated mRNA reference sequences for the Homo sapiens (AK313033.1) and Drosophila melanogaster (BT016028.1) mtSSB as queries. The alignment was performed using the MUSCLE algorithm built into the MEGA6 software [59]. The outgroup sequences used here were the Saccharomyces cerevisiae mtSSB (S43128.1) and the Escherichia coli SSB (J01704.1). Red (gray in the print version) and light green (light gray in the print version) arrows above the alignment indicate the residues that are crucial for ssDNA binding and subunit interactions in the E. coli SSB protein, respectively [60,61]. Gray, purple (dark gray in the print version), and dark green (dark gray in the print version) bars below the alignment indicate, respectively, three mtSSB regions grouped by their functional properties: N- and C-termini (which have negative effects on Pol γ stimulation [48]); loop 2,3, alpha-1, and loop 4,5-1 (which have positive effects on Pol γ stimulation [8]); and loop 1,2 and loop 4,5-2 (which have positive effects on mtDNA helicase stimulation [8]). The cyan (light gray in the print version) arrows and helix below the alignment indicate the residues that form the β sheets and the only α-helix in the mtSSB polypeptide, according to the crystal structure (PDB: 3ULL, [62]). The residues highlighted in yellow (light gray in the print version) were identified by the Evolutionary Trace Server [63,64] as conserved in metazoans. These have been mapped onto the crystal structure of the human mtSSB

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Fig. 4—Cont'd (B) and onto a model of ssDNA-bound mtSSB (C). The model is as reported by Oliveira and Kaguni [8]. D52 and D105 are the only residues identified by the Evolutionary Trace analysis that are most likely not involved in ssDNA binding (see text for details).

We recently analyzed the largest number of Pol γ-α and β sequences currently available for animal species, taking advantage of the ever-increasing volume of genomic and transcriptomic data in public databases [20]. We showed that the mitochondrial replicase presents distinct patterns of molecular evolution throughout the animal phylogenetic distribution, which might reflect distinct mechanisms for replicating mtDNA (see Section 3). The most striking finding relates to the oligomeric composition of Pol γ, which extends beyond a simplistic view of one Pol γ-α and one or two Pol γ-β protomers forming the holoenzyme. The presence of the helixloop-helix (HLH)-3β domain in Pol γ-β (Fig. 5) that enables its homodimerization in the human and mouse replicases [18,71] argues that the heterotrimeric nature is conserved in all vertebrate species. On the other hand, a holoenzyme comprising one Pol γ-α and one Pol γ-β, represented by the D. melanogaster enzyme [27,28], is most likely the ancestral form for all animals. However, our analysis revealed exceptions to this ancestral

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Fig. 5 Dimerization of vertebrate Pol γ-β via the formation of the four-helix bundle structure. (A) Amino acid sequence alignment indicates the presence of the HLH-3β domain (boxed) in all species of Vertebrata and possibly in a few other animal groups. (B) Comparison of the crystal structure of the human Pol γ-β dimer and structural models for Pol γ-β of Trichoplax adhaerens, Strongylocentrotus purpuratus, Drosophila melanogaster, and Ciona intestinalis, showing that only vertebrate Pol γ-β can fold into a HLH-3β structure and therefore, form the four-helix bundle dimerization interface. The inset shows the three short β-sheets at the base of the HLH-3β structure. Reproduced with permission from M.T. Oliveira, J. Haukka, L.S. Kaguni, Evolution of the metazoan mitochondrial replicase, Genome Biol. Evol. 7 (2015) 943–959.

heterodimeric form. Specifically, we were unable to find the Pol γ-β gene in the genome and/or transcriptome of any nematode species, a finding that correlates with the absence of the AID domain in the Pol γ-α polypeptide, indicating that the mitochondrial replicase in this animal group does not

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have a β accessory subunit, and may be a single-subunit enzyme [20]. The nematode mtDNA polymerase resembles the enzyme of other eukaryotic taxa such as Saccharomyces cerevisiae. Notably, mtDNA replication in the nematode C. elegans proceeds via a rolling circle mechanism [72], which is apparently the main mode of mtDNA replication in S. cerevisiae [73], and represents a mechanism distinct from that shown for other animals (see Section 3). Clearly, the evolution of the oligomeric form of Pol γ has taken several routes that may correlate with variations in the mechanisms of mtDNA replication in vivo [20]. We reported recently a direct comparison of recombinant Pol γ holoenzymes from a vertebrate (human) and an insect (D. melanogaster) species, which showed that the extent of DNA synthesis on a singly primed, circular ssDNA template is fivefold higher with the human enzyme [49] (Fig. 6C). In comparison, when variants of the human and mouse Pol γ-β that are unable to homodimerize were examined in similar in vitro assays in a heterodimeric state with Pol γ-α, thus resembling insect Pol γ, the decrease in DNA polymerase activity observed as compared to that of their native heterotrimeric forms ranged from four to sevenfold [71,74,75]. The heterotrimeric holoenzyme form might provide a functional advantage to vertebrates during mtDNA replication to increase the rate of nucleotide incorporation. In comparing the crystal structure of the human Pol γ apo-holoenzyme with that of the replicase in a ternary complex with a primer-template DNA and nucleotide, Yin and collaborators noted that the Pol γ-β dimer is rotated 22 degrees toward the pol domain of Pol γ-α upon DNA binding, allowing the catalytic subunit to interact extensively with the distal Pol γ-β protomer, and enabling the dimeric accessory subunit to regulate DNA synthesis allosterically [7,19]. In the ternary complex, human Pol γ-α undergoes conformational changes in several subdomains including the fingers, thumb, L-helix, and IP subdomain. These changes are perhaps more limited in the single β-containing animal Pol γs, including that of D. melanogaster, such that its intrinsic DNA synthesis capacity is lower in the absence of other replisome proteins. Comparison of the heterodimeric and heterotrimeric Pol γs in in vitro DNA polymerase assays in the absence of other components of the mtDNA replisome provides an incomplete picture. Indeed, mtSSB has been shown to stimulate both the DNA polymerase and exonuclease activities of Pol γ 20-fold, although this value varies substantially between the human and D. melanogaster systems [8,46–48]. To explore this, we pursued a comparative analysis of the human and D. melanogaster Pol γ holoenzymes with their cognate and heterologous mtSSBs using a combination of

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Fig. 6 Stimulation of DNA synthesis catalyzed by Pol γ correlates with specific ssDNA template organization by mtSSB. (A and B) DNA polymerase assays were performed using 58.5 fmol of singly primed M13 DNA, 35 fmol of human Pol γ-α, 220 fmol of human Pol γ-β (A), or 40 fmol of D. melanogaster Pol γ (as Pol γ-α) (B), and 0, 6.4, 10.7, 16, or 32 pmol of either human (open circles) or D. melanogaster (closed circles) wild-type mtSSB. Assays were performed at 30 mM KCl and 4 mM MgCl2. The data were normalized to the amount of nucleotide incorporated by human Pol γ in the absence of mtSSB (arbitrarily set to 1 in each case). (C) Comparison of nucleotide incorporation by human and D. melanogaster Pol γ in the absence or presence of their cognate mtSSBs at the concentrations resulting in maximal stimulation. (D) Electron microscopy of human (top) and D. melanogaster (bottom) wild-type mtSSB proteins bound to M13 DNA. The binding reaction was performed at 30 mM KCl and 4 mM MgCl2. The images are representative of template species formed at the following ratios of mtSSB tetramers per 100 nucleotides, which correspond to the indicated individual phases of the stimulation of human Pol γ activity: limiting mtSSB, 1.6 human and 1.2 D. melanogaster mtSSB; initial stimulation, 3.2 human and 1.8 D. melanogaster mtSSB; maximal stimulation, 3.8 human and 2.5 D. melanogaster mtSSB; inhibition, 6.4 human and 7 D. melanogaster mtSSB. Reproduced with permission from G.L. Ciesielski, O. Bermek, F.A. Rosado-Ruiz, S.L. Hovde, O.J. Neitzke, J.D. Griffith, L.S. Kaguni, Mitochondrial single-stranded DNA-binding proteins stimulate the activity of DNA polymerase γ by organization of the template DNA, J. Biol. Chem. 290 (2015) 28697–28707.

biochemical assays and transmission electron microscopy [49]. Under optimal conditions for each, both holoenzymes showed comparable DNA synthesis in the presence of their cognate SSBs. D. melanogaster Pol γ alone is fivefold less efficient than the human replicase. Notwithstanding their

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different oligomeric forms, the human and fly Pol γs behave similarly at maximum stimulation by mtSSB. In addition, D. melanogaster mtSSB stimulates human Pol γ as efficiently as does human mtSSB, and vice versa (Fig. 6A and B). However, the stimulation profile is mtSSB specific: both Pol γs reach their maximal activity at a concentration of the insect mtSSB lower than that required in the presence of human mtSSB. This corresponds to a similar shift in the concentrations required for generation of specific mtSSB-template complexes (Fig. 6D) that correlate with individual phases of the stimulatory effect [49]. That the ssDNA-binding affinity of both mtSSBs is similar suggests the more efficient DNA template organization by D. melanogaster mtSSB compensates for the lower DNA synthetic capacity of D. melanogaster Pol γ. In composite, our results suggest that structural differences between the human and D. melanogaster mtSSBs give rise to their differential stimulatory effects on Pol γ. We identified by site-directed mutagenesis regions of human mtSSB that are important for maximal stimulation of human Pol γ in vitro, including loop 2,3 (S51-L59), the α-helix 1 (Y83-Q84), and loop 4,5 (Y100-E102) [8]. In fact, loop 2,3 and α-helix 1 show little sequence similarity between vertebrate and invertebrate species, and an extension of 6–7 amino acid residues in loop 2,3, which is disordered in the crystal structure of the human mtSSB, appears predominantly in vertebrate species (Fig. 4A). Our recent electron microscopic study shows that loop 2,3 may be important for human mtSSB to organize the template in a competent configuration for Pol γ [49]. The extended loop 2,3 of vertebrate mtSSB, together with the presence of the additional Pol γ-β protomer in the vertebrate mitochondrial replicase, and other putative structural evolutionary novelties described for the catalytic subunit [20] may then explain the biochemical differences in the vertebrate and insect systems. This may in turn correlate with different modes of mtDNA replication in vivo (see Section 3). Animal mtSSBs lack the long acidic C-terminal tail present in bacterial SSBs (Fig. 4A), which protrudes into the ssDNA-binding channel in the absence of DNA and is then rendered available upon ssDNA binding for protein–protein interactions with other components of the replication machinery (reviewed in Ref. [45]). By contrast, animal mtSSBs contain an N-terminal extension (notwithstanding the mitochondrial presequence) that is absent in the eubacterial homologues (Fig. 4A, gray bars). Removal of both termini of the human mtSSB, independently or in combination, revealed that they regulate negatively the stimulation of Pol γ while exerting no effects on ssDNA-binding affinity [48]. It remains to be determined what

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effects the termini of mtSSB might exhibit within the mtDNA replisome, and because the terminal regions are extremely variable among metazoans, molecular evolutionary analysis fails to shed light on their specific roles in other taxa. At present, site-directed mutagenesis of the human protein suggests that amino acid residues in mtSSB that are important for the functional interactions with Pol γ and the mtDNA helicase are distinct (Fig. 4A, purple and dark green bars, respectively) [8]. Maximal stimulation of the human mtDNA helicase requires residues E33-K35 in loop 1,2 and K106-N108 in loop 4,5 of the human mtSSB. dsDNA unwinding produces ssDNA that is bound by mtSSB (consistent with recent data in vivo [76]). In contrast, in mtSSB–Pol γ interactions, mtSSB first binds to the template DNA strand, organizes it in a competent conformation for Pol γ, and then is displaced by the replicase as it synthesizes the complementary strand. This is consistent with the fact that excess mtSSB inhibits DNA synthesis by Pol γ, but not dsDNA unwinding by the mtDNA helicase [8,49]. Falkenberg and coworkers first reconstituted a minimal mtDNA replisome, combining human Pol γ, mtDNA helicase and mtSSB in in vitro DNA synthesis using a 70-nt single-stranded circular DNA template annealed to a primer with a 50 -extension [51]. In this system, DNA synthesis occurs by a rolling circle-like mechanism, producing linear fragments much longer than the template (2000 nt). The substantial stimulation by mtSSB may reflect functional interactions with Pol γ and/or its binding to the leading and lagging strands in a competent conformation at the fork. Stranddisplacement DNA synthesis by the minimal mtDNA replisome has been employed to elucidate the replication defects in a number of disease-related enzyme variants of both human Pol γ-α and mtDNA helicase in the context of a functional mtDNA replication fork [32,77,78]. Furthermore, addition of the mitochondrial RNA polymerase engenders priming and DNA synthesis on the lagging DNA strand, to generate nicked dsDNA products [79–81]. In subsequent studies, a double-stranded circular DNA template containing a displacement loop (bubble template) was used to show that the mtDNA helicase can load in the absence of a helicase loader such as those required in prokaryotic and nuclear systems, leading to DNA synthesis by the concerted actions of Pol γ, mtDNA helicase, and mtSSB [43]. Notably, the reaction efficiency on this template was low as compared with the earlier substrates. To date, only the elongation phase of mtDNA replication has been reconstituted. Thus, there remains much to learn about the initiation and termination of mtDNA replication, and the proteins that function in those processes.

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In sum, biochemical studies have demonstrated important functional interactions at the mtDNA replication fork, and we have attempted to provide an overview with evolutionary perspective. At present, we know little about how these functional interactions rely on physical contacts, and if and how they may vary in different animals. We also acknowledge the importance of other factors in mtDNA maintenance: TFAM in organization of mtDNA nucleoids, RNase H1, FEN1, DNA2, and MGME1 in primer processing, Top1mt, Top2β, and Top3α in altering mtDNA topology, DNA ligase III, mitochondrial RNA polymerase, and PrimPol (possible primase activity), and a diverse array of transcription and repair proteins (reviewed in Ref. [11]). With the diversity in the protein requirements for mtDNA synthesis in replication, repair, and recombination, it is likely that other factors also participate and as a result, the approach of large-scale screening for new proteins with mtDNA maintenance functions, such as that recently reported by Fukuoh et al. [55], is especially compelling.

3. MECHANISMS OF MITOCHONDRIAL DNA REPLICATION IN VIVO The recent application of diverse techniques to evaluate mtDNA replication in various physiological systems has prompted new models for animal mtDNA replication, highlighting the mechanistic diversity found in vivo to ensure appropriate mtDNA copy number, mitochondrial gene expression, and ATP production via oxidative phosphorylation. Here, we provide an overview of the current models of mtDNA replication in nematodes, insects, echinoderms, and vertebrates, though these may not represent all that exist throughout the animal phylogenetic distribution.

3.1 The Rolling Circle Model in C. elegans Although the round worm C. elegans has been used as a model organism in many studies, including those on mitochondrial diseases [82–84], replication of its mtDNA has only recently been investigated. Two-dimensional agarose gel electrophoresis (2DAGE) of mitochondrial nucleic acids (mtNA) isolated from sucrose gradient-purified mitochondria revealed prominent signals for Y and X arcs. These are representative of fragments containing elongating replication forks and cruciform structures, respectively [85–88]. No structure corresponding to a replication initiation bubble (bubble arc) was detected [72]. Direct observation of the isolated mtDNA by transmission electron microscopy (TEM) identified branched-circular lariat

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Fig. 7 Models of invertebrate mtDNA replication. (A) The rolling circle mechanism of nematode mtDNA replication proposed by Lewis et al. [72]. Circular progeny mtDNA molecules are proposed to be formed by a recombination-based resolution that involves the major noncoding region (NCR). (B) The strand-coupled theta-like model of insect mtDNA replication. In the predominant mode described by Joers and Jacobs [89], replication initiates within the noncoding A + T region (A + T) and proceeds unidirectionally. The partially or completely strand-uncoupled models for Drosophila mtDNA replication [89,90] (see text for details) are not represented. Arrows associated with replicating mtDNA indicate the 50 - to 3-0 direction of DNA synthesis. The gray arrowhead indicates the number and directionality of replication forks generated at the origin.

molecules with concatemeric tails, which are characteristic of intermediates during rolling circle replication (Fig. 7). The X arcs observed by 2DAGE, which traditionally correspond to various cruciform structures that may result from recombination or replication termination [87,91–93], were abundant primarily when the fragment harboring the major noncoding region (NCR) of the nematode mtDNA was probed. Treatment of the mtNA with a combination of S1 nuclease and E. coli resolvase RusA, highly specific for resolving Holiday junctions, reduced the X arc signal substantially, suggesting that a subfraction of the X-shaped DNA molecules are possibly hemicatenanes, a DNA species resulting from the convergence of two Holiday junctions, or replication fork stalling [94]. The rolling circle mechanism is a robust mechanism to assure efficient production of genomes as is exploited by various bacteriophages, such as Phi29, T4, λ, and M13 [95–98]. Interestingly, mitochondria of plants [99,100] and fungi [73,101] employ

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this mechanism of DNA replication, which suggests its ancestral origin. Nematodes represented by C. elegans appear to have maintained it [72], although all other animals analyzed to date seem to have evolved distinct mechanisms to replicate their mtDNA (see below).

3.2 The Theta Model in D. melanogaster EM studies as early as the 1970s identified replication bubble-like structures, indicative of the initiation site within the repetitive noncoding A + T region in replicating mtDNA isolated from ovaries and embryos of various Drosophila species, including D. melanogaster [90,102]. Recent analysis of D. melanogaster mtDNA from S2 cells in culture and whole animals using the 2DAGE technique corroborated these findings [89], and both the early and current reports concluded that replication starts at and proceeds unidirectionally from the A + T region in a theta-like fashion (Fig. 7). The relative contribution of replication intermediates (RIs) that are products of uncoupled or coupled leading- and lagging-strand synthesis is a point of disagreement; whereas the EM studies reported that an asynchronous mode of DNA synthesis predominates (with up to 99% of leading DNA strand synthesis completed before the initiation of the lagging-strand synthesis), the 2DAGE analyses suggest the presence of only a subset of replicating molecules with stretches of ssDNA, arguing that D. melanogaster mtDNA replicates via a mechanism of coupled leading- and lagging-strand synthesis. Because duplex DNA RIs observed by EM in an earlier study [103] are consistent with the 2DAGE data, the differences reported do not appear to be related to the techniques used; rather differences may lie in the preparation of mtDNA, and remain to be reconciled. The 2DAGE analysis also revealed strong signals within the D. melanogaster mtDNA for two distinct replication pause sites or slowreplicating zones, which were mapped to the binding sites of the transcription termination factor DmTTF [89,104]. This implies that as with mammalian mtDNA [105], the transcription apparatus in Drosophila mitochondria also regulates mtDNA replication rate. Similar to the analysis of C. elegans mtDNA RIs, cruciform species (X arcs) are prominent intermediates in the Drosophila 2DAGE analysis, and map to the middle of the A +T region [89,104]. In 1977, Rubenstein et al. reported that a significant fraction of D. melanogaster mtDNA was interlinked physically in two predominant forms of either supercoiled or relaxed catenated molecules. Interestingly, the frequency of catenanes increased during development from 6.2% of the closed circular mtDNA molecules in 1–6-h-old embryos to

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9.5% in 20-h-old embryos, and 12.5% in tissue culture cells in which mtDNA is replicated actively [103]. Thus, it appears that catenanes are related to the mtDNA replication process, suggesting that the cruciform intermediates observed by 2DAGE are derived by their digestion with the restriction endonucleases used in mtNA processing. Prominent RusAsensitive X arcs and the presence of catenanes among the D. melanogaster mtRIs might be explained by a template switching-like mechanism, which has been proposed recently for replication of the 2 μ plasmid-based high copy number minichromosome of the yeast S. cerevisiae [93]. Templateswitching replication mediates damage bypass via a recombination-based mechanism in which a dynamic range of cruciform structures ultimately result in formation of hemicatenanes or similar forms. Further studies that engage both 2DAGE and EM are needed to explain the presence of cruciform structures and/or catenanes in D. melanogaster mtDNA.

3.3 The Theta Model in Sea Urchin Sea urchins, such as Strongylocentrotus purpuratus, have been used as invertebrate models of the Deuterostome superphylum for many decades [106]. Their mtDNA contains a structure called the D-loop (alternately, R-loop) that is formed by the stable association of transcripts of 60 bp with the major NCR, to generate a short triplex structure [107]. The D-loop RNA segments are found covalently linked to an 15-bp long DNA, implying that the D-loop is the site of transition from priming to nascent leading DNA strand synthesis [107]. As with vertebrates (see below), D-loop strand synthesis events terminate frequently at the replication termination sequence downstream of the D-loop, but the mechanism allowing read-through and subsequent processive DNA synthesis remains unknown [108]. Using EM and 2DAGE, replication was shown to initiate in the D-loop region and proceed unidirectionally, implying that echinoderm mtDNA replication advances by D-loop expansion in a theta-like fashion [108,109]. Characterization of RIs showed a high frequency of multiple duplex DNA segments on the lagging strand, and a less abundant fraction of ssDNA-containing species. Replication pause sites were found at widely scattered positions in the genome. The most prominent are at a distance of 1/3 from the leading-strand origin that coincides with the proposed origin of lagging-strand synthesis. Interestingly, lagging-strand synthesis appears to pause at the leading-strand origin, suggesting that synthesis from, and pausing events in both sites could produce species that would appear as Cairns’ forms in EM analysis [108]. By contrast, Matsumoto et al. [109] observed RIs with expanded D-loops that were exclusively single-stranded.

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3.4 mtDNA Replication in Vertebrates 3.4.1 Common Concepts We begin our discussion of vertebrate mtDNA replication with a description of initiation of leading-strand synthesis, its relation to transcriptional start sites, and the structure and function of the D-loop and 7S DNA. Our focus will be primarily on mammals because they are represented by the bulk of the reported results, and we cite data available on other systems where relevant. The complementary strands of vertebrate mtDNA are denoted as heavy (H) and light (L) due to their distinct base composition, which results in different sedimentation patterns in CsCl gradients [110]. The single NCR of the mammalian mitochondrial genome spans 1 kb on average and contains three conserved sequence blocks (CSBs), assigned such that CSB1 is located near the tRNAPro gene and CSB3 lies toward the middle of the NCR [111–113]. In 1971, Kasamatsu and colleagues demonstrated that up to 50% of closed circular mtDNA molecules isolated from mouse L cells in culture contain a D-loop structure comprising a displaced heavy strand, and a light strand hybridized to a DNA segment with a sedimentation velocity of 7S [114] (Fig. 8A). Further analysis demonstrated that the D-loop is located in the NCR [111–113]. The actual length of the D-loop depends on the length of the 7S DNA, which in humans is generally 600 nt but may vary in size at its 50 -end that lies adjacent to CSB1 [115]. The 30 -end of the 7S DNA maps specifically between nucleotides 16104–16106, adjacent to the termination-associated sequence (TAS) (Fig. 8A) on the other end of the NCR, near the tRNAPhe gene [116,117]. The TAS is likely engaged in formation of a secondary structure, and represents a termination site for both RNA and DNA synthesis in vivo [116–118]. Several comparative studies of the mitochondrial NCR region demonstrated that the presence of individual CSB sites or length of the D-loop region vary among mammals [119,120]. For example, the mouse and bovine D-loop regions are 200 nt shorter than in human mtDNA, and the bovine NCR appears to lack CSB1 and 3 [121]. These differences extend to other classes of Vertebrates. The D-loop region of Xenopus laevis mtDNA is 1.6 kb long, and CSB1 appears to be absent [122]. The galliform species of birds contain a mitochondrial genome 200 nt larger than that of most mammals, and a D-loop of 780 nt accounts for much of the length difference [123,124]. The chicken NCR has been shown to contain a single bidirectional promoter as compared to the two separate transcriptional promoters as in mammals [125].

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Fig. 8 Current models of vertebrate mtDNA replication. (A) Structural organization of the D-loop and the adjacent cis-elements present within the noncoding region of vertebrate mtDNA. CSB1, 2, and 3, conserved sequence blocks 1, 2, and 3; TAS, termination-associated sequence; LSP, light-strand promoter; OH, origin of heavy-strand DNA synthesis. The numbers below each element represent genomic positions in the human mtDNA reference sequence. The schematic is to scale, except for the region represented by the dashed lines. (B) Strand-displacement, RITOLS/bootlace and strandcoupled models of vertebrate mtDNA replication (see text for details). The sites OH and OL (origin of light-strand DNA synthesis) are represented as reference points on the genome map, although these sites are important primarily for the strand-displacement model. Arrows associated with replicating mtDNA indicate the 50 - to 30 - direction of DNA synthesis; continuous and dotted lines represent DNA and RNA, respectively. Only the long stretches of RNA described in the RITOLS/bootlace model are represented; the putative short RNA primers of the other models are not shown. Gray arrowheads indicate the number and directionality of replication forks generated at the origin according to each model. Adapted from E.A. McKinney, M.T. Oliveira, Replicating animal mitochondrial DNA, Genet. Mol. Biol. 36 (2013) 308–315.

It is commonly accepted that synthesis of the nascent heavy strand in mammals, which is the leading strand in mtDNA replication [126,127], originates in the NCR with the majority of replication initiation events occurring downstream of CBS1 at OH (nt position 191) [88,111–113]. The free 50 -end of the longest 7S DNA identified maps to OH [115,128], implying that 7S DNA is a prematurely terminated leading strand. Consistent with this, the 50 -end of nascent (elongated beyond NCR) and total

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(including 7S DNA) heavy strands have been mapped to OH (nucleotide position 190  1) [128]. Synthesis of the leading strand from OH is primed from the light-strand transcriptional promoter (LSP) located at nucleotide position 400 [129,130]. Indeed, in addition to the long transcripts produced from LSP, transcripts of around 200 nt have also been documented as 7S RNA. The location of the free 50 -end of 7S RNA corresponds to the LSP, whereas its 30 -ends map to all three CSB sites with CSB1 predominating [113,117,131]. That mitochondrial transcripts are typically of genome length suggests a mechanism exists to inhibit processive synthesis, thereby enabling synthesis of short primers for replication initiation. It was shown recently that up to two-thirds of all transcription events initiated at LSP are terminated prematurely due to stacking interactions of guanine residues of nascent RNA and the nontemplate displaced heavy strand, forming a G-quadruplex at the CSB2 site [132,133]. In vitro, the mitochondrial transcription elongation factor (TEFM) promotes processive synthesis by the mitochondrial RNA polymerase (POLRMT), abolishing premature termination at CSB2 and increasing dramatically the abundance of longer transcripts [134–136]. It has been proposed that TEFM binds the G-quadruplex region directly, serving as a molecular switch between primer synthesis and processive transcription [136]. In contrast, RNA primers remain in the NCR upon depletion of mitochondrial RNase H in mouse embryonic fibroblasts, resulting in generation of double-stranded breaks at the origin, thus highlighting the essential role of RNase H in primer processing at the OH site [137]. To date, the roles of the R-loop at CSB2, and the mechanism(s) for bypassing it by the priming apparatus are not fully understood, and warrant further investigation. Because the half-life of the 7S DNA in rodent cells has been documented to be 45 min [138], one can argue that the frequent replacement of the 7S DNA contradicts the notion of it serving to “prime” further synthesis. If so, a high rate of initiation events from OH rarely results in synthesis through the TAS site. The terminated DNA is likely removed by the mitochondrial ssDNA nuclease MGME1 [139,140]; MGME1 knockdown in cultured cells or MGME1 deficiency in human patients results in a large accumulation of 7S DNA [139,140]. Pol γ appears to bind preferentially at OH in vivo, and at the site corresponding to the 30 -end of the 7S DNA, which suggests a mechanism for replisome restart from the latter [117]. Upon ddCTP treatment, increased binding of the mtDNA helicase at the TAS site is also observed, suggesting that it also represents replication restart from the 30 -end of 7S DNA. However, this treatment leads to substantially reduced occupancy of Pol γ

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at this site, which would in any case be nonproductive due to its documented sensitivity to ddCTP incorporation. Perhaps the mtDNA helicase simply accumulates at the transition point from ssDNA to dsDNA when DNA synthesis stalls or is terminated. In support, data indicates that mtDNA helicase is enriched at the OH region and not at the 30 -end of the 7S DNA under normal conditions. The reason for the rapid turnover of 7S DNA remains unclear, but as suggested previously [128] may relate to functions of the D-loop in processes not associated with replication initiation. In that regard, a novel mitochondrial protein, ATAD3, binds the D-loop specifically and facilitates the interaction and segregation of mtDNA molecules in dividing mitochondria, and possibly anchors the mtDNA molecules to a portion of the mitochondrial inner membrane that is cholesterol-rich [141,142]. A site for protein complex formation and plausibly for interactions of mtDNA with the mitochondrial inner membrane, the D-loop is also a natural substrate for mtSSB binding [143]. Given that the presence of mtSSB is a hallmark of replicating mtDNA molecules, the D-loop likely serves as the site for replisome assembly [76]. Moreover, Pol γ-β binds dsDNA [144] and has been demonstrated to bind with high affinity to the D-loop [145]. Furthermore, cells depleted of mtSSB and mtDNA helicase show loss of 7S DNA [56,146], implying that the D-loop structure is maintained by the replisome, akin to the formation of the replication initiation bubble in other replication systems [147]. 3.4.2 The Strand-Displacement, RITOLS/Bootlace, and Strand-Coupled Models On the basis of EM analyses of mtDNA isolated from mouse cells in culture [110], it was proposed over three decades ago that mammalian mtDNA is replicated via a strand-displacement mechanism that is unidirectional and asynchronous [148] (Fig. 8B). In this model, replication proceeds from OH continuously and unidirectionally, displacing the parental heavy strand, which remains single-stranded and bound by mtSSB [81,117,148,149]. When the replication fork approaches approximately two-thirds of the genome length, the initiation site for lagging-strand synthesis, OL, is exposed forming a stem-and-loop structure [148,150,151]. POLRMT has been shown to bind to the OL structure to initiate primer synthesis up to 25 nts, followed by binding by Pol γ to catalyze light-strand DNA synthesis [80,150–152]. The specific mechanism by which POLRMT recognizes OL and activates primer synthesis in vivo remains unknown, although a role for POLRMT in priming DNA synthesis from both strands has been shown

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in vitro [79,80]. After initiation at OL, synthesis on both strands proceeds continuously, until two fully replicated daughter molecules are formed and segregated (Fig. 8B). Notably, a stem-loop structure equivalent to OL appears to be absent in the mtDNA of galliform species, which is also the case for species from many classes of Vertebrates, including for example, short-beaked dolphins and snake-neck turtles [124,152]. Thus one might expect variations in the replication process among vertebrates, and in lagging DNA strand synthesis in particular. In the year 2000, Holt and colleagues published the first results involving analyses of human and rodent RIs by 2DAGE [86]. They demonstrated that RIs contain a ssDNA-specific nuclease-resistant Y arc, suggesting that the long single-stranded stretches of the parental heavy strand predicted by the strand-displacement model were absent. Instead, the authors proposed that the RIs observed in their analysis resulted from a strand-coupled replication mechanism. Holt and colleagues noted that a small subset of the Y arc intermediates were sensitive to ssDNA-specific nucleases. Thereafter, a refined protocol for isolation of mtNA to include a step of sucrose density-gradient purification led to the elimination of ssDNA-containing species in 2DAGE analysis, revealing a substantial presence of ribonucleotides in mtDNA RIs [153]. Novel, unusually large DNA molecules forming slow-moving arcs in RIs were also identified. Whereas they were found to be largely resistant to restriction endonuclease digestion, they were sensitive to RNase H, suggesting that the incorporated RNA tracts were inhibiting the restriction enzymes. This led to a new proposed mechanism of mtDNA replication, similar in principle to the strand-displacement model, but positing ribonucleotides incorporated through out the lagging strand (RITOLS) prior to initiation of the light-strand synthesis [127,154] (Fig. 8B). Later, it was demonstrated that inhibition of transcription with cordycepin triphosphate did not affect replication, arguing that during leading-strand synthesis preexisting RNA is incorporated on the lagging strand via a “bootlace” strategy, rather than being synthesized concomitant with leading-strand synthesis [155]. Indeed, mature RNA is stored in mammalian mitochondria in RNA granules juxtaposed to the mitochondrial nucleoids [156,157], and could feasibly serve as a source of the RITOLS. It remains unclear how the incorporated RNA is removed in maturation of progeny molecules [155,158]. Although mitochondrial RNase H may be involved, recent data on the effects of its depletion in mouse embryonic fibroblasts show accumulation of RNA at OH and OL, suggesting a role in primer removal, though accumulation of the bootlace RNA was not apparent [137].

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The 2DAGE technique has been used extensively in the analysis of various plasmid replication mechanisms [85,87,91,159–163], but it differs from the traditional, well-established EM analyses using CsCl-EtBr gradientpurified mtDNA [110,114]. Providing experimental support for the RITOLS model, Pohjoisma¨ki and colleagues demonstrated that various methods of sample preparation affected substantially the composition of RIs. In particular, the use of CsCl gradient sedimentation and E. coli SSB binding (common in EM analysis) lead to systematic loss of RNA from the RNA/DNA hybrids. Moreover, EM analyses of the samples prepared as for 2DAGE showed that intact mtDNA RIs are predominantly fully duplex. Rather, RNase H treatment resulted in accumulation of singlestranded intermediates consistent with those observed earlier by EM analysis [164]. In contrast, analysis by atomic force microscopy of rat liver mtNA that was isolated by a protocol similar to that used for the 2DAGE analysis, but involving CsCl-EtBr gradient sedimentation, lead to the conclusion that the majority of mtDNA RIs are formed by the strand-displacement mechanism [149]. Fuste´ et al. recently evaluated existing 2DAGE data in comparison with mtNAs treated with RNases or with DNase I; they observed that simple mixing of either RNA- or DNA-free mtNA samples restores the bubble arc that had disappeared upon removal of the RNA, implying that the RIs giving rise to the RITOLS model could be artifacts of the isolation procedure. This evidence does not disprove the presence of the RNA/DNA hybrids in vivo [81], and this study did not reconstitute fully the slowmoving species that represent a hallmark of the RITOLS model [154]. Compelling evidence for the strand-displacement model derives from recent data generated using ChIP-Seq on the distribution of mtSSB bound to mtDNA [81]. mtSSB was found bound almost exclusively to the heavy strand with high concentrations found in the D-loop, and decreasing proportionally toward OL, which is consistent with it being single-stranded during nascent heavy-strand synthesis. In contrast, Reyes et al. used psoralen/ UV cross-linking and in organello labeling to provide strong evidence that RNA/DNA hybrids do indeed form in vivo during the process of mammalian mtDNA replication and are later matured into fully duplex DNA [155]. Clearly, further experimentation is required to resolve these issues. The findings published in the early 2000s suggest that a fraction of the mtRIs comprise fully duplex dsDNA, the presumptive products of thetalike replication produced by coupled leading- and lagging-strand DNA synthesis [86,165]. This mode, however, appears to occur primarily when cells are recovering from EtBr-induced mtDNA depletion, a condition that

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stimulates mtDNA replication. In this case, a broad zone containing the CYTB, ND5, and ND6 genes constitutes the initiation site from which replication proceeds bidirectionally until termination occurs in the the D-loop region [165] (Fig. 8B). The model invokes the presence of Okazaki fragments, and though they have not yet been demonstrated directly, Okazaki fragmentprocessing enzymes including Pif1, FEN1, and Dna2 have been documented in mitochondria [166–168]. One might argue that the multiple sites of lagging-strand initiation found by Brown et al. [149] are evidence for Okazaki fragment-like species in replicating mtDNA. Although the strand-coupled model of mammalian mtDNA replication is now considered to be a secondary mechanism, 2DAGE analysis of mtRIs from the chicken Gallus gallus suggested that a strand-coupled theta replication mode predominates, and similar to that in mammals, initiates over a broad zone, ori-Z [165,169]. In contrast to mammals, in which the majority of the replication initiation events occur in the OH region, the prominent replication initiation site in chicken maps to the ND6 gene, and OH likely serves a role as a termination site. Interestingly, the ND6 and CYTB genes are transposed in bird mtDNA, such that ND6 maps adjacent to the NCR [123,169]. Termination of mtDNA replication in vertebrates is a subject of current studies that suggest a possible involvement of four-way junctions. These are proposed to arise when replication forks arrest in the NCR [165]. Three proteins of the mTERF family may contribute to the termination of mtDNA replication in human cells in culture [105,170]. Four-way structures that resemble Holliday junctions have been shown by EM to occur frequently in replicating mtDNA from human hearts, which also appears to be organized in multimeric catenated networks [171], suggestive of a recombination-based replication mechanism. The increased abundance of Holliday junctions and complex mtDNA forms obseved upon overexpression of TFAM or the mtDNA helicase in the mouse heart [171] suggests a similar recombinational DNA replication pathway. Similarly to that found in D. melanogaster, formation of more complex forms of mtDNA in the human heart correlates with high mtDNA copy number [172]. Interestingly, the mtDNA from mouse, rat, rabbit, and infant human hearts has a less complex organization [171,172].

4. PERSPECTIVES Since the discovery of mitochondrial DNA 50 years ago and the identification of the mitochondrial replicase a decade later, a broad base of

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knowledge has been established on key elements of mtDNA replication. Yet, many fascinating questions remain relevant to the expanding repertoire of proteins involved in mtDNA metabolism and its regulation. Understanding the interplay of key proteins at the replication fork with new protein players will require development of new substrates for in vitro assays that are validated by novel approaches to probe the changing physiological environment in mitochondria in both normal and disease states. A major hurdle remains to elucidate the mechanism(s) of replication initiation across taxa. The processes of termination and segregation to ensure the integrity of mtDNA inheritance also warrant future study. Exploring the roles of recombinational intermediates and catenanes in replication may link the physiological processes of DNA replication and recombination, and perhaps also postreplicational repair.

ACKNOWLEDGMENTS We thank Dr. Jon Kaguni for critical reading of the chapter. Research cited from the L.S.K. lab was supported by Grant GM45295 from the National Institutes of Health. L.S.K. was supported partially by the Academy of Finland. G.L.C. was supported by the University of Tampere. M.T.O. acknowledges support from the Fundac¸a˜o de Amparo à Pesquisa do Estado de Sa˜o Paulo (Grant 2014/02253-6).

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CHAPTER NINE

Fidelity of Nucleotide Incorporation by the RNA-Dependent RNA Polymerase from Poliovirus C.E. Cameron1, I.M. Moustafa, J.J. Arnold The Pennsylvania State University, University Park, PA, United States 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 1.1 Four Classes of Nucleic Acid Polymerases Exist 1.2 Universal and Adapted Features Revealed by Polymerase Structures 1.3 All Polymerases Employ a Two-Metal-Ion Mechanism for Catalysis 1.4 Two Rate-Limiting Conformational Changes Bracket Phosphoryl Transfer in the Kinetic Mechanism for Single-Nucleotide Incorporation 1.5 Views on the Kinetic and Structural Basis for Polymerase Fidelity Are Still Evolving 1.6 A Role for Dynamics in Enzymic Catalysis 2. Nucleic Acid Polymerases Employ General Acid Catalysis 3. Kinetic and Structural Determinants of PV RdRp Fidelity 4. Remote-Site Control of an Active-Site Fidelity Checkpoint 5. RdRp Fidelity Is a Determinant of Viral Virulence 6. RdRp Fidelity Mutants Are Vaccine Candidates 7. Correlated Motions of Functionally Important Motifs of the RdRp may be a Determinant of Nucleotide Incorporation Fidelity 8. Nucleotide Binding-Occluded and Binding-Competent States of the RdRp–RNA Complex as Determinants of Fidelity 9. Conserved, Active-Site Determinants of RdRp Incorporation Fidelity may Exist that can be Targeted for Viral Attenuation Acknowledgments References

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Abstract Using poliovirus (PV) and its RNA-dependent RNA polymerase (RdRp) as our primary model system, we have advanced knowledge fundamental to the chemistry and

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fidelity of nucleotide addition by nucleic acid polymerase. Two fidelity checkpoints exist prior to nucleotide addition. The first toggles the enzyme between a nucleotide binding-occluded state and a nucleotide binding-competent state. The second represents an ensemble of conformational states of conserved structural motifs that permits retention of the incoming nucleotide in a state competent for phosphoryl transfer long enough for chemistry to occur. Nucleophilic attack of the alpha-phosphorous atom of the incoming nucleotide produces a pentavalent transition state, collapse of which is facilitated by protonation of the pyrophosphate leaving group by a general acid. All of the relevant conformational states of the enzyme are controlled by a network of interacting residues that permits remote-site residues to control active-site function. The current state of the art for PV RdRp enzymology is such that mechanisms governing fidelity of this enzyme can now be targeted genetically and chemically for development of attenuated viruses and antiviral agents, respectively. Application of the knowledge obtained with the PV RdRp to the development of vaccines and antivirals for emerging RNA viruses represents an important goal for the future.

1. INTRODUCTION 1.1 Four Classes of Nucleic Acid Polymerases Exist Nucleic acid polymerases catalyze the template-dependent polymerization of (deoxy)nucleoside triphosphates and are essential for transcription, replication, and repair of the genomes of all organisms, including viruses [1,2]. Studies of nucleic acid polymerases in vitro have permitted the enzymes to be classified into four categories based upon the specificity for template and (deoxy)nucleoside triphosphate (NTP): (1) DNA-dependent DNA polymerase (DdDp); (2) RNA/DdDp (reverse transcriptase, RT); (3) DNAdependent RNA polymerase (DdRp); and (4) RNA-dependent RNA polymerases (RdRps). It should be noted that the template and substrate specificity of the various classes of polymerases is not absolute and is very easily manipulated in vitro by subtle changes in solution conditions, for example, by changing the divalent cation cofactor employed in the reaction from Mg2+ to Mn2+ [3–8]. Model systems have emerged for the study of the kinetic, thermodynamic, and structural basis for the specificity and activity of the various classes of polymerases. We have established the RdRp from poliovirus (PV), as a model system for understanding the chemical, kinetic, thermodynamic, structural, and dynamical mechanisms employed by this class of polymerases. Studies presented in subsequent sections of this chapter will hopefully leave you convinced that the PV RdRp system is not only a useful model for the RdRp but for all classes of polymerases.

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1.2 Universal and Adapted Features Revealed by Polymerase Structures Crystal structures have been solved for single-subunit polymerases representing all four classes of polymerases [9–16]. In general, the overall topology of polymerases resembles a cupped, right hand with fingers, palm, and thumb subdomains. The RdRp contains an extension of the fingers, the so-called fingertips, that interacts with the thumb subdomain, leading to a completely encircled active site in this class of polymerases (Fig. 1A).

Fig. 1 Universal and adapted features of polymerase structures. (A) PV RdRp structure. Conserved structural motifs in the palm subdomain are colored as follows: motif A, red (dark gray in the print version); motif B, green (dark gray in the print version); motif C, yellow (light gray in the print version); motif D, blue (dark gray in the print version); motif E, purple (gray in the print version); motif F, orange (gray in the print version); and motif G, black. (B) The NTP-binding site. The “universal” interactions are observed in all polymerases. The “adapted” interactions dictate the sugar configuration recognized by the polymerase. Reprinted with permission from D.W. Gohara, J.J. Arnold, C.E. Cameron, Poliovirus RNA-dependent RNA polymerase (3Dpol): kinetic, thermodynamic, and structural analysis of ribonucleotide selection, Biochemistry 43 (18) (2004) 5149–5158. Copyright (2004) American Chemical Society.

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The palm subdomain is the most conserved subdomain of polymerases and can be divided into several conserved structural motifs. In the case of the RdRp, seven conserved structural motifs exist: A–G (Fig. 1A). Motifs A and C contain conserved aspartic acid residues involved in binding the divalent cation cofactors required for nucleotidyl transfer (Fig. 1B). Motif A extends from the (deoxy)ribose-binding pocket to the catalytic center (Fig. 1B). Elements of this motif interacting with the triphosphate moiety and divalent cation superimpose exactly in different polymerases. Elements of this motif interacting with the sugar have evolved to sense the appropriate sugar configuration, thus linking the NTP-binding site to the catalytic site. Motif B contains residues that interact with the (deoxy)ribose moiety of the nucleotide substrate. Motif D was thought to provide structural integrity to the palm subdomain; however, the studies described herein have altered substantially this long-standing view by showing a role for this motif in the chemistry of nucleotidyl transfer. The remaining motifs are unique to RdRps, although some overlap with RTs also exists. Motif E provides interaction between the enzyme and primer. Motif F interacts with the triphosphate moiety of NTP and most likely functions at the earliest step of NTP binding. Motif G has no known function. Structural information is also available on polymerase elongation complexes [11,17–21]. Upon NTP binding, the fingers subdomain often moves relative to the thumb subdomain, causing an open-to-closed transition that has been considered a rate-limiting step in the nucleotide-addition cycle. This view is now being reconsidered [22,23].

1.3 All Polymerases Employ a Two-Metal-Ion Mechanism for Catalysis Given the conserved nature of the catalytic center of polymerases, the chemical mechanism is also conserved. The chemical mechanism is often referred to as the two-metal-ion mechanism (Fig. 2) [24,25]. Two divalent cations are required for polymerase-catalyzed nucleotidyl transfer; Mg2+ is the biologically relevant cofactor. One metal ion (metal B) enters the active site in complex with the nucleotide substrate. The other metal ion (metal A) likely enters from solution after binding of the Mg2+–NTP complex. The metals are stabilized in the active site by coordination to conserved residues in motifs A and C, oxygens from all three phosphates of NTP, and the oxygen from the primer terminus. It is generally accepted that the role of metal A is to increase the nucleophilicity of the primer 30 -OH by lowering the pKa of the hydroxyl. Metals A and B interact with oxygens of the α-phosphate

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Fig. 2 Two-metal-ion mechanism for nucleotidyl transfer. Nucleoside triphosphate (green, dark gray in the print version) enters the active site with a divalent cation (Mg2+, metal B). A second divalent cation (Mg2+, metal A) is coordinated by the 30 OH of primer terminus (cyan, light gray in the print version), the nucleotide α-phosphate, as well as Asp residues of structural motifs A and C. The unidentified proton acceptor and proton donor are indicated as A and B, respectively. The proton from the 30 -OH is indicated as Ha, and the proton provided by the unidentified donor is indicated as Hb. Adapted from C. Castro, E.D. Smidansky, J.J. Arnold, K.R. Maksimchuk, I. Moustafa, A. Uchida, M. Gotte, W. Konigsberg, C.E. Cameron, Nucleic acid polymerases use a general acid for nucleotidyl transfer, Nat. Struct. Mol. Biol. 16 (2) (2009) 212–218.

so these metals could presumably increase the electrophilicity of the α-phosphorous atom. Finally, metal B may also organize the triphosphate into a conformation that is essential for catalysis—that is, permit the optimal distance between the 30 -OH and the α-phosphate to be achieved. Nucleotidyl transfer reactions likely go through a pentavalent transition state [24–27]. At least one-proton-transfer reaction must occur during the reaction: deprotonation of the 30 -OH nucleophile. As described later, we have altered this view of polymerase chemistry by revealing two proton-transfer reactions in the transition state and the use of a general acid for protonation of the PPi leaving group.

1.4 Two Rate-Limiting Conformational Changes Bracket Phosphoryl Transfer in the Kinetic Mechanism for Single-Nucleotide Incorporation Presteady-state kinetic analysis of DdDps and RTs has produced complete kinetic mechanisms for incorporation of correct and incorrect nucleotides (Fig. 3) [28–32]. The nucleotide-addition cycle requires five steps, with

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Fig. 3 Complete kinetic mechanism for single-nucleotide incorporation for all classes of nucleic acid polymerases studied to date.

the nucleotidyl-transfer step bracketed by pre and postchemistry, conformational change steps. There is a consensus on the number of steps employed during the nucleotide-addition cycle. However, the rate-limiting step differs among polymerases and a physical description of the conformational changes is debated or not known. Experiments described later for the PV model system have begun to address these issues.

1.5 Views on the Kinetic and Structural Basis for Polymerase Fidelity Are Still Evolving There is no debate that polymerase fidelity represents an important, and perhaps the final, frontier for the field of polymerase enzymology. From a kinetic point of view, there is no debate that much of the observed difference between correct and incorrect nucleotide incorporation is expressed in the observed rate constant for the reaction (kpol) rather than the equilibrium constant for NTP binding (Kd) [22,23,33]. The problem has been assignment of kpol to one or more of the microscopic rate constants for the elementary steps of the reaction. The reason for this problem is the absence of a universal approach to determine if or the extent to which the nucleotidyl-transfer step contributes to kpol. It was once thought that the use of the phosphorothioate elemental (thio) effect was a useful tool [33,34]. However, the absence of a clear value for the theoretical maximum that could be used to determine the extent to which chemistry is expressed in kpol has made “thio effect” a naughty pair of words in the field [23,33]. We have made two significant advances in the PV RdRp system that have begun to resolve this problem. We have uncovered solution conditions to make chemistry completely rate limiting, permitting empirical determination of the maximal thio effect [35,36]. In addition, we have shown that the solvent deuterium isotope effect can be used to probe for chemistry in kpol [37,38]. It is now becoming clear that both the prechemistry, conformational change step and chemistry are reflected in kpol. The question remains whether one or both steps contribute to fidelity. If both steps contribute, how are they coupled? Moreover, the possibility that steps after chemistry contribute to nucleotide incorporation fidelity has really not been addressed.

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The structural basis for the conformational change preceding the chemical step is not known. Three hypotheses have been put forward. The first hypothesis is that this step reflects orientation of the triphosphate into the catalytically active conformation [30,39,40]. The second is that this step reflects the open-to-closed transition observed crystallographically after nucleotide binding [11,15,31,41]. The third is that this step reflects binding of metal A [19,42]. Published structures of DNA polymerase show that the open-to-closed transition can occur prior to binding of metal A, an observation interpreted to be inconsistent with this transition reflecting the conformational change observed kinetically [19].

1.6 A Role for Dynamics in Enzymic Catalysis Conformational changes required for an enzyme to progress successfully through its reaction coordinate are now believed to be influenced by dynamics on timescales ranging from picoseconds to milliseconds [43–47]. Dynamics on the fastest timescales have been observed to be coordinated by networks that include not only residues present at the catalytic site but also at remote sites, and perhaps explain allostery [43–46]. Very little is known about polymerase dynamics. We have obtained evidence for dynamics on the nanosecond timescale as a determinant of nucleotide incorporation fidelity for the PV RdRp. We suggest that dynamics–function relationships for all polymerase will yield insight into the mechanism of nucleotide incorporation fidelity that structure–function relationships have been unable to achieve.

2. NUCLEIC ACID POLYMERASES EMPLOY GENERAL ACID CATALYSIS Nucleotidyl transfer requires, minimally, one-proton-transfer reaction. The proton from primer 30 -OH must be removed for catalysis to occur. The function of one (metal A) of the two divalent cations required for activity is to lower the pKa of this proton to facilitate catalysis. However, it was not known if the pyrophosphate leaving group needed to be protonated for efficient nucleotidyl transfer. Most structures of polymerases poised for or undergoing catalysis showed a basic amino acid near the β-phosphate, consistent with this possibility [19,48–52]. We have employed the PV RdRp to address the question of protontransfer reactions during nucleotide addition. This system is uniquely suited to the characterization of the chemical mechanism of nucleotidyl transfer for

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two reasons. First, the stability of PV RdRp on our model primed template (sym/sub or s/s) is independent of pH [38,53]. Second, chemistry is the sole rate-limiting step when Mn2+ is employed as the divalent cation cofactor, thus permitting us to characterize this step without complications from conformational change steps [36]. We showed that the maximal rate constant for nucleotide incorporation (kpol) exhibited a bell-shaped dependence on pH (Fig. 4A). The pKa for the descending limb was 10.5, consistent with a Lys residue serving as a general

Fig. 4 Lys-359 participates directly in nucleotidyl transfer and functions as a general acid. (A) Values for kpol plotted as a function of pH for WT and K359L PV RdRp. RdRp–sym/sub complexes were rapidly mixed with varying concentrations of ATP at different pH values using the stopped-flow assay. At each pH, time courses at a fixed nucleotide concentration were fit to a single exponential to obtain kobs. The value for kobs was then plotted as a function of ATP concentration and fit to a hyperbola to obtain kpol at a given pH value. In the case of WT PV RdRp, the solid line shows the fit of the data to a model describing two ionizable groups, yielding pKa values of 7.0  0.1 and 10.5  0.1. Error bars indicate the standard deviation. (B) Proton-inventory plots for WT and K359L PV RdRp. kn values were obtained in the presence of different mole fractions (n) of D2O. The ratio of kn/kpol (kpol is in H2O) is then plotted as a function of mole fraction of D2O. WT data (filled circles) were fit to a two-proton model (solid line) with straight, dashed line shown for comparison. K359L data (filled squares) were fit to a one-proton model. Error bars indicate the standard deviation. Adapted from C. Castro, E.D. Smidansky, J.J. Arnold, K.R. Maksimchuk, I. Moustafa, A. Uchida, M. Gotte, W. Konigsberg, C.E. Cameron, Nucleic acid polymerases use a general acid for nucleotidyl transfer, Nat. Struct. Mol. Biol. 16 (2) (2009) 212–218.

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acid. We observed a solvent deuterium kinetic isotope effect (SDKIE) on kpol, permitting us to count the number of protons transferring in the transition state for nucleotidyl transfer by performing a proton-inventory experiment [37,38,54,55]. In this experiment, the observed rate constant for nucleotide incorporation (kn) is determined at various mole fractions of D2O (n). Values for kn/kpol are plotted as function of n. If two protontransfer reactions occur, then the data will fit best to a second-order polynomial. If one-proton-transfer reaction occurs, then the data will fit best to a line. For WT 3Dpol, the data fit best to a two-proton-transfer model (Fig. 4B). We identified Lys-359 on conserved structural motif D of 3Dpol as the putative general acid. If Lys-359 is the general acid, then by changing this residue to Leu, we should lose the descending limb of the pH rate profile and observe only one proton transferring in the transition state for nucleotidyl transfer. Both of these predictions were observed experimentally (see K359L in Fig. 4A and B). Since the first report of the two-metal-ion mechanism for nucleotidyl transfer was proposed, it has been generally accepted that active-site residues only contribute indirectly to the chemistry of nucleotidyl transfer by serving as ligands to one or more of the divalent cations required for catalysis. Our observation of a general acid in PV RdRp represents the first example of an active-site residue contributing directly to nucleotidyl transfer. This discovery is the first extension of the chemical mechanism for nucleotidyl transfer in almost three decades. Therefore, we felt obligated to determine if this observation was unique to the RdRp. We established three additional polymerase systems in the lab for this purpose [37]: the RT from human immunodeficiency virus (HIV RT), the DdDp from bacteriophage RB69 (RB69 DdDp), and the DdRp from bacteriophage T7 (T7 DdRp). T7 DdRp is a model for all A-family polymerases, and RB69 is a model for all B-family polymerases. All of these enzymes exhibited a SDKIE that was dependent on a conserved Lys residue (Table 1) [37]. Moreover, by changing this Lys residue to Leu, the proton inventory went from two to one (Table 1). These and other experiments lead us to conclude that all nucleic acid polymerases employ general acid catalysis for nucleotidyl transfer. Together, these studies validate the PV RdRp system as a powerful model system not only for RdRps but for all classes of nucleic acid polymerases. Our studies have revealed an unexpected role for the general acid in nucleotide incorporation fidelity (Fig. 5A and B) [37,56,57]. We envision other possible roles for this residue in coupling the chemical step to other aspects of the kinetic mechanism for nucleotide incorporation.

Table 1 Kinetic Analysis of PV RdRp, HIV-1 RT, RB69 DdDp, and T7 DdRp Supports General Acid Catalysis in Nucleotidyl Transfer [37] PV RdRp HIV RT RB69 DdDp T7 DdRp Parameter Measured

Lys-359

Leu-359

Lys-220

Leu-220

Lys-560

Leu-560

Lys-631

Leu-631

50  5

1  0.1

60  5

0.3  0.1

200  10

0.10  0.01

60  5

0.6  0.1

Kd,app (μM)

200  20

700  80

71

52

40  5

1000  100

300  30

5.0  1.0  104b

SDKIEc

3.0  0.3

2.5  0.3

2.2  0.4

1.8  0.4

4.2  0.2

1.8  0.2

5.2  0.5

2.6  0.5b

PId

2

1

2

1

2

1

2

nde

1

kpol (s ) a

Kd,app is for (d)ATP. Kd,app, kpol, and SDKIE values listed for T7 K631L were obtained by using data collected with 80 mM ATP, a subsaturating concentration. c SDKIE is solvent deuterium kinetic isotope effect, calculated as kobs in H2O/kobs in D2O at saturating [(d)ATP]. d PI is proton inventory, calculated from a plot of kn/k0 as a function of n. The data were fit to a modified Gross–Butler equation for either a two-proton-transfer model or a one-proton-transfer model. The value reported is the proton-transfer model that best fits the data. e Not determined. a

b

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Fig. 5 PV RdRp Lys-359 in structural motif D is a determinant of RdRp catalytic efficiency and fidelity. (A and B) K359R PV RdRp is slower and more faithful than the WT lysine-containing enzyme. RTP stands for ribavirin triphosphate. (C) Motif D changes conformation upon nucleotide binding. (D) The Met-354 resonance serves as a reporter of the catalytically competent complex. A substantial change in this resonance is only observed when the correct NTP is bound. Panels (A and B): Adapted from S.A. Weeks, C.A. Lee, Y. Zhao, E.D. Smidansky, A. August, J.J. Arnold, C.E. Cameron, A polymerase mechanism-based strategy for viral attenuation and vaccine development, J. Biol. Chem. 287 (38) (2012) 31618–31622. Panel (D): Adapted with permission from X. Yang, J.L. Welch, J.J. Arnold, D.D. Boehr, Long-range interaction networks in the function and fidelity of poliovirus RNA-dependent RNA polymerase studied by nuclear magnetic resonance, Biochemistry 49 (43) (2010) 9361–9371. Copyright (2010) American Chemical Society.

Collectively, our mechanistic studies were consistent with an active-site general acid that not only contributed to the efficiency of nucleotidyl transfer but also the fidelity of nucleotide selection. There was one problem. None of the beautiful structures of the picornaviral RdRps solved by Peersen and his group showed Lys-359 in a position consistent with hydrogen bonding to the incoming NTP [58–60]. In contrast, structures of the human norovirus RdRp in the absence and presence of NTP showed a dramatic change in the conformation of motif D, which was modeled by us for PV RdRp (Fig. 5C) [61]. This observation inspired the hypothesis that the rate-limiting conformational changes observed kinetically might actually correspond to structural changes that may not be revealed by X-ray crystallography and led to our pursuit of dynamics–function relationships of the RdRp. To this end, we established a collaboration with Dr. David Boehr in our Department of Chemistry at PSU. Together, our laboratories succeeded in applying the structural and dynamical wherewithal of multidimensional heteronuclear nuclear magnetic resonance spectroscopy to

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the study of PV RdRp, a 52 kDa protein [57,62,63]. Our initial experiments monitored 13C-labeled methionines in the protein. This choice was made because we hoped to use Met-354 in motif D as a reporter for any conformational changes of this motif, which contains Lys-359 (Fig. 5C). In going from unbound enzyme to the enzyme–RNA complex, a small change in the resonance for Met-354 was noted (Fig. 5D) [57,63]. Importantly, a huge change was observed for this resonance in the enzyme–RNA–NTP ternary complex [57,63], consistent with a large conformational change of motif D in the catalytically competent complex. Additional studies further implicated a hydrogen bonding interaction between the epsilon amino group of Lys-359 and the beta phosphate of the NTP [57].

3. KINETIC AND STRUCTURAL DETERMINANTS OF PV RdRp FIDELITY We have solved the complete kinetic mechanism for correct and incorrect nucleotide incorporation catalyzed by PV RdRp. The kinetic mechanism for correct nucleotide incorporation is shown in Fig. 6. At least five steps are employed for a single-nucleotide-addition cycle. Nucleotide, in this case ATP, binds (step 1) followed by a conformational change step (step 2) that produces a complex competent for nucleotidyl transfer. Chemistry occurs (step 3) followed by a second conformational change step (step 4) that we currently assign to translocation. The cycle ends with release of PPi (step 5). Qualitatively, this mechanism is identical to that observed for other classes of nucleic acid polymerases [28–32]. However, this kinetic mechanism differs from others in that both the prechemistry conformational change step (step 2) and chemistry (step 3) are partially rate limiting for nucleotide addition. We have employed two methods to interrogate step 2 (Fig. 7A): isotope trapping (Fig. 7B) and EDTA vs HCl quenching

Fig. 6 Complete kinetic mechanism for single-nucleotide incorporation catalyzed by PV RdRp. Steps 2 and 3 are both partially rate limiting for nucleotide incorporation. Adapted from J.J. Arnold, M. Vignuzzi, J.K. Stone, R. Andino, C.E. Cameron, Remote site control of an active site fidelity checkpoint in a viral RNA-dependent RNA polymerase, J. Biol. Chem. 280 (27) (2005) 25706–25716.

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Fig. 7 The prechemistry, conformational change step as a fidelity checkpoint for PV 3Dpol. (A) Minimal mechanism for pulse-chase analysis. (B) Kinetics of pulse-chase (•) and pulse-quench () using ATP. 4 μM PV RdRp was incubated with 20 μM sym/sub (10 μM duplex) and rapidly mixed with 130 μM [α-32P]ATP (3.8 Ci/mmol). At the indicated times, reactions were either chased by addition of 20 mM ATP final concentration or quenched by addition of 1 N HCl. After addition of the chase solution, the reaction was allowed to proceed for an additional 30 s at which time the reaction was quenched (Continued)

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(Fig. 7C and D). These experiments reveal a reduction in the equilibrium constant for step 2 when nucleotides with an incorrect sugar configuration or incorrect base are incorporated, thus revealing the use of the prechemistry, conformational change step as a kinetic determinant of nucleotide incorporation fidelity. Subsequent studies showed that fidelity was determined in large part by the stability of the *ERnNTP complex, which is determined by the rate constant, k2 [64]. All nucleic acid polymerases have a motif equivalent to conserved structural motif A found in RdRps and RTs. This motif extends from the (deoxy)ribose-binding pocket to the top of the catalytic site (Fig. 1B). In the ribose-binding pocket, at least one residue (Asp-238) stabilizes the ribose configuration so that the 30 -OH of incoming nucleotide can interact with the motif A backbone. In the catalytic site, both the backbone and side chains of motif A serve as ligands to metal A, the divalent cation required for catalysis. We have shown that both amino acid substitutions in motif A and the nature of the divalent cation can change the equilibrium constant for the prechemistry, conformational change step [65]. We currently assign step 2 to reorientation of the triphosphate into a position competent for nucleotidyl transfer. These and other studies link motif A to step 2 of the kinetic mechanism, thus revealing conserved structural motif A as a structural determinant of nucleotide incorporation fidelity.

Fig. 7—Cont'd by addition of 1 N HCl. Immediately after addition of HCl, the solution was neutralized by addition of 1 M KOH and 300 mM Tris. The solid line represents the kinetic simulation of the data to the mechanism shown in (A) with step 2 (K2) equal to 3 and k+3 equal to 30 s1. The simulated curve of the pulse-quench data predicts the rate of formation of ERn+1; the simulated curve of the pulse-chase data predicts the rate of formation of *ERnNTP and ERn+1. (C) 20 -dATP. A surrogate for the pulse-chase pulse-quench reaction was used for both 20 -dATP and GTP whereby the reaction was quenched by either EDTA (•) or HCl (). The solid line represents the kinetic simulation of the data to a mechanism with step 2 (K2) equal to 0.4 and k+3 equal to 30 s1. The dotted line represents the kinetic simulation of the data fit to a mechanism with step 2 (K2) equal to 3 and k+3 equal to 10 s1. (D) GTP. The solid line represents the kinetic simulation of the data fit to a mechanism with step 2 (K2) equal to 0.05 and k+3 equal to 30 s1. The dotted line represents the kinetic simulation of the data to a mechanism with step 2 (K2) equal to 3 and k+3 equal to 3 s1. Reprinted with permission from J.J. Arnold, D.W. Gohara, C.E. Cameron, Poliovirus RNA-dependent RNA polymerase (3Dpol): pre-steady-state kinetic analysis of ribonucleotide incorporation in the presence of Mn2+, Biochemistry 43 (18) (2004) 5138–5148. Copyright (2004) American Chemical Society.

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4. REMOTE-SITE CONTROL OF AN ACTIVE-SITE FIDELITY CHECKPOINT We have isolated two PV mutants: one with an antimutator phenotype [64] and the other with a mutator phenotype [66]. The antimutator phenotype was conferred by a change of Gly-64 to Ser (G64S); the mutator phenotype was conferred by a change of His-273 to Arg (H273R). Surprisingly, both Gly-64 and His-273 are located in the fingers domain of PV RdRp, not the active site (Fig. 8). We will use antimutator synonymously with high fidelity and mutator synonymously with low fidelity. As expected, G64S PV was less sensitive than WT PV to ribavirin, a mutagenic ribonucleoside [64], and H273R PV was more sensitive to ribavirin than WT PV [66]. The mutation frequency observed in cell culture was consistent with the mutation frequency observed for the corresponding PV RdRp derivatives determined biochemically [64,66]. The surprise was that the change in fidelity appeared to be no greater than threefold. Detailed kinetic analysis, in particular isotope-trapping experiments, of these PV RdRp derivatives revealed a change in the equilibrium constant for the prechemistry, conformational change step (step 2) as the kinetic basis for the change in nucleotide incorporation fidelity PV RdRp. These data

Fig. 8 Fidelity determinants located at sites remote from the catalytic site. The location of Gly-64 (green, light gray in the print version) and His-273 (red, dark gray in the print version) is indicated. Reprinted from V.K. Korboukh, C.A. Lee, A. Acevedo, M. Vignuzzi, Y. Xiao, J.J. Arnold, S. Hemperly, J.D. Graci, A. August, R. Andino, et al., RNA virus population diversity, an optimum for maximal fitness and virulence, J. Biol. Chem. 289 (43) (2014) 29531–29544.

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provide indisputable, genetic evidence for a role of the prechemistry, conformational change step as a determinant of PV RdRp fidelity. Moreover, these data reveal remote-site control of this active-site fidelity checkpoint.

5. RdRp FIDELITY IS A DETERMINANT OF VIRAL VIRULENCE Our results suggested that PV RdRp fidelity altered viral population dynamics in cell culture. We were therefore interested in determining the impact of altered fidelity on virus multiplication in a more complex environment. For these experiments, we collaborated with Dr. Raul Andino (UCSF). Dr. Andino developed a new strain of mice expressing the PV receptor, cPVR mice [67]. Infection of these mice with WT PV at a dose of 108 plaque-forming units (pfu) kills all mice in 5 days, with 50% of the mice developing paralysis (PD50) at a dose 107 pfu (Table 2). The lowfidelity mutant was attenuated in this highly permissive animal model (Table 2). The PD50 was increased on the order of two logs for H273R PV (Table 2). These data suggest that RdRp fidelity is optimized. Moreover, these data show that RdRp fidelity is a determinant of virulence, a completely novel concept at the time [66]. A great deal of data exist that point to the RdRp as a virulence determinant, even in the case of the 1918 Spanish flu strain [69–71]; however, the mechanism is unknown. What an impact two mutants can have on our understanding of the chemistry and biology of a virus.

6. RdRp FIDELITY MUTANTS ARE VACCINE CANDIDATES The practical application of any attenuated virus is vaccination. Therefore, mice were inoculated with the high-fidelity or low-fidelity virus. Four Table 2 RdRp Fidelity Is a Determinant of Viral Virulence [66,68] Virus PD50a

WT

2  107

G64S

1  107

H273R

1  109

a Dose of virus in plaque-forming units that leads to paralysis in 50% of inoculated animals.

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weeks or 6 months thereafter, these mice were challenged with five times the lethal dose of WT PV. In most cases, 100% of the mice were protected [56,66,68]. We conclude that an understanding of RdRp fidelity is no longer just a fundamental question of polymerase enzymology but now underpins a novel strategy for vaccine development. The fact that the first PV fidelity mutants were caused by amino acid substitutions in the RdRp that were not only remote from the active site but were also not conserved across RdRp family of enzymes. In contrast, the general acid of PV, Lys-359, is conserved (Fig. 9A). The K359R PV mutant produced infectious virus [56]. There was a longer eclipse phase of the life cycle without a substantial change in the kinetics of the exponential phase of the life cycle and culminating with a virus yield one log lower than wild type (Fig. 9B). The ability to produce reasonable titers of virus permitted us to determine if the reduced kinetics of virus production caused attenuation of the virus in the cPVR transgenic mouse model [67]. To evaluate virus multiplication and virulence in this model, we performed an intraperitoneal inoculation as this permits the largest virus inoculum to be used (3 mL). K359R PV was attenuated relative to WT PV (Fig. 9C) [56]. The meaning of this phenotype was unclear. Was K359R PV noninfectious or just incapable of causing disease? We reasoned that if K359R PV was infectious, then these infected animals might be protected from challenge with WT PV. Indeed, at a dose of 1  108, K359R PV protected 100% of animals from WT PV-induced paralysis (Fig. 9D). This observation supports the conclusion that we have identified a polymerase mechanism-based strategy for PV attenuation and vaccine development [56]. We compared protection by K359R PV to the type 1 Sabin strain of PV and found that there was no more than a one-log difference in the dose required for protection (Fig. 9D). The K359R PV has one amino acid substitution, whereas Sabin 1 PV has 21 amino acid substitutions [72]. In collaboration with Dr. Andrew Macadam (National Institute of Biological Standards and Control, UK), we showed that K359R PV was as attenuated as Sabin 1 PV using an intraspinal route of inoculation, providing some evidence for the safety of this approach to viral attenuation (unpublished observations). We have realized a polymerase mechanism-based strategy for viral attenuation. The question now is whether or not this knowledge can be applied to other viral systems.

Fig. 9 Conserved structural motif D as a target for viral attenuation and vaccine development. (A) Conservation of the motif D lysine in viral RdRps. (B) K359R PV exhibits a delayed growth phenotype. (C) K359R PV is attenuated. (D) K359R PV elicits a protective immune response. Adapted from S.A. Weeks, C.A. Lee, Y. Zhao, E.D. Smidansky, A. August, J.J. Arnold, C.E. Cameron, A polymerase mechanism-based strategy for viral attenuation and vaccine development, J. Biol. Chem. 287 (38) (2012) 31618–31622.

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7. CORRELATED MOTIONS OF FUNCTIONALLY IMPORTANT MOTIFS OF THE RdRp MAY BE A DETERMINANT OF NUCLEOTIDE INCORPORATION FIDELITY Our studies of incorporation fidelity revealed the ability of remote sites of PV RdRp to control conformational changes that occur at the active site. We made a similar observation in our studies of dihydrofolate reductase [73]. In that system, it turned out that remote-site control of active-site function was mediated by protein dynamics on multiple timescales [74]. Therefore, our observations with 3Dpol inspired the provocative hypothesis that polymerase incorporation fidelity might be controlled by dynamics as well. As a first step in this direction, we turned to molecular dynamics simulation (MD). MD is a powerful, computational approach to study protein dynamics [75,76]. In spite of all of the structural information available for polymerases, MD has primarily been applied to polymerases involved in DNA repair [77–80]. We have exploited the availability of structures for polymerases from multiple picornaviruses, including PV [60], Coxsackievirus B3 (CVB3) [81], human rhinovirus type 16 (HRV16) [82], and foot-and-mouth disease virus (FMDV) [83] to determine if conserved dynamical properties exist for these enzymes that could be important for function. We performed an all-atom (or atomistic) MD study of the four picornaviral RdRps listed earlier using the protocol illustrated in Fig. 10 [84,85]. All simulations were performed without substrates or metal ions. First, we performed the simulation of PV RdRp for 25 ns. The overall structure was maintained on the timescale of this experiment. The flexibility of the enzyme structure was determined by using the MD data to calculate B-factors. These data were in good agreement with B-factors obtained from X-ray data, suggesting that the force field and the protocol employed for the simulation are reasonable. Next, we performed MD on the other picornaviral RdRps. However, in each case we evaluated an incrementally shorter timescale: FMDV 3Dpol (20 ns), CVB3 3Dpol, and HRV16 3Dpol (14 ns). The shortest timescale (14 ns) did not alter the reliability of the data, decreasing substantially the amount of computational time required to perform the experiment. All systems reached equilibrium in 2–4 ns. The temperature, energy, and density of the system were monitored throughout the simulation, and all of these parameters displayed well-behaved dynamic trajectories.

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Fig. 10 Molecular dynamics protocol. Flow chart describing the protocol used to perform the all-atom MD simulations. Programs LEaP, SANDER, PMEMD, and PTRAJ are modules in the AMBER simulation package [84].

Principal component analysis (PCA) permits us to separate large, concerted motions from the noise [86,87]. The analysis is carried out by calculating the correlation matrix for the displacements of Cα atoms from the averaged structure. Treating the matrix as an eigenvalue problem results in a set of eigenvectors (or principal components), defining a new coordinate space to describe the observed motions in the simulation. The number of PCs is equivalent to three times the number of amino acid residues in the data set. However, when the PCs are arranged in descending order, most of the variance (displacements/motion) in the data can be accounted for in a small fraction of the PCs. In the case of the simulations for the four picornaviral RdRps (461 residues), the first six PCs accounted for more than 50% of the global motion in the polymerases. Thus, we restricted our analysis to these PCs. In doing so, we restricted our focus to the major motions of each polymerase, facilitating comparative analysis. Our premise was that if motion is important, then it should be a major motion and should be conserved. Because the magnitude of the values for the displacements varied for each polymerase, direct visual comparison of the top six PCs for each polymerase was difficult. To address this issue, the sum of the top six PCs were normalized to the average of the sum of the five least flexible residues. This analysis revealed a remarkable conservation in the dynamics of the four polymerases [85]. In order to determine if correlated motions existed in the polymerases, we calculated dynamics cross-correlation maps (DCCMs) [88].

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The DCCMs for three of the four picornaviral RdRps are shown in Fig. 11. As observed for the PCA, the DCCMs were remarkably similar. Correlated motions were observed between several conserved structural motifs, including A and D (region I in Fig. 11) and F and G (region II in Fig. 11). The amino acid sequences of CVB3 RdRp and FMDV RdRp are 74% and 30%, respectively, identical to that of PV RdRp. Interestingly, the DCCM for PV G64S RdRp exhibited substantial differences from PV RdRp [85]. The ability for MD to reveal conserved dynamics within this family of polymerases engenders confidence in our protocol. Moreover, these results provide evidence for evolution of structure and dynamics, a very exciting finding with significant implications for the existence of conserved dynamics in other polymerase systems. In other systems in which correlated motions have been observed, the correlated motions are often coordinated by virtue of the existence of a global network of interacting amino acid residues [89,90]. We have been able to identify such a network in PV RdRp [85]. Interestingly, Gly-64 and His-273, remote-site residues that influence the prechemistry, conformational change step, are located in this network. Importantly, disruptions in the network resulting from amino acid substitutions at these positions cause changes in correlated motions observed between conserved structural motifs (unpublished observations). Together, these observations lead us to propose that this network may be responsible for this and perhaps other conformational changes required for RdRp function.

8. NUCLEOTIDE BINDING-OCCLUDED AND BINDINGCOMPETENT STATES OF THE RdRp–RNA COMPLEX AS DETERMINANTS OF FIDELITY Our MD experiments have revealed two states of the PV RdRp– RNA complex based on the conformation and dynamics of conserved residues in conserved structural motifs A (Asp-238), B (Asn-297), D (Lys-359), E (Lys-375), and F (Arg-174) [91]. The first state occludes binding of the incoming NTP because of an ionic interaction between Asp-238 and Arg-174 (binding occluded in Fig. 12A). The second state is competent for NTP binding as the 238–174 interaction is broken, causing a rearrangement of the pocket that includes movement of Lys-359 toward the active site (binding competent in Fig. 12A). Closure of motif D to position Lys-359 to serve as the general acid would then produce the catalytically competent state (catalytic in Fig. 12A) [57]. The distance between

Fig. 11 See legend on opposite page.

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the Asp-238 and Arg-174 side chains as a function of time is indicative of the interconversion between the NTP-binding occluded (points below the red, gray in the print version, line in Fig. 12B) and competent (points above the red, gray in the print version, line in Fig. 12B) states. In the absence of RNA and NTP, the enzyme samples both states, but the occluded state is favored 9:1 (see WT in Fig. 12B). Addition of RNA increases the sampling frequency such that the competent state is present 40% of the time (see WT-RNA in Fig. 12B). In the mutator polymerase, the competent state is present 98% of the time (see H273R-RNA in Fig. 12B). By analyzing the differences in the average structures for the WT and H273R enzymes, it is evident that accommodation of the larger side chain at position 273 leads to a chain reaction that disconnects the network of interactions stabilizing the occluded conformation (direction of red, gray in the print version, arrows in Fig. 12C indicate the change from WT, gray, to H273R, black). From these data, we suggest that at least two structural changes comprise the fidelity checkpoint observed kinetically and that the equilibrium between occluded and competent state may be a determinant of fidelity.

Fig. 11 Conservation of correlated motions in picornaviral RdRps. The cross-correlation of the atomic displacement vectors of Cα atoms was examined. The pair-wise correlation of the displacements of Cα atoms relative to a reference structure was calculated and used to build the dynamic cross-correlation matrix (DCCM). Each element of the matrix is correlation between displacements of Cα atoms of a residue pair (ij). The matrix is plotted as a color-coded map, shown at the top panel for the three polymerases: PV, CVB3, and FMDV. For completely correlated motions the correlation score measures (+1), whereas for completely uncorrelated motions the score is (1). The maps are coded with color gradient for the correlation scores (+1 more red, gray in the print version; 1 more blue, dark gray in the print version). The similarity between the correlations observed in the three simulated polymerases can be inferred from the comparable patterns of the color distributions in the three maps. Two regions I and II with positive correlations are labeled in the maps. Region I corresponds to the coupled motion of motifs A and D; region II refers to motifs F and G. The locations of the different motifs are indicated by the gray bars at the top and right of the maps. The bottom panel shows the structures of the three polymerases with the fingers, palm, and thumb subunits colored green (light gray in the print version), blue (dark gray in the print version), and red (gray in the print version), respectively. Despite the diverse sequence identity between the different picornaviral polymerases (30–74%), the similarity of the overall structure and dynamics is noteworthy. The top panel of the figure was created using MATLAB 7.6. Adapted from I.M. Moustafa, H. Shen, B. Morton, C.M. Colina, C.E. Cameron, Molecular dynamics simulations of viral RNA polymerases link conserved and correlated motions of functional elements to fidelity, J. Mol. Biol. 410 (1) (2011) 159–181. Copyright (2011), with permission from Elsevier.

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Fig. 12 Physical mechanism of PV RdRp fidelity: an hypothesis. (A) Active-site conformational changes revealed by MD simulations of PV RdRp–RNA binary complex. (B) Time evolution of the distance between NH2 of Arg-174 and OD1 of Asp-238 atoms during 150 ns MD simulations of the free WT and the binary complexes of WT and H273R RdRps. (C) The average structures of WT-RNA (gray) and H273R-RNA (black) observed during the MD simulations are superimposed. Direction of arrows indicates the change from WT-RNA to H273R-RNA. Adapted from I.M. Moustafa, V.K. Korboukh, J.J. Arnold, E.D. Smidansky, L.L. Marcotte, D.W. Gohara, X. Yang, M.A. Sanchez-Farran, D. Filman, J.K. Maranas, et al., Structural dynamics as a contributor to error-prone replication by an RNA-dependent RNA polymerase, J. Biol. Chem. 289 (52) (2014) 36229–36248.

Worth noting, the dynamics sampled by the PV WT RdRp or complexes thereof are restricted to those of relevance to the next step in the reaction coordinate [91]. For example, PV WT RdRp samples conformations required for nucleic acid binding. Once nucleic acid is bound, however, PV WT RdRp–RNA complex samples states competent for nucleotide binding [91].

9. CONSERVED, ACTIVE-SITE DETERMINANTS OF RdRp INCORPORATION FIDELITY MAY EXIST THAT CAN BE TARGETED FOR VIRAL ATTENUATION One of the most exciting advances made during our studies of RdRp fidelity has been the formulation of a hypothesis linking the structural dynamics and accessibility of the NTP-binding site to fidelity [91]. WT RdRp cycles between occluded (Fig. 13A) and competent (Fig. 13B) conformations. Side chains indicated interact with one or more portions of the incoming nucleotide. In the binding-occluded conformation, all of these residues interact with each other. Therefore, a stable shift of the equilibrium

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Fig. 13 Proposed structural dynamics in the transition from NTP binding-occluded state to NTP binding-competent state employs conserved residues of the RdRp. All residues shown are conserved among RdRps of positive-strand viruses. (A) NTP bindingoccluded state. (B) NTP binding-competent state. Adapted from I.M. Moustafa, V. K. Korboukh, J.J. Arnold, E.D. Smidansky, L.L. Marcotte, D.W. Gohara, X. Yang, M.A. SanchezFarran, D. Filman, J.K. Maranas, et al., Structural dynamics as a contributor to error-prone replication by an RNA-dependent RNA polymerase, J. Biol. Chem. 289 (52) (2014) 36229–36248.

to the binding-competent conformation requires binding of a correct nucleotide. When a nucleotide with an incorrect sugar configuration or base binds, one or more interactions in the binding-occluded conformation will not be disrupted. Partial disruption may preclude a stable shift to the binding-competent conformation. Therefore, an RdRp derivative that favors the binding-occluded state should exhibit a fidelity higher than WT. Conversely, one that favors the binding-competent state should exhibit a fidelity lower than WT. A central, stabilizing determinant of the occluded state is the interaction between Asp-238 (motif A) and Arg-174 (motif F) (Fig. 13A). These same residues are critical for stabilizing the bound NTP (Fig. 13B), consistent with our suggestion that binding of a correct nucleotide might diminish return to the occluded state and nucleotide release. Importantly, essentially all of these residues are conserved among the RdRps of positive-strand RNA viruses. It may be possible to identify substitutions of one or more of these residues that create fidelity variants that can serve as vaccine candidates for any RNA virus, especially those that have yet to emerge.

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ACKNOWLEDGMENTS For nearly 20 years, C.E.C. and J.J.A. have worked to elucidate the fundamental principles responsible for the specificity, chemistry, and biology of the viral RdRp. The summary presented here would not have been possible without the collaborations that we have had with graduate and postdoctoral students and colleagues both here at PSU and elsewhere. Worth special mention are Raul Andino (UCSF), David Boehr (PSU), Coray Colina (PSU) as these scientists have both enabled our technical capabilities and have enriched our research. Our studies of the RdRp have been supported by Grant AI045818 from NIAID, NIH.

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AUTHOR INDEX Note: Page numbers followed by “f ” indicate figures, and “t” indicate tables.

A Abdulrehman, S.A., 37 Abe, Y., 4, 11–13, 16, 34t Abeyrathne, P.D., 182–184 Acevedo, A., 307–309, 308t Aebersold, R., 34t, 53, 57 Agaronyan, K., 278–279 Agarwal, P.K., 313 Agarwal, T., 194, 210–212, 218 Aggarwal, A.K., 207–208 Aguilar, P.V., 308 Ahnert, P., 104–105 Aitchison, J.D., 193, 215 Akabayov, B., 95, 99, 102–107, 110, 112–114, 120–121, 123 Akabayov, S.R., 95, 99, 114 Akeson, M., 146 Akimitsu, N., 11–13 Akiyama, M.T., 44 Albert, A., 160–161 Alberts, B.M., 43, 61, 109–110, 119, 216–217, 273–275 Alcamı´, A., 149 Ali, S.S., 92 Allemand, J.F., 123–124, 215–216 Allen, G.J., 16–18 Allen, J.L., 95 Allen, K.N., 296–297 Allen, L.M., 34t, 67 Almeida, D.V., 48 Almiron, M., 13 Alonso, J.M., 303–304 Altschul, S.F., 266f Altuvia, S., 13 Alwood, A., 58 Amadei, A., 312–313 Amal, I., 48 Amarasiriwardena, C.J., 91–92 Anand, R.P., 243–244 Anderson, D.L., 138–139 Anderson, K.S., 297–298, 304–306 Anderson, S., 277–279

Anderson, S.G., 57–58 Andino, R., 304–309, 308t Andjelkovic, J., 42, 57 Anjum, R.S., 186–187 Ansari, A., 41 Anstey-Gilbert, C.S., 67 Antonicka, H., 281 Antony, E., 42 Arad, G., 34t Arai, K., 14, 34t, 37, 41 Arai, N., 34t Araki, H., 192, 194–196, 203–206, 215 Aravind, L., 38–39, 54–55, 193–194, 215–216 Archer, R.H., 13 Argiriadi, M.A., 48 Arias, A., 311 Arias, E.E., 232, 237 Arias-Gonzalez, J.R., 144–145 Arias-Palomo, E., 14, 18–19, 37–38 Armstrong, A.A., 236, 239 Arneodo, A., 193 Arnold, E., 295–296 Arnold, J.J., 294–317, 302t, 308t Arora, K., 311 Arunkumar, A.I., 184 Arvai, A.S., 103 Asai, T., 4–5 Asensio, J.L., 160–161 Ason, B., 54–55 Atack, J.M., 67 Atanassova, N., 272 Atkinson, J., 34t, 68 Atlung, T., 10 Auchtung, J.M., 10 Audit, B., 193 August, A., 301, 307–309, 308t Augustin, M.A., 202 Ayyagari, R., 203, 205–206

B Backert, S., 273–275 Bae, B., 182 325

326 Bae, S.J., 92 Bahng, S., 42, 56f, 57–58 Bai, L., 108, 197–198 Bailey, S., 12f, 14, 19–20, 37–39, 39f, 49–53, 104–105 Bajic, G., 19–20 Baker, A., 193 Baker, T., 294 Baker, T.A., 34t, 36–37, 40, 45, 66–67, 138, 193–194, 201, 216–217 Bakht, S., 149 Balistreri, L., 311 Ballesteros-Plaza, D., 149–150, 159–160 Balvocˇiu¯t_e, M., 49 Bambara, R.A., 44–45 Bamford, D.H., 138, 153–155 Bamford, J.K., 138 Bando, M., 203–205 Banks, G.R., 201–202 Baradaran, K., 39–40, 100–101 Barcena, M., 14 Barclay, E., 205 Barnoux, C., 53 Barrett, M.P., 102 Barry, E.R., 170–171 Barry, J., 43, 61 Basler, C.F., 308 Basta, T., 138 Bates, D.B., 4–5, 103 Bath, C., 138 Bauer, A., 48 Bazin, A., 19–20 Beach, D., 232 Beard, W.A., 299, 311 Beattie, T.R., 36 Beauchamp, B.B., 91, 93–95 Beaudry, A.A., 58, 294 Beck, E., 2–3 Beck, J.L., 34t, 38, 42–43, 45–48, 51–53 Beckett, E., 92 Beckman, R.A., 294 Bedinger, P., 43, 61 Beerbaum, M., 34t, 42 Beese, L.S., 34t, 54, 66, 147–148, 295–296, 299 Beeson, D., 102 Bekker-Jensen, S., 48 Bell, H., 68

Author Index

Bell, L., 273–275, 282 Bell, S.D., 170–187, 173f, 193, 198, 201–203 Bell, S.P., 171–172, 174, 177, 185–186, 196–197, 236 Benkovic, P.A., 297–299, 304–306 Benkovic, S.J., 59–60, 63, 104–105, 108–110, 113–114, 121–122, 124, 193–194, 216–219, 295–299, 304–306, 311, 313 Bensimon, D., 108 Bentley, D., 205 Ben-Yehuda, S., 160 Berendsen, H.J., 312–313 Berger, J.M., 7–8, 11, 12f, 14–15, 18–19, 34t, 36–40, 62, 115–117, 177–178, 185–186, 192–194, 196, 198–200, 208–209, 212–215, 214f Berghuis, B.A., 68 Bergmans, H.E., 2–3 Berkowitz, S.A., 95, 102–107, 110, 112–114, 120–121, 123 Berlatzky, I.A., 160 Berman, A.J., 141–152, 155–156 Bermudez, V.P., 197–198, 200, 212–215 Bernad, A., 54, 138, 140–145, 153, 155–157 Bernander, R., 16–18, 174–176 Bernstein, J.A., 100–102, 107, 115–117 Berriman, M., 172 Bertram, J.G., 54–55 Besnard, M., 58 Bessman, M.J., 66, 91 Betterton, M.D., 107 Beuning, P.J., 49 Bhatnagar, S.K., 91 Bhattacharyya, S., 39–40, 62–63 Bi, L.-J., 48 Bi, X., 239–240 Bienko, M., 232–233, 236, 239–240 Bienstock, R.J., 260 Billeter, S.R., 313 Bird, L.E., 20 Bird, R.E., 2–3 Biswas, E.E., 39–40 Biswas, S.B., 18, 34t, 37–40 Biswas, T., 37 Biswas-Fiss, E.E., 18 Bjornson, K.P., 107

Author Index

Blaesing, F., 11 Blanco, L., 54, 138–158, 273–275 Blasco, M.A., 54, 138–139, 141–145, 193 Blattner, F.R., 32 Bleichert, F., 199–200 Blinkova, A., 218 Blinkowa, A.L., 55 Blinov, V.M., 58–59 Bloch, C.A., 32 Bloom, L.B., 54–55, 57–58 Blundell, T.L., 103, 265, 266f Boardman, A.P., 219 Bobst, A.M., 34t Bobst, E.V., 34t Bochman, M.L., 174, 194–196, 199–200 Bocquier, A.A., 201 Boehm, E.M., 232–248 Boehr, D.D., 301, 303–304, 313–315 Boezi, J.A., 201–202 Bogden, C.E., 34t, 37–38 Bogenhagen, D.F., 100, 277, 280 Bogutzki, A., 34t, 42 Boitano, M., 10 Bond, M.W., 34t Boneca, I.G., 13–14 Bonhoeffer, F., 34t Bonilla, C.Y., 13–14 Bonnin, A., 158 Bortner, C., 109–110 Bostina, M., 95, 102–107, 110, 112–114, 120–121, 123 Botchan, M.R., 182, 194–196, 198–200, 212–215, 214f, 219 Bouche, J.P., 38–39 Bourne, H.R., 265, 266f Bowmaker, M., 282–283 Bowman, G.D., 58–59, 208–210 Boye, E., 4–5, 8–9 Bradshaw, R.A., 193–194 Braman, J.C., 295–296 Bramhill, D., 3–4, 7, 11–13 Branscomb, E.W., 294 Brassinga, A.K., 10 Bratic, A., 260–261 Bratic, I., 273–275 Braun, R.E., 36 Bravo, A., 138–139 Bravo, R., 232

327 Bray-Ward, P., 149 Bressani, R., 39–40 Brewer, B.J., 193, 273–275, 282 Brewster, A.S., 182, 197–198 Brick, P., 66, 96–97, 112–113, 295–296, 299 Bricogne, G., 311 Brieba, L.G., 99 Brinkman, K., 259 Brissett, N.C., 38–39 Broda, P., 2–3 Broderick, P., 205 Brooke, R.G., 202 Brown, S.E., 14, 34t, 37–38, 51–52 Brown, T.A., 280–283 Brown, W.M., 256, 277–279 Browning, D.F., 8–9 Broyde, S., 311 Bruand, C., 218–219 Bruck, I., 48, 218–219 Bruice, T.C., 313 Brun, C., 282 Bruning, J.B., 235 Br€ uning, J.G., 68 Bruno, J.F., 149 Brusilow, W.S., 4, 9 Brutlag, D., 275–276 Bryan, S.K., 45–48, 206 Brzoska, A.J., 46–49, 58 Buc, H., 66 Buck, K.W., 294 Bueno, A., 219 Bugreev, D.V., 103 Bujalowska, D., 14, 37 Bujalowski, W., 14–15, 18, 36–38, 41–42 Bullard, J.M., 49–51, 58 Bulloch, E.M., 34t, 51–53 Bunting, K.A., 46–48 Burgers, P.M.J., 34t, 51–52, 60, 203, 205–208, 232, 238, 240 Burke, R.L., 109–110 Burkholder, A.B., 205–206 Burland, V., 32 Burnouf, D.Y., 46–48 Busby, S.J., 8–9 Bushnell, D.A., 299 Bussetta, C., 311 Bustamante, C.J., 37–38, 147–148

328 Byers, B., 273–275, 282 Bylund, G.O., 207–208

C Cable, M.B., 295–296 Cadman, C.J., 34t Cadwell, G.W., 4–5 Cai, J., 238 Cairns, J., 32 Caldentey, J., 153–155 Calhoun, L.N., 13 Calzada, A., 203–205, 219 Camacho, A., 158 Camara, J.E., 8–9 Cameron, C.E., 294–317, 302t, 308t Campbell, C.S., 203 Campbell, J.L., 206 Canard, B., 311 Cann, I.K., 103, 201 Cano, M.I., 193 Cantor, A.J., 34t, 57–59, 209–210 Cao, F.J., 144–145 Cao, Y., 4–5 Carazo, J.M., 14, 37–38, 197–198 Carballido-Lo´pez, R., 149–150, 159–161 Carl, P.L., 16–18 Caro, L., 2–3 Carr, A.M., 205–206 Carr, K.M., 15–18 Carr, P.D., 34t, 51–52 Carrascosa, J.L., 144–145 Carrodeguas, J.A., 100, 265, 269, 280 Carter, J.R., 34t, 53, 57 Caruthers, J.M., 261 Carvajal-Carmona, L.G., 205 Carver, T.E., 121–122 Case, D.A., 311, 312f Cassler, M.R., 8–9 Castelli, V., 58 Castilla-Llorente, V., 160–161 Castro, C., 298–301, 302t Causer, R.J., 36, 46–48, 57 Cazier, J.B., 205 Cech, T.A., 298 Ceci, P., 13 Celis, J.E., 232 Center, M.S., 92, 95 Cerami, A., 66

Author Index

Ceska, T., 68 Chaban, Y.L., 103 Chabes, A., 205–206 Chait, B.T., 193, 200, 210–215, 214f, 218 Chaloin, O., 48 Chalov, S., 53–54 Chambers, M.W., 4–5 Chang, D.D., 278–279 Chang, F., 185–186 Chang, P., 20 Chao, M.C., 9 Chapados, B.R., 238–239 Chapuis, J., 218–219 Chase, J.W., 34t, 41–42, 109–110 Chase, M.R., 49 Chastain, P.D., 61, 108–109, 119–122, 124 Chattoraj, D.K., 6f, 9 Chaurasiya, K.R., 34t Chazin, W.J., 201–202 Cheatham, T.E., 311, 312f Cheetham, G.M., 295–296 Chemla, Y.R., 147–148 Chen, C.L., 193 Chen, H., 45–48 Chen, J., 49 Chen, W.T., 12f, 14 Chen, X.S., 37, 197–198 Chen, Y.-Y., 48 Cheng, H.M., 8 Cheng, Y., 95, 102–105 Cherf, G.M., 146 Cherrier, M.V., 19–20 Chiancone, E., 13 Chikova, A.K., 53–54 Chilkova, O., 237–238 Chintakayala, K., 39–40 Chiraniya, A., 45–46 Chiu, W., 108–109 Cho, E., 13–14 Cho, W.-K., 60 Chodavarapu, S., 2–21, 36 Chong, J.P., 194–196 Chopra, R., 295–296, 299 Choudhury, N.R., 19–20 Chowdhury, K., 117–119 Christensen, J., 49–51 Chrysogelos, S., 41–42 Chuang, K., 7–8

329

Author Index

Chubb, D., 205 Chui, G., 201 Chung, Y.J., 295–296 Churchich, J.E., 99, 108–110 Ciesielski, G.L., 256–284 Ciesla, Z., 53 Clark, A.B., 205–206 Clark, A.D., 295–296 Clark, D., 245–246 Clark, P., 295–296 Clark, S., 205 Clark, T.A., 10 Clark-Walker, G.D., 267–269, 273–275 Clausen, A.R., 205–206 Clausen, M.F., 205–206 Clayton, D.A., 275–281 Cleary, J.M., 4, 9 Cohen, F.E., 265, 266f Coker, J.A., 175 Colina, C.M., 311–313 Collier, J., 10 Collingwood, D., 193 Comi, G., 102 Conaway, R.C., 201–202 Conrad, J., 49, 50f, 52–53 Conway, A., 193 Cook, P.R., 247–248, 281 Cook, T.E., 99, 108–110, 112, 114 Coons, M.M., 15, 18–19 Cooper, D.L., 34t Copeland, W.C., 201–202, 257–260 Corn, J.E., 34t, 39–40, 62, 115–117 Correa, R., 44, 55 Costa, A., 177–178, 182, 186, 192, 196, 198–200, 203–205, 212–215, 214f Coster, G., 185–186 Cotterill, S., 201 Coutard, B., 311 Cox, M.M., 34t, 41–42 Cox, N.J., 308 Craig, L., 103 Crampton, D.J., 102–107, 120–121, 263 Crews, S., 277–279 Crick, F.H., 43 Crombie, R., 34t, 55–57 Crooke, E., 4, 8–9 Crooke, S.T., 68 Croquette, V., 63, 108, 218

Cross, B., 68 Cross, H.F., 68 Crotty, S., 308–309 Crow, W., 44, 48–49 Crowther, J.A., 34t, 51–53, 57 Croxen, R., 102 Cruaud, C., 58 Cseresnyes, Z., 247–248 Cukalac, T., 138 Cullmann, G., 208–209 Cupp, J.D., 273–275 Curth, U., 34t, 42, 57–58, 265 Czerwonka, C., 3–4

D da Silva, M.S., 193 Dahl, J.M., 146, 148–149 Dahlberg, M.E., 297–298, 304–306 Dai, X., 219 Dallmann, H.G., 34t, 54–58, 218 Dalrymple, B.P., 34t, 45–48, 51 Dammerova, N., 37–38 Daniel, R., 160–161, 218–219 Daniels, D.S., 103 Danilova, O., 55 Darden, T., 52, 311, 312f Datta, S., 41–42 Dattilo, B.M., 201–202 d’Aubenton-Carafa, Y., 193 Davern, C.I., 16–18 Davey, M.J., 15–16, 18, 54–55, 194–200, 215 Davies, D.R., 49–51 Davis, B.M., 9 Davis, R.E., 303–304 Davis, R.W., 193 Day, B.J., 259 de Berardinis, V., 58 De Falco, M., 212 de Lange, T., 203 De Palma, A.M., 311 De Piccoli, G., 215 de Vega, M., 138–161, 142t de Vegabe, M., 138 Dean, F., 45–46 Dean, F.B., 149 Debyser, Z., 61, 95, 119 Deegan, T.D., 196

330 del Prado, A., 142t, 149–150, 152–153 Delagoutte, E., 107, 197–198 Demarre, G., 6f, 9 DePamphilis, M., 138–139 DePhamphilis, M.L., 193 Dephoure, N., 219 Derbyshire, V., 34t, 54, 66, 147–148 DeRose, E.F., 52–54 Dervan, J.J., 68 Dervyn, E., 218–219 Desai, A., 203 Desjardins, P., 277, 280–281 Deutscher, M.P., 66 Devos, K.M., 149 DeWyngaert, M.A., 92, 95 Dhar, S.K., 19–20, 37 Di Re, M., 280 Diaz, A., 67 Dieckman, L.M., 235, 240–241, 245–246 Diehl, W., 311 Dietrich, M., 54 Diffley, J.F.X., 172, 196, 236 Dillingham, M.S., 14–15, 193–194, 196–197 Dilworth, D.J., 215 Ding, F., 218 Dionne, I., 247 DiRuggiero, J., 103 Dixon, N.E., 14, 32–69, 34t, 50f Dobbins, S., 205 Doda, J.N., 277 Doherty, A.J., 38–39 Dohrmann, P.R., 44, 51, 55, 64–65 Domingo, E., 205, 311 Donate, L.E., 14 Donchenko, A.P., 58–59 Donczew, R., 9–10, 13–14 Dong, F., 37, 218 Dong, Z., 34t, 55–57 Donmez, I., 63, 100–101, 105–108, 112–113, 123–124, 197–198 Donnelly, P., 205 Doolittle, W.F., 170 Dorman, C.J., 92 Doublie`, S., 95–97, 99, 112–113, 117, 147–148, 299 Douzery, E., 277 Downey, C.D., 64–65

Author Index

Dragovich, P.S., 311 Dressler, D., 93 Drlica, K., 11–13 Drummond, M.H., 52–53 Du, J., 149 Du, Y., 149 Dua, R., 206 Duderstadt, K.E., 7–8, 36, 192 Dueber, E.L.C., 170–171, 176–178 Dufour, E., 141–144 Duggin, I.G., 175–176, 181 Dulin, D., 68 Dumas, L.B., 202 Dumas, P., 46–48 Dunaway-Mariano, D., 296–297 Dunn, J.J., 90, 101–102, 149 Dutta, A., 236 Duzen, J.M., 34t Dyall-Smith, M.L., 138 Dzantiev, L., 245

E Early, A., 196 Echols, H., 34t, 51–52 Eddy, M.J., 7 Eddy, S.R., 266f Edgell, D.R., 170 Edmondson, R.D., 200–201, 203–205, 215, 218–219 Egan, E.S., 9 Egan, J.B., 8–9 Egelman, E.H., 14, 37–38, 103–107, 120–121 Ehrlich, S.D., 13–14, 218–219 Eichman, B.F., 99 Eki, T., 201–202 El Houry Mignan, S., 58 Elia, A.E., 219 Eliason, W.K., 12f, 14–15, 19–20, 37–39, 39f, 104–105 Elledge, S.J., 219 Ellenberger, T., 95–97, 99, 102–110, 112–113, 115–121, 147–148, 299 Elshenawy, M.M., 34t, 44–48, 51–52, 68 Enemark, E.J., 14–15, 196–197, 200 Engler, M.J., 93–94, 96 Ennifar, E., 48 Enomoto, R., 103

Author Index

Eom, S.H., 54, 66–67, 147–148 Epa, V.C., 34t, 46–48 Epling, L.B., 196–197 Erie, D.A., 245 Ernst, S., 58 Errington, J., 13–14, 160–161, 218–219 Erzberger, J.P., 7, 11, 12f, 15, 18–19, 37–38, 178, 185–186, 193–194, 208–209 Escarmı´s, C., 138–139, 311 Estabrook, R.A., 313 Esteban, J.A., 138–140, 144–145, 153, 155, 157–158 Ettema, T.J., 170 Euro, L., 259–260 Evans, E.A., 92 Evans, R.J., 49–51 Everley, R.A., 219 Evrin, C., 185–186, 203–205, 215

F Fabrizi, G.M., 102 Falkenberg, M., 94, 100, 102, 264 Fan, L., 100, 265 Fang, L., 15–16, 18, 55–59, 149 Fangman, W.L., 193, 273–275, 282 Fanning, E., 8–9, 201–202 Fanning, T.G., 308 Farge, G., 94, 261–263 Fargo, D.C., 205–206 Farina, A., 194–198 Farnum, G.A., 259–260 Farr, C.L., 100, 264, 269–271 Farrugia, G., 205–206 Faruqi, A.F., 149 Fass, D., 34t, 37–39 Fay, P.J., 44–45 Fazio, N., 42 Feig, M., 15, 18–19 Felczak, M.M., 4, 11–14, 16 Feng, M., 68 Fenyo, D., 193 Fernandez-Cid, A., 199–200 Fernandez-Leiro, R., 49, 50f, 52–53 Fernandez-Millan, P., 261, 263 Fernendez-Cid, A., 174 Ferrari, E., 295–296 Ferrari, M.E., 41–42 Ferre, R.A., 311

331 Ferrer-Orta, C., 311 Ferrin, L.J., 299 Ferris, A.L., 295–296 Fien, K., 208–209 Fijalkowska, I.J., 32, 53 Filee, J., 193–194, 215–216 Filman, D., 313–317 Finkel, S.E., 8–9 Finkelstein, J., 44–45, 55, 57–58, 194, 210–218 Firshein, W., 6f, 158–159 Fischer, M.G., 138 Fishel, R., 245 Fisher, P.A., 238 Flamm, E.L., 8–9 Fleck, R.A., 212–215, 214f Flemming, C.S., 67 Flick, K., 206 Flores-Rozas, H., 245–246 Flower, A.M., 55 Flowers, S., 18 Fogg, M.J., 13–14 Foltman, M., 203–205, 215 Fontaine, E., 48 Ford, C.B., 49 Formosa, T., 43, 61, 203–205 Forterre, P., 193–194, 201, 215–216 Fox, G.E., 170 Franden, M.A., 34t, 53, 57, 61, 122 Frank, E.G., 219 Franklin, M.C., 49, 54, 147–148, 299 Freed, E.O., 295–296 Freedman, L.P., 13 Freire, R., 138–139, 153 French, S.L., 181 Frenkel, G.D., 91 Freudenthal, B.D., 234f, 235–236, 240–241 Frick, D.N., 39–40, 62–63, 100–101, 115–117, 123, 193–194, 201–202 Friedberg, E.C., 68 Friedman, A.M., 108–109 Friedman, J.M., 295–296 Frigola, J., 174, 184 Frisch, R.L., 44, 55 Fritzler, M.J., 232 Froelich, C.A., 196–197 Frols, S., 175–176 Fu, Y.V., 197–198, 200, 212–215

332 Fuchs, R.P.P., 46–48, 68 Fuhrman, S.A., 311 Fujii, S., 46–48, 68 Fujikawa, N., 3–4, 8, 11 Fujimitsu, K., 13 Fujiyama, A., 92 Fukuhara, N., 215 Fukuoh, A., 264, 273 Fukushima, S., 67–68 Fuller, C.W., 93–94 Funnell, B.E., 36–37 Furlong, D., 13 Furuichi, Y., 282–283 Fuste, J.M., 94, 272, 280–281 Futami, K., 282–283 Futterer, K., 41–42, 108–109

G Gaal, T., 13 Gabdoulkhakov, A., 37 Gabel, S., 52 Galal, W.C., 194–198 Gale, M.D., 149 Galletto, R., 14–15, 18, 36–37 Gambus, A., 200–201, 203–205, 218–219 Gan, G.N., 207 Gan, H., 205–206 Gan, P.H.P., 38–39 Ganai, R.A., 207–208 Gao, D., 34t, 49, 55, 57, 215–216 Gao, N., 199–200 Gao, S., 149 Garalde, D.R., 146 Garcia, A.E., 312–313 Garcia, J., 54 Garcı´a, J.A., 152, 159–160 Garcı´a, P., 153–155 Garcia-Alvarez, B., 202 Garcia-Sastre, A., 308 Garesse, R., 256, 273 Garg, P., 205–206, 240 Garmendia, C., 138, 140–141, 156–157 Garrett, R.A., 138 Garrido, N., 102 Gary, R., 232 Gary, S., 208 Gaspari, M., 264 Gaudier, M., 177–181

Author Index

Gawande, R., 49 Gawel, D., 91 Gay, N.J., 58–59 Geertsema, H.J., 95, 115, 122 Gefter, M.L., 53 Geider, K., 45–46 Gella, P., 158 Genau, H.M., 207 Genba, Y., 309 Genschel, J., 34t, 42 Gensler, S., 279–280 George, M., 256 George, N.P., 42, 56f, 57–58 Georgescu, R.E., 15, 18, 34t, 44–51, 55, 64–65, 113–114, 121–122, 193–194, 200, 203–205, 208, 210–218, 214f, 238 Gerding, M.A., 9 Gerhold, J.M., 279–280 Gerik, K.J., 207 Gerrick, E.R., 49 Ghirlando, R., 6f, 9 Ghodgaonkar, M.M., 246 Ghosh, S., 95, 100–101, 105, 110, 112–114 Ghosh, S.K., 294 Giannattasio, M., 273–276 Giardini, M.A., 193 Gibbs, P.E., 240–241 Gibbs-Seymour, I., 48 Giesler, T.L., 149 Gildenberg, M.S., 232–248 Gille, H., 3–4, 8–9 Gillis, A.J., 299 Gissi, C., 256 Glassberg, J., 41–42 Glover, B.P., 34t, 42, 54–57 Goddard, J.M., 274f, 275 Godson, G.N., 34t Goedken, E.R., 34t, 48, 57–59, 209–210 Goellner, E.M., 245–246 Gogol, E.P., 37 Gohara, D.W., 298–300, 306, 313–317 Gohlke, H., 311, 312f Gold, L., 109–110 Goldar, A., 193 Gomes, X.V., 203, 238 Gomez, R., 13 Gomez-Llorente, Y., 186 Gong, P., 303–304

333

Author Index

Gonzalez, D., 10 Gonza´lez-Huici, V., 150–153 Goodey, N.M., 299 Goodman, H.M., 277–279 Goodman, J.L., 141–148 Goodman, M.F., 51, 64–65, 113–114, 218–219, 294 Gorbalenya, A.E., 58–59, 100–101, 262f Gorbatyuk, B., 13–14 Gordenin, D.A., 203, 205–206 Goret, G., 19–20 Gorman, M., 205 Gotte, M., 298–301, 302t Gotz, D., 175–176 Gourinath, S., 37 Gourse, R.L., 13 Grabowski, B., 197–198 Graci, J.D., 307–309, 308t Grad, L.I., 273–275 Graham, B., 41–42 Grahl, A., 13–14 Grainge, I., 179–181 Grainger, D.C., 8–9 Graumann, P.L., 13–14 Graves, S.W., 100 Gray, M.W., 90, 102, 263–265 Green, C.M., 247–248 Green, L.S., 49–51, 109–110 Greenhaw, G.A., 95 Greenleaf, W.B., 197–198 Greipel, J., 42 Griep, M.A., 38–40, 62–63 Griffey, R., 68 Griffin, K., 105–107 Griffith, J., 4, 61 Griffith, J.D., 41–42, 44–45, 55, 61, 105–110, 119–122, 124 Grimes, J., 205 Grimwade, J.E., 2–4, 8–9, 11–13 Grindley, N.D., 67 Grippo, P., 96 Grochulski, P., 303–304 Groger, P., 8 Grosse, F., 201 Grossman, A.D., 10, 13–14 Gruez, A., 311 Guainazzi, A., 197–198, 200, 212–215 Guarino, E., 205

Guenther, B., 58–59, 208–209 Guggenheimer, R.A., 41–42, 109–110 Gui, W.-J., 48 Guidotti, S., 294 Guilbaud, G., 193 Guiles, J.W., 49–51 Guilliam, T.A., 38–39 Gulbis, J.M., 57, 208, 209f, 235 Guo, C., 239–240 Guo, S., 95, 100–107 Gupta, M.K., 34t, 68 Gupta, R., 52 Gustafsson, C.M., 94, 100, 102 Gutierrez, C., 157–158 Gutierrez, J., 152–153 Gutsche, I., 11–13, 19–20 Guy, C.P., 34t, 68 Guy, L., 170 Gwynn, E.J., 34t Gygi, S.P., 219 Gygli, G., 48

H Ha, T., 41–43, 63, 123–124, 198 Haber, J.E., 243–244 Hacker, K.J., 197–198, 216–217 Hall, D.R., 11–13 Hall, R.J., 202 Halvas, E.K., 295–296 Hamatake, R.K., 205–206 Hamdan, S.M., 34t, 43, 49, 50f, 51–53, 55, 61, 63, 65, 68, 95, 99, 102–103, 108–117, 119, 121–124, 216–218 Hamilton, M.D., 91 Hamlin, R., 66, 96–97, 112–113, 295–296, 299 Hammes, G.G., 299 Hammes-Schiffer, S., 299, 313 Han, E.S., 34t Han, J., 205–206 Handayani, R., 54–55 Hansen, F.G., 2–5, 10 Hansen, J.L., 295–296 Haq, I., 68 Haracska, L., 232, 240 Harada, Y., 13 Harding, A.E., 256 Harding, N.E., 4, 9, 159–160

334 Haring, M., 138 Harki, D.A., 303–304 Harrington, J., 203 Harris, L.D., 41–42 Harrison, S.C., 38–39, 295–296, 299 Harry, E.J., 46–49, 58 Hartmann, A., 8 Hartmann, K.H., 4 Harvey, S., 34t, 52–54 Hatano, M., 11–13 Haugen, E., 58 Haukka, J., 259–260, 265, 267–271 Hauschka, P.V., 201 Hauswirth, W.W., 278–279 Havens, C.G., 237 Hawkins, M., 175 Hayashi, I., 103 Hayashi, M.K., 194–196 Hayner, J.N., 45–46 He, J., 279–280 Headlam, M.J., 36 Hedglin, M., 193–194 Hefferon, K.L., 141, 142t Helleday, T., 243 Heller, R.C., 68 Heltzel, J.M.H., 52 Hemperly, S., 307–309, 308t Hemsworth, G.R., 67 Hench, J., 273–275 Hendel, A., 34t Hengge, A.C., 296–297 Henzler-Wildman, K., 299 Herbert, E.R., 276 Hermoso, J.A., 160–161 Hermoso, J.M., 150–153 Herrera, M.C., 199–200 Herschlag, D., 298 Hervas, C., 218 Hickman, D.D., 138–139 Higashi, M., 11–13, 16 Hill, F.R., 34t, 49, 52–53 Hill, T.M., 68 Himawan, J.S., 99–100 Hindges, R., 235 Hine, A.V., 117–119 Hingorani, M.M., 34t, 45–48, 54–57, 59–60, 103–107, 120–121 Hinkle, D.C., 44, 92, 95

Author Index

Hinrichs, S.H., 39–40 Hiraga, S., 2–3 Hirota, Y., 2–4, 53 Hirshberg, D., 54–55 Hirst, J., 201–202 Hishiki, A., 235 Hite, R.K., 63, 95, 99, 112, 115–117, 119, 123–124 Hitomi, C., 103 Hix, L., 308–309 Hixson, J.E., 280–281 Ho, T.V., 197–198, 200, 212–215 Hodgkinson, J., 103 Hodgman, T.C., 58–59 Hodgson, B., 219 Hodskinson, M.R.G., 34t, 67 Hoege, C., 232–233, 235–236, 239–242 Hogg, M., 207–208 Hogrefe, H.H., 68 Hogrefe, R.I., 68 Holden, L.G., 37 Holding, A.N., 45–48, 51–52 Holguera, I., 149–150, 159–160 Holley-Shanks, R., 103 Hollis, T., 95, 108–110 Holmes, C.C., 205 Holmes, J.B., 278–279, 281 Holmlund, T., 272 Holt, I.J., 256, 273–275, 278–279, 281–283 Holz, A., 3–4 Hombauer, H., 203 Hong, Z., 295–296 Hood, I.V., 14, 18–19, 37–38, 177–178, 192, 196 Horan, N.P., 34t, 44–49, 51–53 Hori, K., 99 Horie, H., 309 Hosoda, J., 109–110 Hosono, S., 149 Houlston, R.S., 205 Hounslow, A.M., 40 Howard, J.L., 68 Howarth, K.M., 205 Hozak, P., 247–248 Hsiao, C.D., 12f, 14 Hua, Y.-J., 49 Huang, C.Y., 12f, 14 Huang, H., 201–202, 295–296, 299

335

Author Index

Huang, L., 18 Huang, Y., 294 Huber, H.E., 91, 96, 99, 141 Huber, R., 202 H€ ubscher, U., 141, 193–194, 232, 235 Hughes, A.J.J., 45–48, 61, 122 Hughes, S.H., 295–296 Huisman, G., 13 Humphray, S., 205 Hunt, S., 193 Hupert-Kocurek, K., 18 Hura, G.L., 39–40, 62, 115–117 Hurwitz, J., 14–15, 34t, 194–198, 200–202, 208, 209f, 212–215 Huttener, M., 54 Huttlin, E.L., 219 Hwang, D.S., 4, 8–9, 11–13 Hyberts, S.G., 117–119 Hyland, E.M., 109–110 Hyrien, O., 193 Hyvarinen, A.K., 275–276, 283

I Iannelli, F., 256 Ibarra, B., 144–145, 147–148 Iborra, F.J., 281 Ichiye, T., 312–313 Iida, T., 215 Illana, B., 152–155 Ilsley, D., 201–202 Ilves, I., 194–196, 199–200, 212–215, 214f, 219 Ilyina, T., 140–141, 144–145 Ilyina, T.V., 100–101, 262f Imamoto, F., 8–9 Imura, N., 309 Inciarte, M.R., 157–160 Indiani, C., 48–49, 59, 64, 113–114, 194–196, 199–200, 210–215, 218 Innis, C.A., 265, 266f Iqbal, M., 3–4 Ira, G., 243–244 Ishida, T., 11–13 Ishigo-Oka, D., 10 Ishikawa, S., 13–14 Ishino, S., 201 Ishino, Y., 103, 201, 209–210 Ishmael, F.T., 104–105

Isoz, I., 205–206 Itaya, M., 67–68 Ito, J., 138–139, 159–160 Ito, T., 95, 102–103, 110, 115–119 Itoh, T., 203–205 Itsathitphaisarn, O., 14–15 Ivanov, I., 233 Ivanova, M.E., 203–205 Iyer, R.R., 245

J Jabafi, I., 311 Jacob, D.T., 52 Jacobo-Molina, A., 295–296 Jacobs, H.T., 102, 273–276, 274f, 281–283 Jacobs, M.A., 58 Jameson, K.H., 13–14 Jamin, M., 19–20 Janissen, R., 68 Janniere, L., 218–219 Janska, A., 196 Janssen, C.S., 102 Jarvis, T.C., 58 Jaworski, P., 10 Jemt, E., 263, 272, 277–281 Jennings, P.A., 34t, 45–48, 51 Jentsch, S., 233 Jeong, Y.J., 100–101, 105–107, 112–113, 197–198 Jergic, S., 32–69, 34t, 50f Jeruzalmi, D., 34t, 45–48, 54–55, 58–60, 93, 208–209, 295–296 Jerva, L.F., 297–298, 304–306 Jessop, L.L., 54–55 Jetha, N.N., 146 Jezewska, M.J., 14–15, 18, 36–38 Jha, J.K., 6f, 9 Jia, S., 205–206 Jiang, T., 48 Jiang, Y., 197–198 Jimenez, M., 160–161 Jin, G., 308 Jin, Y.H., 205–206 Jiricny, J., 245 Jo, T., 4, 34t Joers, P., 274f, 275–276 Johanson, K.O., 34t, 44–45 Johansson, E., 54, 202, 205–208

336 Johne, R., 149 Johnson, A., 55, 64, 215–216, 236 Johnson, A.A., 100 Johnson, D.E., 95, 99, 102–103, 105, 110, 112–113, 115, 124, 218 Johnson, D.S., 108, 197–198 Johnson, K.A., 48, 100, 107, 112, 197–198, 295–299, 304–306 Johnson, R.C., 8–9 Johnson, R.E., 205–208, 239, 245 Johnson, S.J., 295–296, 299 Johnson, S.K., 39–40, 62–63 Jokela, M., 202 Jonczyk, P., 32, 53 Jones, A.D., 15, 18–19 Jones, A.M., 205 Jones, C.E., 121–122 Jones, L.J., 160–161 Jones, R.C., 200–201, 203–205, 215, 218–219 Jongeneel, C.V., 43, 61 Jonsson, Z.O., 141, 235 Jose, D., 197–198 Joshua-Tor, L., 14–15, 200 Joyce, C.M., 67, 295–298 Jueterbock, W.R., 3–4 Julicher, F., 107 Jurka, J., 138

K Kadyrov, F.A., 245 Kagawa, W., 8, 103 Kaguni, J.M., 2–21, 36, 171–172, 174, 177, 280 Kaguni, L.S., 100, 102, 201–202, 256–284, 257f, 266f Kaidow, A., 4–5 Kaiser, J.T., 202 Kajander, O.A., 273–275 Kakuda, T.N., 259 Kalkkinen, N., 138 Kamada, K., 212 Kamimura, Y., 194–196 Kamtekar, S., 141–152, 155–156 Kanemaki, M., 200–201, 203–205, 218–219 Kang, D., 278–280 Kang, S., 185–186, 196–197 Kang, Y.H., 194–198

Author Index

Kannouche, P.L., 239–240 Kano, Y., 8–9 Kantake, N., 34t Kapitonov, V.V., 138 Kaplan, C.D., 299 Kaplan, D.L., 14, 105, 197–198 Kapp, U., 11–14, 19–20 Karam, J., 49 Karlseder, J., 138 Karplus, M., 311–313 Kasamatsu, H., 277, 280–282 Kashav, T., 37 Kashioka, T., 11–13 Kasho, K., 13 Kataoka, Y., 309 Katayama, T., 3–5, 7–8, 10–13, 16, 34t Kath, J.E., 52 Kati, W.M., 297–298, 304–306 Kato, H., 67–68 Kato, M., 95, 102–103, 110, 115–119 Katou, Y., 203–205, 215 Katz, F., 55, 64, 215–216 Katz, J.M., 308 Kaufmann, G., 238 Kaur, K., 205 Kawabata, T., 215 Kawai, Y., 160–161 Kawakami, H., 3–4, 8, 13 Kawata, M., 67 Kaziro, Y., 34t Kazmirski, S.L., 34t, 57–59, 209–210 Kearsey, S.E., 205 Keck, J.L., 34t, 38–42, 56f, 57–58, 193–194 Keen, B.A., 38–39 Keener, S., 294 Kelch, B.A., 34t, 57–60, 193–194, 205, 208–210 Kelly, R.M., 175–176 Kelly, T., 232 Kelly, T.J., 95, 205 Kelman, Z., 34t, 42, 45–46, 55–58, 64, 141, 194–198, 208, 209f Kelso, M.J., 46–48 Kemp, Z., 205 Keniry, M.A., 34t, 40, 50f, 52–55 Kennedy, W.D., 100, 259–260, 267–269 Kennelly, P.J., 186–187 Kent, H., 45–48, 51–52

Author Index

Keranen, S., 206 Kern, D., 299 Kerr, C., 92 Kerr, D.J., 205 Kesti, T., 206 Keyamura, K., 3–4, 11–13, 16 Khopde, S.M., 34t, 37–40 Kiefer, J.R., 295–296 Kilkenny, M.L., 202–205 Kilpelainen, S., 202 Kim, D., 60 Kim, D.E., 105–107, 112 Kim, D.R., 34t, 51, 53 Kim, S., 100, 218 Kim, S.-S., 34t, 45–48 Kim, Y., 66–67 Kim, Y.T., 99, 108–110 Kimura, H., 281 Kinebuchi, T., 41–42, 103 King, A.J., 153–155 King, T.C., 277 Kinney, J.B., 219 Kirby, T.W., 53–54 Kirk, B.W., 193–194 Kisker, C., 100 Kissling, G.E., 246 Kitani, T., 39–40 Kjaer, S., 203–205 Klassen, R., 205–206 Klein, M.G., 37, 197–198 Klett, R.P., 66 Kling, A., 48 Klinge, S., 201–202, 207–208 Klonowska, M.M., 37–38 Knippers, R., 8–9 Kobayashi, R., 208–209 Kobori, J.A., 15 Kochaniak, A.B., 234–235 Kogoma, T., 4–5, 175 Kohara, Y., 92, 94 Kohda, D., 201 Kohlstaedt, L.A., 295–296 Kois, A., 13–14 Kokoska, R.J., 99 Kolodner, R.D., 203, 245–248 Kolter, R., 13 Komori, K., 103, 201 Kong, D., 107–109

337 Kong, X.-P., 34t, 45–48, 141, 193–194, 205, 208, 232 Kongsuwan, K., 34t, 45–48, 51 Konigsberg, W.H., 49, 108–109, 298–301, 302t Konopa, G., 273–275 Kool, E.T., 114 Koonin, E.V., 38–39, 54–55, 58–59, 100–101, 193–196, 215–216, 262f, 265 Kopf, G., 207 Korboukh, V.K., 307–309, 308t, 313–317 Koren, I., 219 Korhonen, J.A., 102, 261, 264, 272 Korlach, J., 10 Korn, D., 238 Kornberg, A., 3–4, 7–9, 11–18, 34t, 36–42, 44–46, 54–55, 66–67, 91, 138, 193–194, 201, 216–217, 294 Kornberg, R.D., 299 Kornberg, S.R., 91 Kornberg, T., 53 Kornblum, C., 279–280 Korneeva, V.S., 298–301 Korolev, S., 41–42, 147–148 Kortus, M.G., 303–304 Kovac, M.B., 205 Kowalczykowski, S.C., 34t, 103, 109–110 Kowalski, D., 7 Kozdon, J.B., 10 Kozlov, A.G., 34t, 41–43, 42f, 108–109 Kralicek, A.V., 68 Krassa, K.B., 109–110 Krause, M., 6f Kraut, J., 299 Krejci, L., 239 Kreuzer, K.N., 43, 61 Krishna, T.S., 208, 232–233, 234f Krude, T., 201–202 Kubota, H., 215 Kubota, T., 11–13 Kuchta, R.D., 193–194, 201–202, 297–299, 304–306 Kuge, S., 309 Kujoth, G.C., 259–261 Kukkaro, P., 138 Kulczyk, A.W., 90–124 Kumar, R., 193–194 Kumar, S., 62–63, 123

338 Kunkel, T.A., 99, 203, 205–207, 245–246 Kunz, C., 245 Kuriyan, J., 34t, 38–40, 45–51, 54–55, 57–60, 141, 193–194, 205, 208–210, 209f Kurth, I., 48–49, 55, 64–65, 121–122, 215–217 Kurumizaka, H., 3–4, 8, 11, 103 Kusakabe, T., 62–63, 108–109, 117–120, 122–124 Kuzmic, P., 54–55 Kuznetsov, S.V., 41 Kuznetsov, V.B., 100 Kwon, Y.M., 13

L Labaw, L.W., 92 L’Abbe, D., 277 Labib, K., 200–201, 203–205, 215, 218–219 Labigne, A., 13–14 Lacks, S.A., 67 Lahiri, S.D., 296–297 Lahue, R., 245 Laine, P.S., 34t, 41–42 Lamers, M.H., 34t, 45–53, 50f Lang, U.F., 49 Langer, U., 3–4 Langston, L.D., 113–114, 192, 194, 196, 210–215, 218 Lao-Sirieix, S.H., 201–203 Larrea, A.A., 203 Larson, M.A., 39–40 Larsson, C., 102 Larsson, N.G., 94, 100, 102 Lasken, R.S., 15–18, 149 Latham, G.J., 60 Laurinavicius, S., 138 Lawrence, C.W., 240–241 La´zaro, J.M., 138–158, 142t Leake, M.C., 48–49, 55, 215–216 Learn, B.A., 18 LeBowitz, J.H., 14, 34t Lechner, R.L., 93–94, 96 LeClerc, J.E., 92 Lee, C.A., 301, 303–304, 307–309, 308t Lee, D.D., 95 Lee, D.S., 66–67 Lee, J., 61, 102–103, 108–109, 119–122, 124

Author Index

Lee, J.-B., 60, 63, 95, 99, 102–103, 110, 112–113, 115–117, 119, 123–124, 218 Lee, J.K., 194–196 Lee, M.Y., 207 Lee, S.-G., 34t, 49–51 Lee, S.H., 232, 238, 273–275 Lee, S.-J., 91–92, 95, 99–121, 123 Lee, Y.S., 100, 259–260, 267–269 Lehman, I.R., 34t, 66–67, 91, 201–202 Lehmann, A.R., 68, 239–240 Lehnherr, H., 53–54 Lei, J., 199–200 Leipe, D.D., 38–39, 193–194, 215–216 Leis, J., 138–139 Leman, A.R., 236 Lemire, B.D., 273–275 Lemonidis, K.M., 68 Lengronne, A., 219 Leonard, A.C., 2–4, 8–9, 11–13, 172 Lesburg, C.A., 295–296 Lesnik, E.A., 68 Leu, F.P., 34t, 59, 64–65 Levin, D.S., 232 Levin, M.K., 100–101, 112–113, 197–198 Levy, D.L., 206 Lewis, J.S., 32–69 Lewis, P.J., 46–49, 58 Lewis, P.W., 194–196 Lewis, S.C., 267–269, 273–275, 274f Lewis, W., 259 Li, D., 53–54 Li, F.Y., 102 Li, G.M., 245 Li, H., 199–200, 210–215, 214f, 218 Li, J.J., 203, 205 Li, N., 199–200 Li, W., 199–200 Li, X., 121–122, 203 Li, Y., 95, 102–105, 147–148 Li, Z., 103 Lia, G., 123–124, 215–216 Lian, K., 48 Liang, J., 198 Liao, J.C., 105–107 Liberti, S.E., 203 Lichtarge, O., 265, 266f Lieber, M.R., 203 Lieberman, K.R., 146, 148–149

339

Author Index

Liepinsh, E., 34t, 37–40 Lifsics, M.R., 34t, 53 Lill, R., 207 Lilley, P.E., 34t, 38–40, 46–48 Lima, C.D., 236, 239 Lin, S.-Q., 48 Lin, T., 49 Lindahl, L., 13 Lindahl, T., 66–67 Ling, F., 273–275 Lingardo, L.K., 311 Lingner, J., 193 Link, A.J., 13 Linssen, A.B., 312–313 Lionnet, T., 108 Lippincott, J.A., 57 Liu, B., 14 Liu, J., 294 Liu, J.Y., 177–178 Liu, L., 201, 207 Liu, M., 48 Liu, P., 282–283 Liu, Q., 92 Liu, X., 303–304 Liu, Y., 311 Livingston, D.M., 44 Livneh, Z., 34t Llorca, O., 14, 202, 207–208 Llorente, B., 243–244 Lloyd, R.G., 68 Lo, A.T.Y., 42–43 Lo, Y.H., 12f, 14 Lobner-Olesen, A., 4–5 Łobocka, M.B., 53–54 Lockhart, D.J., 193 Loeb, L.A., 49–51, 294 Loebner-Olesen, A., 8–9 Lohman, T.M., 34t, 41–43, 42f, 107–109 Lonberg, N., 109–110 London, R.E., 52–54 Lone, S., 34t Long, A.M., 95–97, 99, 117, 147–148, 295–296 Long, D.T., 197–198, 200, 212–215 Longa´s, E., 149–156 Longley, M.J., 259–261 Longo, M.A., 202

Loparo, J.J., 32–33, 43–45, 55, 61, 65, 95, 97–99, 110, 113–115, 119, 121–123, 216–217 Lo´pez de Saro, F.J., 34t, 45–48, 51, 64–65, 67, 121–122 Lopez, P., 67 Lo´pez-Bueno, A., 149 LoPresti, M.B., 41–42, 109–110 Lorimer, H.E., 273–275, 281–283 Loscha, K.V., 34t, 37–40, 63 Losick, R., 13–14 Louarn, J., 2–3 Lourens, R.M., 308 Love, R.A., 311 Loveland, A.B., 197–198 Lovett, S.T., 34t, 243–244 Low, R.L., 277 Lu, C., 34t, 51–52 Lu, D., 34t, 42 Lucassen, A., 205 Ludlam, A.V., 15–18 Luft, B.J., 149 Lujan, S.A., 205–206 Lukat, P., 48 Lum, D., 301, 303–304, 313–315 Lundgren, M., 170–171, 175–176 Lundquist, R.C., 66–67 Lundstrom, E.B., 205–208 Lunyak, V.V., 282 Luo, J., 313 Luo, R., 311, 312f Luong, K., 10 Lupski, J.R., 34t Lurz, R., 6f, 10, 13–14 Lutz, S., 311 Lydeard, J.R., 243–245 Lynch, A.S., 34t, 38–40, 193–194 Lyngstadaas, A., 8–9 Lyubimov, A.Y., 14–15, 37–38, 199–200

M Ma, C., 48 Ma, Y., 205 Macdonald, P., 158–159 Mackiewicz, P., 13–14 MacLellan-Gibson, K., 212–215, 214f Macneill, S.A., 202 Maculins, T., 219

340 Madiraju, M.V.V.M., 34t Madsen, P., 232 Maegley, K.A., 311 Maga, G., 193–194, 235 Magazin, M., 93 Mahdi, A.A., 34t Mahlow, E., 4 Mahoungou, C., 265 Mahy, B.W.J., 138 Mai, A.H., 146 Maier, D., 264 Mailand, N., 48 Majka, J., 117 Makarova, A.V., 207 Makarova, K.S., 193–196 Makhov, A.M., 61, 121–122 Maki, H., 34t, 44–45, 54, 215 Maki, S., 215 Makiniemi, M., 202 Makino, D.L., 59–60, 193–194, 205, 208–210 Makowska-Grzyska, M., 16, 18–19 Maksimchuk, K.R., 298–301, 302t, 303–304, 313–315 Malc, E.P., 205–206 Maleszka, R., 267–269, 273–275 Malkova, A., 243–244 Maman, J.D., 201–202 Manhart, C.M., 42, 64–65 Mannarino, A.F., 295–296 Manosas, M., 63, 218 Mao, C., 295–296 Maranas, J.K., 313–317 Marceau, A.H., 42, 56f, 57–58 Marcotte, L.L., 313–317 Marczynski, G.T., 9–10 Margulies, C., 3–4, 10 Marians, K.J., 20, 34t, 38–39, 42, 56f, 57–58, 61, 65, 68, 121–122, 216–218 Marintchev, A., 109–110 Marintcheva, B., 99, 102–103, 108–112 Marinus, M.G., 34t Mark, D.F., 96, 99 Marsh, R.C., 2–3 Marszalek, J., 16 Martı´n, A.C., 153–155 Martı´n, G., 138, 140–141, 156–158 Martin, L., 205

Author Index

Martinez, P., 193 Martin-Parras, L., 282 Martuscelli, J., 2–3 Marzahn, M.R., 45–46 Marziali, A., 146 Masai, H., 212 Masamune, Y., 91 Mashimo, K., 67 Masker, W., 108–109 Mason, C.E., 34t, 41–48, 51–52 Masson, J.Y., 103 Massoni, S.C., 42, 56f, 57–58 Masters, M., 2–3 Matak-Vinkovic, D., 203–205 Mathews, M.B., 232 Matson, S.W., 93–94, 100–101, 103–105 Matsuda, Y., 4, 34t Matsui, M., 3–4 Matsumoto, L., 276 Matsumoto, T., 41–42 Matsunaga, C., 4, 13, 34t Matsunaga, F., 175, 179–181, 201 Matsushima, Y., 261–263 Maupin-Furlow, J.A., 186–187 Maurer, R., 34t, 53 Mayanagi, K., 103, 209–210 Mayhook, A.G., 276 Mazin, A.V., 103 McAdams, H.H., 10 McCallum, S.A., 175–176, 181 McCammon, J.A., 311 McCauley, M.J., 49 McEachern, M.J., 243–244 McGarry, K.C., 3–4 McGlynn, P., 34t, 68 McGovern, S., 13–14 McHenry, C.S., 20, 34t, 42, 44–49, 51–58, 61, 64–66, 121–122, 215–219 McInerney, P., 15, 18, 38–40, 55, 64, 113–114, 193–194, 215–216 McIntyre, J., 219 McKay, D.B., 261 McKinney, E.A., 256 McLaughlin, L., 95 McLenigan, M.P., 219 McMacken, R., 14, 18, 34t McNally, R., 34t, 57–59, 209–210, 233 McNatt, M.W., 15–18

341

Author Index

McNeill, S.M., 34t McSwiggen, J., 294 McVean, G., 205 Mechali, M., 172, 193 Mehra, P., 19–20 Mehta, A., 243–244 Meijer, M., 2–3 Meijer, W.J., 152, 160–161 Melero, R., 202 Melfi, M.D., 10 Mellado, R.P., 138–139, 157–158 Mencı´a, M., 149, 158–160 Mendez, E., 152 Mendez, J., 138–139, 141–144, 153–155 Meng, Q., 100 Mer, G., 184–185 Merrikh, H., 13–14 Merz, K.M., 311, 312f Meselson, M., 43 Messer, W., 2–4, 6f, 8–11, 16, 36 Meyer, R.R., 34t, 41–42 Michel, B., 123–124, 215–216 Mieczkowski, P.A., 205–206 Mielke, T., 10 Miki, T., 2–3 Mildvan, A.S., 294, 299 Milenkovic, D., 280 Miles, C.S., 34t, 37–38 Miles, J., 203–205 Millar, D., 59–60 Miller, D., 92 Miller, J., 138–139 Miller, J.M., 182, 183f Minczuk, M., 277–279 Miralles Fuste, J., 272, 280–282 Mitkova, A.V., 34t, 37–39 Mitsunobu, H., 92 Mittal, A., 42 Miyabe, I., 205–206 Miyachi, K., 232 Miyagawa, K., 103 Miyata, T., 103, 209–210 Mizrahi, V., 297–299, 304–306 Mo, J., 207 Moarefi, I., 59–60 Model, P., 96 Modrich, P., 34t, 96, 245 Moe, E., 48

Moffatt, B.A., 93 Mohideen, F., 236, 239 Moldovan, G.L., 233 Molineux, I.J., 95 Monia, B.P., 68 Montello, M.L., 34t Montoya, J., 278–279 Mooney, P.Q., 95 Morais, R., 277, 280–281 Morandi, L., 102 Morgan-Hughes, J.A., 256 Morikawa, K., 209–210 Morimoto, T., 13–14 Morimoto, Y., 41–42 Morin, J.A., 144–145 Moriya, S., 6f, 10 Morris, C.F., 43, 61, 119, 273–275 Morrison, A., 205–206, 232 Morton, B., 311–313 Morvillo, M.V., 277 Moses, R.E., 45–48, 206 Mosig, G., 158–159 Mott, M.L., 7, 15, 18–19, 178 Moustafa, I.M., 294–317, 302t Moya, A., 149 Moyer, S.E., 194–196 Mueller, G.A., 53–54 Mukherjee, S., 105–107, 120–121 Mukhopadhyay, A., 102 Mukhopadhyay, G., 19–20 Mulcahy, H.L., 102–103 Mulcair, M.D., 68 M€ uller, H., 149 Mun˜oz-Espı´n, D., 149–150, 159–161 Muramatsu, S., 194–196 Murray, H., 13–14 Murzin, A.G., 49 Myers, J.A., 91 Myllykallio, H., 170–171, 175, 193–194, 201, 215–216

N Nagata, T., 2–3 Nagata, Y., 67 Nair, P.A., 34t Nakai, H., 108–110 Nakamura, K., 8 Nakamura, T., 49

342 Nakano, Y., 309 Nakato, R., 215 Nakayama, N., 34t Naktinis, V., 51, 55–59 Nandakumar, J., 34t Nanni, R.G., 295–296 Narayan, M., 112 Nass, M.M., 256, 277, 282–283 Nass, S., 256 Natrajan, G., 11–14 Naue, N., 34t, 42, 58 Navarre, W.W., 92 Neilson, L., 68 Nelson, J.R., 149 Nelson, S.W., 121–122, 124, 216–217 Nethanel, T., 238 Netz, D.J., 207 Neuwald, A.F., 54–55 Nevin, P., 34t Newport, J.W., 109–110 Neylon, C., 68 Neyts, J., 311 Ng, K.K., 303–304 Nguyen, B., 42 Nicholls, T.J., 277–279 Nick McElhinny, S.A., 205–206, 237–238, 246 Nielsen, B.L., 273–275 Nielsin, L.D., 2–3 Nievera, C.J., 8–9 Niiranen, L., 48 Nikali, K., 102 Nilges, M.J., 201–202 Nishikawa, H., 219 Nitharwal, R., 37 Niwa, O., 206 Nix, J.C., 303–304 Nkosi, P.J., 219 Nogales, E., 199–200, 202 Noguchi, E., 236 Noirot, P., 13–14 Noirot-Gros, M.F., 11–14 Nolan, J.P., 67 Nomoto, A., 309 Nomura, N., 273–275 Norais, C., 175, 179–181, 201 Norcum, M.T., 104–105 North, R., 95

Author Index

Northam, M.R., 240–241 Nossal, N.G., 61, 108–109, 121–122 Notarnicola, S.M., 102–103, 105–107 Nowicka, A., 53 Nunes, V.S., 193 Nunez-Ramirez, R., 202, 207–208 Nureki, O., 3–4, 11 Nurminen, A., 259–260

O Oakley, A.J., 34t, 37–42, 44–48, 68 Ochsner, U., 58 O’Day, K., 36 Ode, K., 215 O’Donnell, M.E., 15–16, 18, 20, 34t, 38–40, 42, 44–51, 53–60, 64–65, 67–68, 113–114, 121–122, 141, 192–220, 209f, 214f, 236 Ogasawara, N., 6f, 10, 13–14, 67–68 Ogata, Y., 11–13 Ogawa, T., 39–40 Ogura, T., 185–186 Ohi, M., 102–105 Ohmori, H., 247–248 Oka, A., 3–4 Okawa, M., 194–196 Okazaki, R., 32, 34t, 67 Okazaki, T., 32, 34t, 38–40, 67, 92, 94 Okorokov, A.L., 103 Okumura, K., 44 Olieric, V., 46–48 Oliveira, M.T., 102, 256–284, 257f, 266f Olivera, B.M., 66–67 Ollis, D.L., 34t, 51–52, 66, 96–97, 112–113, 295–296, 299 Olson, M.W., 55–57, 260–261, 267–269 Omata, T., 309 Onesti, S., 182, 186 Onrust, R., 34t, 45–46, 55–59, 141, 193–194, 205, 208–209, 218 Onufriev, A., 311, 312f Ordog, T., 205–206 Orebaugh, C.D., 205–206 Orlova, E.V., 103 Ortmann, A.C., 175–176 Osbourn, A., 149 O’Shea, V.L., 14, 18–19, 37–38 Oster, G., 105–107

Author Index

Osterman, P., 207–208 Otting, G., 34t, 37–42, 50f, 51–55, 57 Otto, B., 34t Ouimet, M.C., 9–10 Overman, L.B., 41–42 Owen, E.A., 34t Oyama, T., 209–210 Ozaki, S., 7–8, 13 Ozawa, K., 34t, 41–42, 44–49, 50f, 51–53, 55, 57

P Palese, P., 308 Palles, C., 205 Palm, L., 92 Pan, H., 20, 40 Pandey, M., 63, 68, 123–124 Papouli, E., 232–233, 236, 239 Pappas, C., 308 Pappin, D.J., 66–67 Pares, E., 141–144 Parge, H.E., 311 Park, A.Y., 34t, 41–42, 50f, 51–55 Park, K., 61, 105–107, 119 Park, M.S., 67 Park, S.Y., 8 Parker, J.L., 239 Parra, F., 303–304 Parsons, M.R., 108–109 Pascal, J.M., 238–239 Paschall, C.O., 45–46 Pasero, P., 68, 219 Passy, S.I., 103 Patel, D., 68 Patel, G., 63, 100, 123–124 Patel, P.H., 49–51 Patel, S.S., 63, 68, 100–101, 103–108, 112–113, 120–121, 123–124, 197–198, 297–299, 304–306 Pathak, V.K., 295–296 Patoli, A.A., 46–48 Paul, B.J., 13 Paul, L.S., 109–110 Pausch, M.H., 202 Paytubi, S., 54 Pearl, L.H., 46–48 Pease, P.J., 39–40, 62, 115–117 Pedro, L., 54

343 Peersen, O.B., 303–304, 311 Pellegrini, L., 103, 201–205, 207–208 Pellegrini, M., 102 Pelletier, H., 299 Pelton, J.G., 34t Pelve, E.A., 170–171, 175 Pen˜alva, M.A., 152, 157–160 Peng, X., 138 Perera, R.L., 202–205 Perez-Arnaiz, P., 140–141, 142t, 147–153, 155–156 Perez-Luque, R., 311 Perna, N.T., 32 Perrino, F.W., 34t, 52–54 Perrone, L.A., 308 Persky, N.S., 34t Pesavento, J.J., 194–196, 198, 212–215, 214f Pesole, G., 256, 277 Petermann, E., 243 Peterson, B.R., 303–304 Petojevic, T., 194–196, 198–200, 212–215, 214f Petridis, C., 205 Petsko, G.A., 299 Pfander, B., 232–233, 236, 239 Pfeiffer, V., 193 Pham, T.M., 44 Pham, X.H., 102, 278–279 Phillips, R.S., 102 Piccirilli, J.A., 298 Picha, K.M., 104–105 Pierik, A.J., 207 Pietroni, P., 60 Pinet, A., 58 Pintacuda, G., 34t, 37–40, 50f, 52–54 Pinz, K.G., 280 Piotr, J., 13–14 Pirruccello, M.M., 7, 11, 12f Pisani, F.M., 212 Pluciennik, A., 246 Plunkett, G., 32, 53–54 Plyasunov, S., 147–148 Podobnik, M., 38–40, 58–59, 193–194 Podust, V.N., 232 Pohjoismaki, J.L., 282–283 Polaczek, P., 8–9 Polard, P., 13–14 Poli, J., 68

344 Politis, A., 54–55 Polychronopoulos, D., 203–205 Pomerantz, R.T., 68 Pons, M., 54 Ponting, C.P., 194–196 Porter, K., 138 Pospiech, H., 202 Posse, V., 278–279 Possoz, C., 55 Poulton, J., 102 Prakash, L., 205–208, 239 Prakash, S., 205–208, 239 Prangishvili, D., 138 Prasad, B.V., 108–109 Prasad, R., 299 Prelich, G., 232 Prieto, I., 152–153 Pritchard, A.E., 34t, 49, 54–57 Prosselkov, P., 34t, 45–48 Pryor, J.M., 239 Purdy, M.M., 313 Pursell, Z.F., 205–207, 237–238 Putnam, C.D., 247–248 Putnam, F.W., 92 Pyle, A.M., 197–198

Q Qi, X., 149 Qimron, U., 95, 99, 102–107, 110, 112–113, 115–117, 123–124, 218 Quesada, A., 149

R Rabkin, S.D., 93–94 Radding, C.M., 103 Raghunathan, S., 34t, 41–42, 42f, 108–109, 266f Raghuraman, M.K., 193 Rahn-Lee, L., 13–14 Rajagopal, V., 197–198 Rajagopalan, P.T., 311, 313 Rajala, N., 271–272, 280 Rajendran, S., 14–15, 18, 37 Randi, E., 277 Randolph, K.M., 100 Rankine, J., 276 Rappailles, A., 193 Rast, J.P., 276

Author Index

Ravantti, J.J., 138 Ray, D.S., 273–275 Rea, S.L., 273–275 Rector, A., 149 Recuero-Checa, M.A., 202 Redrejo-Rodrı´guez, M., 149–150, 159–160 Reems, J.A., 65, 121–122 Regenmortel, M.H.V.v., 138 Register, J., 41–42 Reha-Krantz, L.J., 14, 34t Reich, E., 66 Reich, N.O., 313 Reid, A.H., 308 Reilly, B.E., 138–139 Rein, D.C., 34t Reinbolt, J., 46–48 Renault, L., 203–205, 212–215, 214f Resnick, M.A., 205–206 Reyes, A., 281–283 Reyes-Lamothe, R., 36, 48–49, 55, 192, 215–216 Rezende, L.F., 108–110 Ricard, C.S., 41–42 Ricchetti, M., 66 Rice, P.A., 8–9, 11–13, 295–296 Richardson, C.C., 39–40, 43–44, 61–63, 65, 90–124, 141, 147–148, 193–194, 201–202, 216–218, 294 Richter, S., 3–4 Riehle, M.O., 102 Riera, A., 192, 196, 199–200 Riise, E., 2–3 Riley, M., 32 Rinaldi, A.M., 276 Ringe, D., 299 Robberson, D.L., 277, 280–282 Roberti, M., 277 Roberts, M., 311 Robins, P., 66–67 Robinson, A., 34t, 36, 46–49, 52–53, 57–58 Robinson, N.P., 170–171, 175–176, 178–181 Roccasecca, R.M., 113–114 Roche, D.D., 34t, 38–40, 193–194 Rock, J.M., 49 Rodrı´guez, I., 141–144, 147–150 Rodriguez-Belmonte, E.M., 207 Roe, S.M., 46–48

Author Index

Rogers, R.S., 193 Rokop, M.E., 10 Romano, L.J., 94, 102 Ronnholm, G., 138 Rose, D.J., 53–54 Rose, J.P., 295–296 Rosenberg, A.H., 105–107 Rosenberg, S.M., 44, 55 Ross, W., 13 Rossignol, J.M., 201–202 Rostami, N., 13–14 Roth, A., 3–4, 8–9 Rothenberg, E., 198 Rout, M.P., 215 Rouviere-Yaniv, J., 11–13 Rouvinski, A., 160 Roux, B., 311 Rowen, L., 34t Roy, R., 41–43 Roylance, R., 205 Rozgaja, T.A., 3–4 Rubenstein, J.L., 275–276 Ruckert, B., 6f, 10 Rudner, D.Z., 197–198 Rudolph, C.J., 34t Ruhanen, H., 264, 280 Runswick, M.J., 58–59 Ruotolo, B.T., 54–55 Rupp, W.D., 34t, 41–42 Rusin, M., 53–54 Ruslie, C., 34t Russel, M., 96 Ryan, V.T., 3–4, 8–9, 11–13

S Sabouri, N., 205–206 Saccone, C., 277 Sadowski, P.D., 92, 95 Saenger, W., 37 Sage, J.M., 18 Saito, H., 91, 93–94 Saito, Y., 245 Saitoh, K., 203–205 Sakabe, K., 32 Sakakibara, Y., 34t Sakumura, Y., 44 Sakurai, S., 234f, 235–236 Salas, M., 54, 138–161, 142t

345 Sale, J.E., 239 Salguero, I., 205 Sali, A., 58–59, 208–209 Salinas, F., 109–110, 121–122, 219 Samel, S.A., 199–200 Samojedny, A., 53–54 Sampoli Benitez, B.A., 311 Samson, R.Y., 170–187, 173f San Martin, M.C., 37–38 Sancar, A., 34t, 41–42 Sanchez-Diaz, A., 200–201, 203–205, 218–219 Sanchez-Farran, M.A., 313–317 Sanchez-Pulido, L., 194–196 Sandler, S.J., 42, 56f, 57–58 Saneto, R.P., 257–259 Santoro, L., 102 Santos, E., 140–141, 142t Saraste, M., 58–59 Satapathy, A.K., 95, 100–101, 105, 112–113 Sato, S., 102 Sauer, B., 201–202 Sauer-Eriksson, A.E., 207–208 Saugar, I., 243 Sauguet, L., 202 Savvides, S.N., 41–42, 108–109 Sawaya, M.R., 95, 102–107, 120–121, 299 Sawyer, E.J., 205 Sayers, J.R., 34t, 67–68 Sayood, K., 39–40 Sbisa, E., 277 Schaaper, R.M., 32, 34t, 49, 52–54, 91 Schaechter, M., 158–159 Schaefer, C., 3–4 Schaefer, K.T., 100 Schaeffer, P.M., 34t, 36–40, 63, 68 Schaller, H., 2–3, 34t Schaper, S., 3–4 Schar, P., 245 Scharer, O.D., 197–198, 200, 212–215 Schauer, G.D., 194, 210–212, 218 Schekman, R., 45–46 Scheres, S.H., 49, 50f, 52–53 Scheuermann, R., 34t, 51–52 Schlick, T., 311 Schlierf, M., 8 Schmidt, A., 10 Schmieder, P., 34t, 42

346 Scholefield, G., 13–14 Schowen, R.L., 300–301 Schubach, W.H., 16–18 Schuller, A.P., 299 Schultz, S.C., 295–296 Schwacha, A., 174, 194–196, 199–200 Schwarz, U., 247–248 Scott, D.D., 34t, 49, 50f, 55 Sedelnikova, S.E., 67 Sefcikova, J., 49 Segatto, M., 193 Seitz, E.M., 103 Seitz, H., 8, 16 Sekedat, M.D., 193 Sekimizu, K., 3–4, 7, 11–13 Selisko, B., 311 Sengerova´, B., 67 Seow, F., 102 Serna-Rico, A., 152, 160–161 Serrano, M., 138–139, 153 Serwer, P., 95 Seto, D., 91 Sexton, D.J., 121–122 Seyedin, S.N., 209–210 Shadel, G.S., 278–279 Shamoo, Y., 54, 108–109, 147–148, 235 Shapiro, L., 10 Sharma, V., 313 She, Q., 175–176 Sheaff, R.J., 201–202 Shechter, D., 45–46 Sheil, M.M., 38, 52 Shell, S.S., 247–248 Shen, B., 67 Shen, H., 311–313 Sherborne, A., 205 Shereda, R.D., 34t, 41, 264, 271–272 Sherratt, D.J., 48–49, 55, 192, 215–216 Sheu, Y.J., 219 Shi, J., 265, 266f Shi, Y., 94, 200, 210–215, 214f, 218 Shibahara, K., 232, 247 Shibata, N., 41–42 Shibata, T., 103, 273–275 Shimamoto, A., 282–283 Shimamoto, N., 41–42 Shimuta, T.R., 13 Shin, D.S., 103

Author Index

Shin, J.H., 197–198 Shinagawa, H., 103 Shine, J., 277–279 Shirahige, K., 203–205, 215 Shirouzu, M., 3–4, 11 Shishmarev, D., 41–42 Shivji, M.K., 103 Shokri, L., 49 Showalter, A.K., 298 Shuman, S., 34t Shumate, C., 100 Shutt, T.E., 90, 102, 263–265 Siam, R., 10 Siegel, P.J., 158–159 Sigal, L.J., 308–309 Silva, M.C., 34t Simmerling, C., 311, 312f Simmons, L.A., 4, 11, 16 Simms, E.S., 66, 91 Simon, A.C., 203–205 Simon, M.N., 194–196 Simonetta, K.R., 34t, 57–59, 209–210 Singh, H., 202 Singleton, M.R., 14–15, 95, 102–107, 120–121, 177–178, 193–194, 196–197 Sinha, N.K., 43, 61, 119, 273–275 Skangalis, M., 34t, 55 Skarstad, K., 4–5, 8–10, 16–18 Skelly, P.J., 267–269, 273–275 Skibbens, R.V., 232 Sklar, L.A., 67 Skordalakes, E., 299 Skovgaard, O., 13–14 Skrobuk, P., 13–14 Slater, S.C., 34t, 53 Smidansky, E.D., 298–301, 302t, 303–304, 308–309, 313–317 Smiley, B.L., 34t Smith, B.Y., 108, 197–198 Smith, C.E., 203, 243–244 Smith, D.J., 205–206 Smith, D.W., 4, 9 Smith, S.B., 147–148 Snyder, A.K., 54–55 Sobhy, M.A., 44, 68 Soengas, M.S., 138–140, 144–145, 157–158 Sogo, J.M., 138–139, 157–160 Sohl, C.D., 259–260, 269

347

Author Index

Solorzano, A., 308 Somer, H., 102 Somerharju, P., 138 Soni, R.K., 19–20 Sordello, S., 48 Soufo, C.D., 13–14 Soufo, H.J., 13–14 Soultanas, P., 20, 39–40 Sousa, R., 294–296, 311 Spain, S.L., 205 Speck, C., 8, 192, 196, 199–200 Spelbrink, J.N., 102 Spencer, D., 58 Spicer, E.K., 41–42, 109–110 Spiering, M.M., 63, 104–105, 108, 218 Spies, M., 245 Spouge, J.L., 54–55 Stahl, F.W., 43 Stahl, P.D., 193–194 Stamford, N.P.J., 37–40 Stano, N.M., 100–101, 112–113 Stattel, J.M., 95, 108–110 Steen, H.B., 4–5 Steindorf, A., 13–14 Steitz, T.A., 12f, 14–15, 19–20, 34t, 37–39, 39f, 49–54, 58–59, 66–67, 93, 96–97, 103–105, 108–109, 112–113, 141–152, 155–156, 193–194, 207–208, 295–297, 299 Stelter, M., 19–20 Stelter, P., 232–233, 236, 239 Stengel, G., 201–202 Stepankiw, N., 4–5 Stevens, H., 149 Stewart, J., 34t, 45–48, 59–60, 64–65, 121–122 Stiban, J., 261–264 Stillman, B., 192, 194–196, 202–203, 205–206, 208–209, 219, 232, 238, 247 Stingl, K., 13–14 Stith, C.M., 205–207 Stodola, J.L., 207 Stone, J.K., 304–309, 308t Story, R.M., 58–59, 103 Storz, G., 13 Strycharska, M., 14–15 Strycharska, M.S., 37–38

Studier, F.W., 90, 92–93, 95, 101–102, 105–107 Studwell, P.S., 57 Studwell-Vaughan, P.S., 34t, 44–49, 53–55, 193–194, 205, 208 Stukenberg, P.T., 44–48, 64–65, 121–122, 193–194, 205, 208, 216–218 Stumpf, J.D., 257–259 Stumpfig, M., 207 Su, X.-C., 34t, 38–42, 49, 50f, 55 Suck, D., 68 Su’etsugu, M., 13 Sugimoto, K., 32, 34t, 67, 94 Sugimoto, T., 264 Sugino, A., 32, 194–196, 205–206 Sugiyama, T., 34t Suh, S.W., 66–67 Sun, J., 199–200, 210–215, 214f, 218 Sun, Y.J., 12f, 14 Sun, Z., 149 Suomalainen, A., 102 Suski, C., 34t, 218–219 Sutera, V.A., 34t Suttle, C.A., 138 Sutton, M.D., 11, 16, 34t, 52 Suzuki, H., 209–210 Svarovskaia, E.S., 295–296 Svec, P.S., 34t Swan, M.K., 207–208 Swanson, L., 67 Swart, J.R., 38–40, 62–63 Swayne, D.E., 308 Swinger, K.K., 8–9, 11–13 Swuec, P., 212–215, 214f Syed, S., 63, 123–124 Symington, L.S., 243–244 Syson, K., 40, 67 Syvaoja, J.E., 202, 206 Szafranski, S., 13–14 Szczesny, R.J., 279–280 Szymanski, M.R., 100, 257–258f, 259–260, 269

T Tabor, S., 61, 91–115, 117–119, 124, 141, 147–148, 218, 294 Tachdjian, S., 175–176 Tackett, A.J., 193, 215

348 Taft-Benz, S.A., 34t, 49, 52–53 Tainer, J.A., 100, 103 Tajima, R., 215 Takahashi, M., 43–44, 61, 65, 68, 95, 110, 112–113, 119, 121–123, 216–217 Takamatsu, C., 280 Takanami, M., 3–4 Takayama, Y., 194–196 Tam, S., 34t, 51–52 Tamames, J., 149 Tamanoi, F., 93–94 Tamberg, N., 199–200, 219 Tamura, K., 266f Tan, C.K., 232 Tan, E.M., 232 Tan, K.W., 44 Tan, M., 92, 100 Tan, X., 201–202 Tanaka, H., 203–205 Tanaka, K., 103 Tanaka, S., 192, 196 Tanner, N.A., 44–45, 55, 63, 95, 99, 102–103, 110, 112–113, 115, 124, 218 Tappin, I., 194–198 Tariq, M., 102 Taubenberger, J.K., 308 Taylor, J., 205 Taylor, J.A., 9–10 Taylor, J.S., 295–296, 299 Terada, T., 3–4, 11 Terradot, L., 11–14, 19–20 Thaipisuttikul, I., 58 Theile, C., 95 Theis, K., 100 Thermes, C., 193 Thirlway, J., 40 Thomas, C.A., 95 Thomas, H.J., 205 Thommes, P., 264, 269–271 Thompson, A.A., 303–304, 311 Thompson, A.C., 11–13 Thompson, J.A., 45–46 Thompson, P.R., 34t, 51–53, 57 Thomsen, N.D., 14–15 Thomson, D.P., 34t Thresher, R., 4 Ticau, S., 174 Tiengwe, C., 172

Author Index

Timinskas, A., 49 Timinskas, K., 49 Timmins, J., 19–20 Tiranti, V., 102 Tisma, J.K., 138–139 Tjajadi, M., 197–198 Tognetti, S., 192, 196, 199–200 Tokonzaba, E., 37 Tolun, G., 44–45, 55 Tomlinson, C.G., 67 Tomlinson, I., 205 Torella, R., 202 Torres-Ramos, C.A., 242–243 Toscano, A., 102 Toste R^ego, A., 45–48, 51–52 Toth, E.A., 95, 102–105, 261 Tougu, K., 20, 38–39, 65, 121–122, 216–217 Toyoda, H., 309 Trainor, C., 202 Trakselis, M.A., 59–60, 104–105, 113–114, 198 Tran, N.Q., 91–92, 108–113, 115 Trifunovic, A., 259–261, 273–275 Tsai, K.L., 12f, 14 Tsai, M.-D., 294, 298 Tsai, Y., 100 Tsodikov, O.V., 37 Tsubota, T., 215 Tsuchihashi, Z., 34t, 55 Tsukihara, T., 41–42 Tsurimoto, T., 232, 238 Tsuruta, H., 103 Tsutakawa, S.E., 240 Tummalapalli, P., 100–101, 112–113 Tumpey, T.M., 308 Turkenburg, J.P., 13–14 Turner, J., 34t, 45–46, 51, 55–60, 64–65, 121–122, 208, 216–217 Turner, K.M., 46–49, 58 Turner, S.W., 10 Tye, B.K., 199–200

U Uchida, A., 298, 300–301, 302t Ueda, T., 4, 11–13, 16, 34t Ulrich, H.D., 232–233, 236, 239 Um, S.J., 18

349

Author Index

Umar, A., 232, 245 Unk, I., 241 Urathamakul, T., 34t, 38, 44–48, 51–52 Urbanke, C., 34t, 42, 57 Urlacher, T.M., 38–39 Urmoneit, B., 4, 8–9

V Valentine, A.M., 109–110, 121–122, 219 Vallenet, D., 58 Valpuesta, J.M., 144–145 van der Vliet, P.C., 153–155 van Deursen, F., 200–201, 203–205, 218–219 van Laar, T., 68 van Oijen, A.M., 32–33, 43–45, 55, 61, 63, 65, 69, 95, 97–103, 110, 112–117, 119, 121–124, 192, 197–198, 200, 212–218 van Ranst, M., 149 Vande Berg, B.J., 299 VanLoock, M.S., 14 Veaute, X., 239 Vela´zquez, D., 149 Velten, M., 13–14 ˇ ., 49 Venclovas, C Venkatasubban, K.S., 300–301 Venkitaraman, A.R., 103 Venkova-Canova, T., 6f, 9 Ventura, N., 273–275 Verdaguer, N., 311 Verdine, G.L., 295–296, 299 Verdun, R.E., 138 Vicente, M., 13, 16 Vignuzzi, M., 304–309, 308t Vihinen, M., 202 Vijayakumar, S., 234f Villar, L., 142t, 149–156, 160–161 Vinograd, J., 277, 280–282 Vino´s, J., 152 Vin˜uela, E., 138–139 Vogel, D., 207 Volkmann, N., 103 von Hippel, P.H., 37, 60, 107, 109–110, 197–198, 218 von Meyenburg, K., 2–5 Voortman, L., 34t Vora, M., 3–4

W Waga, S., 203, 205, 208–209, 232 Wagner, G., 95, 102–103, 109–110, 115–119 Wagner, J., 46–48 Wahle, E., 15–18 Waksman, G., 34t, 41–42, 42f, 108–109, 147–148 Walder, J.A., 68 Walder, R.Y., 68 Waldman, V., 42 Waldor, M.K., 9 Walker, G.C., 34t Walker, J.E., 58–59 Walker, J.R., 55, 218 Wallace, D.C., 256 Wallen, J.R., 117 Walter, J.C., 174, 197–198, 200, 212–215, 232, 237 Walther, D.S., 95, 108–110 Waltho, J.P., 40 Walz, T., 102–105 Wang, B., 311, 312f Wang, B.C., 295–296 Wang, D., 299 Wang, D.C., 219 Wang, E.D., 92, 100 Wang, G., 37, 197–198 Wang, H., 146, 148–149 Wang, J., 14–15, 49, 54, 66–67, 141–145, 147–152, 155–156, 295–296, 299 Wang, J.C., 38–39 Wang, M.D., 108, 197–198 Wang, R., 308 Wang, S., 10 Wang, T.S., 201–202, 238 Wang, X., 197–198 Wang, Y., 41–43, 48, 219, 264, 269–271 Wang, Z., 198 Wanrooij, P.H., 278–279 Wanrooij, S., 94, 102, 261–263, 272, 280–281 Warbrick, E., 232, 235 Wargachuk, R., 9–10 Warner, M.D., 185–186 Warrington, J.A., 104–105 Washington, M.T., 105–107, 232–248 Watanabe, K., 239–240 Watson, J.D., 43

350 Watt, S.J., 38 Weakliem, P., 313 Weber, I.T., 103 Weber, P.C., 295–296 Wechsler, J.A., 16–18, 53 Weeks, S.A., 301, 308–309 Wei, T.F., 41–42 Weigel, C., 3–4, 8, 10–11, 16 Weigelt, J., 34t, 37–38 Weiland, E., 42 Weiner, B.E., 201–202 Weisberg, R.A., 8–9 Weitze, T.F., 58–59 Weitzel, S.E., 197–198, 218 Welch, J.L., 301, 303–304, 313–315 Welzeck, M., 11 Wernette, C.M., 260–261, 267–269 West, S.C., 103 Westover, K.D., 299 Whitaker, R.D., 34t White, J.H., 93–95 Whiteheart, S.W., 185–186 Whitmer, J.D., 16–18 Whittell, L.R., 48 Wickner, S., 15, 34t Wickner, W., 45–46 Wieczorek, A., 49, 52 Wiedemann, C., 182, 183f, 184 Wigley, D.B., 14–15, 20, 40, 95, 102–107, 120–121, 193–194, 196–197 Wijffels, G., 34t, 45–48, 51, 57 Wilce, M.C.J., 34t, 37–40, 45–46 Wild, R., 104–105, 120–121 Wilkinson, A.J., 13–14, 185–186 Willcox, S., 108–109 Williams, C.R., 54–55, 57–58 Williams, G.T., 202 Williams, J.S., 205–206 Williams, K.R., 34t, 41–42, 109–110 Williams, M.C., 49 Williams, N.K., 34t, 38, 49, 50f, 55 Williams, R.L., 295–296 Williams, T.A., 170 Wilson, A.C., 256 Wilson, R.J., 102 Wilson, S., 311 Wilson, S.H., 299, 311 Wing, J., 239–240

Author Index

Wing, R.A., 14–15, 49–53 Winnacher, E.L., 8–9 Winter, J.A., 46–48 Winzeler, E.A., 193 Withers, H.L., 16–18 Withers, R., 46–49, 58 Witte, G., 42, 57–58 Wittenberg, C., 206 Wittschieben, B.O., 207 Wittschieben, J.P., 207 Wodicka, L., 193 Woelker, B., 4 Woese, C.R., 170 Wolanski, M., 9 Wold, M.S., 232 Wold, S., 8–9, 16–18 Wolf, Y.I., 265 Wolff, P., 48 Wolfson, J., 93 Wolstenholme, D.R., 274f, 275 Wong, I., 112, 297–299, 304–306 Wong, T.W., 280–281 Woo, H.J., 311 Wood, R.D., 66–67, 207 Woodgate, R., 219, 239 Woods, R.J., 311, 312f Wootton, J.C., 52–53 Worcel, A., 2–3 Wright, A., 36 Wright, C.T., 277 Wright, M., 34t Wu, C.A., 61, 65, 121–122 Wu, X., 149 Wu, Z., 175, 177 Wurgler, S.M., 91

X Xiao, H., 34t, 55–57 Xiao, Y., 307–309, 308t Xie, S., 184 Xie, X.S., 63, 95, 99, 112, 115–117, 119, 123–124 Xiong, Y., 232 Xu, R.M., 194–196 Xu, Y., 149 Xu, Z.-Q., 44, 68 Xuong, N.G., 66, 96–97, 112–113, 295–296, 299

351

Author Index

Y Yagi, H., 34t, 49, 52–53 Yagura, M., 203–205 Yakubovskaya, E., 267–269 Yamagata, Y., 49 Yamamoto, K., 67 Yang, C., 265, 266f Yang, G., 49 Yang, H., 175 Yang, J., 113–114, 121–122, 124, 216–217 Yang, J.Y., 34t, 51–53 Yang, K., 242 Yang, L., 311 Yang, M., 199–200 Yang, M.Y., 281 Yang, S., 14, 103 Yang, W., 49, 193–194, 207–208, 219 Yang, X., 301, 303–304, 313–317 Yang, X.M., 99 Yaniv, J.R., 11–13 Yao, N., 45–46 Yao, N.Y., 20, 44–45, 48–49, 64–65, 121–122, 193–194, 210–218 Yardimci, H., 174, 197–198, 200, 212–215 Yasuda, S., 2–4, 14 Yasukawa, T., 273–275, 278–279, 281 Yasuoka, N., 41–42 Yeeles, J.T., 68, 196, 243 Yelent, B., 103 Yin, Y.W., 100, 259–260, 267–269, 295–296, 299 Yin, Z., 46–48 Yoda, K., 38–40 Yoder, B.L., 60 Yokoyama, S., 3–4, 8, 11, 103 Yoshikawa, H., 6f, 10, 67–68, 138–139 Yoshimura, M., 13–14 You, Z., 212 Young, M.C., 34t, 45–48, 59–60 Young, M.J., 259–260 Yu, C., 205–206 Yu, D.S., 103 Yu, M., 108–109 Yu, X., 14, 37–38, 103–107, 120–121, 197–198, 311 Yu, Y., 110

Yuan, Q., 42, 65–66, 216–217 Yuan, Q.P., 102 Yuan, Z., 200, 210–215, 214f, 218 Yung, B.Y., 8 Yura, T., 2–3 Yurieva, O., 34t, 45–49, 59–60, 64–65, 113–114, 121–122, 194, 210–215, 218 Yuzhakov, A., 34t, 42, 57–58, 64

Z Zaballos, A., 140, 152, 159–160 Zahn, G., 36 Zakrzewska-Czerwinska, J., 9–10, 13–14 Zamyatkin, D.F., 303–304 Zawilak-Pawlik, A., 9–14 Zechel, K., 38–39 Zechner, E.L., 61, 65, 121–122 Zeng, H., 308 Zengel, J.M., 13 Zerbe, L.K., 201–202 Zeviani, M., 102 Zhai, Y., 199–200 Zhang, D., 192–220 Zhang, G., 296–297 Zhang, H., 101–102, 108–113, 115, 232, 240–241 Zhang, J., 41–42, 67 Zhang, S.S., 201 Zhang, X., 93, 95 Zhang, X.-E., 48 Zhang, Y., 37, 199–200 Zhang, Z., 205–206, 232, 247 Zhao, Y., 34t, 45–49, 59–60, 301, 308–309 Zheng, L., 282–283 Zhou, B., 10 Zhou, C., 219 Zhou, J.C., 203–205 Zhou, R., 41–42 Zhou, Z.X., 205–206 Zhu, B., 92, 100, 108–113, 115, 117–119 Zhuang, Z., 60, 63, 113–114 Ziebarth, T.D., 263 Zlotkin, T., 238 Zong, Q., 149 Zweiger, G., 10 Zyskind, J.W., 4, 9

SUBJECT INDEX Note: Page numbers followed by “f ” indicate figures, and “t” indicate tables.

A Accessory-interacting determinant (AID) subdomain, 259–260, 267–269 Acinetobacter baylyi, 58 Active-site fidelity checkpoint, 307–308 Aeropyrum pernix, DNA replication, 177–178, 179f Animal phylogenetic distribution, 267–269, 273 Aquifex aeolicus DnaA, 7–8 DnaB, 14–15 DnaC, 16 Archaeal DNA replication eukaryotic model systems, 171–174 helicase recruitment, 174 initiator proteins, 170–171, 176–177 MCM ATP in, 181–182 cartoon model for, 183f ORC1–1, 182–186, 185f structure of, 183f Orc1/Cdc6 proteins, 177–181 origin specification, 175–177 prereplicative complex proteins, 173f Pyrococcus, 170–171 revolution, 170 Arginine finger, 105, 106f ATP, in MCM loading, 181–182

B Bacillus subtilis, 6f, 158–160 replication origins, 10 SirA, 13–14 Bacterial replication origin assembly of replisome, 20 B. subtilis, 10 C. crescentus, 9–10 DnaA domain I interacts, 11–14, 12f replication initiation, 11 DnaB, 14–15

DnaB–DnaC complex formation, 15 helicase activation, 19–20 prepriming complex, 16–19, 17f E. coli bacterial growth and cell division, 2–3 DnaA boxes, 3–4 dnaC(Ts) mutant, 2–3 DNA motifs, 4–5, 6f DNA unwinding element, 7–8 DUE, 8 Fis and IHF, 8–9 helicase loading, 16, 19 H. pylori oriC, 10 V. cholerae oriC, 9 Bacterial replisome, 192, 215–219, 216f Bacteriophage T7, DNA replication chromosome, initiation, 93–94, 94f concatemers, 95 discrimination against ddNTPs, 97–99 dsDNA unwinding, 107–108 Escherichia coli, 90 exchange of polymerases, 113–115 gene 1.7, 91–92 gene 4 protein, 93 gene 5 protein, 93 gene 5.5, 91–92 genetic map, 91f gp2.5 (see Gene 2.5 protein (gp2.5)) gp4 oligomerization, 102–103 human mitochondrial and, 90 intermediates, 95 minimal, 95–110, 96f nucleotide hydrolysis, 105–107 nucleotide polymerization, 96–97 polymerase-lysozyme complex, 93 primase-helicase, 100–108, 101f proteins in, 91–93 ring-opening model, 104f single-stranded protein, 108–110 ssDNA, 103–105 trombone model of, 119 353

354 Bacteriophage T7, DNA replication (Continued ) trx binding, 99–100 β2 sliding clamp, 45–48, 47f Bootlace strategy, 280–283 Break-induced replication, PCNA, 243–245, 244f

C Caenorhabditis elegans, rolling circle model, 273–275 Catalysis enzymic catalysis, 299 general acid catalysis, 299–304 two-metal-ion mechanism, 296–297, 297f Chromatin association factor-1 (CAF-1), 246–247 Clamp-binding motifs (CBMs), 46–52 Clamp loader complex (CLC), 33f, 54–60, 56f CMG (Cdc45, RecJ, and GINS complex) helicase, 194–201, 197f Drosophila melanogaster, 199f, 200 formation and activation, 199–200 function, 210–215 GINS component, 204f leading strand, 203–205, 218 Pol alpha functions with, 212–215 in Xenopus, 198 CMG–Pol epsilon (CMGE), 210–215, 213f Collision model, E. coli replisome, 64–65 Conserved sequence blocks (CSBs), 277 Cryptic cytosine, 115–119 C-terminal helicase domain (CTD), 261 C-terminal segment of ε (εCTS), 51–52

D

20 , 30 -Dideoxynucleoside triphosphate (ddNTP), 96–97 discrimination against, 97–99, 98f Dideoxythymidylate (ddTMP), 91–92 DnaA at oriC domain I interacts, 11–14, 12f replication initiation, 11 DNA amplification, 158 DnaB, 14–15, 36–40, 39f DnaB–DnaC complex

Subject Index

formation, 15 helicase activation, 19–20 prepriming complex, 16–19, 17f DnaB helicase, 33f, 62, 215–217 DNA-dependent DNA polymerase (DdDp), 297–298 DnaG, 36–40 DNAP heterodimer, 152 DNA polymerase-binding protein 2 (Dpb2), 207 DNA polymerase γ, 257–261, 258f DNA polymerase III holoenzyme (Pol III HE), 33f assembly and action, 44–45 β2 clamp, 45–48, 47f clamp loader complex, 54–60, 56f dnaQ49 gene, 53 holE gene, 53 αPHP domain, 52–53 Pol III core (aεθ), 48–54, 50f DNA polymerases, 232 exchange of, 113–115 replicative, 241 translesion synthesis, 239 DNA primase-helicase, 100–108, 101f DNA-primed elongation, 156–158, 157f DNA replication, bacteriophage T7 chromosome, initiation, 93–94, 94f concatemers, 95 discrimination against ddNTPs, 97–99 dsDNA unwinding, 107–108 Escherichia coli, 90 exchange of polymerases, 113–115 gene 1.7, 91–92 gene 4 protein, 93 gene 5 protein, 93 gene 5.5, 91–92 genetic map, 91f gp2.5 (see Gene 2.5 protein (gp2.5)) gp4 oligomerization, 102–103 human mitochondrial and, 90 intermediates, 95 minimal, 95–110, 96f nucleotide hydrolysis, 105–107 nucleotide polymerization, 96–97 polymerase-lysozyme complex, 93 primase-helicase, 100–108, 101f proteins in, 91–93

355

Subject Index

ring-opening model, 104f single-stranded protein, 108–110 ssDNA, 103–105 trombone model of, 119 trx binding, 99–100 Drosophila melanogaster mtDNA helicase, 261–263 theta model in, 275–276 ZBD, 263–264 dsDNA, unwinding by gp4, 107–108 Dynamics-function relationship, 299, 303–304

E Enzymic catalysis, 299 Error-free damage bypass, PCNA, 241–243, 242f Escherichia coli deoxyguanosine triphosphatase (dGTPase), 91–92 Escherichia coli DNA replication DnaB, 36–40, 39f DnaG, 36–40 elongation stage, 43 [3H]-thymidine-labeled E. coli chromosome, 32 lagging strand collision model, 64–65 dynamics, 60–61 Okazaki fragment maturation, 66–68 priming, 62–64 signaling model, 65–66 leading strand DNA Pol III HE, 44, 60–61 dsDNA, 62 Okazaki fragments, 32 at oriC, 40 primer, 37 synthesis, 57–58 oriC, 36 bacterial growth and cell division, 2–3 DnaA boxes, 3–4 dnaC(Ts) mutant, 2–3 DNA motifs, 4–5, 6f DNA unwinding element, 7–8 DUE, 8 Fis and IHF, 8–9 Pol III HE, 33f assembly and action, 44–45

β2 clamp, 45–48, 47f clamp loader complex, 54–60, 56f dnaQ49 gene, 53 holE gene, 53 αPHP domain, 52–53 Pol III core (aεθ), 48–54, 50f primosome, 36–40 replisomes, 32, 33f, 34t, 36 SSB role in, 41–43 trombone model, 43 Eukaryotic DNA replication, PCNA in, 232–233, 247–248 break-induced replication, 243–245, 244f error-free damage bypass, 241–243, 242f function, 233–236 mismatch repair, 245–246 in normal DNA replication, 236–239 processivity factor, 232 replication-coupled nucleosome assembly, 246–247 separation-of-function mutations, 241f structure, 233–236, 234f in translesion synthesis, 239–241 Eukaryotic replisome architecture of, 214f bacterial and, 192, 215–219, 216f CMG helicase, 194–201, 197f core components of, 192–194, 195t DNA damage, 219 DNA polymerases, 193–194 Drosophila, EM structure f, 200 genome sequencing, 193–194 leading and lagging strand synthesis, 205–208 Mcm2–7, 199–200, 199f PCNA/RFC clamp loader, 208–210 polymerase alpha-primase, 201–205, 203–204f reconstitution, 211f ring-shaped helicases, 197–198 structure and function, 210–215 Evolutionary Trace Server, 265, 266f

F Factor for inversion stimulation (Fis), 8–9 Φ29 DNA replication protein-primed mechanism binary complex structures, 146f

356 Φ29 DNA replication (Continued ) biotechnological applications, 149 DNA amplification, 158 DNAP heterodimer, 152 DNA-primed elongation, 156–158, 157f DNA synthesis coupling to strand displacement, 140–145 initiation, 139–152, 139f N-terminal and C-terminal motifs, 140–141, 142t polymerization and proofreading activities, 147–149, 148f processivity factors, 141, 149, 157f schematic representation, 144f sliding-back mechanism, 153–155, 154f three-dimensional structures, 143f TP, 149–152, 151f TP heterodimer, 152 transition, 155–156, 156f translocation mechanism, 145–146 in vivo compartmentalization, 158–161, 160f Full-length flap endonuclease 1 (FEN1), 235–236

G Gene 2.5 protein (gp2.5) binding to ssDNA, 109–110 C-terminal tail, 109–110 interactions with replication proteins, 110 Gene 4 protein (Gp4) C-terminal tail, 102–103 dsDNA unwinding by, 107–108 interactions in leading-strand synthesis, 110–113, 111f 56-kDa protein, 101–102 63-kDa protein, 101–102 loading to ssDNA, 103–105 lock-washer conformation, 104–105 N-terminal helix, 102–103 nucleotide-binding sites, 105, 106f, 107 oligomerization of, 102–103 primer synthesis by, 116f sedimentation velocity, 104–105 General acid catalysis, 299–304 Geobacillus stearothermophilus, 38–39

Subject Index

Gp5 crystal structures of, 100 interactions in leading-strand synthesis, 110–113, 111f

H Haloferax mediterranei, 175 Haloferax volcanii, 175 Helicobacter pylori, 13–14 H-NS protein, 92 holE gene, 53–54 Holliday junction, 243–244

I Integration host factor (IHF), 8–9 Inverted terminal repetitions (ITR), 138

K Kinetic mechanism, RdRp, 297–298, 298f, 304f

L Lagging-strand replication collision model, 64–65 coordination, 123–124 dynamics, 60–61 eukaryotes, 205–208 formation and release, 119–122 Okazaki fragment maturation, 61, 66–68 oligoribonucleotide, 115–117 primer synthesis by gp4, 116f priming, 62–64 release of replication loop, 121f RNA primer handoff, 117–119 signaling model, 65–66 Leading-strand replication DNA Pol III HE, 44, 60–61 dsDNA, 62 eukaryotes, 205–208 gp4-gp5/trx interactions in, 110–113 Okazaki fragments, 32 at oriC, 40 primer, 37 synthesis, 57–58 Light-strand transcriptional promoter (LSP), 278–279

357

Subject Index

M MCM ATP in, 181–182 cartoon model for, 183f ORC1–1, 182–186, 185f structure of, 183f Methanopyrus kandleri, 170 Mismatch repair, PCNA, 245–246 Mitochondrial DNA (mtDNA) replication C. elegans, rolling circle model, 273–275 invertebrate, 274f structure–function relationships, 256 DNA polymerase γ, 257–261, 258f Evolutionary Trace Server, 265, 266f four-helix bundle structure, 268f functional interactions, 265–273, 270f mtSSB, 264–265 oligomerization, 261, 263 proteins, 257f replicative mtDNA helicase, 261–264, 262f theta model D. melanogaster, 275–276 sea urchin, 276 vertebrates, 277–283 Mitochondrial helicase (Twinkle), 102 Mitochondrial nucleic acids (mtNA), 273–275 Mitochondrial polymerase (Pol-γA), 100 Mitochondrial single-stranded DNAbinding protein (mtSSB), 256, 264–265 Mitochondrial transcription elongation factor (TEFM), 278–279 Molecular dynamics protocol, 312f mtDNA replication. See Mitochondrial DNA (mtDNA) replication mtSSB. See Mitochondrial single-stranded DNA-binding protein (mtSSB) Multiple displacement amplification (MDA), 149 Mycobacterium tuberculosis β2 clamp, 48

N

N-terminal domain of ε, 51–52 Nucleic acid polymerase, 294, 299–304

Nucleoside analog reverse transcriptase inhibitors (NRTIs), 259–260 Nucleotide-addition cycle, 297–298 Nucleotide hydrolysis coupling, 105–107 DnaB, 14–15 duplex DNA, 197–198 Nucleotide incorporation fidelity, RdRp, 311–313 Nucleotide polymerization, 96–97, 97f Nucleotidyl transfer, 297f

O Okazaki fragments E. coli DNA replication fork, 61, 66–68 PCNA, 238–239, 246, 248 Oligoribonucleotides, 100–101 Origin of replication (oriC), 36 DnaA domain I interacts, 11–14, 12f replication initiation, 11 E. coli, 36 bacterial growth and cell division, 2–3 DnaA boxes, 3–4 dnaC(Ts) mutant, 2–3 DNA motifs, 4–5, 6f DNA unwinding element, 7–8 DUE, 8 Fis and IHF, 8–9 H. pylori, 10 leading-strand replication at, 40 V. cholerae, 9 Origin recognition complex (ORC) in eukaryotes, 171–174 MCM recruitment by, 182–186, 185f Orc1/Cdc6 proteins, 177–181 prereplicative complex proteins, 173f

P PCNA clamp loader, 208–210, 209f PCNA-interacting protein (PIP), 235–236 Phosphorothioate elemental (thio) effect, 298 Phosphoryl transfer, RdRp, 297–298 Plasmodium falciparum, 102 Pol III core (aεθ), 48–54, 50f Pol III HE. See DNA polymerase III holoenzyme (Pol III HE)

358 Polymerase fidelity, 298–299 Polymerase structure, RdRp, 295–296, 295f Posttranslational modifications, PCNA, 232–233, 236, 239 Primosome, 36–40 Principal component analysis (PCA), 312–313 Proliferating cell nuclear antigen (PCNA), 48, 232–233, 247–248 break-induced replication, 243–245, 244f error-free damage bypass, 241–243, 242f full-length flap endonuclease 1, 235–236 function, 233–236 mismatch repair, 245–246 in normal DNA replication, 236–239 posttranslational modifications, 236 processivity factor, 232 replication-coupled nucleosome assembly, 246–247 separation-of-function mutations, 241f structure, 233–236, 234f sumoylation, 232–233, 236, 239 in translesion synthesis, 239–241 ubiquitylation, 232–233, 236 Protein-nucleic acid complex, 117 Protein-primed replication, ϕ29 DNA binary complex structures, 146f biotechnological applications, 149 DNA amplification, 158 DNAP heterodimer, 152 DNA-primed elongation, 156–158, 157f DNA synthesis coupling to strand displacement, 140–145 initiation, 139–152, 139f N-terminal and C-terminal motifs, 140–141, 142t polymerization and proofreading activities, 147–149, 148f schematic representation, 144f sliding-back mechanism, 153–155, 154f three-dimensional structures, 143f TP, 149–152, 151f TP heterodimer, 152 transition, 155–156, 156f translocation mechanism, 145–146 Pseudomonas aeruginosa, 58 PV RdRp fidelity, 304–306, 316f Pyrobaculum aerophilum, 177–178

Subject Index

Pyrococcus abyssi, 175 Pyrococcus furiosus RadA protein (PfRad51), 103

R RecA protein, 103 Remote-site control, RdRp, 307–308 Replication factor C (RFC), 232, 238 Replication forks, 32 Reverse transcriptase (RT), 297–298 RFC clamp loader, 208–210, 209f Ribonucleotides incorporated through out the lagging strand (RITOLS), 280–283 RNA population, 307f, 316–317 primer handoff, 117–119, 118f RNA-dependent RNA polymerase (RdRp) determinant of nucleotide incorporation fidelity, 311–313 fidelity as determinant of viral virulence, 308 fidelity mutants, 308–310 incorporation fidelity, 316–317 RdRp-RNA complex, 313–316 RNA polymerase domain (RPD), 100–101, 112–113

S Saccharomyces cerevisiae, 195t, 267–269 Sea urchins, theta model in, 276 Signaling model, E. coli replisome, 65–66 Simian virus 40 (SV40) replication, 205–206 Single-nucleotide incorporation, 297–298, 298f, 304f Single-stranded DNA (ssDNA), 138 gp4 loading to, 103–105 nucleotide hydrolysis and, 105–107 Single-stranded DNA-binding protein (SSB), 95, 96f, 108–110, 108f EcSSB, 265 in replication fork, 41–43 structure of, 42f Sliding-back mechanism, ϕ29 DNA replication, 153–155, 154f Sliding clamps, 232–233 Small-angle X-ray scattering (SAXS) model, 112

359

Subject Index

ssDNA. See Single-stranded DNA (ssDNA) Strand-coupled replication, 280–283 Strand displacement DNA synthesis coupling to, 140–145 mtDNA replication, 280–283 Strongylocentrotus purpuratus, 268f, 276 Structural dynamics, RdRp, 316–317, 316–317f Sulfolobus acidocaldarius, 176 Sulfolobus islandicus, 176–177 Sulfolobus solfataricus, 103, 170–171 Surface plasmon resonance (SPR), 112–113

Twinkle, 102 Two-dimensional agarose gel electrophoresis (2DAGE), 273–275 Tyrosine (Tyr) 526 enzymatic property of, 97–99 phenylalanine substitution of, 97–99 Tyr254/Tyr390, 145

T

Vertebrate mtDNA replication bootlace, 280–283 conserved sequence blocks, 277 leading-strand synthesis, 277 light-strand transcriptional promoter, 278–279 RITOLS, 280–283 strand-coupled models, 280–283 structural organization, 278f Viral virulence, RdRp, 308

TBD. See Thioredoxin-binding domain (TBD) TEFM. See Mitochondrial transcription elongation factor (TEFM) Terminal protein (TP) heterodimer, 152 heterologous systems, 150–151 independent expression, 151–152 N-terminal domain, 149–150 parental TP, 149–150 placement, 151f Thermus aquaticus α, 49–51 Thioredoxin A (trx) binding of, 99–100 interactions in leading-strand synthesis, 110–113, 111f Thioredoxin-binding domain (TBD), 99 TP. See Terminal protein (TP) TPR1/TPR2, 141–144 Translesion synthesis, PCNA, 239–241 Trombone model, 43

U Ubiquitin-modified PCNA (Ub-PCNA), 239–242

V

X Xenopus laevis, 200, 212–215 CMG, 198 D-loop region, 277

Z Zinc-binding domain (ZBD) in binding to gp5/trx, 112–113 DNA primase-helicase, 100–101 interaction with RPD, 115–117 RNA primer handoff, 117–119